Peroxisome proliferator-activated receptor a–retinoid X receptor agonists induce beta-cell protection against palmitate toxicity Karine Hellemans1, Karen Kerckhofs1, Jean-Claude Hannaert1, Geert Martens1, Paul Van Veldhoven2 and Daniel Pipeleers1 1 Diabetes Research Center (DRC), Brussels Free University-VUB, Belgium 2 Afdeling Farmakologie, Departement Moleculaire Celbiologie, K. U. Leuven, Belgium
Keywords cytotoxicity; fibrate; free fatty acid; pancreatic beta-cells; peroxisome proliferator-activated receptor a Correspondence K. Hellemans, Diabetes Research Center, Brussels Free University-VUB, Laarbeeklaan 103, 1090 Brussels, Belgium Fax: +32 2 4774545 Tel: +32 2 4774541 E-mail:
[email protected] (Received 11 July 2007, revised 1 October 2007, accepted 8 October 2007) doi:10.1111/j.1742-4658.2007.06131.x
Fatty acids can stimulate the secretory activity of insulin-producing betacells. At elevated concentrations, they can also be toxic to isolated betacells. This toxicity varies inversely with the cellular ability to accumulate neutral lipids in the cytoplasm. To further examine whether cytoprotection can be achieved by decreasing cytoplasmic levels of free acyl moieties, we investigated whether palmitate toxicity is also lowered by stimulating its b-oxidation. Lower rates of palmitate-induced beta-cell death were measured in the presence of l-carnitine as well as after addition of peroxisome proliferator-activated receptor a (PPARa) agonists, conditions leading to increased palmitate oxidation. In contrast, inhibition of mitochondrial b-oxidation by etomoxir increased palmitate toxicity. A combination of PPARa and retinoid X receptor (RXR) agonists acted synergistically and led to complete protection; this was associated with enhanced expression levels of genes involved in mitochondrial and peroxisomal b-oxidation, lipid metabolism, and peroxisome proliferation. PPARa–RXR protection was abolished by the carnitine palmitoyl transferase 1 inhibitor etomoxir. These observations indicate that PPARa and RXR regulate beta-cell susceptibility to long-chain fatty acid toxicity by increasing the rates of b-oxidation and by involving peroxisomes in fatty acid metabolism.
Under normal circumstances, long-chain fatty acids serve as regulators of beta-cell function [1,2]. At sustained, elevated concentrations, they can exert cytotoxic actions on beta-cells, and this has led to the view that they could be, at least in part, responsible for the loss of beta-cells in diabetes [3,4]. Several in vitro and in vivo studies have supported this lipotoxicity concept and extended into experiments aimed at pharmacologic prevention of this process [3,5]. We previously reported that pancreatic beta-cells possess defense mechanisms against oxidative damage [6] and could be induced to
provide cytoprotection [7]. It is still uncertain whether they also exhibit such properties when exposed to cytotoxic fatty acid concentrations, and if so, whether they can be activated through similar or other mechanisms. Indirect evidence for the presence of such a protective mechanism comes from the observation that fatty acid-induced toxicity was limited to a subpopulation of beta-cells, and apparently related to the cellular ability to accumulate neutral lipids in the cytoplasm [7]. We thus proposed that the formation of cytoplasmatic lipids could reduce fatty acid-induced toxicity by
Abbreviations CPT1, carnitine palmitoyl transferase 1, liver; GPAT, glycerol-3-phosphate acyltransferase, mitochondrial; Pex2, peroxisomal biogenesis factor 2; Pex3, peroxisomal biogenesis factor 3; Pex11a, peroxisomal biogenesis factor 11a; Pex14, peroxisomal biogenesis factor 14; Pex16, peroxisomal biogenesis factor 16; PPARa, peroxisome proliferator-activated receptor a; PMP70, peroxisomal membrane protein 70; RA, retinoic acid; RXRa, retinoid X receptor; SCD1, stearoyl-CoA desaturase 1; SCD2, stearoyl-CoA desaturase 2.
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Results Specificity of palmitate toxicity When rat beta-cells were cultured with palmitate, timeand concentration-dependent cytotoxicity was measured. At 50 and 100 lm, no toxic effect was detected after 2 days, and only 10–20% cells were damaged after 8 days (data not shown). At 250 and 500 lm, cytotoxicity was noticed after 2 days and resulted, after 8 days, in, respectively, 38 ± 2% and 75 ± 4% dead cells (Fig. 1). It was noticed that with exposure longer than 3 days at 500 lm, the slope of the toxicity curve declined. The percentage of surviving cells tended to stabilize around 25% after 6 days, despite administration of a new bolus of fatty acid every 48 h. With 250 lm, the fraction of cell survival stabilized around 60%. The cytotoxic effect of palmitate did not vary with the glucose concentration in the medium (comparison of 5, 10 and 20 mm glucose, data not shown). Subsequent studies were conducted at 10 mm glucose for 2 days with 500 lm palmitate (acute toxicity) and for 8 days at 250 lm (chronic toxicity). Islet endocrine nonbeta-cells exhibited lower susceptibility to palmitate toxicity: cytotoxicity was 9 ± 0.5% after 2 days at 500 lm, and 16 ± 3% after
100 Cytotoxicity (%)
preventing a rise in toxic free acyl moieties [7] and ⁄ or fatty acid metabolites such as ceramides [8,9]. Along this line, one can further hypothesize that an increased rate of fatty acid oxidation also lowers the formation of these cytotoxic mediators and could thus also act as a cytoprotective mechanism. To test this hypothesis, we examined whether palmitate toxicity can be reduced by increasing its oxidation rates. We first assessed whether protection could be conferred by l-carnitine, a rate-limiting component for long-chain fatty acid transport into the mitochondria, or suppressed by etomoxir, an irreversible carnitine palmitoyl transferase 1 (CPT1) inhibitor [10]. In a second set of experiments, we assessed the effects of agonists for peroxisome proliferator-activated receptor a (PPARa) and retinoid X receptor (RXR). PPARa–RXR dimers can be activated by both PPARa and RXR agonists. PPARa–RXR dimers typically regulate the expression of multiple genes involved in mitochondrial and peroxisomal b-oxidation as well as lipoprotein metabolism [11]. PPARa is expressed in primary rat beta-cells [12], and has been shown to activate fatty acid oxidation [13,14]. PPARa agonists have been reported to prevent fatty acid-induced beta-cell dysfunction and apoptosis in human islets [15], and improve beta-cell function in insulin-resistant rodent models [16].
Beta-cell protection against palmitate toxicity
75
250 M C16:0 500 M C16:0
50 25 0
2
4 6 8 Days of exposure
10
Fig. 1. Time course analysis for palmitate cytotoxicity. Primary rat beta-cells were exposed to 250 or 500 lM palmitate. Cytotoxicity was measured on a daily basis (n ¼ 4, vertical bars represent SEM).
8 days at 250 lm. After 8 days at 500 lm, the cytotoxicity increased to 46 ± 8% (results not shown). In the latter condition, more than 95% of purified beta-cells died, which means that approximately 20% of the dead cells in the nonbeta-cell fraction correspond to beta-cells, as the nonbeta-cell fraction is contaminated to this extent by beta-cells; consequently, the palmitate toxicity for the islet nonbeta-cells is calculated to be about 25% after 8 days at 500 lm. Palmitate toxicity in beta-cells was compared with that of equimolar concentrations of other fatty acids in the presence of 1% BSA (Table 1). Oleate, an unsaturated long-chain fatty acid, was less toxic, and the shorter-chain molecules butyrate (C4), hexanoate (C6) and octanoate (C8) were only marginally toxic. The 2-methyl and 3-methyl derivatives of palmitate were virtually nontoxic: < 5% after 2 days and < 10% after 8 days, both at 250 and at 500 lm, whereas Table 1. Cytotoxicity of fatty acids for rat beta-cells. Beta-cells were exposed to the following fatty acids at 500 lM for 2 days or at 250 lM for 8 days: palmitate (C16:0), oleate (C18:1), butyrate (C4:0), hexanoate (C6:0), octanoate (C8:0), 2-methylhexadecanoic acid (2-Me-C16:0), 3-methylhexadecanoic acid (3-Me-C16:0), 2-bromopalmitate (Br-C16:0). Data represent means ± SD (n ¼ 3–6). *P < 0.001 as compared to C16:0 (student’s t-test). Percentage cytotoxicity Fatty acid
500 lM for 2 days
250 lM for 8 days
C16:0 C18:1 C4:0 C6:0 C8:0 2-Me-C16:0 3-Me-C16:0 Br-C16:0
24 11 5 1 9 4 5 58
38 28 4 5 11 4 6 92
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± ± ± ± ± ± ± ±
2 2* 2* 1* 1* 1* 4* 6*
± ± ± ± ± ± ± ±
2 3* 1* 1* 1* 4* 4* 3*
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Table 2. Metabolism of [14C]palmitate and [14C]glucose in beta-cells and islet nonbeta-cells at 3.3 and 20 mM glucose. Palmitate and glucose metabolism was measured in freshly isolated beta-cells and islet nonbeta-cells incubated for 2 h with 50 lM [14C]palmitate or 5 lCi of 14 14 D-[U- C]glucose. C incorporation was measured in CO2 and small metabolic intermediates, as well as in the lipid and protein fractions and expressed as (pmol per 2 h per 103 cells). Data are indicated as mean ± SD (n ¼ 5). Italic: P < 0.05 for 20 mM glucose as compared to 3.3 mM glucose. *P < 0.05 for nonbeta-cells as compared to beta-cells, **P < 0.001 for nonbeta-cells as compared to beta-cells (Student’s t-test). 14
C recovery as:
CO2 From [14C]palmitate Beta-cells Glucose 3.3 mM Glucose 20 mM Nonbeta-cells Glucose 3.3 mM Glucose 20 mM From [14C]glucose Beta-cells Glucose 3.3 mM Glucose 20 mM Nonbeta-cells Glucose 3.3 mM Glucose 20 mM
Intermediates
Protein
0.52 ± 0.04 0.10 ± 0.01
0.62 ± 0.05 0.06 ± 0.01
1.71 ± 0.11 1.73 ± 0.04
0.21 ± 0.06 0.13 ± 0.08
0.63 ± 0.04* 0.46 ± 0.04*
0.18 ± 0.003* 0.10 ± 0.01
1.18 ± 0.04* 1.06 ± 0.07*
0.13 ± 0.08 0.07 ± 0.03
0.29 ± 0.08 2.16 ± 0.72
0.40 ± 0.02 3.16 ± 0.99
0.13 ± 0.03* 1.13 ± 0.55
0.19 ± 0.04* 0.45 ± 0.07**
5.49 ± 0.31 31.15 ± 1.86
4.41 ± 0.17 19.90 ± 1.96
0.96 ± 0.22** 3.82 ± 0.95**
1.20 ± 0.11** 3.78 ± 0.48**
2-bromopalmitate was markedly more toxic than palmitate. There was no difference in the percentage of living cells following vehicle treatment when compared to the standard control. Palmitate and glucose metabolism by beta-cells and islet nonbeta-cells Palmitate and glucose metabolism was measured in freshly isolated beta-cells and in islet nonbeta-cells that were incubated for 2 h with 50 lm [14C]palmitate or 5 lCi of d-[U-14C]glucose. 14C incorporation was measured in CO2 and small metabolic intermediates, as well as in the lipid and protein fractions (Table 2). After incubation with [14C]palmitate at 3.3 mm glucose, the largest amount was recovered in the lipid-soluble fraction (55%) and the lowest in the protein fraction, whereas comparable amounts were converted to CO2 (17%) and to small metabolic intermediates (20%); at 20 mm glucose, the total amounts in the lipid fraction (representing 82%) and protein fraction were not significantly different from those at 3 mm glucose, but the production of 14CO2 and of 14C intermediates was, respectively, five- and 10-fold lower (Table 2). Analysis of the fate of [14C]glucose indicated that, at 3.3 mm glucose, the tracer was converted to CO2 and to small intermediates, and that high glucose increased this rate five-fold, and also increased seven-fold the 14C incorporation into the lipid and protein fraction. Islet endocrine nonbeta-cells exhibit much lower rates of glucose oxidation and utilization than islet 6096
Lipid
beta-cells; the values shown in Table 2 are an overestimation, in view of the contamination of this fraction by 20–25% beta-cells; the CO2 production from glucose that is calculated for islet nonbeta-cells is thus less than 10% of that in beta-cells. On the other hand, their level of CO2 production from palmitate is higher, in particular at 20 mm glucose, where four-fold higher rates were measured than in beta-cells (Table 2). In contrast to the situation in beta-cells, 30% of the labeled palmitate was converted to CO2 independently of the glucose concentration. Effect of regulators of palmitate metabolism on palmitate toxicity in beta-cells When beta-cells were precultured for 24 h with the CPT1 activator l-carnitine before measurement of their palmitate oxidation during 2 h of incubation in the further presence of l-carnitine, 14CO2 formation was six-fold higher (to 0.83 ± 0.14 pmol per 2 h per 103 cells, n ¼ 4, P < 0.05) than in control cells cultured and incubated with the solvent (0.12 ± 0.03). This stimulatory effect was preserved when 250 lm palmitate was added to the preculture medium (0.78 ± 0.19 pmol per 2 h per 103 cells, n ¼ 4, P < 0.01) (Fig. 2A). It was associated with an eight-fold elevation of 14C incorporation into metabolic intermediates (from 0.15 ± 0.05 to 0.95 ± 0.07 pmol per 2 h per 103 cells, P < 0.05) (results not shown). Preculture with 1 mm l-carnitine protected beta-cells from palmitate toxicity during a
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**
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Effect of PPARa–RXR agonists on palmitate cytotoxicity
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Palmitate oxidation (pmol/Kc/2h)
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Beta-cell protection against palmitate toxicity
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** ***
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500
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250 C4:0
M
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Fig. 2. Effect of regulators of fatty acid metabolism on fatty acid toxicity. (A) Effect of L-carnitine on palmitate oxidation. Beta-cells were precultured for 24 h with the CPT1 activator L-carnitine (1 mM) in the absence or presence of 250 lM palmitate before measurement of [14C]palmitate oxidation during a 2 h incubation in the further presence of L-carnitine. (B) Effect of L-carnitine on palmitate cytotoxicity. Cells were exposed for 2 or 8 days to 500 or 250 lM palmitate in the presence of L-carnitine following 24 h of preculture with 1 mM L-carnitine. (C) Effect of a CPT1 inhibitor on fatty acid toxicity. Cells were exposed for 2 or 8 days to 500 or 250 lM butyrate or palmitate alone, or in combination with 200 lM etomoxir. Data are presented as mean ± SEM (n > 4); **P < 0.01, ***P < 0.001 (Student’s t-test).
subsequent exposure to 500 lm palmitate for 2 days or 250 lm for 8 days by, respectively, 70% and 40% (Fig. 2B). On the other hand, when beta-cells were exposed to palmitate in the presence of the CPT1 inhibitor etomoxir (200 lm) (Fig. 2C), their survival was further decreased. Lower concentrations of etomoxir (1, 5 and 50 lm), which are known to stimulate PPARa, failed to show an effect on palmitate toxicity. Etomoxir (200 lm) did not affect beta-cell survival in the presence of butyrate (C4), which is known to enter mitochondria independently of CPT1. Addition of l-cycloserine (100 lm), an inhibitor of serine palmitoyl transferase, was also found to reduce palmitate toxicity, both after 2 days at 500 lm (from 24 ± 2% to 10 ± 1%), and after 8 days at 250 lm (from 38 ± 2% to 20 ± 4%), but not that of oleate (results not shown).
The effects of PPARa–RXR agonists clofibrate, ciprofibrate and 9-cis-retinoic acid (9-cis-RA) were examined by adding these compounds to the 2 and 8 day culture media. Clofibrate alone (tested at 100, 250 and 500 lm, two-way ANOVA, P < 0.001) reduced palmitate toxicity at both time points, with a maximal effect at 250 lm (Fig. 3A). Protection from palmitate toxicity was also observed after treatment with 9-cis-RA alone at all tested concentrations (0.5, 2 or 5 lm, two-way ANOVA, P < 0.001); at 5 lm, the effect was comparable to that of 250 lm clofibrate, namely 60% protection after 2 days at 500 lm and 70% after 8 days at 250 lm (Fig. 3A). Combinations of different concentrations of both agents further reduced palmitate toxicity; the maximal level of protection, reducing palmitate-induced cell death to only 5%, was reached using a combination of 5 lm 9-cis-RA and 100 lm clofibrate. Higher clofibrate concentrations in the presence of 9-cis-RA did not lower palmitate toxicity any further (Fig. 3A). For all further experiments, a combination of 250 lm clofibrate and 2 lm 9-cis-RA was used, resulting in a reduction of palmitate toxicity by 90% (Fig. 3B). The efficacy of this combination did not significantly differ from that of 250 lm clofibrate plus 5 lm 9-cis-RA. The same level of protection as found in the presence of 10 mm glucose was found at low (5 mm) and high (20 mm) glucose concentrations (results not shown). Under the same conditions, clofibrate and 9-cis-RA were also found to protect against oleate toxicity (from 25 ± 3% to 13 ± 2% for 500 lm after 2 days, and from 32 ± 3% to 14 ± 4% for 250 lm after 8 days, P < 0.01, results not shown). Ciprofibrate (10, 50 and 100 lm) mimicked the effect of clofibrate, with a maximal effect at 100 lm, and similar additive protection by 9-cis-RA (Fig. 3B). Addition of fibrate and ⁄ or 9-cis-RA to palmitate-free control medium did not influence cell survival during culture (data not shown). When endocrine nonbeta-cells were exposed to 500 lm palmitate for 8 days with or without 250 lm clofibrate plus 2 lm 9-cis-RA, no differences in toxicity were noticed, indicating the absence of a protective effect of the supplement (results not shown). Effect of PPARa–RXR agonists on palmitate metabolism Preculture (24 h) of beta-cells with 250 lm clofibrate plus 2 lm 9-cis-RA increased palmitate oxidation during a subsequent 2 h incubation with 50 lm
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#$
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10 0
500 0
250 100 clofibrate (
M
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500 M)
Fig. 3. Effect of PPARa and RXR agonists on palmitate toxicity. (A) Effect of clofibrate and 9-cis-RA on palmitate toxicity. Primary rat betacells were exposed to 500 lM palmitate for 2 days, or to 250 lM palmitate for 8 days, in the presence or absence of clofibrate (100, 250, 500 lM) and ⁄ or 9-cis-RA (0.5, 2, 5 lM). (n ¼ 4–8). Vertical bars represent SEM. (B) Comparison between the protective effect of clofibrate (250 lM) and of ciprofibrate (100 lM) against palmitate toxicity, alone or in combination with 2 lM 9-cis-RA. Data are indicated as mean ± SEM (n > 5). #P < 0.001 as compared to palmitate; $P < 0.001 as compared to single agonist treatment (clofibrate, ciprofibrate or 9-cis-RA) (two-way ANOVA). Table 3. Effect of palmitate and PPARa–RXR agonists on [14C]palmitate metabolism. Beta-cells were precultured for 24 h in the presence or absence of 250 lM palmitate and ⁄ or 250 lM clofibrate (Clof) plus 2 lM 9-cis-RA (RA). Incorporation of the 14C label into CO2, lipid intermediates, lipids and proteins was measured during a 2 h incubation with 50 lM [14C]palmitate, and expressed as a percentage of the control condition (vehicle only, 10 mM glucose). *P < 0.001 as compared to control, **P < 0.001 as compared to cytochrome P250, Student’s t-test (n ¼ 5). 14
C-recovery as
Treatment
CO2
Intermediates
Lipid
Protein
C16:0 C16:0 + Clof ⁄ RA Clof ⁄ RA
59 ± 13* 94 ± 12** 181 ± 27*
80 ± 18 78 ± 10 94 ± 27
101 ± 9 94 ± 7 108 ± 9
93 ± 12 77 ± 27 83 ± 24
[14C]palmitate by 80% (P < 0.001) (Table 3). This effect was also seen when palmitate (250 lm) was present during preculture (Table 3). The lower 14CO2 values measured in this condition when compared with the control reflect isotopic dilution in palmitate-pretreated cells. No differences were found for the incorporation of label in lipids, intermediates or proteins. Effect of PPARa–RXR agonists on gene expression of enzymes involved in fatty acid metabolism and peroxisomal membrane proteins RT-PCR analysis was performed to investigate whether the agonist combination (250 lm clofibrate 6098
plus 2 lm 9-cis-RA) increased expression of genes coding for enzymes involved in peroxisomal or mitochondrial lipid metabolism or of peroxisomal membrane proteins (Table 4). Palmitate 250 lm alone induced the mRNA expression of CPT1 (two-fold increase over control, P < 0.05), and decreased that of PPARa, stearoyl-CoA desaturase 1 (SCD1), stearoyl-CoA desaturase 2 (SCD2), and the peroxisomal membrane proteins peroxisomal biogenesis factor 2 (Pex2) and peroxisomal biogenesis factor 14 (Pex14) (P < 0.05). Addition of clofibrate plus 9-cis-RA further increased mRNA levels of CPT1 (1.7-fold, P < 0.001) and induced the expression of the mitochondrial enzymes acyl-CoA dehydrogenase (medium chain), acyl-CoA dehydrogenase (long chain), and mitochondrial acetylCoA acetyltransferase 2 (P < 0.01), as well as of the peroxisomal enzymes peroxisomal acetyl-CoA acetyltransferase 1, palmitoyl-CoA oxidase 1, and pristanoyl-CoA oxidase (P < 0.01), and the prolipogenic endoplasmatic reticulum enzymes glycerol-3-phosphate acyltransferase, mitochondrial (GPAT), SCD1 and SCD2 (P < 0.01). This combination was also found to increase the mRNA levels of peroxisomal biogenesis factor 3 (Pex3), peroxisomal biogenesis factor 16 (Pex16), and peroxisomal biogenesis factor 11a (Pex11a), as well as of Pex2, Pex14 and peroxisomal membrane protein 70 (PMP70). Comparable changes in expression were also found after stimulation with clofibrate and 9-cis-RA in the absence of palmitate.
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Beta-cell protection against palmitate toxicity
Table 4. Effect of palmitate and PPARa–RXR agonists on mRNA expression levels. Beta-cells were exposed for 2 days to 250 lM palmitate and ⁄ or 250 lM clofibrate plus 2 lM 9-cis-RA or vehicle (control). qPCR values were normalized to actin and calculated as DDCt values relative to the indicated control conditions. Unpaired student t-test, two-tailed, mean ± SD, n ¼ 4–6, *P < 0.05, **P < 0.01, ***P < 0.001.
Protein Gene transcription Peroxisome proliferator-activated receptor a Mitochondrial b-oxidation Carnitine palmitoyl transferase 1 Acyl-CoA dehydrogenase, long chain Acyl-CoA dehydrogenase, medium chain Acyl-CoA dehydrogenase, short chain Mitochondrial acetyl-CoA acetyltransferase 2 Peroxisomal fatty acid oxidation Peroxisomal acetyl-CoA acetyltransferase 1 Palmitoyl-CoA oxidase 1 Pristanoyl-CoA oxidase a-Methylacyl-CoA racemase Lipid synthesis Stearoyl-CoA desaturase 1 Stearoyl-CoA desaturase 2 Glycerol-3-phosphate acyltransferase Peroxisomal membrane proteins Peroxisomal biogenesis factor 3 Peroxisomal biogenesis factor 16 Peroxisomal biogenesis factor 11a Peroxisomal membrane protein 70 Peroxisomal biogenesis factor 14 Peroxisomal biogenesis factor 2
C16:0, compared to control
Clofibrate +9-cis-RA, compared to control
C16:0 + clofibrate + 9-cis-RA, compared to C16:0
0.7 ± 0.2
1.6 ± 0.2***
1.9 ± 0.4 ***
1.9 1.4 0.8 1.0 1.3
± ± ± ± ±
0.6** 0.7 0.2 0.1 0.4
2.6 1.3 1.3 1.8 2.5
± ± ± ± ±
0.4*** 0.2 0.3 0.6* 1.0**
1.9 1.5 1.6 1.1 1.6
± ± ± ± ±
0.2*** 0.2** 0.3** 0.2 0.1***
0.8 1.0 1.4 1.1
± ± ± ±
0.2 0.3 0.9 0.2
0.9 1.6 2.0 0.9
± ± ± ±
0.5 0.2*** 0.8* 0.1
1.6 1.6 1.5 1.2
± ± ± ±
0.1*** 0.4** 0.2*** 0.2
0.5 ± 0.2*** 0.7 ± 0.1*** 1.1 ± 0.3
4.5 ± 2.5** 1.8 ± 0.3*** 1.5 ± 0.2**
5.8 ± 2.7*** 2.6 ± 0.6*** 1.5 ± 0.2**
1.2 0.9 1.0 0.9 0.7 0.7
1.2 1.7 1.3 1.4 2.6 1.4
1.5 1.4 1.6 2.0 1.4 1.4
± ± ± ± ± ±
0.5 0.1 0.2 0.3 0.1** 0.2*
Effect of etomoxir on palmitate toxicity in the presence of PPARa–RXR agonists
Discussion This study confirms that sustained exposure to palmitate causes time- and dose-dependent toxicity on rat beta-cells. Over an 8 day culture period, palmitate, at 250 or 500 lm, progressively reduced the number of surviving cells, involving both necrotic and apoptotic pathways [7], but a significant fraction remained resistant. These survival curves show that primary betacells can be less susceptible or even resistant to fatty
0.6 0.5** 0.2* 0.3** 1.0* 0.5
± ± ± ± ± ±
0.3* 0.2** 0.3** 0.6** 0.2** 0.2*
C16:0 C16:0 + Clofibrate / 9-cis RA C16:0 + Clof / 9RA + etomoxir
30 % Cytotoxicity
In the presence of etomoxir, the protective effect of clofibrate and 9-cis-RA was abolished, and the cytotoxicity of palmitate 500 lm for 2 days increased four-fold, i.e. from 4 ± 3% to 17 ± 5%. (Fig. 4). This toxicity was also increased when the cells were precultured for 24 h with 200 lm etomoxir prior to their incubation with the palmitate ⁄ clofibrate ⁄ RA mixture for 2 days (19 ± 3%, P < 0.05 versus 5 ± 2%). The effect of preculture with etomoxir was lost after prolonged subsequent culture in its absence (250 lm palmitate for 8 days).
40
± ± ± ± ± ±
$ 20
$
*** 10
*** ***
0 500
M
- 2d
250
M
- 8d
Fig. 4. Effect of etomoxir on PPARa–RXR protection against palmitate toxicity. Primary beta-cells were cultured with 500 lM palmitate for 2 days, or 250 lM palmitate for 8 days, in the absence or presence of 250 lM clofibrate plus 2 lM 9-cis-RA, or in combination with 200 lM etomoxir. Data are presented as mean ± SEM (n ¼ 4). ***P < 0.001 as compared to palmitate; $P < 0.01 for etomoxir as compared to clofibrate plus 9-cis-RA (Student’s t-test).
acid toxicity, and that this property is heterogeneously expressed, like other cell functions [17,18]. We previously reported that addition of oleate increases the
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resistance of rat beta-cells to palmitate-induced cell death [7]. This oleate effect was also observed in human beta-cells [19] and in other cell types [20–23]. It appeared to be correlated with the formation of triglycerides, supporting the view that fatty acid incorporation into neutral lipids prevents the accumulation of toxic free palmitoyl acyl moieties [22], and ⁄ or ceramide derivatives [24,25]. A role of ceramides in palmitate toxicity is supported by the observed protection by l-cycloserine, an inhibitor of serine palmitoyl transferase and thus of ceramide synthesis. At variance with other cytotoxic conditions [26], no protective effect could be attributed to glucose, as similar palmitate toxicities were measured following culture at 5 or 10 mm glucose. The percentage of dead cells was not increased when palmitate exposure was assessed at excessive glucose levels (20 mm), which contrasts with observations in beta-cell lines [27–30]. The latter discrepancy might be related to differences in experimental protocols, such as free fatty acid concentrations, free fatty acid ⁄ BSA ratios [7], or the use of serum, but could also result from differences between primary beta-cells and cell lines. Glucose cytotoxicity has also been seen in cell lines [27,31], whereas increased glucose exerts cytoprotective effects in primary beta-cells, at least in conditions that cause an oxidative shift in their metabolic state [32–35]. Our data suggest that glucose-induced changes in the cellular metabolic redox state do not alter cellular susceptibility to palmitate toxicity. At a nontoxic palmitate concentration (50 lm), glucose suppresses its oxidation, probably due to dynamic regulation of the malonyl-CoA ⁄ CPT1 axis [3], but this seems not to be accompanied by a toxic effect; in fact, this may lead to the formation of fatty acid derivatives with physiologic action in the presence of glucose [36– 38], or in protective accumulation in the form of neutral lipids [7,39]. Furthermore, palmitate induced the expression of CPT1 two-fold, suggesting that beta-cells have the inherent capacity to adapt their oxidation rate to elevated fatty acid levels, independently of the suppressive effect of glucose. Our data indicate that palmitate toxicity can be reduced by increasing its oxidation through mitochondrial and ⁄ or peroxisomal pathways. Mitochondrial oxidation of long-chain fatty acids is known to be rate limited at the level of CPT1 [10]. Viral overexpression of CPT1 has been shown to enhance palmitate oxidation in INS-1 cells and islets [40,41]. In our study, culture with l-carnitine, an essential component of CPT1, stimulated palmitate oxidation and reduced its toxicity, while etomoxir, an inhibitor of CPT1, increased the toxicity of palmitate (C16:0) but not of butyrate (C4:0), which enters mitochondria independently of 6100
CPT1. In fact, no toxicity was measured for any of the tested shorter-chain fatty acids, raising the possibility that shortening the palmitate chain represents another and perhaps more important mechanism for inducing cytoprotection. It is so far unclear to what extent peroxisomes in beta-cells contribute to fatty acid metabolism. That they could be involved is suggested by the absence of cytotoxicity for the 2-methyl and 3-methyl derivatives of palmitate. Like other branched fatty acids, these compounds are known to be preferentially transported to the peroxisomes, where 2-methyl-C16 undergoes b-oxidation and 3-methyl-C16 will be a-oxidized before being transported to the mitochondria [42]. Fibrates are known to regulate genes involved in mitochondrial, as well as peroxisomal, fatty acid oxidation and to induce peroxisome proliferation and maturation in multiple cell types [11]. Both clofibrate and ciprofibrate were found to increase palmitate breakdown and to reduce its toxicity; their combination with 9-cis-RA resulted in complete protection. This protective action of 9-cis-RA against palmitate toxicity contrasts with the proapoptotic effect observed in MIN6 cells at a 10-fold higher concentration [43]. Clofibrate plus 9-cis-RA was found to provide the same level of protection at all examined glucose concentrations. This effect correlated with induced expression of CPT1 and mitochondrial and peroxisomal b-oxidation enzymes, and resulted in normalization of palmitate oxidation. In further support of this, inhibition of CPT1 by etomoxir was found to abolish the protective action of clofibrate plus 9-cis-RA. PPARa:RXR agonists also induced expression of GPAT, SCD1 and SCD2 mRNA, which might mediate incorporation of the fatty acid into (phospho)lipids [44]. Increased SCD1 expression has been previously noticed in palmitate-resistant MIN6 cells but has not yet been directly correlated with a cytoprotective diversion of palmitate into lipid formation [39]; esterification of palmitate was shown to result in accumulation of insoluble tripalmitin and to correlate with endoplasmic reticulum stress and apoptosis [45,46]. The PPARa–RXR agonists were also found to act at a third level of potential relevance, namely the expression of proteins involved in peroxisome biogenesis (Pex3 and Pex16), proliferation (Pex11a) and maturation (PMP70, Pex2 and Pex14) [47]. By inducing the peroxisomal compartment, they might indeed increase the channeling of palmitate through the first cycles of chain shortening before further breakdown in mitochondria. This view is consistent with the virtual absence of toxicities for short-chain fatty acids. Peroxisomes might thus represent a key site for reducing the
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toxicity of palmitate in primary beta-cells. Their activity might be low in normal circumstances, as judged by the low catalase activity in islet beta-cells [48,49]. Addition of clofibrate was shown to increase catalase activity in INS-1 cells together with palmitate oxidation [50]. Clofibrate plus 9-cis-RA did not provide protection in the nonbeta-cells where, independently of glucose, a substantially higher proportion of palmitate was converted to CO2. Our data supplement previous in vivo findings or observations in cell lines, and more specifically indicate to what extent they reflect effects on the survival of primary beta-cells and, if so, through which mechanism these can then be explained. Although fatty acids, and particularly palmitate, are classically seen as mediators of lipotoxicity at the level of the beta-cells, it is often not clear whether the reported derangements are the result of beta-cell death and ⁄ or dysfunction. Consequently, the protective action of PPARa agonists with or without RXR agonists was not always well specified in these terms [51]. Adenoviral coexpression of PPARa and RXRa synergistically – and in a dose- and liganddependent manner ) potentiated glucose-stimulated insulin secretion from INS-1E cells while increasing their expression of genes involved in free fatty acid uptake and b-oxidation [52,53]. An increase of PPARadriven b-oxidation in response to topiramate was also found to protect INS-1E cells from oleate toxicity [54]. When administered in vivo, PPAR–RXR ligands induced expression of b-oxidation enzymes and stimulated palmitate oxidation in isolated islets [13]. Fibrate treatment restored the coupling between insulin secretion and action in glucose-intolerant rats on a high-fat diet [55] and prevented diabetes in obese OLETF rats [56,57]. Combination therapy with PPARa and a-agonists, or dual agonists, ameliorated insulin secretion and increased insulin stores in genetically obese diabetic db ⁄ db mice [58]. The present work has shown that PPARa–RXR agonists can protect primary beta-cells against the cytodestructive effects of palmitate. It has provided evidence that this protection is achieved by stimulating mitochondrial and peroxisomal pathways for palmitate breakdown. Further work is needed to assess the functional properties of these protected beta-cells and to evaluate the influence of the agonists at nontoxic palmitate concentrations.
Experimental procedures Materials Palmitate, oleate, butyrate, hexanoate, octanoate (sodium salts), 2-bromohexadecanoic acid, clofibrate, ciprofibrate,
Beta-cell protection against palmitate toxicity
9-cis-RA, l-carnitine and l-cycloserine were purchased from Sigma-Aldrich (Bornem, Belgium). Branched 2-methylhexadecanoic acid and 3-methylhexadecanoic acid were prepared as described previously [59,60]. Stock solutions of fatty acids (25 mm, 50 mm) were made in 90% ethanol by heating to 60 C, except for 2-bromopalmitate, which was dissolved at room temperature. Stock solutions of clofibrate (200 mm), ciprofibrate (20 mm) and 9-cis-RA (10 mm) were dissolved in absolute ethanol. Etomoxir was a gift from V. Grill (Trondheim University, Norway) and dissolved in saline. d-[U-14C]glucose (287–311 mCiÆmmol)1; 1 mCi per 5 mL) was purchased from Amersham Biosciences (Roosendaal, Belgium), and [U-14C]palmitic acid (850 mCiÆ mmol)1; 0.1 mCiÆmL)1) from Perkin Elmer Life Sciences (Zaventem, Belgium).
Preparation and culture of rat beta-cells Adult male Wistar rats were bred according to Belgian regulations on animal welfare. Experiments were carried out in accordance with the European Communities Council Directive (86/609/EEC). Pancreatic islets were isolated, dissociated and purified into single beta-cells (purity 88 ± 4% insulin-positive cells) and endocrine nonbeta-cells (70 ± 11% alpha-cells, 23 ± 3% beta-cells) by autofluorescence-activated cell sorting [61]. For studies on cytoxicity, isolated cells were cultured in polylysine-coated microtiter plates (2500–3000 cells per well) with Ham’s F10 medium containing 10 mm glucose (unless stated otherwise), 1% charcoal-extracted BSA (fraction V, radioimmunoassay grade; Sigma-Aldrich), 2 mm l-glutamine, 50 mm 3-isobutyl-1-methylxanthine, 0.075 mgÆmL)1 penicillin and 0.1 mgÆmL)1 streptomycin [7]. Test reagents were added to the culture medium, with control conditions receiving similar dilutions of solvent. After 2 and 5 days of culture, the medium was changed and fresh reagents were added. Percentages of living and dead cells were determined by vital staining using neutral red [7]. For metabolic and gene expression studies, freshly isolated cells were reaggregated and cultured in suspension as previously described [62].
Measurement of glucose and palmitate metabolism Duplicate samples of 5 · 104 rat beta-cells were incubated for 2 h at 37 C using Ham’s F10 medium containing 0.5% BSA, 2 mm l-glutamine and 10 mm Hepes for measuring glucose metabolism (5 lCi of d-[U-14C]glucose with different concentrations of unlabeled d-glucose) [63]. Palmitate metabolism was measured using KRBH medium, containing 0.2% BSA (fraction V), 2 mm calcium chloride, and 10 mm Hepes (0.5 lCi of [U-14C]palmitic acid, with unlabeled palmitate up to 50 lm in order to achieve the same ratio of free fatty acid over BSA as in the cytotoxicity experiments with 250 lm palmitate). The rate of
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d-[U-14C]glucose or [U-14C]palmitic acid oxidation was assessed through the formation of 14CO2 [63]. Cells were incubated in a siliconized tube trapped in an airtight glass vial. After 2 h, 20 lL of 1 m HCl was injected, and 250 lL of Hyamine (Packard Bioscience, Groningen, the Netherlands) added to capture 14CO2 for 1 h at room temperature. The 14C incorporation into lipids, proteins and metabolic intermediates was measured as previously described [64].
Gene expression analysis Total beta-cell RNA was extracted with TRIzol Reagent (Gibco BRL, Carlsbad, CA, USA) and its quality was assessed on a 2100 Bioanalyzer (Agilent, Waldbronn, Germany), taking a minimal cutoff RNA integrity number of 8. RNA clean-up was performed with the Turbo DNA Free Kit (Ambion, Austin, TX, USA) and cDNA prepared with the High-Capacity cDNA Archive Kit (Applied Biosystems, Foster City, CA, USA). Real-time PCR was performed using an ABI Prism Sequence Detector (Applied Biosystems). Primers were obtained from Applied Biosystems (Table 4). For each RT-PCR reaction, the cycle threshold (Ct) was determined with sds 1.9.1 software. DDCt values were calculated versus b-actin. Fold changes were calculated starting from DDCt values of a minimum of four independent experiments performed in duplicate.
Data analysis Data are presented as mean ± SEM, or as mean ± SD of n independent experiments. Statistical analysis was performed using Student’s t-test, unless stated otherwise. Differences were considered significant for P < 0.05.
Acknowledgements This work was supported by the Research Foundation Flanders (Fonds Voor Wetenschappelijk OnderzoekVlaanderen, Grant FWO-G.0357.03, Grant FWO-1.5.195.05 and PhD grant FWOTM277 to K. Kerckhofs) and by the Inter-University Poles of Attraction Program (IUAP P5 ⁄ 17) from the Belgian Science Policy. The Diabetes Research Center is a partner of the Juvenile Diabetes Research Center for Beta Cell Therapy in Diabetes.
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