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© The Rockefeller University Press, 0021-9525/1998//765 $5.00
The Journal of Cell Biology, Volume 141, Number 3, , 1998 765-777

Effects of Forced Expression of an NH₂-terminal Truncated β-Catenin on Mouse Intestinal Epithelial Homeostasis

Melissa H. Wong^*, Bonnee Rubinfeld, and Jeffrey I. Gordon^*

^* Department of Molecular Biology and Pharmacology, Washington University School of Medicine, St. Louis, Missouri 63110; and Onyx Pharmaceuticals, Richmond, California 94806

β-Catenin functions as a downstream component of the Wnt/Wingless signal transduction pathway and as an effector of cell–cell adhesion through its association with cadherins. To explore the in vivo effects of β-catenin on proliferation, cell fate specification, adhesion, and migration in a mammalian epithelium, a human NH₂-terminal truncation mutant (N89β-catenin) was expressed in the 129/Sv embryonic stem cell–derived component of the small intestine of adult C57Bl/6–ROSA26 129/Sv chimeric mice. N89β-Catenin was chosen because mutants of this type are more stable than the wild-type protein, and phenocopy activation of the Wnt/Wingless signaling pathway in Xenopus and Drosophila. N89β-Catenin had several effects. Cell division was stimulated fourfold in undifferentiated cells located in the proliferative compartment of the intestine (crypts of Lieberkühn). The proliferative response was not associated with any discernible changes in cell fate specification but was accompanied by a three- to fourfold increase in crypt apoptosis. There was a marked augmentation of E-cadherin at the adherens junctions and basolateral surfaces of 129/Sv (N89β-catenin) intestinal epithelial cells and an accompanying slowing of cellular migration along crypt-villus units. 1–2% of 129/Sv (N89β-catenin) villi exhibited an abnormal branched architecture. Forced expression of N89β-catenin expression did not perturb the level or intracellular distribution of the tumor suppressor adenomatous polyposis coli (APC). The ability of N89β-catenin to interact with normal cellular pools of APC and/or augmented pools of E-cadherin may have helped prevent the 129/Sv gut epithelium from undergoing neoplastic transformation during the 10-mo period that animals were studied. Together, these in vivo studies emphasize the importance of β-catenin in regulating normal adhesive and signaling functions within this epithelium.

Abbreviations used in this paper: β-gal, β-galactosidase; N89β-catenin, NH₂-terminal truncation mutant of human β-catenin lacking amino acid residues 1–89; APC, adenomatous polyposis coli protein or gene; BrdU, 5'-bromo-2'deoxyuridine; Cy3, indocarbocyanine; DBA, Dolichos biflorus agglutinin; ES cell, embryonic stem cell; Fabpl, fatty acid binding protein gene; GSK-3, glycogen synthase kinase-3; hGH, human growth hormone gene; LEF-1, lymphocyte enhancing factor-1; PLP, periodate-lysine-paraformaldehyde; RT, reverse transcriptase; TAg, T antigen; Tcf, T-cell factor; X-Gal, 5-bromo-4-chloro-3-indolyl β-D-galactoside.

β-CATENIN plays important roles in cell adhesion and cell signaling (for review see Miller and Moon, 1996; Nusse, 1997). The protein influences adhesion by providing a functional bridge between cadherins and the actin cytoskeleton. Calcium-dependent homotypic interactions between the extracellular domains of cadherins result in the formation of adhesion "zippers" between adjacent cells (Overduin et al., 1995; Shapiro et al., 1995). Although these interactions help define the specificity of cellular interactions, they are not sufficient for productive adhesion. Productive adhesion at the adherens junction is accomplished by the binding of β-catenin to the conserved cytoplasmic domains of cadherins and to the cytoplasmic protein -catenin. -Catenin, in turn, is linked to the cytoskeleton via its interactions with other proteins (e.g., actinin; Nagafuchi and Takeichi, 1988; Ozawa et al., 1989, 1990; Aberle et al., 1994; Hinck et al., 1994; Jou et al., 1995; Rimm et al., 1995).

β-Catenin is also a critical downstream component of the Wnt signal transduction pathway in vertebrates. In the absence of a Wnt signal, serine/threonine phosphorylation by glycogen synthase kinase-3 (GSK-3)¹ leads to rapid degradation of cytoplasmic pools of β-catenin through a ubiquitin–proteosome pathway (Miller and Moon, 1996; Munemitsu et al., 1996; Yost et al., 1996; Aberle et al., 1997; Cadigan and Nusse, 1997). In contrast, stimulation of the Wnt pathway leads to repression of GSK-3 (Noordermeer et al., 1994; Cook et al., 1996), decreased β-catenin phosphorylation, and enhanced protein stability. The resulting augmentation of β-catenin pools facilitates formation of complexes between β-catenin and high mobility group box transcription factors (T-cell factor [Tcf] and lymphocyte enhancing factor-1 [LEF-1]); Behrens et al., 1996; Huber et al., 1996). In the nucleus, β-catenin functions to coactivate transcription of largely unspecified gene targets (Behrens et al., 1996; Huber et al., 1996; Molenaar et al., 1996; Brunner et al., 1997; Riese et al., 1997; van de Wetering et al., 1997).

Studies in genetically manipulatable nonvertebrate species as well as nonmammalian vertebrate organisms have shown that β-catenin–mediated signaling affects axis formation and cell fate specification (McCrea et al., 1993; Heasman et al., 1994; Funayama et al., 1995; Cox et al., 1996; Molenaar et al., 1996). However, attempts to test the in vivo functions of β-catenin in mammals have been hampered by the fact that mice homozygous for a genetically engineered null allele die during early embryogenesis (Haegel et al., 1995).

One function of β-catenin in mammalian cell lineages that has been recently explored is its role in oncogenic transformation. Several reports have emphasized that the tumor suppressor adenomatous polyposis coli (APC) functions to affect intestinal tumorigenesis through its regulation of β-catenin signaling. Mutations in APC lead to intestinal adenomas and adenocarcinoma in mice and humans (Su et al., 1992; for review see Kinzler and Vogelstein, 1996; Shibata et al., 1997). The interaction between β-catenin and APC is enhanced when APC is phosphorylated by GSK-3, thereby promoting β-catenin turnover (Su et al., 1993; Rubinfeld et al., 1993, 1996). When APC is absent, or is mutated so that β-catenin binding is impaired, cytosolic pools of β-catenin are elevated, and β-catenin–Tcf signaling is induced (Munemitsu et al., 1995; Korinek et al., 1997; Morin et al., 1997). Additional evidence for the importance of β-catenin signaling in tumorigenesis comes from the observation that some human colorectal neoplasms with wild-type APC genes have mutations in β-catenin (Ilyas et al., 1997; Morin et al., 1997). Some of these mutations affect potential sites for GSK-3 phosphorylation and therefore are likely to lead to augmented pools of β-catenin.

In addition to its role in gut neoplasia, there are other reasons why the self-renewing adult intestinal epithelium represents an attractive mammalian model for studying the in vivo functions of β-catenin. Fundamental developmental processes such as cellular proliferation, lineage allocation, differentiation, migration/adhesion, and death are continuously expressed in well-defined domains of the small intestine's anatomically distinct crypt-villus units. Proliferation is confined to flask-shaped mucosal invaginations known as crypts of Lieberkühn. In the adult mouse, each of the small intestinal crypts contains one or more active multipotent stem cells functionally anchored near its base (Loeffler et al., 1993; Potten et al., 1997). These stem cells give rise to daughters that undergo four to six rounds of cell division in the midportion of the crypt and are allocated to four principal epithelial cell lineages. Each lineage completes its differentiation during a highly organized, rapid migration. Absorptive enterocytes (accounting for >80% of the epithelial population), mucus-producing goblet cells, and enteroendocrine cells exit the crypt and move up an adjacent finger-like structure (the villus) in vertical coherent columns (Schmidt et al., 1985; Hermiston et al., 1996). Cells are removed from the villus tip by apoptosis and/or extrusion into the lumen (Hall et al., 1994). The entire sequence is completed in 3–5 d (Cheng and Leblond, 1974a–c; Cheng, 1974a). Members of the Paneth cell lineage differentiate as they move downward to the base of the crypt where they are removed after an 20-d residence (Cheng, 1974b). E-Cadherin, the principal intestinal epithelial cadherin, β-catenin, -catenin, and APC are expressed in cells distributed along the length of crypt-villus units (Hermiston et al., 1996; Näthke et al., 1996; Wong et al., 1996). The importance of cadherins and their related proteins to normal intestinal epithelial homeostasis was established when a dominant-negative cadherin was expressed in the 129/Sv component of C57Bl/6 129/ Sv chimeric mouse gut. Cell–cell and cell–substratum interactions were disrupted leading to alterations in migration rates, diminished cell survival, breakdown of mucosal barrier function, inflammatory bowel disease, and adenoma formation (Hermiston et al., 1995a,b).

In the present report, we have examined the effects of forced expression of a human β-catenin lacking its NH₂-terminal 89 residues (N89β-catenin) on the small intestinal epithelium of C57Bl/6 129/Sv chimeric mice. N89β-Catenin was chosen for several reasons. Studies in cultured cell lines indicated that this mutant, which lacks putative GSK-3 phosphorylation sites (Yost et al., 1996), has a longer half-life than the wild-type protein, leading to its accumulation both as a stable monomer and as a complex with APC (Munemitsu et al., 1996; Barth et al., 1997; Rubinfeld et al., 1997). The NH₂-terminal truncation does not appear to affect binding to E-cadherin, does not involve the domains involved in binding -catenin or Tcf (Munemitsu et al., 1996; Orsulic and Peifer, 1996), and preserves all of the protein's structurally and functionally important armadillo repeats (Huber et al., 1997). Finally, studies in Drosophila and Xenopus had shown that NH₂-terminal truncation mutants of β-catenin promote signaling (Yost et al., 1996; Pai et al., 1997), whereas expression in cultured mammalian epithelial cells affects cell migration (Pollack et al., 1997). Thus, we anticipated that N89β-catenin would allow us to examine the effects of β-catenin on both signaling and adhesion/migration in the mouse intestine.

	Materials and Methods

Top Abstract Materials and Methods Results Discussion Prospectus References

 
Generation of Chimeric-Transgenic Mice
pL596hGHpNeoB₂ (Hermiston et al., 1996) contains nucleotides -596 to +21 of the rat fatty acid binding protein gene (Fabpl) upstream of nucleotides +3 to +2150 of the human growth hormone gene (hGH). A phosphoglycerate kinase (pgk) neomycin resistance selection cassette is located just downstream of Fabpl-hGH. pL596hGHpNeoB₂ was cleaved at its unique BamHI site located at nucleotide +3 of hGH, and a double-stranded oligonucleotide containing flanking BamHI sites and internal KpnI and XbaI sites (5'-GATCCGGTACCATCGATGGGTCTAGAG-3') was inserted. The resulting DNA, pLFhGHpNeoK contains a unique KpnI site for subcloning cDNAs into exon 1 of hGH. pCANβ-cateninN₉₀c.62 (provided by P. Polakis, Onyx Pharmaceuticals, Richmond, CA) contains a DNA insert encoding myc-tagged human N89β-catenin. A 2.2-kb KpnI/XbaI fragment from pCANβ-cateninN₉₀c.62 was subcloned into KpnI/XbaI–digested pLFhGHpNeoK DNA, creating pLFN89βcat. The 6.9-kb Fabpl–N89βcat-hGH–pgkneo insert in pLFN89βcat was excised with SacII, purified by gel electrophoresis, and then electroporated into D3 129/Sv ES cells (Hermiston et al., 1995a). Stably transfected embryonic stem (ES) cell clones were injected into C57Bl/6 ROSA26^+/– blastocysts (Wong et al., 1996) to produce B6-ROSA26 129/Sv(N89β-catenin) chimeric-transgenic mice. Chimeras generated from three of the cell lines were analyzed at 6 wk, 6 mo, and 10 mo of life. Their 129/Sv contribution ranged from 20 to 80% based on coat color. The presence of ROSA26 in chimeras was established using a PCR protocol (The Jackson Laboratory, Bar Harbor, ME) and tail genomic DNA. Independent confirmation was obtained by staining segments of tail with 5-bromo-4-chloro-3-inodyl β-D-galactoside (X-Gal) (Wong et al., 1996). Nontransfected D3 ES cells were used to generate B6-ROSA26 129/Sv chimeras (referred to as "normal control" chimeras in this study).

All animals were maintained in microisolator cages under a strict 12-h light cycle and fed a standard chow diet ad libitum (Pico Lab Rodent Diet 20; Purina Mills, Inc., St. Louis, MO). Routine screening of sentinel animals indicated that all chimeric mice were free of pathogens.

Assays for Transgene Expression
Reverse Transcriptase-PCR.
Total cellular RNA was isolated from the proximal, middle, and distal thirds of the small intestine and the skeletal muscle of 6-wk-old B6-ROSA26 129/Sv(N89β-catenin) mice using the RNeasy kit (QIAGEN Inc., Santa Clarita, CA). Oligo dT-primed cDNA was generated from DNA-free intestinal RNA. Four separate PCR reactions contained one of four primers that recognize human β-catenin sequences (5'-TTGATGGGCTGCCAGTACTG-3', 5'-CTACCAGTTGTGGTTAAGC-3', 5'-TGCACATCAGGATACCCAGC-3', or 5'-TATTGAAGCTGAGGGAGCCACAGC-3'), and a primer that recognizes sequences derived from exon 2 of the hGH gene (5'-GGCAGAGCAGGCCAAAAGCC-3'). The following conditions were used for PCR: denaturation at 95°C for 1 min; annealing at 65°C for 1 min; and extension at 72°C for 2 min using standard buffer and salt conditions (total = 30 cycles). Control reactions contained intestinal RNA prepared from normal control B6-ROSA26 129/Sv chimeras.

Immunoblots.
A segment was taken from the middle of the small intestine of B6-ROSA26 129/Sv(N89β-catenin) and B6-ROSA26 129/Sv mice and frozen immediately. After lyophilization, total cellular proteins were extracted by homogenizing the sample at 60°C in 1 ml of denaturation buffer (Laemmli, 1970) supplemented with protease inhibitors (1 µg/ml leupeptin, 1 µg/ml pepstatin, 4 mM 4-[2-aminoethyl]-benzensulfonyl fluoride hydrochloride, 1 µg/ml aprotinin, 1 mM EDTA). Insoluble debris was removed by centrifugation at 12,000 g for 10 min. The protein concentration in the resulting supernatant was defined (Hill and Straka, 1988). Samples of protein (100 µg) were fractionated by SDS-PAGE and electrophoretically transferred to polyvinylidene difluoride membranes. The membranes were stained with Ponceau S to verify successful transfer, preincubated for 1 h at 24°C with blocking buffer (PBS supplemented with 1% wt/vol gelatin and 0.5% vol/vol Tween-20) (Sigma Chemical Co., St. Louis, MO), and then probed for 1 h at 24°C with one of the following antibody preparations: (a) affinity-purified rabbit anti–c-myc epitope (MEQKLISEEDLN, 1:50 in PBS blocking buffer; Upstate Biotechnology Inc., Lake Placid, NY); (b) rabbit antisera raised against the COOH-terminal 14 residues of human/mouse β-catenin (PGDSNQLAWFDTDL, 1:200; provided by J. Nelson, Stanford University, Stanford, CA); and (c) rabbit anti-actin (1:5,000; Sigma Chemical Co.). Antigen–antibody complexes were detected using the reagents and protocols included in the Western Light kit^® from Tropix, Inc. (Bedford, MA).

Preparation and Staining of Intestinal Wholemounts
The small intestine was removed en bloc from mice immediately after killing and divided in half. Each half was gently flushed with ice-cold PBS, followed by periodate-lysine-paraformaldehyde (PLP; McLean and Nakane, 1974). Each half was opened with a single longitudinal incision along its mesenteric side and then subdivided into three equal parts. Each part was pinned onto a wax substrate and then fixed in PLP for 1 h at 24°C. The samples were washed with PBS (three cycles of 5 min each), incubated in 20 mM DTT/20% ethanol per 150 mM Tris-HCl, pH 8.0, for 45 min to remove mucus, and then washed again with PBS (three cycles). The epithelium was then genotyped by incubating all parts of the small intestine for 12 h at 4°C in PBS containing 2 mM X-Gal, 4 mM potassium ferricyanide, 4 mM potassium ferrocyanide, and 2 mM MgCl₂, final pH 7.6. The six regions of the small intestine were analyzed and photographed using a stereoscope (model SHZ10; Olympus, Tokyo, Japan), and then placed in 2% agar/PBS. Care was taken to preserve the orientation of crypt-villus units in each region while it was embedded in paraffin. Serial sections (5–8-µm-thick) were prepared along the cephalocaudal axis of each region (n = 300–500/region per animal).

Quantitation of Apoptotic and M-Phase Cells
Sections, prepared from the two middle regions of X-Gal–stained small intestine (jejunum), were counterstained with hematoxylin and eosin. M-phase and apoptotic cells were identified in each of these regions by their characteristic morphology (Hall et al., 1994; Coopersmith et al., 1997). Two 6-mo-old animals from each of three B6-ROSA26 129/ Sv(N89β-catenin) lines and three 6-mo-old B6-ROSA26 129/Sv mice were studied. A minimum of 200 well-oriented B6-ROSA26 and 200 well-oriented 129/Sv crypts were counted per mouse (well-oriented = crypt-sectioned parallel to the crypt-villus axis with Paneth cells represented at the crypt base and an unbroken epithelial column extending from the crypt base to the villus tip). A crypt had to be located within juxtaposed patches of B6-ROSA26 and 129/Sv crypt-villus units to be tallied. Each animal served as its own control; data from individual mice were expressed as the average number of M-phase (or apoptotic) cells per 129/Sv crypt divided by the average number of M-phase or apoptotic cells per B6-ROSA26 crypt. All slides were analyzed in a single-blinded fashion. Values obtained from animals from each line were averaged. Statistical analyses were conducted using unpaired Student's t-test.

Quantitation of Branched Villi
Sections obtained from the middle portions of the small intestine (see above), were stained with nuclear fast red and scored for the presence of branched villi. A minimum of 500–1000 villi in juxtaposed patches of B6-ROSA26 and 129/Sv crypt-villus units were scored per animal (n = two 6-mo-old mice/line of B6-ROSA26 129/Sv[N89β-catenin] chimeric-transgenic animals; n = three age-matched B6-ROSA26 129/Sv normal control chimeras). Branched villi were identified using the following criteria: (a) a villus had to have a continuous layer of epithelium over all of its branches; (b) immunostaining of an adjacent serial section with rabbit anti-laminin sera had to reveal a continuous ribbon of basement membrane underlying the uninterrupted epithelial sheet; and (c) the branched villus had to be present in adjacent serial sections.

Single and Multilabel Immunohistochemistry
Mice were injected with 5'-bromo-2'deoxyuridine (BrdU, 120 mg/kg body weight; Sigma Chemical Co.) and 5'-fluoro-2'-deoxyuridine (12 mg/kg; Sigma Chemical Co.) 60 h before killing. The entire small intestine was removed en bloc and wholemounts were prepared, fixed in PLP, stained with X-Gal, embedded in paraffin as described above, and then cut into 5-µm-thick sections. Alternatively, the middle third of the small intestine was flushed with PBS, fixed in PLP for 1 h at 24°C, and then flushed and embedded with O.C.T.(Miles Inc., Kankakee, IL). The embedded tissue was then frozen in Cytocool (VWR, St. Louis, MO) and 6–8-µm-thick sections were prepared.

Protocols used for single- and multilabel immunohistochemical studies have been described in our earlier publications (Hermiston et al., 1995a; 1996). PLP-fixed frozen sections of jejunum were stained with a 19-member panel of antibodies: (a) affinity-purified rabbit anti–Escherichia coli β-galactosidase (β-gal) (1:500; 5' 3' Inc., Boulder, CO); (b) rabbit anti– β-catenin sera (see above, final dilution in PBS/blocking buffer = 1:500); (c) affinity-purified rabbit anti–c-myc (see above, 1:100); (d) affinity-purified rabbit antibodies raised against amino acids 1034–2130 of human APC (APC2, a gift of P. Polakis; Näthke et al., 1996; Wong et al., 1996); (e) affinity-purified rabbit anti–-catenin (1:500; gift of J. Nelson); (f) a monoclonal rat antibody to E-cadherin (1:1,000; Sigma Chemical Co.; Hermiston et al., 1995a); (g) rat anti–ZO-1 (polyclonal antibodies, 1:50; Chemicon International, Inc., Temecula, CA); (h) rabbit anti-laminin (1: 1,000; Chemicon International Inc.); (i) rabbit anti-mouse fibronectin (1: 1,000; Chemicon International Inc.); (j) rabbit anti-mouse collagen type IV (1:1,000; Chemicon International Inc.); (k) rat anti-mouse β₁ integrin (1:500; PharMingen, San Diego, CA); (l) rat anti-mouse β₇ integrin (1:500; PharMingen); (m) rat anti-mouse β₄ integrin (1:500; PharMingen); (n) rat anti-mouse ₆ integrin (1:500; PharMingen); (o) goat anti-BrdU (1:1,000; Cohn et al., 1992); (p) rabbit anti-serotonin (1:1,000, a marker of the predominant enteroendocrine subpopulation in the adult mouse intestine; Incstar, Stillwater, MN); (q) rabbit anti-chromagranin A (1:1,000, a general marker of enteroendocrine cells; Incstar); and (r) rabbit anti-liver fatty acid binding protein (1:1,000, an enterocyte lineage marker; Sweetser et al., 1988).

Antigen–antibody complexes were detected with indocarbocyanine (Cy3)- or FITC-conjugated donkey anti–rabbit, anti–rat, or anti–goat Ig (1:500; Jackson ImmunoResearch Laboratories, Inc., West Grove, PA).

Analysis of the components of the diffuse gut–associated lymphoid tissue was performed on PLP-fixed frozen sections of jejunum. Antigen– antibody complexes were visualized using tyramide signal amplification as described previously (Garabedian et al., 1997).

PLP-fixed and X-Gal–stained sections of paraffin-embedded wholemounts were also incubated with a series of lectins (all used at a final concentration of 5 µg/ml PBS blocking buffer and detected according to Falk et al., 1994): (a) Ulex europaeus agglutinin 1 (carbohydrate specificity = Fuc1, 2Gal epitopes; lineage specificity = Paneth, goblet, and enteroendocrine cells; Sigma Chemical Co.); (b) Peanut (Arachis hypogaea) agglutinin (Galβ3GalNAc epitopes; all four epithelial lineages; Sigma Chemical Co.); (c) Dolichos biflorus agglutinin (GalNAc3GalNAc and GalNAc3Gal epitopes; Paneth and goblet cells plus enterocytes; Sigma Chemical Co.); (d) Helix pomentia agglutinin (-GalNAc; GalNAcβ4Gal epitopes; Paneth and goblet cells; Sigma Chemical Co.); (e) Jacalin-1 (Artrocarpus integrifolia agglutinin; Gal6Gal; Galβ3GalNAc epitopes; enterocytes and goblet cells; E.Y. Laboratories, Inc., San Mateo, CA).

Stained sections were viewed with an Axiophot microscope (Carl Zeiss, Inc., Thornwood, NY) and/or model 2001 confocal microscope (Molecular Dynamics, Inc., Sunnyvale, CA). Scans in the confocal microscope were performed at 1-µm intervals.

	Results

Top Abstract Materials and Methods Results Discussion Prospectus References

 
Generation of B6-ROSA26129/Sv(N89β-Catenin) Chimeric Mice
129/Sv ES cells were stably transfected with a recombinant DNA consisting of nucleotides -596 to +21 of Fabpl (Sweetser et al., 1988) positioned upstream of an open reading frame encoding N89β-catenin with an NH₂-terminal myc tag. The Fabpl transcriptional regulatory elements were selected because previous studies in neonatal and adult transgenic mice had shown that they could be used to direct expression of a variety of proteins to the region of small intestinal crypt containing the multipotent stem cell, and to all four of the stem cell's descendant cell lineages throughout the course of their differentiation (Trahair et al., 1989; Kim et al., 1993; Simon et al., 1993; Hermiston et al., 1996). Moreover, Fabpl-reporter transgene expression commences coincidently with initial cytodifferentiation of the intestinal epithelium in late fetal life and is sustained throughout adulthood, with highest levels of expression occurring in the middle third of the small bowel (jejunum).

Individual clones of stably transfected, Fabpl–N89β-catenin ES cells, or control nontransfected ES cells, were injected into C57Bl/6-ROSA26 (B6-ROSA26) blastocysts (Friedrich and Soriano, 1991; Wong et al., 1996). There are several reasons why the resulting chimeras are well- suited for studying the effects of transgene expression on the gut. First, intestinal epithelial cells of each genotype are separated into anatomically distinct units. By the time gut morphogenesis is completed, all crypts in B6-ROSA26 129/ Sv chimeras are monoclonal; they contain either B6-ROSA26 or 129/Sv epithelial cells, but not a mixture of both (Wong et al., 1996). Several crypts surround the villus base and supply epithelial cells to each villus. Thus, the chimeric small intestine will contain patches of transgenic 129/Sv crypt-villus units and adjacent patches of normal (nontransgenic) B6-ROSA26 crypt-villus units. A villus located at the border of a patch of B6-ROSA26 crypts and a patch of 129/Sv crypts will be polyclonal, containing a vertical coherent column of transgenic epithelial cells emanating from a monoclonal 129/Sv crypt and an adjacent column of normal epithelial cells emanating from a monoclonal B6 crypt (Fig. 1, A and B). Second, genotyping is simple. All undifferentiated and differentiated epithelial cells in B6-ROSA26 patches produce E. coli β-gal (Wong et al., 1996). Therefore, the B6-ROSA26 component can be identified by staining the opened chimeric small intestine with X-Gal (see blue-stained villi in Fig. 1 A). The 129/Sv epithelium does not produce β-gal and can be identified, even in chimeras with a low percentage 129/Sv contribution, by its white appearance. Third, regions of nontransgenic B6 epithelium serve as a critical internal control for defining the effects of the transgene's product. Since the small intestine is characterized by complex cephalocaudal variations in cell production and differentiation, it is critical that the control epithelial population be represented in a similar location as the genetically manipulated population. In the case of a polyclonal villus, the effect of the transgene can be defined by comparing 129/Sv and B6 cells located at a given cell stratum of a single villus positioned at a unique location along duodenal-ileal axis of an individual animal (Fig. 1 B).

View larger version (246K): [in this window] [in a new window] [Download PPT slide]   Figure 1 Jejunum from normal B6-ROSA26 129/Sv chimeric mice. (A) The jejunum has been opened, stained with X-Gal, and then photographed looking down on the tips of villi. The B6-ROSA26–derived epithelium expresses β-gal and appears blue after incubation with X-Gal. The 129/Sv component lacks β-gal (white). Closed arrow, wholly 129/Sv villus that receives cells from several monoclonal 129/Sv crypts positioned around its base. Open arrow, polyclonal villus. It appears striped because it receives columns of β-gal–negative (white) cells from monoclonal 129/Sv crypts and β-gal–positive cells from monoclonal B6-ROSA26 crypts. (B) Paraffin-embedded section prepared from the same wholemount and stained subsequently with nuclear fast red. The villus on the left is polyclonal: it is supplied by a monoclonal B6-ROSA26 crypt that contains an entirely β-gal–positive population of blue cells and by a monoclonal 129/ Sv crypt that only contains β-gal–negative cells. The villus on the right is only supplied by 129/Sv crypts. Bar, 25 µm.

View larger version (163K): [in this window] [in a new window] [Download PPT slide]   Figure 2 Evidence for expression of the Fabpl–N89β-catenin transgene. (A) RT-PCR analysis of RNAs isolated from the jejunum and skeletal muscle of a 6-wk-old-chimeric-transgenic mouse (N89β-catenin) and from the jejunum of a 6-wk-old chimera generated using nontransfected ES cells (nl chimera). The arrow points to the expected size of the product generated from N89β-catenin mRNA. (B) Duplicate immunoblots are shown each containing total cellular proteins isolated from the jejunum of a 6-wk-old normal control B6-ROSA26 129/Sv chimeric mouse and a 6-wk-old B6-ROSA26 129/Sv(N89β-catenin) chimeric-transgenic animal (both with 30–40% 129/Sv contribution based on coat color). The blot on the left of the panel was probed with antibodies that recognize the myc epitope tag present at the NH2 terminus of the 80-kD N89β-catenin mutant (arrow). The blot on the right of the panel was probed with antibodies raised against the COOH terminus of β-catenin (molecular mass of wild-type β-catenin = 91 kD). (C) PLP-fixed frozen section of a polyclonal villus from a B6-ROSA26 129/Sv(N89β-catenin) mouse. The section was incubated with affinity-purified rabbit antibodies to the myc tag, Cy3-tagged donkey anti–rabbit Ig, and the nuclear stain bis-benzimide (dark blue). Myc-tagged N89β-catenin (magenta) is prominently represented in 129/Sv but not B6-ROSA26 villus epithelial cells. (Genotyping was accomplished by staining an adjacent serial section with antibodies to β-gal.) (D) Section of a polyclonal villus from a normal chimeric mouse stained with the same reagents as in C. 129/Sv epithelial cells lack the transgene, and therefore do not contain any myc-tagged protein. Bars: (C and D) 25 µm.

A final confirmation of transgene expression was provided by immunohistochemistry. Serial sections of chimeric-transgenic and normal chimeric jejunums were stained with antibodies raised against the myc tag and β-gal. In chimeric-transgenic mice, immunoreactive myc was present in β-gal–negative 129/Sv epithelial cells, but not in β-gal–positive B6-ROSA26 epithelial cells (Fig. 2 C). In normal control chimeras, the myc tag was not detectable in either 129/Sv or B6-ROSA26 epithelium (Fig. 2 D).

Villus Branching in 129/Sv(N89β-Catenin) Epithelium
Chimeric-transgenic animals generated from three of the four ES cell lines were characterized further. All had similar phenotypes. There were no statistically significant differences between the growth rates or adult body weights of chimeric-transgenic and normal chimeric mice (n = 72).

Surveys of X-Gal–stained intestinal wholemounts from 1.5-, 6-, and 10-mo-old chimeric-transgenic mice (n = 1–8 animals/line per time point) revealed that the β-gal–negative 129/Sv(N89β-catenin) epithelium contained branched villi. Typically, there were two branches per villus, each of equivalent height. Branched villi were similar in height to surrounding unbranched villi (Fig. 3, A and B). When polyclonal villi were encountered with branching, there was always a wholly 129/Sv(N89β-catenin) branch (Fig. 3 C).

View larger version (83K): [in this window] [in a new window] [Download PPT slide]   Figure 3 Villus branching in 129/Sv(N89β-catenin) epithelium. (A) Wholemount preparation of the jejunum from a 6-mo-old chimeric-transgenic mouse. Arrow, branched 129/Sv villus. (B) Section from the same wholemount stained with nuclear fast red. Each limb of the branched villus is equivalent in height. There are no obvious morphologic abnormalities in the crypts associated with this villus (arrows). (C) A branched polyclonal villus. Open arrow, a stripe of β-gal–positive (blue) B6-ROSA26 epithelium in one of the branches. The other branch is wholly 129/Sv (closed arrow; white). Inset, section of the branched villus incubated with rabbit antibodies to β-gal and Cy3-conjugated donkey anti–rabbit Ig. The stripe of B6-ROSA26 epithelium in one of the branches appears red. Closed arrow, the other, β-gal–negative, 129/Sv branch. Bars: (B and C) 25 µm.

View larger version (12K): [in this window] [in a new window] [Download PPT slide]   Figure 4 Quantitation of villus branching. Branching was defined and quantitated as described in Materials and Methods. Branched villi were scored in the B6-ROSA26 and 129/Sv jejunal components of 6-mo-old normal chimeras (B6-ROSA26 129/ Sv) and chimeric-transgenic (B6-ROSA26 129/Sv[N89β-catenin]) mice. Each line of chimeric-transgenic mice was derived from an independent clone of ES cells stably transfected with Fabpl–N89β-catenin DNA. Mean values ±1 SD are plotted. Asterisk, the frequency of branched villi was <0.01%.

N89β-Catenin Expression Is Associated with Alterations in Proliferation, Apoptosis, and Migration but Not in Cell Fate Specification or Differentiation

View larger version (102K): [in this window] [in a new window] [Download PPT slide]   Figure 5 Forced expression of N89β-catenin has no discernible effects on epithelial cell differentiation. Frozen sections were prepared from PLP-fixed jejunums of 6-mo-old chimeric-transgenic animals. (A) A polyclonal villus stained with biotin-conjugated Dolichos biflorus agglutinin (DBA), Cy3-conjugated avidin, rabbit anti–β-gal, and FITC-conjugated donkey anti–rabbit Ig. Glycoconjugates containing GalNAc3GalNAc and GalNAc3Gal recognized by DBA appear yellow-orange; β-gal appears green. The polarity and differentiation of enterocytes appears to be unaffected, as judged by the distribution of these glycoconjugates in apical membranes and supranuclear Golgi apparatus (closed arrow). Similarly, based on their reaction with DBA, the number and differentiation of goblet cells (open arrows) is equivalent in the 129/Sv and B6-ROSA26 components of the polyclonal villus. (B) The base of a polyclonal villus with its crypt-villus junction indicated by closed arrows. Three crypts are seen (open arrows at their base): the one on the left is supplying cells to another villus. The section was incubated with rat anti-β4 integrin subunit, Cy3 donkey anti–rat Ig, rabbit anti–β-gal, FITC donkey anti–rabbit Ig, and bis-benzimide. Nuclei (blue); the β4 integrin subunit (orange); β-gal (green-brown). The location of β4 integrin at the base of epithelial cells and its distribution along the crypt-villus unit are unaffected by N89β-catenin. (C) Villi sectioned perpendicular to their crypt-villus axis. The tight junction protein ZO-1 (orange) was detected with rat anti-ZO-1 and Cy3 donkey anti–rat Ig. β-Gal (green) was visualized with the same reagents used in the preceding sections. The levels and location of ZO-1 in the 129/Sv(N89β-catenin) and B6-ROSA26 components of polyclonal villi are similar (e.g., open arrows). (D) Villi sectioned perpendicular to their crypt-villus axis as in C. The section was incubated with rat anti-β7 integrin, Cy3-donkey anti–rat Ig, rabbit anti-laminin, rabbit anti–β-gal, FITC donkey anti–rabbit Ig, and bis-benzimide. B6-ROSA26 cells exhibit diffuse staining of their cytoplasm due to the presence of β-gal (green). β7 integrin is confined to intraepithelial lymphocytes (orange). Comparable numbers of these cells are seen in the B6-ROSA26 and 129/ Sv(N89β-catenin) components of polyclonal villi and in wholly 129/Sv(N89β-catenin) villi. Laminin appears as linear green immunoreactivity underlying 129/Sv and B6-ROSA26 epithelium (e.g., closed arrows). The intensity of staining is similar under cells of both genotypes. Bars, 25 µm.

View larger version (19K): [in this window] [in a new window] [Download PPT slide]   Figure 6 N89β-catenin stimulates proliferation and apoptosis in 129/Sv crypts. (A) The number of M-phase cells in juxtaposed B6-ROSA26 and 129/Sv jejunal crypt sections were scored in 6-mo-old normal chimeras, in two lines of chimeric-transgenic mice generated using separate ES cell clones stably transfected with Fabpl–N89β-catenin DNA, and in two lines of chimeric-transgenic mice produced from independent ES cells clones stably transfected with Fabpl-mouse E-cadherin DNA (Hermiston et al., 1996). Each mouse served as its own control: the average number of M-phase cells in their 129/Sv crypts were divided by the average number of M-phase cells in their B6-ROSA26 crypts. Mean values ±1 SD obtained from animals in each group are plotted except for line 2 of B6 129/Sv (E-cadherin) chimeras where only two animals were examined. Note that the stimulation of crypt proliferation is similar in mice belonging to a given line of B6-ROSA26 129/Sv(N89β-catenin) chimeras. Differences between lines produced from different Fabpl–N89β-catenin ES cell clones were attributed to differences in the levels of expression of this transgene. (B) Apoptotic cells were scored in jejunal sections prepared from the same animals used to define the mitotic index in A.

The effect of N89β-catenin expression on cell migration was also examined. 6-mo-old chimeric-transgenic and normal control chimeric mice were injected with BrdU to label crypt epithelial cells in S phase. Animals were killed 60 h later (n = two mice/line). Serial sections were prepared from their jejunums and then the sections were stained with antibodies to β-gal and BrdU. Analysis of polyclonal villi present in chimeric-transgenic mice revealed a marked difference in migration between 129/ Sv(N89β-catenin) and B6-ROSA26 epithelial cells. 60 h after pulse labeling, the leading edge of BrdU-positive B6 cells had moved from the crypt to the upper quarter of the villus, whereas the leading edge of BrdU-positive 129/ Sv(N89β-catenin) cells had only reached the middle portion of the villus (Fig. 7). This effect was related to forced expression of N89β-catenin: control studies revealed that there were no detectable differences in migration between 129/Sv and B6-ROSA26 epithelial cells located in the polyclonal villi of normal chimeras (for illustration see Hermiston et al., 1996). Based on previous determinations of the time it takes BrdU-tagged cells to move from the crypt to the villus tip (Hermiston et al., 1996), the observed difference represents a slowing of 129/Sv(N89β-catenin) cell migration by 12–24 h.

View larger version (59K): [in this window] [in a new window] [Download PPT slide]   Figure 7 Forced expression of N89β-catenin is associated with a slowing of epithelial migration. A 6-mo-old chimeric-transgenic mouse was pulse labeled with BrdU 60 h before killing. Left, a section stained with rabbit anti–β-gal and Cy3 donkey anti–rabbit Ig. The villus encompassed by the box is polyclonal. Right, an adjacent section of the polyclonal villus, stained with goat anti-BrdU, Cy3 donkey anti–goat Ig, and bis-benzimide. The leading edge of the column of BrdU–positive 129/Sv(N89β-catenin) epithelial cells (magenta-colored nuclei) has only reached the middle portion of the villus (arrow), whereas the leading edge of the BrdU–positive B6-ROSA26 column is already near the villus tip. Bars, 25 µm.

The slowing of migration was not accompanied by any detectable perturbations in contacts between epithelial cells. There was no evidence of disrupted mucosal barrier function: (a) surveys of hematoxylin- and eosin-stained jejunal sections from 6-mo-old chimeric-transgenic mice failed to show any signs of inflammatory bowel disease; and (b) immunohistochemical analyses indicated that the number and crypt-villus distributions of several components of the diffuse gut–associated lymphoid tissue were unperturbed (e.g., β₇-positive intraepithelial lymphocytes, CD4⁺ T cells, and CD8⁺ T cells; refer to Fig. 5 D plus data not shown).

Because aberrant β-catenin signaling has been implicated in the pathogenesis of intestinal neoplasia (refer to Introduction), we carefully surveyed X-Gal–stained wholemounts and serial sections of the small intestine from chimeric-transgenic animals for evidence of dysplasia, adenoma formation, or adenocarcinoma. None of the mice had any evidence of these pathologic changes (n = 72; ages 1.5–10 mo).

Mechanistic Analyses: N89β-Catenin Expression Is Associated with an Augmentation in Cellular Pools of E-Cadherin
Conventional light microscopic and confocal microscopic surveys of jejunal sections, prepared from chimeric-transgenic mice and stained with antibodies against the myc tag or β-catenin, failed to disclose N89β-catenin and endogenous β-catenin in the nuclei of transgenic 129/Sv or normal B6-ROSA26 epithelial cells (refer to Fig. 2 C and see Fig. 8 A). β-Catenin was present at the adherens junctions and basolateral surfaces of epithelial cells (Fig. 8 A).

View larger version (67K): [in this window] [in a new window] [Download PPT slide]   Figure 8 N89β-Catenin expression results in augmented cellular pools of E-cadherin. Frozen sections were prepared from PLP-fixed jejunum recovered from a 6-mo-old B6-ROSA26 129/ Sv(N89β-catenin) chimeric-transgenic animal (A–D) and a 6-mo-old normal B6-ROSA26 129/Sv chimera (E and F). (A) Polyclonal villus incubated with rabbit anti–β-catenin, Cy3 donkey anti–rabbit Ig, and bis-benzimide. Prominent β-catenin staining (red) is evident at the adherens junctions and basolateral surfaces of epithelial cells. No immunoreactive protein is detectable in 129/Sv or B6-ROSA26 nuclei. (B) Polyclonal villus incubated with rabbit anti-APC, Cy3 donkey anti–rabbit Ig, and bis-benzimide. The levels and intracellular distribution of APC (red) are similar in B6-ROSA26 and 129/Sv(N89β-catenin) epithelial cells. (C and D) Villi from the chimeric-transgenic mouse that have been sectioned perpendicular their crypt-villus axis. (C) Section stained with rat anti–E-cadherin and Cy3-donkey anti– rat Ig. (D) Dual exposure of the same section after incubation with rabbit anti–β-gal and FITC donkey anti–rabbit Ig. Steady-state levels of E-cadherin are markedly increased in the 129/ Sv(N89β-catenin) component of polyclonal villi. (E and F) Villi from a control normal chimera processed as in (C and D). E-Cadherin levels are comparable in the B6-ROSA26 and 129/Sv villus epithelium. Bars, 25 µm.

In contrast, the steady-state level of E-cadherin was markedly increased in N89β-catenin–producing epithelial cells (Fig. 8, C and D). The augmented concentration of E-cadherin was not accompanied by detectable changes in its intracellular location (adherens junctions and basolateral surfaces). Analysis of polyclonal jejunal villi from normal chimeras established that the increased level of E-cadherin was not simply due to genotypic differences between 129/Sv and B6-ROSA26 epithelium (Fig. 8, E and F).

Previous studies, using the same Fabpl transcriptional regulatory elements used in this report, had shown that forced expression of wild-type E-cadherin in the jejunal crypt-villus units of chimeric-transgenic mice produced a slowing of epithelial migration equivalent to that observed with N89β-catenin (Hermiston et al., 1996). To determine whether augmented levels of E-cadherin were responsible for the other phenotypic changes observed B6-ROSA26 129/Sv(N89β-catenin) chimeras, we defined the jejunal crypt mitotic and apoptotic indices in 6-mo-old chimeric-transgenic mice generated using ES cells stably transfected with Fabpl mouse E-cadherin DNA (Hermiston et al., 1996). The increase in total M-phase cells/ crypt section associated with N89β-catenin expression was not evident in E-cadherin overexpressing crypts (refer to Fig. 6 A), indicating that this feature was directly related to N89β-catenin production rather than to a secondary effect of elevated E-cadherin concentrations. We could not make such a statement in the case of the apoptotic response, because similar elevations in crypt apoptosis were associated with forced expression of wild-type E-cadherin and N89β-catenin (Fig. 6 B). Forced expression of E-cadherin did not produce villus branching (data not shown).

	Discussion

Top Abstract Materials and Methods Results Discussion Prospectus References

 
Expression of N89β-catenin, which lacks sites for GSK-3 phosphorylation, can be viewed as simulating at least one effect of active Wnt signaling: generation of augmented cellular pools of stable, hypophosphorylated β-catenin. Comparable NH₂-terminal truncation mutants are known to phenocopy activation of the Wnt signaling pathway in Xenopus and Drosophila. In these nonmammalian systems, the consequences include axis duplication and changes in cell fate specification (Yost et al., 1996; Pai et al., 1997). Moreover, components in the Wnt signaling pathway play critical roles in specifying endoderm and gut differentiation during early Caenorhabditis elegans development (Han, 1997; Rocheleau et al., 1997; Thorpe et al., 1997). N89β-Catenin production in the small intestine of chimeric-transgenic mice was not only designed as an in vivo test of the role of Wnt signaling in establishing and maintaining a self-renewing mammalian epithelium, but also as a test of the effects of β-catenin on a system with complex adhesive requirements (i.e., preservation of cell– cell contacts in a mucosal barrier, orderly yet rapid cell migration, and exfoliation at a defined point in the cellular life cycle).

N89β-Catenin Expression Results in a Proliferative Response Restricted to Crypts
Forced expression of Wnt-1 causes mammary epithelial hyperplasia and tumor formation (Brown et al., 1986; Tsukamoto et al., 1988). In addition, ectopic expression of Wnt-1 enhances proliferation in the developing mammalian central nervous system (Dickinson et al., 1994). The mitogenic response of crypt epithelial cells to N89β-catenin is consistent with the observed mitogenic effects of active Wnt signaling in these other lineages. Surprisingly, this proliferative response was observed in the absence of detectable myc-tagged N89β-catenin (or endogenous β-catenin) within the nucleus of crypt epithelial cells. Forced expression of wild-type β-catenin or N89β-catenin in Drosophila, Xenopus, two-cell mouse embryos, and cultured mammalian cells has been reported to result in redistribution of β-catenin to the cytoplasm and nucleus (Funayama et al., 1995; Huber et al., 1996; Munemitsu et al., 1996; Yost et al., 1996; Pai et al., 1997). We assume that the amount of immunoreactive β-catenin in the nuclei of crypt epithelial cells was below the limits of detection of our staining techniques.

The proliferative response did not involve villus epithelial cells despite sustained expression of N89β-catenin from the crypt to the villus tip. Studies of nontransgenic mice have shown that these short-lived postmitotic cells retain many critical regulators that allow entry into S phase (e.g., cyclin E and cdk4; Chandrasekaran et al., 1996). Moreover, the villus microenvironment does not impose an absolute prohibition on crossing the G1/S boundary: e.g., villus enterocytes can be induced to undergo a pRB-dependent reentry into the cell cycle by forced expression of SV40 large T antigen (TAg; Chandrasekaran et al., 1996; Coopersmith et al., 1997). The reason why N89β-catenin fails to evoke a proliferative response in villus cells remains unresolved. However, terminal differentiation of these cells may change the availability or function of downstream effectors of the Wnt pathway, such as high motility group box transcription factors.

Primary Versus Secondary Responses: Stimulation of Apoptosis in N89β-Catenin–Producing Crypts and Villus Branching
The intestinal epithelium has a great capacity to initiate compensatory responses that preserve the steady-state cellular census when that census is threatened by changes in proliferative status or cell survival (Hermiston et al., 1995a; Coopersmith et al., 1997). As a consequence, it is often difficult to distinguish between primary responses to an applied stimulus and responses that compensate for an effect produced by that stimulus. At this point, we cannot determine whether the apoptosis reflects a direct effect of augmented cytosolic β-catenin pools (and signaling), or a secondary compensatory response to augmented crypt proliferation. Whatever the underlying mechanism, given the lack of discernible effects on villus height, we conclude that the observed changes in cell death are generally able to compensate for the observed changes in cell production.

A quandary exists regarding the interpretation of villus branching. There is little information available about the determinants of villus geometry (Totafurno et al., 1990). For example, are the determinants of morphology primarily epithelial-based or do they also involve mesenchymal signals? The normal variation in villus height observed along the duodenal-ileal axis of the adult mouse intestine can be correlated with the number of crypts that surround the base of each villus (Wright and Irwin, 1982). Totafurno et al. (1987) examined rare branched villi in normal mice and found that the number of crypts that surrounded their base was approximately twice the normal number, suggesting that villus branching may be a compensatory response to increased epithelial cell input. Increased cell production may not have to come from crypts: SV40 TAg expression in villus enterocytes leads to reentry into the cell cycle and branching in a subset of villi (Coopersmith et al., 1997).

We do not view branching as analogous to the axis duplication observed in Xenopus when the Wnt pathway is stimulated. It is unclear how a branched shape can be maintained given the perpetual epithelial cell renewal that occurs in crypt-villus units. In fact, the absence of a notable increase in the frequency of villus branching as B6-ROSA26 129/Sv(N89β-catenin) mice age suggests that branched villi have a limited life span. This is consistent with the view that normal intestine continually produces new crypts and villi as a byproduct of its dynamic cell renewal (Totafurno et al., 1987). Branching in our chimeric-transgenic mice may be a default response, reflecting the inability of some of their villi to adequately compensate for a threatened or real increase in their steady-state epithelial cell census beyond some critical threshold value. Such a threat may arise from the increased cell production observed in their crypts, from the inability of crypt apoptosis to compensate for this proliferative response, and/or from the effects of slowed epithelial cell migration. Branching could also represent a direct response to aberrant signals, originating from a genetically manipulated epithelium and operating through interactions with the underlying mesenchyme. The significance of branching is that it provides a potential inroad to deciphering how villus structure is preserved in the face of perpetual, rapid renewal of its principal cellular component.

N89β-Catenin Produces an Increase in Steady-State Levels of E-Cadherin
Forced expression of N89β-catenin results in a marked increase in E-cadherin levels at the adherens junction and basolateral surfaces of intestinal epithelial cells. Several recent studies suggest that this is due to activation of E-cadherin gene expression. The mouse E-cadherin gene contains a 7-bp binding sequence for Lef-1/Tcf (Huber et al., 1996). Yanagawa and co-workers (1997) used a Drosophila wing imaginal disc cell line to show that stimulation of the Wingless pathway induces Drosophila E-cadherin gene transcription and accumulation of DE-cadherin at cellular junctions. They also found that forced expression of Dishevelled, an inhibitor of Zeste-white 3 (the Drosophila GSK-3 homologue), led to accumulation of Armadillo (the β-catenin homologue) in the cytosol of these cells, a marked increase in Drosophila E-cadherin mRNA, and an elevation in junctional DE-cadherin. In addition, they noted that forced expression of an NH₂-terminal truncation mutant of Armadillo produces elevated DE-cadherin mRNA and protein levels.

The N89β-catenin–mediated increase in E-cadherin levels is likely to counteract the signaling activity of N89β-catenin within the intestinal epithelium. Studies in Xenopus have shown that the binding of β-catenin to cadherins opposes signaling by sequestering β-catenin (Heasman et al., 1994; Fagotto et al., 1996). The location of myc-tagged N89β-catenin within intestinal epithelial cells provides evidence that a similar process occurs in a mammalian system.

The augmentation in cellular E-cadherin pools is also likely to contribute to the slow migratory phenotype of 129/Sv(N89β-catenin) epithelium. When forced expression of E-cadherin was limited to villus enterocytes in transgenic mice, enterocytic migration from the villus base to villus tip was slowed (Hermiston et al., 1996). In this previous study, there was no augmentation of E-cadherin in the crypt epithelium, no perturbations in crypt proliferation or death, and therefore little likelihood that the observed effect on migration was due to a diminution in net cellular output from the crypt.

APC and the Effects of N89β-Catenin on Intestinal Epithelial Biology
N89β-Catenin production was not associated with any detectable abnormalities in the intracellular distribution or levels of APC within crypt-villus units. This may have contributed to two phenotypic features of our chimeric-transgenic mice: (a) slowed migration and (b) lack of neoplastic transformation.

Studies in MDCK cells have indicated that β-catenin affects APC's ability to organize microtubules (Näthke et al., 1996; Pollack et al., 1997). N89β-catenin–APC complexes are more stable than wild-type β-catenin–APC complexes, leading to accumulation of N89β-catenin at clusters of APC positioned at the tips of MDCK cell membrane extensions (Näthke et al., 1996; Barth et al., 1997; Pollack et al., 1997). Based on these findings, Pollack and co-workers (1997) proposed that this stabilization of N89β-catenin–APC complexes inhibits (MDCK) migration by somehow perturbing APC function.

Regulation of β-catenin degradation appears to play an important role in colorectal tumorigenesis and in the formation of melanomas (Korinek et al., 1997; Morin et al., 1997; Rubinfeld et al., 1997). Levels of soluble β-catenin are elevated in colon tumor cells (Munemitsu et al., 1995). In addition, mutations in the NH₂ terminus of β-catenin are oncogenic in cultured cells and are associated with intestinal neoplasms (Morin et al., 1997). The stabilized N89β-catenin mutant was produced in intestinal epithelial cells that were able to maintain normal levels of APC. This may account for the failure of these cells to undergo neoplastic transformation during the 10-mo period that animals were studied. In addition, the β-catenin mutations that have been associated with human colorectal cancer involve alterations in NH₂-terminal serine residues. Although it may seem, superficially, that NH₂-terminal truncations would have similar biochemical effects as serine substitution (i.e., protein stabilization), NH₂-terminal phosphorylation may have other roles, including affects on protein–protein interactions that modulate signaling.

	Prospectus

Top Abstract Materials and Methods Results Discussion Prospectus References

 
The self-renewing intestinal epithelium is able to invoke a variety of compensatory responses when its census is threatened. This capacity to compensate makes sense given the enormous energetic investments required to sustain normal self-renewal. Compensation provides an experimental challenge when trying to decipher the function of molecules such as β-catenin, since dramatic or even noticeable phenotypes may be difficult to elicit. Further analysis of the effects of active β-catenin–mediated signaling on epithelial homeostasis may require forced expression of β-catenin–Lef-1 (or Tcf) fusion proteins or dominant-negative Tcf/Lef-1 mutants (Molenaar et al., 1996). The chimeric-transgenic system described above provides a way of assaying the effects of these mutants in vivo under well-controlled conditions.

	Acknowledgments

 
We thank D. O'Donnell (Washington University School of Medicine) for expert technical assistance. We are grateful to J. Nelson and P. Polakis for supplying reagents and for their many valuable suggestions during the course of this work.

These studies were supported in part by grants from the National Institutes of Health (DK37960, DK30292, and T32 HlO7275).

Submitted: 4 February 1998

Revised: 11 March 1998

Address all correspondence to Jeffrey I. Gordon, Department of Molecular Biology and Pharmacology, Box 8103, Washington University School of Medicine, 660 South Euclid Ave., St. Louis, MO 63110. Tel.: (314) 362-7243. Fax: (314) 362-7047. E-mail: jgordon{at}pharmdec.wustl.edu

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