1Henry Wellcome Signalling Laboratories and Department of Biochemistry, School of Medical Sciences, University of Bristol, Bristol, BS8 1TD United Kingdom; and 2GlaxoSmithKline, Department of Metabolic Diseases, Research Triangle Park, North Carolina 27709
Submitted 9 January 2004 ; accepted in final form 26 April 2004
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ABSTRACT |
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-cell; insulin; secretion; peroxisome proliferator-activated receptor-
; sterol regulatory element-binding protein-1c; thiazolidinedione; glucolipotoxicity
PPAR, a member of the nuclear hormone receptor family, is translated from an alternately spliced mRNA in humans and rodents, generating three (
1,
2, and
3) or two (
1 and
2) isoforms, respectively, in these species (see Fig. 1A) (16, 17, 45, 48). Whereas PPAR
1 is present in many mammalian tissues, the expression of PPAR
2 is confined largely to adipose tissue (48). Both PPAR
mRNA and protein have been detected in human (13) and rodent (4, 53) pancreatic islets, although the relative levels of the
1 and
2 isoforms have yet to be established in this tissue.
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Recent data have suggested that changes in PPAR activity in the islet may also influence the development of diabetes. First, of four morbidly obese individuals carrying a rare activating mutation (Pro113Glu) of PPAR
2 (38), three were diabetic with decreased circulating insulin levels. Second, activation of PPAR
1 has been shown to inhibit glucose-stimulated insulin release ex vivo both from a tumoral
-cell line (34) and from primary islets (3). Third, PPAR
expression is increased more than fivefold in islets from ZDF rats (53), suggesting that PPAR
induction may also negatively regulate insulin release in this model. In contrast to these observations, PPAR
activation has been suggested to preserve
-cell function, morphology, and mass in other rodent models of diabetes. Thus TZDs, presumably acting by binding to PPAR
(29), reduce islet TG content and promote glucose- and arginine-stimulated insulin secretion in the fat-laden islets of ZDF rats (26, 43). Furthermore, TZD treatment enhances insulin release in glucose-intolerant human subjects (7). One possible explanation for these discrepancies is that changes in
-cell function in vivo may not result from a direct effect of TZDs on the islet but may instead be a consequence of improved whole body metabolic parameters (e.g., blood glucose and circulating free fatty acid levels).
In light of such contrasting observations, the present study was designed to evaluate the effects of PPAR activation and/or overexpression on 1) the gene expression profile, 2) glucose and fatty acid metabolism, and 3) glucose-stimulated insulin secretion in primary isolated rat islets. Using a combination of agonist treatment and adenoviral gene delivery to achieve a graded activation of PPAR
, we show that this factor unexpectedly enhances the expression of genes responsible for fatty acid oxidation and disposal in islets, including those involved in mitochondrial and peroxisomal
-oxidation, fatty acid transport, and binding. Although the consequent metabolic changes are associated with an acute suppression of glucose oxidation and glucose-stimulated insulin release, they may also serve to preserve islet
-cell mass in the longer term by attenuating the proapoptotic effects of fatty acids and their nonoxidative metabolites (43).
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MATERIALS AND METHODS |
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Collagenase (1.2 pzu/mg) was obtained from Boehringer Mannheim (Mannheim, Germany). Culture medium (Dulbecco's modified Eagles medium; DMEM), and heat-inactivated fetal bovine serum (FCS), were obtained from Life Science Technologies (Paisley, UK). Histopaque gradient solutions, penicillin, streptomycin, glutamine, and TRI Reagent were obtained from Sigma (Poole, Dorset, UK).
Amplification of Recombinant Adenoviruses
Recombinant adenoviruses encoding 1) constitutively active SREBP-1c (amino acids 1403, wild-type sequence; SREBP CA) (2, 18), 2) enhanced green fluorescent protein (eGFP; null), or 3) PPAR1 (wild-type sequence; Ad.PPAR
), were generated and amplified as described (22).
Islet Isolation and Culture
Pancreatic islets were isolated from male Wistar rats (225250 g) and killed according to locally approved animal procedures. Perfusion of the pancreatic duct and in situ collagenase digestion were performed as described (19). Islets were subsequently purified on either Histopaque gradient solutions (10 ml of 1.119 g/l; 6 ml of 1.083 g/l, 6 ml of 1.077 g/l) or gradients of bovine serum albumin (35%, 31% and 27% wt/vol). Isolated islets were cultured in suspension for 16 h in DMEM containing 10% (vol/vol) FCS, 11 mM glucose, 2 mM glutamine, 100 IU/ml penicillin, and 100 mg/ml streptomycin and incubated at 37°C with 95% air-5% CO2. Islets were then hand picked the next day for further experiments.
Adenoviral Infection
Islets were infected with SREBP CA, Ad.PPAR, or null adenoviruses at a multiplicity of infection of 50 viral particles/cell for 32 h. Islets were then placed in medium containing 3 mM glucose in the presence of the PPAR
agonist GW-347845 (200 nM) or its vehicle (dimethyl sulfoxide at a final concentration of 0.004%) for a further 16 h before use.
Western Blot Analysis
Total protein extracts (25 µg) were resolved by SDS-PAGE (10% bisacrylamide-acrylamide, vol/vol) and transferred to polyvinylidene difluoride membranes (50), followed by immunoblotting with rabbit polyclonal anti-PPAR antibody raised against a peptide corresponding to amino acids 6105 of PPAR
(Santa Cruz Biotechnology, Santa Cruz, CA). Secondary antibodies were revealed using BM Chemiluminescence blotting substrate (Roche Diagnostics, Mannheim, Germany).
Immunocytochemistry
Islet slices were prepared essentially as described (12). Briefly, islets were fixed with Zamboni's fixative overnight at 4°C, and sections of islets (10 µm) were obtained using a cryostat (Bright OTF5000; Jencons, Leighton Buzzard, UK). Islets were permeabilized with 0.3% (vol/vol) Triton X-100 overnight and then incubated with primary antibodies at 4°C. Rabbit polyclonal anti-PPAR antibody was used at 1:50 dilution and guinea pig anti-insulin antibody at 1:500 dilution. Primary antibodies were revealed using TRITC- or FITC-conjugated secondary antibodies against rabbit or guinea pig IgG (1:500 dilution), respectively.
Microarray Analysis
Sample preparation and processing. Total RNA samples from four separate islet cultures per condition were isolated by TRI Reagent and purified on an RNeasy column (Qiagen, Valencia, CA) before labeling for hybridization to Affymetrix RAE230 rat arrays (Affymetrix, Santa Clara, CA). Total RNA was prepared for hybridization and subsequently hybridized according to recommendations from Affymetrix (Genechip Expression Analysis Technical Manual [online] http://www.affymetrix.com/support/technical/manual/expression_manual.affx) (2003), based on a small-sample labeling protocol. Briefly, each sample was processed from 100 ng of total RNA that were first converted to cDNA. All of the cDNA was subsequently converted to cRNA in the first round of linear amplification. The resulting cRNA (400 ng) was used as a template for second-round cDNA synthesis that was again converted to cRNA. The final cRNA product (20 µg) was fragmented, and 15 µg of the fragmented cRNA were hybridized to the Affymetrix chips. Hybridized chips were washed and then scanned on an Affymetrix GeneChip 3000 confocal scanner. Gene expression data were generated in the GeneChip MAS 5.0 software from Affymetrix. Both total RNA and cRNA samples were checked for quality on an RNA 6000 Nano LabChip with the Bioanalyzer system (Agilent Technologies, Palo Alto, CA).
Microarray data analysis. From the GeneChip MAS 5.0 software, we obtained an expression signal and a Present/Absent call status for every probe set on each of the hybridized Affymetrix chips. Multiple probe sets often encode a single gene. All expression data were normalized by global scaling to a trimmed average intensity of 150 per chip. The Present/Absent call status was used as a criterion for gene filtering; probe sets with a Present call in three of the four replicates in at least one of the four treatments were kept for subsequent analysis. Control, noneukaryotic genes were also removed. After filtering, 9,563 of a total of 15,923 probe sets were retained for analysis.
A principal-components analysis was performed on the 16 (4 treatments x 4 replicates per treatment) samples, using gene signals as sample descriptors to identify outlier samples. None of the samples behaved as outliers (data not shown); consequently, the full set of 16 samples was retained for analysis.
Gene signals were logged (to base 2). The mean signal for all genes within replicate samples was calculated on the logged signals. Log ratios of the average signals were calculated to estimate the fold change in gene expression between treatments. Statistical significance of the difference in expression for a gene between two treatments was assessed by the heteroscedastic two-tailed Student's t-test for unpaired samples. Because the t-test was performed over thousands of genes, a false discovery rate (46) was also calculated to adjust for multiple testing using the Q-value program in the R statistical package.
Analysis of clusters.
Genes were ranked in order of increasing P value according to differences between the PPAR plus GW-347845 vs. null adenovirus groups. Among those genes that showed highly significant (P < 0.01) changes (
300 in total), clusters were identified for all groups (null, agonist alone, PPAR
alone, or combined treatment) according to expression changes (Gene Tree, GeneSpring; SiliconGenetics, Redwood City, CA; www.silicongenetics.com) (15) (see Fig. 3D) and functional similarity (GeneSpring and GeneMAPP, Gladstone Institutes, University of California at San Francisco, www.genemapp.org; see Table 2).
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Semiquantitative and Real-Time Quantitative RT-PCR
Total RNA was extracted using TRI Reagent from 100 islets. RNA samples were treated with DNA-free (Ambion, Austin, TX) to remove any contaminating genomic DNA, and the concentration was determined using RiboGreen assay (Molecular Probes, Eugene, OR). Complementary DNA was synthesized from 1 µg of total RNA in 100 µl by using random hexamers and Moloney murine leukemia virus reverse transcriptase. All genes, primers, and probes used in real-time RT-PCR analysis are described in Table 1. PCR was performed using 20 ng of reverse-transcribed total RNA with 900 nM of sense and antisense primers, 5.5 mM MgCl2, 300 µM dNTP, 1.25 U of Taq polymerase and 1x Taqman buffer A (Applied Biosystems, Foster City, CA) in a total volume of 25 µl in an ABI PRISM 7700 Sequence Detection System instrument. Standard curves were constructed by amplifying serial dilutions of untreated rat islet cDNA (50 ng to 0.64 pg) and plotting cycle threshold values as a function of starting reverse-transcribed RNA, the slope of which was used to calculate relative expression of the target gene.
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Total lipids were extracted from 150 islets by use of a chloroform-methanol (2:1 vol/vol) mixture. The extracted lipids were air dried, and 10 µl of a detergent (Thesit; Fluka, UK) were added to the dry pellet. Samples were air dried again and resuspended in 50 µl of water. TG was measured using a commercial kit (Infinity Triglyceride Reagent, Sigma) and a standard curve of triolein (Sigma) treated in parallel with the samples.
Insulin Secretion
Forty-eight hours after adenoviral infection, islets were incubated for 60 min at 37°C in 2 ml of Krebs-Ringer bicarbonate-HEPES buffer (KRBH) [in mM: 130 NaCl, 3.6 KCl, 1.5 CaCl2, 0.5 MgSO4, 0.5 KH2PO4, 2.0 NaHCO3, and 10 HEPES] supplemented with 3 mM glucose, 0.1% (wt/vol) bovine serum albumin preequilibrated with 95% O2-5% CO2, pH 7.4. Islets were separated into three groups of five islets per condition and incubated as above for 30 min in 1 ml of KRBH containing either 3 or 17 mM glucose or 3 mM glucose plus KCl (35 mM). Total insulin was extracted in acidified ethanol (1). Insulin was measured using radioimmunoassay by competition with 125I-labeled rat insulin (Linco Research, St. Charles, MO) according to the manufacturer's instructions. Insulin secretion was calculated as the percentage of total islet insulin content and normalized as the fold change between individual experiments. Typically, insulin release in the presence of 17 mM glucose represented 23% of total islet insulin content/30 min.
ATP Assay
Rat islets were prepared and incubated as described above for the assay of insulin secretion. After experimental manipulation, islets were lysed in ice-cold perchloric acid (20% vol/vol, 125 µl), rapidly frozen, and stored at 80°C. Samples were thawed and adjusted to pH 7.4 with a known volume of neutralization mixture (0.5 M triethanolamine, 2 M KOH, 100 mM EDTA). Precipitated potassium perchlorate was removed by centrifugation (10,000 g, 2 min) and the supernatant stored on ice before assay. Neutralized sample (10 µl) was added to 970 µl of assay buffer (130 mM NaHAsO4, 17 mM MgSO4, 4 µM NaH2PO4, pH 7.4) and 10 µg of firefly lantern extract (10 mg/ml stock, Sigma). The reaction was initiated with 10 µg of firefly luciferase, and light emission was recorded for 30 s using a photon counting luminometer. ATP concentration was obtained by comparison to standards (025 pmol) prepared in parallel with the cell extracts.
[1-14C]Palmitate Oxidation
Islets were preincubated for 30 min at 37°C in KRBH supplemented with 3 mM glucose and 0.5% fatty acid-free BSA (wt/vol). Triplicate groups of 100 islets were then placed in 250 µl of oxidation mix (KRHB containing 0.2 mM unlabeled palmitate, 1 mM carnitine, 0.4 µCi/ml [1-14C]palmitate, 0.5% fatty acid-free BSA and 3 or 17 mM glucose) in a 24-well plate. A rubber gasket the size of the 24-well plate and containing 0.5-cm holes was aligned over the plate. A UniFilter-24 GF/B Plate (PerkinElmer, Beaconsfield, UK) was sealed with an adhesive sheet, and 100 µl of 40% (wt/vol) KOH were pipetted onto each filter. The filter plate was inverted and aligned over the rubber gasket to form a small CO2 capture chamber. Finally, the chamber was sealed with a 6-mm glass plate, a 6-mm metal plate, and a lead weight to ensure an airtight seal. The apparatus was incubated for 2 h at 37°C. Filters were removed, and captured 14CO2 was measured by scintillation counting. Control incubations lacking islets were included in each incubation series.
[U-14C]Glucose Oxidation
Islets were preincubated for 30 min at 37°C in KRBH supplemented with 3 mM glucose and 0.5% BSA (wt/vol). Triplicate groups of 100 islets were then placed in 250 µl of KRHB containing 3 or 17 mM unlabeled glucose, 1.7 µCi [U-14C]glucose, and 0.5% BSA (wt/vol) in a 24-well plate. A CO2 capture chamber was created as above for palmitate oxidation and incubated for 2 h at 37°C. Captured 14CO2 was measured by scintillation counting.
Statistical Analysis
All functional analyses were performed at least three times in triplicate. Data are presented as means ± SE. Statistical significance was assessed by Student's t-test for unpaired comparison and two-tailed analysis.
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RESULTS |
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Consistent with earlier reports (13), mRNAs encoding both PPAR1 and -
2 (Fig. 1A) were detectable in rat islets by semiquantitative and quantitative RT-PCR (Fig. 1, B and C). The expression of both isoforms was further induced (
1 by
7-fold and
2 by
2.5-fold; Fig. 1, B and C) by adenoviral transduction of islets with the active nuclear fragment of SREBP-1c (2, 11, 51). Confirming the likely expression of PPAR
in
-cells within the islet: 1) primers able to amplify either isoform generated the expected product by PCR using clonal MIN6
-cell-derived cDNA (Fig. 1D), and 2) PPAR
immunoreactivity, significantly above that in slices incubated in the absence of primary antibody (not shown), was associated with insulin-containing structures in null adenovirus-infected islets (see below; Fig. 2B).
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In an effort to identify the principal target genes for PPAR in the islet, we next explored the impact of adenoviral transduction of islets with PPAR
1. As shown in Fig. 2A, infection of islets with PPAR
1-bearing virus increased levels of PPAR
protein
10-fold. PPAR
1 protein expression was detected in 3075% of islet cells as assessed by immunocytochemical analysis of islet slices. Although low levels of PPAR
overexpression could be detected throughout both small (
50 µm diameter) and larger islets (Fig. 2B, bottom), the highest levels were achieved in cells at the islet periphery, of which the majority were insulin positive (Fig. 2B, bottom, "zoom"). This pattern is similar to that previously reported in islets infected with either null (eGFP-expressing) or SREBP-1c-encoding adenoviruses (12) and was not associated with any detectable increase in cellular apoptosis (assessed by annexin V binding) or necrosis (12) at the viral titer deployed (data not shown).
Oligonucleotide microarray analysis (Fig. 3 and Table 2) revealed that, of 15,000 mRNAs and expressed sequence tags present on the arrays used, 9,563 were present in islets (see MATERIALS AND METHODS). To test the effect on gene expression of the progressive activation of PPAR
, we employed three protocols: A) treatment with a PPAR
-specific agonist (200 nM GW-347845), which displays >103-fold selectively over PPAR
or PPAR
(for the human and mouse isoforms, log EC50 values for transactivation are, respectively, 9.2 and 8.9 for PPAR
, 5.5 and 5.0 for PPAR
, and 4.6 and 5.0 for PPAR
) (31), B) adenoviral transduction with PPAR
1, and C) the combination of both treatments (Fig. 3 and Table 2). These three protocols aimed to examine the effects of relatively small, physiological increases in endogenous PPAR
activity (2- to 10-fold; treatment A), similar to those achieved after the overexpression of active SREBP-1c (Fig. 1) (12) or in the islets of ZDF rats (53), as well as the impact of much larger increases in PPAR
activity (treatments B and C). By providing much more substantial increases in the transcription of target genes, the latter protocols (B and C) were expected to permit the unambiguous identification of genes whose changes may be too small to detect by microarray analysis when PPAR
was weakly overexpressed.
Of more than 15,000 endogenous mRNAs examined, the levels of 284 were affected very significantly (P < 0.0099) by infection with the PPAR1-encoding adenovirus and incubation with GW-347845 (treatment C). By contrast, 119 and 49 genes, respectively, were altered by expression of PPAR
1 alone or treatment with GW-347845 after null virus infection (treatments B and A, respectively; Fig. 3). Analysis of clusters (15) revealed three distinct groups of genes that were progressively downregulated [a and b (i)] or upregulated [b (ii) and (iii), c (i) and (ii)] moving through treatments A to C (Fig. 3). Maximal stimulation of PPAR
1 (treatment C) had no effect on the expression of a number of genes involved in glucose sensing by islet cells, including glucokinase or the ATP-sensitive potassium channel subunits Kir6.2 or SUR1, but did cause a small but significant decrease in mRNA encoding pancreatic duodenum homeobox-1 (PDX-1) (Fig. 3 and Table 2). Similarly, maximal PPAR
1 activation had no or minor effects on the levels of cell cycle-dependent genes, including cyclin D1, and did not affect the expression of several proapoptotic (Bcl-x, BAD) or antiapoptotic (p53, Bcl2) genes (data not shown and Table 3).
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Changes in the expression of key genes detected by microarray analysis were quantified by real-time PCR analysis (TaqMan; Table 3). This analysis confirmed that lipogenic genes (acetyl-CoA carboxylase, fatty acid synthase) were minimally affected by PPAR1/GW-347845 treatment, whereas GLUT2/Scl2a2 and PDX-1 expression was slightly decreased at 3 and 17 mM glucose, respectively (Table 3). By contrast, RT-PCR analysis revealed that expression of mitochondrial carnitine palmitoyltransferase I and the plasma membrane FAT/CD36 (9) were both strongly induced by PPAR
1/GW-347845 (Table 3). Conversely, pyruvate dehydrogenase kinase-4 was upregulated in the presence of activated PPAR
1 (Table 3), a change likely to inhibit mitochondrial pyruvate oxidation (35).
Effects of PPAR Activation on Islet Glucose and Fatty Acid Metabolism
As shown in Fig. 4A, the combination of transduction with PPAR1 and incubation with GW-347845 (treatment C) significantly diminished islet TG levels. By contrast, the oxidation of exogenously added palmitate was significantly increased by this combined treatment (Fig. 4B), whereas the oxidation of 17 mM glucose was inhibited by >70% under these conditions (Fig. 4C). Correspondingly, glucose-stimulated increases in islet ATP content were completely inhibited by GW-347845 or expression of PPAR
, in either the presence or absence of the agonist (Table 4), with no associated changes in the expression of mitochondrial uncoupling protein-2 (Table 3). By contrast, culture of nontransfected islets in the presence of the positive agonist alone had no impact on the oxidation of exogenous palmitate or of glucose (Fig. 4, D and E).
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We next determined whether the marked changes in islet glucose metabolism caused by PPAR activation were associated with changes in insulin release. As shown in Fig. 5A, glucose-stimulated insulin secretion was significantly reduced after infection of islets with adenoviral PPAR
1 in either the presence or absence of GW-347845. In contrast, incubation with the PPAR
agonist alone had no effect on insulin secretion at 17 mM glucose but significantly enhanced release at 3 mM glucose (Fig. 5A). The metabolism-independent stimulation of secretion with 35 mM KCl was unaffected by either PPAR
1 alone or PPAR
1 plus GW-347845 (Fig. 5B), consistent with a preserved insulin content and activity of the basal exocytotic machinery under these conditions. Correspondingly, no changes in the levels of mRNAs encoding proteins involved in exocytosis (24), including members of the syntaxin or synaptotagmin families, synaptosomal protein-25 (SNAP25), or Munc18 were revealed by microarray analysis (data not shown), although the expression of Rim1 (Rab3-interacting molecule 1), a known regulator of exocytosis in neurons (24), was decreased (Table 2). Further indicating that the actions of GW-347845 were likely to be mediated by PPAR
, the effects of the drug on glucose-induced insulin secretion (Fig. 5C) and on the expression of two target genes (Fig. 6) in islets infected with adenoviral PPAR
were completely reversed with the PPAR
-selective antagonist GW-9662 (Fig. 5, legend).
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DISCUSSION |
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We demonstrated first that mRNAs encoding both PPAR1 and PPAR
2 isoforms are present in primary rodent islets and that PPAR
1 and -
2 levels are considerably increased after the forced expression of SREBP-1c. The latter factor has previously been shown to be upregulated severalfold by glucose in clonal
-cells (2, 51) but more modestly (
60%) in isolated islets (11, 12). Indeed, in the present studies, SREBP-1c expression was not significantly affected by elevated glucose concentrations (Table 3), a difference that may reflect a shorter preincubation period (16 h) at low glucose concentrations than in our previous reports (24 h) (11, 12).
In contrast to the effects of overexpressing activated SREBP-1c in islets (26), increases in PPAR1 activity over a broad range had no effect on, or even reduced, the expression of lipogenic genes in the islet (Tables 2 and 3). Moreover, because the expression of key "glucose-sensing" genes (e.g., Kir6.2, SUR1, and glucokinase) was not affected by PPAR
1 activation, whereas the expression of GLUT2 and PDX-1 was only weakly reduced (Fig. 7, legend), PPAR
activation alone would appear insufficient to promote the transdifferentiation of islet cells toward an adipocyte phenotype (30, 48). On the contrary, the presence of activated PPAR
1 decreased islet TG content and glucose oxidation while stimulating fatty acid oxidation (Fig. 4). Thus PPAR
would seem to play little if any role in mediating the effects of SREBP-1c on gene expression but may, rather, act as a "brake" to oppose the lipogenic effects of the latter factor (see below).
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PPAR agonists including troglitazone (32) have previously been reported to activate insulin release acutely, an effect possibly mediated by binding to ATP-sensitive K+ channels (47). A direct action of GW-347845 on these channels may thus contribute to the stimulation of insulin release observed here in response to the agonist alone (Fig. 5A). Although the longer-term effects of PPAR
activation are consistent with the previously described action of troglitazone to reduce the lipid content of ZDF rat islets (26, 43), the favoring of fatty acid oxidation over synthesis was unexpected. Indeed, in other tissues such a change is more reminiscent of the activation of PPAR
than PPAR
(10, 39). Importantly, these effects cannot be ascribed to an upregulation of PPAR
by overexpressed PPAR
, since the level of PPAR
expression was found to be low in control islets and unaffected by any of the treatments used (data not shown). It should be stressed, however, that GW-347845 is expected to be highly selective for the activation of PPAR
compared with other PPAR isoforms at the concentration used here (see RESULTS). We therefore speculate that the apparent tissue-specific differences in PPAR
function revealed here may reflect differing levels of PPAR
-binding partners such as retinoic acid receptors or of coactivators such as CREB-binding protein/p300 or PPAR
coactivator (PGC)-1 (37). In this context, PGC-1
was recently shown to be expressed at low levels in nondiabetic islets but induced in two models of diabetes and associated with an inhibition of glucose oxidation and insulin secretion (52). In addition, the complete absence from the islet of adipocyte-specific transcription factors such as CCAAT box enhancer-binding protein-
/
(8) may limit the ability of PPAR
to induce the expression of lipid-synthesizing enzymes in islet cells.
Recent studies have indicated that an accumulation of free fatty acids or nonoxidative metabolites, including ceramides (49), rather than the formation of TG (5), may be most closely correlated with apoptotic changes and loss of -cell function. We show here that the forced activation of PPAR
in the islet leads to the stimulation of multiple metabolic pathways that favor the disposal of fatty acids (mitochondrial and peroxisomal oxidation, cellular export, and incorporation into diglycerides). Such mechanisms may oppose the deleterious long-term consequences for islet
-cell survival and function of high levels of circulating fatty acids and thus restrict the impact of the glucose-induced upregulation of lipogenic transcription factors including SREBP-1c (2, 51).
A recent report (40) has suggested that PPAR in islet
-cells has only a limited role in the antidiabetic effects of TZDs. In the latter studies, the normal expansion of
-cell mass that occurs in control mice in response to high-fat feeding was markedly blunted in PPAR
-deleted animals. However, PPAR
deletion in
-cells had no impact on the improvement in glucose tolerance elicited by rosiglitazone in high-fat-fed mice. Whether deletion of PPAR
in islet
-cells affects the efficacy of TZDs in the context of other models of type 2 diabetes remains to be established.
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GRANTS |
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ACKNOWLEDGMENTS |
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FOOTNOTES |
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The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
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REFERENCES |
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