Dehydroepiandrosterone Stimulates Glucose Uptake in Human and Murine Adipocytes by Inducing GLUT1 and GLUT4 Translocation to the Plasma Membrane

Polyclonal insulin receptor β-subunit, monoclonal phosphotyrosine (PY99), monoclonal PKC-β₁, and polyclonal PKC-β₂ antibodies were purchased from Santa Cruz Biotechnology (Santa Cruz, CA). Polyclonal antibodies against Akt, phospho-Akt (Thr 308), phospho-Akt (Ser-473), phospho-GSK-3α/β (Ser-9/Ser-21), mitogen-activated protein (MAP)/extracellular signal-related kinase (ERK) kinase (MEK)-1/2, phospho-MEK-1/2 (Ser-217/Ser-221), and phospho-p42/p44 MAP kinase (Thr-202/Tyr-204) were from Cell Signaling Technology (Beverly, MA). Polyclonal antibodies against p85, GSK-3, IRS-1, and IRS-2 were purchased from Upstate Biotechnology (Saranac Lake, NY). MEK-1/2 antibodies were from Zymed Laboratories (San Francisco, CA). A monoclonal antibody against clathrin heavy chain was from Transduction Laboratories (Lexington, KY). Rabbit polyclonal antibodies against rat GLUT4 were provided by Dr. R.J. Smith (Brown University, Providence, RI) or purchased from Diagnostic International (Schriesheim, Germany). Sheep polyclonal antibodies against Akt2 were provided by Dr. D.R. Alessi (University of Dundee, Dundee, U.K.). GLUT1 antibody was from Charles River (Southbridge, MA). Antibodies against PKC isozymes (α, β, ζ/λ) were from Life Technologies (Gibco BRL). Wortmannin, LY294002, GF109203X, U73122, Calphostin C, Gö6,976, and PD098059 were from Calbiochem (La Jolla, CA). The PKC-ζ/λ inhibitor cell-permeable myristoylated PKC-ζ/λ pseudosubstrate (myr-PKC-ζ/λ) was from Quality Controlled Biochemicals (Hopkington, MA). Porcine insulin and LY379196, a specific PKC-β inhibitor, were gifts from Eli Lilly (Indianapolis, IN). The PKC-β inhibitor cell-permeable myristoylated PKC-β pseudosubstrate was from Calbiochem.

3T3-L1 fibroblasts (American Type Culture Collection, Rockville, MD) were grown in Dulbecco’s modified Eagle’s medium (DMEM) supplemented with 10% FCS, 2 mmol/l l-glutamine, 100 units/ml penicillin, and 100 mg/ml streptomycin at 37°C in an atmosphere of 5% CO₂. Differentiation into adipocytes was induced as described previously (18).

Specimens of human subcutaneous adipose tissue were obtained from the abdominal region of nondiabetic subjects undergoing elective surgery for nonmalignant diseases. The study had the approval of the local ethical committee. The subcutaneous adipose tissue (∼1 g) was removed and placed in Krebs-Ringer buffer (136 mmol/l NaCl, 4.7 mmol/l KCl, 1.25 mmol/l MgSO₄, 5 mmol/l Na₂HPO₄, 1.25 mmol/l CaCl₂ [pH 7.4]), containing 40 mg/ml BSA (fraction V) and 5.5 mmol/l glucose, at 37°C. Isolated adipocytes were obtained using a method modified from Rodbell (19) and characterized by evaluating the insulin receptor and GLUT4 content in total cell lysates.

For measuring glucose transport rates in 3T3-L1 adipocytes, cells were grown in serum-free DMEM for 16 h and then incubated in the absence or presence of insulin or DHEA for the indicated times at 37°C. Transport was started by adding 50 μmol/l [³H]2-deoxy-d-glucose (NEN, Boston, MA) and 1 μCi in 1 ml of Krebs-Ringer phosphate buffer (pH 7.4) for 5 min at 37°C and stopped by placing the cells on ice and rapidly washing them three times with ice-cold buffer. Cells were lysed in 1 ml of lysis buffer containing 0.1% Triton X-100 for 45 min. Aliquots of the cell lysates were used for liquid scintillation counting and determination of protein content by the Bradford method, respectively. Nonspecific transport was assayed in the presence of 10 μmol/l cytochalasin B. Glucose transport measurements in human adipocytes were performed as described by Ciaraldi et al. (20).

For obtaining total membranes from 3T3-L1 adipocytes, cells were collected into 10 ml of ice-cold HES buffer (250 mmol/l sucrose, 1 mmol/l EDTA, 1 mmol/l phenylmethylsulfonyl fluoride [PMSF], 1 μmol/l pepstatin, 1 μmol/l aprotinin, 1 μmol/l leupeptin, and 20 mmol/l HEPES, pH 7.4) and subsequently homogenized with 20 strokes in a glass Dounce homogenizer (Type C; Thomas, Philadelphia, PA) at 4°C. After centrifugation at 1,000g for 3 min at 4°C to remove large cell debris and unbroken cells, the supernatant was then centrifuged at 245,000g for 90 min at 4°C to yield a pellet of total cellular membranes and a supernatant representing the cytosolic fraction (21). For yielding total cellular membranes from human adipocytes, cells were homogenized with 20 strokes in a glass Dounce homogenizer in ice-cold HES buffer. The homogenate was then centrifuged at 1,000g for 3 min at 4°C, and the postnuclear supernatant was separated from the pellet and fat cake and then centrifuged at 245,000g for 90 min at 4°C. The resulting pellet representing the total cellular membrane fraction (22) was resuspended in HES buffer before use. High-density microsome, low-density microsome (LDM), and plasma membrane (PM) subcellular fractions from 3T3-L1 adipocytes were obtained by differential ultracentrifugation, as described previously (23). PM, LDM, and high-density microsome fractions from human adipocytes were prepared according to Garvey et al. (22).

The preparation of PM lawns was performed as described by Robinson et al. (23). Briefly, after incubating cells on coverslips with the appropriate treatment, adipocytes were sonicated, yielding a lawn of PM fragments attached to the coverslip. Coverslips were then incubated with GLUT4 or GLUT1 antibodies and then with Alexa 546- or Alexa 488-conjugated secondary antibodies, respectively. The fluorescence intensity of individual PM fragments from five random fields for each experimental condition was analyzed using a Leica TCS SP2 laser confocal microscope.

For preparing total cell lysates, 3T3-L1 adipocytes were washed with Ca²⁺/Mg²⁺-free PBS and then mechanically detached in ice-cold lysis buffer containing 50 mmol/l HEPES (pH 7.5), 150 mmol/l NaCl, 1 mmol/l MgCl₂, 1 mmol/l CaCl₂, 10% glycerol, 10 mmol/l sodium pyrophosphate, 10 mmol/l sodium fluoride, 2 mmol/l EDTA, 2 mmol/l PMSF, 5 μg/ml leupeptin, 2 mmol/l sodium orthovanadate, and 1% Nonidet p-40. After incubation for 45 min at 4°C, the preparation was centrifuged at 12,000g for 10 min at 4°C. The resulting supernatant was assayed for determination of protein concentration using the Bradford method and subjected overnight to immunoprecipitation at 4°C with antibodies against the insulin receptor, IRS-1, IRS-2, PKC-β₁, or PKC-β₂, as indicated. The resulting immune complexes were adsorbed to protein A-Sepharose beads for 2 h at 4°C, and the pelleted beads were washed three times in lysis buffer and then incubated in Laemmli buffer for 5 min at 100°C. Protein samples were resolved by electrophoresis on 6%, 7%, 10%, or 12% SDS-polyacrylamide gels, as appropriate, directly or after immunoprecipitation and subjected to immunoblotting with the appropriate antibodies, as described previously (24). The proteins were quantified by densitometric analysis using Optilab image analysis software (Graftek SA, Mirmande, France). For measurements of PI 3-kinase activity, cell lysates (∼1–2 mg) were subjected overnight to immunoprecipitation at 4°C with antibodies against IRS-1 or IRS-2. The activity of PI 3-kinase in the immunoprecipitates was determined as described previously (25).

Cytosolic-free calcium concentrations were evaluated in single adipocytes loaded with the intracellular Ca²⁺ indicator fura-2 (26). Isolated cells seeded onto 24-mm round glass coverslips were loaded with 10 μmol/l fura-2/AM in serum-free DMEM for 1 h at 37°C. Coverslips were washed three times with PBS and transferred to a Sykes Bellco open chamber (Bellco Biotechnology, Vineland, NJ) containing 1 ml of Krebs-Ringer-HEPES buffer (125 mmol/l NaCl, 5 mmol/l KCl, 1.2 mmol/l KH₂PO₄, 1.2 mmol/l MgSO₄, 2 mmol/l CaCl₂, 265 mmol/l HEPES, and 6 mmol/l glucose). [Ca²⁺]_i-dependent fluorescence was measured with a microfluorometer (Cleveland Bioinstrumentation, Cleveland, OH) connected with a Zeiss IM35 inverted microscope equipped with a Nikon GFX40 fluor objective. Recordings were performed at dual excitation wavelength (340 and 380 nm, bandwidth 0.5 nm) using an air turbine high-speed rotating wheel carrying the two excitation filters. At the end of each experiment, calibration was performed by adding 5 μmol/l ionomycin followed by 7.5 mmol/l EGTA to obtain Ca²⁺-saturated and nominally Ca²⁺-free fura-2 fluorescence, respectively.

All data are expressed as mean ± SE. Statistical analyses were performed by unpaired Student’s t tests.

For investigating the effects of DHEA on the glucose transport system, 3T3-L1 adipocytes were incubated in the presence of various concentrations of DHEA for different times, and glucose transport was measured by determining the rates of [³H]2-deoxy-d-glucose uptake. DHEA induced a time- and dose-dependent increase in glucose transport rates in 3T3-L1 adipocytes. An initial statistically significant effect of DHEA on transport was observed at the concentration of 1 μmol/l (35% of basal; P < 0.05) and the maximal effect at 100 μmol/l (300% of basal; P < 0.05; Fig. 1A). With 100 μmol/l DHEA, glucose transport was maximally increased after 120 min of incubation with the steroid (Fig. 1A). The effects of DHEA on glucose transport were specific because treatment of 3T3-L1 adipocytes with equimolar concentrations of other steroid hormones, including 17-β-estradiol, progesterone, Δ-4-androstenedione, testosterone, and dihydrotestosterone, had no effects on 2-deoxy-d-glucose uptake rates (Fig. 1A). However, DHEA was less potent than insulin, used at maximally effective concentrations, in stimulating glucose transport in 3T3-L1 adipocytes (P < 0.05 vs. insulin; Fig. 1A). For assessing whether the ability of DHEA to enhance glucose transport in 3T3-L1 adipocytes could be explained by DHEA-induced changes in the amounts of glucose transporter protein at the cell surface, the protein levels of GLUT1 and GLUT4, the two predominant glucose transporter isoforms expressed in 3T3-L1 adipocytes, were measured in LDM and PM fractions in the basal state or after treatment with DHEA, or insulin for comparison. PM and LDM fractions prepared from basal, DHEA-treated, and insulin-treated adipocytes contained comparable amounts of clathrin (27) (Fig. 2A). DHEA induced a significant increase in the PM content of both GLUT1 and GLUT4 proteins (respectively, 180 and 160% of basal; P < 0.05; Fig. 2A and B). DHEA and insulin stimulated GLUT1 translocation to the PM to a similar extent, whereas DHEA was less effective than insulin in inducing PM translocation of GLUT4 (P < 0.05 vs. insulin; Fig. 2A and B). Similar results were obtained by measuring the amounts of GLUT4 and GLUT1 in adipocyte PM fragments from 3T3-L1 adipocytes by immunofluorescence analysis (Fig. 2C). In addition, a modest but significant decrease in the amount of GLUT4 and GLUT1 in the LDM fraction was observed after insulin treatment (P < 0.05 vs. basal) but not in response to DHEA (Fig. 2A and B). The increase in PM GLUT1 and GLUT4 induced by DHEA was not the consequence of higher levels of these transporters in the cell because total GLUT1 and GLUT4 protein levels were not altered by DHEA (Fig. 2D).

For assessing whether the glucose transport enhancement by DHEA was a metabolic response restricted to the 3T3-L1 adipocyte cell line or could also be observed in human fat cells, 2-deoxy-d-glucose uptake rates were measured in adipocytes isolated from abdominal subcutaneous fat tissue of normal subjects and incubated in the absence or presence of DHEA. As shown in Fig. 1B, DHEA treatment of human adipocytes resulted in a dose- and time-dependent augmentation of 2-deoxy-d-glucose uptake (P < 0.05 vs. basal). The glucose transport system was more sensitive to DHEA in human as compared with 3T3-L1 adipocytes, because a significant increase in glucose uptake occurred with DHEA concentrations as low as 0.1 μmol/l in human adipocytes (compare A and B in Fig. 1). Similar to the results in 3T3-L1 adipocytes, in human adipocytes, DHEA showed maximal effects on glucose transport after 120 min of treatment and was less effective than insulin (P < 0.05 vs. insulin; Fig. 1B). The DHEA-induced increase in glucose transport was associated with twofold higher GLUT4 levels in the PM fraction (P < 0.05 vs. basal; Fig. 3A and B). Insulin augmented GLUT4 levels in the PM to a greater extent than DHEA (P < 0.05) and, in contrast to DHEA, significantly decreased GLUT4 in the LDM (Fig. 3A and B). The protein content of GLUT1 in the PM was slightly higher in DHEA- and insulin-treated than in control adipocytes, but these changes did not reach statistical significance (Fig. 3A and B). DHEA did not modify the total levels of GLUT4 (Fig. 3C). The total levels of GLUT1 in human adipocytes were very low, in agreement with previously reported findings (22), and did not show any significant changes with DHEA treatment (Fig. 3C).

Insulin stimulation of glucose transport in adipocytes requires insulin receptor-mediated tyrosine phosphorylation of IRS-1 and IRS-2 and subsequent activation of PI 3-kinase (1,2). Thus, whether increased glucose uptake in response to DHEA was associated with increased insulin receptor and/or IRS tyrosine phosphorylation and PI 3-kinase activity was determined next. As expected, insulin stimulation of cells resulted in augmentation of insulin receptor, IRS-1 and IRS-2 tyrosine phosphorylation, association of p85 with IRS-1 and IRS-2, and increased PI 3-kinase activity in IRS-1 and IRS-2 immunoprecipitates (Fig. 4A and B). By contrast, DHEA treatment of adipocytes for various times had no effects on tyrosine phosphorylation of the insulin receptor (Fig. 4A). However, DHEA significantly enhanced tyrosine phosphorylation of IRS-1 and IRS-2 (respectively, 150 ± 5% and 180 ± 6% of basal; P < 0.05), induced the association of p85 with IRS-1 and IRS-2, and increased PI 3-kinase activity in IRS-1 and IRS-2 signaling complexes (Fig. 4B). As compared with insulin, DHEA was less effective in stimulating IRS-1 tyrosine phosphorylation (150 ± 5% vs. 220 ± 7% of basal in response to DHEA and insulin, respectively; P < 0.05) and the associated PI 3-kinase activity but equally effective on IRS-2 tyrosine phosphorylation (180 ± 6% and 190 ± 10% of basal in response to DHEA and insulin, respectively; NS) and IRS-2-associated PI 3-kinase activity (Fig. 4B). For investigating the possibility that DHEA activates insulin-signaling intermediates distal to PI 3-kinase, the phosphorylation of the serine/threonine kinase Akt was evaluated next. Whereas treatment of 3T3-L1 adipocytes with insulin for 15 min resulted in marked stimulation of Akt phosphorylation on both Thr-308 and Ser-473, treatment with 100 μmol/l DHEA for 5–120 min did not modify Akt phosphorylation (Fig. 4C). Furthermore, DHEA treatment did not activate Akt2, assessed by evaluating Thr-308 phosphorylation specifically in Akt2 immunoprecipitates (Fig. 4D), and did not induce phosphorylation of GSK-3β, which reflects Akt activity in intact cells (Fig. 4E). Finally, DHEA did not change the phosphorylation state of MEK and ERK-1/2 kinases, which were markedly activated in response to insulin (Fig. 4C). The levels of Erk-1 and Erk-2 phosphorylation were, respectively, 750 ± 47% and 588 ± 49% of basal after insulin stimulation (P < 0.05), 113 ± 23% and 113 ± 26% of basal after DHEA stimulation for 5 min (P = 0.64), and 87 ± 25% and 85 ± 26% of basal after DHEA stimulation for 15 min (P = 0.66).

Because DHEA was found to activate PI 3-kinase in IRS-1 and IRS-2 signaling complexes, the involvement of PI 3-kinase in the DHEA-induced augmentation of glucose transport was assessed next. For this purpose, 2-deoxy-d-glucose uptake rates were measured in 3T3-L1 adipocytes after pretreatment with PI 3-kinase inhibitors. Both Wortmannin and LY294002 completely abrogated DHEA-stimulated glucose uptake in 3T3-L1 adipocytes (P < 0.05 vs. DHEA control; Fig. 5A). In addition, these compounds markedly inhibited transport stimulation by insulin (P < 0.05 vs. insulin control; Fig. 5A), as reported previously (28). Pretreatment of adipocytes with LY294002 resulted in marked inhibition of the ability of DHEA to increase GLUT1 and GLUT4 in PM fractions (P < 0.05 vs. DHEA control; Fig. 5B and C). LY294002 also prevented insulin-induced translocation of GLUT1 and GLUT4 from the LDM to PM fractions (P < 0.05 vs. insulin control; Fig. 5B and C), as shown previously (28). Glucose transport stimulation by DHEA in 3T3-L1 adipocytes was not affected by preincubation of cells with PD098059, an inhibitor of MEK. Glucose transport (in pmol 2-DG · μg protein⁻¹ · min⁻¹) was 34 ± 2, 54 ± 12, 38 ± 3, and 56 ± 1 under basal conditions and after incubation with 100 μmol/l DHEA, 20 μmol/l PD098059, and 100 μmol/l DHEA plus 20 μmol/l PD098059, respectively. In addition, it was not affected by 100 nmol/l rapamycin, a p70S6 kinase inhibitor (data not shown). Thus, PI 3-kinase activity is required for DHEA-induced translocation of GLUT1 and GLUT4 and subsequent enhancement of glucose uptake by 3T3-L1 adipocytes.

DHEA has reportedly been shown to activate PKC in rat adipocytes (16,17). For evaluating the ability of DHEA to activate specific PKC isoforms in 3T3-L1 adipocytes, cytosolic and membrane fractions from DHEA- and insulin-treated cells were analyzed by immunoblotting with antibodies specific to PKC-α, PKC-β, and PKC-ζ/λ, three PKC isoforms that are expressed in 3T3-L1 adipocytes (29,30). DHEA induced membrane translocation of PKC-β, with maximal effect after 30 min (P < 0.05 vs. basal), and did not modify the distribution of either PKC-α or PKC-ζ/λ in cytosolic and membrane fractions (Fig. 6A). Because tyrosine phosphorylation reflects the activation state of specific PKC isozymes (31,32), PKC-β tyrosine phosphorylation was assessed in these experimental conditions by immunoprecipitation with antibodies against PKC-β₁ or PKC-β₂ followed by immunoblotting with phosphotyrosine antibody. DHEA induced tyrosine phosphorylation of PKC-β₂ (Fig. 6B), and this effect was abrogated by pretreatment of cells with the PKC-β inhibitor LY379196 (33) (Fig. 6B). Conversely, no effects of DHEA on tyrosine phosphorylation of PKC-β₁ were observed (data not shown). In contrast to DHEA, insulin promoted membrane translocation of PKC-α and PKC-ζ/λ and did not affect the cellular distribution of PKC-β (Fig. 6A) or tyrosine phosphorylation of PKC-β₂ (Fig. 6B) or PKC-β₁ (data not shown) in 3T3-L1 adipocytes.

Because membrane translocation of conventional PKC isoforms, such as PKC-β, follows changes in the intracellular concentrations and localization of Ca²⁺ (34), cytosolic-free Ca²⁺ concentrations were measured at multiple time points after treatment of 3T3-L1 adipocytes with 100 μmol/l DHEA. Cytosolic Ca²⁺ concentrations were increased by DHEA with an initial statistically significant effect at 33 min (P < 0.05 vs. control; Fig. 6C), which was concomitant with the DHEA-induced membrane translocation of PKC-β (Fig. 6A). For assessing whether the ability of DHEA to activate PKC-β required PI 3-kinase or phospholipase C-γ (PLC-γ) activity, membrane translocation of PKC-β in response to DHEA was assessed in adipocytes that had been pretreated with the PI 3-kinase inhibitor LY294002 or the PLC-γ inhibitor U73122 (35), respectively. DHEA stimulation of PKC-β translocation, which was observed in control cells, could not be demonstrated after preincubation with LY294002 or U73122 (Fig. 6D). As expected, PKC-β translocation in response to DHEA was prevented by the general PKC inhibitor bisindolylmaleimide GF109203X (36) and the specific PKC-β inhibitor LY379196 (Fig. 6D); however, it occurred normally in adipocytes that had been pretreated with the MEK inhibitor PD098059 (Fig. 6D). These results demonstrate that membrane translocation of PKC-β in response to DHEA requires the activities of PI 3-kinase and PLC-γ, as well as the activity of PKC-β itself, independent of MEK/MAP kinase activity.

Specific PKC isoforms and PLC-γ have been shown to mediate, at least in part, insulin stimulation of glucose transport in adipocytes (30,37,38). For investigating the potential involvement of PKC in the DHEA-dependent enhancement of glucose uptake, the effects of the general PKC inhibitor GF109203X on DHEA-stimulated 2-deoxy-d-glucose uptake and transporter translocation were examined. Pretreatment of 3T3-L1 adipocytes with 20 μmol/l GF109203X for 30 min abolished DHEA stimulation of glucose transport (P < 0.05 vs. control DHEA; Fig. 7A). However, insulin-stimulated glucose transport was inhibited 50% by this PKC inhibitor (P < 0.05 vs. control insulin; Fig. 7A). In a similar manner, GF109203X abrogated DHEA-induced translocation of GLUT4 and GLUT1 (P < 0.05 vs. DHEA control; Fig. 7B and C) and partially inhibited insulin stimulation of these responses (Fig. 7B and C). Insulin-stimulated GLUT4 and GLUT1 protein levels in PM fractions from GF109203X-treated adipocytes were 70 and 50%, respectively, of control (Fig. 7C). DHEA stimulation of glucose transport was also abrogated after treatment of 3T3-L1 adipocytes with 30 μmol/l staurosporine, another PKC inhibitor (data not shown). Thus, PKC activity is required for DHEA-dependent glucose transporter translocation and enhancement of glucose uptake. The role of PLC-γ in glucose transport stimulation by DHEA was investigated by using the specific PLC-γ inhibitor U73122. Pretreatment of 3T3-L1 adipocytes with 10 μmol/l U73122 for 30 min completely inhibited stimulation of 2-deoxy-d-glucose uptake by DHEA (P < 0.05 vs. control DHEA) and, as reported previously (35), reduced insulin-induced 2-deoxy-d-glucose uptake 50% (P < 0.05 vs. control insulin; Fig. 7A).

The potential contribution of distinct PKC isoforms to DHEA-induced glucose transport was evaluated by pretreating the adipocytes with Gö6976, an inhibitor of conventional PKCs, Calphostin C, an inhibitor of novel PKCs, myr-PKC-ζ/λ, an inhibitor of atypical PKCs, or the PCK-β inhibitor LY379196 (33,37–39). DHEA stimulation of glucose transport was completely blocked by pretreatment with 50 nmol/l Gö6976 and 30 nmol/l LY379196 (P < 0.05 vs. control DHEA), whereas Calphostin C and myr-PKC-ζ/λ had no effect (Fig. 8). By contrast, insulin-stimulated glucose transport was reduced 60% after pretreatment with myr-PKC-ζ/λ (P < 0.05 vs. control insulin), in agreement with previously reported results (38,39), and not altered by Gö6976, Calphostin C, or LY379196 (Fig. 8). Furthermore, DHEA stimulation of glucose transport was abrogated in the presence of 100 μmol/l myristoylated PKC-β inhibitory peptide, which had no effect on the insulin response (data not shown). Altogether, these results suggest that different PKC isoforms are involved in glucose transport stimulation by insulin and DHEA in 3T3-L1 adipocytes.

We show that DHEA is capable of inducing a rapid stimulation of cellular glucose uptake in both human and murine adipocytes by activating signaling responses that lead to glucose transporter translocation to the cell surface. The effects of DHEA occur within minutes of adipocyte exposure to this steroid hormone and do not involve changes in cellular content of GLUT1 or GLUT4. No stimulation of glucose uptake is observed in response to other steroid hormones. The rapid onset and specificity of DHEA action may be explained by DHEA binding to a specific DHEA receptor at the cell surface. Evidence for high-affinity DHEA binding to isolated plasma membranes from endothelial cells has recently been provided, and it has been shown that DHEA increases binding of GTPγS to Gα_i2,3, suggesting that the DHEA receptor may be coupled to this G protein (40). However, the molecular nature and tissue distribution of such DHEA receptors are still unknown. DHEA treatment of 3T3-L1 adipocytes resulted in IRS tyrosine phosphorylation and stimulation of PI 3-kinase activity in the absence of changes in tyrosine phosphorylation of the insulin receptor that could be detected by phosphotyrosine antibody immunoblotting. PI 3-kinase activation in an insulin-independent manner has been demonstrated in adipocytes after overexpression of the constitutively active Gα_i2, and it has been shown that this may occur via inhibition of protein tyrosine phosphatase 1B and subsequent enhancement of IRS tyrosine phosphorylation (41). This suggests that DHEA activation of the IRS/PI 3-kinase pathway may occur by DHEA binding to specific, although yet uncharacterized, Gα_i2-coupled cell-surface receptors.

The glucose transport enhancement by DHEA was associated with an approximately twofold increase in GLUT4 and GLUT1 content in adipocyte PM fractions. In addition, DHEA treatment resulted in increased GLUT4 and GLUT1 immunostaining in PM lawns from 3T3-L1 adipocytes. Evidence that DHEA enhances glucose uptake in rat adipocytes has recently been provided (16,17). However, these previous studies did not assess whether the effects of DHEA occur through translocation of glucose transporters or by an increase in transporter intrinsic activity. Oral administration of DHEA to insulin-resistant GK and OLETF rats for 2 weeks resulted in increased glucose uptake by adipocytes compared with untreated animals (17). In addition, DHEA has been shown to restore insulin sensitivity in obese Zucker rats (42) and to protect against the development of insulin resistance in rats fed a high-fat diet (14). Whereas the data in rodents clearly indicate a beneficial effect of DHEA on insulin sensitivity, studies in humans have yielded less clear results (43). For the first time, this study shows a stimulatory effect of DHEA on glucose uptake by human fat cells that occurs through GLUT4 translocation to the cell surface. In human adipocytes in vitro, DHEA was effective at the concentration of 100 nmol/l, which is close to the physiologic circulating DHEA levels in humans (0.7–15 nmol/l in childhood, 1.5–30 nmol/l in puberty, 5–50 nmol/l in adulthood, and 1–10 nmol/l after age 50). These findings may be important to foster further studies in human tissues, including the investigation of DHEA actions in adipose tissue from different body locations and/or insulin-resistant subjects.

In this study, maximal glucose transport stimulation by DHEA was lower compared with insulin, because the effects of DHEA were ∼30% and ∼60% of those of insulin in 3T3-L1 and human adipocytes, respectively. In addition, whereas DHEA and insulin were equally effective in inducing GLUT1 translocation to the PM, DHEA was less effective than insulin in promoting GLUT4 translocation and, in contrast to insulin, did not decrease GLUT4 in the LDM fraction. The lower magnitude of transport stimulation by DHEA compared with insulin thus may be explained by the lower extent of GLUT4 translocation to the PM in response to this steroid hormone, possibly as a result of differential regulation of specific intracellular transporter pools. Multiple other hormones and drugs, such as endothelin-1, arachidonic acid, and troglitazone, that are capable of stimulating glucose uptake by adipocytes also show a somewhat limited effectiveness on this metabolic response (6,7,9). The stimulation of glucose transport by arachidonic acid was found to be less than twofold over basal, this compound being less efficient than insulin in causing GLUT4 translocation to the PM (7). Similarly, endothelin-1 was shown to induce an approximately twofold increase in 2-deoxy-d-glucose uptake in 3T3-L1 adipocytes and to promote GLUT4 translocation to a lesser extent than insulin (6). These results suggest that not all of the intracellular GLUT4 vesicle pools are recruited by DHEA, arachidonic acid, or endothelin-1, respectively, and this may explain the relatively low glucose transport response to these molecules in adipocytes. Another potential explanation for the lower effect of DHEA on transport compared with insulin may involve differential regulation of transporter intrinsic activity. Compared with insulin, DHEA was less effective on transport by 70% and on GLUT4 translocation by 30%. Insulin may increase glucose transport in 3T3-L1 adipocytes by a mechanism that is independent of GLUT4 translocation to the cell surface (44). The lack of complete correlation between GLUT4 translocation and glucose transport in response to insulin and DHEA, respectively, may be potentially explained by DHEA causing GLUT4 translocation without increasing transporter intrinsic activity.

In 3T3-L1 adipocytes, DHEA was found to enhance PI 3-kinase activity in IRS-1 and IRS-2 signaling complexes but did not produce detectable activation of Akt, including Akt2 that is the presumably more relevant isoform for stimulation of GLUT4 translocation (45,46), as assessed by measurements of Akt and GSK-3 phosphorylation. In addition, DHEA did not activate MEK or ERK kinases. Akt activation can be triggered in the presence of very limited PI trisphosphate (PIP₃) levels, as it occurs after activation of some G protein-coupled receptors (47), or in cells treated with PI 3-kinase inhibitors in the presence of angiotensin (5). The mechanism of this PI 3-kinase-independent activation of Akt remains to be elucidated. Conversely, generation of PIP₃ in IRS signaling complexes may be not sufficient for Akt activation, as was observed in response to DHEA. Basic fibroblast growth factor, platelet-derived growth factor, and epidermal growth factor activate PI 3-kinase in vivo to a similar extent but differ in their ability to activate PI 3-kinase-dependent signaling, including Akt phosphorylation (48). These findings may be potentially explained by the existence of distinct subcellular compartments of active PI 3-kinase that are linked to specific downstream signaling molecules. Thus, the DHEA-induced PIP₃ generation may occur within a subcellular compartment excluding Akt, to which other PIP₃-regulatable signaling molecules, such as PLC-γ (49,50), are recruited upon stimulation.

Multiple PKC isoforms are activated by insulin in adipocytes. Insulin was shown to activate PKC-α and, to a lesser extent, PKC-β and PKC-ζ/λ in 3T3-L1 adipocytes (29,30) and PKC-ζ/λ in rat adipocytes (37). However, PKC-α and PKC-ζ activation by insulin has not been observed in all studies (51,52). In this study, insulin was found to affect the subcellular distribution of PKC-α and PKC-ζ/λ, whereas it did not influence translocation or tyrosine phosphorylation of PKC-β. By contrast, DHEA induced activation of PKC-β₂ but did not affect the α and ζ/λ isoforms. The ability of DHEA to activate specific classic isoforms of PKC provides an example of selective activation of PKC-β in the absence of detectable PKC-α or PKC-γ stimulation, and a similar selectivity has been reported in mesangial cells exposed to glucosamine (53). DHEA-induced tyrosine phosphorylation and membrane translocation of PKC-β were blocked by the PKC inhibitors GF109203X and LY379196, indicating that these effects require PKC-β activity. Inhibitors of PI 3-kinase and PLC-γ also blocked PKC-β activation by DHEA. It has recently been demonstrated that insulin-induced membrane translocation of PKC-β in rat skeletal muscle cells requires PI 3-kinase activity (32), but it is the first time that PKC-β translocation is found to be downstream of both PI 3-kinase and PLC-γ.

Inhibition of PI 3-kinase activity in 3T3-L1 adipocytes resulted in abrogation of glucose transport stimulation by DHEA and DHEA-induced translocation of GLUT4 and GLUT1. In addition, DHEA effects on the glucose transport system were prevented by a general PKC inhibitor and a PLC-γ inhibitor, both of which partially inhibited the effects of insulin by 50%. Finally, inhibitors of conventional PKCs and the specific PKC-β inhibitor blocked DHEA-induced glucose transport but had no effect on the transport response to insulin. By contrast, insulin stimulation of glucose transport was inhibited 50% by the myr-PKC-ζ/λ compound, which blocks atypical PKCs, in agreement with previous reports (37–39). Altogether, these results suggest that both insulin and DHEA stimulate glucose transport by activating PI 3-kinase and PLC-γ, the latter playing a partial role in the insulin effect. Signaling by these hormones may then diverge at the level of atypical or classical PKCs for insulin and DHEA, respectively. However, in one study in rat adipocytes (16), the Gö6976 compound was shown to have no effect on DHEA-stimulated glucose transport, which was blocked by the myr-PKC-ζ/λ compound. The opposite results found in our study could be potentially explained by species-related differences in signaling reactions regulating glucose transport. Indeed, in 3T3-L1 adipocytes, DHEA did not induce membrane translocation of PKC-ζ/λ, and its effect on transport was not prevented by the myr-PKC-ζ/λ compound.

In conclusion, DHEA treatment of human and 3T3-L1 adipocytes results in enhanced glucose transport rates through GLUT4 and GLUT1 transporter translocation to the cell surface. This effect seems to involve stimulation of IRS tyrosine phosphorylation and the associated PI 3-kinase activity. Then, generation of PIP₃ products in specific cell compartments may lead to the activation of PLC-γ resulting in increased calcium flux and PKC-β₂ activation. These processes depict a novel insulin-independent signaling pathway potentially relevant for the regulation of glucose uptake by fat cells.

This work was supported by grants from the Ministero dell’Istruzione, Università e Ricerca (Italy), the Cofinlab 2000-Centro di Eccellenza “Genomica comparata: geni coinvolti in processi fisiopatologici in campo biomedico e agrario” (Italy), the Ministero della Salute (Italy), the Società Italiana di Diabetologia (Italy), and an educational grant from Pfizer Italia srl (ARADO Program) to F. Giorgino.