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Altered NAD(P)H production in neonatal rat islets resistant to H2O2
Life Sciences 83 (2008) 709–716
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Life Sciences j o u r n a l h o m e p a g e : w w w. e l s ev i e r. c o m / l o c a t e / l i f e s c i e
Altered NAD(P)H production in neonatal rat islets resistant to H2O2 Luiz F. Stoppiglia a, Luiz F. Rezende b, Ana P.G. Cappelli b, Fabiano Ferreira c, Antonio C. Boschero b,⁎ a b c
Departamento de Química, Instituto de Ciências Exatas e da Terra, Universidade Federal de Cuiabá, MT, Brazil Departamento de Fisiologia e Biofísica, Instituto de Biologia, Universidade Estadual de Campinas (UNICAMP), Brazil Departamento de Fisiologia e Farmacologia, Centro de Ciências Biológicas, Universidade Federal de Pernambuco, PE, Brazil
a r t i c l e
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Article history: Received 4 January 2008 Accepted 4 September 2008 Keywords: Glucose oxidation H2O2 NADH shuttles Neonatal pancreatic islets Pentose phosphate pathway
a b s t r a c t Aims: We determined the involvement of NAD(P)H generation ability on the resistance of pancreatic islets Bcells to oxidative stress caused by culture exposition to H2O2. Main methods: We cultured isolated neonatal Wistar rat islets for four days in medium containing 5.6 or 20 mM glucose, with or without H2O2 (200 μM), and analyzed several parameters associated with islet survival in different media. High glucose was used since it protects neonatal islets against the loss of GSIS. Key findings: While none of the culture conditions increased the rate of NAD(P)H content at 16.7 mM glucose, the islets resistant to H2O2 and those exposed to 20 mM glucose showed a greater use of the pentose phosphate pathway and increased ATP synthesis from glucose. Significance: Oxidative stress contributes to the loss of glucose-induced insulin secretion (GSIS) during the onset of diabetes mellitus. Although immature rat islets have reduced GSIS compared to mature islets, they adapt better to oxidative stress and are a good model for understanding the causes involved in the destruction or survival of islet cells. These data support the idea that GSIS and resistance against oxidative stress in immature islets rely on NADH shuttle activities, with little contribution of reduced equivalents from the tricarboxylic acid cycle (TCAC). © 2008 Elsevier Inc. All rights reserved.
Introduction Malfunction of pancreatic islets impairs glucose homeostasis and can lead to diabetes mellitus. Insulin secretion occurs after an increase in the ATP/ADP ratio in B-cells via the Ca2+ dependent and independent pathways. Ca2+-dependent insulin secretion is referred to as the triggering pathway, whereas Ca2+-independent secretion accounts for the amplifying pathway (Asanuma et al., 1997). In humans and experimental animals, a combination of chronic hyperglycemia, high fatty acid concentration, cytokines and reactive oxygen species (ROS) reduce the responsiveness of islets to glucose by affecting both pathways. Impairment of GSIS is mainly assigned to depletion of insulin-promoter ligands, B-cell death, inhibition of glucolytic key enzymes, mitochondria damage, shift of glucose oxidation to other pathways, and defects in cellular calcium handling. On the other hand, chronic exposure to high levels of glucose increases ATP/ADP ratio and NAD(P)H concentrations, UCP2 activity and basal cytoplasmic Ca2+. Chronic hyperglycemia is believed to be a main cause of the loss of GSIS (Mears, 2004; Khaldi et al., 2004; Dubois et al., 2007).
⁎ Corresponding author. Departamento de Fisiologia e Biofísica, Instituto de Biologia, Universidade Estadual de Campinas (UNICAMP), CP 6109, Campinas, 13083-970, SP, Brazil. Fax: +55 19 3521 6185. E-mail address: [email protected] (A.C. Boschero). 0024-3205/$ – see front matter © 2008 Elsevier Inc. All rights reserved. doi:10.1016/j.lfs.2008.09.012
Mitochondrial oxidation of glycolysis-derived NADH is an essential trigger for insulin secretion (Eto et al., 1999). NADH is transported actively against its concentration gradient from cytosol the mitochondrial matrix by NADH shuttles, as a required complement to pyruvatederived NADH/FADH2 (Dukes et al., 1994). This is provided by the malate/aspartate and glycerol-phosphate shuttles. The first consumes the proton gradient through an electrogenic glutamate/aspartate antiporter at the inner mitochondrial membrane; the second wastes the O2 motive force by use of a FADH2-linked dehydrogenase to bypass proton-pumps and insert electrons directly in the respiratory chain. The oxidation of cytoplasmic NADH by mitochondria gives more energy equivalents than what is lost by the shuttles and equilibrates the necessary NADH/NAD ratio, accounting for 50% of the glycolytic flux (MacDonald et al., 2005a,b; Westermark et al., 2007). In rat islets, a counter-current mechanism uses the pyruvate and NAD (P)H gradients through the mitochondrial inner membrane to export NADH equivalents from matrix to cytoplasm, provided mainly by the pyruvate/malate shuttle. Malate is exported in large amounts by the mitochondria of mouse and rat islets and signs for TCAC cycle activity. Some malate is converted to pyruvate by a NADP-linked malic enzyme, which maintains the high cytosolic NADPH/NADP ratio in B-cells. The citrate/isocitrate shuttle may be not critical for insulin secretion, since insulin secretagogues do not cause exportation of isocitrate from rat mitochondria (MacDonald et al., 2005b). Neonatal rat islets, cultured in high concentrations of glucose, but not L-leucine or other secretagogues, present augmented glucose-
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stimulated insulin secretion (GSIS) and resistance to the deleterious effects of H2O2 (Stoppiglia et al., 2002; Anello et al., 2001). Despite the similarity between insulin secretion induced by glucose and L-leucine, the latter lowers cytoplasmic glutamate and increases aspartate levels through allosteric activation of mitochondrial glutamate dehydrogenase (Gao et al., 2003). Rat islets apparently have a sufficiently high glutamate content to maintain insulin secretion for 2–3 h through glutamate dehydrogenase and mal/asp shuttle activities. However, the chronic use of non-pyruvate-producing metabolites impairs the shuttle activity by depleting mitochondrial aspartate (MacDonald et al., 2005a,b). The activity of these NADH shuttles is amplified several times when islets progress from a low-responsiveness fetal state to an adult condition (Tan et al., 2002). We have previously suggested that neonatal rat islets have a different mechanism of defense against ROS by offering high concentrations of NAD(P)H to a scavenger enzyme-poor system (Stoppiglia et al., 2004). This mechanism may indicate that β-cells can actively modify the production of reduced equivalents from their high glucolytic rate and direct it to survival purposes, possibly altering insulin secretion. In this work, we investigated whether the defense status against H2O2 is sufficient to increase cytoplasmic NAD(P)H production and how this affects glucose-stimulated insulin secretion. Materials and methods H2O2 concentrations All H2O2 concentrations cited have been carefully tested by H2O2 colorimetric reaction with a horseradish peroxidase/4-aminoantipyridine system and with potassium iodide as described (Stoppiglia et al., 2004). In previous experiments, we tested different concentrations of H2O2 for deleterious activity in cultured islets (Stoppiglia et al., 2002). A borderline deleterious activity with observable alteration of metabolic activity was found to happen with 200 μM of H2O2 after a 4 day incubation period. The same H2O2 concentration appeared to cause much more damage to adult than to neonatal rat islets. This concentration was used for the experiments described, else being described in each case. Chemicals Radiochemicals were from G.E. Heath Care (UK), the aminotransferase inhibitor amino-oxyacetate (carboxymethoxylamine) was from Sigma-Aldrich (St. Louis/MO, USA), the MTS/PMS preparation was from a CellTitter96 aqueous assay of Promega (Madison/WI, USA) and all RT-PCR reagents were from Invitrogen (Carlsbad/CA, USA). The other reagents were from Sigma-Aldrich, whenever specified. Islet isolation and culture Neonatal (1–2-day-old) Wistar rats were from the animal facilities at the State University of Campinas. After decapitation, the islets were isolated by collagenase (EC 3.4.24.3) digestion of pancreata in Hank's balanced salt solution (in mM: 137 NaCl, 5.5 KCl, 4.5 NaHCO3, 0.4 KH2PO4, 0.4 Na2HPO4, 0.8 MgSO4, 1.5 CaCl2, pH 7.4). The islets were extensively washed in sterile Hank's solution and cultured in RPMI 1640 medium supplemented with 2 g NaHCO3/l, 1% (v/v) penicillin/streptomycin and 10 mM D-glucose, pH 7.4. Approximately 1000 islets/dish were maintained at 37 °C in a humidified atmosphere with 3% CO2 for two days before further additions. The culture medium contained glucose (5.6 mM or 20 mM) with or without 200 μM H2O2, for an additional four days. The medium was renewed every 24 h. The islet experimental groups were assigned according to the culture conditions: G5.6 (islets cultured in 5.6 mM glucose), P5.6 (5.6 mM glucose and 200 μM H2O2), G20 (20 mM glucose) and P20 (20 mM glucose and 200 μM H2O2). A similar procedure is described (Stoppiglia et al., 2002).
Insulin secretion Batches of 10 islets were each incubated for 30 min at 37 °C in Krebs–Hepes buffered saline (KHBS, in mM: 115 NaCl, 10 NaHCO3, 5 KCl, 1 MgCl2, 2.5 CaCl2, 15 Hepes) containing 0.5 g BSA/l and 5.6 mM glucose, pH 7.4, and equilibrated with 95% O2 and 5% CO2. The medium was discarded and the islets were incubated for a further 1 h in 1 ml of KHBS containing 2.8 or 16.7 mM glucose with the additions described in the corresponding figure legends. The supernatant was collected and the insulin concentration was measured by radioimmunoassay (Delghingaro-Augusto et al., 2004). Glucose uptake and metabolism Batches of 50 islets each were incubated for 2 h at 37 °C in KHBS containing 2.8 or 16.7 mM glucose with trace amounts of either 14 14 14 D-[U- C]glucose or D-[1- C]glucose to measure CO2 production. HCl (1 N) was added to the batches to stop respiration and the 14CO2 formed was collected for 1 h at 4 °C in 1 N NaOH. For the glucose uptake assays, the batches were incubated under similar conditions with 2-deoxy-D-[U-14C]glucose for 2 h, after which the islets were washed twice with KHBS at 4 °C, sedimented and homogenized in the presence of Trizol (Invitrogen). The partitioning of glucose between the TCAC and the pentose phosphate pathway (PPP) was calculated by assuming that the total 14CO2 produced from labeled glucose was from the TCAC and the PPP. A detailed description of the data processing is given by Katz and Wood, 1963. In short, if we define D-[1-14C]glucose and D-[U-14C]glucose conversion to CO2 as G1 and GU, respectively, it is possible to construct a linear system of two equation were G1 is the sum of TCAC and PPP; because only one 14 C carbon is detected per glucose molecule. However, GU is obtained as the sum of PPP glucose utilization and 6 times the TCAC glucose utilization, because TCAC generates 6 14C atoms per glucose molecule oxidized. Then, the amount of glucose that passed through the TCAC was calculated as (GU − G1)/5, and that which passed through the PPP as (6 × G1 − GU)/5. RT-PCR Groups of 1000 islets were homogenized in Trizol followed by phenol–chloroform RNA extraction, according to the manufacturer's instructions. The quality of the RNA was assessed by agarose gel electrophoresis. One to five micrograms of total RNA were transferred to reaction vials containing 1 μg of dNTP mix. The reactions were incubated for 5 min at 65 °C before adding 150 ng of random primers (10 min at 25 °C) followed by 14.3 mM MgCl2, 2.8 mM DTT and 0.4 U/μl RNase-out (2 min at 42 °C), and 1.25 U/μl RNA Super Script II. The mixtures were then incubated at 42 °C for 50 min and at 70 °C for 15 min, followed by cooling to 4 °C. The resulting cDNAs were diluted in PCR buffer (60 mM Tris–HCl, 1.5 mM MgCl2, 15 mM NH4SO4, pH 10) with 50 mM MgCl2, 0.3 mM each of dATP, dCTP, dGTP and dTTP and 2.5 U/μl Taq DNA polymerase (Gibco/BRL), followed by the addition of 10 mM of forward and reverse primers. PCR amplification of cDNA was carried out with a GeneAmp PCR System 9700 (Applied Biosystems, Foster City, CA, USA) and involved 32 cycles of denaturation for 1 min at 94 °C, annealing for 1 min at referred temperatures, extension for 1.5 min at 72 °C, and a final extension for 7 min at 72 °C. The annealing temperatures and the number of cycles used for amplification are shown in the corresponding legends. Following amplification, the cDNA fragments were analyzed on 1.6% agarose gels containing a 100 bp DNA molecular weight ladder (Gibco/BRL). The PCR products were detected by staining with ethidium bromide and were documented with a Gel Doc EQ photodocumentation system (BioRad). Internal control reactions were performed for the RPS-29 subunit of the small ribosomal unit 40S, since this showed the best homogeneity
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between treated and non-treated groups. The primers were designed and tested against the Rattus norvergicus genome (Gene Bank) to ensure no amplification of other cDNAs. The oligonucleotide primers used were as follows: RPS-29 (forward) 5′AGG CAA GAT GGG TCA CCA GC3′; RPS-29 (reverse) 5′AGT CGA ATC CAT TCA GGT CG3′; rat pro-insulin 2 (forward) 5′TTG CAG TAG TTC TCC AGT T3′; rat pro-insulin 2 (reverse) 5′ATT GTT CCA ACA TGG CCC TGT3′; GLUT2 (forward) 5′CAT TGC TGG AAG CGT ATC AG3′; GLUT2 (reverse) 5′GAG ACC TTC TGC TCA GTC GAC G3′; mitochondrial pyruvate dehydrogenase subunit E1 alpha 2 (mPDH) (forward) 5′TCA AGT ACT ACA GGA TGA TG3′; mPDH (reverse) 5′GGC GTA CAT GTG CAT TGA TC3′; mitochondrial glycerol-3-phosphate dehydrogenase (mGPDH) (forward) 5′AGA AAG TCT GCA TCG TAG GCT3′; mGPDH (reverse) 5′GGA AGT TGG GTG TTT GCA TCA3′; mitochondrial malate dehydrogenase (mMDH) (forward) 5′CCT GAA GCC ATG ATT TGC ATC3′; mMDH (reverse) 5′TTC TTG ATG GAG GCT TTC AGC3′; glucose-6-phosphate dehydrogenase (G6PDH) (forward) 5′AGC TCC AAT CAA CTG TCG AAC3′; G6PDH (reverse) 5′TCC TCA GGG TTG AAG AAC ATG3′. All annealing temperatures and number of cycles were chosen to provide maximum sensitivity in the assay. A similar procedure is described (Delghingaro-Augusto et al., 2004). NAD(P)H determination NAD(P)H was measured by the coupled reduction of phenazine methosulfate (PMS) and subsequent transfer of electrons to the tetrazolium salt MTS, both of which are membrane permeable. The MTS and PMS solutions were provided as parts of the CellTitter96 aqueous assay (Promega) and were mixed according to the manufacturer's instructions. There was little change in the absorbance at 492 nm absorbance with concentrations of NAD(P)H b10 μmol/l. Because of the important interference by proteins bound to NAD(P)H, the standard curves were less accurate at low concentrations of NAD(P)H. However, spectroscopic analysis showed an increase in light absorption at 650 nm of ~ 10− 3 cm− 1 islet− 1 when different numbers of islets where disrupted in MTS/PMS solution containing 2 mg of BSA/ml. Heat-denaturated islet homogenates were used as negative controls. Static measurements of NAD(P)H were done by incubating groups of 200 islets in KHBS containing 2.8 or 16.7 mM of glucose, which reproduced the same conditions used in the insulin secretion experiments. The islets were then washed in ice-cold Hanks solution and immediately sonicated in 150 μl of this same solution. The homogenates were centrifuged (10 000 g × 2 min) to remove islet debris and aliquots of the supernatants were then added to the MTS/ PMS solution and incubated for 30 min at room temperature before recording the absorbances at 650 nm and 405 nm (background). Mixtures with no islets were used as blanks. In each experiment, the changes in the absorbance at 650 nm were normalized to the mean absorbance of G5.6 samples incubated with 16.7 mM of glucose. Dynamic measurements of NAD(P)H were done by incubating groups of 20 islets in 200 μl of KHBS containing 0 or 20 mM of glucose and 5% (v/v) of MTS/PMS. This concentration of MTS/PMS provided the best sensitivity and was chosen after testing concentrations from 1% to 20% (v/v). As judged by the transient rate of MTS/PMS reduction, concentrations of MTS/PMS N7% caused cell death through NAD(P)H depletion. Samples were incubated for 3 h in a 95% O2 + 5% CO2 atmosphere (140 μM O2 in KHBS) and then changed to a 100% N2 atmosphere (90 μM O2) for a further 3 h to cause hypoxia. Some of the samples were returned to the 95% O2 atmosphere and cell death caused by hypoxia was estimated to be b20%. Samples with no islets were used as blanks and the absorbances at 492 nm were recorded every 30 min. Standard curves of NADPH were used to calculate the decrease in the NAD(P)H content of the samples. The NAD(P)+ reduction rate (NRR) was determined from the increase in MTS/PMS absorbance in each sample.
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Partitioning of the NAD(P)+ reduction rate The partitioning of the NRR was calculated by assuming that the NRR consisted of four components: cytoplasmic NRR from endogenous sources (endo.cyto), cytoplasmic NRR from glucose oxidation (gluc. cyto), mitochondrial NRR from endogenous sources (endo.mito) and mitochondrial NRR from glucose-mediated metabolite oxidation (gluc. mito). The mitochondrial components were assumed to depend on an adequate oxygen supply, whereas the cytoplasmic components were essentially anaerobic. The NRR from endogenous sources was estimated from values observed in the absence of glucose. These endogenous fuels were presumed to be mostly fatty acids and amino acids. Statistics The results were expressed as means ± S.E.M. Point-to-point comparisons were done using Students t-test. The groups were compared by two-way ANOVA followed by the unpaired Tukey– Kramer test. A value of P b 0.05 indicated significance. However, in the RT-PCR experiments, significance was only considered when P b 0.001. In each figure, different letters were used to indicate significant differences in comparisons between the groups plotted. Identical letters above two groups indicate that no statistical difference was seen. Greek letters were used to indicate a separated set of results compared only between them. Results At the end of the culture period, the GSIS of the control group G5.6 was similar to that of fresh isolated islets (not shown). Islets cultured
Fig. 1. (A) Insulin secretion by islets cultured 4 days with 5.6 or 20 mM glucose (G5.6 and G20), in the absence or presence of 200 μM H2O2 (P5.6 and P20). Islets were preincubated for 30 min in KHBS containing 5.6 mM glucose, as described, and then incubated either with 2.8 or 16.7 mM glucose for 1 h. The bars represent means ± S.E.M. of 8 independent experiments, with different letters indicating P b 0.05. (B) Partitioning of metabolized glucose between the TCAC (open part of bars) and the PPP (filled bars). The values were calculated from the amount of 14CO2 produced by D-[U-14C]glucose and 14 D-[1- C]glucose metabolism in vials containing KHBS with 2.8 or 16.7 mM glucose, as described. All of the samples gave similar results in the presence of 2.8 mM glucose. The bars are the means ± S.E.M. of 8 experiments performed simultaneously with D-[U-14C] glucose and D-[1-14C]glucose. Different letters indicate P b 0.05 for comparisons within the TCAC and within the PPP.
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Fig. 2. (A) Glucose uptake in islets cultured in the same conditions as in Fig. 1. Islets were incubated for 120 min in KHBS with 2.8 or 16.7 mM glucose containing equal amounts of 2-deoxy-D-[U-14C]glucose, and then washed and homogenized with Trizol. The bars are the means± S.E.M. of at least 8 experiments. Different letters indicate P b 0.05. (B) Effect of incubation with high glucose/H2O2 on islet GLUT2 mRNA levels. The annealing temperatures and cycle numbers were 55 °C and 29 cycles for GLUT2 and 57 °C and 29 for RPS-29, respectively. RPS-29 was used as an internal control showing no variation among the conditions tested. The bars are the mean ± S.E.M. of 12 experiments. There were no statistical differences in mRNA content.
with high glucose or H2O2 showed increased glucose oxidation and higher GSIS compared to the controls (Fig. 1), but reduced coupling of GSIS to glucose oxidation. When islets are incubated in 2.8 or 16.7 mM
glucose, there are increases in glucose oxidation and insulin secretion. Comparing the increase of insulin secretion/increase of glucose oxidation, we have the following values (in Δng insulin/Δpmol glucose): a b b ab G5.6 1.4 ± 0.1 , P5.6 0.5 ± 0.1 , G20 0.7 ± 0.2 and P20 1.1 ± 0.2 (see Materials and methods for meaning of letters). This suggests that glucose and H2O2 may share a mechanism that diverts carbons from ATP production. Despite a significant loss of islets when cultured with H2O2, treatments with high glucose or H2O2 did not change GLUT2 mRNA content (Fig. 2B) nor decrease glucose uptake (Fig. 2A). In addition, in the post-culture islet population, there was no significant change in the insulin content (data not shown). Exposure to high glucose and H2O2 selected islets with increased oxidation of glucose through the PPP (Fig. 1B), which produces cytoplasmic NADPH. This selection was consistent with an increased expression of G6PDH mRNA (Fig. 3A). The expression of genes encoding proteins related to NAD(P)H that transit between mitochondria and the cytoplasm was also modified by glucose and H2O2 (Fig. 3B,C): mGPDH expression was increased in islets exposed to high glucose, whereas mMDH expression was higher in islets resistant to H2O2. The expression of mPDH was unaffected by either of these treatments (Fig. 3D). When challenged with glucose, islets resistant to H2O2 were less dependent on glucose oxidation via the mal/asp shuttle, as indicated by the use of the transaminase inhibitor amino-oxyacetate (Fig. 4A). Remarkably, the participation of the mal/asp shuttle in glucose oxidation was activated by a glucose challenge only in the P5.6 group, whereas this shuttle was required for GSIS in all other groups (Fig. 4B). To induce insulin release, NADH from the TCAC and cytoplasm must enter the respiratory chain and donate electrons to ATP synthesis. Electrons from NADH are donated to complex I or directly to complex II through the activity of a FADH2-linked glycerolphosphate shuttle. When complex I was inhibited with rotenone, exposure to high glucose and H2O2 increased the use of mitochondrial NADH for GSIS, although these effects were not synergic or additive
Fig. 3. Effect of exposure to high glucose and H2O2 on the mRNA levels of (A) G6PDH, (B) mG3PDH, (C) mMDH and (D) mPDH. The RT-PCR annealing temperatures and cycle numbers used were 57 °C and 31 cycles (G6PDH and mG3PDH), 57 °C and 35 cycles (mMDH) and 55 °C and 32 cycles (mPDH). RPS-29 cDNA was amplified using 57 °C and 29 cycles, and showed no variation among the conditions tested. Bars are the means ± S.E.M. of 12 experiments. Different letters indicate P b 0.001.
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(Fig. 5). The IC50 values for GSIS (glucose raised from 2.8 to 16.7 mM) were G5.6 43 ± 1a, P5.6 29 ± 1b, G20 20 ± 2c and P20 18 ± 1c nM rotenone (different letters indicate P b 0.05). To further understand the NAD(P)+ reduction within islets, timecourse and static determinations of NAD(P)H were performed in which the islets were incubated with no glucose to estimate the NAD(P)+ reduction rate (NRR) from endogenous sources. Incubations in 100% N2 atmosphere were used to inhibit mitochondrial activity (hypoxia). In a similar series of experiments, islets were incubated with 200 μM H2O2 to localize the inhibition/activation of NAD(P)+ reduction pathways. Fig. 6 shows that most islet NAD(P)H came from endogenous fuels. When challenged with 20 mM glucose the islets in the G20 group, which were the most glucose-responsive, doubled their NRR while maintaining the NAD(P)H content (Fig. 6E). In the other groups, the NRR increased by only 25–50% when exposed to the same concentration of glucose. Only in islets from the 5.6 group was there an increase in the NAD(P)H content in response to glucose stimulation (Fig. 6E), indicating a poor transfer of NADH electrons to the respiratory chain. High glucose and treatment with H2O2 increased the aerobic contribution to NRR. In all groups of islets, glucose had only a small stimulatory effect on the NRR under hypoxia (Fig. 6A–D). The results in Fig. 6 were used to estimate the cytoplasmic and mitochondrial NRRs from endogenous sources (fatty acids, amino acids, etc) or from glucose oxidation. The contribution of endogenous fuels to NRR differentiated islets in the poor glucose-responsive group G5.6 from those in the high glucose-responsive groups, G20 and P20 (Fig. 7A). A high NRR, induced by glucose, similar to that normally seen in adult islets was only observed in group G20. This glucose-responsive NRR was abolished when islets from group G20 were incubated with 200 μM H2O2 (Fig. 7A). When islets in groups G5.6 and G20 were incubated for 3 h in 200 μM H2O2 there was a reversible decrease in the NRR from
Fig. 4. (A) Glucose metabolism in islets exposed to the aminotransferase inhibitor, AOA (5 mM). Islets were incubated in KHBS with 2.8 or 16.7 mM glucose containing D-[U-14C] glucose, as described. (B) Insulin secretion of islets incubated for 1 h in KHBS with 2.8 or 16.7 mM glucose and 5 mM AOA. The values were normalized relative to those for control experiments (no AOA) performed simultaneously. The bars are the means ± S.E.M. of 12 experiments. Different letters indicate P b 0.05.
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Fig. 5. Inhibition of GSIS by rotenone. Islets were incubated in KHBS buffer containing 2.8 or 16.7 mM glucose and different concentrations of rotenone. The values are the fold increase in insulin secretion when the glucose concentration was raised from 2.8 to 16.7 mM. Each point is the mean ± S.E.M. of 8 independent experiments. Different letters indicate P b 0.05 between the entire curves.
endogenous fuels and from glucose. However, incubation with H2O2 increased the cytoplasmic glucose-derived NRR in islets from group G5.6; this increase was not seen in group P5.6 after long-term exposure to H2O2 (Fig. 7B). Except for group P20, incubation with H2O2 abolished the mitochondrial glucose-derived NRR in islets of all groups. Others have observed a similar H2O2-mediated inhibition of mitochondrial activity in mature islets exposed to this oxidant. Discussion Hydrogen peroxide is a small nonpolar molecule able to diffuse across lipid membranes and is actually known as an important signaling/reactive oxidant. Inside the cells, it targets mainly thiol-and Fe2+-containing molecules, resulting in H2O production and forming disulfide bonds and Fe3+ ions. Several proteins are sensitive to these modifications, shifting metabolic processes, signaling phosphorylation cascades and even binding to DNA, thus altering gene expression. The answer to H2O2 varies within mammalian cell types, as the concentrations at which modifications occur. These modifications range from mitotic stimulus to activation of apoptotic genes or celldefense ones (Veal et al., 2007). In pancreatic β-cells, H2O2 and other oxidants inhibit glucokinase and phosphofructokinase, target mitochondria for aconitase and uncouple ATP production/glucose oxidation, possibly resulting in cytochrome C leakage and activation of celldeath caspases (Fujimoto et al., 2007). In this study, we examined the mechanism by which a concentration of H2O2 borderline deleterious to β-cells (200 μM) and a supraphysiological concentration of glucose (20 mM) select islets with increased GSIS, and how both mechanisms are interconnected. A critical coupling step between glucose oxidation and insulin secretion by rat β-cells is the introduction of cytoplasmic NAD(P)H into the mitochondria matrix by the NADH shuttles (pyr/mal, glycerolphosphate and mal/asp shuttles). Indeed, the glucose-responsiveness of islets is thought to be triggered by NAD(P)H generated in the cytoplasm (Eto et al., 1999). Although these islets showed a high degree of integration between glucose oxidation and GSIS, inhibition of the mal/asp shuttle with AOA blocks glucose oxidation, but not insulin secretion (Fig. 4). Islets from adult rats have a marginal use of glucose via the PPP and NADPH in the cytoplasm derives mostly from the pyr/mal and mal/asp shuttles (MacDonald, 2003). The activity of the glycerol-phosphate and mal/asp shuttles increases the frequency of Ca2+ oscillations in the cytoplasm of glucose-stimulated islets and triggers Ca2+ entry into mitochondria to activate three Ca2+-dependent dehydrogenases of the TCAC (MacDonald et al., 2005a,b). In contrast to mature islets, the group P5.6 culture-selected islets from neonatal rats make extensive
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Fig. 6. (A, B, C and D) NAD(P)+ reduction rate. Islets were incubated for 3 h in KHBS containing 5% (v/v) MTS/PMS and 0 mM glucose (G0), 20 mM glucose (G20), 0 mM glucose + 200 μM H2O2 (H0) and 20 mM glucose + 200 μM H2O2 (H20). The incubations were carried out in a 95% O2 atmosphere that produced 140 μM O2 in KHBS solution, and then switched to a 100% N2 atmosphere that reduced the O2 concentration in the solution to 90 μM. The islets were subsequently incubated for another 3 h under hypoxia (hypox). The NRR of each sample was calculated as the change in NAD(P)H that caused a reduction in MTS/PMS over time. The values are the means ± SEM of 8 experiments. (E) NAD(P)H content of entire islets incubated for 1 h in KHBS containing 2.8 or 16.7 mM glucose. The values are the mean ± S.E.M. of 10 experiments. Different letters indicate P b 0.05.
use of the PPP. This metabolic difference increases the activity of PPP in response to high glucose and H2O2: in islets cultured with 5.6 mM glucose, H2O2 increased 3× the use of PPP, whereas in islets cultured with high glucose (20 mM), the use of PPP is already 2× high and was not altered (Fig. 1B). Fetal and neonatal islets have a deficient glycolysis when compared to mature islets, but normal mitochondrial processing of metabolites (Weinhaus et al., 1995). The increased activity of PPP can feed with NADPH the antioxidant and reductive biosynthesis pathways, altering the overall reducing status of the cell and mitochondria. The increased mal/asp shuttle activity also suggests that the islets cultivated with H2O2 (groups P5.6 and P20) have an increased provision of fuel for ATP production (Fig. 3C), what is a different condition of islets cultivated with high glucose (groups G20 and P20) (Fig. 3B). The pyr/mal shuttle is highly active in rat islets and exports malate from mitochondria, thereby removing reduced equivalents from the
mitochondrial matrix when β-cells are stimulated by glucose. Hence, this shuttle does not contribute directly to insulin secretion. The mal/asp and glycerol-phosphate shuttles insert NADH into the mitochondrial matrix or directly at complex II of the respiratory chain, which are inhibited by rotenone and antimicyn A, respectively (MacDonald et al., 2005a,b). The mal/asp shuttle appeared to be critical for glucose oxidation and GSIS in the control islets (G5.6), however (Fig. 4). In other groups, the increased expression of mMDH in islets cultured with H2O2 paralleled its requirement for GSIS, but not for glucose oxidation. Such apparent paradox reinforces the hypothesis that GSIS of neonatal islets is triggered by cytoplasmic NAD(P)H. This conclusion was particularly demonstrated by the fact that hypoxia provoked only a marginal effect on the NRR of control islets (G5.6), which was the less glucose-responsive (Fig. 6A). Islets were selected throughout the culture period, with necrosis and apoptosis leading to the destruction of those islets/cells unable to
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Fig. 7. NAD(P)+ reduction from endogenous fuels in the cytoplasm (endo.cyto), from endogenous fuels in mitochondria (endo.mito), from glucose in the cytoplasm (gluc.cyto) or from glucose-derived fuels in mitochondria (gluc.mito). The calculations were made as described in Materials and methods using values from Fig. 6 for islets incubated in the absence (A) or presence (B) of 200 μM H2O2. The total NRRs shown in the upper panel are the sum of the four fractions of each group. Statistical analyses were done only between islet culture groups. Different letters indicate P b 0.05.
counteract the damage imposed by H2O2 and/or high glucose concentrations (Stoppiglia et al., 2002). Although a higher NAD(P)H generation is expected in H2O2-selected islets (P5.6 and P20 groups), the NRRs of islets cultured with H2O2 were not higher than in other groups. Conversely, the stimulation of these islets with 20 mM glucose seems to increase a previously high mitochondrial NRR generated from endogenous substrates (Fig. 6B,D). This NRR does not support a high NAD(P)H content in group P5.6 islets, however (Fig. 6E), possibly due to the use of NAD(P)H equivalents to sustain repair mechanisms such as thiol reduction (Brennan et al., 2003). In H2O2-selected islets, it is agreeable that a high baseline mitochondrial NRR should protect from the imposed oxidative damage. A physiological importance of the mal/asp shuttle was suggested by the exposure of adult islets to the complex I inhibitor rotenone, what doubled the formation of non-superoxide radicals from glyceraldehyde. Exposure of the islets to the complex II inhibitor myxothiazol had no effect (Takahashi et al., 2004), indicating that mature islets mitochondria preferentially use electrons inserted through complex I instead of complex II. In the neonatal islets cultured with 20 mM glucose (G20 and P20 groups), the expression of mGPDH was increased 2–3 fold compared to their respective controls G5.6 and P5.6 (Fig. 3B). The glucose oxidation and insulin release of these “glucose-selected islets” became less dependent of the mal/asp shuttle (Fig. 4B), thus suggesting a higher use of the glycerolphosphate shuttle in islets cultured in high glucose media. The activity of the glycerol-phosphate shuttle has been associated with GSIS as a modulator of the mitochondrial activity, and a deficiency in this shuttle has been observed in models of type 2 diabetes (Giroix et al., 1992). All islets cultured with 20 mM glucose showed similar NRR from endogenous substrates (Fig. 7A) and an increased NAD(P)H
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content (Fig. 6E). Part of this higher NAD(P)H generation is thought to depend on an increased activity of the PPP (Fig. 1B). Both the PPP and the glycerol-phosphate shuttle, however, are known to participate in lipogenic pathways, the first providing reduced equivalents and the former generating glycerol from glucolytic derivatives. It is very plausible, too, that the so called mitochondrial endogenous fuels protecting mitochondria against H2O2 are in someway lipids. According to some works, the acyl-CoA synthesis correlates with alterations of the activity of the pyr/mal shuttle and is necessary for GSIS (Boucher et al., 2004; Remedi et al., 2006). On the other hand, the maximal NRR observed in islets stimulated with 20 mM glucose clearly did not match the GSIS (Fig. 1A), although it parallels glucose oxidation rate (Fig. 1B). This supports the idea of NADH shuttles limiting the carbon flux into the TCAC of neonatal islets (Fig. 7A, upper table and Fig. 4A). In acute exposition to H2O2, we observed that H2O2 targets preferentially the glucose-derived NRR, even abolishing it in islets of the G20 group. Islets from the group P20 (cultured with H2O2) did not show such increased NRR from glucose, although it kept GSIS and glucose oxidation rate responsive to glucose (Fig. 7B). Considering the highly glucose-sensitive processing of glucose derivatives in mitochondria of mature islets (Remedi et al., 2006), it is possible that these islets develop a highly glucose-sensitive mitochondrial NRR that is easily damaged by H2O2. Islets selected by cultured with H2O2 showed an increased cytoplasmic production of NAD(P)H from glucose and a significant mitochondrial processing of endogenous fuels, these islets being resistant to hydrogen peroxide. Since the chronic exposure of islets to high glucose concentrations may generate oxidative stress (see Fig. 1B, G20), the activation of PPP, cytoplasmic glucose-dependent NRR and glycerol-phosphate NADH shuttle appear to be characteristic features of islets that resist H2O2 (Figs. 1B and 7B) and can be assigned as suitable mechanisms for preserving the glucose-responsiveness of pancreatic islets. Finally, our results suggest that immature islets have the plasticity to modify the site and source of NAD(P)+ reduction, thereby changing the glucose oxidation rate and the coupling to ATP production. Islets cultured at low glucose concentrations are unable to maintain high levels of NAD(P)H. When challenged with H2O2, these islets increase the mal/asp shuttle activity and generate ATP from reduced equivalents in the cytoplasm. In contrast, islets cultured at high glucose concentrations generate ATP from cytoplasmic processing of glucose and aerobic oxidation of endogenous fuels, possibly through the glycerol-phosphate shuttle, with a consequent increase in GSIS. Contrary to glucotoxicity in adult islets, the keeping of a higher mitochondrial generation of reduced equivalents appears to protect neonatal islets against the deleterious effects of H2O2. Acknowledgments This work was partially supported by the Brazilian foundations: CAPES, CNPQ and FAPESP. We are grateful to Dr. Nicola Conran for editing the English. References Anello, M., Ucciardello, V., Piro, S., Patane, G., Frittitta, L., Calabrese, V., Giuffrida, S.A.M., Vigneri, R., Purrello, F., Rabuazzo, A.M., 2001. Chronic exposure to high leucine impairs glucose-induced insulin release by lowering the ATP-to-ADP ratio. American Journal of Physiology — Endocrinology and Metabolism 281, 1082–1087. Asanuma, N., Aizawa, T., Sato, Y., Schermerhorn, T., Komatsu, M., Sharp, G.W.G., Hashizume, K., 1997. Two signaling pathways, from the upper glycolytic flux and from the mitochondria, converge to potentiate insulin release. Endocrinology 138, 751–755. Boucher, A., Lu, D., Burgess, S.C., Potts, S.T., Jensen, M.V., Mulder, H., Wang, M.Y., Unger, R.H., Sherry, A.D., Newgard, C.B., 2004. Biochemical mechanism of lipid-induced impairment of glucose-stimulated insulin secretion and reversal with a malate analogue. Journal of Biological Chemistry 279, 27263–27271. Brennan, L., Corless, M., Hewage, C., Malthouse, J.P.G., McClenaghan, N.H., Flatt, P.R., Newsholme, P., 2003. 13C NMR analysis reveals a link between L-glutamine
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metabolism, D-glucose metabolism and γ-glutamyl cycle activity in a clonal pancreatic beta cell line. Diabetologia 46, 1512–1521. Delghingaro-Augusto, V., Ferreira, F., Bordin, S., Amaral, M.E.C., Toyama, M.H., Boschero, A.C., Carneiro, E.M., 2004. A low protein diet alters gene expression in rat pancreatic islets. Journal of Nutrition 134 (2), 321–327. Dubois, M., Vacher, P., Roger, B., Huyghe, D., Vandewalle, B., Kerr-Conte, J., Pattou, F., Moustaid-Moussa, N., Lang, J., 2007. Glucotoxicity inhibits late steps of insulin exocytosis. Endocrinology 148 (4), 1605–1614. Dukes, I.D., McIntyre, M.S., Mertz, R.J., Philipson, L.H., Roe, M.W., Spencer, B., Worley III, J.F., 1994. Dependence on NADH produced during glycolysis for β-cell glucose signaling. Journal of Biological Chemistry 269, 10979–10982. Eto, K., Tsubamoto, Y., Terauchi, Y., Sugiyama, T., Kishimoto, T., Takahashi, N., Yamauchi, N., Kubota, N., Murayama, S., Aizawa, S., Kasai, H., Yazaki, Y., Kadowakil, T., 1999. Role of NADH shuttle system in glucose-induced activation of mitochondrial metabolism and insulin secretion. Science 283, 981–985. Fujimoto, S., Nabe, K., Takehiro, M., Shimodahira, M., Kajikawa, M., Takeda, T., Mukai, E., Inagaki, N., Seino, Y., 2007. Impaired metabolism-secretion coupling in pancreatic β-cells: role of determinants of mitochondrial ATP production. Diaberes Research & Clinical Practice 77, 2–10. Gao, Z., Young, R.A., Li, G., Najafi, H., Buetther, C., Sukumvanich, S.S., Wong, R.K., Wolf, B.A., Matschinsky, F.M., 2003. Distinguishing features of leucine and α-ketoisocaproato sensing in pancreatic β-cells. Endocrinology 144, 1949–1957. Giroix, M.H., Rasschaert, J., Bailbe, D., Leclercq-Meyer, V., Sener, A., Portha, B., Malaisse, W.J., 1992. Study of hexose transport, glycerol phosphate shuttle and Krebs cycle in islets of adult rats injected with streptozotocin during the neonatal period. Molecular and Cellular Endocrinology 83, 95–104. Katz, J., Wood, H.G., 1963. The use of C14O2 yields from glucose-1- and -6-C14 for the evaluation of the pathways of glucose metabolism. Journal of Biological Chemistry 238, 517–523. Khaldi, M.Z., Guiot, Y., Gilon, P., Henkin, J.C., Jonas, J.C., 2004. Increased glucose sensitivity of both triggering and amplifying pathways of insulin secretion in rat islets cultured for 1 wk in high glucose. American Journal of Physiology — Endocrinology and Metababolism 287, 207–217.
MacDonald, M.J., 2003. The export of metabolites from mitochondria and anaplerosis in insulin secretion. Biochimica et Biophysica Acta 1619 (1), 77–88. MacDonald, M.J., Fahien, L.A., Brown, L.J., Hasan, N.M., Buss, J.D., Kendrick, M.A., 2005a. Perspective: emerging evidence for signaling roles of mitochondrial anaplerotic products in insulin secretion. American Journal of Physiology — Endocrinology and Metabolism 288, E1–E15. MacDonald, P.E., Joseph, J.W., Rorsman, P., 2005b. Glucose-sensing mechanisms in pancreatic β-cells. Philosophical transactions of the royal society B 360 (1464), 2211–2225. Mears, D., 2004. Regulation of insulin secretion in islets of Langerhans by Ca2+ channels. Journal of Membrane Biology 200 (2), 57–66. Remedi, M.S., Nichols, C.G., Koster, J.C., 2006. The mitochondria and insulin release: Nnt just a passing relationship. Cell Metabolism 3 (1), 5–7. Stoppiglia, L.F., Nogueira, T.A., Leite, A.R., Carneiro, E.M., Boschero, A.C., 2002. Protective effect of D-glucose, L-leucine and fetal calf serum against oxidative stress in neonatal pancreatic islets. Biochimica et Biophysica Acta 1588 (2), 113–118. Stoppiglia, L.F., Rezende, L.F., Ferreira, F., Filiputti, E., Carneiro, E.M., Boschero, A.C., 2004. Characterization of the peroxidase system at low H2O2 concentrations in isolated neonatal rat islets. Biochimica et Biophysica Acta 1690 (2), 159–168. Takahashi, H., Tran, P.O.T., Leroy, E., Harmon, J.S., Tanaka, Y., Robertson, R.P., 2004. DGlyceraldehyde causes production of intracellular peroxide in pancreatic islets, oxidative stress, and defective beta cell function via non-mitochondrial pathways. Journal of Biological Chemistry 279 (36), 37316–37323. Tan, C., Tuch, B.E., Too, J., Brown, S.A., 2002. Role of NADH shuttles in glucose-induced insulin secretion from fetal β-cells. Diabetes 51, 2989–2996. Veal, E.A., Day, A.M., Morgan, B.A., 2007. Hydrogen peroxide sensing and signaling. Molecular Cell 26, 1–14. Weinhaus, A.J., Poronnik, P., Cook, D.I., Tuch, B.E., 1995. Insulin secretagogues, but not glucose, stimulate an increase in (Ca2+)i in fetal rat β-cell. Diabetes 44 (1), 118–124. Westermark, P.O., Kotaleski, J.H., Björklund, A., Grill, V., Lansner, A., 2007. A mathematical model of the mitochondrial NADH shuttles and anaplerosis in the pancreatic β-cell. American Journal of Physiology — Endocrinology and Metabolism 292, E373–E393.
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Life Sciences j o u r n a l h o m e p a g e : w w w. e l s ev i e r. c o m / l o c a t e / l i f e s c i e
Altered NAD(P)H production in neonatal rat islets resistant to H2O2 Luiz F. Stoppiglia a, Luiz F. Rezende b, Ana P.G. Cappelli b, Fabiano Ferreira c, Antonio C. Boschero b,⁎ a b c
Departamento de Química, Instituto de Ciências Exatas e da Terra, Universidade Federal de Cuiabá, MT, Brazil Departamento de Fisiologia e Biofísica, Instituto de Biologia, Universidade Estadual de Campinas (UNICAMP), Brazil Departamento de Fisiologia e Farmacologia, Centro de Ciências Biológicas, Universidade Federal de Pernambuco, PE, Brazil
a r t i c l e
i n f o
Article history: Received 4 January 2008 Accepted 4 September 2008 Keywords: Glucose oxidation H2O2 NADH shuttles Neonatal pancreatic islets Pentose phosphate pathway
a b s t r a c t Aims: We determined the involvement of NAD(P)H generation ability on the resistance of pancreatic islets Bcells to oxidative stress caused by culture exposition to H2O2. Main methods: We cultured isolated neonatal Wistar rat islets for four days in medium containing 5.6 or 20 mM glucose, with or without H2O2 (200 μM), and analyzed several parameters associated with islet survival in different media. High glucose was used since it protects neonatal islets against the loss of GSIS. Key findings: While none of the culture conditions increased the rate of NAD(P)H content at 16.7 mM glucose, the islets resistant to H2O2 and those exposed to 20 mM glucose showed a greater use of the pentose phosphate pathway and increased ATP synthesis from glucose. Significance: Oxidative stress contributes to the loss of glucose-induced insulin secretion (GSIS) during the onset of diabetes mellitus. Although immature rat islets have reduced GSIS compared to mature islets, they adapt better to oxidative stress and are a good model for understanding the causes involved in the destruction or survival of islet cells. These data support the idea that GSIS and resistance against oxidative stress in immature islets rely on NADH shuttle activities, with little contribution of reduced equivalents from the tricarboxylic acid cycle (TCAC). © 2008 Elsevier Inc. All rights reserved.
Introduction Malfunction of pancreatic islets impairs glucose homeostasis and can lead to diabetes mellitus. Insulin secretion occurs after an increase in the ATP/ADP ratio in B-cells via the Ca2+ dependent and independent pathways. Ca2+-dependent insulin secretion is referred to as the triggering pathway, whereas Ca2+-independent secretion accounts for the amplifying pathway (Asanuma et al., 1997). In humans and experimental animals, a combination of chronic hyperglycemia, high fatty acid concentration, cytokines and reactive oxygen species (ROS) reduce the responsiveness of islets to glucose by affecting both pathways. Impairment of GSIS is mainly assigned to depletion of insulin-promoter ligands, B-cell death, inhibition of glucolytic key enzymes, mitochondria damage, shift of glucose oxidation to other pathways, and defects in cellular calcium handling. On the other hand, chronic exposure to high levels of glucose increases ATP/ADP ratio and NAD(P)H concentrations, UCP2 activity and basal cytoplasmic Ca2+. Chronic hyperglycemia is believed to be a main cause of the loss of GSIS (Mears, 2004; Khaldi et al., 2004; Dubois et al., 2007).
⁎ Corresponding author. Departamento de Fisiologia e Biofísica, Instituto de Biologia, Universidade Estadual de Campinas (UNICAMP), CP 6109, Campinas, 13083-970, SP, Brazil. Fax: +55 19 3521 6185. E-mail address: [email protected] (A.C. Boschero). 0024-3205/$ – see front matter © 2008 Elsevier Inc. All rights reserved. doi:10.1016/j.lfs.2008.09.012
Mitochondrial oxidation of glycolysis-derived NADH is an essential trigger for insulin secretion (Eto et al., 1999). NADH is transported actively against its concentration gradient from cytosol the mitochondrial matrix by NADH shuttles, as a required complement to pyruvatederived NADH/FADH2 (Dukes et al., 1994). This is provided by the malate/aspartate and glycerol-phosphate shuttles. The first consumes the proton gradient through an electrogenic glutamate/aspartate antiporter at the inner mitochondrial membrane; the second wastes the O2 motive force by use of a FADH2-linked dehydrogenase to bypass proton-pumps and insert electrons directly in the respiratory chain. The oxidation of cytoplasmic NADH by mitochondria gives more energy equivalents than what is lost by the shuttles and equilibrates the necessary NADH/NAD ratio, accounting for 50% of the glycolytic flux (MacDonald et al., 2005a,b; Westermark et al., 2007). In rat islets, a counter-current mechanism uses the pyruvate and NAD (P)H gradients through the mitochondrial inner membrane to export NADH equivalents from matrix to cytoplasm, provided mainly by the pyruvate/malate shuttle. Malate is exported in large amounts by the mitochondria of mouse and rat islets and signs for TCAC cycle activity. Some malate is converted to pyruvate by a NADP-linked malic enzyme, which maintains the high cytosolic NADPH/NADP ratio in B-cells. The citrate/isocitrate shuttle may be not critical for insulin secretion, since insulin secretagogues do not cause exportation of isocitrate from rat mitochondria (MacDonald et al., 2005b). Neonatal rat islets, cultured in high concentrations of glucose, but not L-leucine or other secretagogues, present augmented glucose-
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stimulated insulin secretion (GSIS) and resistance to the deleterious effects of H2O2 (Stoppiglia et al., 2002; Anello et al., 2001). Despite the similarity between insulin secretion induced by glucose and L-leucine, the latter lowers cytoplasmic glutamate and increases aspartate levels through allosteric activation of mitochondrial glutamate dehydrogenase (Gao et al., 2003). Rat islets apparently have a sufficiently high glutamate content to maintain insulin secretion for 2–3 h through glutamate dehydrogenase and mal/asp shuttle activities. However, the chronic use of non-pyruvate-producing metabolites impairs the shuttle activity by depleting mitochondrial aspartate (MacDonald et al., 2005a,b). The activity of these NADH shuttles is amplified several times when islets progress from a low-responsiveness fetal state to an adult condition (Tan et al., 2002). We have previously suggested that neonatal rat islets have a different mechanism of defense against ROS by offering high concentrations of NAD(P)H to a scavenger enzyme-poor system (Stoppiglia et al., 2004). This mechanism may indicate that β-cells can actively modify the production of reduced equivalents from their high glucolytic rate and direct it to survival purposes, possibly altering insulin secretion. In this work, we investigated whether the defense status against H2O2 is sufficient to increase cytoplasmic NAD(P)H production and how this affects glucose-stimulated insulin secretion. Materials and methods H2O2 concentrations All H2O2 concentrations cited have been carefully tested by H2O2 colorimetric reaction with a horseradish peroxidase/4-aminoantipyridine system and with potassium iodide as described (Stoppiglia et al., 2004). In previous experiments, we tested different concentrations of H2O2 for deleterious activity in cultured islets (Stoppiglia et al., 2002). A borderline deleterious activity with observable alteration of metabolic activity was found to happen with 200 μM of H2O2 after a 4 day incubation period. The same H2O2 concentration appeared to cause much more damage to adult than to neonatal rat islets. This concentration was used for the experiments described, else being described in each case. Chemicals Radiochemicals were from G.E. Heath Care (UK), the aminotransferase inhibitor amino-oxyacetate (carboxymethoxylamine) was from Sigma-Aldrich (St. Louis/MO, USA), the MTS/PMS preparation was from a CellTitter96 aqueous assay of Promega (Madison/WI, USA) and all RT-PCR reagents were from Invitrogen (Carlsbad/CA, USA). The other reagents were from Sigma-Aldrich, whenever specified. Islet isolation and culture Neonatal (1–2-day-old) Wistar rats were from the animal facilities at the State University of Campinas. After decapitation, the islets were isolated by collagenase (EC 3.4.24.3) digestion of pancreata in Hank's balanced salt solution (in mM: 137 NaCl, 5.5 KCl, 4.5 NaHCO3, 0.4 KH2PO4, 0.4 Na2HPO4, 0.8 MgSO4, 1.5 CaCl2, pH 7.4). The islets were extensively washed in sterile Hank's solution and cultured in RPMI 1640 medium supplemented with 2 g NaHCO3/l, 1% (v/v) penicillin/streptomycin and 10 mM D-glucose, pH 7.4. Approximately 1000 islets/dish were maintained at 37 °C in a humidified atmosphere with 3% CO2 for two days before further additions. The culture medium contained glucose (5.6 mM or 20 mM) with or without 200 μM H2O2, for an additional four days. The medium was renewed every 24 h. The islet experimental groups were assigned according to the culture conditions: G5.6 (islets cultured in 5.6 mM glucose), P5.6 (5.6 mM glucose and 200 μM H2O2), G20 (20 mM glucose) and P20 (20 mM glucose and 200 μM H2O2). A similar procedure is described (Stoppiglia et al., 2002).
Insulin secretion Batches of 10 islets were each incubated for 30 min at 37 °C in Krebs–Hepes buffered saline (KHBS, in mM: 115 NaCl, 10 NaHCO3, 5 KCl, 1 MgCl2, 2.5 CaCl2, 15 Hepes) containing 0.5 g BSA/l and 5.6 mM glucose, pH 7.4, and equilibrated with 95% O2 and 5% CO2. The medium was discarded and the islets were incubated for a further 1 h in 1 ml of KHBS containing 2.8 or 16.7 mM glucose with the additions described in the corresponding figure legends. The supernatant was collected and the insulin concentration was measured by radioimmunoassay (Delghingaro-Augusto et al., 2004). Glucose uptake and metabolism Batches of 50 islets each were incubated for 2 h at 37 °C in KHBS containing 2.8 or 16.7 mM glucose with trace amounts of either 14 14 14 D-[U- C]glucose or D-[1- C]glucose to measure CO2 production. HCl (1 N) was added to the batches to stop respiration and the 14CO2 formed was collected for 1 h at 4 °C in 1 N NaOH. For the glucose uptake assays, the batches were incubated under similar conditions with 2-deoxy-D-[U-14C]glucose for 2 h, after which the islets were washed twice with KHBS at 4 °C, sedimented and homogenized in the presence of Trizol (Invitrogen). The partitioning of glucose between the TCAC and the pentose phosphate pathway (PPP) was calculated by assuming that the total 14CO2 produced from labeled glucose was from the TCAC and the PPP. A detailed description of the data processing is given by Katz and Wood, 1963. In short, if we define D-[1-14C]glucose and D-[U-14C]glucose conversion to CO2 as G1 and GU, respectively, it is possible to construct a linear system of two equation were G1 is the sum of TCAC and PPP; because only one 14 C carbon is detected per glucose molecule. However, GU is obtained as the sum of PPP glucose utilization and 6 times the TCAC glucose utilization, because TCAC generates 6 14C atoms per glucose molecule oxidized. Then, the amount of glucose that passed through the TCAC was calculated as (GU − G1)/5, and that which passed through the PPP as (6 × G1 − GU)/5. RT-PCR Groups of 1000 islets were homogenized in Trizol followed by phenol–chloroform RNA extraction, according to the manufacturer's instructions. The quality of the RNA was assessed by agarose gel electrophoresis. One to five micrograms of total RNA were transferred to reaction vials containing 1 μg of dNTP mix. The reactions were incubated for 5 min at 65 °C before adding 150 ng of random primers (10 min at 25 °C) followed by 14.3 mM MgCl2, 2.8 mM DTT and 0.4 U/μl RNase-out (2 min at 42 °C), and 1.25 U/μl RNA Super Script II. The mixtures were then incubated at 42 °C for 50 min and at 70 °C for 15 min, followed by cooling to 4 °C. The resulting cDNAs were diluted in PCR buffer (60 mM Tris–HCl, 1.5 mM MgCl2, 15 mM NH4SO4, pH 10) with 50 mM MgCl2, 0.3 mM each of dATP, dCTP, dGTP and dTTP and 2.5 U/μl Taq DNA polymerase (Gibco/BRL), followed by the addition of 10 mM of forward and reverse primers. PCR amplification of cDNA was carried out with a GeneAmp PCR System 9700 (Applied Biosystems, Foster City, CA, USA) and involved 32 cycles of denaturation for 1 min at 94 °C, annealing for 1 min at referred temperatures, extension for 1.5 min at 72 °C, and a final extension for 7 min at 72 °C. The annealing temperatures and the number of cycles used for amplification are shown in the corresponding legends. Following amplification, the cDNA fragments were analyzed on 1.6% agarose gels containing a 100 bp DNA molecular weight ladder (Gibco/BRL). The PCR products were detected by staining with ethidium bromide and were documented with a Gel Doc EQ photodocumentation system (BioRad). Internal control reactions were performed for the RPS-29 subunit of the small ribosomal unit 40S, since this showed the best homogeneity
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between treated and non-treated groups. The primers were designed and tested against the Rattus norvergicus genome (Gene Bank) to ensure no amplification of other cDNAs. The oligonucleotide primers used were as follows: RPS-29 (forward) 5′AGG CAA GAT GGG TCA CCA GC3′; RPS-29 (reverse) 5′AGT CGA ATC CAT TCA GGT CG3′; rat pro-insulin 2 (forward) 5′TTG CAG TAG TTC TCC AGT T3′; rat pro-insulin 2 (reverse) 5′ATT GTT CCA ACA TGG CCC TGT3′; GLUT2 (forward) 5′CAT TGC TGG AAG CGT ATC AG3′; GLUT2 (reverse) 5′GAG ACC TTC TGC TCA GTC GAC G3′; mitochondrial pyruvate dehydrogenase subunit E1 alpha 2 (mPDH) (forward) 5′TCA AGT ACT ACA GGA TGA TG3′; mPDH (reverse) 5′GGC GTA CAT GTG CAT TGA TC3′; mitochondrial glycerol-3-phosphate dehydrogenase (mGPDH) (forward) 5′AGA AAG TCT GCA TCG TAG GCT3′; mGPDH (reverse) 5′GGA AGT TGG GTG TTT GCA TCA3′; mitochondrial malate dehydrogenase (mMDH) (forward) 5′CCT GAA GCC ATG ATT TGC ATC3′; mMDH (reverse) 5′TTC TTG ATG GAG GCT TTC AGC3′; glucose-6-phosphate dehydrogenase (G6PDH) (forward) 5′AGC TCC AAT CAA CTG TCG AAC3′; G6PDH (reverse) 5′TCC TCA GGG TTG AAG AAC ATG3′. All annealing temperatures and number of cycles were chosen to provide maximum sensitivity in the assay. A similar procedure is described (Delghingaro-Augusto et al., 2004). NAD(P)H determination NAD(P)H was measured by the coupled reduction of phenazine methosulfate (PMS) and subsequent transfer of electrons to the tetrazolium salt MTS, both of which are membrane permeable. The MTS and PMS solutions were provided as parts of the CellTitter96 aqueous assay (Promega) and were mixed according to the manufacturer's instructions. There was little change in the absorbance at 492 nm absorbance with concentrations of NAD(P)H b10 μmol/l. Because of the important interference by proteins bound to NAD(P)H, the standard curves were less accurate at low concentrations of NAD(P)H. However, spectroscopic analysis showed an increase in light absorption at 650 nm of ~ 10− 3 cm− 1 islet− 1 when different numbers of islets where disrupted in MTS/PMS solution containing 2 mg of BSA/ml. Heat-denaturated islet homogenates were used as negative controls. Static measurements of NAD(P)H were done by incubating groups of 200 islets in KHBS containing 2.8 or 16.7 mM of glucose, which reproduced the same conditions used in the insulin secretion experiments. The islets were then washed in ice-cold Hanks solution and immediately sonicated in 150 μl of this same solution. The homogenates were centrifuged (10 000 g × 2 min) to remove islet debris and aliquots of the supernatants were then added to the MTS/ PMS solution and incubated for 30 min at room temperature before recording the absorbances at 650 nm and 405 nm (background). Mixtures with no islets were used as blanks. In each experiment, the changes in the absorbance at 650 nm were normalized to the mean absorbance of G5.6 samples incubated with 16.7 mM of glucose. Dynamic measurements of NAD(P)H were done by incubating groups of 20 islets in 200 μl of KHBS containing 0 or 20 mM of glucose and 5% (v/v) of MTS/PMS. This concentration of MTS/PMS provided the best sensitivity and was chosen after testing concentrations from 1% to 20% (v/v). As judged by the transient rate of MTS/PMS reduction, concentrations of MTS/PMS N7% caused cell death through NAD(P)H depletion. Samples were incubated for 3 h in a 95% O2 + 5% CO2 atmosphere (140 μM O2 in KHBS) and then changed to a 100% N2 atmosphere (90 μM O2) for a further 3 h to cause hypoxia. Some of the samples were returned to the 95% O2 atmosphere and cell death caused by hypoxia was estimated to be b20%. Samples with no islets were used as blanks and the absorbances at 492 nm were recorded every 30 min. Standard curves of NADPH were used to calculate the decrease in the NAD(P)H content of the samples. The NAD(P)+ reduction rate (NRR) was determined from the increase in MTS/PMS absorbance in each sample.
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Partitioning of the NAD(P)+ reduction rate The partitioning of the NRR was calculated by assuming that the NRR consisted of four components: cytoplasmic NRR from endogenous sources (endo.cyto), cytoplasmic NRR from glucose oxidation (gluc. cyto), mitochondrial NRR from endogenous sources (endo.mito) and mitochondrial NRR from glucose-mediated metabolite oxidation (gluc. mito). The mitochondrial components were assumed to depend on an adequate oxygen supply, whereas the cytoplasmic components were essentially anaerobic. The NRR from endogenous sources was estimated from values observed in the absence of glucose. These endogenous fuels were presumed to be mostly fatty acids and amino acids. Statistics The results were expressed as means ± S.E.M. Point-to-point comparisons were done using Students t-test. The groups were compared by two-way ANOVA followed by the unpaired Tukey– Kramer test. A value of P b 0.05 indicated significance. However, in the RT-PCR experiments, significance was only considered when P b 0.001. In each figure, different letters were used to indicate significant differences in comparisons between the groups plotted. Identical letters above two groups indicate that no statistical difference was seen. Greek letters were used to indicate a separated set of results compared only between them. Results At the end of the culture period, the GSIS of the control group G5.6 was similar to that of fresh isolated islets (not shown). Islets cultured
Fig. 1. (A) Insulin secretion by islets cultured 4 days with 5.6 or 20 mM glucose (G5.6 and G20), in the absence or presence of 200 μM H2O2 (P5.6 and P20). Islets were preincubated for 30 min in KHBS containing 5.6 mM glucose, as described, and then incubated either with 2.8 or 16.7 mM glucose for 1 h. The bars represent means ± S.E.M. of 8 independent experiments, with different letters indicating P b 0.05. (B) Partitioning of metabolized glucose between the TCAC (open part of bars) and the PPP (filled bars). The values were calculated from the amount of 14CO2 produced by D-[U-14C]glucose and 14 D-[1- C]glucose metabolism in vials containing KHBS with 2.8 or 16.7 mM glucose, as described. All of the samples gave similar results in the presence of 2.8 mM glucose. The bars are the means ± S.E.M. of 8 experiments performed simultaneously with D-[U-14C] glucose and D-[1-14C]glucose. Different letters indicate P b 0.05 for comparisons within the TCAC and within the PPP.
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Fig. 2. (A) Glucose uptake in islets cultured in the same conditions as in Fig. 1. Islets were incubated for 120 min in KHBS with 2.8 or 16.7 mM glucose containing equal amounts of 2-deoxy-D-[U-14C]glucose, and then washed and homogenized with Trizol. The bars are the means± S.E.M. of at least 8 experiments. Different letters indicate P b 0.05. (B) Effect of incubation with high glucose/H2O2 on islet GLUT2 mRNA levels. The annealing temperatures and cycle numbers were 55 °C and 29 cycles for GLUT2 and 57 °C and 29 for RPS-29, respectively. RPS-29 was used as an internal control showing no variation among the conditions tested. The bars are the mean ± S.E.M. of 12 experiments. There were no statistical differences in mRNA content.
with high glucose or H2O2 showed increased glucose oxidation and higher GSIS compared to the controls (Fig. 1), but reduced coupling of GSIS to glucose oxidation. When islets are incubated in 2.8 or 16.7 mM
glucose, there are increases in glucose oxidation and insulin secretion. Comparing the increase of insulin secretion/increase of glucose oxidation, we have the following values (in Δng insulin/Δpmol glucose): a b b ab G5.6 1.4 ± 0.1 , P5.6 0.5 ± 0.1 , G20 0.7 ± 0.2 and P20 1.1 ± 0.2 (see Materials and methods for meaning of letters). This suggests that glucose and H2O2 may share a mechanism that diverts carbons from ATP production. Despite a significant loss of islets when cultured with H2O2, treatments with high glucose or H2O2 did not change GLUT2 mRNA content (Fig. 2B) nor decrease glucose uptake (Fig. 2A). In addition, in the post-culture islet population, there was no significant change in the insulin content (data not shown). Exposure to high glucose and H2O2 selected islets with increased oxidation of glucose through the PPP (Fig. 1B), which produces cytoplasmic NADPH. This selection was consistent with an increased expression of G6PDH mRNA (Fig. 3A). The expression of genes encoding proteins related to NAD(P)H that transit between mitochondria and the cytoplasm was also modified by glucose and H2O2 (Fig. 3B,C): mGPDH expression was increased in islets exposed to high glucose, whereas mMDH expression was higher in islets resistant to H2O2. The expression of mPDH was unaffected by either of these treatments (Fig. 3D). When challenged with glucose, islets resistant to H2O2 were less dependent on glucose oxidation via the mal/asp shuttle, as indicated by the use of the transaminase inhibitor amino-oxyacetate (Fig. 4A). Remarkably, the participation of the mal/asp shuttle in glucose oxidation was activated by a glucose challenge only in the P5.6 group, whereas this shuttle was required for GSIS in all other groups (Fig. 4B). To induce insulin release, NADH from the TCAC and cytoplasm must enter the respiratory chain and donate electrons to ATP synthesis. Electrons from NADH are donated to complex I or directly to complex II through the activity of a FADH2-linked glycerolphosphate shuttle. When complex I was inhibited with rotenone, exposure to high glucose and H2O2 increased the use of mitochondrial NADH for GSIS, although these effects were not synergic or additive
Fig. 3. Effect of exposure to high glucose and H2O2 on the mRNA levels of (A) G6PDH, (B) mG3PDH, (C) mMDH and (D) mPDH. The RT-PCR annealing temperatures and cycle numbers used were 57 °C and 31 cycles (G6PDH and mG3PDH), 57 °C and 35 cycles (mMDH) and 55 °C and 32 cycles (mPDH). RPS-29 cDNA was amplified using 57 °C and 29 cycles, and showed no variation among the conditions tested. Bars are the means ± S.E.M. of 12 experiments. Different letters indicate P b 0.001.
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(Fig. 5). The IC50 values for GSIS (glucose raised from 2.8 to 16.7 mM) were G5.6 43 ± 1a, P5.6 29 ± 1b, G20 20 ± 2c and P20 18 ± 1c nM rotenone (different letters indicate P b 0.05). To further understand the NAD(P)+ reduction within islets, timecourse and static determinations of NAD(P)H were performed in which the islets were incubated with no glucose to estimate the NAD(P)+ reduction rate (NRR) from endogenous sources. Incubations in 100% N2 atmosphere were used to inhibit mitochondrial activity (hypoxia). In a similar series of experiments, islets were incubated with 200 μM H2O2 to localize the inhibition/activation of NAD(P)+ reduction pathways. Fig. 6 shows that most islet NAD(P)H came from endogenous fuels. When challenged with 20 mM glucose the islets in the G20 group, which were the most glucose-responsive, doubled their NRR while maintaining the NAD(P)H content (Fig. 6E). In the other groups, the NRR increased by only 25–50% when exposed to the same concentration of glucose. Only in islets from the 5.6 group was there an increase in the NAD(P)H content in response to glucose stimulation (Fig. 6E), indicating a poor transfer of NADH electrons to the respiratory chain. High glucose and treatment with H2O2 increased the aerobic contribution to NRR. In all groups of islets, glucose had only a small stimulatory effect on the NRR under hypoxia (Fig. 6A–D). The results in Fig. 6 were used to estimate the cytoplasmic and mitochondrial NRRs from endogenous sources (fatty acids, amino acids, etc) or from glucose oxidation. The contribution of endogenous fuels to NRR differentiated islets in the poor glucose-responsive group G5.6 from those in the high glucose-responsive groups, G20 and P20 (Fig. 7A). A high NRR, induced by glucose, similar to that normally seen in adult islets was only observed in group G20. This glucose-responsive NRR was abolished when islets from group G20 were incubated with 200 μM H2O2 (Fig. 7A). When islets in groups G5.6 and G20 were incubated for 3 h in 200 μM H2O2 there was a reversible decrease in the NRR from
Fig. 4. (A) Glucose metabolism in islets exposed to the aminotransferase inhibitor, AOA (5 mM). Islets were incubated in KHBS with 2.8 or 16.7 mM glucose containing D-[U-14C] glucose, as described. (B) Insulin secretion of islets incubated for 1 h in KHBS with 2.8 or 16.7 mM glucose and 5 mM AOA. The values were normalized relative to those for control experiments (no AOA) performed simultaneously. The bars are the means ± S.E.M. of 12 experiments. Different letters indicate P b 0.05.
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Fig. 5. Inhibition of GSIS by rotenone. Islets were incubated in KHBS buffer containing 2.8 or 16.7 mM glucose and different concentrations of rotenone. The values are the fold increase in insulin secretion when the glucose concentration was raised from 2.8 to 16.7 mM. Each point is the mean ± S.E.M. of 8 independent experiments. Different letters indicate P b 0.05 between the entire curves.
endogenous fuels and from glucose. However, incubation with H2O2 increased the cytoplasmic glucose-derived NRR in islets from group G5.6; this increase was not seen in group P5.6 after long-term exposure to H2O2 (Fig. 7B). Except for group P20, incubation with H2O2 abolished the mitochondrial glucose-derived NRR in islets of all groups. Others have observed a similar H2O2-mediated inhibition of mitochondrial activity in mature islets exposed to this oxidant. Discussion Hydrogen peroxide is a small nonpolar molecule able to diffuse across lipid membranes and is actually known as an important signaling/reactive oxidant. Inside the cells, it targets mainly thiol-and Fe2+-containing molecules, resulting in H2O production and forming disulfide bonds and Fe3+ ions. Several proteins are sensitive to these modifications, shifting metabolic processes, signaling phosphorylation cascades and even binding to DNA, thus altering gene expression. The answer to H2O2 varies within mammalian cell types, as the concentrations at which modifications occur. These modifications range from mitotic stimulus to activation of apoptotic genes or celldefense ones (Veal et al., 2007). In pancreatic β-cells, H2O2 and other oxidants inhibit glucokinase and phosphofructokinase, target mitochondria for aconitase and uncouple ATP production/glucose oxidation, possibly resulting in cytochrome C leakage and activation of celldeath caspases (Fujimoto et al., 2007). In this study, we examined the mechanism by which a concentration of H2O2 borderline deleterious to β-cells (200 μM) and a supraphysiological concentration of glucose (20 mM) select islets with increased GSIS, and how both mechanisms are interconnected. A critical coupling step between glucose oxidation and insulin secretion by rat β-cells is the introduction of cytoplasmic NAD(P)H into the mitochondria matrix by the NADH shuttles (pyr/mal, glycerolphosphate and mal/asp shuttles). Indeed, the glucose-responsiveness of islets is thought to be triggered by NAD(P)H generated in the cytoplasm (Eto et al., 1999). Although these islets showed a high degree of integration between glucose oxidation and GSIS, inhibition of the mal/asp shuttle with AOA blocks glucose oxidation, but not insulin secretion (Fig. 4). Islets from adult rats have a marginal use of glucose via the PPP and NADPH in the cytoplasm derives mostly from the pyr/mal and mal/asp shuttles (MacDonald, 2003). The activity of the glycerol-phosphate and mal/asp shuttles increases the frequency of Ca2+ oscillations in the cytoplasm of glucose-stimulated islets and triggers Ca2+ entry into mitochondria to activate three Ca2+-dependent dehydrogenases of the TCAC (MacDonald et al., 2005a,b). In contrast to mature islets, the group P5.6 culture-selected islets from neonatal rats make extensive
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Fig. 6. (A, B, C and D) NAD(P)+ reduction rate. Islets were incubated for 3 h in KHBS containing 5% (v/v) MTS/PMS and 0 mM glucose (G0), 20 mM glucose (G20), 0 mM glucose + 200 μM H2O2 (H0) and 20 mM glucose + 200 μM H2O2 (H20). The incubations were carried out in a 95% O2 atmosphere that produced 140 μM O2 in KHBS solution, and then switched to a 100% N2 atmosphere that reduced the O2 concentration in the solution to 90 μM. The islets were subsequently incubated for another 3 h under hypoxia (hypox). The NRR of each sample was calculated as the change in NAD(P)H that caused a reduction in MTS/PMS over time. The values are the means ± SEM of 8 experiments. (E) NAD(P)H content of entire islets incubated for 1 h in KHBS containing 2.8 or 16.7 mM glucose. The values are the mean ± S.E.M. of 10 experiments. Different letters indicate P b 0.05.
use of the PPP. This metabolic difference increases the activity of PPP in response to high glucose and H2O2: in islets cultured with 5.6 mM glucose, H2O2 increased 3× the use of PPP, whereas in islets cultured with high glucose (20 mM), the use of PPP is already 2× high and was not altered (Fig. 1B). Fetal and neonatal islets have a deficient glycolysis when compared to mature islets, but normal mitochondrial processing of metabolites (Weinhaus et al., 1995). The increased activity of PPP can feed with NADPH the antioxidant and reductive biosynthesis pathways, altering the overall reducing status of the cell and mitochondria. The increased mal/asp shuttle activity also suggests that the islets cultivated with H2O2 (groups P5.6 and P20) have an increased provision of fuel for ATP production (Fig. 3C), what is a different condition of islets cultivated with high glucose (groups G20 and P20) (Fig. 3B). The pyr/mal shuttle is highly active in rat islets and exports malate from mitochondria, thereby removing reduced equivalents from the
mitochondrial matrix when β-cells are stimulated by glucose. Hence, this shuttle does not contribute directly to insulin secretion. The mal/asp and glycerol-phosphate shuttles insert NADH into the mitochondrial matrix or directly at complex II of the respiratory chain, which are inhibited by rotenone and antimicyn A, respectively (MacDonald et al., 2005a,b). The mal/asp shuttle appeared to be critical for glucose oxidation and GSIS in the control islets (G5.6), however (Fig. 4). In other groups, the increased expression of mMDH in islets cultured with H2O2 paralleled its requirement for GSIS, but not for glucose oxidation. Such apparent paradox reinforces the hypothesis that GSIS of neonatal islets is triggered by cytoplasmic NAD(P)H. This conclusion was particularly demonstrated by the fact that hypoxia provoked only a marginal effect on the NRR of control islets (G5.6), which was the less glucose-responsive (Fig. 6A). Islets were selected throughout the culture period, with necrosis and apoptosis leading to the destruction of those islets/cells unable to
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Fig. 7. NAD(P)+ reduction from endogenous fuels in the cytoplasm (endo.cyto), from endogenous fuels in mitochondria (endo.mito), from glucose in the cytoplasm (gluc.cyto) or from glucose-derived fuels in mitochondria (gluc.mito). The calculations were made as described in Materials and methods using values from Fig. 6 for islets incubated in the absence (A) or presence (B) of 200 μM H2O2. The total NRRs shown in the upper panel are the sum of the four fractions of each group. Statistical analyses were done only between islet culture groups. Different letters indicate P b 0.05.
counteract the damage imposed by H2O2 and/or high glucose concentrations (Stoppiglia et al., 2002). Although a higher NAD(P)H generation is expected in H2O2-selected islets (P5.6 and P20 groups), the NRRs of islets cultured with H2O2 were not higher than in other groups. Conversely, the stimulation of these islets with 20 mM glucose seems to increase a previously high mitochondrial NRR generated from endogenous substrates (Fig. 6B,D). This NRR does not support a high NAD(P)H content in group P5.6 islets, however (Fig. 6E), possibly due to the use of NAD(P)H equivalents to sustain repair mechanisms such as thiol reduction (Brennan et al., 2003). In H2O2-selected islets, it is agreeable that a high baseline mitochondrial NRR should protect from the imposed oxidative damage. A physiological importance of the mal/asp shuttle was suggested by the exposure of adult islets to the complex I inhibitor rotenone, what doubled the formation of non-superoxide radicals from glyceraldehyde. Exposure of the islets to the complex II inhibitor myxothiazol had no effect (Takahashi et al., 2004), indicating that mature islets mitochondria preferentially use electrons inserted through complex I instead of complex II. In the neonatal islets cultured with 20 mM glucose (G20 and P20 groups), the expression of mGPDH was increased 2–3 fold compared to their respective controls G5.6 and P5.6 (Fig. 3B). The glucose oxidation and insulin release of these “glucose-selected islets” became less dependent of the mal/asp shuttle (Fig. 4B), thus suggesting a higher use of the glycerolphosphate shuttle in islets cultured in high glucose media. The activity of the glycerol-phosphate shuttle has been associated with GSIS as a modulator of the mitochondrial activity, and a deficiency in this shuttle has been observed in models of type 2 diabetes (Giroix et al., 1992). All islets cultured with 20 mM glucose showed similar NRR from endogenous substrates (Fig. 7A) and an increased NAD(P)H
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content (Fig. 6E). Part of this higher NAD(P)H generation is thought to depend on an increased activity of the PPP (Fig. 1B). Both the PPP and the glycerol-phosphate shuttle, however, are known to participate in lipogenic pathways, the first providing reduced equivalents and the former generating glycerol from glucolytic derivatives. It is very plausible, too, that the so called mitochondrial endogenous fuels protecting mitochondria against H2O2 are in someway lipids. According to some works, the acyl-CoA synthesis correlates with alterations of the activity of the pyr/mal shuttle and is necessary for GSIS (Boucher et al., 2004; Remedi et al., 2006). On the other hand, the maximal NRR observed in islets stimulated with 20 mM glucose clearly did not match the GSIS (Fig. 1A), although it parallels glucose oxidation rate (Fig. 1B). This supports the idea of NADH shuttles limiting the carbon flux into the TCAC of neonatal islets (Fig. 7A, upper table and Fig. 4A). In acute exposition to H2O2, we observed that H2O2 targets preferentially the glucose-derived NRR, even abolishing it in islets of the G20 group. Islets from the group P20 (cultured with H2O2) did not show such increased NRR from glucose, although it kept GSIS and glucose oxidation rate responsive to glucose (Fig. 7B). Considering the highly glucose-sensitive processing of glucose derivatives in mitochondria of mature islets (Remedi et al., 2006), it is possible that these islets develop a highly glucose-sensitive mitochondrial NRR that is easily damaged by H2O2. Islets selected by cultured with H2O2 showed an increased cytoplasmic production of NAD(P)H from glucose and a significant mitochondrial processing of endogenous fuels, these islets being resistant to hydrogen peroxide. Since the chronic exposure of islets to high glucose concentrations may generate oxidative stress (see Fig. 1B, G20), the activation of PPP, cytoplasmic glucose-dependent NRR and glycerol-phosphate NADH shuttle appear to be characteristic features of islets that resist H2O2 (Figs. 1B and 7B) and can be assigned as suitable mechanisms for preserving the glucose-responsiveness of pancreatic islets. Finally, our results suggest that immature islets have the plasticity to modify the site and source of NAD(P)+ reduction, thereby changing the glucose oxidation rate and the coupling to ATP production. Islets cultured at low glucose concentrations are unable to maintain high levels of NAD(P)H. When challenged with H2O2, these islets increase the mal/asp shuttle activity and generate ATP from reduced equivalents in the cytoplasm. In contrast, islets cultured at high glucose concentrations generate ATP from cytoplasmic processing of glucose and aerobic oxidation of endogenous fuels, possibly through the glycerol-phosphate shuttle, with a consequent increase in GSIS. Contrary to glucotoxicity in adult islets, the keeping of a higher mitochondrial generation of reduced equivalents appears to protect neonatal islets against the deleterious effects of H2O2. Acknowledgments This work was partially supported by the Brazilian foundations: CAPES, CNPQ and FAPESP. We are grateful to Dr. Nicola Conran for editing the English. References Anello, M., Ucciardello, V., Piro, S., Patane, G., Frittitta, L., Calabrese, V., Giuffrida, S.A.M., Vigneri, R., Purrello, F., Rabuazzo, A.M., 2001. Chronic exposure to high leucine impairs glucose-induced insulin release by lowering the ATP-to-ADP ratio. American Journal of Physiology — Endocrinology and Metabolism 281, 1082–1087. Asanuma, N., Aizawa, T., Sato, Y., Schermerhorn, T., Komatsu, M., Sharp, G.W.G., Hashizume, K., 1997. Two signaling pathways, from the upper glycolytic flux and from the mitochondria, converge to potentiate insulin release. Endocrinology 138, 751–755. Boucher, A., Lu, D., Burgess, S.C., Potts, S.T., Jensen, M.V., Mulder, H., Wang, M.Y., Unger, R.H., Sherry, A.D., Newgard, C.B., 2004. Biochemical mechanism of lipid-induced impairment of glucose-stimulated insulin secretion and reversal with a malate analogue. Journal of Biological Chemistry 279, 27263–27271. Brennan, L., Corless, M., Hewage, C., Malthouse, J.P.G., McClenaghan, N.H., Flatt, P.R., Newsholme, P., 2003. 13C NMR analysis reveals a link between L-glutamine
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