Regulation of β-Catenin Signaling in the Wnt Pathway

Regulation of β-Catenin Signaling in the Wnt Pathway

Biochemical and Biophysical Research Communications 268, 243–248 (2000) doi:10.1006/bbrc.1999.1860, available online at on ...

157KB Sizes 2 Downloads 149 Views

Biochemical and Biophysical Research Communications 268, 243–248 (2000) doi:10.1006/bbrc.1999.1860, available online at on

BREAKTHROUGHS AND VIEWS Regulation of ␤-Catenin Signaling in the Wnt Pathway Akira Kikuchi 1 Department of Biochemistry, Hiroshima University School of Medicine, 1-2-3, Kasumi, Minami-ku, Hiroshima 734-8551, Japan

Received November 11, 1999

␤-Catenin not only regulates cell to cell adhesion as a protein interacting with cadherin, but also functions as a component of the Wnt signaling pathway. The Wnt signaling pathway is conserved in various organisms from worms to mammals, and plays important roles in development, cellular proliferation, and differentiation. Wnt stabilizes cytoplasmic ␤-catenin and then ␤-catenin is translocated into the nucleus where it stimulates the expression of genes including c-myc, c-jun, fra-1, and cyclin D1. The amounts and functions of ␤-catenin are regulated in both the cytoplasm and nucleus. Its molecular mechanisms are becoming increasingly well understood. © 2000 Academic Press

phorylation of ␤-catenin is reduced and ␤-catenin is no longer degraded, resulting in its accumulation in cytoplasm. Accumulated ␤-catenin is translocated into the nucleus where it binds to Tcf/Lef, a transcription factor, and stimulates gene expression. In the nucleus several proteins that bind to ␤-catenin and Tcf/Lef regulate the complex formation of ␤-catenin-Tcf-DNA. Therefore, it appears that ␤-catenin signaling is regulated in both the cytoplasm and nucleus. In this review, regulation of ␤-catenin signaling is described. 2. REGULATION OF ␤-CATENIN SIGNALING BY AXIN IN THE CYTOPLASM 2.1. Identification of Axin

1. Wnt SIGNALING PATHWAY Wnt proteins constitute a large family of cysteinerich secreted ligands that control development in organisms ranging from nematode worms to mammals (1). The outlines of the Wnt signal transduction pathway were first elucidated by a genetic analysis of Wingless signaling during the development of segmental polarity in Drosophila, and extended through studies of embryonic axis formation in Xenopus. In vertebrates, a number of components of the Wingless pathway are conserved and the Wnt signaling pathway regulates organ development and cellular proliferation, morphology, motility, and fate (2– 4). In the current model, the serine/threonine kinase glycogen synthase kinase-3␤ (GSK-3␤) targets cytoplasmic ␤-catenin for degradation in the absence of Wnt (Fig. 1). As a result, cytoplasmic ␤-catenin levels are low. When Wnt acts on its cell-surface receptor Frizzled, Dvl, a cytoplasmic protein with unknown functions, is activated. Dvl antagonizes the action of GSK-3␤, perhaps by modulating its enzymatic activity. The phosFax: ⫹81-82-257-5134. E-mail: [email protected] 1

Axin was originally identified as the product of the mouse gene Fused (5). An intriguing feature of many homozygous axin mutant embryos is a duplication of the embryonic axis (6, 7), suggesting that Axin normally plays a negative regulatory role in the response to an axis-inducing signal. Indeed, when Axin is injected into Xenopus embryos, most embryos develop with strong axial defects (5). Several experiments indicate that Axin exerts its function on axis formation by specifically inhibiting the Wnt signaling (5). Independent biochemical approaches have demonstrated the existence of Axin in various species. Axin and its homolog, Axil (Axin like), have been identified as proteins that bind to GSK-3␤ (8, 9). Another Axin homolog, conductin, has been identified as a ␤-cateninbinding protein (10). Conductin and Axil are the same protein, implying that there are at least two Axin family members in mammals. The Axin gene is conserved in humans, rats, mice, chickens, and Xenopus (5, 8, 11). Human Axin and Axil are located in 16q13-3 and 17q23-q24, respectively (12). When the amino acid numbers of Axin are indicated in this review, they refer to rat Axin (rAxin). A homolog of the vertebrate Axin has been also found in Drosophila, and named D-Axin (13, 14). Genetic evidence demonstrates that Wingless


0006-291X/00 $35.00 Copyright © 2000 by Academic Press All rights of reproduction in any form reserved.

Vol. 268, No. 2, 2000


FIG. 1. Mechanism by which Wnt regulates the stability of ␤-catenin. ␤-Catenin is present in the Axin complex in the absence of Wnt. In this complex, ␤-catenin is phosphorylated, ubiquitinated, and degraded by proteasome. Dvl antagonizes Axin activity in response to Wnt, and ␤-catenin is dissociated and accumulated in the cytosol. The accumulated ␤-catenin is translocated into the nucleus, and binds to and activates Tcf/Lef, resulting in expression of the target genes. E1, ubiquitin-activating enzyme; E2, ubiquitin-conjugating enzyme; E3, ubiquitin ligase; Ub, ubiquitin.

signaling is activated in embryos lacking D-Axin and that the ectopic expression of D-Axin suppressed Wingless signaling. Therefore, the functions of Axin are conserved in mammals and flies. 2.2. Complex Formation of GSK-3␤ and ␤-Catenin with Axin The central region of Axin contains the GSK-3␤- and ␤-catenin-binding sites, and residues 353-437 and 437506 of rAxin are responsible for the binding of GSK-3␤ and ␤-catenin, respectively (8) (Fig. 2). GSK-3 was originally characterized as a serine/threonine kinase that phosphorylates and inactivates glycogen synthase and is now implicated in the regulation of several physiological responses in mammalian cells by phosphorylating many substrates (15). The cDNAs of mammalian GSK-3␣ and GSK-3␤ have been isolated (16). Although both GSK-3␣ and GSK-3␤ form complexes with Axin (8), GSK-3␤ is mainly used in analyses of the Wnt signaling pathway, since it but not GSK-3␣ rescues the phenotype of the Drosophila zw3/shaggy gene product, a GSK-3 homolog (17). ␤-catenin was originally identified as a protein which interacts with the cytoplasmic domain of cadherin and links cadherin to ␣-catenin,

which in turn mediates the anchorage of the cadherin complex to the cortical actin cytoskeleton (18). Genetic and embryological studies have revealed that ␤-catenin is a component of the Wnt signaling pathway and that it exhibits signaling functions (2– 4). Axin interacts directly with the region containing Armadillo repeats 2 to 7 of ␤-catenin (8). GSK-3␤ and ␤-catenin bind simultaneously to different sites of Axin, forming a ternary complex (8). ␤-catenin has a consensus sequence of a phosphorylation site for GSK-3␤ (19, 20),

FIG. 2. Structure of Axin and its binding proteins. Axin possesses binding sites for APC, GSK-3␤, ␤-catenin, Dvl, and PP2A. In the Axin complex, GSK-3␤ efficiently phosphorylates ␤-catenin, APC, and Axin itself, while Dvl and PP2A prevent the phosphorylation.


Vol. 268, No. 2, 2000


and Axin greatly enhances the phosphorylation of ␤-catenin by GSK-3␤ under conditions in which these three proteins form a complex (8, 21). Axil also enhances the phosphorylation of ␤-catenin by GSK-3␤ (9). Further, Axin and Axil are substrates of GSK-3␤ and the phosphorylation sites are close to the GSK-3␤binding region. It has been shown that phosphorylation of Axin regulates its stability and its affinities for ␤-catenin and GSK-3␤ (22–24). ␤-Catenin is a target for the ubiquitin-proteasome pathway and the phosphorylation by GSK-3␤ is required for the ubiquitination (25). In general, degradation of proteins by the ubiquitin-proteasome pathway involves a ubiquitin-activation enzyme (E1), a ubiquitin-conjugating enzyme (E2), and a ubiquitin ligase (E3) (26). An F-box protein is a component of E3 and serves as a receptor for the target proteins which are usually phosphorylated (27, 28). In Drosophila, a mutation of F-box protein Slimb leads to accumulation of Armadillo, a ␤-catenin homolog (29). Consistent with these genetic findings, ␤TrCP/FWD1, a mammalian homolog of Slimb, associates with ␤-catenin in the presence of Axin and stimulates ubiquitination (30 – 32). Thus, the phosphorylation of ␤-catenin by GSK-3␤ and its ubiquitination are enhanced in the Axin complex. The observations that Axin promotes the phosphorylation and ubiquitination of ␤-catenin suggest that Axin regulates the stability of ␤-catenin. Indeed, expression of Axin and Axil (conductin) in SW480 or COS cells stimulates the degradation of ␤-catenin (10, 33, 34). Wnt-3a induces the accumulation of ␤-catenin in mouse fibroblast L cells, and the Wnt-3a-dependent increase of ␤-catenin is inhibited in L cells stably expressing Axin (35). Wnt-3a stimulates Tcf-4 transcriptional activity through ␤-catenin, and expression of Axin inhibits Wnt-3a-dependent Tcf-4 activity (35, 36). Furthermore, expression of Axin in SW480 cells inhibits cellular proliferation (21). Therefore, Axin negatively regulates the Wnt-signaling pathway by downregulating ␤-catenin. Axin also binds to plakoglobin, a homolog of ␤-catenin, and enhances its phosphorylation by GSK-3␤, resulting in the degradation of plakoglobin (37). 2.3. Interaction of APC with Axin APC is a tumor suppressor linked to familial adenomatous polyposis coli and to the initiation of sporadic human colorectal cancer (38). APC encodes a 300-kDa multifunctional protein with several structural domains. The middle portion of APC contains three successive 15-amino-acid (15-aa) repeats followed by seven related but distinct 20-aa repeats, both of which are able to bind independently to ␤-catenin (39 – 41). The APC activity to downregulate the level of cytoplasmic ␤-catenin is localized to the central region of the

protein, and at least three 20-aa repeats are necessary for the degradation of ␤-catenin (38, 42). Axin and Axil (Conductin) have the RGS domains in their N-termini (5, 8 –10). The RGS domains of Axin and conductin interact directly with the region containing the third to seventh 20-aa repeats of APC (10, 33, 34) (Fig. 2). Most of the APC mutants observed in human colon cancer cells, where cytoplasmic ␤-catenin is accumulated, have lost the region containing 20-aa repeats and do not bind to Axin (34). Therefore, the interaction of APC with Axin may be important for the APC activity to downregulate ␤-catenin. GSK-3␤ phosphorylates APC directly (43), and the binding of APC to Axin enhances GSK-3␤-dependent phosphorylation (33, 44). Although the significance of the phosphorylation of APC is not known, the phosphorylation of APC appears to be important for the degradation of ␤-catenin (42, 43). Thus, GSK-3␤ and its substrates, ␤-catenin, APC, and Axin itself, are simultaneously present in the Axin complex and their phosphorylation occurs efficiently in the complex. The protein stability and functions of ␤-catenin, APC, and Axin are regulated in the complex. 2.4. Regulation of Phosphorylation by Dvl and PP2A in the Axin Complex Dvl is a mammalian homolog of fly dishevelled (dsh) and acts negatively upstream of shaggy (2, 3, 45, 46). Dvl-1, -2, and -3 genes have been isolated (47– 49). All Dsh and Dvl family members contain three highly conserved domains: an N-terminal DIX domain; a central PDZ domain, and a DEP domain (2, 3). The PDZ domain is essential for the roles of Dvl in the Wnt/ Wingless signaling pathway (50 –52). The DEP domain is critical for rescue of the dsh planar polarity defect and for the activation of the Jun-N-terminal kinase (53, 54). Dvl binds to Axin (21, 55–57) and inhibits GSK-3␤-dependent phosphorylation of ␤-catenin and APC in the presence of Axin (21). Furthermore, Dvl inhibits the phosphorylation of Axin by GSK-3␤ (22). These results suggest that Dvl modulates Axin activity by their interaction and are consistent with the observation that Dvl antagonizes the ability of Axin to inhibit axis formation in Xenopus embryos (5). The binding of Dvl to Axin may induce conformational changes of the Axin complex, which result in ineffective phosphorylation by GSK-3␤ of its substrates. Protein phosphatase 2A (PP2A) is one of the four major serine/threonine protein phosphatases (58). The catalytic subunit of PP2A (PP2Ac) is always associated with a regulatory subunit of 65 kDa (PR65 or A subunit). To this dimeric core, various third or variable regulatory subunits (B subunits) bind and modulate the enzymatic activity of PP2Ac (59). Axin forms a complex with PP2A, and PP2Ac binds directly to the region containing amino acids 298-506 or 508-832 of rAxin, but the A subunit of PP2A alone does not (44,


Vol. 268, No. 2, 2000


60) (Fig. 2). PP2A bound to Axin dephosphorylates APC and Axin phosphorylated by GSK-3␤ in the Axin complex (44). The B⬙ subunit of PP2A interacts with APC, and expression of the B⬙ subunit reduces the level of ␤-catenin and inhibits the transcription of the ␤-catenin target gene (61). These results suggest that the B⬙ subunit enhances the phosphorylation of ␤-catenin by inhibiting the catalytic activity of PP2Ac. Therefore, PP2A is present in the Axin complex and may regulate the phosphorylation of the substrates of GSK-3␤. 2.5. Axin as a Scaffold Protein Axin may be a scaffold protein, in that it binds to several signaling molecules to create a multi-enzyme complex (Fig. 1). In mammalian cells such as COS or L cells, Axin interacts with GSK-3␤, ␤-catenin, and APC in a high molecular mass complex of more than 10 3 kDa on gel filtration column chromatography (35). ␤-catenin is present in the high molecular mass complex in the absence of Wnt-3a, while addition of Wnt-3a to the cells increases ␤-catenin in a lower molecular mass complex of 200-300 kDa (35). In cells overexpressing Axin, the Wnt-3a-induced increase of ␤-catenin in the low molecular mass complex is not observed (35). These results suggest that the balance between the high and low molecular mass complexes containing ␤-catenin is tightly regulated, and that Axin plays a role in limiting the accumulation of ␤-catenin in the low molecular mass complex. Therefore, Wnt may regulate the assembly of the complex consisting of Axin, APC, ␤-catenin, and GSK-3␤, and induce the dissociation of ␤-catenin from the complex. It is possible that ␤-catenin free from the complex is accumulated, binds to different partners such as Tcf/ Lef, and thereby transmits transcription regulatory signals. Three possible mechanisms by which ␤-catenin is dissociated from the complex have been proposed. The first one is that Wnt-dependent dephosphorylation of Axin induces the degradation of Axin (22). Degradation of Axin due to hypophosphorylation may induce the dissociation of ␤-catenin from the complex. The second one is that Wnt-induced dephosphorylation of Axin decreases its affinities for ␤-catenin and GSK-3␤ (23, 24). The third possibility is that Frat1 is involved in the Wnt signaling (57). Frat1 was originally cloned for its tumor promoting activity in lymphocytes (62), and is a homolog of Xenopus GBP, which binds to GSK-3␤ and inhibits GSK-3␤-dependent phosphorylation (63). Dvl interacts with Axin and Frat1. Wnt-1 promotes the disintegration of the Frat1/Dvl/GSK-3␤/Axin complex, resulting in the dissociation of GSK-3␤ from Axin and in the stabilization of ␤-catenin. In any case, ␤-catenin is dissociated from the Axin complex in response to

Wnt. Therefore, Axin may act as a scaffold protein to selectively channel the signal from Wnt to ␤-catenin. 3. REGULATION OF ␤-CATENIN SIGNALING IN THE NUCLEUS 3.1. Tcf-Binding Proteins

␤-catenin dissociated from the high molecular weight Axin complex enters the nucleus and forms a complex with Tcf to activate transcription of Wnt target genes (Fig. 3). Several target genes have been identified, including siamois in Xenopus (64), Ubx in Drosophila (65), and c-myc, c-jun, fra-1, and cyclin D1 in mammals (66 – 68). These genes have Tcf-binding sites near or in their promoters. It is thought that Tcf may be a transcriptional repressor rather than an activator, because it binds to proteins that can mediate repression. One repressor is Groucho in Drosophila (69). The binding sites for Armadillo and Groucho on Tcf do not overlap, but whether or not Armadillo and Groucho bind simultaneously to Tcf is not clear. It is possible that expression of Tcf-target genes is regulated by a balance between Armadillo and Groucho. Another Tcfbinding protein is Drosophila CBP (70). CBP interacts with the high-mobility group domain of Tcf and acetylates a conserved lysine in the Armadillo-binding domain of Tcf. This acetylation lowers the affinity of Tcf for Armadillo. It is possible that similar repressors regulate the Tcf activity in mammals. It has been shown that NEMO-like kinase (NLK) binds directly to and phosphorylates Tcf and that the phosphorylation of Tcf inhibits the binding of the ␤-catenin/Tcf complex to DNA (71). Therefore, these Tcf-binding proteins negatively regulate the ␤-catenin signaling. 3.2. ␤-Catenin-Binding Proteins in the Nucleus Pontin52 is a nuclear protein which binds to ␤-catenin (72). Pontin52 can be coimmunoprecipitated within a large complex containing ␤-catenin and Lef-1 and is proposed to provide a bridging function between Lef-1/␤-catenin complexes and transcriptional machinery (72). However, whether Pontin52 affects the ␤-catenin signaling remains to be clarified. Duplin is a new ␤-catenin-binding protein found in the nucleus 2 (Fig. 3). Duplin does not affect the stability or subcellular localization of ␤-catenin, but competes with Tcf-4 for the binding to ␤-catenin. Consistent with these characteristics, Duplin inhibits Wnt-3a- and ␤-catenindependent Tcf-4 activation in L cells and suppresses Wnt- and ␤-catenin-induced axis formation in Xenopus embryos. It appears that Duplin inhibits the ␤-catenin signaling in a manner different from Groucho, CBP, and NLK. 2

Sakamoto, I., Kishida, S., Fukui, A., Kishida, M., Yamamoto, H., Michiue, T., Takada, S., Asashima, M., Kikuchi, A., submitted.


Vol. 268, No. 2, 2000


FIG. 3. ␤-Catenin complexes in the cytoplasm and nucleus. Axin regulates the intracellular distribution of ␤-catenin between the cytosol and nucleus.

4. CONCLUSION There are multiple mechanisms to inhibit ␤-catenin signaling. The findings suggest that two complexes containing ␤-catenin, the Axin and Tcf complexes, exist in the cytoplasm and nucleus, respectively (Fig. 3). Wnt may regulate the subcellular distribution of ␤-catenin between the cytoplasm and nucleus. In the cytoplasm, the amount of ␤-catenin is negatively regulated by the degradation of ␤-catenin in the Axin complex. In the nucleus, gene expression induced by ␤-catenin is negatively regulated by inhibiting the complex formation of ␤-catenin, Tcf, and DNA. Mutations in ␤-catenin have been found in human cancers, including colon cancer and melanoma, and the mutations result in the accumulation of ␤-catenin (73, 74). Since ␤-catenin functions as an oncogene, it is speculated that there are several mechanisms for protecting against abnormal cellular proliferation by inhibiting ␤-catenin signaling. ACKNOWLEDGMENTS I thank my colleagues, S. Koyama, S. Kishida, M. Kishida, S. Ikeda, H. Yamamoto, S. Hino, I. Sakamoto, S. Kodama, K. Matsubara, and S. Nakashima.

REFERENCES 1. Nusse, R., and Varmus, H. E. (1992) Cell 69, 1073–1087. 2. Cadigan, K. M., and Nusse, R. (1997) Genes Dev. 11, 3286 –3305.

3. Dale, T. C. (1998) Biochem. J. 329, 209 –223. 4. Miller, J. R., and Moon, R. T. (1996) Genes Dev. 10, 2527–2539. 5. Zeng, L., Fagotto, F., Zhang, T., Hsu, W., Vasicek, T. J., Perry, W. L., III, Lee, J. J., Tilghman, S. M., Gumbiner, B. M., and Costantini, F. (1997) Cell 90, 181–192. 6. Gluecksohn-Schoenheimer, S. (1949) J. Exp. Zool. 110, 47–76. 7. Jacobs-Cohen, R. J., Spiegelman, M., Cookingham, J. C., and Bennett, D. (1984) Genet. Res. 43, 43–50. 8. Ikeda, S., Kishida, S., Yamamoto, H., Murai, H., Koyama, S., and Kikuchi, A. (1998) EMBO J. 17, 1371–1384. 9. Yamamoto, H., Kishida, S., Uochi, T., Ikeda, S., Koyama, S., Asashima, M., and Kikuchi, A. (1998) Mol. Cell. Biol. 18, 2867– 2875. 10. Behrens, J., Jerchow, B.-A., Wu¨rtele, M., Grimm, J., Asbrand, C., Wirtz, R., Ku¨hl, M., Wedlich, D., and Birchmeier, W. (1998) Science 280, 596 –599. 11. Hedgepeth, C. M., Deardorff, M. A., and Klein, P. S. (1999) Mech. Dev. 80, 147–151. 12. Mai, M., Qian, C., Yokomizo, A., Smith, D. I., and Liu, W. (1999) Genomics 55, 341–344. 13. Hamada, F., Tomoyasu, Y., Takatsu, Y., Nakamura, M., Nagai, S.-I., Suzuki, A., Fujita, F., Shibuya, H., Toyoshima, K., Ueno, N., and Akiyama, T. (1999) Science 283, 1739 –1742. 14. Willert, K., Logan, C. Y., Arora, A., Fish, M., and Nusse, R. (1999) Development 126, 4165– 4173. 15. Plyte, S. E., Hughes, K., Nikolakaki, E., Pulverer, B. J., and Woodgett, J. R. (1992) Biochim. Biophys. Acta 1114, 147–162. 16. Woodgett, J. R. (1990) EMBO J. 9, 2431–2438. 17. Ruel, L., Bourouis, M., Heitzler, P., Pantesco, V., and Simpson, P. (1993) Nature 362, 557–560. 18. Takeichi, M. (1991) Science 251, 1451–1455. 19. Munemitsu, S., Albert, I., Rubinfeld, B., and Polakis, P. (1996) Mol. Cell. Biol. 16, 4088 – 4094.


Vol. 268, No. 2, 2000


20. Yost, C., Torres, M., Miller, J. R., Huang, E., Kimelman, D., and Moon, R. T. (1996) Genes Dev. 10, 1443–1454. 21. Kishida, S., Yamamoto, H., Hino, S., Ikeda, S., Kishida, M., and Kikuchi, A. (1999) Mol. Cell. Biol. 19, 4414 – 4422. 22. Yamamoto, H., Kishida, S., Kishida, M., Ikeda, S., Takada, S., and Kikuchi, A. (1999) J. Biol. Chem. 274, 10681–10684. 23. Willert, K., Shibamoto, S., and Nusse, R. (1999) Genes Dev. 13, 1768 –1773. 24. Ruel, L., Stambolic, V., Ali, A., Manoukian, A. S., and Woodgett, J. R. (1999) J. Biol. Chem. 274, 21790 –21796. 25. Aberle, H., Bauer, A., Stappert, J., Kispert, A., and Kemler, R. (1997) EMBO J. 16, 3797–3804. 26. Ciechanover, A. (1994) Cell 79, 13–21. 27. Skowyra, D., Craig, K. L., Tyers, M., Elledge, S. J., and Harper, J. W. (1997) Cell 91, 209 –219. 28. Feldman, R. M., Correll, C. C., Kaplan, K. B., and Deshaies, R. J. (1997) Cell 91, 221–230. 29. Jiang, J., and Struhl, G. (1998) Nature 391, 493– 496. 30. Kitagawa, M., Hatakeyama, S., Shirane, M., Matsumoto, M., Ishida, N., Hattori, K., Nakamichi, I., Kikuchi, A., Nakayama, K.-I., and Nakayama, K. (1999) EMBO J. 18, 2401–2410. 31. Winston, J. T., Strack, P., Beer-Romero, P., Chu, C. Y., Elledge, S. J., and Harper, J. W. (1999) Genes Dev. 13, 270 –283. 32. Latres, E., Chiaur, D. S., and Pagano, M. (1999) Oncogene 18, 849 – 854. 33. Hart, M. J., de los Santos, R., Albert, I. N., Rubinfeld, B., and Polakis, P. (1998) Curr. Biol. 8, 573–581. 34. Kishida, S., Yamamoto, H., Ikeda, S., Kishida, M., Sakamoto, I., Koyama, S., and Kikuchi, A. (1998) J. Biol. Chem. 273, 10823– 10826. 35. Kishida, M., Koyama, S., Kishida, S., Matsubara, K., Nakashima, S., Higano, K., Takada, R., Takada, S., and Kikuchi, A. (1999) Oncogene 18, 979 –985. 36. Sakanaka, C., Weiss, J. B., and Williams, L. T. (1998) Proc. Natl. Acad. Sci. USA 95, 3020 –3023. 37. Kodama, S., Ikeda, S., Asahara, T., Kishida, M., and Kikuchi, A. (1999) J. Biol. Chem. 274, 27682–27688. 38. Polakis, P. (1997) Biochim. Biophys. Acta 1332, F127–F147. 39. Rubinfeld, B., Souza, B., Albert, I., Mu¨ller, O., Chamberlain, S. H., Masiarz, F. R., Munemitsu, S., and Polakis, P. (1993) Science 262, 1731–1734 40. Su, L. K., Vogelstein, B., and Kinzler, K. W. (1993) Science 262, 1734 –1737. 41. Rubinfeld, B., Souza, B., Albert, I., Munemitsu, S., and Polakis, P. (1995) J. Biol. Chem. 270, 5549 –5555. 42. Rubinfeld, B., Albert, I., Porfiri, E., Munemitsu, S., and Polakis, P. (1997) Cancer Res. 57, 4624 – 4630. 43. Rubinfeld, B., Albert, I., Porfiri, E., Fiol, C., Munemitsu, S., and Polakis, P. (1996) Science 272, 1023–1026. 44. Ikeda, S., Kishida, M., Matsuura, Y., Usui, H., and Kikuchi, A. (2000) Oncogene, in press. 45. Klingensmith, J., Nusse, R., and Perrimon, N. (1994) Genes Dev. 8, 118 –130. 46. Theisen, H., Purcell, J., Bennett, M., Kansagara, D., Syed, A., and Marsh, J. L. (1994) Development 120, 347–360. 47. Sussman, D. J., Klingensmith, J., Salinas, P., Adams, P. S., Nusse, R., and Perrimon, N. (1994) Dev. Biol. 166, 73– 86. 48. Klingensmith, J., Yang, Y., Axelrod, J. D., Beier, D. R., Perrimon, N., and Sussman, D. J. (1996) Mech. Dev. 58, 15–26.

49. Pizzuti, A., Amati, F., Calabrese, G., Mari, A., Colosimo, A., Silani, V., Giardino, L., Ratti, A., Penso, D., Calza, L., Palka, G., Scarlato, G., Novelli, G., and Dallapiccola, B. (1996) Hum. Mol. Genet. 5, 953–958. 50. Sokol, S. Y. (1996) Curr. Biol. 6, 1456 –1467. 51. Yanagawa, S., van Leeuwen, F., Wodarz, A., Klingensmith, J., and Nusse, R. (1995) Genes Dev. 9, 1087–1097. 52. Yanagawa, S., Lee, J., Haruna, T., Oda, H., Uemura, T., Takeichi, M., and Ishimoto, A. (1997) J. Biol. Chem. 272, 25243– 25251. 53. Boutros, M., Paricio, N., Strutt, D. I., and Mlodzik, M. (1998) Cell 94, 109 –118. 54. Li, L., Yuan, H., Xie, W., Mao, J., Caruso, A. M., McMahon, A., Sussman, D. J., and Wu, D. (1999) J. Biol. Chem. 274, 129 –134. 55. Smalley, M. J., Sara, E., Paterson, H., Naylor, S., Cook, D., Jayatilake, H., Fryer, L. G., Hutchinson, L., Fry, M. J., and Dale, T. C. (1999) EMBO J. 18, 2823–2835. 56. Fagotto, F., Jho, E., Zeng, L., Kurth, T., Joos, T., Kaufmann, C., and Costantini, F. (1999) J. Cell Biol. 145, 741–756. 57. Li, L., Yuan, H., Weaver, C. D., Mao, J., Farr, I. G. H., Sussman, D. J., Jonkers, J., Kimelman, D., and Wu, D. (1999) EMBO J. 18, 4233– 4240. 58. Cohen, P. (1989) Annu. Rev. Biochem. 58, 453–508. 59. Wera, S., and Hemmings, B. A. (1995) Biochem. J. 311, 17–29. 60. Hsu, W., Zeng, L., and Costantini, F. (1999) J. Biol. Chem. 274, 3439 –3445. 61. Seeling, J. M., Miller, J. R., Gil, R., Moon, R. T., White, R., and Virshup, D. M. (1999) Science 283, 2089 –2091. 62. Jonkers, J., Korswagen, H. C., Acton, D., Breuer, M., and Berns, A. (1997) EMBO J. 16, 441– 450. 63. Yost, C., Farr, G. H., III, Pierce, S. B., Ferkey, D. M., Chen, M. M., and Kimelman, D. (1998) Cell 93, 1031–1041. 64. Brannon, M., and Kimelman, D. (1996) Dev. Biol. 180, 344 –347. 65. Riese, J., Yu, X., Munnerlyn, A., Eresh, S., Hsu, S. C., Grosschedl, R., and Bienz, M. (1997) Cell 88, 777–787. 66. He, T., Sparks, A. B., Rago, C., Hermeking, H., Zawel, L., da Costa, L. T., Morin, P. J., Vogelstein, B., and Kinzler, K. W. (1998) Science 281, 1509 –1512. 67. Mann, B., Gelos, M., Siedow, A., Hanski, M. L., Gratchev, A., Ilyas, M., Bodmer, W. F., Moyer, M. P., Riecken, E. O., Buhr, H. J., and Hanski, C. (1999) Proc. Natl. Acad. Sci. USA 96, 1603–1608. 68. Tetsu, O., and McCormick, F. (1999) Nature 398, 422– 426. 69. Cavallo, R. A., Cox, R. T., Moline, M. M., Roose, J., Polevoy, G. A., Clevers, H., Peifer, M., and Bejsovec, A. (1998) Nature 395, 604 – 608. 70. Waltzer, L., and Bienz, M. (1998) Nature 395, 521–525. 71. Ishitani, T., Ninomiya-Tsuji, J., Nagai, S., Nishita, M., Meneghini, M., Barker, N., Waterman, M., Bowerman, B., Clevers, H., Shibuya, H., and Matsumoto, K. (1999) Nature 399, 798 – 802. 72. Bauer, A., Huber, O., and Kemler, R. (1998) Proc. Natl. Acad. Sci. USA 95, 14787–14792. 73. Morin, P. J., Sparks, A. B., Korinek, V., Barker, N., Clevers, H., Vogelstein, B., and Kinzler, K. W. (1997) Science 275, 1787– 1790. 74. Rubinfeld, B., Robbins, P., El-Gamil, M., Albert, I., Porfiri, E., and Polakis, P. (1997) Science 275, 1790 –1792.