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Laboratory of Endocrine Pathophysiology and
1 Laboratory of Cellular Biology of Hypertension and Molecular Medicine, Department of Medicine, Centre Hospitalier de l’Université de Montréal, Montreal, Quebec, Canada
(Requests for offprints should be addressed to A Lacroix, Hôtel-Dieu du Centre Hospitalier de l’Université de Montréal, 3850 St Urbain Street, Montreal, Quebec, Canada, H2W 1T7; Email: andre.lacroix{at}umontreal.ca)
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Abstract |
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 Top Abstract IntroductionExperimental proceduresResultsDiscussionReferences |
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Introduction |
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TopAbstractIntroductionExperimental proceduresResultsDiscussionReferences |
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Hamet et al.(1987) were the first to identify food-dependent cortisol production in a patient with unilateral adrenal adenoma and Cushing’s syndrome. Further studies demonstrated the involvement of ectopic GIP-R expression in the adrenal tissues of patients with GIP-dependent Cushing’s syndrome and bilateral macronodular hyperplasia (Lacroix et al. 1992, Reznik et al. 1992). GIP-dependent Cushing’s syndrome has now been identified in at least 17 patients with adrenocorticotropin (ACTH)-independent macro-nodular adrenal hyperplasia (Chabre et al. 1998, Lebrethon et al. 1998, N’Diaye et al. 1999, Pralong et al. 1999, Croughs et al. 2000, Gerl et al. 2000, Lacroix et al. 2001, Groussin et al. 2002, Bertagna et al. 2003) and seven with unilateral adenoma (Hamet et al. 1987, De Herder et al. 1996, Chabre et al. 1998, Luton et al. 1998, Tsagarakis et al. 2001). Sequence analysis of the full-length cDNA of GIP-dependent adrenal tissues revealed no GIP-R mutation in the affected adenomas or macronodular hyperplasia of these patients (N’Diaye et al. 1998). Recent analysis also showed that no mutation was present in the promoter region of the human (h) GIP-R gene in the adrenal tissues of patients with GIP-dependent Cushing’s syndrome compared with normal controls (Antonini et al. 2004). This suggests that abnormalities in transcription factors or co-factors could be responsible for the ectopic GIP-R expression in these patients.
The recently cloned rat promoter does not have TATA or CAAT boxes but contains potential consensus sequences for Sp1, CREB and Oct-1 transcriptions factors (Boylan et al. 1999). The molecular mechanisms responsible for human GIP-R expression are still unknown. To identify some of the transcription factors involved in the expression of the hGIP-R gene, we partially characterized its proximal promoter region.
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Experimental procedures |
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A membrane containing mRNA from a large number of adult and fetal tissues was purchased from Clontech (BD Bioscience, Palo Alto, CA, USA). An hGIP-R cDNA probe corresponding to nucleotides 387–1195 and a ubiquitin probe (Clontech) were labeled with [
-32P]dCTP, using the Klenow fragment (Invitrogen, Carlsbad, CA, USA). Hybridization was carried out under conditions described in the protocol from Clontech. Detection was conducted with a Phospho-Imager after a 3 day exposure for the GIP-R probe and a 2 h exposure for the ubiquitin probe.
Functional analysis of the hGIP-R gene promoter
Deleted fragments of the hGIP-R gene promoter were amplified by genomic PCR. Mutated fragments were prepared by two rounds of PCR as described by Chen & Przbyla (1994). The fragments were sub-cloned into a pGL-3 Basic vector (Promega Biosciences, San Luis Obispo, CA, USA) and the constructs were verified by sequencing. Human gastric HGT-1 (kindly gifted by Dr C L Laboisse, INSERM, Nantes, France) and mouse adrenocortical Y1 (American Type Culture Collection, Mannassas, VA, USA) cell lines were plated in 60 mm dishes in DMEM+ (Invitrogen) supplemented with 10% fetal bovine serum (BioMedia, Drummondville, QC, Canada) and containing 100 µg/ml penicillin/streptomycin (Sigma-Aldrich). The different promoter reporter gene constructs (2.5 µg) were transiently co-transfected with an RSV ß-gal-containing plasmid (2.5 µg) by the calcium phosphate precipitation method. The media were changed 24 h after transfections. ß-Gal and luciferase activities were measured 48 h after transfection. Luciferase activity was normalized with respect to ß-gal activity.
Nuclear extracts and electrophoretic mobility shift assay (EMSA)
Nuclear extracts were prepared according to the method of Andrews & Faller (1991). The probes (see Fig. 3
) were labeled with [-32P]dCTP, using the Klenow fragment. Binding reactions were prepared in a final volume of 30 µl (24 mM Hepes, 3 mM MgCl2, 70 mM KCl, 24% glycerol, 3 mM dithiothreitol (DTT), 600 µg/µl BSA, 1 µg/µl poly(dI-dc), 5 µg nuclear extracts, 150 000 c.p.m. labeled probe) and incubated for 30 min at room temperature. For competition assays, unlabeled specific or non-specific (5' TCTGGAAGGGGATCCGTGTAC ACAGGAAGTGACAATTTTC 3') fragments were added simultaneously with the labeled fragment. When antibody (Santa Cruz Biotechnology, Santa Cruz, CA, USA) was included in the reaction, nuclear extracts and antibody were pre-incubated for 15 min at room temperature before addition of the probe. Complexes were separated on non-denaturing 4% polyacrylamide gel in 0.5xTris–borate–EDTA buffer. Detection was undertaken with a PhosphorImager after a 24 h exposure.
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Adrenal tissues were collected for this study from patients with Cushing’s syndrome associated with either GIP-dependent Cushing’s syndrome with confirmed ectopic GIP-R expression (one patient with unilateral adenoma and six patients with bilateral macronodular adrenal hyperplasia), non-GIP-dependent unilateral adrenal adenoma (seven patients), ACTH-dependent Cushing’s disease (seven patients) or adrenal carcinoma (two patients), and from three normal adrenal of patients undergoing radical nephrectomy. Commercially available RNA (from a pool of 52 adrenals and from two other normal individuals) was also used as normal control (Clontech and Ambion (Austin, TX, USA) respectively). Informed consent was obtained from each patient and this study was approved by the institutional ethics committee. Tissue specimens obtained at surgery were immediately frozen in liquid nitrogen and stored at –80 °C until RNA extraction. Total RNA was extracted with Trizol reagent (Invitrogen). RNA (3 µg) was reverse-transcribed using M-MLV RT (Invitrogen) in a final volume of 20 µl (1 x first-strand buffer; 10 mM DTT; 0.5 mM dNTPs; 0.01 µg/µl random primers; 1 U/µl RNAse out; 1 U/µl M-MLV RT). PCR was performed using TAQ DNA polymerase (Invitrogen) in a final volume of 50 µl (1x TAQ DNA polymerase buffer; 0.2 µM specific sense and antisense probes; 0.2 µM 18S probe (Ambion); 0.2 mM dNTPs; 2 mM MgCl2; 2 µl cDNA; 0.05 U/µl TAQ DNA polymerase). The linear range for PCR amplification was determined as described in the Quantum RNA 18S internal standards protocol from Ambion. The number of cycles used for each PCR amplification are indicated in the figure legends. PCR products were separated in a 2% agarose gel and detected using a PhosphorImager. Data are expressed as means ± S.D. Student’s t-test was used in the statistical analysis and a P< 0.05 was considered statistically significant.
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Results |
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Distribution of GIP-R mRNA in human tissues was determined by a multiple-expression tissue assay with a blot containing mRNA from 65 adult and seven fetal tissues. In adult tissues (Fig. 1A, lanes A–I), maximal expression was found in the pancreas and trachea. GIP-R was also detected, at a lower level, in the brain, heart, gut, spleen, thymus, blood cells, lung and kidney, whereas it was not seen in the liver, placenta, testis, uterus and adrenal gland. In fetal tissues (Fig. 1A, lane J), the messenger for GIP-R was found in the lung, heart and kidney, but not in the brain, liver, spleen and thymus. Specificity of the probe was confirmed by the absence of signal in the control samples (Fig. 1A, lanes K1–K6). Some of these results were also confirmed by semi-quantitative RT-PCR (Fig. 1B). The expected bands for GIP-R were clearly present in pancreas, brain, trachea, kidney and small intestine, but were only faintly detected in adrenal gland.
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The sequences for the hGIP-R mRNA and promoter are available from GeneBank (NM000164 and AC006132 [GenBank] respectively). The exact localization of the transcription start site is still unknown but we used the 5'-end of the published cDNA sequence as the –1 nucleotide to identify the constructs.
To identify regions of interest, we transfected deleted fragments of the hGIP-R promoter up to 2 kb in HGT-1 and Y1 cells expressing respectively high and low levels of GIP-R mRNA (Fig. 2A). Transfection of HGT-1 cells with deleted fragments from –2068 to –595 bp showed an increase of ~5 fold of the control vector in relative luciferase activity whereas, transfection of Y1 cells with these fragments failed to elevate the luciferase activity higher than the control vector. These results suggest that different transcription factors could be able to bind that region of the promoter depending of the cell lines. Deletion of the region between –595 and –473 bp led to a decrease in relative luciferase activity to levels comparable with the control luciferase vector in HGT-1 cells. This observation suggests the presence of a potential response element for an activating factor between –595 and –473 bp. In Y1 cells, no modification of the relative luciferase activity was found. Further deletion until –336 bp induced significant elevation of relative luciferase activity in HGT-1 and Y1 cells. The binding of an inhibitor between –473 and –336 bp is probably responsible for that effect. Our hypothesis to explain the effect observed between –336 and –595 bp is that the loss of inhibition with the construct GIPR–595/+73 is due to the binding of an activator that interferes with the factors capable of interaction with the inhibitory binding site. Relative luciferase activity was maintained with fragments up to –100 bp, but was not detected with the –49 bp fragment. To ensure that the differences observed in the promoter activity in HGT-1 and Y1 cell lines was not due to the species used, we also transfected some of the constructs in the human HeLa cell line that expresses a low level of GIP-R mRNA. Relative luciferase activity was similar in Y1 and HeLa cell lines.
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The proximal promoter region of the hGIP-R gene revealed a GC-rich region without TATA or CAAT boxes, but contains multiple GC (–237/–242, –77/–82, –65/–70, –48/–53) or GT (–223/–229, –150/–156) boxes (Fig. 3
). The sites are potential response elements for transcription factor Sp1.
To better characterize this region of the promoter, we transfected constructs that contained an increasing number of GC/GT boxes in HGT-1 cell lines (Fig. 2B). We also transfected constructs with punctual mutations within the consensus sequences (Fig. 2C). The short 49 bp fragment of the hGIP-R promoter, which does not contain any consensus for transcription factors, did not confer transcriptional activity compared with the control. Minimal basal luciferase activity was observed with the construct GIP-R–66/+73 that includes the GC-4 box. We also observed that single mutation of GC-4 led to the abolition of transcriptional activity of the promoter. Taken together, these results suggest that the GC-4 site is essential but not sufficient to confer high luciferase activity. Maximal luciferase activity is detected with the addition of the GC-3 box (construct GIP-R–75/+73). Mutation of the GC-3 binding site also led to the abolition of transcriptional activity of the promoter showing that GC-3 is also necessary to confer luciferase activity. The presence of other GC and GT boxes caused only a small change in the promoter activity. Surprisingly, an inactivating mutation of GC-1, GC-2, GT-1 and GT-2 increased the luciferase activity of the promoter. Interactions between protein complexes could be an explanation of these two discordant observations.
EMSA
To identify proteins capable of binding to the GC/GT boxes, we performed EMSAs. Nuclear extracts from HGT-1 cells were incubated with DNA fragments containing one of the GC or GT boxes. The resulting complexes were resolved on non-denaturing polyacrylamide gel (Fig. 4). We observed the formation of four complexes in the presence of the GT-1 probe (Fig. 4D, lane 1) and GC-4 probe (data not shown), five complexes with the GC-2 probe (Fig. 4B, lane 1) and six complexes with the GC-1 (Fig. 4A, lane 1), GC-3 (Fig. 4C, lane 1) and GT-2 probes (Fig. 4E, lane 1). The specificity of binding was demonstrated by competition assay. The complexes were displaced by an increasing amount of unlabeled probe (data not shown). A 100-fold excess of unlabeled specific probe completely abolished the DNA–protein interactions (lane 2). A 100-fold excess of an unlabeled non-specific probe had no effect (lane 3) on the complexes. GC/GT boxes are potential binding sites for the Sp1 transcription factor. To determine if Sp1 was capable of binding to these probes, we incubated nuclear extracts with Sp1 antibody (lane 4). A supershifted band was produced with a parallel decrease in the abundance of slowly migrating C1 complex using the GC-1, GC-3 and GT-2 probes, whereas the complex C1 disappeared when the Sp1 antibody was pre-incubated with nuclear extracts prior to addition of the GC-2 and GT-1 probes. Transcription factor Sp3 also binds GC/GT-rich element (lane 5). Pre-incubation with Sp3 antibody blocked the formation of the C2 and C3 complexes for the GC-2, GC-3 and GT-1 probes, whereas only the C2 complex was lost in the presence of the GT-2 probe. Pre-incubation of the nuclear extracts with the Sp3 antibody prior to incubation with the GC-1 probe leads to loss of C2 and C4 complexes. Sp1 and Sp3 antibodies were unable to displace the complexes observed in the presence of the GC-4 probe. Pre-incubation of the nuclear extracts with pre-immune serum had no effect on migration of the complexes showing the specificity for the interactions between the antibodies and the shifted complexes (lane 6). Incubating recombinant Sp1 protein in the presence of the wild-type GIP-R promoter probes leads to the formation of only one complex that corresponds to the complex C1 shifted by the Sp1 antibody (data not shown). These data confirm the presence of the Sp1 transcription factor in the slowly migrating complex.
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To identify whether Sp transcription factor family genes are regulated in adrenal tissues of patients with Cushing’s syndrome, we performed further analysis by gene array of samples from patients with macronodular adrenal hyperplasia previously described (Bourdeau et al. 2004). We found overexpression of the transcription factor Sp3 and the co-factor CRSP9 in adrenal tissues from three patients with GIP-dependent Cushing’s syndrome compared with a pool of normal adrenal tissues. These preliminary results were extended by semi-quantitative RT-PCR in a larger number of adrenal tissues from patients with GIP-dependent Cushing’s syndrome, non-GIP-dependent adrenal cortisol-secreting tumors, ACTH-dependent Cushing’s disease and controls. We found a 2-fold increase in the expression of Sp3 and CRSP9 in all seven cases of GIP-dependent Cushing’s syndrome compared with a normal control (Fig. 5A, Fig 6A). We also observed a 1.5-fold increase in the expression of Sp3 and CRSP9 in four out of seven cases of Cushing’s disease (Fig. 5B, Fig. 6B). In non-GIP-dependent cortisol-secreting adrenal adenoma, Sp3 mRNA was increased 1.3-fold in three out of seven patients (Fig. 5C), while the level of CRSP9 mRNA was not changed (Fig. 6C). No significant difference was observed in two adrenal carcinomas (Fig. 5D, Fig. 6D). GIP-R was not expressed in non-GIP-dependent Cushing’s syndrome patients overexpressing CRSP9 and/or Sp3 (Fig. 7).
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Discussion |
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GIP-R has been identified and shown to be coupled to steroidogenesis in rat adrenocortical cells (Mazzocchi et al. 1999). In this study, we show that GIP-R mRNA is not or very weakly expressed in the normal human adult adrenal gland. The faint band detected after PCR amplification is not efficiently coupled to steroidogenesis (Lacroix et al. 1992) and may reflect the presence of GIP-R in endothelial cells rather than in adrenocortical cells (Zhong et al. 2000). This finding is consistent with previous reports demonstrating that GIP-R is expressed in adrenal adenomas or hyperplasias in GIP-dependent Cushing’s syndrome, but not in patients with non-GIP-dependent Cushing’s syndrome or in adrenal tissues from normal controls (Lacroix et al. 1992, Reznik et al. 1992, De Herder et al. 1996, Chabre et al. 1998, Lebrethon et al. 1998, Luton et al. 1998, N’Diaye et al. 1998, 1999).
Sequence analysis of the full-length cDNA or promoter region of the hGIP-R gene revealed no mutation in the affected unilateral adenomas or bilateral macronodular hyperplasias (N’Diaye et al. 1998, Antonini et al. 2004), suggesting that abnormalities in transcription factors or co-factors could be responsible for the aberrant GIP-R expression in these patients. GIP-R belongs to the family of secretin/glucagon/vasoactive intestinal polypeptide (VIP) receptors, as do receptors for glucagon-like peptide-1 (GLP-1) and parathyroid hormone (PTH). In recent years, it has been shown that the promoter region of this family of genes is TATA-less and contains a GC-rich region capable of binding to Sp1 transcription factors which are involved in the activation of their transcription (McCuaig et al. 1994, Sreedharan et al. 1995, Lankat-Buttgereit & Göke 1997, Buggy et al. 1995, Wildhage et al. 1999). Cloning of the rat GIP-R gene and its promoter revealed that its promoter was TATA-less and GC-rich, suggesting a mode of regulation similar to that described for other members of the family of secretin/glucagon/VIP receptors (Boylan et al. 1999). To better understand the regulation of the expression of hGIP-R gene in normal cells, we first characterized its promoter region. Analysis of the DNA sequence 350 bp upstream to the 5'-end of GIP-R cDNA revealed a GC-rich region containing no TATA or CAAT boxes. We identified six consensus sequences for the Sp1 transcription factors in this region and found them to be functionally important in regulating luciferase reporter gene activity in luciferase assays. Using EMSAs, we confirmed that Sp1 and Sp3 are capable of binding to these sites.
Characterization of the promoter up to 2 kb indicates others regions of potential interest for the regulation of the hGIP-R gene. Deletion of the region between –595 and –473 bp decreased relative luciferase activity to a level similar to the control, suggesting the presence of an activating factor in the region. We also observed that deleted fragments from –2 kb to –473 bp failed to elevate relative luciferase activity in Y1 cells, while activity was found in HGT-1. The difference between the relative luciferase activity may indicate that different transcription factors could be able to bind that region of the promoter depending of the cell lines. Further studies will be necessary to identify other response elements that could be important for the specific expression of the hGIP-R gene.
We also found that Sp3 and CRSP9, but not Sp1, are overexpressed in the adrenal tissues of all GIP-dependent Cushing’s syndrome patients studied as compared with normal adrenal. CRSP9 is an important regulator of Sp1 transcription factor (Ryu et al. 1999) and its overexpression could stimulate the promoter of hGIP-R by increasing the activity of Sp1. By contrast, Sp3 is well known to bind DNA with similar specificities and affinities to Sp1. However, while Sp1 acts only as an activator, Sp3 may also exert a repressor activity (Majello et al. 1997). As Sp3 and CRSP9 were also overexpressed in the adrenals of some patients with non-GIP-dependent adenomas and with Cushing’s disease, who do not express the GIP-R, we conclude that the overexpression of these transcription factors is not sufficient to explain the ectopic expression of GIP-R in food-dependent Cushing’s syndrome.
In a recent report, GIP-R was found to be overexpressed in the adrenals of five patients with Cushing’s disease and in one with primary pigmented nodular adrenal disease, suggesting that chronic ACTH stimulation or constitutive activation of the ACTH receptor signaling pathway may modulate the expression of GIP-R (Swords et al. 2005). Our study does not confirm this hypothesis as we failed to identify significant expression of GIP-R above normal in the adrenals of seven patients with Cushing’s disease and in seven patients with unilateral adrenal adenoma. This is consistent with a previous report in one patient with Cushing’s disease (N’Diaye et al. 1998) and in two patients with ectopic ACTH secretion (Chabre et al. 1998).
In conclusion, the hGIP-R gene promoter is TATA-less and contains multiple Sp1/Sp3 binding sites, which appear to be involved in the cellular expression of the receptor. This is similar to previous studies of other members of this family of seven trans-membrane hormone receptors such as VIP, glucagon, PTH and GLP-1 receptors. Further studies will be necessary to identify the tissue-specific transcription factors that are involved in the normal and ectopic expression of hGIPR resulting in GIP-dependent Cushing’s syndrome.
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Acknowledgements |
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References |
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Antonini SR, N’Diaye N, Baldacchino V, Hamet P, Tremblay J & Lacroix A 2004 Analysis of the putative regulatory region of the gastric inhibitory polypeptide receptor gene in food-dependent Cushing’s syndrome. Journal of Steroid Biochemistry and Molecular Biology 91 171–177.[CrossRef][Medline]
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Bourdeau I, Antonini S, Lacroix A, Kirschner LS, Lorang D, Libutti SK & Stratakis CA 2004 Gene array analysis of macronodular adrenal hyperplasias confirms clinical heterogeneity and identifies several genes as molecular mediators. Oncogene 23 1575–1585.[CrossRef][ISI][Medline]
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Received 6 March 2005
Accepted 18 April 2005
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