Identification of Transacting Factors Responsible for the Tissue-specific Expression of Human Glucose Transporter Type 2 Isoform Gene - Ji-Young Cha; Hail Kim; Kyung‐Sup Kim; Man‐Wook Hur; Yong‐Ho Ahn (2000)
We investigated transacting factors binding to the cis-element important in tissue-specific expression of the human glucose transporter type 2 isoform (GLUT2) gene. By transient transfection assay, we determined that the 227-base pair fragment upstream of the ATG start site contained promoter activity and that the region from +87 to +132 (site C) was responsible for tissue-specific expression. DNase I footprinting and electrophoretic mobility shift assay indicated that site C contained one binding site for hepatocyte nuclear factor 1 (HNF1) and two binding sites for HNF3. The mutations at positions +101 and +103, which are considered to be critical in binding HNF1 and HNF3, resulted in a 53% decrease in promoter activity, whereas the mutation of the proximal HNF3 binding site (+115 and +117) reduced promoter activity by 28%. The mutations of these four sites resulted in marked decrease (70%) in promoter activity as well as diminished bindings of HNF1 and HNF3. A to G mutation, which causes conversion of the HNF1 and HNF3 binding sequence to the NF-Y binding site, resulted in a 22% decrease in promoter activity. We identified that both HNF1 and HNF3 function as transcriptional activators in GLUT2 gene expression. Coexpression of the pGL+74 (+74 to +301) construct with the HNF1α and HNF3β expression vectors in NIH 3T3 cells showed the synergistic effect on GLUT2 promoter activity compared with the expression of HNF1α, HNF3β, or a combination of HNF1β and HNF3β. These data suggest that HNF1α and HNF3β may be the most important players in the tissue-specific expression of the human GLUT2 gene. We investigated transacting factors binding to the cis-element important in tissue-specific expression of the human glucose transporter type 2 isoform (GLUT2) gene. By transient transfection assay, we determined that the 227-base pair fragment upstream of the ATG start site contained promoter activity and that the region from +87 to +132 (site C) was responsible for tissue-specific expression. DNase I footprinting and electrophoretic mobility shift assay indicated that site C contained one binding site for hepatocyte nuclear factor 1 (HNF1) and two binding sites for HNF3. The mutations at positions +101 and +103, which are considered to be critical in binding HNF1 and HNF3, resulted in a 53% decrease in promoter activity, whereas the mutation of the proximal HNF3 binding site (+115 and +117) reduced promoter activity by 28%. The mutations of these four sites resulted in marked decrease (70%) in promoter activity as well as diminished bindings of HNF1 and HNF3. A to G mutation, which causes conversion of the HNF1 and HNF3 binding sequence to the NF-Y binding site, resulted in a 22% decrease in promoter activity. We identified that both HNF1 and HNF3 function as transcriptional activators in GLUT2 gene expression. Coexpression of the pGL+74 (+74 to +301) construct with the HNF1α and HNF3β expression vectors in NIH 3T3 cells showed the synergistic effect on GLUT2 promoter activity compared with the expression of HNF1α, HNF3β, or a combination of HNF1β and HNF3β. These data suggest that HNF1α and HNF3β may be the most important players in the tissue-specific expression of the human GLUT2 gene. glucose transporter type 2 isoform hepatocyte nuclear factor nuclear factor-Y electrophoretic mobility shift assay base pair(s) The glucose transporter type 2 isoform (GLUT2)1 is a transmembrane protein involved in maintaining glucose homeostasis by facilitating the entry of glucose through the plasma membrane. GLUT2 is predominantly expressed in hepatocytes, β-cells of the pancreas, and to a lesser extent in absorptive epithelia of the intestine and kidney (12.Tronche F. Ringeisen F. Blumenfeld M. Yaniv M. Pontoglio M. J. Mol. Biol. 1997; 266: 231-245Crossref PubMed Scopus (156) Google Scholar). GLUT2 plays an important role in absorbing glucose after meals in hepatocyte (3.Axelrod J.L. Pilch P.F. Biochemistry. 1983; 22: 2222-2227Crossref PubMed Scopus (60) Google Scholar) as well as in insulin secretion in pancreatic β-cells (4.Orci L. Ravazzola M. Baetens D. Imnan L. Amhersdt M. Peterson R.G. Newgard C.B. Johnson J.H. Unger R.H. Proc. Natl. Acad. Sci. U. S. A. 1990; 87: 9953-9957Crossref PubMed Scopus (126) Google Scholar). The GLUT2 mRNA level is influenced by the blood glucose level and insulin (5.Chen L. Alam T. Johnson J.H. Hughes S. Newgard C.B. Unger R.H. Proc. Natl. Acad. Sci. U. S. A. 1990; 87: 4088-4092Crossref PubMed Scopus (91) Google Scholar, 6.Postic C. Burcelin R. Rencurel F. Pegorier J.P. Lorzeau M. Girard J. Biochem. J. 1993; 293: 119-124Crossref PubMed Scopus (72) Google Scholar). The GLUT2 expression between hepatocytes and pancreatic β-cells is contradictory in diabetic animal model; GLUT2 expression is increased in hepatocytes, whereas it is decreased in β-cells (7.Rencurel F. Waeber G. Antoine B. Rocchiccioli F. Maulard P. Girard J. Leturque A. Biochem. J. 1996; 314: 903-909Crossref PubMed Scopus (95) Google Scholar, 8.Thorens B. Wu Y.J. Leahy J.L. Weir G.C. J. Clin. Invest. 1992; 90: 77-85Crossref PubMed Scopus (142) Google Scholar, 9.Ohneda M. Johnson J.H Inman L.R. Chen L. Suzuki K. Goto Y. Alam T. Ravazzola M. Orci L. Unger R.H. Diabetes. 1993; 42: 1065-1072Crossref PubMed Scopus (79) Google Scholar). Although the transcriptional regulation of GLUT2 expression under various metabolic states has been studied, the transacting factors regulating the tissue-specific expression of the GLUT2 gene are relatively unknown. So far, Pdx-1, a pancreatic homeodomain transcription factor, has been proposed to regulate the expression of the GLUT2 gene (10.Waeber G. Thompson N. Nicod P. Bonny C. Mol. Endocrinol. 1996; 10: 1327-1334Crossref PubMed Scopus (326) Google Scholar). However, this factor is not expressed in hepatocytes. Thus, liver-specific transcription factors, such as hepatocyte nuclear factor 1 (HNF1) (11.De Simone V. Cortese R. Curr. Opin. Cell Biol. 1991; 3: 960-965Crossref PubMed Scopus (60) Google Scholar, 12.Tronche F. Ringeisen F. Blumenfeld M. Yaniv M. Pontoglio M. J. Mol. Biol. 1997; 266: 231-245Crossref PubMed Scopus (156) Google Scholar), CCAAT/enhancer-binding protein (13.Kim J.W. Ahn Y.H. Biochem. J. 1998; 336: 83-90Crossref PubMed Scopus (31) Google Scholar), HNF3 (14.Duncan S.A. Navas M.A. Dufort D. Rossant J. Stoffel M. Science. 1998; 281: 692-695Crossref PubMed Scopus (296) Google Scholar) and HNF4 (15.Stoffel M. Duncan S.A. Proc. Natl. Acad. Sci. U. S. A. 1997; 94: 13209-13214Crossref PubMed Scopus (341) Google Scholar) have been expected to regulate GLUT2 gene expression. None of these transcription factors are found exclusively in hepatocytes; rather, they are all expressed, although to a lesser degree, in other tissues. Thus, the tissue specificity and levels of gene expression depend on the concerted actions of several factors working together. HNF1 is a homeodomain protein that plays a key role in the liver-specific expression of many genes during differentiation and development (16.Miura N. Tanaka K. Nucleic Acids Res. 1993; 21: 3731-3736Crossref PubMed Scopus (74) Google Scholar). HNF1 is required for the expression of GLUT2 and other liver-specific genes such as albumin, α1-antitrypsin, and fibrinogen (17.Wang H. Maechler P. Hagenfeldt K.A. Wollheim C.B. EMBO J. 1998; 17: 6701-6713Crossref PubMed Google Scholar, 18.Courtois G. Morgan J. Campbell L.A. Fourel G. Crabtree G.R. Science. 1987; 238: 688-692Crossref PubMed Scopus (282) Google Scholar, 19.Lai E. Darnell J.E. Trends Biochem. Sci. 1991; 16: 427-430Abstract Full Text PDF PubMed Scopus (112) Google Scholar). HNF1α associates with the highly related HNF1β (vHNF1) to form homo- or heterodimers (20.Mendel D.B. Crabtree G.R. J. Biol. Chem. 1991; 266: 677-680Abstract Full Text PDF PubMed Google Scholar). The expression of the HNF1α dominant negative form resulted in reduction of GLUT2 gene expression (21.Wang H. Maechler P. Hagenfeldt K.A. Wollheim C.B. EMBO J. 1998; 17: 6701-6713Crossref PubMed Google Scholar). Mutation of HNF1β resulted in decreased transcription of GLUT2 and was ultimately related to familial type 2 diabetes mellitus (22.Tomura H. Nishigori H. Sho K. Yamagata K. Inoue I. Takeda J. J. Biol. Chem. 1999; 274: 12975-12978Abstract Full Text Full Text PDF PubMed Scopus (40) Google Scholar). The human GLUT2 promoter has a binding site for HNF1 at the −1030 bp region. However, deletion of the HNF1 binding site did not abolish GLUT2 promoter activity mediated by HNF1β (22.Tomura H. Nishigori H. Sho K. Yamagata K. Inoue I. Takeda J. J. Biol. Chem. 1999; 274: 12975-12978Abstract Full Text Full Text PDF PubMed Scopus (40) Google Scholar). Thus, it should be determined whether other cis-elements for HNF1 exist in the human GLUT2 promoter or whether HNF1 indirectly regulates GLUT2 gene expression by other transcription factors. HNF3, a member of the forkhead winged helix family, has been known to have a function in liver and gut development (23.Lai E. Prezioso V.R. Tao W.F. Chen W.S. Darnell J.E. Genes Dev. 1991; 5: 416-427Crossref PubMed Scopus (436) Google Scholar). HNF3α, -3β, and -3γ bind to the same DNA sequence with different affinities. It was shown that HNF3 could activate the albumin gene expression by repositioning the nucleosomes in the albumin enhancer (24.McPherson C.E. Shim E.Y. Friedman D.S. Zaret K.S. Cell. 1993; 75: 387-398Abstract Full Text PDF PubMed Scopus (284) Google Scholar, 25.Shim E.Y. Woodcock C. Zaret K.S. Genes Dev. 1998; 12: 5-10Crossref PubMed Scopus (101) Google Scholar). In this study, we showed that human GLUT2 expression was regulated directly by HNF1 and HNF3. HNF1α and HNF3β synergistically activated transcription in NIH 3T3 cells. Thus, we propose that the combined action of the two liver-enriched factors, HNF1α and HNF3β, plays a critical role in the tissue-specific expression of the GLUT2 gene. Furthermore, we showed that the promoter activity of the mutG construct containing the +103A→G mutation, which was found in a Korean non-insulin-dependent diabetes mellitus patient, was in part compensated by the binding of NF-Y instead of HNF1 or HNF3. The promoter region (−294 to +301 bp) of the human GLUT2 gene was cloned into the SmaI site of pGL3-basic reporter vector (Promega, Madison, WI) to generate pGL-294. Serial deletions from the 5′-end of pGL3-294 were created by polymerase chain reaction. The mutations were generated by polymerase chain reaction-based site-directed mutagenesis (Stratagene, La Jolla, CA; for sequences, see below). For pGL+74m3, the mutant pGL+74m1 was mutated again with the oligonucleotide m2. The sequences of constructs were confirmed by DNA sequencing. pRSV-HNF1 and pRSV-vHNF1 were gifts from Dr. M. Yaniv. pGem-HNF3α and pGem-HNF3β were provided by Dr. R. H. Costa. Nuclear extracts from liver of male Sprague-Dawley rats or cell lines were prepared as described by Gorski et al. (26.Gorski K. Carneiro M. Schibler U. Cell. 1986; 47: 767-776Abstract Full Text PDF PubMed Scopus (973) Google Scholar) or Dignam et al. (27.Dignam J.D. Methods Enzymol. 1990; 182: 194-203Crossref PubMed Scopus (223) Google Scholar). Protein concentration was determined according to Bradford (28.Bradford M.M. Anal. Biochem. 1976; 72: 248-254Crossref PubMed Scopus (217544) Google Scholar). The extracts were frozen in aliquots and stored at −70 °C. DNA fragments (−294 to +16 bp or +46 to +301 bp) were labeled in each strand and purified as follows. The promoter region was amplified by polymerase chain reaction to obtain −294 to +16 and +48 to +301 promoter fragments, and the resulting fragments were subcloned into the EcoRV site of pT7Blue (R) vector. The promoter fragments were isolated by double digestion with EcoRI (BamHI) and PstI (HindIII) to obtain 5′-overhanging ends and 3′-overhanging ends. The fragments were labeled using Klenow fragment and [α-32P]dATP and then purified from agarose gel electrophoresis by a gel extraction kit (Qiagen, Valencia, CA). DNA-protein binding reactions were performed using 50,000 cpm (approximately 1 ng) of probe per reaction in a solution containing 10 mm HEPES, pH 7.9, 60 mm KCl, 7% glycerol, 1 mm EDTA, 1 mm dithiothreitol, 2 μg of poly(dI-dC), and the indicated amount of nuclear extracts. After a 20-min incubation on ice, 5 μl of DNase I freshly diluted in a solution containing 10 mm HEPES, pH 7.9, 60 mmKCl, 25 mm MgCl2, 5 mmCaCl2, and 7% glycerol was added to the reaction and kept at room temperature for 2 min. Dilution folds of DNase I were ranged from 1:200 to 1:2000 of stock (10 units/μl) depending on the amount of protein present in the reaction. Digestion reactions were stopped by adding 80 μl of a stop solution containing 20 mm Tris-Cl, pH 8.0, 20 mm EDTA, 250 mm NaCl, 0.5% SDS, 4 μg of yeast tRNA, and 10 μg of proteinase K. The samples were incubated for 30 min at 45 °C, extracted with phenol/chloroform, ethanol-precipitated, and resuspended in formamide dye. The samples were resolved in 6% polyacrylamide, 7 m urea sequencing gel. The protected regions were mapped with reference to the migration of Maxam-Gilbert A + G sequencing products (29.Sambrook J. Fritsch E.F. Maniatis T. Molecular Cloning: A Laboratory Manual. 2nd Ed. Cold Spring Harbor Laboratory, Cold Spring Harbor, NY1989: 13.78-13.101Google Scholar). Probes for gel shift assays were labeled with32P in the presence of [γ-32P]ATP and T4 polynucleotide kinase. Labeled double-stranded oligonucleotides were prepared by mixing a 5-fold molar excess of the complementary single-stranded DNAs in 50 mm NaCl, heating to 90 °C for 5 min, and then cooling to room temperature. The oligonucleotides used in these assays were as follows (the mutated nucleotides are underlined): C-1, 5′-GGCAAAGCACTTATTGATTAGATTCCCATC-3′; C-2, 5′-GATTCCCATCAATATTCAGCTGCCGC-3′; mutC, 5′- GGCAAAGCACTTATTGCTTAGATTCCCATC-3′; mutG, 5′- GGCAAAGCACTTATTGTTAGATTCCCATC-3′; m1, 5′- GGCAAAGCACTTATCGCTTAGATTCCCATC-3′; m2, 5′-GATTCCCAGCCATATTCAGCTGCCGC-3′. The oligonucleotides for HNF1 (30.Vaulont S. Puzenat N. Levrat F. Cognet M. Kahn A. Raymondjean M. J. Mol. Biol. 1989; 209: 205-219Crossref PubMed Scopus (113) Google Scholar), HNF3 (31.Costa R. Grayson D. Darnell J.E. Mol. Cell. Biol. 1989; 9: 1415-1425Crossref PubMed Scopus (428) Google Scholar), HNF4 (32.Sladek F.M. Zhong W. Tal E. Darnell J.E. Genes Dev. 1990; 4: 2353-2365Crossref PubMed Scopus (861) Google Scholar), and NF-Y (33.Boam D.S.W. Clark A.R. Docherty K. J. Biol. Chem. 1990; 265: 8285-8296Abstract Full Text PDF PubMed Google Scholar) were synthesized, and Oct-1 (5′-TGTCGAATGCAAATCACTAGAA-3′) and Sp1 consensus oligonucleotides (5′-ATTCGATCGGGGCGGGGCGAGC-3′) were purchased from Promega. The labeled probe (50,000 cpm) was combined with nuclear proteins in 25 mm Tris/HCl, pH 7.4, 80 mm KCl, 0.1 mm EDTA, 1 mmdithiothreitol, and 10% (v/v) glycerol. The nonspecific competitor, 1.5 μg of poly(dI-dC), was added to each binding reaction. In the supershift assays, anti-HNF1α and/or anti-HNF1β antibodies were purchased from Santa Cruz Biotechnology, Inc. (Santa Cruz, CA), and the antibody against the B subunit of NF-Y was provided by Dr. R. Mantovani. Antibodies were added to the binding reaction 10 min before the addition of the labeled probe. Binding reaction mixtures were incubated for 20 min on ice and resolved on a nondenatured (5% w/v) acrylamide gel (29:1 (w/w) acrylamide/bisacrylamide) in 0.5× TBE at 4 °C. For competition assays, a 50–200-fold molar excess of various unlabeled competitor DNAs was added to the reaction mixture prior to the addition of the labeled probe. The dried gels were exposed to x-ray film at −70 °C with an intensifying screen. Cells used in this experiment were maintained as monolayer cultures and grown in appropriate media. Plasmid DNAs were purified on Qiagen Midiprep kit columns (Qiagen) at least twice independently. Cells were plated in six-well tissue culture plates at a density of 1 × 106 cells/well in 2 ml of medium. After a 20-h attachment period, transfections were performed with LipofectAMINE PLUS reagent (Life Technologies, Inc.), following the manufacturer's protocol. Briefly, 0.5 μg of each construct of GLUT2 promoter; 0.1 μg of pCMV-β-galactosidase, with or without 20 ng of HNF1α, HNF1β, HNF3α, and/or HNF3β; and 4 μl of PLUS reagent and 2 μl of LipofectAMINE in 200 μl of OPTI-MEM I (Life Technologies, Inc.) medium lacking serum were mixed and added to cells. In order to transfect constant amounts of DNA, sample DNAs were supplemented with an appropriate control vector. After 3 h, the medium containing the LipofectAMINE-DNA complex was removed and replaced by appropriate medium (containing serum and antibiotics). Cells were then cultured further for 48 h and harvested in reporter lysis buffer (Promega). The lysed cells were centrifuged to remove cell debris, and the supernatant was collected. Luciferase assays were conducted with 10 μl of cell extracts and 50 μl of luciferase assay reagent (Promega). β-Galactosidase activity was determined with 10 μl of cell extract and 190 μl of assay reagent containingO-nitrophenol-β-d-galactopyranoside in a colorimetric assay. Luciferase data were expressed as luciferase activity corrected by β-galactosidase activity in the cell lysate. Each transfection was performed in triplicate and repeated 3–5 times. All transfection studies were performed in 3–5 separate experiments, where triplicate dishes were transfected. The data were represented as mean ± S.D. Statistical analysis was carried out using Microsoft Excel®(Microsoft). To identify the important region in GLUT2 transcription, a series of sequential 5′-deletions of the GLUT2 294-bp 5′-flanking region contiguous to the promoter and 301-bp 5′-untranslated region were generated and fused to a luciferase reporter gene (Fig.1 A). These constructs were transiently transfected in HIT-T15 cells, which express GLUT2, or NIH 3T3 cells, which do not express GLUT2, and luciferase activity was determined. Luciferase activity for all constructs was very high in HIT-T15 but low in NIH 3T3 cells (Fig. 1 A). These differences in the activities indicated that the 5′-flanking region of the GLUT2 promoter might be responsible for tissue-specific expression. In HIT-T15 cells, luciferase activities for the pGL+34 construct remained high (Fig. 1 A). However, deletion down to +73 resulted in a marked increase, whereas further deletion to +188 caused a drastic decrease in luciferase activity. These results indicated the presence of negative and positive regulatory elements in the 5′-untranslated region. To generalize that the 5′-flanking region of human GLUT2 promoter was responsible for tissue-specific expression, we have transfected the reporter constructs to MIN6, HepG2 (GLUT2-expressing) cells and HeLa (GLUT2-nonexpressing) cells (Fig.1 B). As expected, the promoter activity was high in MIN6 and HepG2 cells, whereas HeLa cells showed very low promoter activity. Since the promoter activities remained high down to the transcription start site (Fig. 1 A), two promoter fragments (from −294 to +16 or from +46 to +301) were analyzed. Eight sites were protected with rat liver nuclear extract. These sites were designated as site H (−294/−281), site G (−274/−247), site F (230/−196), site E (−180/−161), site D (−154/−135), site C (+87/+132), site B (+190/+216), and site A (+229/+257) (Fig.2). Computer-based analysis of the sequence revealed several motifs resembling consensus sequences of many known nuclear factors (Fig.3 A). Site C contained three putative transacting factor-binding sites for HNF1 and two HNF3s in reverse orientation. A sequence corresponding to the consensus Oct-1-binding site was also present in site C. Site C and site A contained consensus sequences for HNFs that were mainly expressed in the liver, pancreas, and kidney. The deletion of the promoter region from +74 to +188 bp greatly decreased the promoter activity, suggesting that the factors binding to the footprinting region of site C may act as positive regulators.Figure 3Summary of the consensus sequences in the human GLUT2 promoter. A, the transcription initiation site is designated +1 (35.Lichtsteiner S. Schibler U. Cell. 1989; 57: 1179-1187Abstract Full Text PDF PubMed Scopus (160) Google Scholar). The boxed areasrepresent the protected regions of human GLUT2 promoter identified by the DNase I footprinting experiment. The known consensus sequences areunderlined. B, comparison of potential HNF1 and HNF3 binding sites in human, mouse, and rat GLUT2 promoter sequences.View Large Image Figure ViewerDownload Hi-res image Download (PPT) As shown in Fig. 3 B, site C is well conserved between species. The +87/+132 region of the human GLUT2 promoter matched well with the −131/−86 region of mouse (34.Bonny C. Thompson N. Nicod P. Waeber G. Mol. Endocrinol. 1995; 9: 1413-1426PubMed Google Scholar) and the +492/+541 region of rat (13.Kim J.W. Ahn Y.H. Biochem. J. 1998; 336: 83-90Crossref PubMed Scopus (31) Google Scholar). A computer search suggested that these regions are potential binding sites for HNFs in these species. Also, it is interesting to note that site B of human GLUT2 promoter region was also conserved both in mouse (34.Bonny C. Thompson N. Nicod P. Waeber G. Mol. Endocrinol. 1995; 9: 1413-1426PubMed Google Scholar) and rat (13.Kim J.W. Ahn Y.H. Biochem. J. 1998; 336: 83-90Crossref PubMed Scopus (31) Google Scholar), although the transcription factors binding to the site B are unknown. On the basis of DNase I footprinting and computer-based analysis, we attempted to identify the nuclear factor(s) binding to site C by EMSA. Site C was divided into two overlapping halves, site C-1 and site C-2 (Fig. 4 A). Site C-1 (+87 to +116) included putative HNF1 and HNF3 binding sites. The HNF1 binding site covers the HNF3 binding site. Site C-2 (+107 to +132) included putative HNF3 and Oct-1 consensus sites. In site C, two HNF3 binding sites were inversely oriented. For site C-1, multiple bands were observed by the addition of rat liver nuclear extracts (Fig.4 B, lanes 1 and 2). These bands were specific, since they competed by an excess of unlabeled oligonucleotide site C-1 (lane 3).Band b was selectively abolished by an oligonucleotide HNF1 site of the l-pyruvate kinase promoter (lane 4). Band e decreased in the presence of an excess of the unlabeled oligonucleotide corresponding to HNF3 sequences in the transthyretin promoter (lane 5). The addition of the HNF1 and HNF3 consensus oligonucleotides abolished bands b ande (lane 6). The unrelated HNF4 oligonucleotide did not affect protein binding (lane 7). Band c represented the complex containing Oct-1 that was competed by Oct-1 consensus oligonucleotide (data not shown). Supershift experiments using anti-HNF1α and/or β antibodies revealed that HNF1α and HNF1β were binding (lanes 8 and 9), although HNF1α is the predominant member of the HNF1 family in hepatocytes (35.Lichtsteiner S. Schibler U. Cell. 1989; 57: 1179-1187Abstract Full Text PDF PubMed Scopus (160) Google Scholar). When both antibodies against HNF1α and HNF1β were used together,band b was completely supershifted (lane 10). Gel retardation using oligonucleotide HNF1 as a probe gave band b, and it was shifted by anti-HNF1α antibody (lanes 12 and13). The identity of band f, the fastest migrating band, was not determined in this study. It's interesting to note that an A to G mutation at +103 in site C-1, which was found in the course of screening GLUT2 promoter mutations of non-insulin-dependent diabetes mellitus patients, resulted in the appearance of band d (lane 11 and Fig. 4 C). To identify the transcription factor binding to this mutated region, we compared this sequence to a number of transcription factor consensus sequences and found that the sequence was identical to the NF-Y consensus sequences. This was confirmed by a cold competitor and NF-Y consensus oligonucleotide (Fig.4 C, lane 3 and lanes 5 and 6, respectively). Site C-1 abolishedbands e and f, but not band d (lane 4). The identity of band d was further confirmed by an anti-NF-Y B subunit antibody (lane 7). HNF3 cold oligonucleotide abolishedband e (lane 8). Gel retardation assay of a site C-2 probe revealed two shifted bands (Fig. 4 D, lane 2). These bands were competed out by an excess of unlabeled probe (lane 3). The lower one had the same mobility as band e in the site C-1 probe and was competed by oligonucleotide HNF3 (lane 5). The slow mobility band (band c) represented a complex containing Oct-1, because it was completely competed out by the consensus sequence for Oct-1 (lane 6). The consensus oligonucleotide for Oct-1 also abolished band d. The unrelated Sp1 oligonucleotide did not compete out any protein binding (lane 7). Next, we determined the effect of mutations on the binding affinity of HNF1, HNF3, and Oct-1 with the promoter. The HNF-binding sites in site C were disrupted by site-directed mutagenesis, and the resulting changes in binding activity were determined (Fig.5). Using oligonucleotide C-1 as a probe, the retarded bands disappeared with a 50-fold molar excess of cold competitors (Fig. 5 A, lanes 3 and4). The bands b and e were slightly competed out by the addition of a 200-fold molar excess of the unlabeled mutC (A to C mutation at +103) or mutG (A to G mutation at +103) (lanes 5–8). However, the mutant m1 (T to C mutation at +101 and A to G mutation at +103) showed decreased competition when compared with a self-competitor, mutC or mutG, suggesting that the base changes at +101 and +103 in site C-1 may decrease HNF1 and HNF3 binding (lanes 9 and10). In the case of the site C-2 probe, Oct-1 and HNF3 bindings were efficiently competed out by C-2 and partially competed out by C-1, which contained HNF3 and Oct-1 binding sites too (Fig. 5 B,lanes 2–4 and lane 7). Mutant oligonucleotide m2 (T to G mutation at +115 and A to C mutation at +117) could not compete out protein binding even in the presence of a 200-fold molar excess of C-2 probe (lanes 5 and6). To correlate the decreased HNF1 and HNF3 binding with GLUT2 promoter activity, we introduced the mutC, mutG, m1, m2, and m3 substitutions at their respective positions in the pGL+74 (+74 to +301) construct. The effects of the mutations within the promoter were tested by transient transfection assay in HIT-T15 cells (Fig.6). The C or G mutation of the +103 base reduced GLUT2 promoter activity to 70 and 78% of wild type pGL+74 expression, respectively. Two-base substitutions of site C-1 (pGL+74m1) resulted in a 53% (p < 0.05) decrease in activity, whereas two-base mutations of site C-2 (pGL+74m2) resulted in a 28% decrease in GLUT2 promoter activity. The pGL+74m3 mutant, in which both site C-1 and C-2 were mutated, reduced transcriptional activity by 70% (p < 0.05). These data showed that the HNF1 and HNF3 binding sites in site C act as positive cis-elements in HIT-T15 cells and that site C-1 is more important than site C-2 in the expression of the GLUT2 gene. Finally, we examined whether these transcription factors were able to transactivate the human GLUT2 gene in a non-HNF-expressing system. As shown in Fig.7, the HNF1 and HNF3 were not expressed in NIH 3T3 cells, whereas they were expressed in HIT-T15 cells as well as in hepatocytes. Therefore, we overexpressed the subtypes of HNF1 and HNF3 alone or in various combinations in NIH 3T3 cells to investigate their effects on GLUT2 gene expression. Except for HNF1β, the subtypes of HNFs showed positive transacting effects on the GLUT2 promoter. HNF1α, HNF3α, and HNF3β resulted in 1.9-, 2.0-, and 3.6-fold increases in promoter activity, respectively. Coexpression of HNF1α and HNF3β proteins showed the highest activation (8.7-fold) of promoter activity, whereas other coexpressions of HNF1α and HNF3α or of HNF1β and HNF3β showed activation effects by 3.1- and 3.4-fold, respectively (Fig. 8).Figure 8Cooperative activation of HNF1α and HNF3β on human GLUT2 promoter activity. NIH 3T3 cells were transiently transfected with 0.5 μg of pGL+74 construct and 20 ng of HNF1 and/or HNF3 expression vectors, as indicated. In all cases, equal amounts of DNA (0.64 μg) were transfected into cells by adding control expression vectors devoid of insert DNA. Cells were harvested 48 h after transfection, and the luciferase activity was measured. Luciferase activities were normalized on the basis of β-galactosidase activity produced by the co-transfected control plasmid, pCMV-β-galactosidase. Results are the mean ± S.D. of three independent experiments in triplicate.View Large Image Figure ViewerDownload Hi-res image Download (PPT) Although the GLUT2 gene is mainly expressed in liver and pancreatic β-cells, the transcription factors and cis-elements responsible for the tissue-specific expression are largely unknown. PDX-1, a pancreatic homeodomain transcription factor, has been showed to activate the expression of the mouse GLUT2 gene (10.Waeber G. Thompson N. Nicod P. Bonny C. Mol. Endocrinol. 1996; 10: 1327-1334Crossref PubMed Scopus (326) Google Scholar). However, its expression is not correlated with reduced GLUT2 gene expression in diabetes, which suggests that transacting factors other than PDX-1 may play some roles in GLUT2 gene expression. In this study, we showed that the 5′-untranslated region of the human GLUT2 gene plays a critical role in transcriptional activation, and specifically, site C (+87 to +132) is responsible for tissue-specific expression of the human GLUT2 gene. Also, a serial deletion study revealed that deletion down to the +189 region abolished GLUT2 promoter activity, which suggested that multiple transcription initiation sites exist in this 5′-untranslated region (36.Takeda J. Kayano T. Fukumoto H. Bell G.I. Diabetes. 1993; 42: 773-777Crossref PubMed Scopus (59) Google Scholar). In EMSA, site C-1 (+87 to +116), which contained overlapping HNF3 and HNF1 binding sites, provided the HNF1/probe and HNF3/probe complexes (Fig. 4 B). Competition experiments indicated that HNF1 and HNF3 bound to site C-1 in a mutually exclusive manner, although HNF1 binding seemed to be preferential. In supershift experiments, most of the HNF1 subtypes present in hepatocytes or HIT-T15 cells may be the HNF1α form rather than HNF1β (Fig. 4 B). This result suggests that most of HNF1 acts as HNF1α homodimeric forms in the cells where GLUT2 is mainly expressed. In contrast to hepatocytes or HIT-T15 cells, HNF1β is the predominant form in the kidney (37.Mendel D.B. Hansen L.P. Graves M.K. Conley P.B. Crabtree G.R. Genes Dev. 1991; 5: 1042-1056Crossref PubMed Scopus (251) Google Scholar), where GLUT2 is expressed at a lower level. Thus, it could be assumed that the differential expression of these HNF1 subtypes may be one of the factors regulating tissue-specific GLUT2 expression. We have compared the sequences of site C in human promoter with that of mouse and rat. As shown in Fig. 3 B, these sequences were well conserved in mouse (GTII region; Ref. 34.Bonny C. Thompson N. Nicod P. Waeber G. Mol. Endocrinol. 1995; 9: 1413-1426PubMed Google Scholar) and rat (13.Kim J.W. Ahn Y.H. Biochem. J. 1998; 336: 83-90Crossref PubMed Scopus (31) Google Scholar) promoters and may be real HNF1 and HNF3 binding sites. Bonny et al. (34.Bonny C. Thompson N. Nicod P. Waeber G. Mol. Endocrinol. 1995; 9: 1413-1426PubMed Google Scholar) suggested that the GTII region might be a cis-element having a stimulatory role on GLUT2. Thus far, transacting factors binding to this region in mice have not been known. Based on our experiment and the homology relationship between human and mouse sequences of the site C and GTII region, it is assumed that HNFs might bind to the GTII region and be working as positive regulators. Transfection studies using HIT-T15, MIN6, HepG2, NIH 3T3, and HeLa cells revealed the differences in promoter activities (Fig. 1,A and B). These activities were well correlated with the status of HNF1 and HNF3 mRNA expression (data not shown). From these observations, it is suggested that GLUT2 promoter could be activated by the expression of HNF1 and HNF3 regardless of the origins or types of cells. It is possible that the subtypes of HNFs may directly or indirectly interact with each other in controlling GLUT2 gene expression, because site C contains both HNF1 and HNF3 binding sites that are located closely together. Previous studies have suggested that an important role of HNF3 could be to cooperate with other factors bound to closely oriented cis-elements (38.Nitsch D. Schutz G. Mol. Cell. Biol. 1993; 13: 4494-4504Crossref PubMed Scopus (38) Google Scholar, 39.Espinas M.L. Roux J. Pictet R. Grange T. Mol. Cell. Biol. 1995; 15: 5346-5354Crossref PubMed Scopus (49) Google Scholar, 40.Jackson D.A. Rowader K.E. Stevens K. Jiang C. Milos P. Zaret K.S. Mol. Cell. Biol. 1993; 13: 2401-2410Crossref PubMed Scopus (134) Google Scholar, 41.Braun H. Suske G. J. Biol. Chem. 1998; 273: 9821-9828Abstract Full Text Full Text PDF PubMed Scopus (58) Google Scholar). In the case of the aldolase B gene, HNF3 competitively antagonized HNF1-dependent transactivation (42.Gregori C. Kahn A. Pichard A.L. Nucleic Acids Res. 1994; 22: 1242-1246Crossref PubMed Scopus (33) Google Scholar). Here, we report another type of HNF1 and HNF3 relationship in activating the GLUT2 promoter, in which we assume there is cooperative activation. Recent papers have shown that HNF3 may be involved in the transition of chromatin from an inactive to an active conformation (24.McPherson C.E. Shim E.Y. Friedman D.S. Zaret K.S. Cell. 1993; 75: 387-398Abstract Full Text PDF PubMed Scopus (284) Google Scholar, 25.Shim E.Y. Woodcock C. Zaret K.S. Genes Dev. 1998; 12: 5-10Crossref PubMed Scopus (101) Google Scholar). In the GLUT2 promoter, binding HNF3 to site C-2 may contribute to a higher affinity for HNF1, perhaps by opening the chromatin. Interestingly, in the course of screening non-insulin-dependent diabetes mellitus patients who may have possible mutations at the upstream region of the GLUT2 gene, we found A to G mutation in the +103 region, which belongs to site C-1. By this mutation, the consensus sequence for HNF1 and HNF3 is changed to that of NF-Y. Thus, it was expected that this mutation definitely caused a decrease in promoter activity. However, functional assay of this mutant showed only a 22% decrease in the promoter activity, whereas the m1 construct, which has lost its ability to bind HNF1 and HNF3, showed a 50% decrease in the promoter activity (Fig. 6). These results can be explained by assuming that NF-Y binding instead of HNFs binding to site C-1 might compensate the promoter activity, because NF-Y is known to act as a positive transacting factor in many other genes (43.Maity S.N. De Crombrugghe B. Trends Biochem. Sci. 1998; 23: 174-178Abstract Full Text Full Text PDF PubMed Scopus (312) Google Scholar). In conclusion, we identified that the liver-enriched transcription factors HNF1 and HNF3 are two of the main stimulatory trans-acting factors, and they may be responsible for tissue-specific expression of the GLUT2 gene. Furthermore, our results emphasize a new type of functional interplay between HNF1 and HNF3 to drive human GLUT2 gene expression. We thank Dr. M. Yaniv for providing pRSV-HNF1 and pRSV-vHNF1, Dr. R. H. Costa for providing pGem-HNF3α and pGem-HNF3β, Dr. R. Mantovani for providing the antibody against B subunit of NF-Y, and Dr. Jun-ichi Miyazaki for providing MIN6 beta cells.