Serum- and Glucocorticoid-Inducible Kinase 1 (SGK1) Mediates Glucocorticoid-Induced Inhibition of Insulin Secretion

MSD-D, a cardiac K_v1.5 channel inhibitor (patent no. WO9818475), and full-length hKv2.1 and hKv3.1 plasmids suitable for expression in Xenopus oocytes were a generous gift of Aventis Pharma Deutschland (Frankfurt, Germany). All other chemicals were from Sigma (Deisenhofen, Germany) and of analytical grade, unless otherwise stated.

Homozygous SGK1^(−/−) and SGK1^(+/+) littermate mice with 129/SvJ background were generated as previously described (30). Animal protocols were approved by the animal research committees of the respective institutions.

INS-1 cells (kindly provided by C.B. Wollheim, University of Geneva, Geneva, Switzerland) derived from a rat insulinoma were cultured in HEPES-buffered RPMI 1640 supplemented with 10% FCS (Biochrom, Berlin, Germany), 1 mmol/l HEPES, 1 mmol/l Na pyruvate, 10 μmol/l β-mercaptoethanol (Sigma, Munich, Germany), and antibiotics as described elsewhere (31,32). Cells were seeded at a cell density of 2.0–2.5 × 10⁵ cells/ml in 24-well culture plates and cultured for 2 days before the experiment. Cells were washed twice with incubation buffer containing (in mmol/l) 140 NaCl, 5.6 KCl, 1.2 MgCl₂, 2.6 CaCl₂, 0.5 glucose, 10 HEPES, and 5 g/l bovine serum albumin (fraction V; Sigma, Deisenhofen, Germany), pH 7.4, and preincubated for 30 min at 37°C. Thereafter, cells were incubated for 30 min at 37°C in fresh medium containing the test substances at the appropriate concentrations. Incubations were stopped on ice, and insulin released into the supernatant and insulin content after extraction with acid ethanol (1.5% [vol/vol] HCl/75% [vol/vol] ethanol) were measured by radioimmunoassay using rat insulin antiserum (Linco, Biotrend Chemikalien, Cologne, Germany), I¹²⁵-insulin (CIS Diagnostik, Dreieich, Germany), and rat insulin (Novo Nordisk, Mainz, Germany) as standard or an insulin enzyme-linked immunosorbent assay kit (Mercodia, Uppsala, Sweden).

Islets were isolated by collagenase digestion (1 mg/ml; Serva, Heidelberg, Germany) for 10 min at 37°C and isolated from exocrine tissue under the dissection microscope. Islets were cultured overnight in RPMI 1640 containing 11 mmol/l glucose. Dexamethasone (100 nmol/l) or DMSO (control) were added during the last 4 h of culture. Culture islets were then preincubated for 1 h at 37°C in incubation buffer containing 2.8 mmol/l glucose. Thereafter, batches of five islets/0.5 ml were incubated for 30 min at 37°C in the presence of test substances as indicated for each experiment.

INS-1 cells and mouse islet cells after digestion of isolated islets with trypsin (1 mg/ml) were cultured for 2–4 days at subconfluent cell densities. The cells were superfused with incubation buffer containing 0.5 mmol/l glucose. For patch clamp experiments, pipettes (Clark-Medical, Reading, U.K.) with a resistance of 4–6 MΩ were pulled using a DMZ-universal puller (Zeitz, Augsburg, Germany). They were filled with an internal solution containing (in mmol/l) 30 KCl, 95 K⁺ gluconate, 1 MgCl₂, 1.2 NaH₂PO₄, 4.8 Na₂HPO₄, 5 Na₂ATP, 1 Na₃GTP, and 5 EGTA, pH 7.2. An EPC9 patch clamp amplifier (Heka Electronic, Lambrecht, Germany) was used for current measurements. Only stable current measurements, i.e., when currents reached at least 90% of control current after removal of the respective inhibitory drug, were used for analysis. INS-1 cells were transfected with pIRES2SGK1 using a TransFast kit (Promega, Mannheim, Germany). One to 2 days after transfection, cells were treated with IGF-1 (50 ng/ml) for 4 h to activate SGK1 before patch clamp experiments were performed on green fluorescent protein-positive cells.

For [Ca²⁺]_i measurements, cells were loaded with fura-2/AM: 10 μmol/l, for 10 min at 37°C. Fluorescence measurements were performed using an inverted microscope (Axiovert 100 TV; Carl Zeiss Jena, Jena, Germany), a monochromator (Till Photonics, Martinsried, Germany), and a digital camera (Micromax 5 MHz; Princeton Instruments, Trenton, NJ). The fluorescence emission ratio at 340/380-nm excitation was calculated as a measure of [Ca²⁺]_i.

SGK1^+/+ and SGK1^−/− littermates were treated with dexamethasone for 24 h by subcutaneous injection of 10 mg/kg body wt. Pancreata were fixed with 4% phosphate-buffered paraformaldehyde. Subsequent sections of tissue were stained for SGK1 and insulin with rabbit anti-hSGK1 antibodies (1:400) and guinea pig anti-insulin serum antibodies (1:100; Linco, St. Charles, MO; diluted with PBS containing 0.3% Triton-X and 1% DMSO). Binding of anti-SGK1 antibodies was visualized with anti-rabbit antibodies conjugated to Alexa546, and anti-insulin antibody binding was visualized with anti-guinea pig conjugated to Alexa488 (molecular probes). Sections were analyzed on a confocal laser-scanning microscope (Zeiss LSM 510) using a He/Ne at 543 nm and an Ar-laser at 488-nm excitation wavelengths with appropriate filter sets. Unspecific staining was controlled by omitting the primary antibody.

INS-1 cells were cultured in 75-cm (2) flasks and treated with dexamethasone and RU486 as indicated. Cells were lysed, total RNA was isolated (Mini kit; Qiagen, Hilden, Germany), and 2 μg were transcribed into cDNA using RT M-MuLV (Roche Diagnostics, Roche Applied Science, Mannheim, Germany). Aliquots of cDNA, corresponding to equal amounts of RNA, were used for quantification of mRNA. The following specific primers were used: for detection of rat K_v1.5 channel mRNA: sense: 5′-ACTTCGCAGAGGCAGACAATCA-3′; antisense: 5′-GGTTGCCTTGTTCTTCCTTCAG-3′. For rat K_v2.1 channel: sense: 5′-GGATCCCCCGAAAAGGCCAG-3′ and antisense: 5′-GTTGGTCTCCGTGAGCCTCC-3′; for rat K_v2.2 channel: sense: 5′-GAGGTTAGCCAAAAAGACTC-3′ and antisense: 5′-TTGCAAAGGACCATGGGAAG-3′; and for rat K_v3.2 channel: sense: 5′-GCAAGCTCACCTACATTTTG-3′ and antisense: 5′-TCCATCAGAAGCGCACGTGT-3′. Primers used for rat SGK1 mRNA quantification were sense: 5′-TTTTTTTTCCCAACCCTTGC-3′ and antisense: 5′-AATGAACAAAGGTTGGGGGG-3′. SGK1 and K_v channel mRNAs were quantified using a light cycler system (Roche Diagnostics, Roche Applied Science, Mannheim, Germany). PCR reactions were performed in a final volume of 20 μl containing 2 μl cDNA, 3 mmol/l MgCl₂, 0.5 μmol/l of each primer, and 2 μl cDNA Master Sybr Green I mix (Roche Molecular Biochemicals, Mannheim, Germany). The transcript levels of the housekeeping gene glyceraldehyde-3-phsophate dehydrogenase (GAPDH) of each sample was taken as reference using a commercial primer kit (Search LC, Heidelberg, Germany). Amplification of the target DNA was performed during 35 cycles, each 10 s at 95°C, 10 s at 68°C, and 16 s at 72°C. Melting curve and agarose gel analyses confirmed the specificity of amplified products. Results were calculated as a ratio of the target versus housekeeping gene transcripts.

After cell culture with dexamethasone, mouse islets were collected and dissolved in lysis buffer (Mini kit; Qiagen, Hilden, Germany) and disrupted mechanically. RT-PCR for detection of SGK1 mRNA was performed using as sense primer: 5′-TGTCTTGGGGCTGTCCTGTATG-3′ and an antisense primer: 5′-GCTTCTGCT GCTTCCTTCACAC-3′.

INS-1 cells were cultured in 75-cm (2) flasks without (control) or with 100 nmol/l dexamethasone for the indicated period of time. Cells were lysed in a solution containing 300 mmol/l NaCl, 20 mmol/l Tris HCl, pH 7.4, 1% (vol/vol) Triton X-100, 1% sodiumdeoxycholate, 0.1% SDS, 2.5 mmol/l EDTA, 10 μg/ml pepstatin A, 10 μg/ml aprotinin, and 0.1 mmol/l phenylmethylsulfonyl fluoride. Total cellular protein, 50 μg quantified by Coomassie blue G staining (Bradford dye assay; Biorad Laboratories, Munich, Germany), was subjected to SDS-PAGE (10%) and blotted onto a nitrocellulose membrane (Schleicher and Schuell, Dassel, Germany). Blots were incubated with antibodies against SGK1 (New England Biolabs, Beverly, MA). Bound antibody was visualized using a secondary antibody coupled to horseradish peroxidase.

Dissection of Xenopus laevis ovaries has been described elsewhere (33). Oocytes were injected with 0.12 ng cRNA of human K_v1.5, K_v2.1, or K_v3.1 alone or together with 4 ng hSGK1 cDNA in a volume of 60 nl. Three days after injection, two-electrode voltage clamp recordings were performed at room temperature using a TurboTEC 10CX amplifier (NPI Electronics, Tamm, Germany). Currents were elicited with voltage steps from a holding potential of −60 to +60 mV for 3 s with an interval of 10 s. The data were filtered at 1 KHz and recorded with an interface DIGIDATA 1322a (Axon Instruments, San Diego, CA) and the software Clampex 9.0 (Axon instruments). The external control solution (superfusate/ND96) contained (in mmol/l) 96 NaCl, 2 KCl, 1.8 CaCl₂, 1 MgCl₂, and 5 HEPES, pH 7.4.

Data are presented as means ± SE. ANOVA for multiple groups and Student’s t tests were used for statistical analysis. P values <0.05 were accepted to indicate statistical significance.

To examine dexamethasone-induced transcription in insulin-secreting cells by semiquantitative RT-PCR, we have chosen the clonal rat insulinoma cell line INS-1 (34). We first examined whether dexamethasone affects transcription of K_v channels. Previously, we have found that INS-1 cells express a variety of K_v channels, including K_v1.5, K_v2.1, K_v2.2, and K_v3.2. Real-time PCR reveals that dexamethasone (100 nmol/l) treatment increases the abundance of K_v1.5 mRNA approximately sevenfold (Fig. 1A). The expression of other K_v channels, K_v2.1, K_v2.2, and K_v3.2, was not significantly stimulated by dexamethasone (Fig. 1B–D). In mouse islets treated for 4 h with 100 nmol/l dexamethasone, K_v1.5 mRNA increased fourfold from 0.11 ± 0.02 to 0.41 ± 0.05 copies K_v1.5/GAPDH*1,000 (n = 3, P < 0.005).

Next, we searched for SGK1 transcripts in INS-1 cells. According to real-time PCR, the SGK1 transcript level is low in untreated INS-1 cells (Fig. 2A). However, incubation of INS-1 cells with 100 nmol/l dexamethasone for 2–23 h increased mRNA transcript levels, an effect that was completely abrogated by the glucocorticoid receptor antagonist RU486 (Fig. 2A). Within 23 h, dexamethasone increased the cellular SGK1/GAPDH mRNA ratio by a factor of 146 ± 58 (range 28- to 275-fold, n = 4). When SGK1 transcript levels were measured in isolated mouse islets, SGK1 mRNA increased by a factor of 13.6 ± 3.4 (n = 6) following treatment with dexamethasone. This induction of SGK1 in mouse islets is larger than the sixfold increase of human SGK1 in epithelial cells (35).

To confirm the expression of SGK1 on protein level, Western blotting has been performed with INS-1. SGK1 protein was not detectable in untreated INS-1 cells but appeared within 2 h and increased further within the next 23 h of exposure to dexamethasone (100 nmol/l) (Fig. 2B). The increase in SGK1 protein was fully inhibited by RU486. Immunostaining of SGK1 in mouse pancreas revealed that SGK1 is expressed in insulin-containing cells of wild-type mice but not SGK1^−/− mice after dexamethasone treatment (Fig. 2C). These experiments demonstrate that dexamethasone stimulates the expression of both K_v1.5 and SGK1 in insulin-secreting cells. To analyze whether changes in K_v1.5 mRNA depend on synthesis of SGK1 protein, INS-1 cells were incubated with protein synthesis inhibitor cycloheximide. Incubation of the cells with 1 μmol/l cycloheximide for 20 h inhibited SGK1 protein synthesis completely (Fig. 3B) but led to a further increase of mRNA levels for K_v1.5 and SGK1 (Fig. 3A). Dexamethasone increased K_v1.5 mRNA in the absence as well as in the presence of cycloheximide. Thus, expression of the SGK1 protein is not required for the transcriptional stimulation of K_v1.5, pointing to a more direct effect of glucocorticoids on K_v1.5 transcription. Indeed, the K_v1.5 gene contains a glucocorticoid response element (29).

To examine whether SGK1 modulates K_v channel activities, we used the Xenopus leavis expression system. While water-injected Xenopus leavis oocytes did not show appreciable voltage-gated K⁺ currents (data not shown), those currents were elicited in oocytes injected with 0.12 ng cRNA encoding K_v1.5 (Fig. 4A). Coexpression of SGK1 and K_v1.5 doubled K_v1.5-mediated current, pointing to upregulation of K_v1.5 channel activity by SGK1. As shown earlier, SGK1 similarly stimulates K⁺ channels including K_v (17,36).

To characterize K_v1.5 currents in more detail, we used the K⁺ channel inhibitor MSD-D (Fig. 4B). MSD-D dose-dependently inhibited K_v1.5 channel currents with a K_i of 0.16 ± 0.02 μmol/l. The drug (0.3 μmol/l) had no effect on currents of hK_v2.1 and hK_v3.1 channels (data not shown). Thus, MSD-D is indeed more selective for K_v1.5 than 4-aminopyridine (4-AP) and tetraethylammonium (TEA) and about ∼1,000 times and 9,000 times more potent than 4-AP and TEA, respectively (25).

To examine whether K_v channel activity is increased after dexamethasone treatment, we analyzed outward currents in INS-1 cells, which are sensitive to TEA and 4-AP (25). The activation of K_v1.5 channels should result in a selective increase of 4-AP-sensitive outward current. As illustrated in Fig. 5A–C, treatment with dexamethasone increased 4-AP-sensitive voltage-gated outward current. In untreated cells, the K⁺ channel blocker 4-AP inhibited 10% (at 0.1 mmol/l) and 28% (at 1 mmol/l) of outward current. Following a 4-h treatment with 100 nmol/l dexamethasone, the 4-AP-sensitive current increased to 28% (by 0.1 mmol/l 4-AP) and 40% (by 1 mmol/l 4-AP) of total current. TEA inhibited significantly more current at 1 mmol/l but not at 10 mmol/l after dexamethasone. The role of SGK1 was further examined in cells overexpressing hSGK1. INS-1 cells were transiently transfected with hSGK1 and the enzyme activated by 50 ng/ml IGF-1 (Fig. 5D and ref. 37). In cells expressing hSGK1, 1 mmol/l 4-AP inhibited 29.2% of whole-cell current, whereas in cells transfected with the empty GFP vector, it inhibited 26.7% of current (P < 0.03).

In mouse islet cells treated with dexamethasone, whole-cell current was significantly larger, and 1 mmol/l 4-AP inhibited 28% of outward current and 21% in control cells. These data suggest that parallel to increased expression, the activity of K_v1.5 channels is augmented by dexamethasone and activated by SGK1 in insulin-secreting cells.

To examine whether dexamethasone alters [Ca²⁺]_i, INS-1 cells were loaded with fura-2 after a 4-h treatment with 100 nmol/l dexamethasone or DMSO (0.1%, control). Average fluorescence ratio (340/380 nm) reflecting Ca²⁺ concentration was at 0.5 mmol/l glucose, 0.86 ± 0.03 in control cells, and not significantly modified by dexamethasone treatment (0.87 ± 0.04). The addition of glucose led to the appearance of irregular Ca²⁺ spikes, with a minor increase of average fluorescence ratio in both control (1.05 ± 0.05) and dexamethasone-treated (1.01 ± 0.01) cells. Peak values of Ca²⁺ oscillations (reaching values >1.2) were observed in 29.0 ± 2.8% of control (166 cells of n = 3 independent experiments) and in 19.7 ± 4.2% of dexamethasone-treated (145 cells of n = 3 independent experiments, Fig. 6) cells. Large magnitude [Ca²⁺]_i spikes (reaching a ratio value >1.4) were significantly more frequent in control (18.0 ± 2.3%) than in dexamethasone-treated (4.0 ± 1.8%) cells. This observation suggests that increased K_v channel activity results in a reduction of Ca²⁺ peaks after dexamethasone. The difference was not reflected by changes of average Ca²⁺ concentrations.

The following experiments were performed to elucidate the impact of K_v channels on the blunting of insulin release by dexamethasone. As illustrated in Fig. 7A–C, pretreatment of INS-1 cells with dexamethasone (100 nmol/l) significantly (*P < 0.05) inhibited glucose-induced insulin secretion. This inhibition was reversed by K_v channel blockers 10 mmol/l TEA and 5 mmol/l 4-AP, suggesting that dexamethasone-mediated inhibition of insulin secretion depends on K_v channel activity (Fig. 7A). However, higher concentrations of K⁺ channel inhibitors were required to counteract secretion than those significantly decreasing whole-cell current. Therefore, a more selective K_v1.5 channel inhibitor, MSD-D, was used in the following. The inhibition of K_v1.5 channel activity by 0.3 μmol/l MSD-D had no effect on secretion at low glucose but augmented glucose-stimulated insulin secretion (#P = 0.008). MSD-D reversed the inhibitory effects of dexamethasone on glucose-induced insulin release (§P = 0.0002, Fig. 7B). Next, insulin secretion was stimulated in the presence of 30 mmol/l KCl and diazoxide, thus bypassing K⁺ channel regulation (Fig. 7C). Glucose at high KCl significantly (#P = 0.004) stimulated secretion to the same extent in control and dexamethasone-treated cells. These data support the idea that increased K_v channel activity is involved in dexamethasone-mediated inhibition of glucose-induced insulin secretion. Finally, we tested whether inhibition of K_v channels counteracts dexamethasone-induced inhibition of insulin secretion in mouse islets as it does in INS-1 cells. Glucose significantly (§P = 0.0085) stimulated secretion fourfold in untreated but not in dexamethasone-treated mouse islets. In the presence of MSD-D (0.3 μmol/l), glucose stimulated secretion 14-fold (#P = 0.0028). MSD-D completely reversed the inhibition of secretion by dexamethasone (^aP = 0.0054, Fig. 7D). Thus, as in INS-1 cells, inhibition of K_v1.5 channels improves glucose-induced insulin secretion in dexamethasone-treated mouse islets.

To estimate the contribution of SGK1 to the inhibitory effect of dexamethasone on insulin secretion, we studied the effects of dexamethasone in SGK1 knockout mice (SGK1^−/−) compared with that in wild-type littermates (SGK1^+/+). Without dexamethasone pretreatment, insulin secretion following exposure to glucose (16.7 mmol/l), activation of adenylate cyclase (5 μmol/l forskolin), or stimulation of protein kinase C (100 nmol/l phorbol myristic acid) was not significantly different in islets isolated from SGK1^−/− and SGK1^+/+mice (Fig. 8A and B, black bars). Dexamethasone treatment, however, significantly decreased the stimulatory effect of glucose, forskolin, or phorbol myristic acid on insulin secretion in SGK1^+/+ islets but not in SGK1^−/− islets. These data strongly suggest that SGK1 participates in the downregulation of insulin secretion by dexamethasone.

We demonstrate here that the synthetic glucocorticoid dexamethasone induces the expression of SGK1 in insulin-secreting cells. This kinase is only slightly expressed in untreated β-cells (Fig. 1), and low transcript levels have been reported previously for human pancreatic islets (38). The expression after glucocorticoid treatment of INS-1 cells is high; the amount of SGK1 mRNA reaches 80% of the abundance of GAPDH mRNA. Similarly, strong stimulation of SGK1 transcription by glucocorticoids was observed in other cell types (39,40). In native mouse islets, the copy numbers of SGK1 mRNA compared with GAPDH mRNA were much smaller. However, in both cell preparations, INS-1 cells and mouse islets, SGK1 protein was detectable after dexamethasone treatment. The amount of protein detected by western blotting also increased with time. This induction of SGK1 expression may contribute to the phenotype of impaired glucose-induced insulin secretion after glucocorticoid treatment and may contribute to the development of diabetes.

The endogenous cortisol plasma concentrations are governed by a circadian rhythm, reaching maximal concentrations of 700 nmol/l in the morning (41). Dexamethasone concentrations used for studying metabolic parameters range from 10 nmol/l to 1μmol/l, concentrations encountered in glucocorticoid excess (6,42,43). Dexamethasone is ∼30 times more potent than cortisol. We have chosen a concentration (100 nmol/l) that significantly and reproducibly inhibited insulin secretion in a variety of systems including INS-1 cells and isolated mouse islets (ref. 6 and Fig. 7 and 8).

The pleiotropic effects of glucocorticoids on insulin-secreting cells include an increase of α₂-adrenoceptor mRNA and binding sites (2,9) and a decrease of Glut2 transporter expression at the plasma membrane (8,11). Both proteins are regulated at a transcriptional and a posttranslational level. Here, we present two additional proteins, SGK1 and K_v1.5 channels, whose expression is regulated by dexamethasone at the transcriptional level. Both genes contain a glucocorticoid responsive element in their respective promoter region (15,29). Increased transcription is accompanied by increased translation of SGK1 as judged by Western blotting. K_v1.5 channels are minor components of plasma membrane proteins of insulin-secreting cells. Although the protein was detected by Western blotting, the abundance was too low to be quantified, since detection was largely limited by the quality of the antibody (data not shown). Functionally, 4-AP-sensitive outward currents were increased in INS-1 cells, suggesting increased K_v1.5 channel activity after dexamethasone treatment. K_v1.5 currents have been described to be insensitive to TEA (27). Since, under control conditions, the major part of voltage-activated outward current is TEA sensitive, it is afforded by K_v2.1 and/or K_v2.2. and K_v3.4 (22,24). The effect of dexamethasone was reflected by a decrease of Ca²⁺ spikes rather than a decrease of average Ca²⁺ concentration. It should be kept in mind that the local increase of Ca²⁺ and not the mean cytosolic Ca²⁺ concentration matters for insulin release (44,45). The mechanism by which such changes in [Ca²⁺]_i oscillations might affect insulin secretion remains to be elucidated. In mouse islets, K_v1.5 mRNA was indeed low under control conditions. A fourfold increase of K_v1.5 mRNA levels after dexamethasone treatment might, however, change the ratio of K_v channel expression and function. Changes in the function of K_v channels have previously been described to alter glucose-induced insulin release (24). We thus propose that changes in K_v1.5 channel function might contribute to impaired insulin secretion after dexamethasone treatment. Indeed, blocking K_v channels in INS-1 cells and mouse islets counteracts dexamethasone-induced inhibition of insulin secretion. The sensitivity of insulin secretion to 4-AP and TEA is lower than that of whole-cell currents, indicating that other mechanisms may contribute to impairment of release. Dexamethasone did not inhibit secretion induced by 15 mmol/l glucose in the presence of 30 mmol/l KCl and 0.25 μmol/l diazoxide, i.e., when K⁺ channel regulation was bypassed. This observation strongly suggests that dexamethasone influences insulin release indeed by modifying K⁺ channel activity. Previously, Henquin et al. (6) described that dexamethasone inhibits secretion induced by glucose and KCl in mouse islets. Thus, under the experimental conditions of these previous experiments, dexamethasone may exert additional inhibitory effects on insulin release that are independent from K⁺ channel activation. Whether these additional mechanisms are mediated by SGK1 cannot be answered from our observations.

Pertussis toxin and cAMP have been found to counteract dexamethasone inhibition of insulin secretion (6). Thus, incretins and glucagon, i.e., mediators increasing cAMP in β-cells might have a protective effect against glucocorticoid-induced diabetes. Whether an impaired function of incretins is responsible for increased susceptibility to glucocorticoid-induced diabetes especially in humans needs further examinations. Indeed, incretins not only potentiate glucose-induced insulin secretion but also activate cell proliferation and inhibit apoptosis (46).

The involvement of SGK1 in dexamethasone-induced inhibition of insulin secretion is suggested by experiments in SGK1^−/− islets. Dexamethasone did not significantly affect secretion from islets of SGK1^−/− mice, while secretion of the wild-type littermates was impaired. These results clearly show that SGK1 activation is involved in dexamethasone-induced inhibition of insulin secretion.

Impairment of insulin release contributes to but presumably does not fully account for the diabetic effect of glucocorticoid excess. The actions of glucocorticoids also include stimulation on hepatic glucose mobilization and peripheral insulin resistance (4).

In conclusion, the present experiments consider two proteins that are upregulated by glucocorticoids in insulin-secreting cells. The glucocorticoid dexamethasone enhances the transcription and expression of SGK1 and K_v1.5 channels. The kinase is able to activate K_v1.5 channels. Overexpression of K_v channels hyperpolarizes the β-cell plasma membrane, thus impeding the activation of voltage-gated Ca²⁺ channels. Accordingly, the kinase may contribute to the inhibition of insulin release during glucocorticoid excess and thus to the well-known aggravation of diabetes by stress conditions (47). Whether SGK1 is also involved in translational changes of α₂-adrenoceptors and Glut2 is an attractive hypothesis and should be examined.

The study was supported by DFG (German Research Foundation) Grants UL 140/4-1 and LA 315/4-6, a grant from the German Diabetes Society, and grants from the Köln Fortune and Tübingen Fortüne programs. S.U. was a recipient of a Heisenberg fellowship of the DFG (UL 140/2-1).

We are grateful to Sanofi-Aventis for providing us with MSD-D, hKv2.1, and hKv3.1.