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Endocrinology Vol. 141, No. 8 2822-2828
Copyright © 2000 by The Endocrine Society

ARTICLES

Interleukin-1ß Regulates Phospholipase D-1 Expression in Rat Pancreatic ß-Cells1

Meng-Chi Chen, Veronica Paez-Espinosa, Nils Welsh and Décio L. Eizirik

Gene Expression Unit (M.-C.C., V.P.-E., D.L.E.), Diabetes Research Center, Vrije Universiteit Brussel, Laarbeeklaan 103, B-1090 Brussels, Belgium; and Department of Medical Cell Biology (N.W., D.L.E.), Uppsala University, S-751 23 Uppsala, Sweden

Address all correspondence and requests for reprints to: D. L. Eizirik, Gene Expression Unit, Diabetes Research Center, Vrije Universiteit Brussel, Laarbeeklaan 103, B-1090, Brussels, Belgium. E-mail: deizirik{at}mebo.vub.ac.be


   Abstract
Top
 Abstract
Introduction
 Materials and Methods
 Results
 Discussion
 References
 
The cytokine interleukin (IL)-1ß induces a biphasic effect in rat pancreatic islets, with an early and transitory stimulation of insulin release followed by progressive functional suppression. To clarify the mechanisms involved in these effects, we have recently performed a differential display of messenger RNA (mRNA) by RT-PCR (DDRT-PCR) on rat ß-cells exposed for 6 or 24 h to IL-1ß. Among the different IL-1ß-induced genes, there was an early and transient increase in phospholipase D-1 (PLD1) expression. PLD1 can induce phosphatidic acid formation and subsequent activation of protein kinase C, a process which stimulates insulin release. In the present study, we characterized the regulation of PLD isoforms by IL-1ß in pancreatic ß-cells. By using different combinations of primers and RT-PCR, we observed that IL-1ß induces an early increase (2 and 6 h) in the expression of both alternatively spliced isoforms of PLD1 (PLD1a and 1b). Prolonged exposure to IL-1ß (12 and 24 h) caused a decrease of PLD1a mRNA expression compared with control ß-cells, and lead to a return of PLD1b mRNA to basal level. NG-methyl-L-arginine (L-MA), a blocker of the inducible form of nitric oxide synthase (iNOS), prevented this late inhibitory effect of IL-1ß, suggesting that IL-1ß-induced decrease in PLD1a expression is NO-mediated. IL-1ß induced an early (2–6 h) and sustained (16–24 h) increase in PLD1a mRNA expression in insulin-producing RINm5F cells. This was paralleled by a cytokine-induced increase in PLD1 protein expression and enzyme activity. RINm5F cells, but not primary ß-cells, expressed PLD2, and the expression of this gene was not affected by IL-1ß. In conclusion, we have shown that the cytokine IL-1ß regulates PLD1 expression in primary and clonal ß-cells. The early induction of PLD1 probably contributes to the early stimulatory effects of IL-1ß on islet insulin release.


   Introduction
Top
 Abstract
 Introduction
Materials and Methods
 Results
 Discussion
 References
 
THE CYTOKINE interleukin (IL)-1ß induces an initial phase of functional stimulation in rodent pancreatic islets, which is followed after 4–7 h by a progressive inhibition of insulin release and eventually ß-cell damage (1, 2, 3). Human islets cultured in the presence of IL-1ß alone present a more prolonged stimulatory phase, which may last up to 48 h (4). This initial stimulation of insulin release is associated to increased diacylglycerol (DAG) production and protein kinase C (PKC) activation (5), but it remains to be determined which steps are involved in IL-1ß-induced PKC activity and in the subsequent inhibitory effects on ß-cell function.

To further clarify this issue, we have recently performed a differential display of messenger RNA (mRNA) by RT-PCR (DDRT-PCR) on rat ß-cells exposed for 6 or 24 h to IL-1ß (6, 7). Among the genes whose expression was modified by IL-1ß, we detected a transient increase in the expression of phospholipase D-1 (PLD1) mRNA (6). So far three PLD isoenzymes have been identified: PLD1a, PLD1b, and PLD2 (8, 9, 10, 11). PLD catalyses the hydrolysis of phosphatidylcholine (PC), leading to formation of phosphatidic acid (PA) and choline, and subsequent PKC activation. The activation of PKC may occur via a direct effect of PA or via DAG formed by the cleavage of PA by phosphohydrolase (12, 13, 14). It has been proposed that PLD1 and PLD2 have distinct functions in certain cell types. PLD1 plays an important role in signal transduction, cell growth, and in intracellular protein trafficking and secretion (13, 15, 16). Exogenously added PLD or its product PA activates exocytosis of insulin granules in rat islets (17, 18). PLD1a and PLD1b are formed by alternative splicing of mRNA, with PLD1b missing a 114-bp exon [equivalent to 38 amino acids (aa)]), which is preserved in PLD1a (10, 11). This splicing does not affect catalytic activity, and both isoenzymes show similar properties (10, 11). The other isoenzyme, PLD2, has different regulatory and functional properties, and is mostly involved in cytoskeletal reorganization (9).

It has been previously suggested that pancreatic islet cells exhibit a phospholipase D-like mechanism, which is activated by phorbol esters (19). It remains, however, to be characterized which of the three isoenzymes of PLD are expressed in ß-cells. In the present study we used RT-PCR to determine the effects of IL-1ß on the expression of mRNAs for PLD1a, PLD1b, and PLD2 in FACS-purified primary ß-cells and in insulin-producing RINm5F cells. Total PLD1 protein expression and enzyme activity were also measured in this cell line.


   Materials and Methods
Top
 Abstract
 Introduction
 Materials and Methods
Results
 Discussion
 References
 
Islet ß-cell isolation and culture
Pancreatic islets were isolated from 10-week-old male Wistar rats by collagenase digestion and islet ß-cells were purified by autofluorescence-activated cell sorting (FACStar, Becton-Dickinson and Co., Sunnyvale, CA) (20). The ß-cells were precultured overnight at 37 C in Ham’s F-10 medium (Life Technologies, Inc., Scotland, UK) supplemented with 5 mg/ml charcoal-treated BSA (fraction V, RIA grade, Sigma, St. Louis, MO), 0.1 mg/ml streptomycin, 12.5 U/ml penicillin, 0.3 mg/ml L-glutamine and 10 mmol/liter glucose. This was followed by culture at 37 C in the same medium, but with addition of 50 µmol/liter 3-isobutyl-L-methylxanthine (IBMX; Jansen Chimica, Beerse, Belgium), which is required to preserve ß-cell viability over prolonged periods in culture (21). Insulin-producing RINm5F cells (provided by Å. Lernmark, then at the Hagerdon Institute, Copenhagen, Denmark) were cultured in medium RPMI-1640 supplemented with 10% (vol/vol) FCS, 0.1 mg/ml streptomycin, and 12.5 U/ml penicillin. Recombinant human IL-1ß was kindly provided by Dr. C. W. Reynolds, National Cancer Institute (Frederick, MD). L-{alpha} phosphatidylcholine (from fresh egg yolk), 3,3-dimethylglutaric acid, oleic acid and phosphatidic acid were from Sigma (St. Louis, MO) L-3-phosphatidylcholine and L-palmitoyl-2-[1-14C]oleyl (55 mCi/mmol) were from Amersham Pharmacia Biotech (Uppsala, Sweden).

To characterize PLD1 mRNA expression, rat ß-cells (105 cells) or RINm5F cells (105 cells) were exposed for 2, 6, 12, and 24 h (in some experiments ß-cells were also exposed for 72 h) to IL-1ß alone (30 U/ml), or for 24 h to the cytokine in the presence of the iNOS inhibitor NG-methyl-L-arginine (L-MA, 1.0 mM; Sigma). Culture media were collected in parallel for nitrite measurements (nitrite is a stable product of NO oxidation). Nitrite determination was performed spectrophotometrically at 546 nm wavelength, after reaction with the Griess reagent (22). The choice of cytokine and inhibitor agent concentrations was based on our previous data on insulin-producing cells and pancreatic islets, having as biological end point stimulation (cytokine) or inhibition (L-MA) of cytokine-induced iNOS expression and/or medium nitrite accumulation (23, 24, 25, 26).

Determination of cell viability
Single ß-cells were distributed over polylysine-coated cups (5 x 103 cells/cup in 200 µl medium) and cultured for 72 h with or without IL-1ß. Cell viability was assessed by staining with the nuclear dyes propidium iodide (PI, 10 µg/ml) and Hoechst (HO) 324 (20 µg/ml), as previously described (27). For RINm5F cells (104 cells/cup), viability was assessed by the trypan blue exclusion test after a 24 h culture.

Determination of mRNA expression by RT-PCR
Poly(A)+ RNA was isolated from cell aggregates using oligo(dT)25-coated polystyrene Dynabeads (DynAl, Oslo, Norway) and reverse transcribed into cDNA as previously described (6, 7). Different PLD1 forward primers in combination with a single reverse primer were used to study mRNA expression of the individual PLD1 transcripts (Fig. 1Go). cDNA samples (equivalent to 3 x 103 cells) were amplified by PCR using AmpliTaq Gold DNA polymerase (Perkin-Elmer Corp.). PCR reactions included 12 min predenaturation (hot start PCR) at 95 C and then cycles of: 94 C for 45 sec, 58 C for 45 sec, and 72 C for 80 sec. The numbers of cycles for the PCRs were 28 for GAPDH, 33 for PLDF1.1 and PLDF1a, and 31 for PLDF1.2, and they were selected to allow amplification within the linear range. The gene-specific primer sequences and their respective PCR fragment lengths are GAPDH-F: 5'-TCCCTCAAGATTGTCAGCAA-3', GAPDH-R: 5'-AGATCCACAACGGATACATT-3', (308 bp); rat PLDF1a: 5'-CCTTCCAGTGAGTCTGAGCA-3' (422 bp for PLD1a), rat PLDF1.1: 5'-TAATCTGCACAACTCGGACA-3' (540 and 426 bp for PLD1a and 1b, respectively), rat PLDF1.2: 5'-TCCATCCGTAGTGTGCAGAC-3' (371 bp for PLD1a plus 1b), PLD1-R 5'-GCAGCAGATCGGAGCAACTG-3'; PLD2-F: 5'-CCCTTTCTGGCCATCTATGA-3', PLD2-R: 5'-GCGATAGAAAGACATGGTCA-3' (417 bp). The identity of PCR fragments of each gene were confirmed by DNA sequencing (data not shown). The ethidium bromide-stained agarose gels were photographed under UV-transillumination using Kodak Digital Science DC40 camera (Kodak, Rochester, NY). Abundance of the PCR products of interest was assessed by Biomax 1D Image analysis software (Kodak) and expressed in pixel intensities (OD), normalized for the abundance of the GAPDH signal amplified from the same cDNA sample. We have previously shown that IL-1ß does not modify expression of the "housekeeping" gene GAPDH (28).



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  		Figure 1. Rat PLD proteins and the PCR primers used in the present study. The arrows show the positions of the PCR primers used in this study. Combinations of different PLD1 forward primers and a single reverse primer allowed us to study the mRNA expression of the two alternatively spliced isoforms PLD1a and PLD1b. F1.1 and R1 amplified both 1a and 1b isoforms as two discrete bands. F1a and R1 amplified the PLD1a isoform only. F1.2 and R1 amplified the two isoforms as a single band. F2 and R2 amplified PLD2 mRNA. DD175 indicates the region of induced PLD1 mRNA detected by DDRT-PCR (6 ). The boxes indicate the regions of high homology between three PLD isoenzymes. The numbers shown on the right margin indicate the expected length of the mature peptides. The figure was modified from (10 ).

 
Determination of PLD protein expression and enzyme activity
Due to the difficulty in obtaining enough purified rat ß-cells for these experiments, the immunoprecipitation of PLD1 protein and the determination of PLD enzyme activity were performed in RINm5F cells. For immunoprecipitation, 2 x 106 cells per condition were disrupted in modified radioimmunoprecipitation (RIPA) lysis buffer (1 x PBS, 1% Nonidet P-40, 0.25% sodium deoxycholate, 0.1% SDS, 1 mM PMSF in isopropanol, aprotinin 1 mg/ml, 1 mM sodium orthovanadate, 1 mM NaF). The extracts were centrifuged at 13,000 x g in an Eppendorf centrifuge at 4 C for 20 min to remove insoluble material. Protein quantification was performed using the micro BCA Protein Assay (Pierce Chemical Co., Rockford, IL). Because the antibody for rat PLD1 protein is not available, we used instead rabbit antibody raised against residues 1–15 of human PLD1 (Biosource Technologies, Inc., Nivelles, Belgium; 10). The amino acid sequences of this region are very similar between the two species, allowing good cross-reactivity of the human antibody against the rat PLD1. The antibody was added to 600 µg of protein and incubated overnight. The complex formed by PLD1 and antibody was captured by Dynabeads protein A (DynAl, Oslo, Norway). The unbound proteins were separated by centrifugation and removed. PLD1 protein samples were then run on a 10% SDS-polyacrylamide gel. After electrophoresis, proteins were electrically transferred to nitrocellulose filters and then incubated with the same PLD1 antibody. Horseradish peroxidase-linked goat antirabbit IgG was used as second antibody. Peroxidase activity was detected by enhanced chemiluminescence (Amersham Pharmacia Biotech, Buckinghamshire, UK).

For the determination of phospholipase D enzyme activity, a previously described method (29) was used, with minor modifications. Briefly, groups of 5 x 106 RINm5F cells were mechanically homogenized in 150 µl of ice cold homogenization buffer (pH 7.5) containing 0.32 M sucrose, 5 mM HEPES, 1 mM PMSF, and 1 mM dithiothreitol. The homogenate was centrifuged at 150,000 x g for 20 min and the pellet resuspended by mild sonication in 10 µl of homogenization buffer. An aliquot was used for the determination of protein content according to the Bradford method (30). To the remaining membrane fraction, it was added 2.5 mM [14C]phosphatidylcholine (1.67 Ci/mol), 5 mM sodium oleate, 50 mM ß-dimethylglutaric acid, pH 6.5, 10 mM EDTA, and 25 mM NaF in a total volume of 20 µl. The reaction was carried out at 30 C for 90 min and was terminated by the addition of 400 µl chloroform and 200 µl methanol. Phosphatidic acid (3 µg) was added as a carrier. One hundred fifty microliters of water were added, and the samples were thoroughly mixed. After centrifugation, the chloroform phase was evaporated under a stream of nitrogen and redissolved in 40 µl of chloroform/methanol (2:1, by volume). PBS containing 10 mM EDTA (10 µl) was added, and the samples were again mixed and centrifuged. The resulting chloroform fraction was spotted onto silica gel plates with a solvent system consisting of chloroform/methanol/acetone/acetic acid/water (50:15:15:10:5, by volume). The phosphatidic acid spots were scraped off and the radioactivity was determined by scintillation counting.

Statistical analysis
Values are expressed as means ± SEM and comparisons between groups were performed by Student’s paired t test.


   Results
Top
 Abstract
 Introduction
 Materials and Methods
 Results
Discussion
 References
 
Primary ß-cells express mRNAs encoding for both PLD1a and 1b isoforms (Fig. 2GoA). There was no detectable PLD2 expression in these cells even after 39 cycles of amplification (data not shown). IL-1ß caused a transient (2 and 6 h) induction (2- to 4-fold increase) of the two isoforms. Longer exposure to IL-1ß, however, caused a progressive decrease in PLD1a and PLD1b expression. After 12 h of culture in the presence of IL-1ß the cellular content of PLD1a mRNA declined to levels lower than those observed in control ß-cells, with further decrease after 24–72 h exposure to the cytokine (2- to 5-fold decrease) (Fig. 2Go, A and B). PLD1b mRNA returned to basal levels after a 12 or 24 h exposure of IL-1ß (Figs. 2Go, A and C), but the inhibitory effect of IL-1ß was less pronounced on PLD1b than on PLD1a. After 72 h of culture in the presence of IL-1ß PLD1b mRNA level was also decreased (nearly 50%), but again this inhibition was less marked than that observed on PLD1a (Fig. 2CGo).



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  		Figure 2. IL-1ß affects PLD1 expression in rat ß-cells. Rat ß-cells (5 x 104 cells/well) were exposed to IL-1ß (30 U/ml) or control condition (C, no cytokine added) for 2, 6, 12, 24, or 72 h. At these different time points the cells were harvested, mRNA extracted, and RT-PCR performed with the equivalent of 3 x 103 cells. The different forward primers and a single reverse primer used in these experiments are described in Fig. 1Go. The picture shown in Fig. 2AGo is representative for three to six similar experiments, and the values shown in Figs. 2BGo and 2CGo are means ± SEM of the same three to six experiments, presented as OD corrected per GAPDH expression. *, P < 0.05 vs. control cells.

 
Prolonged ß-cell exposure to IL-1ß leads to iNOS expression and NO production (reviewed in Ref. 2). To determine whether the late inhibitory effect of IL-1ß on PLD1 expression was mediated via NO production, ß-cells were exposed for 24 h to IL-1ß in the presence of the iNOS inhibitor L-MA. Nitrite production was increased by 8-fold following IL-1ß exposure (Fig. 3AGo), an effect abolished by the concomitant addition of L-MA. As described in Fig. 2Go, the cytokine inhibited PLD1a but not PLD1b expression (Fig. 3Go, B–D). LMA prevented both IL-1ß-induced NO production (Fig. 3AGo) and the inhibition of PLD1a expression (Fig. 3Go, B and C), suggesting that this effect of IL-1ß indeed depends on NO formation.



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  		Figure 3. The iNOS inhibitor L-MA prevents IL-1ß-induced decrease in PLD1a mRNA. Rat ß-cells (5 x 104 cells/well) were exposed to the control condition (C, no cytokine added), IL-1ß (30 U/ml), 1 mM L-MA or both for 24 h. The culture media were then collected for nitrite determination (Fig. 3AGo), and the cells harvested for RT-PCR determination of PLD1 mRNA (Fig. 3BGo). RT-PCR experiments were performed as described in Fig. 2Go. The figure is representative of three similar experiments. The values shown in Figs. 3Go, C and D, are the means ± SEM of these three experiments, presented as OD corrected per GAPDH expression. *, P < 0.05 vs. control cells.

 
PLD1a and PLD1b were also expressed in insulin-producing RINm5F cells (Fig. 4AGo). In contrast with the observations in primary ß-cells (see above), RINm5F cells also expressed PLD2 (Fig. 4AGo). IL-1ß increased 2- to 3-fold mRNA expression for both PLD1a and PLD1b (Figs. 4Go, B and C), while PLD2 was not affected by the cytokine. The stimulatory effects of IL-1ß were more prolonged (up to 24 h) in RINm5F cells (Fig. 4Go) than in primary ß-cells (Fig. 2Go). After a 24 h exposure to IL-1ß, there was also induction of NO production. Thus the values for nitrite measurements of RINm5F cells were respectively 9.4 ± 0.6 and 17.4 ± 1.4 pmol/24 h x 103 cells (n = 4; P < 0.05) for control and IL-1ß-treated cells. The observed IL-1ß-induced PLD1 expression in RINm5F cells was paralleled by an increased protein expression (Fig. 5AGo) and enzyme activity (Fig. 5BGo). PLD1 expression was detected by an antibody that recognizes both isoforms of the enzyme. Because the difference in the peptide length of PLD1a and PLD1b is only 38 aa, the immunoblot revealed only one discrete band of about 116 kDa (Fig. 5AGo), corresponding to the expected molecular weight of PLD1 (10). PLD1 was expressed under basal conditions in RINm5F cells, and there was a 2-fold increase in protein expression after 2, 6, and 24 h exposure to IL-1ß (Fig. 5AGo). In line with the protein data, RINm5F cells presented a high basal PLD enzyme activity, which was increased by 50–75% following a 6- to 15-h exposure to IL-1ß (Fig. 5BGo).



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  		Figure 4. Effects of IL-1ß on PLD mRNA expression in insulinoma RINm5F cells. RINm5F (5 x 104 cells/well) were exposed to IL-1ß (30 U/ml) or control condition (C; no cytokine added) for 2, 6, 16, or 24 h. After these time points the cells were harvested, mRNA extracted and RT-PCR performed with the equivalent of 3 x 103 cells (Fig. 4AGo). The different forward primers and reverse primers used in these experiments are described in Fig. 1Go. The pictures shown are representative for three to four similar experiments. The values shown in Fig. 4Go, B and C, are the means ± SEM of these three to four experiments, presented as O. D. corrected per GAPDH expression. *, P < 0.05 vs. control cells.

 


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  		Figure 5. Effects of IL-1ß on PLD protein expression and enzyme activity in RINm5F cells. PLD1 protein expression was measured by immunoprecipitation and enzyme activity was determined as described in Materials and Methods. The picture of immunoprecipitation (5A) is representative of three similar experiments. The values presented below the bands are the mean ± SEM of the band intensities (O. D.; n = 3; control intensity was considered as 1 in each individual experiment). The results of enzyme activity (5B) are the means ± SEM of three to four experiments. *, P < 0.05 vs. control cells.

 
The concentration of IL-1ß used in the present experiments did not decrease the viability of rat ß-cells or RINm5F at the time points studied. After 72 h of culture, the percentages of living, apoptotic or necrotic cells in control ß-cells (n = 6) were respectively: 78 ± 2%, 5 ± 1.5%, and 17 ± 1%, while in IL-1ß-treated groups (n = 6) these values were respectively: 77 ± 1%, 5 ± 1% and 18 ± 1% (P > 0.05 vs. controls). For RINm5F cells, after a 24 h culture (n = 4) there were 98 ± 1% living cells in control and 97 ± 1% living cells in IL-1ß-treated cells (P > 0.05 vs. controls).


   Discussion
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
References
 
In the present study, we report that primary rat ß-cells and clonal RINm5F insulin-producing cells express mRNAs for both isoforms of PLD1, namely PLD1a and PLD1b. RINm5F cells also express PLD2. These findings support previous observations that neonatal rat islets present a "phospholipase-D like mechanism" (19). The cytokine IL-1ß regulates PLD1 expression in primary ß-cells in a time-dependent manner—it causes an early (2–6 h) stimulation, followed by a late (12–24 h) inhibition.

The stimulatory phase of IL-1ß-induced PLD1 mRNA expression is more prolonged in RINm5F cells (up to 24 h), and in these cells it was observed that the increase in PLD1 mRNA expression is paralleled by an increase in PLD1 protein expression and PLD enzyme activity. The increase in mRNA expression was, however, more marked than the increase in protein expression or enzyme activity. The method used to determine PLD1 protein does not allow discrimination between PLD1a and PLD1b, whereas the method for enzyme activity measures both PLD1 and PLD2. Because the main effect of IL-1ß was to increase expression of PLD1a and, to a lesser extent PLD1b mRNA, with little or no effect on PLD2, this may explain why the observed effect on mRNA and protein expression was more marked than the effect on enzyme activity (measuring together PLD1 and PLD2).

After 12–24 h exposures, IL-1ß induced a preferential inhibition on PLD1a mRNA expression in primary ß-cells, with little effect on PLD1b expression. PLD1a and PLD1b are generated by alternative splicing of mRNA (10, 11), and these observations raise the possibility that the cytokine interferes with cis-acting regulatory elements and trans-acting factors that control alternative splicing (31). In line with this, we have previously observed that IL-1ß affects differently mRNA expression for cationic amino acid transporter-2 isoforms (CAT 2A and 2B) in rat ß-cells (Chen and Eizirik, unpublished data). These two CAT isoforms are generated by alternative splicing of mRNA and contain exclusively one of the two isoform-specific exons (32, 33). These data suggest that IL-1ß may affect cellular function by regulating the expression of alternative spliced isoforms.

The biphasic pattern of IL-1ß-induced PLD1 mRNA expression is similar to the biphasic effects of IL-1ß on rat and mouse islet insulin release (1, 34, 35, 36). It has been suggested that the early IL-1ß-induced insulin release is mediated by DAG formation and PKC activation (5). Mouse pancreatic islets exposed to IL-1ß present an early increase in DAG content, which is followed by PKC activation and increased glucose-induced insulin release (5). This initial phase of IL-1ß-induced insulin release is prevented by PKC inhibitors (5). The presently observed IL-1ß-induced increase in PLD1a mRNA and protein expression may provide an explanation for the previous observations that the cytokine induces both an increase in cellular DAG content and PKC activation (5). The PLD1 protein is localized in the endoplasmic reticulum, Golgi apparatus and late endosomes, where it promotes coated vesicle formation and trafficking (15, 16). PA, the product of the reaction catalyzed by PLD has been shown to act directly as a signaling molecule (37, 38), and exogenously added PLD or PA stimulate insulin release by rat pancreatic islets, probably by a direct stimulatory effect on the process of exocytosis (17, 18). These distinct stimulatory mechanisms triggered by PLD1 expression may, at least in part, explain the early stimulatory effects of IL-1ß on insulin release (Fig. 6Go). Interestingly, PKC may also activate PLD1 (39). This could provide a positive feedback on PLD1 activation and preserve IL-1ß-induced insulin release for some hours (1). This "stimulatory" phase would be then interrupted by the increased IL-1ß-induced NO production (see below). Note that IL-1ß activates or inhibits a wide number of biological responses (2). Thus, the observed parallel changes in PLD1a mRNA expression and insulin release do not necessarily indicate a cause-and-effect correlation. Additional experiments, including the blocking of PLD1a expression by antisense RNA, will be required to solve this issue. Long-term exposure of rat pancreatic islets to IL-1ß causes inhibition of glucose oxidation and insulin release (1, 2, 3). These effects are mediated to a large extent by cytokine-induced iNOS expression and NO production (2, 3). Indeed, both chemical iNOS blockers (40, 41) and deficient iNOS expression (iNOS gene knockout) (42) prevent IL-1ß-induced inhibition of insulin release. We have presently shown that the late suppressive effect of IL-1ß on PLD1a expression is also mediated by the radical NO. It remains to be clarified whether the observed decrease in PLD1a expression contributes for the late inhibitory effects of IL-1ß on pancreatic ß-cell function.



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  		Figure 6. Proposed model for the early induction of insulin release by IL-1ß. IL-1ß induces an early increase in PLD1 mRNA, protein and enzyme activity in ß-cells, probably increasing phosphatidic acid (PA) production. PA can be further cleaved by phosphohydrolase to produce diacylglycerol (DAG). The activation of protein kinase C (PKC), which stimulates insulin release in ß-cells, may be either due to the direct effect of PA on PKC activity, or via DAG formation and subsequent PKC activation. PKC may also contribute to PLD-1 activation.

 


   Acknowledgments
 
We are grateful to Professor D. G. Pipeleers for providing access to FACS-sorted ß-cells, to R. Leeman, and the personnel involved in rat islet isolation and ß-cell FACS sorting, for technical assistance.


   Footnotes
 
1 This work was supported by grants from the Juvenile Diabetes Foundation International (JDFI-198202), the Research Program of the Fund for Scientific Research—Flanders (FWO G.0062.00 and FWO Research Prize Pharmacia & Upjohn), by a Shared Cost Action in Medical and Health Research of the European Community (BMHY-CT98–3448), the Swedish Medical Research Council (72X-11564-05 C, 72P-12995–02B), the Nordic Insulin Fund, the Swedish Diabetes Association and the Ernfors Family Fund. Back

Received February 1, 2000.


   References
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 References
 

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