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Endocrinology, doi:10.1210/en.2005-1304
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Endocrinology Vol. 147, No. 4 1685-1696
Copyright © 2006 by The Endocrine Society

The Transcription Factor AP-2ß Causes Cell Enlargement and Insulin Resistance in 3T3-L1 Adipocytes

Yukari Tao, Hiroshi Maegawa, Satoshi Ugi, Kazuhiro Ikeda, Yoshio Nagai, Katsuya Egawa, Takaaki Nakamura, Shuichi Tsukada, Yoshihiko Nishio, Shiro Maeda and Atsunori Kashiwagi

Division of Endocrinology and Metabolism, Department of Medicine (Y.T., H.M., S.U., K.I., Y.Na., K.E., Y.N.i., A.K.), and Department of Anatomy (T.N.), Shiga University of Medical Science, Otsu, Shiga 520-2192, Japan; and Laboratory for Diabetic Nephropathy, SNP Research Center, Institute of Physical and Chemical Research (S.T., S.M.), Kanagawa 230-0045 Japan

Address all correspondence and requests for reprints to: Dr. Hiroshi Maegawa, Division of Endocrinology and Metabolism, Department of Medicine, Shiga University of Medical Science, Seta, Otsu, Shiga 520-2192, Japan. E-mail: maegawa{at}belle.shiga-med.ac.jp.


   Abstract
Top
 Abstract
Introduction
 Materials and Methods
 Results
 Discussion
 References
 
We have reported the association of variations in the activating protein-2ß (AP-2ß) transcription factor gene with type 2 diabetes. This gene was preferentially expressed in 3T3-L1 adipocytes in a differentiation stage-dependent manner, and preliminary experiments showed that subjects with the disease-susceptible allele showed stronger expression in adipose tissue than those without the susceptible allele. Thus, we overexpressed the AP-2ß gene in 3T3-L1 adipocytes to clarify whether AP-2ß might play a crucial role in the pathogenesis of type 2 diabetes through dysregulation of adipocyte function. In cells overexpressing AP-2ß, cells increased in size by accumulation of triglycerides accompanied by enhanced glucose uptake. On the contrary, suppression of AP-2ß expression by small interfering RNA inhibited glucose uptake. Enhancement of glucose uptake by AP-2ß overexpression was attenuated by inhibitors of phospholipase C (PLC) and atypical protein kinase C{zeta}/{lambda} (PKC{zeta}/{lambda}), but not by a phosphatidylinositol 3-kinase (PI3-K) inhibitor. Consistently, we found activation of PLC and atypical PKC, but not PI3-K, by AP-2ß expression. Furthermore, overexpression of PLC{gamma} enhanced glucose uptake, and this activation was inhibited by an atypical PKC inhibitor, suggesting that the enhanced glucose uptake may be mediated through PLC and atypical PKC{zeta}/{lambda}, but not PI3-K. Moreover, we observed the increased tyrosine phosphorylation of Grb2-associated binder-1 (Gab1) and its association with PLC{gamma}, indicating that Gab1 may be involved in AP-2ß-induced PLC{gamma} activation. Finally, AP-2ß overexpression was found to relate to the impaired insulin signaling. We propose that AP-2ß is a candidate gene for producing adipocyte hypertrophy and may relate to the abnormal characteristics of adipocytes observed in obesity.


   Introduction
Top
 Abstract
 Introduction
Materials and Methods
 Results
 Discussion
 References
 
ADIPOSE TISSUE IS recognized as an important source of metabolically active secretory factors, including free fatty acids, leptin, TNF-{alpha}, IL-6, plasminogen activator inhibitor-1, resistin, visfatin, and adiponectin (1). Although small adipocytes secrete the insulin-sensitizing hormone, adiponectin, adipocytes with increased cell size are associated with a decrease in the expression and an increase in the levels of insulin-resistant hormones, resulting in the insulin resistance observed in obesity (2). These phenomena are considered to have crucial roles in the pathogenesis of obesity-related diseases such as metabolic syndrome and type 2 diabetes.

We have recently identified the activating protein-2ß (AP-2ß) transcription factor gene (TFAP2B) located on chromosome 6p12 as a susceptibility gene for type 2 diabetes by conducting a genome-wide association study (3). Several variations in the TFAP2B gene were shown to be significantly associated with obese type 2 diabetes in a Japanese population as well as in a United Kingdom population (3). Furthermore, we discovered that mouse AP-2ß (Tcfap2b) is preferentially expressed in adipose tissue, and its expression is increased during adipocyte differentiation in 3T3-L1 adipocytes (3). Moreover, the preliminary experiments showed that polymorphism in the first intron of TFAP2B may directly affect the transcriptional activity of the gene. In fact, subjects with the disease-susceptible allele have stronger expression of AP-2ß in adipose tissue than those without the susceptible allele (Tsukada, S., Y. Tanaka, H. Maegawa, A. Kashiwagi, R. Kawamori, and S. Maeda, unpublished observations). These results suggest that TFAP2B may play an important role in the pathogenesis of type 2 diabetes through the dysregulation of adipocyte function, and the polymorphisms within TFAP2B may affect the expression of the gene, which thus confers susceptibility to the disease.

AP-2ß has been reported to play an important role in embryonic development (4, 5, 6, 7). Mice lacking AP-2ß die within 1 or 2 d after birth from renal failure due to polycystic kidney disease through enhanced apoptotic renal epithelial cell death (6). In humans, mutations of TFAP2B cause the Char syndrome, characterized by patent ductus arteriosus, variable degrees of facial dimorphism, and hand abnormalities (4, 5, 8). Those findings suggest that AP-2ß plays an important role in the embryonic development of various tissues.

The AP-2 transcription factor family consists of five members, AP-2{alpha}, AP-2ß, AP-2{gamma}, AP-2{delta}, and AP-2{epsilon}, each encoded by a separate gene (9, 10, 11, 12, 13). AP-2 proteins homo- and heterodimerize through a unique C-terminal helix-span-helix motif and bind palindromic DNA recognition sequences (consensus, 5'-GCCN3GGC-3') through the basic domain that lies immediately N terminal of the dimerization motif (11). AP-2{alpha} has been reported to inhibit adipogenesis through inhibition of CCAAT/enhancer-binding protein-{alpha} (C/EBP{alpha}), and the expression of AP-2{alpha} was down-regulated upon induction of differentiation in 3T3-L1 adipocytes (14). However, to date, no report other than our recent paper (3) has emerged to suggest that AP-2ß has a role in adipocyte dysfunction and the pathogenesis of type 2 diabetes.

In the present study we assessed the role of AP-2ß in adipocytes. The results demonstrated that overexpression of AP-2ß leads not only to lipid accumulation, but also to insulin resistance in 3T3-L1 adipocytes. Thus, we propose that AP-2ß is the candidate gene for producing to hypertrophy and dysfunction of adipocyte in obesity.


   Materials and Methods
Top
 Abstract
 Introduction
 Materials and Methods
Results
 Discussion
 References
 
Materials
Human insulin was provided by Eli Lilly & Co. (Indianapolis, IN). Antiphospho-Akt Ser473 antibody, antiphosphoinsulin receptor (antiphospho-IR) antibody, and anti-Akt antibody were obtained from Cell Signaling Technology (Beverly, MA). Antiphospholipase C{gamma}-1 (anti-PLC{gamma}-1) antibody and anti-IR substrate-1 (IRS-1) antibody were obtained from Upstate Biotechnology, Inc. (Lake Placid, NY). Anti-AP-2ß antibody; anti-IRß antibody; antiprotein kinase C{zeta}/{lambda} (anti-PKC{zeta}/{lambda}) antibody; anti-Grb2-associated binder-1 (anti-Gab1) antibody; horseradish peroxidase-linked antirabbit, antigoat, and antimouse antibodies; and protein A/G-agarose were purchased from Santa Cruz Biotechnology (Santa Cruz, CA). Antiphosphotyrosine antibody (RC20) and anti-p85 subunit of phosphatidylinositol 3-kinase (PI3-K) antibody were purchased from BD Biosciences (San Jose, CA). Antiglucose transporter 4 (anti-GLUT4) antibody (1F8) was obtained from Biogenesis, Inc. (Brentwood, NH), and anti-GLUT1 antibody was obtained from Chemicon International, Inc. (Temecula, CA). U73122 and SB203580 were purchased from Calbiochem (San Diego, CA). A myristoylated peptide of the PKC{zeta} pseudosubstrate was purchased from BioSource International (Camarillo, CA). All radioisotopes were obtained from ICN (Costa Mesa, CA). DMEM and fetal calf serum were obtained from Invitrogen Life Technologies, Inc. (Grand Island, NY). XAR-5 film was obtained from Eastman Kodak Co. (Rochester, NY). All other reagents and chemicals were purchased from Sigma-Aldrich Corp. (St. Louis, MO).

Cell culture
3T3-L1 cells, provided by Dr. J. M. Olefsky (University of California, San Diego, CA), were cultured and differentiated as previously described (15). Before each experiment, the adipocytes were trypsinized and reseeded in the appropriate culture dishes. The Ad-E1A-transformed human embryonic kidney cell line, 293 cells, was cultured as described previously (16).

Preparation of recombinant adenovirus and infection
Adenovirus vector encoding human AP-2ß was generated using an Adenovirus Expression Vector Kit (Takara, Shiga, Japan). Adenovirus vector encoding the dominant-negative form of PKC{lambda} (DN-PKC{lambda}) was a gift from Dr. Wataru Ogawa (Kobe University, Kobe, Japan) (17). The plasmid encoding PLC{gamma} was a gift from Dr. Sue Goo Rhee (National Institutes of Health, Bethesda, MD) (18). Adenovirus vector encoding PLC{gamma} was generated by the method previously described (19). LacZ-encoding vector was used for the control. 3T3-L1 adipocytes, on d 10 after induction of differentiation, were infected with adenoviruses at the indicated multiplicity of infection (MOI) for 24 h. Transfected cells were incubated for 48 h at 37 C in an atmosphere of 10% CO2 in high-glucose DMEM with 2% heat-inactivated serum, followed by starvation as required for the assay.

Generation of mutant AP-2ß
Mouse AP-2ß cDNA was subcloned into pcDNA 3.1 (pcDNA 3.1/AP-2ß). A QuikChange site-directed mutagenesis kit (Stratagene, La Jolla, CA) was used for mutagenesis. An oligonucleotide in which arginine 174 of AP-2ß was replaced with cysteine was used as the primer in the in vitro mutagenesis reaction. The resultant mutant AP-2ß cDNA was subcloned into pcDNA 3.1 (pcDNA 3.1/AP-2ß R225C). Arginine 174 is located in the basic domain that is necessary for DNA binding. An R225C mutation has been found in patients with Char syndrome and has been confirmed to extinguish DNA-binding ability (4, 5).

Nuclear extraction
Preparation of nuclear extracts was performed as previously described (20). Briefly, cells were rinsed twice with cold PBS, lightly trypsinized, and pelleted by centrifugation at 3000 rpm for 5 min. The pellet was washed twice with PBS, then suspended in lysis buffer A [10 mM Tris-HCl (pH 7.5), 1.5 mM MgCl2, 10 mM KCl, 1 mM dithiothreitol (DTT), 1 µM phenylmethylsulfonylfluoride (PMSF), 2 µM sodium vanadate, 2 µM leupeptin, 1 µM aprotinin, and 1 µM pepstatin]. The cell suspension was homogenized, and nuclei were pelleted by centrifuging at 10,000 rpm for 5 min. The supernatant was recovered as a cytosol fraction. The pellet was resuspended in buffer C [20 mM Tris-HCl (pH 7.5), 0.42 mM KCl, 20% glycerol, 1.5 mM MgCl2, sodium vanadate, DTT, PMSF, leupeptin, aprotinin, and pepstatin at the concentrations used for buffer A described above]. The lysate was rotated for 30 min at 4 C and centrifuged at 14,000 rpm for 30 min. The supernatant was recovered as the nuclear fraction.

Subcellular fractionation
Cells were washed three times with ice-cold PBS, and scraped into ice-cold HES buffer [225 mM sucrose, 20 mM HEPES (pH 7.4), and 1 mM EDTA] supplemented with 100 mM sodium fluoride, 10 mM sodium pyrophosphate, 1 mM sodium vanadate, 1 µg/ml leupeptin, 1 µg/ml pepstatin, 1 mg/ml benzamidine, and 0.5 mM PMSF. Cells were then homogenized using an LSC homogenizer (Sogorikagaka Grass Works, Kyoto, Japan). Subcellular fractionation was carried out as described previously (21).

Preparation of whole-cell lysates and immunoprecipitation
Cells were lysed in solubilizing buffer [20 mM Tris (pH 7.5), 1 mM EDTA, 140 mM NaCl, 1% Nonidet P-40, 1 mM sodium vanadate, 50 mM sodium fluoride, 1 µM aprotinin, and 1 mM PMSF] for 30 min at 4 C. The cell lysates were centrifuged to remove insoluble materials. For immunoprecipitation, cell lysates were incubated with primary antibody for 5 h at 4 C and with protein A/G-agarose for an additional 2 h. The immunoprecipitates were washed, resuspended in Laemmli sample buffer containing 100 mM DTT, and heated for 5 min at 100 C.

Immunoblotting
Whole-cell lysates and antibody immunoprecipitates were resolved by SDS-PAGE and electrophoretically transferred to polyvinylidene difluoride membranes (Immobilon-P, Millipore Corp., Bedford, MA). Membranes were blocked and probed with the specified antibodies. Blots were then incubated with horseradish peroxidase-linked second antibody, followed by chemiluminescence detection.

Oil red O staining
Infected cells were cultured in medium containing different concentrations of glucose for 88 h. The cells were washed twice with PBS, fixed in 3.7% formaldehyde for 1 h, then stained with 0.6% (wt/vol) oil red O solution (60% isopropanol and 40% water) for 2 h at room temperature. Cells were washed with water to remove unbound dye, then visualized by light microscopy. For quantification, cells in 96-well plates were stained with oil red O, then dye was eluted with isopropanol and quantified by measuring the absorbance at 540 nm.

2-Deoxyglucose uptake
Glucose uptake assay was performed as previously described (15). After 88 h of adenovirus infections, serum- and glucose-deprived cells were incubated in DMEM in the absence or presence of insulin for 60 min at 37 C. Glucose uptake was determined in duplicate at each point after the addition of 10 µl substrate (0.1 µCi 2-[3H]deoxyglucose or L-[3H]glucose; final concentration, 0.01 mM) to provide a concentration at which cell membrane transport was rate limiting. The value for L-glucose was subtracted to correct each sample for the contributions of diffusion and trapping.

PKC{zeta}/{lambda} assay
PKC{zeta}/{lambda} enzyme activity was determined as previously described (15). In brief, starved cells were stimulated with insulin for 30 min at 37 C and lysed in buffer [50 mM Tris (pH 7.5), 150 mM NaCl, 1% Nonidet P-40, 1 mM EGTA, 10 mM 2-glycerol phosphate, 0.1% 2-mercaptoethanol, 1 mM sodium vanadate, 20 mM sodium fluoride, 5 mM sodium pyrophosphate, 0.1 mM PMSF, 10 µg leupeptin/ml, 20 µg aprotinin/ml, and 10 µM microcystin LR]. Clarified supernatants were immunoprecipitated with anti-PKC{zeta}/{lambda} antibody or rabbit IgG and protein A/G-agarose for 4 h at 4 C. Samples were washed twice with the same buffer and twice with kinase assay buffer [50 mM Tris (pH 7.5), 5 mM MgCl2, 1 mM EGTA, 100 µM sodium vanadate, 100 µM sodium pyrophosphate, 1 mM sodium fluoride, and 100 µM PMSF]. Suspensions were then incubated with 4 µCi [{gamma}-32P]ATP, 50 µM ATP, 4 µg phosphatidylserine, and 40 µM [159Ser] PKC (amino acids 153–164)-NH2 for 10 min at 30 C with gentle agitation. Reactions were stopped by the addition of 10 µl 5% acetic acid. Aliquots of the reaction mixtures were spotted on p81 phosphocellulose paper, washed in 5% acetic acid, and quantitated by liquid scintillation counting.

PLC assay
PLC activity was determined as cellular inositol 1,4,5-trisphosphate (IP3) content using the Biotrack assay system (TRK1000, Amersham Biosciences, Piscataway, NJ) according to the manufacturer’s instructions. This assay is based on competition between [3H]IP3 (the tracer) and unlabeled IP3 in standards or samples for binding to a binding protein. Briefly, cells were homogenized in ice-cold 4.2% perchloric acid and centrifuged at 2000 x g for 15 min at 4 C, and the supernatant was quantitatively decanted to a plastic centrifuge tube. The supernatant was neutralized to pH 7.5 with ice-cold 1.8 M KOH containing 60 mM HEPES. KClO4 was sedimented by centrifugation at 2000 x g for 15 min at 4 C, and the supernatant aliquot was quantitatively transferred to a plastic test tube. Bound IP3 was then separated from free IP3 by centrifugation, which sediments the binding protein to the bottom of the tube. The pellet was resuspended in water and quantitated by liquid scintillation counting. Data were evaluated on the basis of percentage binding (bound/free ratio) from the calibration curve.

PI3-K assay
PI3-K enzyme activity was determined as previously described (22). Starved cells were stimulated as described in the figure legends and lysed. Clarified supernatants were immunoprecipitated with anti-Gab1 antibody and protein A/G-agarose for 4 h at 4 C. The washed immunoprecipitants were incubated with phosphatidylinositol (Avanti Polar Lipids, Alabaster, AL) and [{gamma}-32P]ATP (3 mCi/mmol) for 10 min at room temperature. Reactions were stopped with 20 µl 8 N HCl and 160 µl CHCl3/methanol (1:1); the reaction mixtures were centrifuged; and the lower organic phases were removed and applied to a silica gel TLC plate (Merck & Co., Rahway, NJ) that had been coated with 1% potassium oxalate. Thin layer chromatography plates were developed in CHCl3/CH3OH/H2O/NH4OH (60:47:11.3:2), dried, visualized, and quantitated on a PhosphorImager (Amersham Biosciences).

Transfection study
On d 5 after induction of differentiation, cells were trypsinized, and the cell suspension was mixed with 5 µg pcDNA 3.1 empty vector, pcDNA 3.1/AP-2ß, or pcDNA 3.1/AP-2ß (R225C); transferred to a 2-mm electroporation cuvette; and electroporated with a Nucleofector (Amaxa, Cologne, Germany) by using the program U-28. After electroporation, cells were immediately transferred to 1 ml growth medium and cultured in 12-well plates at 37 C. Sample preparation for Western blotting and glucose transport assay was performed after 48 h.

Transfection with small interfering RNAs (siRNAs)
The target sequence for designing the siRNA against AP-2ß was obtained from Hokkaido System Science Co. Ltd. (Hokkaido, Japan), and sequence for scrambled control was designed with four base mutations. Sense and antisense DNA oligonucleotides were inserted into the piGENE mU6 vector. Five micrograms of either expression vector of siRNA against AP-2ß or scrambled control vector was electroporated as described for the transfection study. The target sequences for designing the siRNAs against AP-2ß and scrambled control were as follows: AP-2ß, 5 '-CTACTCAGTTCAACTTCAAAGTACA-3'; and scrambled control, 5'-CTACTCAGCCCAACGGCAAAGTACA-3' (underline indicates mutated bases).

Quantitative real-time PCR
First-strand cDNA was synthesized from total RNA extracted from 3T3-L1 adipocytes infected with LacZ or AP-2ß adenoviruses by oligo(deoxythymidine) priming using SuperScript III reverse transcriptase (Invitrogen Life Technologies, Inc., Carlsbad, CA). Quantitative RT-PCR was performed using SYBR Green detection. The amplifications were carried out in a 25-µl reaction volume containing 1x EX Taq buffer, 200 nM deoxy-NTP mixture, 800 nM of each primer, 1:2000 SYBR Green (for SYBR Green detection), 0.125 U EX Taq HS DNA polymerase (Takara, Shiga, Japan), and 5 ng template. The thermal profile was 50 C for 2 min and 95 C for 10 min, followed by 40 cycles of 95 C for 30 sec, 63 C for 30 sec, and 72 C for 30 sec. Amplification and quantification were performed using the Mx3000P Multiplex Quantitative PCR system (Stratagene, La Jolla, CA). Primer sets were the following: rat/mouse sterol regulatory element binding protein-1a (SREBP-1a), 5'-GTGAGGCGGCTCTGGAACAG-3' and 5'-AGGAAGGCTTCCAGAGAGGA-3'; and rat/mouseSREBP-1c, 5'-GGAGCCATGGATTGCACATT-3' and 5'-AGGAAGGCTTCCAGAGAGGA-3'.

Statistics
Values are expressed as the mean ± SE unless otherwise stated. Scheffé’s multiple comparison test was used to determine the significance of any differences among more than three groups. P < 0.05 was considered significant.


   Results
Top
 Abstract
 Introduction
 Materials and Methods
 Results
Discussion
 References
 
Overexpression of AP-2ß leads to lipid accumulation in adipocytes
Adipocyte hypertrophy has been proposed to be the primary cause of the dysregulation of adipocytokine expression and secretion in obesity, leading to metabolic syndrome and type 2 diabetes (2). To determine whether overexpression of AP-2ß causes lipid accumulation leading to adipocyte enlargement, we constructed an adenovirus vector that encodes AP-2ß and expressed this construct in 3T3-L1 adipocytes. As shown in Fig. 1AGo, endogenous AP-2ß protein was present in the nuclear fraction, but not the cytosolic fraction, from 3T3-L1 adipocytes. AP-2ß expression was increased in a viral dose-dependent manner in the nuclear, but not the cytosolic, fraction, indicating that overexpressed AP-2ß protein localizes specifically to the nucleus.


Figure 1
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  		FIG. 1. Overexpression of AP-2ß leads to lipid accumulation in 3T3-L1 adipocytes. A, Differentiated 3T3-L1 adipocytes infected with adenovirus encoding either LacZ or AP-2ß at the indicated MOI, as described in Materials and Methods. Nuclear and cytosolic fractions were isolated and analyzed by Western blotting with anti-AP-2ß antibody. B, Differentiated 3T3-L1 adipocytes infected with adenovirus encoding either LacZ or AP-2ß at 50 MOI were fixed and stained with oil red O. Micrographs (magnification, x400) of cells are shown. C, Lipid accumulation was assessed by quantification of OD540 in destained oil red O with isopropanol. Data are presented as the increase (n-fold) compared with LacZ control cells and represent the mean ± SE of results from five independent experiments. *, P < 0.01 compared with the LacZ value. D, Differentiated 3T3-L1 adipocytes infected with adenovirus encoding either LacZ or AP-2ß at 50 MOI were cultured in medium containing 5.5, 15.5, and 25.5 mM glucose for 88 h. Lipid accumulation was assessed by quantification of OD540 in destained oil red O with isopropanol. Data are presented as the increase (n-fold) compared with LacZ control cells cultured in 25.5 mM glucose medium and represent the mean ± SE of results from three independent experiments. IB, Immunoblot.

 
When infected cells were stained with oil red O and analyzed by microscopy, cells overexpressing AP-2ß were enlarged (Fig. 1BGo), whereas protein and DNA content per well were not affected (data not shown). Triglyceride (TG) accumulation, assessed by oil red O staining, was also increased in a viral dose-dependent manner (Fig. 1CGo). The dose-response effect of glucose on lipid and TG accumulation was more marked when cells were cultured in high-glucose medium (Fig. 1DGo). These results suggest that enhanced glucose uptake by AP-2ß expression may enhance TG synthesis. We also assessed the expression of SREBP, the key transcription factor for lipogenesis. mRNA expression levels of SREBP-1a and -1c were increased in AP-2ß-overexpressing cells [1.79 ± 0.19-fold (P < 0.05) and 2.58 ± 0.56-fold (P < 0.05), respectively].

Expression of AP-2ß stimulates glucose uptake and GLUT4 translocation
To analyze the mechanisms of AP-2ß-induced lipid accumulation, we next assessed the effect of AP-2ß overexpression on glucose uptake using 2-deoxyglucose. As shown in Fig. 2AGo, glucose uptake was increased in a viral dose-dependent manner in the absence of insulin. To analyze the mechanisms of AP-2ß-enhanced glucose transport, we examined the effects of AP-2ß overexpression on cellular GLUT content (GLUT1 and GLUT4) and on translocation of GLUT protein to the plasma membrane. Infected cells were stimulated with insulin for 30 min, and total cell lysates and plasma membranes were isolated, then analyzed by Western blotting using either anti-GLUT1 or anti-GLUT4 antibody. Although the total cellular expression of both GLUT1 and GLUT4 was unchanged (Fig. 2BGo, lower panel), insulin induced a 3-fold increase in the translocation of GLUT4 to the plasma membrane. Overexpression of AP-2ß also stimulated GLUT4 translocation as potently as maximal stimulation by insulin (Fig. 2BGo, upper panel, and Fig. 2CGo). In contrast, GLUT1 did not translocate to the plasma membrane after either insulin stimulation or AP-2ß overexpression (Fig. 2BGo, upper panel, and Fig. 2CGo). These data suggest that AP-2ß stimulated glucose transport activity through inducing GLUT4 translocation to the plasma membrane in 3T3-L1 adipocytes.


Figure 2
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  		FIG. 2. Expression of AP-2ß enhances glucose transport activity and GLUT4 translocation. A, Differentiated 3T3-L1 adipocytes were infected with adenovirus encoding either LacZ or AP-2ß at the indicated MOI. 2-Deoxyglucose uptake without insulin was measured as described in Materials and Methods. Data are presented as the increase (n-fold) in glucose uptake compared with that of LacZ control cells and represent the mean ± SE of results from three independent experiments. B, Differentiated 3T3-L1 adipocytes infected with adenovirus encoding either LacZ or AP-2ß at 50 MOI were starved, then stimulated with or without insulin (100 ng/ml) for 30 min. Plasma membranes (PM; upper panel) and whole-cell lysates (total; lower panel) were analyzed by Western blotting using anti-GLUT4 and anti-GLUT1 antibodies. C, Graphic representation of the results shown in B (upper panel) generated using a desk scanner. Data are presented as the increase (n-fold) compared with that of unstimulated LacZ control cells and represent the mean ± SE of results from three independent experiments. *, P < 0.05 compared with the LacZ (insulin –) value. IB, Immunoblot.

 
Atypical PKC and PLC were involved in AP-2ß-enhanced glucose uptake
To examine which molecules are involved in AP-2ß-enhanced glucose uptake, we pretreated cells with various inhibitors of signaling molecules involved in activation of glucose transport. As shown in Fig. 3AGo, the PI3-K inhibitor (LY294002) did not suppress AP-2ß-enhanced glucose uptake. In contrast, the atypical PKC inhibitor [myristoylated pseudosubstrate (myr-PS)-PKC{zeta}] almost completely blocked AP-2ß-induced glucose uptake, and the PLC inhibitor (U73122) also suppressed AP-2ß-induced glucose uptake by 70% (Fig. 3AGo). The inhibitor for p38 MAPK (SB203580) was without effect (Fig. 3AGo). These results suggest that atypical PKC and PLC may be involved in AP-2ß-enhanced glucose uptake. To confirm this, we measured the activities of atypical PKC and PLC. As shown in Fig. 3BGo, in the absence of insulin, PKC{zeta}/{lambda} activity was enhanced in AP-2ß-overexpressing cells. PLC activity, assessed by measuring cellular IP3 content, was also enhanced in these cells (Fig. 3CGo), consistent with the results presented in Fig. 3AGo, whereas the protein content of PKC{zeta}/{lambda} or PLC was unchanged (data not shown). To confirm the involvement of atypical PKC, we coexpressed AP-2ß with the DN form of PKC{lambda} (DN-PKC{lambda}) in 3T3-L1 adipocytes. As displayed in Fig. 3DGo, we found that DN-PKC{lambda} can inhibit insulin-stimulated glucose transport by 50% at 50 MOI, and AP-2ß stimulated glucose transport by 67% at the same MOI. These results support the idea that activation of both atypical PKC and PLC is involved in the AP-2ß-stimulated glucose transport.


Figure 3
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  		FIG. 3. Atypical PKC and PLC were involved in AP-2ß-enhanced glucose uptake. A, Differentiated 3T3-L1 adipocytes were infected with adenovirus encoding either LacZ or AP-2ß at 50 MOI. After treatment with 50 µM LY294002, 50 µM myr-PS-PKC{zeta}, 10 µM U73122, or 20 µM SB203580 for 2 h, 2-deoxyglucose uptake was measured as described in Materials and Methods. Data are presented as the increase (n-fold) in glucose uptake compared with that of unstimulated LacZ control cells and represent the mean ± SE of results from four independent experiments. **, P < 0.01 compared with the AP-2ß (–, inhibitor) value. *, P < 0.05 compared with the AP-2ß (–, inhibitor) value. B, Differentiated 3T3-L1 adipocytes infected with adenovirus encoding either LacZ or AP-2ß at 50 MOI were starved and stimulated with or without insulin (100 ng/ml) for 30 min. Whole-cell lysates were immunoprecipitated with anti-PKC{zeta}/{lambda} antibody and assayed for PKC{zeta}/{lambda} activity as described in Materials and Methods. Data are presented as the increase (n-fold) in PKC{zeta}/{lambda} activity compared with unstimulated LacZ control cells and represent the mean ± SE of results from three independent experiments. **, P < 0.01; *, P < 0.05 [compared with the LacZ (insulin –) value. C, Differentiated 3T3-L1 adipocytes infected with adenovirus encoding either LacZ or AP-2ß at 50 MOI. PLC activity was measured as described in Materials and Methods. Data are presented as the increase (n-fold) in PLC activity compared with LacZ control cells and represent the mean ± SE of results from three independent experiments. *, P < 0.01 compared with the LacZ value. D, Differentiated 3T3-L1 adipocytes were infected with adenovirus encoding either LacZ or AP-2ß with DN-PKC at the indicated MOI. 2-Deoxyglucose uptake in the absence or presence of insulin was measured as described in Materials and Methods. Data are presented as the increase (n-fold) in glucose uptake compared with that of unstimulated LacZ control cells and represent the mean ± SE of results from three independent experiments.

 
PLC{gamma}-enhanced glucose uptake is inhibited by the atypical PKC inhibitor
Involvement of PLC{gamma} has been reported in insulin-stimulated glucose transport in several cell types, including 3T3-L1 adipocytes (23, 24, 25). Furthermore, PLC{gamma}-induced activation of atypical PKC has been reported (23). These reports led us to propose that AP-2ß may enhance glucose uptake via stimulation of PLC activity and, subsequently, atypical PKC activation. Thus, we determined whether overexpression of PLC{gamma} stimulates glucose uptake and whether it was inhibited by the atypical PKC inhibitor. For this purpose, we overexpressed PLC{gamma} by adenovirus in 3T3-L1 adipocytes (Fig. 4AGo) and measured glucose transport activity. As shown in Fig. 4BGo, glucose uptake was stimulated by 2-fold in cells overexpressing PLC{gamma}, which was inhibited by myr-PS-PKC{zeta} as potently as by U73122. In contrast, LY294002 was without effect (data not shown). Furthermore, PKC{zeta}/{lambda} activity was enhanced in PLC{gamma}-overexpressing cells (Fig. 4CGo), indicating that atypical PKC is located downstream of PLC{gamma} signaling under these conditions. These results strongly suggest that AP-2ß enhances glucose transport through activation of PLC{gamma} and subsequent atypical PKC activation.


Figure 4
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  		FIG. 4. PLC{gamma}-enhanced glucose uptake is suppressed by atypical PKC inhibitor. A, Differentiated 3T3-L1 adipocytes infected with adenovirus encoding either LacZ or PLC{gamma} at 50 MOI were lysed, and whole-cell lysates were analyzed by Western blotting with anti-PLC{gamma} antibody. B, After treatment with 50 µM myr-PS-PKC{zeta} or 10 µM U73122 for 2 h, 2-deoxyglucose uptake was measured as described in Materials and Methods. Data are presented as the increase (n-fold) in glucose uptake compared with that of unstimulated LacZ control cells and represent the mean ± SE of results from four independent experiments. *, P < 0.01 compared with the PLC{gamma} (–, inhibitor) value. C, Cells were starved, and whole-cell lysates were immunoprecipitated with anti-PKC{zeta}/{lambda} antibody and assayed for PKC{zeta}/{lambda} activity as described in Materials and Methods. Data are presented as the increase (n-fold) in PKC{zeta}/{lambda} activity compared with LacZ control cells and represent the mean ± SE of results from three independent experiments. *, P < 0.05 compared with the LacZ value. IB, Immunoblot.

 
Expression of AP-2ß stimulates Gab1 tyrosine phosphorylation and its association with PLC{gamma}
To investigate the molecular mechanism for activation of PLC{gamma} by AP-2ß expression, we assessed the alteration in tyrosine phosphorylation states of cellular proteins by Western blotting using antiphosphotyrosine antibody in cells overexpressing AP-2ß. We found increased tyrosine phosphorylation in 120- to 130- and 60- to 70-kDa proteins (data not shown). We identified Gab1 as a 120- to 130-kDa tyrosine-phosphorylated protein in AP-2ß-infected cells (Fig. 5AGo, top panel), whereas protein expression was unchanged (Fig. 5AGo, bottom panel). Sorbitol treatment is reported to stimulate Gab1 phosphorylation and increase PLC{gamma} association, and those are proposed as the mechanism of osmotic shock-induced glucose uptake (26, 27). The association of Gab1 with PLC{gamma} was also observed in AP-2ß-infected cells as well as sorbitol treatment (Fig. 5AGo, second panel). Although sorbitol treatment induced the association of Gab1 with p85 of PI3-K, we were not able to observe any association of Gab1 with p85 of PI3-K by AP-2ß expression (Fig. 5AGo, third panel). Consistent with this, PI3-K activity associated with Gab1 was stimulated by sorbitol, but not by AP-2ß (Fig. 5BGo). These results suggest that the precise mechanisms by which osmotic shock and AP-2ß induced Gab1 phosphorylation and glucose uptake are somewhat different.


Figure 5
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  		FIG. 5. Gab1 tyrosine phosphorylation and its association with PLC{gamma} are stimulated in AP-2ß-overexpressing cells. A, Differentiated 3T3-L1 adipocytes infected with adenovirus encoding either LacZ or AP-2ß at 50 MOI were incubated with or without 600 mM sorbitol for 20 min, and whole-cell lysates were immunoprecipitated with anti-Gab1 antibody, followed by Western blotting using anti-phosphotyrosine antibody (top panel), anti-PLC{gamma} antibody (second panel), anti-p85 antibody (third panel), and anti-Gab1 antibody (fourth panel). Whole-cell lysates were also immunoblotted with anti-Gab1 antibody (bottom panel). B, 3T3-L1 adipocytes infected with adenovirus encoding either LacZ or AP-2ß at 50 MOI were incubated with or without 600 mM sorbitol for 20 min. PI3-K activity was measured as described in Materials and Methods.

 
Depletion of endogenous AP-2ß protein attenuates glucose transport
To evaluate the physiological role of AP-2ß, we electroporated siRNA into 3T3-L1 adipocytes to deplete AP-2ß protein. Forty-eight hours after electroporation, the amount of AP-2ß protein in the nuclear fraction was decreased by 70% in AP-2ß siRNA-transfected cells compared with that in control siRNA-transfected cells (Fig. 6AGo). In this condition, knockdown of AP-2ß attenuated glucose transport activity by 30% in the basal state (Fig. 6BGo). In contrast, insulin-stimulated glucose transport was not affected by AP-2ß siRNA (Fig. 6CGo). These results indicate that endogenous AP-2ß actually has a role in modulating glucose transport activity by insulin-independent mechanism in adipocytes.


Figure 6
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  		FIG. 6. Effects of depletion of AP-2ß by siRNA and expression of mutant AP-2ß that lacks DNA-binding ability on glucose transport. A–C, 3T3-L1 adipocytes were nucleofected with 5 µg vector (mock), AP-2ß siRNA, or scrambled control siRNA as described in Materials and Methods. A, Nuclear fractions were prepared and analyzed by Western blotting with anti-AP-2ß antibody. B and C, 2-Deoxyglucose uptake with or without insulin (100 ng/ml) for 60 min was measured as described in Materials and Methods. D and E, 3T3-L1 adipocytes were nucleofected with 5 µg pcDNA 3.1 (mock), pcDNA 3.1/AP-2ß WT, or pcDNA 3.1/AP-2ß (R225C) as described in Materials and Methods. D, Whole-cell lysates were prepared and analyzed by Western blotting with anti-AP-2ß antibody, and 2-deoxyglucose uptake was measured as described in Materials and Methods. E, Cells were immunoprecipitated with anti-Gab1 antibody, followed by Western blotting using antiphosphotyrosine antibody (upper panel) and anti-Gab1 antibody (lower panel). Data are presented as the increase (n-fold) in glucose uptake compared with that of mock control cells and represent the mean ± SE of results from five independent experiments. *, P < 0.05 compared with the mock value. IB, Immunoblot.

 
DNA-binding ability of AP-2ß is necessary for its effect on glucose transport
Because AP-2ß is a transcription factor, it may act through the regulation of gene expression. Thus, to determine whether AP-2ß’s effect of enhancing glucose uptake is exerted through its action as a transcription factor, we prepared a mutant AP-2ß that lacks DNA-binding ability. Arginine 225 of AP-2ß is located in the DNA-binding domain, and replacement of this arginine with cysteine (R225C) extinguishes DNA-binding ability (4, 5). WT and mutant (R225C) AP-2ß were transiently expressed in 3T3-L1 adipocytes, then glucose uptake was assayed. As shown in Fig. 6DGo, glucose uptake was enhanced in WT transfected cells. However, R225C mutant was without effect, whereas the protein expressions of WT- and R225C-expressing cells were identical (Fig. 6DGo). Furthermore, tyrosine phosphorylation of Gab1 was enhanced in WT, but not mutant, transfected cells (Fig. 6EGo). These results suggest that DNA-binding ability is necessary for AP-2ß to enhance both glucose uptake and Gab1 tyrosine phosphorylation, and that AP-2ß may exert its effect through regulating specific gene expression.

Overexpression of AP-2ß causes insulin resistance
We next examined the effect of AP-2ß overexpression on insulin action. We assessed insulin-stimulated glucose uptake and found that it was increased at all insulin concentrations in AP-2ß-overexpressing cells (Fig. 7AGo). However, insulin responsiveness in these cells was reduced compared with that in control cells. Furthermore, overexpression of AP-2ß also led to reduced insulin sensitivity for stimulation of glucose transport, as demonstrated by the rightward shift in the dose-response curve in AP-2ß-overexpressing cells compared with control cells (ED50, 4.53 ± 0.35 and 1.50 ± 0.44 ng/ml, respectively; P < 0.01; Fig. 7BGo). To assess the mechanism by which AP-2ß causes insulin resistance, we analyzed the insulin signaling pathway. As shown in Fig. 7Go, C and D, insulin-stimulated tyrosine phosphorylation of IR and subsequent downstream IRS-1 and Akt were all inhibited in AP-2ß-overexpressing cells. Furthermore, protein expression of IRS-1 was down-regulated in these cells. These results suggest that overexpression of AP-2ß induced insulin resistance mainly through inhibiting the effect of insulin on tyrosine phosphorylation of IR and IRS-1, but also probably through inhibition at multiple steps.


Figure 7
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  		FIG. 7. Overexpression of AP-2ß causes insulin resistance. A, Differentiated 3T3-L1 adipocytes infected with adenovirus encoding either LacZ or AP-2ß at 50 MOI were stimulated with or without insulin at the indicated concentrations for 60 min. 2-Deoxyglucose uptake was measured as described in Materials and Methods. Data are presented as the increase (n-fold) in glucose uptake compared with that of unstimulated LacZ control cells and represent the mean ± SE of results from three independent experiments. B, The graph shows the percentage of the maximal response, and data represent the mean ± SE of results from three independent experiments. *, P < 0.05 compared with the LacZ (1 ng/ml insulin) value. **, P < 0.01 compared with the LacZ (10 ng/ml insulin) value. C, Starved cells were stimulated with or without insulin (100 ng/ml) for 5 min. Whole-cell lysates were analyzed by Western blotting with antiphospho-IR antibody (top panel), IRß antibody (second panel), IRS-1 antibody (fourth panel), phospho-Akt antibody (fifth panel), or Akt antibody (bottom panel). Whole-cell lysates were immunoprecipitated with anti-IRS-1 antibody, followed by immunoblotting with antiphosphotyrosine antibody (third panel). D, Graphic representation of the results shown in C generated using a desk scanner. Data are presented as the increase (n-fold) in protein levels (IRß and IRS-1) or phosphorylation levels (phospho-IR and phospho-IRS-1) compared with unstimulated LacZ control cells and represent the mean ± SE of results from three independent experiments. *, P < 0.01 compared with the LacZ (insulin +) value. IB, Immunoblot.

 

   Discussion
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
References
 
We have recently identified the AP-2ß gene (TFAP2B) as a susceptibility gene for type 2 diabetes by conducting a genome-wide association study (3). Several variations in the TFAP2B were significantly associated with type 2 diabetes. AP-2{alpha} has been reported to inhibit adipogenesis through inhibition of C/EBP{alpha}, and the expression of AP-2{alpha} was down-regulated upon induction of differentiation in 3T3-L1 adipocytes (14). In contrast, we discovered that mouse AP-2ß (Tcfap2b) is preferentially expressed in adipose tissue, and its expression is increased upon induction of differentiation in 3T3-L1 adipocytes (3). Although AP-2{alpha} and AP-2ß recognize the same sequences, disruption of a distinct gene caused death, and the expression patterns of AP-2{alpha} and AP-2ß are distinct (6, 11, 28). Thus, AP-2{alpha} and AP-2ß may have different roles in adipocytes.

The preliminary experiments show that subjects with the disease-susceptible allele have stronger expression in adipose tissue than subjects without the susceptible allele (Tsukada, S., Y. Tanaka, H. Maegawa, A. Kashiwagi, R. Kawamori, and S. Maeda, unpublished observations). Thus, these findings suggest that TFAP2B plays an important role in the pathogenesis of type 2 diabetes through the dysregulation of adipocyte function. In the present study we demonstrated that overexpression of AP-2ß in 3T3-L1 adipocytes leads to TG accumulation and insulin resistance. These phenomena are considered to play crucial role in the pathogenesis of obesity-related disease such as metabolic syndrome and type 2 diabetes.

We found that glucose transport activity and GLUT4 translocation were enhanced in cells overexpressing AP-2ß. On the contrary, knockdown of endogenous AP-2ß inhibited glucose transport activity, indicating that endogenous AP-2ß actually has a role in modulating glucose transport in adipocytes. Furthermore, TG accumulation was dependent on the concentration of glucose in the culture medium. One possible explanation is that an increased amount of glucose is used for TG synthesis, because glucose can be converted to lipid in 3T3-L1 adipocytes (29). In support of this idea, we observed increased SREBP-1a and -1c expression, which induces lipogenic genes, leading to TG accumulation, as observed in liver (30, 31, 32). Additional study is needed to identify the exact mechanism for lipid accumulation induced by AP-2ß.

AP-2ß-enhanced glucose transport was not blocked by the PI3-K inhibitor. The PI3-K dependency of insulin stimulation has been clearly demonstrated in a variety of insulin-responsive tissues and cell lines (33). Akt and atypical PKC are activated downstream of PI3-K, and both are mediators of glucose transport stimulation (34, 35). We observed that AP-2ß’s effect was not inhibited by the PI3-K inhibitor, Akt was not activated, and PI3-K activity was unchanged in cells overexpressing AP-2ß (data not shown), indicating that the signaling pathway in which AP-2ß enhances glucose transport activity is distinct from that of insulin. In contrast, the inhibitor of atypical PKC (chemical and DN forms of PKC{lambda}) markedly blocked AP-2ß-enhanced glucose transport, and it was also suppressed by the PLC inhibitor, indicating that atypical PKC and PLC are involved. This idea was supported by the findings that atypical PKC and PLC activities were enhanced in cells expressing AP-2ß.

The inhibitor for p38 MAPK, SB203580, did not inhibit AP-2ß-enhanced glucose transport. SB203580 has been reported to inhibit insulin-stimulated glucose transport, but not GLUT4 translocation, in 3T3-L1 adipocytes and L6 myotubes, indicating that this reagent inhibits the intrinsic activity of GLUT4 (36, 37). Thus, our results suggest that AP-2ß’s effect on glucose transport was independent of the intrinsic activity of GLUT4.

To explore the interaction between these two molecules, we overexpressed PLC{gamma} and assessed the alteration in glucose transport activity. Overexpression of PLC{gamma} also enhanced glucose transport, which was not inhibited by the PI3-K inhibitor, but was inhibited by the atypical PKC inhibitor, as is the case for AP-2ß overexpression. Furthermore, atypical PKC activity was stimulated by overexpression of PLC{gamma}. These results support our hypothesis that AP-2ß enhances glucose transport through activation of atypical PKC activated by PLC{gamma}. However, the mechanism by which activation of PLC{gamma} leads to the activation of atypical PKC is unclear. PKC{zeta}/{lambda} is known to be a target of both PI3-K and 3-phosphoinositide-dependent protein kinase-1 (19, 38). PKC{zeta}/{lambda} simultaneously and directly interacts with phosphatidylinositol 3,4,5-trisphosphate, which liberates from PS-dependent autoinhibition (39). However, our results showed that AP-2ß-enhanced glucose transport was not inhibited by the PI3-K inhibitor, and PI3-K was not activated by overexpression of AP-2ß, suggesting that PI3-K is not involved in AP-2ß-enhanced atypical PKC activation. PLC catalyzes the hydrolysis of phosphatidylinositol 4,5-bisphosphate to diacylglycerol (DAG) and IP3. DAG is metabolized to phosphatidic acid (PA) by DAG kinase. PKC{zeta}/{lambda} can bind and be activated by PA (40). Thus, it is possible that in our experiment, PA production by PLC activation was involved in the activation of atypical PKC.

PLC signaling has also been implicated in the modulation of GLUT4-mediated glucose transport. The PLC inhibitor, U73122, inhibited insulin-stimulated glucose transport in 3T3-L1 adipocytes (24, 25, 41), brown adipocytes (23), L6 myocytes (42), and rat skeletal muscle (43). U73122 also inhibited epidermal growth factor (EGF)-stimulated glucose transport in 3T3-L1 adipocytes (24) and platelet-derived growth factor-stimulated glucose transport in Chinese hamster ovary cells overexpressing GLUT4 (44). Furthermore, microinjection of the Src homology 2 (SH2) domain of PLC{gamma} inhibited insulin-stimulated GLUT4 translocation in 3T3-L1 adipocytes (25). These data suggest the existence of a PLC-mediated pathway leading to glucose transport. Interestingly, inhibition of insulin-stimulated glucose transport by the PLC inhibitor was restored by the addition of cell-permeable PA in brown adipocytes (23). Taken together, the results suggest that AP-2ß stimulates glucose transport via activation of atypical PKC activated by PLC{gamma}.

Enzymatic activity of PLC{gamma} is known to be stimulated by tyrosine phosphorylation of PLC{gamma} itself after stimulation with EGF, platelet-derived growth factor, and nerve growth factor (45, 46). However, its phosphorylation is not always necessary for activation. PLC{gamma} recognizes phosphotyrosine-containing sequences via its SH2 domain, and association of the SH2 domain with a phosphopeptide or binding of phosphatidylinositol 3,4,5-trisphosphate to the pleckstrin homology domain also stimulates PLC{gamma} activity without tyrosine phosphorylation (45, 46).

Gab1 is one of the adaptor proteins that is phosphorylated on tyrosine in response to insulin, EGF, and several growth factors, creating a number of docking sites to mediate interactions with SH2 domain-containing proteins such as Src homology protein tyrosine phosphatase, p85 subunit of PI3-K, Grb2, Nck, and PLC{gamma} (47). Association with these molecules was found to be critical for the function of Gab protein in mediating intracellular signaling pathways from the receptors. Interestingly, it has been reported that Gab1 is involved in osmotic shock-induced glucose transport through recruiting and activating PI3-K (27) and PLC{gamma} (26) in 3T3-L1 adipocytes. In the present study we found that tyrosine phosphorylation of Gab1 and its association with PLC{gamma} were stimulated by AP-2ß expression. Thus, it is likely that AP-2ß stimulates PLC{gamma} activity through binding to phosphorylated Gab1. It has been reported that osmotic shock induced by sorbitol stimulated Gab1 tyrosine phosphorylation and its association with p85 and PLC{gamma}, and that osmotic shock-induced glucose uptake was inhibited by these inhibitors, suggesting that osmotic shock uses these two pathways. In contrast, we were not able to observe any association of Gab1 with p85 by AP-2ß expression. This was also supported by the findings that PI3-K activity associated with Gab1 was stimulated by osmotic shock, but not by AP-2ß expression. These results are consistent with data showing that AP-2ß-induced glucose uptake is PI3-K independent. PLC{gamma} and p85 recognize different consensus motifs [YXXP for PLC{gamma} (48) and YXXM for p85 (49)]; both are present in Gab1. Thus, it may be possible that osmotic shock and AP-2ß stimulate Gab1 on different tyrosine residues.

Which tyrosine kinases mediate Gab1 tyrosine phosphorylation is currently unknown. In our preliminary experiments we found that expression of Neu, a member of the EGF receptor family, was increased in the cells overexpressing AP-2ß. It has been reported that the promoter region of the Neu gene has an AP-2 binding site, and AP-2 can bind and stimulate its promoter activity (50). Interestingly, Neu contains a potential Gab1 binding site (51), and several reports have indicated the important role of Gab1 in Neu-mediated signaling (51, 52, 53). Thus, we speculate that Neu may be one of the candidate molecules mediating the effect of AP-2ß observed in our experiments. However, we are not able to rule out the possibility that another undefined protein(s) induced by AP-2ß overexpression is involved in AP-2ß’s effects; additional study is needed to clarify the precise mechanism.

We found that overexpression of AP-2ß leads to insulin resistance, as assessed by insulin-stimulated glucose uptake and phosphorylation of target molecules. Tyrosine phosphorylation of IR and subsequent phosphorylation of IRS-1 and Akt were all decreased. These results suggest that the decreased tyrosine phosphorylation of IR and IRS-1 is a major cause of AP-2ß-induced insulin resistance. Serine/threonine phosphorylation of the IR by several kinases has been proposed to exist in insulin-resistant states (54, 55). Serine/threonine phosphorylation of IRS-1 can also serve as a negative modulator of insulin signaling by inhibition of its tyrosine phosphorylation or led to degradation of the protein. Serine/threonine kinases, including PKC{zeta}, that are responsible for phosphorylation of IRS-1 and inhibition of its function have been reported (56). In fact, as shown in Fig. 7Go, protein expression of IRS-1 was decreased by AP-2ß overexpression. The involvement of protein tyrosine phosphatases (PTPases) is also possible, because PTP1B and leukocyte common antigen-related phosphatase are reported to be negative regulators of IR kinase and IRS-1 (16, 57). However, we did not observe any changes in the expression of these PTPases (data not shown). Because AP-2ß modulates the expression of many genes, we speculate that AP-2ß may down-regulate insulin signaling at multiple levels.

In conclusion, AP-2ß overexpression leads to lipid accumulation. Enhanced glucose uptake is one of major mechanisms for lipid accumulation. Furthermore, AP-2ß overexpression induces insulin resistance in 3T3-L1 adipocytes. Thus, we propose that this transcription factor is a candidate gene for causing metabolic syndrome and type 2 diabetes through the induction of adipocyte dysfunction. Additional study is required to clarify the significance of AP-2ß in the pathogenesis of obesity-related disease in vivo.


   Acknowledgments
 
We thank Drs. J. M. Olefsky (University of California, San Diego, CA), S. G. Rhee (National Institutes of Health, Bethesda, MD), and W. Ogawa (Kobe University, Kobe, Japan) for providing 3T3-L1 adipocytes, plasmid encoding PLC{gamma}, and DN-PKC{lambda} adenovirus, respectively. We also thank Ms. Y. Nakai for her technical assistance.


   Footnotes
 
This work was supported in part by a grant-in-aid from the Ministry of Education, Culture, Sports, Science, and Technology, Japan (to H.M.).

First Published Online December 22, 2005

Abbreviations: AP-2ß, Activating protein-2ß; C/EBP{alpha}, CCAAT/enhancer-binding protein-{alpha}; DAG, diacylglycerol; DN, dominant negative; DTT, dithiothreitol; EGF, epidermal growth factor; Gab1, Grb2-associated binder-1; GLUT4, glucose transporter 4; IP3, inositol 1,4,5-trisphosphate; IR, insulin receptor; IRS, insulin receptor substrate; MOI, multiplicity of infection; PA, phosphatidic acid; PI3-K, phosphatidylinositol 3-kinase; PKC, protein kinase C; PLC, phospholipase C; PMSF, phenylmethylsulfonylfluoride; PS, pseudosubstrate; PTPase, protein-tyrosine phosphatase; SH, Src homology; siRNA, small interfering RNA; SREBP, sterol regulatory element-binding protein; TG, triglyceride; WT, wild type.

Received October 13, 2005.

Accepted for publication December 13, 2005.


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 Introduction
 Materials and Methods
 Results
 Discussion
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