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Endocrinology Vol. 145, No. 3 1130-1136
Copyright © 2004 by The Endocrine Society

An Octamer Motif Is Required for Activation of the Inducible Nitric Oxide Synthase Promoter in Pancreatic ß-Cells

Martine I. Darville, Sara Terryn and Décio L. Eizirik

Laboratory of Experimental Medicine, Université libre de Bruxelles, B-1070 Brussels, Belgium

Address all correspondence and requests for reprints to: M. I. Darville, Laboratory of Experimental Medicine, Université libre de Bruxelles, Route de Lennik, 808 CP618, B-1070 Brussels, Belgium. E-mail: mdarvill{at}ulb.ac.be.


   Abstract
Top
 Abstract
Introduction
 Materials and Methods
 Results
 Discussion
 References
 
Nitric oxide, generated by the inducible form of nitric oxide synthase (iNOS), is a potential mediator of cytokine-induced ß-cell dysfunction in type 1 diabetes mellitus. We have previously shown that cytokine-induced iNOS expression is cycloheximide (CHX) sensitive and requires nuclear factor-{kappa}B (NF-{kappa}B) activation. In the present study, we show that an octamer motif located 20 bp downstream of the proximal NF-{kappa}B binding site in the rat iNOS promoter is critical for IL-1ß and interferon-{gamma} induction of promoter activity in rat primary ß-cells and insulin-producing RINm5F cells. In gel shift assays, the octamer motif bound constitutively the transcription factor Oct1. Neither Oct1 nor NF-{kappa}B binding activities were blocked by CHX, suggesting that other factor(s) synthesized in response to IL-1ß contribute to iNOS promoter induction. The high mobility group (HMG)-I(Y) protein also bound the proximal iNOS promoter region. HMG-I(Y) binding was decreased in cells treated with CHX and HMG-I(Y) silencing by RNA interference reduced IL-1ß-induced iNOS promoter activity. These results suggest that Oct1, NF-{kappa}B, and HMG-I(Y) cooperate for transactivation of the iNOS promoter in pancreatic ß-cells.


   Introduction
Top
 Abstract
 Introduction
Materials and Methods
 Results
 Discussion
 References
 
THE RADICAL NITRIC oxide, produced by the inducible form of nitric oxide synthase (iNOS), may contribute to ß-cell dysfunction and damage in early type 1 diabetes mellitus (reviewed in Refs. 1 , 2). Transgenic expression of iNOS in ß-cells leads to diabetes mellitus (3), whereas iNOS-/- mice have decreased sensitivity to diabetes induction by multiple low doses of streptozotocin (4). The cytokine IL-1ß, alone or in combination with interferon (IFN)-{gamma} and TNF{alpha}, induces the expression of iNOS in primary rodent ß-cells or insulin-producing cell lines, and both blocking of iNOS activity, or iNOS gene knockout, prevent some of the deleterious effects of cytokines in ß-cells (5, 6).

We have previously characterized the regulation of iNOS expression by cytokines in ß-cells and observed that a promoter region up to -1002 bp upstream of the transcription start site is necessary to obtain maximal induction of iNOS expression (7). This region contains, among other binding sites for transcription factors, a proximal and distal nuclear factor-{kappa}B (NF-{kappa}B) site, a {gamma}-interferon activated site (GAS), and two adjacent IFN-stimulated response elements (ISREs). Site directed-mutation analysis indicated that both NF-{kappa}B sites and the GAS are required for IL-1ß-induced iNOS expression (7). These observations are in good agreement with our recent finding that inhibition of NF-{kappa}B activation by overexpression of an inhibitor I{kappa}B mutant superrepressor prevents cytokine-induced iNOS expression in rat ß-cells (8). On the other hand, site mutation of the ISRE induced only a minor decrease in the induction of iNOS promoter activity by IL-1ß + IFN{gamma} (7). The lack of a major role for interferon regulatory factor-1 in cytokine-induced iNOS expression was confirmed by the well preserved iNOS expression in ß-cells isolated from interferon regulatory factor-1-/- mice (9).

Studies in macrophage cell lines suggest that an octamer (Oct) motif, located 15 bp downstream of the proximal NF-{kappa}B site in the mouse promoter, is required for iNOS activation by lipopolysaccharide (LPS) (10), IL-6 (11), and IFN{gamma} + LPS (12). In line with these findings, in vivo footprinting analysis of the iNOS promoter shows protein occupation of the Oct site (13), and transfection of fibroblast or neuronal cell lines with expression vectors encoding different Oct motif binding proteins activates the iNOS promoter (14). The Oct transcription factors are members of the Pit-Oct-Unc domain family, exerting their effects through binding to sequences related to the octamer motif (ATGCAAAT) (15). In primary rat aortic smooth muscle cells (RASMCs), the high mobility group (HMG)-I(Y) protein binds to the Oct motif and enhances transactivation of the iNOS promoter by NF-{kappa}B (16). In these cells, IL-1ß induces in parallel the expression of HMG-I(Y) and iNOS (17). HMG-I and -Y are two architectural proteins generated from the same gene by alternative splicing. They bind to AT-rich sequences in the minor groove of the DNA and physically interact with transcription factors to facilitate their recruitment to enhancer regions (Ref. 18 and references therein). It is, however, unclear whether these findings can be extrapolated to primary ß-cells. Indeed, there are important differences in the regulation of iNOS expression between ß-cells and other cell types. Thus, whereas iNOS expression is induced in murine macrophages by LPS and/or IFN{gamma}, ß-cells do not respond to these stimuli. On the other hand, IL-1ß alone induces iNOS expression in rodent ß-cells but not in macrophages (reviewed in Ref. 19). Moreover, regulation of HMG-I(Y) expression in insulin-producing cells has not been reported.

Against this background, the aim of the present study was to determine the role for the octamer motif and HMG-I(Y) in the regulation of iNOS expression by cytokines in both insulin-producing RINm5F cells and fluorescence-activated cell sorter, (FACS)-purified rat primary ß-cells.


   Materials and Methods
Top
 Abstract
 Introduction
 Materials and Methods
Results
 Discussion
 References
 
Cell culture and treatments
RINm5F insulinoma cells were cultured in RPMI 1640 medium with Glutamax-1 (Invitrogen, Paisley, Scotland) supplemented with 10% fetal calf serum. Rat ß-cells were FACS purified from pancreatic islets isolated from male Wistar rats as previously described (20). These preparations contained more than 95% ß-cells and were cultured in HAM’s F-10 medium supplemented with 10 mM glucose, 2 mM glutamine, 0.5% BSA, and 50 µM 3-isobutyl-1-methylxanthine. The presence of 3-isobutyl-1-methylxanthine at this concentration preserves ß-cell survival in culture (21) with minimal effects on cAMP formation in the absence of adenylate cyclase activators (22). The cells were exposed to 30 U/ml recombinant human IL-1ß (kindly provided by Dr. C. W. Reynolds, National Cancer Institute, Bethesda, MD) and/ or 1000 U/ml recombinant mouse IFN{gamma} (Invitrogen). The concentrations of cytokines selected for the experiments were based on previous data from our group (7, 23, 24, 25). When cycloheximide (CHX) was used, it was added to the culture medium at 20 µg/ml 15 min before the cytokines. We have previously shown that this concentration of CHX inhibits total protein synthesis by more than 90% in insulin-producing cells (26).

Plasmids, transfection, and luciferase assay
Plasmid piNOS-1002 luc-containing nucleotides -1002 to +132 of the rat iNOS promoter has been described (7). Mutation of the Oct binding site was introduced by PCR (27) using the Expand high-fidelity PCR system (Roche Molecular Biochemicals, Mannheim, Germany). Primers were 5'-AGAGCGTGGATGGGTATAAATACC (forward primer) and 5'-GCAGAGCTGTCGTGCATAAAGTCAC (reverse primer, with mutated nucleotides underlined). Mutation was confirmed by sequencing. RINm5F and primary ß-cells were cotransfected with the luciferase test plasmids and pRL-cytomegalovirus (CMV) (Promega, Madison, WI) as an internal control by lipofection using lipofectAMINE (Invitrogen, Gaithersburg, MD) as previously described (7). The following modifications were introduced for primary ß-cell transfections: 4 x 104 cells were seeded in 96-well plates and incubated for 3 h with 0.05 µg of test plasmid, 0.005 µg of pRL-CMV, 0.5 µg of PLUS Reagent, and 0.25 µl lipofectAMINE. For transfection with small interfering RNAs (siRNAs), 8 x 104 RINm5F cells were transfected with 0.2 µg piNOS-1002 luc, 0.02 µg pRL-CMV, 0.3 to 10 nM siRNAs, and 0.5 µl of LipofectAMINE 2000. The siRNAs (Eurogentec, Seraing, Belgium) were as followed: siRNA pGL3 luciferase; HMG-I(Y), GCAAGAACAAGGGCACAGCdTdT (sense), GCUGUGCCCUUGUUCUUGCdTdT (antisense); scramble, CACGUAGGUCAGAUCCAGCdTdT (sense), GCUGGAUCUGACCUACGUGdTdT (antisense). The day after transfection, the cells were exposed for 16 h to cytokines. Luciferase activities were assayed with the dual-luciferase reporter assay system (Promega). Luciferase activity values of the test plasmids were corrected for the value of the internal control pRL-CMV and expressed as relative luciferase activity. Treatment of the cells did not significantly affect pRL-CMV activity values, whose means ± SEM (n = 3, experiments performed in duplicates) were 1610 ± 349, 1370 ± 272, 1154 ± 107, and 820 ± 101 for control, IL-1ß-, IFN{gamma}- and IL-1ß+IFN{gamma}-treated RINm5F cells, respectively. For control and IL-1ß-treated ß-cells, values were 333 ± 72 and 357 ± 51, respectively (n = 6, experiments performed in duplicates).

EMSAs
Nuclear extracts were prepared from RINm5F cells as described (28). Nuclear proteins (4 µg) were preincubated with 1 µg poly(dIdC) or poly(dGdC), as indicated in the figure legends, in 20 µl containing 10 mM HEPES (pH 7.9), 50 mM KCl, 5 mM MgCl2, 0.05 mM EDTA, 0.5 mM dithiothreitol, 10% glycerol for 10 min at 0 C before addition of 50 molar excess of competing oligonucleotide, as indicated, and radiolabeled probe (40,000 cpm). The incubation was continued for 20 min at 0 C. Where indicated, 1 µl antibodies (Santa Cruz Biotechnology, Santa Cruz, CA) were added for 30 min at room temperature after or, in the case of anti-HMG-I(Y) antibody, before incubation with the probe. The samples were electrophoresed on 5% polyacrylamide gels in 25 mM Tris, 25 mM boric acid, and 0.5 mM EDTA. Oligonucleotides for EMSAs were as follows (upper strand shown): iNOS NF-{kappa}B (NF)-Oct, 5'-AGTACTGGGGACTCTCCCTTTGGGAACAGTGACTTTATGCAAAACAG; Oct consensus, 5'-TGTCGAATGCAAATCACTAGAA; NF-{kappa}B (NF) consensus, 5'-AGCTTCAGAGGGGACTTTCCGAGA; and activator protein-1 (AP-1), 5'-AGCGCTTGATGACTCAGCCGGAA. For band quantification, the autoradiogram was scanned and analyzed with the Biomax analysis software (Kodak, Rochester, NY). The highest value in each experiment was considered as 100.

RT-PCR analysis
Reverse transcription was performed on poly(A)+ RNA as described (7). Real-time quantitative PCR was performed on rat ß-cell cDNA using a LightCycler instrument (Roche Diagnostics, Mannheim, Germany) and the FastStart Master SYBR Green I (Roche Diagnostics) according to the manufacturer’s instructions. Each 20-µl reaction contained 2 µl reverse transcription reaction, 2 or 3 mM MgCl2 [for HMG-I(Y) or glyceraldehyde-3-phosphate dehydrogenase (GAPDH) amplification, respectively], 500 nM forward and reverse primers, and 2 µl FastStart SYBR Green I mix. The amplification was as follows: 95 C for 10 min, followed by 45 cycles at 95 C for 10 sec, 60 C for 10 sec, and 72 C for 14 sec. Primers were HMG-I(Y): 5'-CGAAGGGAAGCAAGAACAAG (forward) and 5'-GAGATGCCCTCCTCTTCCTC (reverse); and GAPDH: 5'-AGTTCAACGGCACAGTCAAG (forward) and 5'-TACTCAGCACCAGCATCACC (reverse). Serial dilutions of external standards consisting of the corresponding cDNA with known copy number were used as a standard curve. The values for HMG-I(Y) mRNA were normalized to the housekeeping gene GAPDH. Standard PCR was performed on RINm5F cell cDNA as described (7). Primers for HMG-I(Y) were 5'-GCTCAAAGTCCAGCCAGC (forward) and 5'-CTCCTGGGAGATGCCCTCC (reverse). Primers for GAPDH were described elsewhere (7). PCR was performed with 27 cycles at an annealing temperature of 60 or 58 C for HMG-I(Y) or GAPDH, respectively. The results were validated by real-time quantitative RT-PCR.

Western blotting analysis
HMG proteins and histone H1 were copurified by acid extraction with 5% perchloric acid from 40-µg proteins of RINm5F nuclear extracts as described (29). The proteins were resuspended in H2O, electrophoresed on sodium dodecyl sulfate-15% polyacrylamide gel, and transferred onto nitrocellulose membrane. Transferred proteins were stained with Ponceau solution and incubated with an anti-HMG-I(Y) antibody. Horseradish peroxidase-linked antirabbit IgG was used as a second antibody. Peroxidase activity was detected by enhanced chemiluminescence (Amersham Biosciences, Little Chalfont, UK). After scanning, the HMG-I(Y) and the Ponceau-stained histone H1 bands were quantified with the Biomax analysis software. The intensity values for HMG-I(Y) were corrected by the values for histone H1.

Statistical analysis
Results are given as means ± SEM. Multiple comparisons were performed by ANOVA, followed by group comparisons using the Student’s unpaired t test with correction by the Bonferroni method. Comparisons vs. control were performed using the Student’s paired t test.


   Results
Top
 Abstract
 Introduction
 Materials and Methods
 Results
Discussion
 References
 
Role of an Oct binding site in cytokine induction of the iNOS promoter
We have previously shown that the rat promoter region up to -1002 bp, containing a proximal NF-{kappa}B site (-106 to -97) and distal elements located between -965 and -894, namely a NF-{kappa}B site, GAS, and ISRE, is required for maximal induction by cytokines in insulin-producing cells (7). In this study, contribution to iNOS promoter induction by IL-1ß and IFN{gamma} of an Oct binding site, located 20 bp downstream from the proximal NF-{kappa}B site, was investigated by transfection of a construct containing the 1002 bp of the promoter in which the Oct binding site was mutated. In RINm5F cells, inactivation of the Oct site reduced IL-1ß and IL-1ß + IFN{gamma}-induced promoter activity by 80–85% (Fig. 1AGo). The Oct mutation did not affect basal promoter activity. Similar results were obtained in rat primary ß-cells where mutation of the Oct site resulted in a 5-fold decrease of the IL-1ß-induced promoter activity (Fig. 1BGo). These results suggest a critical role for factors binding to the Oct site in cytokine-induction of iNOS promoter activity in insulin-producing cells.



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  		FIG. 1. Effect of mutation of the Oct site on cytokine-induced iNOS promoter activity in RINm5F cells (A) and primary ß-cells (B). The cells were transfected with the promoter construct piNOS-1002 luc, wild-type (WT) or mutated in the Oct site (Oct mut) and either left untreated (control, C) or exposed for 16 h to IL-1ß, IFN{gamma}, or IL-1ß + IFN{gamma}. Relative luciferase activity values are the means ± SEM of three (A) or six (B) independent experiments. **, P < 0.01 vs. WT construct activity exposed to the same cytokine(s) (ANOVA).

 
Identification of factors binding to the proximal region of the iNOS promoter
To identify factors binding to the proximal part of the iNOS promoter, we performed EMSAs with a probe from nucleotides -111 to -65, spanning both the NF-{kappa}B and Oct sites (iNOS NF-Oct) and nuclear extracts from RINm5F cells untreated or treated with IL-1ß. Four complexes (a, b, c1, and c2) were detected (Fig. 2AGo). The strong complex a was constitutively expressed, whereas complex b appeared only in extracts from cells treated with IL-1ß for 30 min (Fig. 2AGo). The minor complexes c were detected at 4 and 8 h (Fig. 2AGo). Complex a was competed by an oligonucleotide containing a consensus octamer motif (Fig. 2BGo, lane 2) and supershifted by an anti-Oct1 antibody (lane 5). Complex b was competed by an oligonucleotide containing a consensus NF-{kappa}B binding site (lane 3) and affected by an anti-NF-{kappa}B p65 antibody (lane 6). A nonrelevant competing oligonucleotide had no effect (lane 4). These experiments thus identified binding of the octamer factor Oct1 and NF-{kappa}B to the iNOS promoter. Consistent with previous observations, NF-{kappa}B binding activity (complex b) was maximal at 30 min (25, 30), whereas Oct1 binding (complex a) was not affected by IL-1ß (present data). Similar binding of both factors to their respective sites was observed when probes containing the NF-{kappa}B site or the Oct site were used independently (data not shown).



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  		FIG. 2. Factors binding to the proximal region of the iNOS promoter. EMSAs were performed with an oligonucleotide spanning from nucleotides -111 to -65 and poly(dIdC) (used as non specific competitor). Arrows a, b, and c indicate specific complexes. A, The probe was incubated with nuclear extracts from RINm5F cells left untreated (control, C) or exposed to IL-1ß for 30 min to 8 h. F, Free probe. The figure is representative of two similar experiments. B, The probe was incubated with nuclear extracts from RINm5F cells exposed to IL-1ß for 30 min and with competing oligonucleotides (lanes 2–4) or specific antibodies (lanes 5 and 6) as indicated above the lanes. Arrowhead indicates supershifted complex. C, The probe was incubated with nuclear extracts from RINm5F cells left untreated (control, C) or treated with CHX, IL-1ß, or both for the indicated time points. The figure is representative of three similar experiments.

 
We have previously shown that the protein synthesis inhibitor CHX blocks IL-1ß-dependent iNOS mRNA expression in insulin-producing cells (26, 31). Thus, we evaluated whether this effect of CHX was secondary to an early decrease in cellular Oct1 content and consequent decrease in Oct1 binding to the iNOS promoter. As shown in Fig. 2CGo, CHX did not decrease DNA binding activity of Oct1 (complex a) and NF-{kappa}B (complex b) in RINm5F cells exposed to IL-1ß. The mean density values ± SEM for the Oct1 complex were as follows: control, 97 ± 2; IL-1ß 30 min, 96 ± 3; IL-1ß + CHX 30 min, 83 ± 13; IL-1ß 6 h, 80 ± 6; IL-1ß + CHX 6 h, 85 ± 8 (n = 3). Instead, exposure of the cells to IL-1ß + CHX for 6 h restored the NF-{kappa}B binding activity (complex b), which was barely detectable in cells exposed to IL-1ß alone for the same time period (Fig. 2CGo). Thus, the inhibition of iNOS mRNA expression by CHX cannot be attributed to a blocking of Oct1 or NF-{kappa}B DNA binding activities and should involve another mechanism.

HMG-I(Y) mRNA and protein expression and binding of HMG-I(Y) protein to the iNOS promoter in insulin-producing cells
In RASMCs, HMG-I(Y) participates in iNOS promoter regulation (16) and HMG-I(Y) mRNA and protein expressions are up-regulated by IL-1ß (17). In insulin-producing cells, IL-1ß-induction of iNOS mRNA content peaks at 6 h and progressively decreases over a 48-h period (32). To test whether HMG-I(Y) mRNA is similarly induced in these cells, we investigated by real-time quantitative RT-PCR the effects of IL-1ß and IFN{gamma} on HMG-I(Y) mRNA levels in primary rat ß-cells and RINm5F cells. In primary ß-cells, there was a slight increase in HMG-I(Y) mRNA content after 24 h exposure to IL-1ß, alone or in combination with IFN{gamma} (Fig. 3AGo). IFN{gamma} alone did not modify HMG-I(Y) mRNA content. Cytokines (IL-1ß or IL-1ß + IFN{gamma}) did not affect HMG-I(Y) mRNA content in RINm5F cells over a time-course period of 2–24 h exposure to cytokines as evaluated by standard RT-PCR and real-time quantitative RT-PCR (data not shown; n = 4). Similarly, HMG-I(Y) protein content was not modified in these cells by exposure to IL-1ß for up to 6 h (Fig. 3BGo). The density values for HMG-I(Y) band detected by Western blotting corrected for histone H1 from two independent experiments were as followed: control, 1.77–2.17; IL-1ß 30 min, 1.29–2.27; IL-1ß 6 h, 2.40–2.32.



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  		FIG. 3. Cytokine regulation of HMG-I(Y) mRNA and protein expression in insulin-producing cells. A, FACS-purified rat ß-cells were cultured for 6 or 24 h in control medium (C, no cytokine added) or with IL-1ß, IFN{gamma}, or IL-1ß + IFN{gamma}. HMG-I(Y) and GAPDH mRNA contents were analyzed by real-time quantitative RT-PCR. Values for HMG-I(Y) mRNA were corrected for GAPDH and normalized considering the maximum signal in each amplification as 100. The values are the means ± SEM of four independent experiments. **, P < 0.01 vs. control (ANOVA). B, RINm5F cells were cultured in control medium (C, no cytokine added) or with IL-1ß for 30 min or 6 h. Extracted HMG-I(Y) protein was analyzed by Western blotting. The blot is representative of two independent experiments.

 
To test whether HMG-I(Y) could account for the formation of some of the complexes binding the proximal iNOS promoter in EMSAs, nuclear extracts from RINm5F cells treated with IL-1ß ± CHX for 30 min or 6 h were incubated with the iNOS NF-Oct probe and poly(dGdC) used as a nonspecific competitor instead of poly(dIdC) because HMG-I(Y) binds preferentially to poly(dIdC) (29). Using these conditions, complexes a (containing Oct1), c1, and c2 were detected, but complex b due to NF-{kappa}B was masked by a nonspecific complex (Fig. 4AGo). The upper complexes c1 and c2 were maximally induced after 6 h of exposure to IL-1ß, and complex c1 was partially abolished by CHX treatment (Fig. 4AGo). To identify the proteins present in complexes c1 and c2, we next performed supershift experiments. Incubation of the RINm5F cell extracts with an anti-HMG-I(Y) antibody interfered with the formation of complex c1 at 6 h exposure to IL-1ß ± CHX (Fig. 4BGo), indicating the presence of HMG-I(Y) in this complex. Complexes a and c2 were not affected by the anti-HMG-I(Y) antibody in any condition. Anti-Oct1 and anti-NF-{kappa}B p65 antibodies disrupted both complex c1 at 30 min or 6 h exposure to IL-1ß ± CHX and complex c2 at 6 h exposure to IL-1ß ± CHX (Fig. 4BGo). This suggests an IL-1ß-induced complex formation among Oct1, NF-{kappa}B, and HMG-I(Y) to the iNOS promoter, whereas CHX reduced the contribution of HMG-I(Y) to complex binding to the promoter.



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  		FIG. 4. Detection of HMG-I(Y) in protein complexes binding to the proximal region of the iNOS promoter. EMSAs were performed with an oligonucleotide spanning from nucleotides -111 to -65 and poly(dGdC) as nonspecific competitor. Arrows a and c indicate specific complexes; ns, nonspecific complex. A, The probe was incubated with nuclear extracts from RINm5F cells left untreated (control, C) or treated with IL-1ß, CHX, or IL-1ß + CHX for the indicated time points. B, The same nuclear extracts as in A were preincubated with anti-Oct1, anti-HMG-I(Y), or anti-NF-{kappa}B p65 antibodies, as indicated above the lanes, before addition of the probe. Arrowheads indicate supershifted complexes. The figure is representative of two or three similar experiments.

 
Effect of HMG-I(Y) silencing on iNOS promoter activity
To assess a role for HMG-I(Y) in the induction of iNOS promoter activity by IL-1ß, we used RNA interference (33) to block the expression of HMG-I(Y). We first tested the feasibility of RNA interference in RINm5F cells by inhibiting luciferase expression in transient transfection. For this purpose, we cotransfected the cells with piNOS-1002 luc and increasing concentrations (up to 10 nM) of a pGL3 luciferase siRNA. We observed a complete inhibition of the IL-1ß-induced luciferase activity driven by the iNOS promoter with 3–10 nM siRNA. Even 0.3 nM was already effective to induce a partial silencing (Fig. 5Go). To block the expression of HMG-I(Y), we used the same range of HMG-I(Y) siRNA concentrations cotransfected with piNOS-1002 luc. A scrambled siRNA was used as a negative control. Addition of HMG-I(Y) siRNA in the range of 3–10 nM reduced by 50% IL-1ß-induced activity of the iNOS promoter, whereas the scrambled siRNA had no effect (Fig. 5Go), suggesting a role for HMG-I(Y) in maximal induction of the iNOS promoter activity by IL-1ß.



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  		FIG. 5. Effect of HMG-I(Y) silencing on iNOS promoter activity in RINm5F cells. The cells were cotransfected with the promoter construct piNOS-1002 luc and pGL3 luciferase (Luc), HMG-I(Y), or scrambled siRNA at concentrations ranging from 0 to 10 nM and exposed to IL-1ß for 16 h. Fold-induction of luciferase activity values, compared with control (no IL-1ß or siRNA added) values, are the means ± SEM of three or four independent experiments. SiRNAs have no effect on the promoter activity in control condition (data not shown). *, P < 0.05 and **, P < 0.01 vs. luciferase activity in the absence of siRNA (paired t test).

 

   Discussion
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
References
 
We show for the first time in the present study that a proximal Oct binding site is essential for induction of rat iNOS promoter activity in response to cytokines in primary ß-cells and an insulinoma cell line. Thus, transfection experiments performed in ß-cells and RINm5F cells indicate that mutation of this Oct site impairs by 80–85% promoter activity after exposure to IL-1ß or IL-1ß + IFN{gamma}. Supershift experiments with a probe containing both the proximal NF-{kappa}B and Oct binding sites identified, besides the IL-1ß-induced binding of NF-{kappa}B, constitutive binding of the ubiquitous transcription factor Oct1. We have previously shown that mutation of the proximal NF-{kappa}B binding site impairs by 50% the iNOS promoter activity induced by IL-1ß or IL-1ß + IFN{gamma} (7). The present data indicate that Oct1 binding is a crucial component for the induction of iNOS expression in pancreatic ß-cells.

IL-1ß-induced expression of iNOS mRNA in insulin-producing cells is CHX sensitive (26, 31). Nevertheless, neither Oct1 nor NF-{kappa}B binding activities were blocked by CHX (present data). Instead, NF-{kappa}B binding activity was enhanced and sustained over 6 h, probably because CHX inhibits de novo synthesis of the NF-{kappa}B inhibitor I{kappa}B, as reported in other cell types (34, 35). This suggests that CHX probably acts by blocking synthesis of other factor(s) involved in iNOS promoter activation in response to IL-1ß. Mouse macrophages treated with LPS contain a CHX-sensitive complex containing NF-{kappa}B, Oct1, and additional unidentified protein(s) required for cooperation between NF-{kappa}B and Oct1 and subsequent stimulation of iNOS transcription (10, 36). We also observed such a complex in RINm5F cells exposed to IL-1ß for 6 h (present data), correlating with the time of maximal induction of iNOS mRNA expression (32). HMG-I(Y) is a candidate protein to interact with NF-{kappa}B and Oct1 to form the ternary complex described above. This architectural protein binds to the minor groove of AT-rich sequences and facilitates the assembly of functional DNA-protein complexes, named enhanceosomes, by modifying the conformation of DNA (18). HMG-I(Y) expression is induced more than 4-fold by IL-1ß in primary RASMCs (17, 37) and by LPS in macrophages (37) and enhances iNOS promoter transactivation by NF-{kappa}B through binding to the Oct site (16).

We presently observed that HMG-I(Y) mRNA content is only slightly increased by IL-1ß in rat primary ß-cells, whereas HMG-I(Y) mRNA expression is constitutive in the insulinoma cell line RINm5F. This is consistent with the fact that proliferating and transformed cells often express high constitutive levels of HMG-I(Y), compared with normal tissues (38, 39). These RT-PCR experiments confirmed the lack of cytokine-induction of HMG-I(Y) expression observed by previous microarray analyses of cytokine-treated rat primary ß-cells (40) and INS-1 cells (41). Thus, whereas our supershift experiments indicate the presence of HMG-I(Y), as well as Oct1 and NF-{kappa}B, in the CHX-sensitive complex detected with the proximal iNOS promoter region, the marginal induction of HMG-I(Y) mRNA and protein expression by IL-1ß in insulin-producing cells suggests that formation of this complex is not due to an increase in HMG-I(Y) protein level. HMG-I(Y) undergoes posttranslational modifications in vivo in response to different stimuli that may modulate DNA binding affinity (18). For instance, depending on which lysine residue of HMG-I(Y) protein is acetylated, the IFNß enhanceosome is either stabilized or disrupted, providing a mechanism of control for the transcription rate of the IFN-ß gene (42). The level of phosphorylation of HMG-I(Y) also influences the ability of this protein to interact with DNA (43). Such modifications induced by IL-1ß might affect HMG-I(Y) binding to the iNOS promoter and/or its ability to interact with Oct1 and NF-{kappa}B. Further studies are required to determine whether the HMG-I(Y) protein is posttranslationally modified in response to IL-1ß.

HMG-I(Y) silencing by RNA interference reduced by 50% IL-1ß-induced iNOS promoter activity (present data), suggesting that HMG-I(Y) is necessary for maximal transactivation of the iNOS promoter in insulin-producing cells. Our gel shift experiments indicate that HMG-I(Y) is not required for Oct1 or NF-{kappa}B binding to the iNOS proximal promoter because binding of these two factors was detected in nuclear extracts from RINm5F cells left untreated or exposed to IL-1ß for 30 min, experimental situations in which HMG-I(Y) is not detected in the complexes, and their binding was not enhanced after 6 h exposure to IL-1ß, an experimental situation in which HMG-I(Y) was detected. This is in contrast to the situation of the IFNß (44) and the insulin receptor (45) promoters in which HMG-I(Y) facilitates the binding of transcription factors to DNA. These studies, however, were performed using recombinant proteins. Thus, it is conceivable that the role for endogenous HMG-I(Y) is to facilitate interaction and functional cooperation between Oct1 and NF-{kappa}B to efficiently transactivate the iNOS promoter.

In conclusion, the present results have broadened our understanding of iNOS gene regulation in insulin-producing cells, suggesting that both Oct1 and HMG-I(Y) are crucial elements for iNOS expression.


   Acknowledgments
 
The technical assistance of R. Leeman and G. Vandenbroeck is gratefully acknowledged.


   Footnotes
 
Present address for S.T.: Department MBW/FBI, Universitaire Campus D, Limburgs Universitair Centrum, 3590-Diepenbeek, Belgium.

This work was supported by grants from the Juvenile Diabetes Foundation International and the Fonds National de la Recherche Scientifique, Belgium, and has been conducted in collaboration with and supported by the JDRF Center for Prevention of ß Cell Destruction in Europe under Grant 4-2002-457.

Abbreviations: CHX, Cycloheximide; CMV, cytomegalovirus; FACS, fluorescence-activated cell sorter; GAPDH, glyceraldehyde-3-phosphate dehydrogenase; GAS, {gamma}-interferon activated site; HMG, high mobility group; IFN, interferon; iNOS, inducible form of nitric oxide synthase; ISRE, IFN-stimulated response element; LPS, lipopolysaccharide; NF-{kappa}B, nuclear factor-{kappa}B; Oct, octamer; RASMC, rat aortic smooth muscle cell; siRNA, small interfering RNA.

Received September 10, 2003.

Accepted for publication November 10, 2003.


   References
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 References
 

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