Reduction of plasma membrane glutamate transport potentiates insulin but not glucagon secretion in pancreatic islet cells

Molecular and Cellular Endocrinology 338 (2011) 46–57

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Reduction of plasma membrane glutamate transport potentiates insulin but not glucagon secretion in pancreatic islet cells Nicole Feldmann a , Rafael Martin del Rio b , Asllan Gjinovci a , Jorge Tamarit-Rodriguez c , Claes B. Wollheim a,∗ , Andreas Wiederkehr a,∗ a b c

Department of Cell Physiology and Metabolism, University Medical Centre, 1, rue Michel Servet, 1211 Geneva 4, Switzerland Research Department, Hospital ‘Ramon y Cajal’, Madrid 28034, Spain Biochemistry Department, Medical School, Complutense University, Madrid 28040, Spain

a r t i c l e

i n f o

Article history: Received 6 January 2011 Received in revised form 21 February 2011 Accepted 21 February 2011 Keywords: Alpha-cells Beta-cells ␣-Ketoglutarate Excitatory amino acid transporter Glutamine GABA Malate–aspartate shuttle

a b s t r a c t Glutamate is generated during nutrient stimulation of pancreatic islets and has been proposed to act both as an intra- and extra-cellular messenger molecule. We demonstrate that glutamate is not cosecreted with the hormones from intact islets or purified ␣- and ␤-cells. Fractional glutamate release was 5–50 times higher than hormone secretion. Furthermore, various hormone secretagogues did not elicit glutamate efflux. Interestingly, epinephrine even decreased glutamate release while increasing glucagon secretion. Rather than being co-secreted with hormones, we show that glutamate is mainly released via plasma membrane excitatory amino acid transporters (EAAT) by uptake reversal. Transcripts for EAAT1, 2 and 3 were present in both rat ␣- and ␤-cells. Inhibition of EAATs by l-trans-pyrrolidine-2,4-dicarboxylate augmented intra-cellular glutamate and ␣-ketoglutarate contents and potentiated glucose-stimulated insulin secretion from islets and purified ␤-cells without affecting glucagon secretion from ␣-cells. In conclusion, intra-cellular glutamate-derived metabolite pools are linked to glucose-stimulated insulin but not glucagon secretion. © 2011 Elsevier Ireland Ltd. All rights reserved.

1. Introduction Insulin secretion is stimulated by an increase in glucose concentration, which is associated with a suppression of glucagon release (Franklin et al., 2005; Olsen et al., 2005; Gromada et al., 2007; Henquin, 2009). The secretion of these hormones is regulated not only by nutrients but further modulated by a large number of hormones, neurotransmitters and metabolites that act either intraor extra-cellular. For instance, glutamate synthesised from glucose in the pancreatic ␤-cells has been proposed to participate in the full activation of the insulin secretory response (Maechler and Wollheim, 1999; Rubi et al., 2001; Hoy et al., 2002; Carobbio et al., 2009). However, the role of glutamate in insulin secretion remains controversial (MacDonald and Fahien, 2000; Bertrand et al., 2002). It has been shown that under permissive conditions glutamate itself or a membrane-permeant glutamate derivative stimulates insulin exocytosis, while being inefficient under non-stimulatory condi-

Abbreviations: EAATs, excitatory amino acid transporters; GSIS, glucosestimulated insulin secretion; ␣KIC, alpha-ketoisocaproate; trans-PDC, l-transpyrrolidine-2,4-dicarboxylate. ∗ Corresponding authors. Tel.: +41 22 3795548; fax: +41 22 3795260. E-mail addresses: [email protected] (C.B. Wollheim), [email protected] (A. Wiederkehr). 0303-7207/$ – see front matter © 2011 Elsevier Ireland Ltd. All rights reserved. doi:10.1016/j.mce.2011.02.019

tions (Bertrand et al., 1992; Rubi et al., 2001; Hoy et al., 2002). The amino acid may thus potentiate nutrient-stimulated insulin secretion in a process referred to as the amplifying pathway (Maechler et al., 2006; Henquin, 2009). Other reports suggest that glutamate functions as an extracellular signal in pancreatic islets, similar to its role in excitatory synapses (Hayashi et al., 2003). Functional intercellular glutamate signalling requires a glutamate system, comprised of the following components: (I) vesicular glutamate transporters (VGLUT 1–3) to store glutamate in synaptic vesicles, (II) glutamate receptors to bind the amino acid and transmit the signal, and (III) an electrogenic plasma membrane glutamate transport system mediated by excitatory amino acid transporters (EAATs 1–5) for the reuptake of the transmitter. These transporters maintain extra-cellular glutamate concentrations low and terminate the process of synaptic transmission (Moriyama and Hayashi, 2003). Except for EAAT expression in ␣- and ␤-cells, all the components necessary for extra-cellular glutamate signalling have been described in pancreatic islets (Inagaki et al., 1995; Weaver et al., 1996, 1998; Storto et al., 2006; Cabrera et al., 2008). For instance, expression of vesicular glutamate transporters was demonstrated in clonal ␣- and ␤-cell lines and in native ␣-cells (Hayashi et al., 2003; Moriyama and Hayashi, 2003; Storto et al., 2006). Based on these results, it has been proposed that glutamate can be taken up into secretory granules for co-storage and co-release with insulin and/or glucagon (Hayashi et al., 2003;

N. Feldmann et al. / Molecular and Cellular Endocrinology 338 (2011) 46–57

Moriyama and Hayashi, 2003). Once secreted, glutamate may then bind to its receptors acting in an autocrine and paracrine manner to modulate exocytosis from ␣- and ␤-cells. Recent studies suggest that glutamate receptor activation stimulates glucagon but not insulin secretion in mouse islet cells (Cabrera et al., 2008; Cho et al., 2010). The present study was designed to elucidate the pathways of glutamate release from pancreatic islets as well as purified ␣- and ␤-cells, the main two islet cell types. We find that glutamate release is mainly mediated via the export through EAATs rather than granule exocytosis. We studied the relevance of this release mechanism on intra-cellular metabolite levels (glutamate and ␣-ketoglutarate) and glucose-stimulated insulin secretion (GSIS). 2. Methods 2.1. Ethical approval Animal care and experimentation were conducted according to the guidelines of the Swiss Academy of Medical Sciences and performed with the permission of the Canton of Geneva Veterinary Office. 2.2. Preparation and culture of rat islets and FACS-purified ˛- and ˇ-cells Male Wistar rats were obtained from Janvier (Le Genest-St-Isle; France). Rats were anesthetized with sodium pentothal 100 mg/kg body weight (intraperitoneal). Islet isolation and purification of islet cells was carried out as described previously (Franklin et al., 2005). Purified ␣- and ␤-cells were seeded onto polyornithinecoated 24-well plates (10,000 cells/well). Sorted cells and islets were cultured overnight in RPMI-1640 medium supplemented with 4 mM glucose, 10% (vol/vol) heat-inactivated foetal calf serum, 100 U/ml penicillin, 100 ␮g/ml streptomycin, 100 ␮g/ml gentamycin and 10 mM HEPES at 37 ◦ C in a humidified atmosphere. 2.3. Hormone assays For the glutamate release study groups of 20 islets were used. In another series using 200 islets per condition amounts of intra-cellular ␣-ketoglutarate and glutamate were determined. Islets and purified ␣- and ␤-cells were washed twice with Krebs Ringer bicarbonate HEPES (KRBH) buffer (Franklin et al., 2005) containing [mM] 130 NaCl, 3.6 KCl, 0.5 NaH2 PO4 , 0.5 MgSO4 , 1.5 CaCl2 , 10 HEPES, 5 NaHCO3 , 2.5 glucose and 0.05% BSA (fractionV; AppliChem, Darmstadt, Germany). After the second wash, cells were incubated at 37 ◦ C for 30 min with KRBH buffer (200 ␮l) supplemented with glucose or other additions as indicated in the figures. All chemicals were from Sigma Chemical Co. (Buchs, Switzerland). At the end of the incubation, supernatants were collected and hormone and glutamate release was determined. For the glutamate release and content, attached cells or whole islets were washed twice with 100 mM Tris–HCl pH 7.5, if applicable detached mechanically and lysed by sonication. Glucagon and insulin were assessed respectively by radioimmunoassay (Franklin et al., 2005) using anti-glucagon serum (Dako Diagnostics, Zug, Switzerland) and by enzyme immunoassay (SPI bio, Montigny le Bretonneux, France). For the combined determination of intra-cellular ␣-ketoglutarate and glutamate contents, islets were washed twice with cold PBS w/o Ca2+ and Mg2+ , thereafter 100 ␮l of 0.6 M perchloric acid (PCA) was added. Islets were disrupted by sonication and metabolites were extracted on ice for at least 1 h. Cell debris and proteins were centrifuged. Cell pellets were dissolved in protein lysis buffer and sonicated. The supernatant containing extracted metabolites was neutralized with 2.7 M K2 CO3 . Precipitates were removed by centrifugation. Cleared metabolite extracts were used for ␣-ketoglutarate and glutamate determination.

47

Table 1 Glucose-induced changes on glutamate, glutamine and alanine in rat islets. Amino acid

2.5 mM glucose

Glutamate (enzyme cycling assay) Glutamate (HPLC) Glutamine (HPLC) Alanine (HPLC)

143.6 102.4 32.8 91.0

± ± ± ±

30.7 17.1 5.6 7.2

16.7 mM glucose 263.3 141.7 47.1 70.8

± ± ± ±

4.4a 22.2b 7.1b 6.7

Rat islets were incubated for 30 min at 37 ◦ C in KRBH buffer containing 2.5 mM or 16.7 mM glucose. Total glutamate, glutamine and alanine in rat islets [pmol/20 islets] were measured with HPLC. For comparison total glutamate determined with the enzyme cycling as shown in Fig. 1 are included. Average values ± S.E.M. are shown. The HPLC results are from 3 independent experiments performed in triplicate. Total glutamate measured by enzyme cycling assay are from the average of the 4 experiments. a p < 0.01 (two-tailed Student’s t-test for unpaired data). b p < 0.01 (two-tailed Student’s t-test for paired data).

were collected. Islets and ␤-cells were washed twice with ice-cold PBS and extracted with 100 ␮l of 35% (w/v) 5-sulfosalicylic acid. 2.5. ˛-Ketoglutarate assay The ␣-ketoglutarate assay is based on two consecutive enzymatic reactions as described (Gamberino et al., 1997; Hazen et al., 1997). In the presence of NAD, NADH is formed stoichiometrically to the ␣-ketoglutarate initially present. The reaction mixture for NADH generation from ␣-ketoglutarate contained: 100 mM KH2 PO4 pH 7.0, 1 mM EGTA pH 8.0; 0.91 mM CaCl2 , 0.05 mM NAD, 0.5 mM dithiothreitol (DTT), 0.025 mM coenzyme A and 5 U/ml ␣-ketoglutarate-dehydrogenase (Sigma Chemical Co., Buchs, Switzerland). 20 ␮l of ␣-ketoglutarate standard (5–100 pmol), diluted in K2 CO3 neutralized 0.6 M PCA, or sample was added to 80 ␮l of the reaction mixture. After 1 h incubation at room temperature, 10 ␮l of this mixture was diluted to 50 ␮l with H2 O and loaded together with 50 ␮l of luciferase reaction mixture into the luminometer. Maximal light emission was recorded 2 min after each addition. The luciferase reaction mixture contained 50 mM KH2 PO4 pH 7.0, 0.2 mM DTT, 10 mg/L photobacteria Fischeri luciferase (Roche Applied Science, Rotkreuz, Switzerland), 160 U/L FMN reductase (NAD(P)H:FMN oxidoreductase; Roche Applied Science, Rotkreuz, Switzerland), 5 ␮M FMN, 0.01% Triton X-100 (Sigma) and 48 ␮M mysteric aldehyde (tetradecanal; ChemSampCo Inc., Trenton, NJ, USA). 2.6. Quantitative real time-PCR (QT-PCR) Total RNA was extracted from FACS-purified pancreatic ␣- and ␤-cells. 0.5 ␮g RNA was converted into cDNA as previously described (Gauthier et al., 1999). Primers for cyclophilin, EAAT1, EAAT2 and EAAT3 were designed using the Primer Express Software (Applera Europe, Rotkreuz, Switzerland). Primers were obtained from Eurogentec (Seraing, Belgium). QT-PCR was performed using an ABI 7000 Sequence Detection System (Applera Europe) and PCR products were quantified using the SYBR Green Core Reagent kit (Gauthier et al., 2004). Three independent experiments were performed in duplicate for each transcript and mean values were normalized to the corresponding cyclophilin value. 2.7. Statistical analysis Results are mean ± S.E.M. Static incubations were performed in triplicate and averaged for each condition. n values refer to separate experiments. The statistical significance of the difference between two groups was assessed by two-tailed Student’s t-test.

3. Results

2.4. Glutamate measurements Glutamate was measured with a modified fluorescence enzyme cycling assay monitoring Amplex Red oxidation (Chapman and Zhou, 1999). Cleared lysates were incubated in the presence of 50 ␮M Amplex Red, 0.04 U/ml glutamate oxidase, 0.25 U/ml glutamic–pyruvic transaminase, 100 ␮M l-alanine, and 0.125 U/ml HRP, in 100 mM Tris–HCl pH 7.5 in a total reaction volume of 70 ␮l. The analysis was performed in black 96 well plates (Greiner bio-one; Huber and Co., AG, Reinach, Switzerland). Fluorescence was measured in a plate reader (Flexstation; Bucher Biotec, Basel Switzerland) every minute during 30 min incubation at 37 ◦ C. The excitation wavelength was 530 nm and the fluorescence emission was monitored at 590 nm. Amplex Red was obtained from Molecular Probes (Basel, Switzerland). All other chemicals were purchased from Sigma Chemical Co. (Buchs, Switzerland). Glutamate measurements were verified by reverse-phase HPLC after pre-column derivatization with o-phthaldialdehyde and quantified by fluorescence detection (Hernandez-Fisac et al., 2006). Glutamine, alanine and GABA were measured by the same HPLC method (Hernandez-Fisac et al., 2006). For glutamate release study 20 islets or 20,000 purified ␤-cells were used. At the end of the incubation, supernatants

3.1. Glucose stimulates glutamate generation in islets and purified ˛- and ˇ-cells Glutamate synthesis and handling was studied in rat islets and purified ␣- and ␤-cells using a fluorescence enzyme cycling assay (Section 2.4). The method was validated by determining glutamate in a set of parallel experiments using HPLC. Both methods revealed glucose-dependent increases of total glutamate levels in islets (Table 1) and purified ␤-cells (Table 2). The similar results obtained with the two methods confirm the reliability of the fluorescence enzyme cycling assay. Measurements by HPLC of other amino acids revealed that alanine was not significantly changed during glucose stimulation of islets or ␤-cells. The concentration of glutamine in islets was much lower than that of glutamate but

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N. Feldmann et al. / Molecular and Cellular Endocrinology 338 (2011) 46–57

Table 2 Glucose-induced changes on glutamate, glutamine and alanine in purified ␤-cells. Amino acid

2.5 mM glucose

Glutamate (enzyme cycling assay) Glutamate (HPLC) Glutamine (HPLC) Alanine (HPLC)

146.1 171.1 605 147.4

± ± ± ±

22 11.5 79 34.5

16.7 mM glucose 169.7 197 556 148.5

± ± ± ±

21.2a 8.3a 155.7 27.8

Purified ␤-cells were incubated for 30 min at 37 ◦ C in KRBH buffer containing 2.5 mM or 16.7 mM glucose. Total glutamate, glutamine and alanine in purified ␤-cells [pmol/10,000 cells] were measured with HPLC. For comparison glutamate determined with the enzyme cycling as used in the rest of this study are also shown. Average values ± S.E.M. are shown. The HPLC results are from 3 independent experiments performed in triplicate. Total glutamate measured by enzyme cycling assay are from the average of the 11 experiments. a p < 0.05 (two-tailed Student’s t-test for paired data).

was also significantly elevated by 16.7 mM glucose. In contrast, glutamine levels in purified ␤-cells were not affected by glucose and glutamine concentrations in these samples were very high. The latter result should be taken with caution as it may reflect carry-over from the culture medium containing 2 mM glutamine. Consistent with this interpretation, glutamine levels were 50 times lower in ␤-cells incubated in medium without glutamine prior to the experiment (data not shown). 3.2. A large fraction of total glutamate is released from islets as well as purified ˛- and ˇ-cells at both resting and stimulatory glucose concentrations Raising the glucose concentration from 2.5 mM to 16.7 mM, increased insulin secretion from rat islets (Fig. 1C) and elevated the islet glutamate content (Fig. 1A) without significantly augmenting glutamate release (Fig. 1B). The fraction of total glutamate released from the islets was much higher than the percentage of insulin secreted. At basal glucose concentration for example 17.7 ± 3.6% glutamate but only 0.33 ± 0.15% insulin was secreted (Fig. 1). As ␣- and ␤-cells constitute respectively 20% and 65% of the rat islet (Baetens et al., 1979; Gromada et al., 2007) and may handle glutamate differently, we studied glutamate production after FACS purification of these cell types. Glucose caused an elevation of intra-cellular glutamate by 44.2 ± 12.9% and 34.3 ± 10.3% in purified ␣- and ␤-cells, respectively (Fig. 2A and D). Glucose stimulation did not significantly affect glutamate release from isolated ␤-cells (Fig. 2B). In contrast, elevated amounts of the amino acid were determined in the supernatant of ␣-cells exposed to high glucose (Fig. 2E). In both ␣- and ␤-cells, more than 60% of the total glutamate was secreted at basal and stimulatory glucose con-

3.3. Glutamate is not primarily formed from glutamine during glucose stimulation Several pathways may contribute to glucose-dependent glutamate production and to replenish intracellular glutamate pools lost as a result of glutamate release. One such pathway is glutaminase catalysed hydrolysis of glutamine to glutamate (Malaisse et al., 1982). Glutamine by itself does not act as a secretagogue but exerts a potentiating effect on insulin secretion induced by other secretagogues (Malaisse et al., 1982; Bertrand et al., 2002; Li et al., 2004). To test the role of glutamine on glutamate production, islet cells were cultured in medium without glutamine 5 h prior to the experiment and compared to cells cultured in medium containing 2 mM glutamine (standard condition). The presence of glutamine in the culture medium did neither affect glutamate content in ␣- (Fig. 4A) nor ␤-cells (Fig. 4B) following glucose stimulation. In rat islets glutamate levels as assessed by HPLC had a tendency to increase when cells were cultured in 2 mM glutamine as compared to 0 mM

B

C

300

200

100

30

insulin secretion (% of total)

**

glutamate release (pmol/20islets 30 minutes)

glutamate content (pmol/20islets)

A

centrations. This fraction is smaller in intact islets, likely because ␣-cells take up extra-cellular glutamate from the interstitial space as suggested previously (Weaver et al., 1998). In agreement with published results, insulin secretion in non-aggregated ␤-cells, was only marginally increased by glucose (Fig. 2C; Halban et al., 1982; Pipeleers et al., 1985; Jaques et al., 2008). This may be explained by the high basal secretion rates of purified ␤-cells. Exocytosis from ␣-cells was stimulated by glucose in the absence of neighbouring ␤-cells (Fig. 2F) consistent with earlier studies (Franklin et al., 2005; Le Marchand and Piston, 2010). Taken together, glutamate release is several-fold higher than the corresponding fraction of hormone secretion and glutamate content rather than extra-cellular glutamate is increased during glucose stimulation. Similar results were obtained using the alternative nutrient secretagogues leucine, ␣-ketoisocaproate (␣KIC) or monomethylsuccinate (Fig. 3). All three substrates stimulated hormone secretion from ␣- and ␤-cells (Fig. 3B and D). On the other hand, glutamate release was unchanged in response to the nutrients leucine and monomethyl-succinate and was even reduced when ␣KIC was used as the secretagogue (Fig. 3A and C). Glutamate is used as an amino group donor during the transamination of ␣KIC to leucine (Hernandez-Fisac et al., 2006). This transamination reaction likely reduces intra-cellular and as a consequence released glutamate (Pizarro-Delgado et al., 2009). The fact that ␣KIC nevertheless acts as a secretagogue supports the notion that glutamate-derived metabolites in the cytosol participate in the amplification of nutrient stimulated insulin secretion.

20

10

0

0

2.5

16.7

2.5

16.7

* 1

0.5

0

2.5

16.7

Fig. 1. Glucose increases glutamate content but not glutamate release from rat islets. Glutamate content of rat islets (A), glutamate released into the supernatant (B) and insulin secretion (C) were measured following a static incubation. Each measurement was performed on 20 islets maintained for 30 min at 37 ◦ C at either 2.5 or 16.7 mM glucose as indicated in the figure. (A) Glutamate content was raised by glucose (n = 4). Glutamate content was determined after lysis as described in Section 2.3. (B) Released glutamate was not significantly increased by glucose. (C) Insulin secretion is presented as the percentage of secreted hormone per total (content plus secreted). The islet insulin content was 434 ± 85 ng insulin/20 islets. Statistical significance of glucose-induced changes was assessed by Students t-test for unpaired data: *p < 0.05; **p < 0.01.

N. Feldmann et al. / Molecular and Cellular Endocrinology 338 (2011) 46–57

B glutamate release

60 40 20 0

D

glutamate release

(pmol/10000 cells)

glutamate content

40

20 10 0

16.7

2.5

40

12

8

4

16.7

2.5

F

*

120 80 40 0

**

0 2.5

E 160

** 30

80

0

16.7

(pmol/10000cells 30 minutes)

2.5

120

glucagon secretion (% of total)

(pmol/10000 cells)

glutamate content

80

C insulin secretion (% of total)

*

(pmol/10000 cells 30 minutes)

A

49

16

**

12 8 4 0

16.7

2.5

16.7

2.5

16.7

Fig. 2. A large fraction of glutamate is secreted from purified ␣- and ␤-cells. Purified ␤-cells (A–C) and ␣-cells (D–F) were incubated for 30 min at 37 ◦ C in the presence of basal (2.5 mM) or high (16.7 mM) glucose, as indicated. Glutamate content (A, D) and glutamate release (B, E) were measured as described in the legend to Fig. 1. (C) Insulin and (F) glucagon secretion are expressed as the percentage of total hormone (total plus secreted; insulin content 61 ± 10 ng/10,000 ␤-cells; glucagon content 7.0 ± 0.5 ng/10,000 ␣-cells). Shown is the average from 11 independent experiments. p-Values were calculated with Student’s t-test for paired data. Parameters significantly elevated by glucose are marked in the figure *p < 0.05; **p < 0.01.

n=10 n.s. n=8 p<0.01

100

n=6 n.s.

n=4 n.s.

50

0 control

leucine

mmsucc tolbutamide

glutamate release (% of control)

C

insulin secretion (% of control)

B 150

n=5 n=8 n.s. p<0.05 n=10 p<0.05 n=6 p<0.01

200

n=6 p<0.01 n=5 n.s.

n=6 n=4 n.s. p<0.05

100

0 control

leucine

mmsucc tolbutamide

D 150

n=5 n.s.

n=5 n.s.

n=6 n.s.

n=6 p<0.05

100

50

0 control

leucine

mmsucc tolbutamide

glucagon secretion (% of control)

glutamate release (% of control)

A

200

n=5 p<0.05

n=6 p<0.05

n=5 p<0.05

n=6 p<0.05

100

0 control

leucine

mmsucc tolbutamide

Fig. 3. Secretagogues differentially affect glutamate release and hormone secretion. Glutamate release (A, C) and hormone secretion (B, D) were measured after stimulation of purified ␤-cells (A, B) or ␣-cells (C, D) for 30 min at 37 ◦ C using the secretagogues leucine (10 mM), ␣-ketoisocaproate (␣-KIC; 10 mM), monomethyl-succinate (mmSucc; 5 mM) or tolbutamide (100 ␮M). The effect of the individual stimuli on glutamate release and hormone secretion was compared pair-wise to cells maintained at 2.5 mM glucose (control). Glutamate release and hormone secretion are expressed as percentage of the control incubation. The number of independent experiments performed (n) and the significance (p) as calculated by Student’s t-test for paired data compared to control are given in the figure.

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N. Feldmann et al. / Molecular and Cellular Endocrinology 338 (2011) 46–57

A

B 100

n=11 content (pmol/10000 cells)

content (pmol/10000 cells)

n=5 40

30

20

10

0

80

60

40

20

0

2 0 glutamine (mM)

C

islet glutamate

D

2 0 glutamine (mM)

islet glutamine

60

content (pmol/20 islets)

n=5 n=11

E

islet alanine

120

*

120

40

80

20

40

80

40

0

0 2 glutamine (mM)

0

0 2 glutamine (mM)

0

0 2 glutamine (mM)

Fig. 4. Glutamine in the culture medium does not raise glutamate in islet cells. Five hours before the experiment cells were grown in medium containing either 0 or 2 mM glutamine. Purified ␣ (A) and ␤-cells (B) were then washed with KRBH and incubated for 30 min in the presence of 16.7 mM glucose. The glutamate content was measured using the enzyme cycling assay. The numbers of independent experiments are given in the figure. (C–E) Rat islets were switched to fresh medium containing either 0 or 2 mM glutamine 5 h before the experiment. Before extraction of metabolites, the islets were washed with KRBH and stimulated with 16.7 mM glucose for 30 min. Glutamate (C), glutamine (D) and alanine (E) content was determined by HPLC. Shown is the average from 2 independent experiments performed in triplicate. Glutamine was significantly increased as calculated by Student’s t-test for unpaired data *p < 0.05.

glutamine but this difference did not reach significance (Fig. 4C). Glutamine in the culture medium did also not affect intracellular alanine levels (Fig. 4E) but caused a marked increase of glutamine (Fig. 4D). Under our conditions, glutaminolysis is not required to maintain the intracellular glutamate concentration in ␣- or ␤-cells. 3.4. Malate–aspartate shuttle activity influences intracellular glutamate levels The mitochondrial NADH shuttles (glycerolphosphate and malate–aspartate shuttle) assure reoxidation of NADH formed during glycolysis and provide additional reducing equivalents for mitochondrial respiration. In ␤-cells, the malate–aspartate-shuttle is of particular importance as it exerts a control function in nutrient-sensing (reviewed in Newsholme et al., 2005; Maechler et al., 2006). Activation of the malate–aspartate shuttle following over-expression of the glutamate/aspartate exchanger aralar increases nutrient-stimulated insulin secretion and glutamate formation (Rubi et al., 2004; Bender et al., 2009). Phenylsuccinate an inhibitor of the malate/␣-ketoglutarate carrier a component of the malate–aspartate shuttle system (McKenna et al., 2006) was used here to test whether malate–aspartate shuttle activity is required to increase glutamate in purified ␣- and ␤-cells.

Indeed, phenylsuccinate reduced glutamate by about 30% in both cell types (Fig. 5A). Inhibition of the malate/␣-ketoglutarate carrier also lowered insulin but not glucagon secretion (Fig. 5B). This confirms that optimal malate–aspartate shuttle activity is necessary for glucose-induced glutamate formation and insulin secretion in primary ␤-cells. 3.5. Stimulation of granule exocytosis does not enhance glutamate release from purified ˛- and ˇ-cells The striking quantitative differences in glutamate versus hormone secretion suggest that the mechanism of glutamate release may differ from hormone exocytosis. To test this hypothesis, secretion was directly stimulated using the sulfonylurea tolbutamide, which closes ATP-sensitive potassium channels (KATP -channels), subsequently causing depolarization of the plasma membrane and Ca2+ -influx triggering secretory granule exocytosis. Tolbutamide significantly increased insulin (Fig. 3B) and glucagon (Fig. 3D) secretion from purified ␤- and ␣-cells (Franklin et al., 2005). Glutamate release however was unaffected by tolbutamide in either preparation (Fig. 3A and C). The glutamate release mechanism was further studied using the catecholamine epinephrine. In ␤-cells, epinephrine activates ␣2-

total glutamate (% of control)

N. Feldmann et al. / Molecular and Cellular Endocrinology 338 (2011) 46–57

100

*

80

*

60 40 20 0 16.7

16.7

16.7

phenylsuccinate

glucagon hormone secretion (% of control)

16.7

phenylsuccinate

51

The compound l-trans-pyrrolidine-2,4-dicarboxylate (transPDC) is a competitive inhibitor of all five EAAT subtypes (Bridges and Esslinger, 2005). Importantly, the compound exhibits little binding and activation of ionotropic excitatory amino acid receptors (Bridges et al., 1991; Garlin et al., 1995). Trans-PDC was shown to inhibit EAATs in rat islets with a half-maximal effect at 134 ␮M with close to maximal inhibition at 500 ␮M (Weaver et al., 1998). At the latter concentration, trans-PDC decreased both basal (−41.1 ± 14.0%; n = 3; data not shown) and glucose-stimulated glutamate release in islets (Fig. 8A; −45.9 ± 5.5%). Similar attenuation was observed in purified ␣- and ␤-cells (Fig. 8C and E). Interestingly, GSIS was potentiated upon addition of trans-PDC in islets and purified ␤-cells (Fig. 8B and D). Basal insulin secretion did not change in the presence of the inhibitor (data not shown). Under the same condition, glucagon exocytosis from purified ␣-cells was not increased (Fig. 8F). These results reveal mechanistical differences of metabolism-secretion coupling in the two cell types.

insulin 3.7. Trans-PDC raises intra-cellular glutamate and ˛-ketoglutarate levels

100

**

80 60 40 20 0 16.7

16.7

phenylsuccinate

16.7

16.7

phenylsuccinate

Fig. 5. Inhibition of the malate–aspartate shuttle lowers insulin but not glucagon secretion from purified islet cells. Purified ␣- and ␤-cells were stimulated with 16.7 mM glucose for 30 min at 37 ◦ C in the presence or absence of 10 mM phenylsuccinate. Total glutamate (A) and secreted hormones (B) were analysed. The data are expressed as % of the control incubation without phenylsuccinate. n = 8 for ␣-cells; n = 6 for ␤-cells. *p < 0.05; **p < 0.01 for paired data.

adrenoceptors (Schuit and Pipeleers, 1986) inhibiting GSIS (Fig. 6B). In ␣-cells epinephrine activates ␣1- and ␤-adrenergic receptors (Schuit and Pipeleers, 1986; Vieira et al., 2004; De Marinis et al., 2010) resulting in stimulation of glucagon secretion (Fig. 6D). Epinephrine reduced glutamate release from ␣- and ␤-cells, despite opposite effects on the secretion of the two hormones (Fig. 6A and C). The experiments with tolbutamide and epinephrine show that amino acid release does not correlate with hormone exocytosis. 3.6. Glutamate release is mediated via excitatory amino acid transporters (EAATs) As the high rates of glutamate release cannot be explained by co-secretion with the hormones, we were looking for an alternative mechanism. In astrocytes, glutamate is taken up by plasma membrane EAATs (Anderson et al., 2001). Depending on the conditions and relative intra- and extra-cellular glutamate concentrations these transporters will mediate either import or export of glutamate (Anderson et al., 2001). To test whether this mechanism could be operative in islet cells, we studied the expression of EAATs in primary ␣- and ␤-cells. Transcript levels of the commonly expressed isoforms EAAT1, EAAT2, and EAAT3 were assessed and compared to brain tissue, which expresses EAATs at very high levels. Both islet cell types express EAAT1, EAAT2, and EAAT3 albeit at much lower levels than in the brain (Fig. 7). EAAT1 and EAAT3 mRNAs were 2.4 respectively 9 times more abundant in ␣- than in ␤-cells (Fig. 7).

Potentiation of GSIS by trans-PDC in islets and purified ␤-cells suggests that maintenance of the intra-cellular glutamate pool or a metabolite linked to glutamate levels is crucial for metabolismsecretion coupling. For instance, ␣-ketoglutarate can be formed directly from glutamate by transamination in a reaction catalysed by aspartate aminotransferase or by oxidation catalysed by glutamate dehydrogenase. We therefore measured the effect of trans-PDC on intra-cellular glutamate as well as ␣-ketoglutarate. More starting material (200 rat islets per condition) was required to determine ␣-ketoglutarate in the lysates. Nevertheless, GSIS was observed and was enhanced by trans-PDC (Fig. 9A). Glucose stimulation caused a significant increase in intra-cellular glutamate, which was less pronounced than under standard conditions. Trans-PDC caused a marked elevation of the islet glutamate content (Fig. 9B). ␣-Ketoglutarate levels were not increased by glucose alone but administration of trans-PDC doubled the level of the TCA cycle intermediate (Fig. 9C). Thus, the potentiation of GSIS is associated with an increase in intra-cellular glutamate and ␣ketoglutarate (Fig. 9). 3.8. Glucose lowers the glutamate-related amino acid GABA Glutamate is the precursor of the amino acid GABA. Synthesis of GABA from glutamate is catalysed by glutamate decarboxylase (Wang et al., 2006; Li et al., 2008). GABA can be further metabolized to succinic acid semialdehyde, which is oxidized to the TCA cycle intermediate succinate (GABA shunt) (Fernandez-Pascual et al., 2004; Wang et al., 2006; Pizarro-Delgado et al., 2010). The deamination of GABA to succinic acid semialdehyde is linked to the regeneration of glutamate from ␣-ketoglutarate catalysed by GABA transaminase (Fernandez-Pascual et al., 2004; Wang et al., 2006). We measured GABA release and content in rat pancreatic islets as well as in isolated rat ␤-cells. In contrast to the effects on glutamate, glucose decreased GABA content in purified ␤-cells and islets (Table 3). Our results strengthen the hypothesis that GABA participates as a substrate in metabolism-secretion coupling (Li et al., 2008). 4. Discussion Glutamate has been proposed to affect islet function in the intraand extra-cellular compartments (Moriyama and Hayashi, 2003; Maechler et al., 2006). We find that a large fraction of the total glutamate content is released from intact islets and purified ␣- and

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B 160

insulin secretion (% of total)

(pmol/10000cells 30 minutes)

glutamate release

A

120

80

40

16

12

8

*

4

0

0 16.7

16.7

16.7

16.7

epinephrine

C

epinephrine

D 160

**

120

80

40

0

2.5

2.5

glucagon secretion (% of total)

(pmol/10000cells 30 minutes)

glutamate release

200

*

2

1.5

1

0.5

0

2.5

2.5

epinephrine

epinephrine

Fig. 6. Differential regulation of glutamate release and hormone secretion by epinephrine. Glutamate release (A, C) and hormone secretion (B, D) were studied as described for Fig. 2. The purified ␣- and ␤-cells were incubated in 2.5 or 16.7 mM glucose and supplemented with epinephrine (1 ␮M) as indicated. Average data from ␣-cells (n = 9) and ␤-cells (n = 5) are presented (p-values were calculated using Student’s t-test for paired data *p < 0.05; **p < 0.01).

B 2

1.5

1

0.5

0

EAAT2 expression % of brain

EAAT1 expression % of brain

A

0.5

0.4

0.3

0.2

0

which may also contribute to basal hypersecretion (Figs. 1 and 2 and values of insulin content in figure legend). The reason for the differences between islets and isolated cells remains unclear but may be attributed to the importance of glutamate handling and cell-cell contacts in the control of nutrient metabolism and hormone exocytosis (Halban et al., 1982; Pipeleers et al., 1985; Jaques et al., 2008).

C EAAT3 expression % of brain

␤-cells. In all three preparations fractional glutamate release was at least many fold higher than the respective hormone secretion. The weak stimulation of insulin secretion by glucose from isolated ␤-cells has been attributed to enhanced basal cytosolic calcium concentrations (Jaques et al., 2008). Moreover comparison of intact islets and purified ␤-cells suggests that glutamate content is elevated in the isolated cells when related to insulin content,

6

4

2

0

Fig. 7. Purified ␣- and ␤-cells express excitatory amino acid transporters. (A) EAAT1, (B) EAAT2, and (C) EAAT3 mRNA levels in ␣- and ␤-cells compared to brain. EAAT1, EAAT2, and EAAT3 expression was quantified using QT-PCR on RNA isolated from purified ␣- and ␤-cells as well as from brain. Data was normalized to the corresponding cyclophilin mRNA levels determined by RT-PCR. EAAT expression in the purified islet cells is expressed as percent expression compared to brain tissue. Data represent the mean ± S.E.M. of three independent experiments performed in duplicate.

N. Feldmann et al. / Molecular and Cellular Endocrinology 338 (2011) 46–57

B

50

islets

40

*

30

20

10

insulin secretion (% of total)

(pmol/20 islets 30 minutes)

glutamate release

A

0

16.7

2.5

1.5

1

0.5

16.7

tPDC

D

160

16

120

*

80

40

0

16.7

12

8

4

0

16.7

*

16.7

tPDC

40

** 20

0

16.7

16.7

tPDC

glucagon secretion (% of total)

(pmol/10000cells 30 minutes)

60

16.7

tPDC

F

E

glutamate release

16.7

tPDC

insulin secretion (% of total)

(pmol/10000cells 30 minutes)

glutamate release

C

*

2

0

16.7

islets

53

10

8

6

4

2

0

16.7

16.7

tPDC

Fig. 8. The EAAT inhibitor trans-PDC diminishes glutamate release and augments insulin secretion. Glutamate release and hormone secretion was studied as described in the legends to Figs. 1 and 2. Islets (A, B), purified ␤-cells (C, D) and purified ␣-cells (E, F) were incubated in medium containing 16.7 mM glucose in the presence or absence of the EAAT inhibitor trans-PDC (500 ␮M). Trans-PDC significantly reduced glutamate release from intact islets (n = 5), ␤-cells (n = 5) and ␣-cells (n = 5) and augmented insulin secretion (n = 5) but not glucagon secretion. p-Values were calculated by Student’s t-test for paired data and indicated by asterisks (*p < 0.05; **p < 0.02).

Tolbutamide augmented hormone secretion from both purified ␣- and ␤-cells while glutamate release was unaffected (Fig. 3). Taken together, the large fractional glutamate release and the results with tolbutamide clearly show that glutamate cosecretion with the hormones does not significantly contribute to the observed glutamate release in either of the two main islet cell types.

The most marked dissociation between hormone exocytosis and glutamate release was observed in purified ␣-cells stimulated with epinephrine. Epinephrine induces glucagon secretion through the activation of ␤-adrenoceptors and the generation of cAMP (Schuit and Pipeleers, 1986; Gromada et al., 2007). Epinephrine stimulated glucagon secretion but lowered glutamate release (Fig. 6C and D). In contrast, the catecholamine inhibits insulin secretion by binding to

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N. Feldmann et al. / Molecular and Cellular Endocrinology 338 (2011) 46–57

Table 3 Effects of glucose on GABA in islets and purified ␤-cells. Glucose (mM)

2.5 16.7

␤-cells GABA (pmol/10,000 cells)

Islet GABA (pmol/20 islets) Release

Content

Total

Release

Content

Total

27.3 ± 3.8 26.1 ± 3.1

169.7 ± 25.6 107.2 ± 18.4 b

197.0 ± 27.8 133.3 ± 21b

9.3 ± 2.2 5.8 ± 1.5a

39.2 ± 10.0 20.3 ± 5.1c

48.5 ± 8.3 26.1 ± 3.9c

GABA release, content and total amino acid amounts were measured after a static incubation of whole islets or purified ␤-cells for 30 min at 37 ◦ C in KRBH buffer containing 2.5 mM or 16.7 mM glucose. Measurements were performed by HPLC (n = 3). a p < 0.05. b p < 0.01. c p < 0.001 (two-tailed Student’s t-test for paired data).

␣2-adrenoceptors, affecting the plasma membrane potential, lowering intra-cellular cAMP and most importantly directly inhibiting exocytosis (Schuit and Pipeleers, 1986; Ullrich and Wollheim, 1988; Sharp, 1996) (Fig. 6B). This inhibition was accompanied by a reduction of net glutamate release. Epinephrine therefore reduces glutamate release in both cell types while having opposite effects on cAMP levels. Hence, we conclude that glutamate release unlike hormone exocytosis is not directly regulated by cAMP signalling. Activation of ␣1-adrenoceptors by norepinephrine has been reported to stimulate glutamate uptake in astrocytes (Fahrig, 1993). The presence of ␣1-adrenoceptors on ␣-cells would argue that acceleration of glutamate re-uptake may explain the observed reduced extra-cellular glutamate concentration (Vieira et al., 2004). It is not known, whether ␤-cells also express this subclass of adrenoceptors. We demonstrate that the bulk of glutamate is not co-released with the islet hormones, which prompted us to search for an alternative mechanism. In the brain, EAATs have been shown to mediate both glutamate uptake and release (uptake reversal) (Anderson et al., 2001). We demonstrate that ␣- as well as ␤-cells express all three common isoforms EAAT1, 2 and 3. Depending on the isoform, expression levels were only 0.4–5% those of brain tissue. The general EAAT inhibitor trans-PDC attenuated glutamate release in whole islets and the two purified cell preparations supporting a functional role of EAAT in mediating glutamate export. Weaver and colleagues reported glutamate uptake mainly by ␣-cells in rat islets (Weaver et al., 1998). Indeed, we find that ␣-cells expressed higher levels of EAATs than ␤-cells. The islet architecture and reuptake of glutamate by ␣-cells may explain why intact islets displayed lower fractional glutamate release than the isolated cell preparations. Intra- and extra-cellular glutamate pools may affect islet hormone secretion. For example, activation of ionotropic glutamate receptors stimulates insulin secretion in the perfused pancreas (Bertrand et al., 1992) and statically incubated mouse islets (Inagaki et al., 1995). On the other hand glutamate receptor ligands did not activate insulin exocytosis in single ␤-cells (Cho et al., 2010) or perfused mouse islets (Cabrera et al., 2008). Paracrine actions of glucagon or truncated GIP (Fujita et al., 2010) may explain the stimulatory effects in the perfused pancreas or when the hormones are allowed to accumulate during static incubations. Indeed, glucagon secretion is invariably stimulated by glutamate and ionotropic receptor ligands (Bertrand et al., 1993; Cabrera et al., 2008; Cho et al., 2010). Interestingly, reduction of glutamate efflux by trans-PDC caused a potentiation of GSIS, both in islets and purified ␤-cells (Fig. 8B and D). A similar stimulatory effect of trans-PDC on insulin secretion has been demonstrated previously in islets stimulated with 8.3 mM glucose (Weaver et al., 1998). These results taken together with the weak or absent effects of glutamate receptor agonists suggest intrarather than extra-cellular action of glutamate in the potentiation of insulin secretion. In contrast, glucagon secretion from ␣-cells was unaffected despite the inhibition of glutamate release by trans-PDC (Fig. 6F).

Regulation of glutamate export may therefore be of functional consequences mainly for the ␤-cell in which maintenance of the intra-cellular concentration of glutamate or metabolites linked to glutamate is crucial for normal GSIS (Maechler and Wollheim, 1999; Rubi et al., 2001; Hoy et al., 2002; Maechler et al., 2006). The intra-cellular glutamate pool is likely of lesser importance in ␣-cells as this cell type depends less on anaplerosis and oxidative metabolism (Schuit et al., 1997; Ishihara et al., 2003; Gromada et al., 2007). The importance of glutamate in ␤-cell metabolism-secretion coupling has been contested mainly on the argument that intracellular glutamate concentrations are not strictly correlated with insulin secretion (Bertrand et al., 2002). However, our results support the hypothesis that intra-cellular glutamate or a derived metabolite is needed to augment GSIS. Indeed we also found some discrepancy between insulin secretion and glutamate formation when applying ␣-KIC. This secretagogue lowers glutamate levels through a transamination reaction forming leucine and ␣ketoglutarate. This suggests that not glutamate but one of its metabolites is implicated in the amplification of secretion. Multiple metabolic pathways influence glucose-dependent glutamate synthesis. Using the inhibitor phenylsuccinate, we provide evidence for the importance of the malate–aspartate shuttle system in glucose stimulated glutamate formation in primary ␣- and ␤cells. Phenylsuccinate lowered glutamate in both cell types, while reducing glucose-stimulated insulin but not glucagon secretion. These results reveal differences in the molecular mechanisms leading to insulin and glucagon secretion. Our findings with purified ␤-cells complement earlier studies demonstrating the importance of the malate–aspartate shuttle in glucose-stimulated glutamate formation and insulin secretion (Rubi et al., 2004; Bender et al., 2009). Glutamine by itself does not act as a secretagogue but potentiates actions of other stimuli (Malaisse et al., 1982; Bertrand et al., 2002; Li et al., 2004). Addition of glutamine during static incubations augments intracellular glutamate (Bertrand et al., 2002). Manipulation of the glutamine concentration in the culture medium prior to the experiment as shown in this study had little or no effect on glutamate levels in rat islets or purified islet cells. We conclude that glutaminolysis is not a main contributor to the nutrient-stimulated glutamate synthesis described in this study. After glucose stimulation of rat islets glutamine levels were significantly increased. These findings are consistent with the possibility that glutamate or glutamine participates in metabolismsecretion coupling (Maechler and Wollheim, 1999; Li et al., 2004, 2008; Maechler et al., 2006). Current knowledge does not allow conclusions on the relative contribution of these two amino acids to the amplification of GSIS. ␣-Ketoglutarate can be formed from glutamate either via transamination or the mitochondrial glutamate dehydrogenase reaction (Newsholme et al., 2005; Li et al., 2006; Wang et al., 2006). Unlike glutamate, ␣-ketoglutarate was not elevated by glucose in agreement with earlier work (MacDonald, 2002, 2003; Liu et al.,

N. Feldmann et al. / Molecular and Cellular Endocrinology 338 (2011) 46–57

A

3

** 2

**

1

0

2.5

16.7

16.7

tPDC glutamate content (nmol/mg protein)

B

*

40

30

*

20

10

0

16.7

16.7

tPDC 0.5

**

The present study shows that glucose stimulates glutamate synthesis in islet ␣- and ␤-cells. Glutamate is not co-secreted with insulin or glucagon but rather released at high rates in part through EAATs. Based on our results with a general EAAT inhibitor, we conclude that intra-cellular glutamate or glutamate-derived metabolites contribute to fuel-induced hormone secretion in ␤but not ␣-cells. Regulated glutamate release or re-uptake, would be a pharmacological target for the modulation of GSIS. Funding sources

0.4

The research was supported by the Swiss National Science Foundation (grant nos. 32-66907.01 and 310000-116750/1). EuroDia (LSHM-CT-2006-518153) a European-Community funded project under framework program 6 as well as by the Bo and Kerstin Hjelt Foundation for Type 2 diabetes research. The work was also partially supported by a grant from Instituto de Salud Carlos III (PI 061744), Madrid (Spain).

0.3

0.2

0.1

0

trans-PDC on GSIS may therefore in part be mediated by raising ␣-ketoglutarate. When compared to glutamate, ␣-ketoglutarate levels in the rat islets are low and similar in concentration to the values published by others (Sener et al., 1981; Malaisse et al., 1982; Liu et al., 2003). Glucose metabolism should increase ␣-ketoglutarate generation in the TCA cycle, catalysed by either isocitrate or glutamate dehydrogenase (Li et al., 2006; Jensen et al., 2008). ␣Ketoglutarate formed from glucose may immediately serve as a substrate for other metabolic pathways to generate further downstream metabolites. This would explain why glucose alone does not raise ␣-ketoglutarate in islets. A possible pathway diminishing ␣-ketoglutarate is the GABA shunt (Fernandez-Pascual et al., 2004; Wang et al., 2006; Li et al., 2008), which contributes to the stimulation of insulin secretion (Li et al., 2008; Pizarro-Delgado et al., 2010). Our results show that GABA content is reduced in both intact islets and isolated ␤-cells following glucose stimulation. These results confirm earlier observations with rat islets (Li et al., 2006; Wang et al., 2006) and demonstrate that GABA shunt activity during glucose stimulation is one pathway that prevents glucosemediated increases of ␣-ketoglutarate in favour of the intra-cellular glutamate pool. 5. Conclusion

2.5

C

55

2.5

16.7

16.7

tPDC Fig. 9. Inhibition of EAAT raises intra-cellular glutamate and ␣-ketoglutarate in rat pancreatic islets. Static incubation of 200 rat islets were carried out in basal (2.5 mM) or stimulatory glucose concentrations (16.7 mM) in the presence or absence of 500 ␮M trans-PDC as shown in the figure. After 30 min at 37 ◦ C insulin secretion (A) was measured and islet lysates were prepared to measure the glutamate (B) and ␣-ketoglutarate (C) content. Values are the mean ± S.E.M. of five independent experiments. Trans-PDC increased insulin secretion, glutamate and ␣-ketoglutarate compared to the incubation with 16.7 mM glucose without the compound. Significance was calculated by Student’s t-test for paired data: *p < 0.05; **p < 0.02.

2003). Inhibition of glutamate export with trans-PDC elevated the concentrations of both metabolites in glucose stimulated islets. ␣Ketoglutarate has been proposed to stimulate insulin exocytosis (Rabaglia et al., 2005; Jensen et al., 2008; Willenborg et al., 2009), possibly by acting at a step distal to the function of plasma membrane ion channels (Willenborg et al., 2009). The importance of the cytosolic ␣-ketoglutarate pool is further suggested by the inhibition of GSIS after suppression of the mitochondrial ␣-ketoglutarate transporter (Odegaard et al., 2010). The stimulatory effects of

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