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Intestine-specific activity of the human guanylyl cyclase C promoter is regulated by Cdx2
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Intestine-Specific Activity of the Human Guanylyl Cyclase C Promoter Is Regulated by Cdx2 JASON PARK, STEPHANIE SCHULZ, and SCOTT A. WALDMAN Division of Clinical Pharmacology, Departments of Medicine and Biochemistry and Molecular Pharmacology, Thomas Jefferson University, Philadelphia, Pennsylvania
Background & Aims: The heat-stable enterotoxin receptor, guanylyl cyclase C, exhibits an intestine-specific pattern of expression. The aim of this study was to identify the transcriptional activator that mediates intestine-specific expression of guanylyl cyclase C. Methods: Fragments of the promoter were assayed to isolate regions directing intestine-specific gene activation. Deoxyribonuclease I footprinting was used to identify a site of intestine-specific protection. Electrophoretic mobility shift assays (EMSAs) and supershift analyses were used to characterize the protein that bound to the protected site. The protected site was mutated to analyze its role in promoter activity. Results: Reporter gene assays revealed that intestine-specific expression of guanylyl cyclase C is directed by the proximal promoter. Deoxyribonuclease I footprinting identified a specific site in the proximal promoter that exhibited intestine-specific protection. EMSAs and supershift analyses revealed that the transcription factor Cdx2 bound to an intestine-specific site of protection. Mutation of the Cdx2-protected site of the promoter eliminated binding of Cdx2 and reduced reporter gene activity to the level of extraintestinal cells. Conclusions: These data show that Cdx2 and its consensus-binding site in the promoter are required for intestine-specific expression of the guanylyl cyclase C gene.
uanylyl cyclase C (GC-C), the receptor for the Escherichia coli heat-stable enterotoxin (STa), catalyzes the conversion of guanosine triphosphate to guanosine 38,58-monophosphate, which initiates a biochemical cascade culminating in the opening of the cystic fibrosis transmembrane conductance regulator, a chloride channel. Efflux of chloride from enterocytes drives intestinal fluid accumulation and results in secretory diarrhea. The intestinal peptides guanylin and uroguanylin are putative endogenous ligands of GC-C, although they bind to and activate GC-C with a lower potency than STa.1,2 Conservation of GC-C throughout the vertebrate subphylum implies that it serves an important physiologic role that remains undefined.3,4 However, targeted disruption of the GC-C gene had no deleterious effects in
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mice, and in fact conferred resistance to STa-induced diarrhea.5,6 In humans, GC-C is expressed exclusively in epithelial cells lining the small and large intestines and in tumors derived from those cells.7,8 Messenger RNA encoding GC-C is localized in crypt and villous cells in the small intestine and in differentiated enterocytes in crypts of the large intestine.9,10 Similar to other intestine-specific genes that are predominately expressed in the small intestine, the level of expression of GC-C decreases along the rostrocaudal axis.11 Tissue specificity of GC-C transcription is directed by the evolutionarily conserved regulatory sequence 1118 to 2257 relative to the transcription start site.12,13 Indeed, GC-C transcription is dependent on a promoter element at 229 to 246 that binds hepatocyte nuclear factor 4a (HNF-4a).13 Although HNF-4a is required for GC-C transcription in the intestine, this transactivating protein is also expressed in liver, kidney, and pancreas. However, GC-C is not expressed in these organs.7,14,15 This wide distribution of HNF-4a expression suggests that additional factors are responsible for expression of GC-C in the intestine. This study identified a site in the promoter of GC-C, 275 to 283, that includes a consensus sequence for binding the intestine-specific homeodomain protein, Cdx2. This site interacts with a protein expressed by intestinal, but not extraintestinal, cells that is functionally, physically, and antigenically similar to Cdx2. This site also interacts specifically with Cdx2 obtained by in vitro transcription and translation. Additionally, this Cdx2 consensus-binding site is required for intestinespecific transcriptional activation of the GC-C promoter. Abbreviations used in this paper: BSA, bovine serum albumin; DMEM, Dulbecco’s modified Eagle medium; EMSA, electrophoretic mobility shift assay; GC-C, guanylyl cyclase C; HNF-4a, hepatocyte nuclear factor 4a; NE, nuclear extract; SDS, sodium dodecyl sulfate; STa, heat-stable enterotoxin; TNT, transcribed and translated. r 2000 by the American Gastroenterological Association 0016-5085/00/$10.00 doi:10.1053/gast.2000.8520
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Materials and Methods Genomic Library Screening and Sequencing The GC-C gene 58 regulatory region was cloned from a lFIXII human genomic library (Stratagene, La Jolla, CA). The library was screened by hybridization with a probe specific for exon 1 of the GC-C complementary DNA. A 2.8-kilobase (kb) XbaI fragment that included 2 kb upstream of the start site of transcription was subcloned into Bluescript KS (Stratagene). All constructs were generated from this Bluescript/human GC-C gene construct. The nucleic acid sequence of each construct was confirmed by BigDye terminator reaction chemistry for sequence analysis on the Applied Biosystems model 377 DNA sequencing system (Perkin-Elmer Applied Biosystems, Foster City, CA; Kimmel Cancer Institute Nucleic Acid Facility, Thomas Jefferson University, Philadelphia, PA).
Reporter Gene Constructs Fragments 2835 to 1117, 2257 to 1117, 2129 to 1117, and 246 to 1117, relative to the start site of transcription, were isolated from Bluescript KS constructs by digestion with selected restriction endonucleases.12 These fragments were blunt-ended and ligated into the EcoRV site of Bluescript KS. Inserts were excised from Bluescript KS with SmaI and KpnI and ligated into the pGL3-Basic Luciferase Vector (Promega, Madison, WI). The pGL3 Control Vector, containing an SV40 promoter with enhancers, was used as a positive control. Mutations were created in the 2835 to 1117 pGL3 construct using the polymerase chain reaction–based Ex-site Mutation Kit (Stratagene). Deletion constructs were created using primers flanking the sites of interest. The FP1 ‘‘CCC’’ mutant was created using the phosphorylated primers: 58GCCCATAGCTCTGACCTTTCTG38 and 58AGAGAGATTAGCTGGGCCTCACCC38.
Cell Culture and Transfection All cell lines were obtained from American Type Culture Collection (Rockville, MD). T84 cells were grown in Dulbecco’s modified Eagle medium (DMEM)/F12 (Life Technologies), Caco2 cells in DMEM (Life Technologies, Rockville, MD), HepG2 and HS766T cells in DMEM High Glucose (Cellgro, Herndon, VA), and HeLa cells in minimum essential medium Eagle (MEM) with glutamine (Life Technologies). All cell lines were maintained at 37°C in a 5% CO2/95% air atmosphere and passaged every 4 days. Assays of reporter gene activity were conducted with cells seeded in 6-well plates at either 5.0 3 105 (T84, Caco2, and HeLa) or 1.0 3 106 cells per well (HepG2 and HS766T). Cells were incubated overnight, washed once with phosphate-buffered saline, and supplemented with fresh media before transfection. Plasmids purified with the Qiafilter Kit (Qiagen, Valencia, CA) were transfected into cells with the nonliposomal lipid transfection reagent Effectene (Stratagene). All cell lines were cotransfected with 0.4 µg of firefly luciferase experimental reporter
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constructs, modified from pGL3-Basic, and 0.1 µg of the Renilla luciferase control reporter, pRL-TK, driven by a viral thymidine kinase promoter (Promega). Cells were incubated with transfection complexes for 24 hours, rinsed with phosphate-buffered saline, then supplemented with appropriate media and incubated for a further 24 hours. After a total of 48 hours, cells were lysed and assayed using the protocol and materials in the Dual-Luciferase Reporter Assay system (Promega). Luminescence was measured with a Bio-Orbit 1251 Luminometer (Bio-Orbit, Turku, Finland). Luciferase expression from pGL3 constructs was normalized to pRL-TK expression.
Nuclear Protein Extraction Nuclear extracts were prepared essentially as previously described.16 Nuclear protein concentration was determined using Coomassie Protein Assay Reagent (Pierce, Rockville, IL).
Deoxyribonuclease I Footprinting A fragment of the GC-C gene regulatory region 246 to 2257 relative to the start of transcription was obtained by digestion with DraIII and AflII, blunt-ended, and subcloned into the Bluescript KS EcoRV site, as described above, and then digested with EcoRI and HindIII to ensure that the coding strand of the probe was singly end-labeled with [a-32P]deoxycytidine triphosphate. The protocol for deoxyribonuclease (DNase) I protection was essentially as described in the SureTrack Footprinting Kit (Amersham, Pharmacia Biotech, Piscataway, NJ), except that CaCl2 was not included in the DNase I digestion buffer, and 2 µg of proteinase K was added to each sample and incubated for 30 minutes at 42°C after the addition of the stop solution (192 mmol/L sodium acetate, 32 mmol/L EDTA, 0.14% sodium dodecyl sulfate [SDS], and 64 µg/mL yeast RNA). Products obtained from footprinting reactions were separated on a denaturing 6% polyacrylamide gel and visualized by a Phosphorimager SI (Molecular Dynamics, Sunnyvale, CA).
Electromobility Shift Assay Protein-DNA binding reactions, performed in the same buffer as the DNase I protection assay (4% glycerol, 10 mmol/L Tris-HCl [pH 7.5], 50 mmol/L NaCl, 2.5 mmol/L MgCl2, and 5 mmol/L dithiothreitol), included 1 µg of poly(dI·dC)-poly(dI·dC) (Amersham Pharmacia Biotech) and 30,000 cpm of probe. Reactions were initiated by the addition of nuclear extract and incubated for 30 minutes at room temperature to produce protein complexes that were separated on a 6% nondenaturing, polyacrylamide (37.5:1) gel in 0.53 TBE running buffer (45 mmol/L Tris borate, 1 mmol/L EDTA, pH 8.3). Gels were dried before visualization of radiolabeled complexes by autoradiography. In competition assays, unlabeled competitor was added to the reaction mixtures at concentrations ranging from 25-fold to 250-fold molar excess of the labeled probe before the addition of the nuclear extract. Supershift assays were performed by adding 2 µL of murine Cdx2 antibody (P. Traber, University of Pennsylvania, Philadelphia, PA) after an initial incubation period of 30 minutes; incubation was then continued for an additional 30 minutes.
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Transcribed and translated (TNT) murine Cdx2 protein was generated in vitro using linearized pRc/CMV-Cdx2 expression vector (P. Traber) as a template for the TNT-Quickcoupled Kit (Promega). Oligonucleotide probes for electrophoretic mobility assay (EMSA) were synthesized by the Kimmel Cancer Institute Nucleic Acid Facility. Complementary oligonucleotides in 10 mmol/L Tris-HCl (pH 7.5) plus 1 mmol/L EDTA were annealed in a Hybaid Thermal Cycler (Teddington, United Kingdom) by a programmed ramp in temperature from 95°C to 25°C over the course of 1 hour. The single stranded sequences of the probes were: FP1: 58 CAGCTAATCTCTCTGTTTATAGCTCTGACCTTTC 38; FP1B: 58 ATCTCTCTGTTTATAGCTCTGACCTTTCTGGGTGC 38; FP1-CCC: 58 CAGCTAATCTCTCTGCCCATAGCTCTGACCTTTC 38; and SIF1: 58 GATCCGGCTGGTGAGGGTGCAATAAAACTTTATGAGTA 38. Bold sequences indicate specific Cdx2 binding sites (see Results). A mutation created in the FP1-protected site is underlined. Five picomoles of annealed oligonucleotide probe was end-labeled using 1 unit of T4 polynucleotide kinase and 2 µL of 7000 Ci/mmol [g-32P]adenosine triphosphate.16 Labeled probes were purified over Qiaquick nucleotide purification columns (Qiagen).
Southwestern and Western Blotting Nuclear extracts were denatured in reducing SDS sample buffer, separated on an 8% Tris-glycine-SDS polyacrylamide gel, and transferred to nitrocellulose. For Southwestern analysis, the blotted proteins were blocked for 1 hour at 4°C in Z8 buffer (25 mmol/L HEPES-KOH [pH 7.6], 12.5 mmol/L MgCl2, 20% glycerol, 0.1% Nonidet P-40, 100 mmol/L KCl, 10 mmol/L ZnSO4, and 1 mmol/L dithiothreitol) containing 3% nonfat dry milk.17 The membrane was rinsed for 5 minutes in EMSA binding buffer and hybridized with 20 mL of EMSA binding buffer with 100,000 cpm/mL of labeled FP1 probe for 1 hour at room temperature. The membrane was then washed for 5 minutes each in 3 changes of EMSA binding buffer, dried, and visualized by autoradiography. Western blots were blocked in Tris-buffered saline/0.1% Tween 20 with 5% nonfat dry milk, and probed with Cdx2 antibody diluted 1:5000. Binding of primary antibody was visualized using goat anti-rabbit alkaline phosphatase– conjugated secondary antibody diluted 1:10,000 (Sigma, St. Louis, MO). Alkaline phosphatase substrates BCIP and NBT were used in an AP Color Kit (Bio-Rad, Hercules, CA).
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Figure 1. Functional characterization of deletion mutants of the human GC-C gene promoter. Deletion mutants of the GC-C gene 58-flanking region were linked to luciferase and cotransfected with the Renilla luciferase control plasmid pRL-TK into intestinal (T84, Caco2) and extraintestinal (HepG2, HeLa, HS766T) cell lines as described in Materials and Methods. Data are expressed as luciferase activity relative to the pGL3 Basic promoterless construct (Relative Activity). Each bar represents the mean 6 SE of at least 3 independent transfections performed in duplicate.
gene constructs (Figure 1). Luciferase activity did not increase when extraintestinal cells were transfected with these constructs (Figure 1). These results are consistent with previous studies of GC-C gene regulation, and suggest that there are one or more tissue-specific regulatory elements within the 1118 to 2257 region.12 Because transfection with the 246 to 2129 construct resulted in a significant increase in activity of the reporter gene in intestinal cells only, and because this region is highly conserved evolutionarily, it was chosen for detailed structure-function analysis.13 DNase I Protection by Intestine-Specific Nuclear Protein Binding to the 58 Regulatory Region of GC-C
Determination of Elements Controlling Intestine-Specific Expression in the 58 Regulatory Region of the GC-C Gene
DNase I protection assay revealed 2 regions (275 to 283, FP1; 2164 to 2178, FP3) that were protected only by nuclear extracts from intestinal cells (T84; Figure 2). Regions 2104 to 2137 (FP2) and 2180 to 2217 (FP4) were protected by nuclear extracts from either intestinal (T84) or extraintestinal (HepG2) cells, although the proximal and distal ends of FP2 showed different patterns of protection. These data suggest that the protected regions designated FP1 and FP3 were specific binding sites for nuclear proteins from intestinal cells. In addition, an intestine-specific site of open chromatin structure in the proximal 58-flanking region of the GC-C gene was identified by a DNase I–hypersensitive site at base 2163 (Figure 2).
Minimal luciferase activity was obtained when various cell lines were transfected with the 246 construct (Figure 1). In contrast, luciferase activity increased in intestinal cells transfected with each of the other reporter
Transfection of T84 cells revealed that deletion of FP3 increased luciferase activity 2.5-fold relative to the
Results
Transcriptional Activity of the 2857 Construct After Deletion of FP1 or FP3
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scription factor that directs intestine-specific expression of several genes, FP1 was more closely examined.11 Specific Complexes Are Formed by Intestinal Nuclear Extract and FP1 Probe
Figure 2. DNase I protection of the proximal human GC-C promoter. Footprinting reactions, as described in Materials and Methods, included the indicated microgram quantities of HepG2 or T84 nuclear extract (NE) and the 246 to 2257 promoter fragment labeled at the 58-end of the coding strand. A control digestion contained 60 µg of bovine serum albumin (BSA). Protected bases were identified by a Maxam–Gilbert sequencing reaction (G-A) of the labeled fragment.16 The sequence of FP1 is given. Arrowhead indicates DNase I hypersensitivity site at base 2163.
wild-type construct (Figure 3). In contrast, elimination of FP1 reduced luciferase activity in T84 cells to levels observed in HepG2 cells (Figure 3). These data suggest that FP3 contains a negative regulatory element, and that FP1 contains an intestine-specific positive regulatory element. Analysis by TRANSFAC, a database of transcription factor binding sites (http://transfac.gbf.de/), revealed that FP1 contains the consensus binding site for the homeodomain protein Cdx2.18 Because Cdx2 is a tran-
Figure 3. Regulation of reporter gene expression by intestine-specific protected elements. FP1 and FP3 were deleted from the 2835 luciferase construct by in vitro mutagenesis, and wild-type and deletion constructs were expressed in HepG2 and T84 cells, as described in Materials and Methods. Results are expressed as luciferase activity relative to a promoterless construct and represent the mean 6 SE of 3 independent transfections performed in duplicate.
The ability of the protected site FP1 to form intestine-specific complexes was determined by incubating an oligonucleotide probe with nuclear extracts prepared from T84, Caco2, HepG2, and HeLa cells. Several complexes were obtained by EMSA when the FP1 probe was incubated with nuclear extracts from those cells (Figure 4). However, only one complex satisfied criteria for intestinal specificity, including formation by nuclear extracts from T84 and Caco2 cells, but not from HepG2 or HeLa cells. Extracts from T84 and Caco2 cells, but not from HepG2 or HeLa cells, also formed complexes with SIF1 that were identical to those previously characterized with that probe, showing the integrity of the extracts used in the present study (data not shown).19 All of the EMSA complexes formed with T84 nuclear extracts were competed with increasing amounts of unlabeled FP1 probe in a concentration-dependent manner (Figure 5A and B). In contrast, an unlabeled competitor in which the Cdx2 binding site was specifically mutated (FP1-CCC probe, see Materials and Methods) did not compete against the intestine-specific complex (Figure 5A). SIF1, an oligonucleotide containing 2 consensus-binding sites for Cdx2, selectively prevented the formation of the FP1-dependent intestine-specific complex with greater potency than unlabeled FP1, but generally did not affect
Figure 4. Intestinal specificity of FP1 probe EMSA. Nuclear extracts from intestinal or extraintestinal cells, or BSA (10 µg), were incubated with labeled FP1 for 30 minutes at room temperature before separation on a nondenaturing 6% polyacrylamide gel. Arrow and asterisk indicate the intestine-specific protein-FP1 complex.
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Figure 5. FP1 probe EMSA competition. Reaction conditions were similar to those in Figure 4, except (A) unlabeled competitors FP1, FP1-CCC, or (B) SIF1 were added in molar concentrations ranging from 253 to 2503 greater than labeled FP1 probe, and were all incubated with 5 µg of T84 nuclear extract. Arrows and asterisks indicate the intestine-specific proteinFP1 complex.
the binding of the remaining T84-EMSA complexes (Figure 5B).19 These data suggest that the intestinespecific factor that binds to the FP1-protected site is most likely Cdx2. Cdx2 Binds Specifically to the FP1 Probe To determine whether FP1 is a binding site for Cdx2, labeled FP1 was incubated with in vitro transcribed and translated murine Cdx2. This resulted in a complex whose mobility was identical to the intestine-
specific complex formed by T84 nuclear extract (Figure 6A and B). In contrast, labeled FP1-CCC did not form the intestine-specific complex with either Cdx2 or T84 nuclear extract (Figure 6A). An antibody against Cdx2 decreased the mobility of the specific complex formed between labeled FP1 and either T84 nuclear extract or in vitro TNT Cdx2 (Figure 6B). In contrast, an antibody against a related homeodomain transcription factor, Cdx1, did not alter the mobility of the intestinespecific complex (Figure 6B). These data led to the
Figure 6. The FP1 intestine-specific complex is Cdx2. (A) T84 nuclear extract (NE in micrograms) or in vitro TNT Cdx2 (2 µL) was incubated with labeled FP1 or FP1-CCC. Control incubations contained a 2-µL aliquot of a TNT reaction performed without Cdx2 complementary DNA. The arrow indicates the intestine-specific protein-FP1 complex. (B) T84 nuclear extract (NE 5 µg) or in vitro TNT Cdx2 (2 µL) or control BSA (10 µg) was incubated with labeled FP1. Anti-Cdx1 or -Cdx2 antibody was included in the incubation where indicated. Arrows and asterisks indicate the intestine-specific protein-FP1 complex. Arrowheads indicate protein-FP1 complexes exhibiting decreased mobility in the presence of specific antibody.
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conclusion that the FP1-protected site is a binding site for Cdx2. Identification of the Intestine-Specific Nuclear Factor by Southwestern and Western Blots Whether the FP1 probe and anti-Cdx2 antibody bound to the same intestine-specific protein was examined. Labeled FP1B, which contains the same DNase I–protected region as the FP1 probe, specifically bound to an intestine-specific protein of approximately 40 kilodaltons in T84 and Caco2, but not HepG2, cell nuclear extracts. In addition, FP1B probe bound to an ,131-kilodalton protein present in all cell lines examined (Figure 7A). Similarly, anti-Cdx2 antibody recognized a protein doublet of approximately 40 kilodaltons expressed in T84, but not in HepG2 or HeLa, cell nuclear extracts, a pattern that is characteristic of Cdx2 (Figure 7B).20 Thus, the FP1-protected region binds to an intestine-specific factor of the same molecular weight and antigenic recognition as Cdx2. Furthermore, Southwestern blots revealed that the FP1 probe binds directly to Cdx2. Role of the Cdx2 Binding Element (FP1) in Intestine-Specific Gene Expression of the GC-C Promoter The ‘‘CCC’’ mutation was introduced into the FP1 element of the 2835 luciferase reporter gene construct
Figure 8. Cdx2 binding element FP1 is required for GC-C reporter gene activation. Putative binding sites for Cdx2 and HNF-4a are indicated on the 2835 construct. T84 and HepG2 cells were transfected with the 2835 reporter construct from which FP1 was deleted or the construct containing the ‘‘CCC’’ mutation. Results are expressed as (Luciferase Activity of Mutant Construct 4 Luciferase Activity of Wild-type Construct) 3 100, and represent the mean 6 SE of 3 independent transfections performed in duplicate. The values expressed as relative luciferase activities are as follows: wild-type: FP1 deletion; ‘‘CCC’’ mutation: T84 (16.2 6 2.7; 1.9 6 0.3; 2.3 6 0.1) and HepG2 (2.1 6 0.1; 2.9 6 0.3; 2.2 6 0.1).
(see Materials and Methods). This mutated reporter gene construct showed reduced activity in T84 cells that was comparable with the construct from which the entire FP1 region was deleted (Figure 8). Neither the FP1 deletion nor the ‘‘CCC’’ mutation in FP1 altered luciferase expression in HepG2 cells (Figure 8). These data show that an intact Cdx2 binding site is required for activity of the GC-C promoter. Indeed, disruption of the Cdx2 binding site resulted in minimal activity.
Discussion
Figure 7. Cdx2 binds directly to the GC-C Promoter. (A) Southwestern blot of 50 or 100 µg of T84, HepG2, or Caco2 nuclear extracts probed with labeled FP1B. The arrow indicates the protein bound by FP1B at approximately 40 kilodaltons as determined from the mobility of prestained molecular-weight markers (Bio-Rad). (B) Western blot of 100 µg of T84, HepG2, or HeLa nuclear extract or in vitro TNT Cdx2 using anti-Cdx2 antibody. The arrow indicates the ,40-kilodalton protein bound by anti-Cdx2 antibody, determined from the mobility of prestained molecular-weight markers (Bio-Rad).
The precise molecular mechanisms underlying tissue-specific expression of particulate guanylyl cyclases remain undefined. In the present study we identified a region of the proximal GC-C promoter required for specific expression in intestinal cells that contains a protected region, FP1, with a consensus binding sequence for Cdx2. FP1 formed a specific complex with nuclear proteins only from intestinal cells, and this complex was recognized by anti-Cdx2 antibody. Elimination or mutation of the Cdx2 consensus binding sequence within FP1 reduced reporter gene activity in intestinal cells to that obtained in extraintestinal cells. These data suggest that Cdx2 activates tissue-specific transcription of GC-C. This is the first identification of a transcriptional activating factor required for intestine-specific expression of GC-C. Previous studies characterizing the transcriptional regulation of GC-C showed that HNF-4a is required for activation of expression.13 HNF-4a, a transcription factor that directs protein expression in hepatocytes, binds to an
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element more proximal to the start of transcription in the GC-C promoter than Cdx2.13,14,21 Although both of these factors bind independently to the GC-C promoter, as shown by DNase I footprint analysis and EMSA, their precise functional relationship to GC-C expression remains to be elucidated. Indeed, activation of expression may involve coordinated interaction of HNF-4a and Cdx2 with the GC-C promoter. However, the specificity of GC-C expression in the intestine likely resides with Cdx2, because HNF-4a is expressed in extraintestinal tissues.13,14,21 Cdx2, a member of the subfamily of homeodomain transcription factors related to the Drosophila melanogaster protein caudal, shows intestine-specific expression and is important in enterocyte development and differentiation.22,23 Caudal-related homeodomain proteins mediate normal intestinal development.24,25 Although Cdx1 and Cdx2 are expressed throughout the intestine, Cdx1 is most abundant in the rectum, whereas Cdx2 is most abundant in the cecum.20 Cdx2 regulates the expression of genes that are selectively expressed in specific regions of the intestine, such as sucrase-isomaltase in the small intestine and carbonic anhydrase I in the colon.19,26 These previous findings suggest that Cdx2, a protein expressed along the entire rostrocaudal axis of the intestine, functions in a coordinated fashion with other transcriptional regulatory factors to mediate gene expression in selective regions of the intestine. GC-C appears to be the first example of a gene regulated by, and exhibiting an overlapping expression pattern with, Cdx2 along the entire rostrocaudal axis of the intestine.20 Thus, neither are expressed in the stomach, but both are transcribed and translated in enterocytes with a rank order of small intestine . large intestine 5 rectum.3,9,20,27,28 Similarly, GC-C and Cdx2 show comparable patterns of expression in enterocytes along the crypt-to-villus axis in the small and large intestine.9,20,27 In conclusion, these studies show that a Cdx2 binding element in the proximal promoter is required to regulate intestine-specific expression of GC-C. Because the pattern of GC-C expression reflects that of Cdx2, the GC-C gene promoter may be a model system in which to study the interaction of Cdx2 with other regulatory proteins in mediating epithelial differentiation. The results suggest that a repressor binds to an element of the GC-C promoter that is 58 to the Cdx2 binding site. Structurefunction relations between such transcriptional activators and repressors in the regulation of GC-C expression are under investigation.
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by the homeodomain protein Cdx2. FEBS Lett 1998;423: 218–222. 27. Swenson ES, Mann EA, Jump ML, Witte DP, Giannella RA. The guanylin/STa receptor is expressed in crypts and apical epithelium throughout the mouse intestine. Biochem Biophys Res Commun 1996;225:1009–1014. 28. Krause G, Bayerl A, Heim JM, Singh S, Gerzer R. Distribution of membrane bound guanylyl cyclases in human intestine. Gut 1994;35:1250–1257.
Received September 10, 1999. Accepted February 23, 2000. Address requests for reprints to: Scott A. Waldman, M.D., Ph.D., 132 South 10th Street, 1170 Main, Philadelphia, Pennsylvania 19107. e-mail: [email protected]; fax: (215) 955-5681. Supported by funding from the National Institutes of Health (grants RO1 HL659214, RO1 CA75123, and R21 CA79663 to S.A.W.), the American Cancer Society (grant EDT-106 to S.A.W.), and Targeted Diagnostics and Therapeutics, Inc. J.P. was supported by NIH pre-doctoral training grant 5 T32 DK07705-05. The authors thank Peter G. Traber for contributing the mouse Cdx1 and Cdx2 expression vectors and antibodies, and Drs. Kenneth P. Chepenik and Martyn Darby for critical review of the manuscript.
Intestine-Specific Activity of the Human Guanylyl Cyclase C Promoter Is Regulated by Cdx2 JASON PARK, STEPHANIE SCHULZ, and SCOTT A. WALDMAN Division of Clinical Pharmacology, Departments of Medicine and Biochemistry and Molecular Pharmacology, Thomas Jefferson University, Philadelphia, Pennsylvania
Background & Aims: The heat-stable enterotoxin receptor, guanylyl cyclase C, exhibits an intestine-specific pattern of expression. The aim of this study was to identify the transcriptional activator that mediates intestine-specific expression of guanylyl cyclase C. Methods: Fragments of the promoter were assayed to isolate regions directing intestine-specific gene activation. Deoxyribonuclease I footprinting was used to identify a site of intestine-specific protection. Electrophoretic mobility shift assays (EMSAs) and supershift analyses were used to characterize the protein that bound to the protected site. The protected site was mutated to analyze its role in promoter activity. Results: Reporter gene assays revealed that intestine-specific expression of guanylyl cyclase C is directed by the proximal promoter. Deoxyribonuclease I footprinting identified a specific site in the proximal promoter that exhibited intestine-specific protection. EMSAs and supershift analyses revealed that the transcription factor Cdx2 bound to an intestine-specific site of protection. Mutation of the Cdx2-protected site of the promoter eliminated binding of Cdx2 and reduced reporter gene activity to the level of extraintestinal cells. Conclusions: These data show that Cdx2 and its consensus-binding site in the promoter are required for intestine-specific expression of the guanylyl cyclase C gene.
uanylyl cyclase C (GC-C), the receptor for the Escherichia coli heat-stable enterotoxin (STa), catalyzes the conversion of guanosine triphosphate to guanosine 38,58-monophosphate, which initiates a biochemical cascade culminating in the opening of the cystic fibrosis transmembrane conductance regulator, a chloride channel. Efflux of chloride from enterocytes drives intestinal fluid accumulation and results in secretory diarrhea. The intestinal peptides guanylin and uroguanylin are putative endogenous ligands of GC-C, although they bind to and activate GC-C with a lower potency than STa.1,2 Conservation of GC-C throughout the vertebrate subphylum implies that it serves an important physiologic role that remains undefined.3,4 However, targeted disruption of the GC-C gene had no deleterious effects in
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mice, and in fact conferred resistance to STa-induced diarrhea.5,6 In humans, GC-C is expressed exclusively in epithelial cells lining the small and large intestines and in tumors derived from those cells.7,8 Messenger RNA encoding GC-C is localized in crypt and villous cells in the small intestine and in differentiated enterocytes in crypts of the large intestine.9,10 Similar to other intestine-specific genes that are predominately expressed in the small intestine, the level of expression of GC-C decreases along the rostrocaudal axis.11 Tissue specificity of GC-C transcription is directed by the evolutionarily conserved regulatory sequence 1118 to 2257 relative to the transcription start site.12,13 Indeed, GC-C transcription is dependent on a promoter element at 229 to 246 that binds hepatocyte nuclear factor 4a (HNF-4a).13 Although HNF-4a is required for GC-C transcription in the intestine, this transactivating protein is also expressed in liver, kidney, and pancreas. However, GC-C is not expressed in these organs.7,14,15 This wide distribution of HNF-4a expression suggests that additional factors are responsible for expression of GC-C in the intestine. This study identified a site in the promoter of GC-C, 275 to 283, that includes a consensus sequence for binding the intestine-specific homeodomain protein, Cdx2. This site interacts with a protein expressed by intestinal, but not extraintestinal, cells that is functionally, physically, and antigenically similar to Cdx2. This site also interacts specifically with Cdx2 obtained by in vitro transcription and translation. Additionally, this Cdx2 consensus-binding site is required for intestinespecific transcriptional activation of the GC-C promoter. Abbreviations used in this paper: BSA, bovine serum albumin; DMEM, Dulbecco’s modified Eagle medium; EMSA, electrophoretic mobility shift assay; GC-C, guanylyl cyclase C; HNF-4a, hepatocyte nuclear factor 4a; NE, nuclear extract; SDS, sodium dodecyl sulfate; STa, heat-stable enterotoxin; TNT, transcribed and translated. r 2000 by the American Gastroenterological Association 0016-5085/00/$10.00 doi:10.1053/gast.2000.8520
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Materials and Methods Genomic Library Screening and Sequencing The GC-C gene 58 regulatory region was cloned from a lFIXII human genomic library (Stratagene, La Jolla, CA). The library was screened by hybridization with a probe specific for exon 1 of the GC-C complementary DNA. A 2.8-kilobase (kb) XbaI fragment that included 2 kb upstream of the start site of transcription was subcloned into Bluescript KS (Stratagene). All constructs were generated from this Bluescript/human GC-C gene construct. The nucleic acid sequence of each construct was confirmed by BigDye terminator reaction chemistry for sequence analysis on the Applied Biosystems model 377 DNA sequencing system (Perkin-Elmer Applied Biosystems, Foster City, CA; Kimmel Cancer Institute Nucleic Acid Facility, Thomas Jefferson University, Philadelphia, PA).
Reporter Gene Constructs Fragments 2835 to 1117, 2257 to 1117, 2129 to 1117, and 246 to 1117, relative to the start site of transcription, were isolated from Bluescript KS constructs by digestion with selected restriction endonucleases.12 These fragments were blunt-ended and ligated into the EcoRV site of Bluescript KS. Inserts were excised from Bluescript KS with SmaI and KpnI and ligated into the pGL3-Basic Luciferase Vector (Promega, Madison, WI). The pGL3 Control Vector, containing an SV40 promoter with enhancers, was used as a positive control. Mutations were created in the 2835 to 1117 pGL3 construct using the polymerase chain reaction–based Ex-site Mutation Kit (Stratagene). Deletion constructs were created using primers flanking the sites of interest. The FP1 ‘‘CCC’’ mutant was created using the phosphorylated primers: 58GCCCATAGCTCTGACCTTTCTG38 and 58AGAGAGATTAGCTGGGCCTCACCC38.
Cell Culture and Transfection All cell lines were obtained from American Type Culture Collection (Rockville, MD). T84 cells were grown in Dulbecco’s modified Eagle medium (DMEM)/F12 (Life Technologies), Caco2 cells in DMEM (Life Technologies, Rockville, MD), HepG2 and HS766T cells in DMEM High Glucose (Cellgro, Herndon, VA), and HeLa cells in minimum essential medium Eagle (MEM) with glutamine (Life Technologies). All cell lines were maintained at 37°C in a 5% CO2/95% air atmosphere and passaged every 4 days. Assays of reporter gene activity were conducted with cells seeded in 6-well plates at either 5.0 3 105 (T84, Caco2, and HeLa) or 1.0 3 106 cells per well (HepG2 and HS766T). Cells were incubated overnight, washed once with phosphate-buffered saline, and supplemented with fresh media before transfection. Plasmids purified with the Qiafilter Kit (Qiagen, Valencia, CA) were transfected into cells with the nonliposomal lipid transfection reagent Effectene (Stratagene). All cell lines were cotransfected with 0.4 µg of firefly luciferase experimental reporter
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constructs, modified from pGL3-Basic, and 0.1 µg of the Renilla luciferase control reporter, pRL-TK, driven by a viral thymidine kinase promoter (Promega). Cells were incubated with transfection complexes for 24 hours, rinsed with phosphate-buffered saline, then supplemented with appropriate media and incubated for a further 24 hours. After a total of 48 hours, cells were lysed and assayed using the protocol and materials in the Dual-Luciferase Reporter Assay system (Promega). Luminescence was measured with a Bio-Orbit 1251 Luminometer (Bio-Orbit, Turku, Finland). Luciferase expression from pGL3 constructs was normalized to pRL-TK expression.
Nuclear Protein Extraction Nuclear extracts were prepared essentially as previously described.16 Nuclear protein concentration was determined using Coomassie Protein Assay Reagent (Pierce, Rockville, IL).
Deoxyribonuclease I Footprinting A fragment of the GC-C gene regulatory region 246 to 2257 relative to the start of transcription was obtained by digestion with DraIII and AflII, blunt-ended, and subcloned into the Bluescript KS EcoRV site, as described above, and then digested with EcoRI and HindIII to ensure that the coding strand of the probe was singly end-labeled with [a-32P]deoxycytidine triphosphate. The protocol for deoxyribonuclease (DNase) I protection was essentially as described in the SureTrack Footprinting Kit (Amersham, Pharmacia Biotech, Piscataway, NJ), except that CaCl2 was not included in the DNase I digestion buffer, and 2 µg of proteinase K was added to each sample and incubated for 30 minutes at 42°C after the addition of the stop solution (192 mmol/L sodium acetate, 32 mmol/L EDTA, 0.14% sodium dodecyl sulfate [SDS], and 64 µg/mL yeast RNA). Products obtained from footprinting reactions were separated on a denaturing 6% polyacrylamide gel and visualized by a Phosphorimager SI (Molecular Dynamics, Sunnyvale, CA).
Electromobility Shift Assay Protein-DNA binding reactions, performed in the same buffer as the DNase I protection assay (4% glycerol, 10 mmol/L Tris-HCl [pH 7.5], 50 mmol/L NaCl, 2.5 mmol/L MgCl2, and 5 mmol/L dithiothreitol), included 1 µg of poly(dI·dC)-poly(dI·dC) (Amersham Pharmacia Biotech) and 30,000 cpm of probe. Reactions were initiated by the addition of nuclear extract and incubated for 30 minutes at room temperature to produce protein complexes that were separated on a 6% nondenaturing, polyacrylamide (37.5:1) gel in 0.53 TBE running buffer (45 mmol/L Tris borate, 1 mmol/L EDTA, pH 8.3). Gels were dried before visualization of radiolabeled complexes by autoradiography. In competition assays, unlabeled competitor was added to the reaction mixtures at concentrations ranging from 25-fold to 250-fold molar excess of the labeled probe before the addition of the nuclear extract. Supershift assays were performed by adding 2 µL of murine Cdx2 antibody (P. Traber, University of Pennsylvania, Philadelphia, PA) after an initial incubation period of 30 minutes; incubation was then continued for an additional 30 minutes.
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Transcribed and translated (TNT) murine Cdx2 protein was generated in vitro using linearized pRc/CMV-Cdx2 expression vector (P. Traber) as a template for the TNT-Quickcoupled Kit (Promega). Oligonucleotide probes for electrophoretic mobility assay (EMSA) were synthesized by the Kimmel Cancer Institute Nucleic Acid Facility. Complementary oligonucleotides in 10 mmol/L Tris-HCl (pH 7.5) plus 1 mmol/L EDTA were annealed in a Hybaid Thermal Cycler (Teddington, United Kingdom) by a programmed ramp in temperature from 95°C to 25°C over the course of 1 hour. The single stranded sequences of the probes were: FP1: 58 CAGCTAATCTCTCTGTTTATAGCTCTGACCTTTC 38; FP1B: 58 ATCTCTCTGTTTATAGCTCTGACCTTTCTGGGTGC 38; FP1-CCC: 58 CAGCTAATCTCTCTGCCCATAGCTCTGACCTTTC 38; and SIF1: 58 GATCCGGCTGGTGAGGGTGCAATAAAACTTTATGAGTA 38. Bold sequences indicate specific Cdx2 binding sites (see Results). A mutation created in the FP1-protected site is underlined. Five picomoles of annealed oligonucleotide probe was end-labeled using 1 unit of T4 polynucleotide kinase and 2 µL of 7000 Ci/mmol [g-32P]adenosine triphosphate.16 Labeled probes were purified over Qiaquick nucleotide purification columns (Qiagen).
Southwestern and Western Blotting Nuclear extracts were denatured in reducing SDS sample buffer, separated on an 8% Tris-glycine-SDS polyacrylamide gel, and transferred to nitrocellulose. For Southwestern analysis, the blotted proteins were blocked for 1 hour at 4°C in Z8 buffer (25 mmol/L HEPES-KOH [pH 7.6], 12.5 mmol/L MgCl2, 20% glycerol, 0.1% Nonidet P-40, 100 mmol/L KCl, 10 mmol/L ZnSO4, and 1 mmol/L dithiothreitol) containing 3% nonfat dry milk.17 The membrane was rinsed for 5 minutes in EMSA binding buffer and hybridized with 20 mL of EMSA binding buffer with 100,000 cpm/mL of labeled FP1 probe for 1 hour at room temperature. The membrane was then washed for 5 minutes each in 3 changes of EMSA binding buffer, dried, and visualized by autoradiography. Western blots were blocked in Tris-buffered saline/0.1% Tween 20 with 5% nonfat dry milk, and probed with Cdx2 antibody diluted 1:5000. Binding of primary antibody was visualized using goat anti-rabbit alkaline phosphatase– conjugated secondary antibody diluted 1:10,000 (Sigma, St. Louis, MO). Alkaline phosphatase substrates BCIP and NBT were used in an AP Color Kit (Bio-Rad, Hercules, CA).
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Figure 1. Functional characterization of deletion mutants of the human GC-C gene promoter. Deletion mutants of the GC-C gene 58-flanking region were linked to luciferase and cotransfected with the Renilla luciferase control plasmid pRL-TK into intestinal (T84, Caco2) and extraintestinal (HepG2, HeLa, HS766T) cell lines as described in Materials and Methods. Data are expressed as luciferase activity relative to the pGL3 Basic promoterless construct (Relative Activity). Each bar represents the mean 6 SE of at least 3 independent transfections performed in duplicate.
gene constructs (Figure 1). Luciferase activity did not increase when extraintestinal cells were transfected with these constructs (Figure 1). These results are consistent with previous studies of GC-C gene regulation, and suggest that there are one or more tissue-specific regulatory elements within the 1118 to 2257 region.12 Because transfection with the 246 to 2129 construct resulted in a significant increase in activity of the reporter gene in intestinal cells only, and because this region is highly conserved evolutionarily, it was chosen for detailed structure-function analysis.13 DNase I Protection by Intestine-Specific Nuclear Protein Binding to the 58 Regulatory Region of GC-C
Determination of Elements Controlling Intestine-Specific Expression in the 58 Regulatory Region of the GC-C Gene
DNase I protection assay revealed 2 regions (275 to 283, FP1; 2164 to 2178, FP3) that were protected only by nuclear extracts from intestinal cells (T84; Figure 2). Regions 2104 to 2137 (FP2) and 2180 to 2217 (FP4) were protected by nuclear extracts from either intestinal (T84) or extraintestinal (HepG2) cells, although the proximal and distal ends of FP2 showed different patterns of protection. These data suggest that the protected regions designated FP1 and FP3 were specific binding sites for nuclear proteins from intestinal cells. In addition, an intestine-specific site of open chromatin structure in the proximal 58-flanking region of the GC-C gene was identified by a DNase I–hypersensitive site at base 2163 (Figure 2).
Minimal luciferase activity was obtained when various cell lines were transfected with the 246 construct (Figure 1). In contrast, luciferase activity increased in intestinal cells transfected with each of the other reporter
Transfection of T84 cells revealed that deletion of FP3 increased luciferase activity 2.5-fold relative to the
Results
Transcriptional Activity of the 2857 Construct After Deletion of FP1 or FP3
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scription factor that directs intestine-specific expression of several genes, FP1 was more closely examined.11 Specific Complexes Are Formed by Intestinal Nuclear Extract and FP1 Probe
Figure 2. DNase I protection of the proximal human GC-C promoter. Footprinting reactions, as described in Materials and Methods, included the indicated microgram quantities of HepG2 or T84 nuclear extract (NE) and the 246 to 2257 promoter fragment labeled at the 58-end of the coding strand. A control digestion contained 60 µg of bovine serum albumin (BSA). Protected bases were identified by a Maxam–Gilbert sequencing reaction (G-A) of the labeled fragment.16 The sequence of FP1 is given. Arrowhead indicates DNase I hypersensitivity site at base 2163.
wild-type construct (Figure 3). In contrast, elimination of FP1 reduced luciferase activity in T84 cells to levels observed in HepG2 cells (Figure 3). These data suggest that FP3 contains a negative regulatory element, and that FP1 contains an intestine-specific positive regulatory element. Analysis by TRANSFAC, a database of transcription factor binding sites (http://transfac.gbf.de/), revealed that FP1 contains the consensus binding site for the homeodomain protein Cdx2.18 Because Cdx2 is a tran-
Figure 3. Regulation of reporter gene expression by intestine-specific protected elements. FP1 and FP3 were deleted from the 2835 luciferase construct by in vitro mutagenesis, and wild-type and deletion constructs were expressed in HepG2 and T84 cells, as described in Materials and Methods. Results are expressed as luciferase activity relative to a promoterless construct and represent the mean 6 SE of 3 independent transfections performed in duplicate.
The ability of the protected site FP1 to form intestine-specific complexes was determined by incubating an oligonucleotide probe with nuclear extracts prepared from T84, Caco2, HepG2, and HeLa cells. Several complexes were obtained by EMSA when the FP1 probe was incubated with nuclear extracts from those cells (Figure 4). However, only one complex satisfied criteria for intestinal specificity, including formation by nuclear extracts from T84 and Caco2 cells, but not from HepG2 or HeLa cells. Extracts from T84 and Caco2 cells, but not from HepG2 or HeLa cells, also formed complexes with SIF1 that were identical to those previously characterized with that probe, showing the integrity of the extracts used in the present study (data not shown).19 All of the EMSA complexes formed with T84 nuclear extracts were competed with increasing amounts of unlabeled FP1 probe in a concentration-dependent manner (Figure 5A and B). In contrast, an unlabeled competitor in which the Cdx2 binding site was specifically mutated (FP1-CCC probe, see Materials and Methods) did not compete against the intestine-specific complex (Figure 5A). SIF1, an oligonucleotide containing 2 consensus-binding sites for Cdx2, selectively prevented the formation of the FP1-dependent intestine-specific complex with greater potency than unlabeled FP1, but generally did not affect
Figure 4. Intestinal specificity of FP1 probe EMSA. Nuclear extracts from intestinal or extraintestinal cells, or BSA (10 µg), were incubated with labeled FP1 for 30 minutes at room temperature before separation on a nondenaturing 6% polyacrylamide gel. Arrow and asterisk indicate the intestine-specific protein-FP1 complex.
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Figure 5. FP1 probe EMSA competition. Reaction conditions were similar to those in Figure 4, except (A) unlabeled competitors FP1, FP1-CCC, or (B) SIF1 were added in molar concentrations ranging from 253 to 2503 greater than labeled FP1 probe, and were all incubated with 5 µg of T84 nuclear extract. Arrows and asterisks indicate the intestine-specific proteinFP1 complex.
the binding of the remaining T84-EMSA complexes (Figure 5B).19 These data suggest that the intestinespecific factor that binds to the FP1-protected site is most likely Cdx2. Cdx2 Binds Specifically to the FP1 Probe To determine whether FP1 is a binding site for Cdx2, labeled FP1 was incubated with in vitro transcribed and translated murine Cdx2. This resulted in a complex whose mobility was identical to the intestine-
specific complex formed by T84 nuclear extract (Figure 6A and B). In contrast, labeled FP1-CCC did not form the intestine-specific complex with either Cdx2 or T84 nuclear extract (Figure 6A). An antibody against Cdx2 decreased the mobility of the specific complex formed between labeled FP1 and either T84 nuclear extract or in vitro TNT Cdx2 (Figure 6B). In contrast, an antibody against a related homeodomain transcription factor, Cdx1, did not alter the mobility of the intestinespecific complex (Figure 6B). These data led to the
Figure 6. The FP1 intestine-specific complex is Cdx2. (A) T84 nuclear extract (NE in micrograms) or in vitro TNT Cdx2 (2 µL) was incubated with labeled FP1 or FP1-CCC. Control incubations contained a 2-µL aliquot of a TNT reaction performed without Cdx2 complementary DNA. The arrow indicates the intestine-specific protein-FP1 complex. (B) T84 nuclear extract (NE 5 µg) or in vitro TNT Cdx2 (2 µL) or control BSA (10 µg) was incubated with labeled FP1. Anti-Cdx1 or -Cdx2 antibody was included in the incubation where indicated. Arrows and asterisks indicate the intestine-specific protein-FP1 complex. Arrowheads indicate protein-FP1 complexes exhibiting decreased mobility in the presence of specific antibody.
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conclusion that the FP1-protected site is a binding site for Cdx2. Identification of the Intestine-Specific Nuclear Factor by Southwestern and Western Blots Whether the FP1 probe and anti-Cdx2 antibody bound to the same intestine-specific protein was examined. Labeled FP1B, which contains the same DNase I–protected region as the FP1 probe, specifically bound to an intestine-specific protein of approximately 40 kilodaltons in T84 and Caco2, but not HepG2, cell nuclear extracts. In addition, FP1B probe bound to an ,131-kilodalton protein present in all cell lines examined (Figure 7A). Similarly, anti-Cdx2 antibody recognized a protein doublet of approximately 40 kilodaltons expressed in T84, but not in HepG2 or HeLa, cell nuclear extracts, a pattern that is characteristic of Cdx2 (Figure 7B).20 Thus, the FP1-protected region binds to an intestine-specific factor of the same molecular weight and antigenic recognition as Cdx2. Furthermore, Southwestern blots revealed that the FP1 probe binds directly to Cdx2. Role of the Cdx2 Binding Element (FP1) in Intestine-Specific Gene Expression of the GC-C Promoter The ‘‘CCC’’ mutation was introduced into the FP1 element of the 2835 luciferase reporter gene construct
Figure 8. Cdx2 binding element FP1 is required for GC-C reporter gene activation. Putative binding sites for Cdx2 and HNF-4a are indicated on the 2835 construct. T84 and HepG2 cells were transfected with the 2835 reporter construct from which FP1 was deleted or the construct containing the ‘‘CCC’’ mutation. Results are expressed as (Luciferase Activity of Mutant Construct 4 Luciferase Activity of Wild-type Construct) 3 100, and represent the mean 6 SE of 3 independent transfections performed in duplicate. The values expressed as relative luciferase activities are as follows: wild-type: FP1 deletion; ‘‘CCC’’ mutation: T84 (16.2 6 2.7; 1.9 6 0.3; 2.3 6 0.1) and HepG2 (2.1 6 0.1; 2.9 6 0.3; 2.2 6 0.1).
(see Materials and Methods). This mutated reporter gene construct showed reduced activity in T84 cells that was comparable with the construct from which the entire FP1 region was deleted (Figure 8). Neither the FP1 deletion nor the ‘‘CCC’’ mutation in FP1 altered luciferase expression in HepG2 cells (Figure 8). These data show that an intact Cdx2 binding site is required for activity of the GC-C promoter. Indeed, disruption of the Cdx2 binding site resulted in minimal activity.
Discussion
Figure 7. Cdx2 binds directly to the GC-C Promoter. (A) Southwestern blot of 50 or 100 µg of T84, HepG2, or Caco2 nuclear extracts probed with labeled FP1B. The arrow indicates the protein bound by FP1B at approximately 40 kilodaltons as determined from the mobility of prestained molecular-weight markers (Bio-Rad). (B) Western blot of 100 µg of T84, HepG2, or HeLa nuclear extract or in vitro TNT Cdx2 using anti-Cdx2 antibody. The arrow indicates the ,40-kilodalton protein bound by anti-Cdx2 antibody, determined from the mobility of prestained molecular-weight markers (Bio-Rad).
The precise molecular mechanisms underlying tissue-specific expression of particulate guanylyl cyclases remain undefined. In the present study we identified a region of the proximal GC-C promoter required for specific expression in intestinal cells that contains a protected region, FP1, with a consensus binding sequence for Cdx2. FP1 formed a specific complex with nuclear proteins only from intestinal cells, and this complex was recognized by anti-Cdx2 antibody. Elimination or mutation of the Cdx2 consensus binding sequence within FP1 reduced reporter gene activity in intestinal cells to that obtained in extraintestinal cells. These data suggest that Cdx2 activates tissue-specific transcription of GC-C. This is the first identification of a transcriptional activating factor required for intestine-specific expression of GC-C. Previous studies characterizing the transcriptional regulation of GC-C showed that HNF-4a is required for activation of expression.13 HNF-4a, a transcription factor that directs protein expression in hepatocytes, binds to an
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element more proximal to the start of transcription in the GC-C promoter than Cdx2.13,14,21 Although both of these factors bind independently to the GC-C promoter, as shown by DNase I footprint analysis and EMSA, their precise functional relationship to GC-C expression remains to be elucidated. Indeed, activation of expression may involve coordinated interaction of HNF-4a and Cdx2 with the GC-C promoter. However, the specificity of GC-C expression in the intestine likely resides with Cdx2, because HNF-4a is expressed in extraintestinal tissues.13,14,21 Cdx2, a member of the subfamily of homeodomain transcription factors related to the Drosophila melanogaster protein caudal, shows intestine-specific expression and is important in enterocyte development and differentiation.22,23 Caudal-related homeodomain proteins mediate normal intestinal development.24,25 Although Cdx1 and Cdx2 are expressed throughout the intestine, Cdx1 is most abundant in the rectum, whereas Cdx2 is most abundant in the cecum.20 Cdx2 regulates the expression of genes that are selectively expressed in specific regions of the intestine, such as sucrase-isomaltase in the small intestine and carbonic anhydrase I in the colon.19,26 These previous findings suggest that Cdx2, a protein expressed along the entire rostrocaudal axis of the intestine, functions in a coordinated fashion with other transcriptional regulatory factors to mediate gene expression in selective regions of the intestine. GC-C appears to be the first example of a gene regulated by, and exhibiting an overlapping expression pattern with, Cdx2 along the entire rostrocaudal axis of the intestine.20 Thus, neither are expressed in the stomach, but both are transcribed and translated in enterocytes with a rank order of small intestine . large intestine 5 rectum.3,9,20,27,28 Similarly, GC-C and Cdx2 show comparable patterns of expression in enterocytes along the crypt-to-villus axis in the small and large intestine.9,20,27 In conclusion, these studies show that a Cdx2 binding element in the proximal promoter is required to regulate intestine-specific expression of GC-C. Because the pattern of GC-C expression reflects that of Cdx2, the GC-C gene promoter may be a model system in which to study the interaction of Cdx2 with other regulatory proteins in mediating epithelial differentiation. The results suggest that a repressor binds to an element of the GC-C promoter that is 58 to the Cdx2 binding site. Structurefunction relations between such transcriptional activators and repressors in the regulation of GC-C expression are under investigation.
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References 1. Currie MG, Fok KF, Kato J, Moore RJ, Hamra FK, Duffin KL, Smith CE. Guanylin: an endogenous activator of intestinal guanylate cyclase. Proc Natl Acad Sci U S A 1992;89:947–951. 2. Hamra FK, Forte LR, Eber SL, Pidhorodeckyj NV, Krause WJ, Freeman RH, Chin DT, Tompkins JA, Fok KF, Smith CE, Duffin KL, Siegel NR, Currie MG. Uroguanylin: structure and activity of a second endogenous peptide that stimulates intestinal guanylate cyclase. Proc Natl Acad Sci U S A 1993;90:10464–10468. 3. Krause WJ, Freeman RH, Eber SL, Hamra FK, Fok KF, Currie MG, Forte LR. Distribution of Escherichia coli heat-stable enterotoxin/ guanylin/uroguanylin receptors in the avian intestinal tract. Acta Anat 1995;153:210–219. 4. Krause WJ, Freeman RH, Eber SL, Hamra FK, Currie MG, Forte LR. Guanylyl cyclase receptors and guanylin-like peptides in reptilian intestine. Gen Comp Endocrinol 1997;107:229–239. 5. Schulz S, Lopez MJ, Kuhn M, Garbers DL. Disruption of the guanylyl cyclase-C gene leads to a paradoxical phenotype of viable but heat-stable enterotoxin-resistant mice. J Clin Invest 1997;100: 1590–1595. 6. Mann EA, Jump ML, Wu J, Yee E, Giannella RA. Mice lacking the guanylyl cyclase C receptor are resistant to STa-induced intestinal secretion. Biochem Biophys Res Commun 1997;239:463–466. 7. Carrithers SL, Barber MT, Biswas S, Parkinson SJ, Park PK, Goldstein SD, Waldman SA. Guanylyl cyclase C is a selective marker for metastatic colorectal tumors in human extraintestinal tissues. Proc Natl Acad Sci U S A 1996;93:14827–14832. 8. Waldman SA, Barber M, Pearlman J, Park J, George R, Parkinson SJ. Heterogeneity of guanylyl cyclase C expressed by human colorectal cancer cell lines in vitro. Cancer Epidemiol Biomarkers Prev 1998;7:505–514. 9. Li Z, Goy MF. Peptide-regulated guanylate cyclase pathways in rat colon: in situ localization of GCA, GCC, and guanylin mRNA. Am J Physiol 1993;265:G394–402. 10. Cohen MB, Mann EA, Lau C, Henning SJ, Giannella RA. A gradient in expression of the Escherichia coli heat-stable enterotoxin receptor exists along the villus-to-crypt axis of rat small intestine. Biochem Biophys Res Commun 1992;186:483–490. 11. Traber PG, Silberg DG. Intestine-specific gene transcription. Annu Rev Physiol 1996;58:275–297. 12. Mann EA, Jump ML, Giannella RA. Cell line-specific transcriptional activation of the promoter of the human guanylyl cyclase C/heatstable enterotoxin/receptor gene. Biochim Biophys Acta 1996; 1305:7–10. 13. Swenson ES, Mann EA, Jump ML, Giannella RA. Hepatocyte nuclear factor-4 regulates intestinal expression of the guanylin/ heat-stable toxin receptor. Am J Physiol 1999;276:G728–736. 14. Sladek FM, Zhong WM, Lai E, Darnell JEJ. Liver-enriched transcription factor HNF-4 is a novel member of the steroid hormone receptor superfamily. Genes Dev 1990;4:2353–2365. 15. Carrithers SL, Parkinson SJ, Goldstein S, Park P, Robertson DC, Waldman SA. Escherichia coli heat-stable toxin receptors in human colonic tumors. Gastroenterology 1994;107:1653– 1661. 16. Ausubel FM, Brent R, Kingston RE, Moore DD, Seidman JG, Smith JA, Struhl K. Current protocols in molecular biology. New York: Wiley, 1999. 17. Hames BD, Higgins SJ. Gene transcription: a practical approach. The practical approach series. New York: Oxford, 1993. 18. Quandt K, Frech K, Karas H, Wingender E, Werner T. MatInd and MatInspector: new fast and versatile tools for detection of consensus matches in nucleotide sequence data. Nucleic Acids Res 1995;23:4878–4884. 19. Suh E, Chen L, Taylor J, Traber PG. A homeodomain protein related to caudal regulates intestine-specific gene transcription. Mol Cell Biol 1994;14:7340–7351.
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Received September 10, 1999. Accepted February 23, 2000. Address requests for reprints to: Scott A. Waldman, M.D., Ph.D., 132 South 10th Street, 1170 Main, Philadelphia, Pennsylvania 19107. e-mail: [email protected]; fax: (215) 955-5681. Supported by funding from the National Institutes of Health (grants RO1 HL659214, RO1 CA75123, and R21 CA79663 to S.A.W.), the American Cancer Society (grant EDT-106 to S.A.W.), and Targeted Diagnostics and Therapeutics, Inc. J.P. was supported by NIH pre-doctoral training grant 5 T32 DK07705-05. The authors thank Peter G. Traber for contributing the mouse Cdx1 and Cdx2 expression vectors and antibodies, and Drs. Kenneth P. Chepenik and Martyn Darby for critical review of the manuscript.