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Ecole Polytechnique Fédérale de Lausanne-ISREC (Swiss Institute for Experimental Cancer Research), Chemin des Boveresses 155, 1066 Epalinges, Switzerland,1 Morphogenesis and Intracellular Signalling, UMR 144, Institut Curie-CNRS, 26 rue d'Ulm, 75248 Paris Cedex 05, France2
Received 12 June 2007/ Returned for modification 19 July 2007/ Accepted 23 August 2007
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ABSTRACT |
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INTRODUCTION |
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TopABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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One of the major players involved in the establishment of tissue architecture during development and in homeostasis of a variety of adult tissues is the canonical Wnt/ß-catenin signaling pathway (8). ß-Catenin is an essential cytoplasmic signal transducer of this canonical Wnt pathway (2). In the absence of pathway stimulation by Wnt ligands, ß-catenin is phosphorylated and targeted for degradation. The degradation complex responsible for ß-catenin destabilization contains the tumor suppressor gene products axin or conductin and adenomatous polyposis coli (APC) as well as glycogen synthase kinase 3ß and casein kinase I (CKI). Upon Wnt ligand binding to Frizzled and/or LRP transmembrane receptors, the cytoplasmic protein Disheveled is activated and blocks the action of the degradation complex. ß-Catenin is then able to enter the nucleus and associate with TCF/LEF transcription factors, thus inducing transcriptional regulation of Wnt target genes (2). Colorectal cancers, the second most common human malignant tumor type, are by and large initiated by mutations that activate the Wnt signaling pathway (2). These tumors are characterized by truncating mutations in APC and axin, as well as mutations in the degradation-inducing phosphorylation sites in ß-catenin, all leading to the formation of constitutive nuclear ß-catenin/TCF complexes (8).
Several studies of Wnt signaling demonstrate its effect on stem cells from various tissues. In mouse epidermis, conditional ablation of the ß-catenin gene blocked the differentiation of the bulge stem cells into follicular lineages (13). TCF3, a transcription factor of the Wnt signaling cascade, is preferentially expressed in the bulge stem cell compartment and has been suggested to maintain the stem cell pool (22). Activation of Wnt signaling in hematopoietic stem cells triggered increased self-renewal, whereas overexpression of axin, which can inactivate the pathway, led to a reduced reconstitution efficiency (31). In neural cells, inactivation of the Wnt pathway decreased the expansion of the progenitor compartment, whereas activation of the pathway increased that compartment (42). Several in vivo approaches based on inactivating and overactivating mutations of various pathway components were undertaken in order to elucidate the role of Wnt/ß-catenin signaling in intestinal epithelium (14, 16, 17, 27, 35, 39, 40). During embryonic development, TCF4 was shown to be required to maintain the proliferative compartment of the intestinal epithelium, as gene ablation leads to neonatal epithelium entirely composed of differentiated, nondividing cells (16). In the adult, alterations in Wnt signaling also indicated an important function of the pathway in intestinal proliferation and Paneth cell differentiation (14, 16, 17, 27, 35, 39, 40). However, all these studies reported rather mild and only transient phenotypes, making it sometimes hard to distinguish whether the observed phenotypes were direct results of pathway inactivation or part of the recovery mechanism. Moreover, the role of Wnt signaling in the control of intestinal stem cells was not directly addressed.
In this study, we determine the immediate consequences of ß-catenin loss in the intestinal epithelium of adult mice. Inactivation of ß-catenin leads to a rapid loss of intestinal epithelial cells, starting with the loss of crypts that concurs with blocked proliferation and increased enterocytic differentiation. Importantly, intestinal stem cells are induced to terminally differentiate in the absence of Wnt signaling, resulting in fatal loss of intestinal function.
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MATERIALS AND METHODS |
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ß-Galactosidase assay. Whole-mount samples from small intestines were isolated; washed with phosphate-buffered saline (PBS); fixed at 4°C with 2.5% glutaraldehyde in PBS, pH 7.4, for 30 min; rinsed in PBS, and incubated in X-Gal solution (1 g/liter X-Gal [5-bromo-4-chloro-3-indolyl-ß-D-galactoside] in PBS, pH 7.4, 5 mmol/liter of potassium ferrocyanide, 5 mmol/liter of potassium ferricyanide, 2 mmol/liter of MgCl2, and 0.2% Triton X-100) overnight in the dark at 37°C. Subsequently, tissue samples were washed with PBS and embedded in plastic (Technovit7100). Sections of 4 µm in thickness were counterstained with eosin, dehydrated through an ethanol series, and embedded in Entellan. Relative LacZ signal intensity was assessed as a percentage of maximal activity. Proliferation was assessed as the number of cells in metaphase/anaphase per position in the crypt. For both analyses, >100 crypts were counted.
Immunohistochemistry and in situ hybridizations. Intestinal tissue for immunohistochemistry and hematoxylin and eosin (H&E) staining was cleaned with PBS, fixed in 4% formaldehyde, and embedded in paraffin. Sections of 4 µm in thickness were incubated with the following primary antibodies: ß-catenin (BD Bioscience), BrdU (Sigma), p21 (BD Bioscience), active caspase 3 (Cell Signaling), CD44 (gift from Andreas Trumpp), lysozyme (DAKO), and FabpL (gift from Jeffrey Gordon). Envision+ (DakoCytomation) was used as a secondary reagent, and stainings were developed with diaminobenzidine. Hematoxylin was used for counterstaining. For in situ hybridization, sense and antisense riboprobes were synthesized from cDNA fragments of mSox4 (NM_009238.2, nucleotides 662 to 1984) and mDiap3 (NM_019670.1, nucleotides 727 to 3516) and hybridized as described previously (11).
Electron microscopy. Intestinal tissue was washed in PBS and fixed in 2.5% glutaraldehyde in 0.1 M sodium cacodylate buffer at 4°C overnight. Tissue was cut into 1-mm3 cubes for fixing. Postfixation was done in 1% osmium tetroxide in 0.1 M cacodylate buffer for 1 h (2 ml of 4% of aqueous OsO4, 4 ml of 0.2 M buffer, and 2 ml H2O were mixed and used immediately). After rinsing in 0.1 M buffer and dehydration, tissue was put in propylene oxide (1.2 epoxy propane) twice for 15 min. Propylene oxide/epoxy resin mixture (50/50) was then applied twice for 1 h at 20°C and then overnight at 4°C. Tissues were embedded in freshly prepared resin and left to polymerize at 60°C for 72 h. Pictures were taken with a Philips CM 10 transmission electron microscope.
Tritiated TdR labeling of LRCs. Three-week-old mice were injected twice daily with 25 µCi of [3H]thymidine ([3H]TdR) for four consecutive days. Three weeks later, animals were injected with tamoxifen for 2, 3, and 4 days, respectively, and sacrificed the following day. Intestines were washed with PBS and fixed in 4% formaldehyde. Sections of 4 µm in thickness were prepared for autoradiography (K5 emulsion; Ilford) and exposed for 15 days. The sections were counterstained with nuclear fast red. Label-retaining cells (LRCs) were counted in transverse sections of the intestine. The threshold for detecting LRCs was set at five or more grains per nucleus.
Transcriptional profiling. Intestinal tissue was frozen in OCT, and sections of 10 µm in thickness were applied on membrane slides for laser capture microdissection (LCM; Molecular Machines and Industries, part no. 50102), stained with eosin, dehydrated, and dissected using an mCut laser microdissection system (Nikon Eclipse TE200). RNA was isolated using the Pico Pure Isolation Kit (Arcturus). After RNA quality control by agarose gel electrophoresis and Agilent Bioanalyzer analysis, amplification was performed using the MessageAmp II aRNA Kit (Ambion) and labeling of the amplified RNA using the IVT Affymetrix kit. Biotin-labeled cRNA was hybridized on 430v 2.0 mouse Affymetrix arrays. Normalization and signal estimation were performed by RMA in the R package using the RACE vignette developed by the DNA Array Facility, Lausanne, Switzerland. Statistical analysis was performed using the Bayes test, and only deregulated genes with changes above 1.5-fold and a P value threshold below 5% were considered for further analysis. Gene ontology and pathway analysis were performed using GenMAPP software.
Nucleotide sequence accession number. The complete microarray data set is deposited in the GEO database (http://www.ncbi.nlm.nih.gov/geo) under accession number GSE8818.
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RESULTS |
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TopABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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The observed changes in crypt morphology coincide with drastic changes in crypt cell proliferation. While 1 day after deletion mutant crypts display unaltered proliferation, a complete block of proliferation is observed after the second day of ß-catenin deletion, as assessed by short-term BrdU incorporation (Fig. 2A and B) (see also the data in the supplemental material). Furthermore, known components of cell cycle control are found to be altered in ß-catenin mutants. Expression of the Wnt target gene c-myc is decreased whereas the cell cycle inhibitor p21 is found to be increased in intervillus regions (data not shown). Increased apoptosis is not responsible for loss of crypts in ß-catenin mutants as analyzed by immunohistochemistry for active caspase 3, a hallmark of apoptotic cell death (Fig. 2C to G). However, mutants show a minor increase of apoptosis at the tips of the villi 4 days after ablation (Fig. 2F and G). This late apoptotic event closely reflects the normal life span of these differentiated cells, which migrate up the villus, undergo apoptosis at the villus tip, and are shed into the lumen of the gut within 4 to 5 days in wild-type animals (cf. Fig. 2C).
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Differentiation of stem cells upon ß-catenin ablation in intestinal epithelium. Loss of progenitor fates suggested a possible defect of intestinal stem cell function upon deletion of Wnt/ß-catenin signaling. Most tissue-specific stem cells divide infrequently, and this characteristic can be employed to identify these cells in situ. After saturated labeling of all dividing cells by tritiated [3H]TdR, cells which divide infrequently will retain the label over long periods of time (so-called LRCs), while cells which cycle more often will lose the label. We injected 3-week-old mice twice daily with tritiated TdR for four consecutive days. Three weeks later we induced the ablation of ß-catenin for 2 to 4 days. As expected, wild-type mouse intestines contain label-retaining stem cells in about every fifth crypt at the previously reported cell position 2, immediately above the Paneth cells (Fig. 5A) (29). In the ß-catenin deletion mutants, total LRC numbers are similar to those in the wild type; however, their distribution becomes aberrant. Observation of over 500 crypt-villus units revealed that LRCs disperse throughout the crypt area and are occasionally found in the villi within 2 days after deletion of ß-catenin. Subsequently, LRCs become evenly distributed throughout the crypt-villus axis within 4 days (Fig. 5B and C). In order to evaluate whether cell migration of the intestinal epithelium is affected by inactivation of ß-catenin, we pulse-labeled control and mutant mice with BrdU shortly before tamoxifen injection for 2 to 3 days. We thereby labeled a cohort of progenitor cell progeny which would differentiate and migrate towards the villus tip within the next few days. BrdU immunohistochemistry revealed that the overall cell migration rate remains unchanged in mutant mice (Fig. 5F to I). We therefore assessed next whether the dispersed LRCs might be migrating, differentiated cells. We performed dual stainings in order to detect LRCs by autoradiography and differentiated enterocytes by immunohistochemistry for the enterocyte marker FabpL. In 4-day mutants, we detected all LRCs colabeled with FabpL (Fig. 5E, n = 30), in contrast to control animals, where LRCs are restricted to crypts and do not express FabpL (Fig. 5D). This strongly suggests that ß-catenin deletion causes forced differentiation of stem cells into the enterocytic lineage. This process of differentiation supports the loss of crypts and contributes to the block of proliferation.
Transcriptional changes underlying the loss of the intestinal progenitor compartment. In order to understand the molecular mechanisms of crypt loss as a result of ß-catenin deletion in the intestinal epithelium, we performed transcriptional profiling of intestinal crypts. We isolated crypt cells by LCM from wild-type and mutant mouse epithelium 2 days after induction of ß-catenin deletion. After RNA isolation and assessment of RNA quality, RNA was amplified, and Affymetrix analysis of differential gene expression was performed (a complete list of differentially expressed genes is found in the supplemental material as Table S1).
Overall, the expression profile matches the phenotype observed by histological analysis. Loss of Wnt signaling activity is reflected by downregulation of ß-catenin itself and of the Wnt target genes conductin and c-myc. It is further reflected by the upregulation of p21, which is transcriptionally repressed by c-myc (37). Indeed, in vitro transfection of colorectal cells with a dominant-negative form of TCF resulted in decreased c-myc, increased p21, and subsequent G1 cell cycle arrest and increased differentiation (37), which is consistent with our results. In line with this, other key genes of cell cycle control (i.e., ccnd2 and ccna2) are downregulated in our transcriptional profiling analysis, as are other known target genes of Wnt signaling such as ephB2 and ephB3. Disruption of these ephrin receptors has been linked to aberrant cell intermingling and mislocalization of Paneth cells (1) in accordance with our immunohistochemical analysis of Paneth cell localization (cf. Fig. 4I to K). Upregulated genes include the enterocytic differentiation marker carbonic anhydrase 4, concurring with our analysis of enhanced terminal differentiation. Together, these results support our conclusion that stem cells and transient-amplifying cells are arrested in the cell cycle and are forced into enterocytic differentiation as a result of ß-catenin deletion (cf. Fig. 5E).
Recently, CD133 has been used as a marker to enrich for cancer stem cells responsible for the formation of human colorectal tumors (24, 32). However, markers which can unambiguously identify normal intestinal stem cells or intestinal cancer stem cells have not been identified. We reasoned that our list of genes functionally implicated in intestinal stem cell maintenance might contain such markers, and we therefore compared our list of genes with genes found to be deregulated in intestinal tumors as well as in other stem cell populations. In detail, we compared our gene list to transcriptional profiles from murine intestinal tumors of PTEN-deficient mice and APC (Min/+) and protein kinase C
-deficient APC (Min/+) mice (12, 18, 25, 26, 30). We identified common genes that were upregulated in those arrays and downregulated in our ß-catenin deletion arrays. Next, we compared the obtained gene list with transcriptomes of over 90 human colon adenomas and adenocarcinomas and narrowed the list down to genes that are also upregulated in at least 20% of those human arrays (4, 10, 23, 43). In order to focus on stemness-related genes, we next determined the overlap with genes that have been shown to be characteristic for various stem cell populations (15, 33). Numerous cell cycle-associated genes are deregulated both in our studies and in those used for comparison, due to either the slow cycling nature of stem cells or the elevated proliferation in tumors. We therefore excluded known cell cycle-related genes from our gene list, based on transcriptional profiling by Whitfield et al. (38) and Cho et al. (7) and on the Celera and Stanford online databases. Common deregulated genes matching all of the criteria described above are summarized in Table 1, which therefore lists candidates for intestinal stem and cancer stem cell markers. For six of these genes, stem cell-specific expression has been described independently based on analyses of human adenomas and in situ hybridization of normal intestines (36). We confirmed stem cell expression by in situ hybridization for several others such as Sox4, which is also expressed by Paneth cells, and Diap3 (Fig. 6A and C; also data not shown). Importantly, we find expression of both genes to be strongly repressed in mutant mouse crypts in agreement with the loss of the stem cell phenotype after loss of ß-catenin signaling (Fig. 6B and D).
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DISCUSSION |
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Previous approaches to studying the role of Wnt signaling in the intestinal epithelium included knocking down or overexpressing different components of the signaling cascade. Overall, these studies induced only minor, transient phenotypes, making it difficult to distinguish consequences of altered Wnt signaling from regeneration mechanisms. Moreover, previous studies did not address the fate of stem cells following Wnt/ß-catenin ablation. Two groups reported overexpression of the secreted Wnt inhibitor Dickkopf1 (Dkk1). In one study, Kuhnert et al. infected mice with adenoviruses expressing Dkk1 (17). Infrequent loss of crypts accompanied by diminished proliferation was observed after 1 week of viral administration. However, decreased Dkk1 expression at later points was followed by epithelial regeneration. In a second study, Pinto et al. generated transgenic mice expressing Dkk1 under the control of the villin promoter (27). Those mice displayed a mild phenotype with reduced epithelial proliferation, partial crypt loss, and increased enterocytic differentiation. In contrast to our mutants, those mice survived, probably due to an incomplete block of Wnt signaling and scattered areas of unaffected crypts due to mosaic expression of the transgene (27). An approach of conditional ablation of ß-catenin was undertaken by Ireland et al., using an Ah promoter-driven cre recombinase (14). Again, this induced only a transient (24-h) deletion of ß-catenin, and complete regeneration was observed within a few days. Partial reduction in the number of crypt cells was followed by recovery from increased proliferation of wild-type cells, which makes comparisons with our results difficult. Several other studies used a complementary approach, i.e., overactivation of the Wnt/ß-catenin pathway (35, 39, 40). As expected, these experiments induced phenotypes opposite from those in our gene ablation model: ß-catenin-overexpressing crypts showed increased proliferation, causing crypt expansion and decreased enterocytic differentiation (35). Based on our novel results, we suggest that this phenotype might reflect an expansion of the intestinal stem cell pool in these mutant animals.
c-Myc is a known target of the Wnt pathway in vitro (8, 37) and was recently confirmed as a critical downstream signal controlling intestinal proliferation in vivo (34). Several groups conditionally deleted c-myc in the intestinal epithelium (3, 21, 34). Surprisingly, in all those studies intestinal function was only transiently perturbed since c-myc-negative crypts were replaced by escaper cells through a crypt fission process, which is reminiscent of postirradiation repair (6). In agreement with our data for an essential requirement of Wnt/ß-catenin signaling in progenitor cell proliferation, this repopulation mechanism involved increased Wnt/ß-catenin activity. However, the preservation of repair mechanisms in these mutants indicates that c-myc is most likely not one of the essential Wnt targets to maintain intestinal stem cell function.
With the aim of identifying novel markers of intestinal stem and cancer stem cells, we performed transcriptional profiling followed by extensive comparisons to other stem cell populations and intestinal tumor studies. This allowed identification of novel marker candidates. In a recent complementary approach, genes upregulated in human intestinal tumors were compared to genes repressed upon a block of Wnt signaling in human colorectal cancer cell lines (36). We note significant overlap between our murine gene list and this human gene list, which together appear to define the set of common intestinal Wnt targets in human and mouse. Importantly, numerous genes that are shown by this study to be specifically expressed in intestinal stem cells, such as apex1, ascl2, gemin4, rhobtb3, sox4, and wdr12, are also deregulated in our arrays. Stem cell-specific expression is also demonstrated in both studies for several genes encoding members of the zinc finger family. Furthermore, we identify additional murine stem cell markers such as msi2, which belongs to the same family as the suggested marker msi1 (28), and in particular diaphanous3 (diap3). Interestingly, the diaphanous gene family has been identified to affect germ cell formation in Drosophila melanogaster and humans and has been implicated in regulating microtubule attachment to kinetochores (41).
Activating mutations of the Wnt/ß-catenin pathway are well-known genetic alterations in premalignant lesions in the intestine (8). This introduced the concept that aberrant Wnt/ß-catenin signaling initiates the transformation process. Based on these results, various efforts have been initiated to identify small-molecule drugs to be used for colorectal cancer therapy which function as inhibitors of Wnt signal transduction. While clinical trials with these inhibitors have not been published, our results challenge this approach and predict serious side effects of such drugs.
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ACKNOWLEDGMENTS |
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T.F. and J.H. were supported in part by the Swiss League against Cancer (OCS 01838-02-2006), the SNF (3100AO-104209), and the Swiss NCCR in Molecular Oncology.
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FOOTNOTES |
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Published ahead of print on 4 September 2007.
Supplemental material for this article may be found at http://mcb.asm.org/.
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REFERENCES |
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2. Behrens, J., and B. Lustig. 2004. The Wnt connection to tumorigenesis. Int. J. Dev. Biol. 48: 477-487.[CrossRef][Medline]
3. Bettess, M. D., N. Dubois, M. J. Murphy, C. Dubey, C. Roger, S. Robine, and A. Trumpp. 2005. c-Myc is required for the formation of intestinal crypts but dispensable for homeostasis of the adult intestinal epithelium. Mol. Cell. Biol. 25: 7868-7878.
4. Bianchini, M., E. Levy, C. Zucchini, V. Pinski, C. Macagno, P. De Sanctis, L. Valvassori, P. Carinci, and J. Mordoh. 2006. Comparative study of gene expression by cDNA microarray in human colorectal cancer tissues and normal mucosa. Int. J. Oncol. 29: 83-94.[Medline]
5. Bjerknes, M., and H. Cheng. 1999. Clonal analysis of mouse intestinal epithelial progenitors. Gastroenterology 116: 7-14.[CrossRef][Medline]
6. Booth, D., J. D. Haley, A. M. Bruskin, and C. S. Potten. 2000. Transforming growth factor-B3 protects murine small intestinal crypt stem cells and animal survival after irradiation, possibly by reducing stem-cell cycling. Int. J. Cancer 86: 53-59.[CrossRef][Medline]
7. Cho, R. J., M. Huang, M. J. Campbell, H. Dong, L. Steinmetz, L. Sapinoso, G. Hampton, S. J. Elledge, R. W. Davis, and D. J. Lockhart. 2001. Transcriptional regulation and function during the human cell cycle. Nat. Genet. 27: 48-54.[Medline]
8. Clevers, H. 2006. Wnt/beta-catenin signaling in development and disease. Cell 127: 469-480.[CrossRef][Medline]
9. el Marjou, F., K. P. Janssen, B. H. Chang, M. Li, V. Hindie, L. Chan, D. Louvard, P. Chambon, D. Metzger, and S. Robine. 2004. Tissue-specific and inducible Cre-mediated recombination in the gut epithelium. Genesis 39: 186-193.[CrossRef][Medline]
10. Gardina, P. J., T. A. Clark, B. Shimada, M. K. Staples, Q. Yang, J. Veitch, A. Schweitzer, T. Awad, C. Sugnet, S. Dee, C. Davies, A. Williams, and Y. Turpaz. 2006. Alternative splicing and differential gene expression in colon cancer detected by a whole genome exon array. BMC Genomics 7: 325.[CrossRef][Medline]
11. Gregorieff, A., D. Pinto, H. Begthel, O. Destree, M. Kielman, and H. Clevers. 2005. Expression pattern of Wnt signaling components in the adult intestine. Gastroenterology 129: 626-638.[CrossRef][Medline]
12. He, X. C., T. Yin, J. C. Grindley, Q. Tian, T. Sato, W. A. Tao, R. Dirisina, K. S. Porter-Westpfahl, M. Hembree, T. Johnson, L. M. Wiedemann, T. A. Barrett, L. Hood, H. Wu, and L. Li. 2007. PTEN-deficient intestinal stem cells initiate intestinal polyposis. Nat. Genet. 39: 189-198.[CrossRef][Medline]
13. Huelsken, J., R. Vogel, B. Erdmann, G. Cotsarelis, and W. Birchmeier. 2001. ß-Catenin controls hair follicle morphogenesis and stem cell differentiation in the skin. Cell 105: 533-545.[CrossRef][Medline]
14. Ireland, H., R. Kemp, C. Houghton, L. Howard, A. R. Clarke, O. J. Sansom, and D. J. Winton. 2004. Inducible Cre-mediated control of gene expression in the murine gastrointestinal tract: effect of loss of beta-catenin. Gastroenterology 126: 1236-1246.[CrossRef][Medline]
15. Kim, C. G., J. J. Lee, D. Y. Jung, J. Jeon, H. S. Heo, H. C. Kang, J. H. Shin, Y. S. Cho, K. J. Cha, C. G. Kim, B. R. Do, K. S. Kim, and H. S. Kim. 2006. Profiling of differentially expressed genes in human stem cells by cDNA microarray. Mol. Cells 21: 343-355.[Medline]
16. Korinek, V., N. Barker, P. Moerer, E. van Donselaar, G. Huls, P. J. Peters, and H. Clevers. 1998. Depletion of epithelial stem-cell compartments in the small intestine of mice lacking Tcf-4. Nat. Genet. 19: 379-383.[CrossRef][Medline]
17. Kuhnert, F., C. R. Davis, H. T. Wang, P. Chu, M. Lee, J. Yuan, R. Nusse, and C. J. Kuo. 2004. Essential requirement for Wnt signaling in proliferation of adult small intestine and colon revealed by adenoviral expression of Dickkopf-1. Proc. Natl. Acad. Sci. USA 101: 266-271.
18. Leclerc, D., L. Deng, J. Trasler, and R. Rozen. 2004. ApcMin/+ mouse model of colon cancer: gene expression profiling in tumors. J. Cell. Biochem. 93: 1242-1254.[CrossRef][Medline]
19. Lustig, B., B. Jerchow, M. Sachs, S. Weiler, T. Pietsch, U. Karsten, M. van de Wetering, H. Clevers, P. M. Schlag, W. Birchmeier, and J. Behrens. 2002. Negative feedback loop of Wnt signaling through upregulation of conductin/axin2 in colorectal and liver tumors. Mol. Cell. Biol. 22: 1184-1193.
20. Marshman, E., C. Booth, and C. S. Potten. 2002. The intestinal epithelial stem cell. Bioessays 24: 91-98.[CrossRef][Medline]
21. Muncan, V., O. J. Sansom, L. Tertoolen, T. J. Phesse, H. Begthel, E. Sancho, A. M. Cole, A. Gregorieff, I. Moreno de Alboran, H. Clevers, and A. R. Clarke. 2006. Rapid loss of intestinal crypts upon conditional deletion of the Wnt/Tcf-4 target gene c-Myc. Mol. Cell. Biol. 26: 8418-8426.
22. Nguyen, H., M. Rendl, and E. Fuchs. 2006. Tcf3 governs stem cell features and represses cell fate determination in skin. Cell 127: 171-183.[CrossRef][Medline]
23. Notterman, D. A., U. Alon, A. J. Sierk, and A. J. Levine. 2001. Transcriptional gene expression profiles of colorectal adenoma, adenocarcinoma, and normal tissue examined by oligonucleotide arrays. Cancer Res. 61: 3124-3130.
24. O'Brien, C. A., A. Pollett, S. Gallinger, and J. E. Dick. 2007. A human colon cancer cell capable of initiating tumour growth in immunodeficient mice. Nature 445: 106-110.[CrossRef][Medline]
25. Oster, H., and M. Leitges. 2006. Protein kinase C alpha but not PKCzeta suppresses intestinal tumor formation in ApcMin/+ mice. Cancer Res. 66: 6955-6963.
26. Paoni, N. F., M. W. Feldman, L. S. Gutierrez, V. A. Ploplis, and F. J. Castellino. 2003. Transcriptional profiling of the transition from normal intestinal epithelia to adenomas and carcinomas in the APCMin/+ mouse. Physiol. Genomics 15: 228-235.
27. Pinto, D., A. Gregorief, H. Begthel, and H. Clevers. 2003. Canonical Wnt signals are essential for homeostasis of the intestinal epithelium. Genes Dev. 17: 1709-1713.
28. Potten, C. S., C. Booth, G. L. Tudor, D. Booth, G. Brady, P. Hurley, G. Ashton, R. Clarke, S. Sakakibara, and H. Okano. 2003. Identification of a putative intestinal stem cell and early lineage marker; musashi-1. Differentiation 71: 28-41.[CrossRef][Medline]
29. Potten, C. S., G. Owen, and D. Booth. 2002. Intestinal stem cells protect their genome by selective segregation of template DNA strands. J. Cell Sci. 115: 2381-2388.
30. Reichling, T., K. H. Goss, D. J. Carson, R. W. Holdcraft, C. Ley-Ebert, D. Witte, B. J. Aronow, and J. Groden. 2005. Transcriptional profiles of intestinal tumors in Apc(Min) mice are unique from those of embryonic intestine and identify novel gene targets dysregulated in human colorectal tumors. Cancer Res. 65: 166-176.
31. Reya, T., A. W. Duncan, L. Ailles, J. Domen, D. C. Scherer, K. Willert, L. Hintz, R. Nusse, and I. L. Weissman. 2003. A role for wnt signalling in self-renewal of hematopoietic stem cells. Nature 423: 409-414.[CrossRef][Medline]
32. Ricci-Vitiani, L., D. G. Lombardi, E. Pilozzi, M. Biffoni, M. Todaro, C. Peschle, and R. De Maria. 2007. Identification and expansion of human colon-cancer-initiating cells. Nature 445: 111-115.[CrossRef][Medline]
33. Rochon, C., V. Frouin, S. Bortoli, K. Giraud-Triboult, V. Duverger, P. Vaigot, C. Petat, P. Fouchet, B. Lassalle, O. Alibert, X. Gidrol, and G. Pietu. 2006. Comparison of gene expression pattern in SP cell populations from four tissues to define common "stemness functions." Exp. Cell Res. 312: 2074-2082.[CrossRef][Medline]
34. Sansom, O. J., V. S. Meniel, V. Muncan, T. J. Phesse, J. A. Wilkins, K. R. Reed, J. K. Vass, D. Athineos, H. Clevers, and A. R. Clarke. 2007. Myc deletion rescues Apc deficiency in the small intestine. Nature 446: 676-679.[CrossRef][Medline]
35. Sansom, O. J., K. R. Reed, A. J. Hayes, H. Ireland, H. Brinkmann, I. P. Newton, E. Batlle, P. Simon-Assmann, H. Clevers, I. S. Nathke, A. R. Clarke, and D. J. Winton. 2004. Loss of Apc in vivo immediately perturbs Wnt signaling, differentiation, and migration. Genes Dev. 18: 1385-1390.
36. Van der Flier, L. G., J. Sabates-Bellver, I. Oving, A. Haegebarth, M. De Palo, M. Anti, M. E. van Gijn, S. Suijkerbuijk, M. van de Wetering, G. Marra, and H. Clevers. 2007. The intestinal Wnt/TCF signature. Gastroenterology 132: 628-632.[CrossRef][Medline]
37. van de Wetering, M., E. Sancho, C. Verweij, W. de Lau, I. Oving, A. Hurlstone, K. van der Horn, E. Batlle, D. Coudreuse, A. P. Haramis, M. Tjon-Pon-Fong, P. Moerer, M. van den Born, G. Soete, S. Pals, M. Eilers, R. Medema, and H. Clevers. 2002. The beta-catenin/TCF-4 complex imposes a crypt progenitor phenotype on colorectal cancer cells. Cell 111: 241-250.[CrossRef][Medline]
38. Whitfield, M. L., G. Sherlock, A. J. Saldanha, J. I. Murray, C. A. Ball, K. E. Alexander, J. C. Matese, C. M. Perou, M. M. Hurt, P. O. Brown, and D. Botstein. 2002. Identification of genes periodically expressed in the human cell cycle and their expression in tumors. Mol. Biol. Cell 13: 1977-2000.
39. Wong, M. H., J. Huelsken, W. Birchmeier, and J. I. Gordon. 2002. Selection of multipotent stem cells during morphogenesis of small intestinal crypts of Lieberkuhn is perturbed by stimulation of Lef-1/ß-catenin signaling. J. Biol. Chem. 277: 15843-15850.
40. Wong, M. H., B. Rubinfeld, and J. I. Gordon. 1998. Effects of forced expression of an NH2-terminal truncated beta-catenin on mouse intestinal epithelial homeostasis. J. Cell Biol. 141: 765-777.
41. Yasuda, S., F. Oceguera-Yanez, T. Kato, M. Okamoto, S. Yonemura, Y. Terada, T. Ishizaki, and S. Narumiya. 2004. Cdc42 and mDia3 regulate microtubule attachment to kinetochores. Nature 428: 767-771.[CrossRef][Medline]
42. Zechner, D., Y. Fujita, J. Huelsken, T. Muller, I. Walther, M. M. Taketo, E. B. Crenshaw III, W. Birchmeier, and C. Birchmeier. 2003. ß-Catenin signals regulate cell growth and the balance between progenitor cell expansion and differentiation in the nervous system. Dev. Biol. 258: 406-418.[CrossRef][Medline]
43. Zhang, J., A. Moseley, A. G. Jegga, A. Gupta, D. P. Witte, M. Sartor, M. Medvedovic, S. S. Williams, C. Ley-Ebert, L. M. Coolen, G. Egnaczyk, M. B. Genter, M. Lehman, J. Lingrel, J. Maggio, L. Parysek, R. Walsh, M. Xu, and B. J. Aronow. 2004. Neural system-enriched gene expression: relationship to biological pathways and neurological diseases. Physiol. Genomics 18: 167-183.
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