Jakob Mejlvang,1,
Shaukat Mahmood,2
Irina Gromova,3
Pavel Gromov,3
Eugene Lukanidin,2
Marina Kriajevska,1
J. Kilian Mellon,1 and
Eugene Tulchinsky1*
Department of Cancer Studies and Molecular Medicine, University of Leicester, Leicester LE1 9HN, United Kingdom,1 Department of Molecular Cancer Biology, Danish Cancer Society, Strandboulevarden 49, Copenhagen 2100, Denmark,2 Department of Proteomics in Cancer, Danish Cancer Society, Strandboulevarden 49, Copenhagen 2100, Denmark3
Received 1 March 2005/ Returned for modification 8 April 2005/ Accepted 21 July 2005
| ABSTRACT |
|---|
|
|
|---|
| INTRODUCTION |
|---|
|
|
|---|
-catenin, which in turn binds
-catenin, providing a link with the actin cytoskeleton and hence strengthening adhesion (9, 16). Disruption of E-cadherin-mediated intercellular adhesion is a hallmark of epithelial-mesenchymal transition (EMT), a phenomenon which occurs at certain stages of normal development and in the malignant progression of carcinoma (59, 60). Different molecular mechanisms including gene mutations (4, 26, 66), hypermethylation of the promoter (17), and transcriptional silencing by transcriptional repressors (Snail, Slug, ZEB-2/SIP1, ZEB-1, and E12/E47) (2, 8, 6, 14, 20, 46) contribute to the inactivation of E-cadherin linked with tumor progression. Reexpression of E-cadherin may induce morphological reversion and suppress cell growth and invasion suggesting an important function for E-cadherin in EMT (24, 56, 58, 67). The mechanism of tumor suppressor function of E-cadherin is not completely understood and may be linked with its role in signal transduction. Indeed, E-cadherin has been implicated not only in epithelial adhesion but also in the regulation of cell signaling. Being an important player in the Wnt signal transduction pathway, ß-catenin links E-cadherin with cellular signaling networks (9, 16, 29, 47). In different systems, sequestration of ß-catenin by the cytoplasmic domain of E-cadherin prevents its nuclear translocation and inhibits ß-catenin/T-cell factor (TCF)-mediated transcriptional activity (42, 51). In a model of Fos protein-induced EMT, loss of E-cadherin activated ß-catenin signaling in murine nontumorigenic Ep-1 cells (19). Inhibition of ß-catenin signaling by E-cadherin may result in suppression of cell growth, providing a molecular basis for adhesion-independent tumor suppression function of E-cadherin (24, 57). A direct link between the functional status of E-cadherin and ß-catenin signaling has been demonstrated in colon carcinoma cells SW480 harboring a mutant APC gene. In these cells, inhibition of adherens junctions by an anti-E-cadherin blocking antibody resulted in activation of ß-catenin/TCF-dependent transcription with subsequent activation of the transcriptional repressor, Slug, and repression of E-cadherin gene transcription (15). However, in other in vitro models of EMT, loss of E-cadherin expression did not result in increased ß-catenin/TCF transcriptional activity (14; J. Mejlvang et al., unpublished data). Moreover, ß-catenin/TCF transcriptional activity does not correlate with E-cadherin status in breast, gastric, and pancreatic carcinoma cell lines (7, 61). It has been suggested that E-cadherin influences cell signaling through receptor tyrosine kinases (RTKs). E-cadherin and epidermal growth factor receptor (EGFR) colocalize to basolateral areas of epithelial cells and have been reported to form multicomponent complexes (28, 44). Formation of adhesive complexes leads to transient ligand-independent activation of EGFR and subsequent activation of mitogen-activated protein kinase (MAPK) (43), phosphatidylinositol 3-kinase (30, 43) signaling cascades, and Rac1 (5, 30). E-cadherin engagement may influence the activity of small GTPases via Src-dependent phosphorylation of RhoA-specific GTPase-activating protein p190RhoGAP (39). In dense epithelial cultures, E-cadherin also activates another RTK, EphA2, and inhibits cell proliferation (68). Recently, E-cadherin-mediated adhesion has been shown to repress ligand-induced activation of several RTKs including EGFR/Neu, insulin-like growth factor 1 receptor and c-Met in Madin-Darby canine kidney (MDCK) (48) but not in SW480 cell lines (15). However, although it is documented that E-cadherin affects cell signaling through RTKs, cytoskeletal reorganization, and ß-catenin/TCF, there is a substantial lack of experimental work investigating the consequence of inhibition of E-cadherin-mediated adhesion for gene regulation. In this report, we demonstrate that prolonged functional inactivation of E-cadherin by a dominant-negative E-cadherin mutant, Ec1WVM, is sufficient to induce full EMT in A431 cells. Short-term inactivation of E-cadherin has a lesser effect on the expression of target genes but is sufficient to activate the transcription factor AP-1. Activation of AP-1 by Ec1WVM appeared to be essential for some of its transcriptional effects. In addition, Ec1WVM regulates tumor cell motility in an AP-1-dependent manner.
| MATERIALS AND METHODS |
|---|
|
|
|---|
Plasmids. To generate pBI-Ec1WVM, the Ec1WVM sequence was excised from pCMVEc1WVM and subcloned into multiple cloning site I of a bidirectional tetracycline (TET)-responsive vector pBI (BD Clontech). To construct a vector with simultaneous expression of Ec1WVM and a dominant-negative AP-1 mutant, a fragment coding for the TAM67-GFP fusion protein was excised from pGFP-TAMpuro (27) (provided by R. Hennigan, University of Cincinnati, Cincinnati, Ohio) and inserted into multiple cloning site II of pBI-Ec1WVM. To generate pUHD-c-Fos, c-Fos cDNA was cut out from pCMV-c-Fos and subcloned into pUHD-10-3.
Gene reporter assays. To determine ß-catenin/TCF/lymphoid enhancer factor (LEF) transcriptional activity, 31D6 cells were transfected with 2 µg of pTOPFLASH or pFOPFLASH reporter constructs. The efficiency of each transfection was monitored using 400 ng of cotransfected ß-galactosidase expression vector, pCMVß-gal (Invitrogen). Cells were maintained with or without DOX for 48 h and lysed, and the luciferase activity was measured with a Lumat LB9501 tube luminometer (Berthold). The lysates obtained were also tested for ß-galactosidase activity by using o-nitrophenyl-ß-D-galactopyranoside (Sigma) as a chromogenic substrate. Results were expressed as a ratio of pTOPFLASH and pFOPFLASH reporter activities normalized to the activity of ß-galactosidase in each experiment. To examine AP-1 activity, cells were transfected with an AP-1-dependent reporter pTREx5Luc containing five copies of an AP-1-binding element upstream of the minimal c-fos promoter (13). To demonstrate the specificity of AP-1 activation, we used a pRSVLuc reporter (13) that is largely AP-1-independent. Transfected cells were incubated for 2 days with or without DOX, and luciferase activity was measured and normalized to the ß-galactosidase activity.
One-dimensional SDS electrophoresis and Western blotting. Proteins (10 or 20 µg) were denatured, separated on precast 4 to 20% gradient sodium dodecyl sulfate (SDS)-polyacrylamide gels (Invitrogen), and then transferred to Immobilon-P membranes (Millipore) by the standard procedure. Following protein transfer, blots were incubated in blocking solution with primary antibody at a dilution of 1:1,000 (for anti-myc tag antibody, clone 9E10; Santa-Cruz Biotech), 1:2,000 (for anti-E-cadherin antibody; BD Biosciences), 1:400 (for anti-GFP antibody; BD Biosciences), or 1:500 (for anti-c-Fos, anti-Fra-1, and anti-c-Jun antibodies; Santa-Cruz Biotech). Immunoreactive proteins were detected using an enhanced chemiluminescence system (Amersham).
Metabolic labeling. Cells were grown to approximately 70% confluence in microtiter 24-well culture dishes and labeled overnight in Dulbecco's modified Eagle's medium lacking methionine and containing 1% dialyzed fetal calf serum and 1 mCi/ml [35S]methionine. Following labeling, cells were gently washed twice with phosphate-buffered saline solution and harvested by solubilization in lysis buffer for two-dimensional polyacrylamide gel electrophoresis (2D PAGE).
2D PAGE and image analysis. After cells were washed, excess phosphate-buffered saline solution was removed from the wells. A total of 50 µl of lysis buffer (40) was overlaid on cell monolayers, and the cells were lysed in solution by gentle pipetting. Samples were kept at 20°C until use. Whole protein lysates were subjected to isoelectrofocusing 2D PAGE as previously described (11). From 20 to 35 µl of sample was applied to the first dimension. Proteins were visualized using autoradiography and/or phosphorimaging followed by a silver staining procedure compatible with mass spectrometry analysis (25). Image analysis was performed using PDQUEST 7.1 software (Bio-Rad). Detection of low-abundant protein spots on silver-stained gels was highly enhanced by the superimposition of the dry silver gel with the corresponding autoradiograph.
Protein identification by mass spectrometry. Protein spots of interest were excised from the dry silver-stained gels, followed by rehydration in water for 30 min at room temperature. Proteins were "in-gel" digested with bovine trypsin for 12 h as previously described (55). The reaction was stopped by adding trifluoroacetic acid up to 0.4%, followed by shaking for 20 min at room temperature to increase peptide recovery. Peptides were concentrated on microcolumns containing C18-based 3M Empore plugs (49) and eluted with 50% acetonitrile-0.2% trifluoroacetic acid directly on the target and cocrystallized with cyano matrix (2 mg/ml cyano-4-hydroxycinnamic acid in 0.5% trifluoroacetic acid-acetonitrile, 1:2 [vol/vol]). The extraction procedure strongly increased the amount of peptides, thus allowing direct sequence analysis of low intensity peptides. Mass spectrometry was performed using a Reflex IV matrix-assisted laser desorption ionization-time of flight (MALDI-TOF) mass spectrometer equipped with a Scout 384 ion source. All spectra were obtained in positive reflector mode with delayed extraction, using an accelerating voltage of 28 kV. Each spectrum represented an average of 100 to 200 laser shots, depending on the signal-to-noise ratio. The resulting mass spectra were internally calibrated by using the autodigested tryptic mass values (805.417, 906.505, 1153.574, 1433.721, 2163.057, and 2273.160) visible in all spectra. Calibrated spectra were processed by the Xmass 5.1.1 and BioTools 2.1 software packages (Bruker Daltonik, GmbH). All spectra were analyzed manually as previously described (10).
Microarray hybridization. Hybridization of Atlas human cDNA expression arrays (Clontech) was performed basically as recommended by the manufacturer. Briefly, filters were prehybridized for 12 to 16 h at 68°C in 10 ml of ExpressHyb solution plus 100 µg/ml denatured sheared salmon sperm DNA. Radiolabeled probes were purified, heat denatured, and then added to 5- to 10-ml aliquots of hybridization buffer containing salmon sperm DNA. The final probe concentration was 5 x 106 to 10 x 106 cpm/hybridization. After extensive washing (three times with 2x SSC [1x SSC is 0.15 M NaCl plus 0.015 M sodium citrate]-1% SDS and two to three times with 0.1x SSC-0.5% SDS, each for 30 min at 68°C), the membranes were subjected to phosphorimaging analysis, and differential signals were identified by AtlasImage software.
Northern blotting. For Northern blot analysis, total RNA was isolated by the guanidine isothiocyanate method and separated in 1.2% agarose gels. RNA blotting and hybridization were performed as previously described (50). Radioactive DNA probes were synthesized using a random-primed labeling kit (Amersham). For radioactive labeling, 200- to 300-bp cDNA fragments corresponding to coding parts of identified genes were generated by reverse transcriptase PCR.
RNA interference. Purified and annealed synthetic oligonucleotides were purchased from Ambion (Austin, TX). The target sequence for Fra-1 was validated previously (63). The target sequence for c-Jun was GAUCCUGAAACAGAGCAUG. A total of 2 x 106 cells were transfected with 2 µg of small interfering RNA (siRNA) by the nucleofection technique in buffer V (nucleofection protocol T-20). The nucleofector device and a nucleofection kit were obtained from Amaxa (Cologne, Germany) and used in accordance with the manufacturer's recommendations. At 30 h after transfection, cells were harvested, counted, and processed for cell motility assays or Western blotting.
Cell motility assays. For wound-healing assays, wounds were generated by 20-µl pipette tips in confluent cultures of cells growing in 6- or 24-well plates. Areas of wounds were marked and photographed at different time points using a digital camera attached to a phase-contrast microscope (Nikon TE 2000-S). Where indicated, cells were maintained in the presence of DOX for 48 h prior to the creation of wounds. A transwell migration assay was performed using 24-well transwell plates containing 8-µm-pore-size polycarbonate filters (Corning Costar Corp., Cambridge, MA). A total of 1 x 105 cells were added to the top chambers and incubated overnight. Adhered cells were allowed to migrate toward serum gradient used as a chemoattractant in the lower chamber for 4 h. Those cells that did not migrate through the pores in the membrane were removed by scraping the membrane with a cotton swab. Cells that migrated to the underside of transwell filters were fixed, stained with a Gurr rapid staining kit (BDH), and counted by bright-field microscopy at a magnification of x200 in four random fields using the ImageJ program.
| RESULTS |
|---|
|
|
|---|
|
|
|
|
Prolonged and short-term (to a lesser extent) expression of Ec1WVM activates tumor cell migration. Prolonged and short-term expression of Ec1WVM in A431 cells resulted in loss of an epithelial pattern of cell growth and in cell dissociation (Fig. 1 and 3). Prolonged Ec1WVM expression down-regulated keratins and activated expression of vimentin (Fig. 2 and Table 1). Since cells undergoing EMT acquire a migratory phenotype, we hypothesized that Ec1WVM may affect cell motility and lead to increased cell migration into a wound. To test this, wounds were created in confluent cultures of NT-2, W2, and 31D6 cells and 31D6 cells pretreated with DOX for 48 h, and closure of wounds was monitored after 8 and 17 h. As expected, cells expressing Ec1WVM displayed accelerated wound closure compared with NT-2 and 31D6 cells maintained in the absence of DOX (Fig. 4). Whereas migration of clones with epithelial morphology closed wounds by approximately 50% in 17 h, wounds disappeared in W2 cell cultures. DOX-treated 31D6 cells exhibited an intermediate motility, and in 17 h they migrated approximately 1.7-fold faster than untreated cells. The moderate activation of cell motility in DOX-treated 31D6 cells was statistically significant. On the other hand, DOX produced no effect on migration of stable W3 and NT-2 clones (data not shown).
|
|
AP-1 is critical for the activation of tumor cell motility by Ec1WVM. We aimed to evaluate whether the effect on tumor cell motility produced by Ec1WVM was AP-1 dependent. Migration into a wound of G10 or B4 cells either maintained without DOX or pretreated with DOX for 48 h was examined. As a positive control, we used a highly motile W3 cell line constitutively expressing Ec1WVM (Fig. 6). Expression of TAM67 not only counteracted the stimulatory effect of Ec1WVM on cell migration observed in 31D6 cells but also almost completely blocked cell motility (Fig. 6, clone G10). Moreover, activation of c-Fos in B4 cells was sufficient to stimulate cell migration into a wound (Fig. 6). Therefore, at early stages of EMT, Ec1WVM-mediated effects on tumor cell motility involve AP-1.
|
|
|
| DISCUSSION |
|---|
|
|
|---|
Transcriptional up-regulation of fra-1 and c-jun and functional activation of AP-1 are early events in Ec1WVM-mediated EMT. Stimulation of fra-1 and c-jun transcription can be blocked by TAM67-GFP (Fig. 5B), suggesting that Ec1WVM activates a positive autoregulatory mechanism that keeps AP-1 activity elevated in cells with compromised cell-cell adhesion.
A431 cells, as other cell lines derived from epithelial cancers, migrate as cell aggregates, sheets, or clusters (collective migration). In this form of migration, aggregated cells move as a functional unit, in which subsets of active cells utilize actin-mediated ruffles and generate integrin-dependent traction. Other cells included in an aggregate are passively dragged forward by means of intercellular adhesion (reviewed in reference 22). Given that induction of c-Fos in clone B4 does not affect epithelial morphology but is sufficient to accelerate cell motility, we conclude that collective migration of epithelial cells is positively regulated by AP-1. This conclusion is consistent with data generated by Malliri et al. showing that prolonged expression of a dominant-negative mutant of c-Jun blocks motility of nonstimulated A431 cells (35). Loss of cell-cell adhesion during EMT results in a switch from collective toward different forms of more efficient individual migration patterns (22). TAM67-GFP effectively blocks cell motility activated by Ec1WVM at an early EMT phase (clone G10). Completion of EMT further contributes to enhanced cell motility (stable clones expressing Ec1WVM are more active in the wounding-healing assay than 31D6 cells pretreated with DOX for 48 h). By RNA interference we demonstrated that enhanced expression of c-Jun and Fra-1 is required for active migration of W3 cells, e.g., at later EMT stages. Taken together, these data clearly demonstrate that the role of AP-1 in cell motility is not restricted to the control of the epithelial type of cell migration. A positive autoregulatory loop, which is triggered by Ec1WVM and activates transcription of fra-1 and c-jun genes, is essential for enhanced cell motility at different stages of EMT.
We were interested to identify Ec1WVM-mediated signaling providing an initial activating stimulus to the preexisting AP-1 complexes. Since abundance, activity, and composition of AP-1 complex is controlled by MAPK, we examined whether expression and phosphorylation levels of MAPK are affected by DOX in 31D6 cells. Even though we did observe a moderate increase in the phosphorylation level of MAPKs in DOX-treated 31D6 cells (data not shown), the exact molecular events triggering induction of AP-1 by Ec1WVM remain unclear. A431 cells express high levels of EGFR and are capable of autocrine stimulation of this receptor. As E-cadherin-mediated adhesion may inhibit ligand-dependent activation of RTK (48), we hypothesized that the application of Ec1WVM would result in activation of EGFR in the A431 cell system. However, Ec1WVM had no effect on phosphorylation of EGFR in DOX-stimulated 31D6 cells (Fig. 8), suggesting that RTK pathways are unlikely to be involved in the activation of AP-1 by Ec1WVM. Nor is ß-catenin signaling, known to activate fra-1 and c-jun gene transcription, involved in Ec1WVM-mediated activation of AP-1 (Fig. 3D). One of the hallmarks of EMT is the reorganization of the actin-based cytoskeleton, which reflects loss of epithelial polarity and a switch from cell-cell to cell-substratum interactions. Recently, we found that expression of Ec1WVM in c-Fos-transformed murine epithelioid carcinoma cells resulted in increased cell adhesion to the extracellular matrix components (J. Mejlvang et al., unpublished data). Therefore, we suggest that Ec1WVM may affect cell-substratum interactions also in the A431 cell system, stimulating integrin signaling and hence triggering the initial AP-1 activation. The documented reciprocity between the level of organization of adherens junctions and focal adhesions (31), as well as previously described cross talks between E-cadherin and specific integrin receptors (65), supports this hypothesis.
EMT-inducing transcription factors Snail, Slug, ZEB-2/SIP1, or E47 directly inhibit the E-cadherin gene promoter. Emerging evidence suggests that these transcriptional repressors act downstream of a variety of EMT-initiating signals to down-regulate E-cadherin gene transcription (15, 20, 23, 45). In addition to transcriptional repression, several other genetic and epigenetic mechanisms may be responsible for inactivation of E-cadherin-dependent cell-cell adhesion in human cancers. E-cadherin function can be inhibited by gene mutations, promoter polymorphisms, promoter hypermethylation, and loss of the E-cadherin locus (4, 17, 26, 66). For instance, in poorly differentiated diffuse-type gastric cancer and lobular breast carcinoma, mutations affecting the extracellular E-cadherin domain have been observed. Our data suggest that structural mutations in the E-cadherin gene or consistent cleavage of E-cadherin extracellular domains chronically exposed to matrix metalloproteinases secreted by stromal cells (32) may be sufficient to trigger a process ultimately leading to EMT in tumor cells. Often, cells respond relatively rapidly to EMT-initiating signals. For example, 5 days of chronic EGF treatment is sufficient to induce morphological transformation and to down-regulate epithelial markers in A431 cells (34). In the same cell line, the transcription factor ZEB-2/SIP1 induced full EMT as rapidly as within 48 h (our unpublished data). In contrast, EMT induced by the dominant-negative E-cadherin mutant is a slow process. Different kinetics of EMTs mediated by an E-cadherin mutant and its transcriptional repressors may indicate that the repressors directly inhibit transcription of other epithelial genes and, therefore, have broader functions in EMT. In support of this, Snail has been shown to down-regulate tight junction components independently of E-cadherin down-regulation (41). Interestingly, rapid EMT of MDCK cells mediated by ectopic expression of Snail involves inhibition of G1/S cell cycle progression (62). A similar effect of exogenous ZEB-2/SIP1 on retinoblastoma protein-dependent cell cycle regulation was observed in the A431/SIP1 model (our unpublished data). This suggests that cells retaining control over G1/S transition and undergoing a rapid EMT acquire a growth disadvantage. In contrast, neither the cell proliferation rate nor cell cycle progression was affected in the EMT model reported here (data not shown). Therefore, it is plausible to speculate that SIP1 or Snail induces either transient EMT or stable EMT only in those cells in which control over G1/S transition has been lost. Gradual EMT initiated by mutations of the components of E-cadherin complex or by cleavage of E-cadherin by proteases may be a prevalent mechanism of stable EMT in cancer cells, in which the control over G1/S transition is not completely compromised (such as A431 or MDCK cells).
Prolonged inhibition of epithelial adhesion alters expression of several genes that are critical players in signal transduction pathways controlling tumor cell motility and invasive growth. The challenge is to further elucidate molecular mechanisms linking inhibition of epithelial cell adhesion with the alterations in cell signaling networks. This may lead to the design of novel methods uncoupling the loss of E-cadherin from tumor cell invasion and metastasis.
| ACKNOWLEDGMENTS |
|---|
We acknowledge financial support from the Prostate Research Campaign UK and the European Association of Urology (European Urology Scholarship Programme).
| FOOTNOTES |
|---|
H.A. and J.M. contributed equally to this work. ![]()
| REFERENCES |
|---|
|
|
|---|
2. Batlle, E., E. Sancho, C. Franci, D. Dominguez, M. Monfar, J. Baulida, A. De Herreros. 2000. The transcription factor snail is a repressor of E-cadherin gene expression in epithelial tumour cells. Nat. Cell. Biol. 2:84-89.[CrossRef][Medline]
3. Bergers, G., P. Graninger, S. Braselmann, C. Wrighton, and M. Busslinger. 1995. Transcriptional activation of the fra-1 gene by AP-1 is mediated by regulatory sequences in the first intron. Mol. Cell. Biol. 15:3748-3758.[Abstract]
4. Berx, G., K. Becker, H. Hofler, and F. van Roy. 1998. Mutations of the human E-cadherin (CDH1) gene. Hum. Mutat. 12:226-237.[CrossRef][Medline]
5. Betson, M., E. Lozano, J. Zhang, and V. Braga. 2002. Rac activation upon cell-cell contact formation is dependent on signaling from the epidermal growth factor receptor. J. Biol. Chem. 277:36962-36969.
6. Bolos, V., H. Peinado, M. Perez-Moreno, M. Fraga, M. Esteller, and A. Cano. 2003. The transcription factor Slug represses E-cadherin expression and induces epithelial to mesenchymal transitions: a comparison with Snail and E47 repressors. J. Cell Sci. 116:499-511.
7. Caca, K., F. Kolligs, J. X. M. Hayes, J. Qian, A. Yahanda, D. Rimm, J. Costa, and E. Fearon. 1999. Beta- and gamma-catenin mutations, but not E-cadherin inactivation, underlie T-cell factor/lymphoid enhancer factor transcriptional deregulation in gastric and pancreatic cancer. Cell. Growth Differ. 10:369-376.
8. Cano, A., M. A. Pérez-Moreno, I. Rodrigo, A. Locascio, M. J. Blanco, M. G. del Barrio, F. Portillo, and M. A. Nieto. 2000. The transcription factor Snail controls epithelial-mesenchymal transitions by repressing E-cadherin expression. Nat. Cell. Biol. 2:76-83.[CrossRef][Medline]
9. Cavallaro, U., and G. Christofori. 2004. Cell adhesion and signalling by cadherins and Ig-CAMs in cancer. Nat. Rev. Cancer. 4:118-132.[Medline]
10. Celis, J. E., P. Gromov, T. Cabezon, J. Moreira, N. Ambartsumian, K. Sandelin, F. Rank, I. Gromova. 2004. Proteomic characterization of the interstitial fluid perfusing the breast tumor microenvironment: a novel resource for biomarker and therapeutic target discovery. Mol. Cell Proteomics 3:327-344.
11. Celis, J. E., S. Trentemølle, and P. Gromov. Gel-based proteomics: high resolution two-dimensional gel electrophoresis of proteins. Isoelectric focusing (IEF) and nonequilibrium pH gradient electrophoresis (NEPHGE). In J. E. Celis, N. Carter, T. Hunter, D. Shotton, K. Simons and J. V. Small (ed.), Cell biology, a laboratory handbook, vol. 4, 3rd ed., in press. Academic Press, San Diego, Calif.
12. Chitaev, N., S. Troyanovsky. 1998. Adhesive but not lateral E-cadherin complexes require calcium and catenins for their formation. J. Cell Biol. 142:837-846.
13. Cohn, M., I. Hjelmso, L.-C. Wu, P. Guldberg, E. Lukanidin, and E. Tulchinsky. 2001. Characterization of Sp1, AP-1, CBF and KRC binding sites and minisatellite DNA as functional elements of the metastasis-associated mts1/S100A4 gene intronic enhancer. Nucleic Acids Res. 29:3335-3346.
14. Comijn, J., G. Berx, P. Vermassen, K. Verschueren, L. van Grunsven, E. Bruyneel, M. Mareel, D. Huylebroeck, and F. van Roy. 2001. The two-handed E box binding zinc finger protein SIP1 downregulates E-cadherin and induces invasion. Mol. Cell 7:1267-1278.[CrossRef][Medline]
15. Conacci-Sorrell, M., I. Simcha, T. Ben-Yedida, J. Blechman, P. Savanger, and A. Ben-Ze'ev. 2003. Autoregulation of E-cadherin expression by cadherin-cadherin interactions: the roles of b-catenin signalling, Slug and MAPK. J. Cell Biol. 163:847-857.
16. Conacci-Sorrell, M., J. Zhurinsky, and A. Ben-Ze'ev. 2002. The cadherin-catenin adhesion system in signalling and cancer. J. Clin. Investig. 109:987-991.[CrossRef][Medline]
17. Di Croce, L., and P. Pelicci. 2003. Tumour-associated hypermethylation: silencing E-cadherin expression enhances invasion and metastasis. Eur. J. Cancer. 39:413-414.
18. Edme, N., J. Downward, J.-P. Thiery, and B. Boyer. 2002. Ras induces NBT-II epithelial cell scattering through the coordinate activities of Rac and MAPK pathways. J. Cell Sci. 115:2591-2601.
19. Eger, A., A. Stockinger, B. Schaffhauser, H. Beug, and R. Foisner. 2000. Epithelial mesenchymal transition by c-Fos estrogen receptor activation involves nuclear translocation of beta-catenin and upregulation of beta-catenin/lymphoid enhancer binding factor-1 transcriptional activity. J. Cell Biol. 148:173-188.
20. Eger, A., K. Aigner, S. Sonderegger, B. Dampier, S. Oehler, M. Schreiber, G. Berx, A. Cano, H. Beug, and R. Foisner. 2005. DeltaEF1 is a transcriptional repressor of E-cadherin and regulates epithelial plasticity in breast cancer cells. Oncogene 24:2375-2385.[CrossRef][Medline]
21. Fialka, I., H. Schwarz, E. Reichmann, M. Oft, M. Busslinger, H. Beug. 1996. The estrogen-dependent c-JunER protein causes a reversible loss of mammary epithelial cell polarity involving a destabilization of adherens junctions. J. Cell Biol. 132:1115-1132.
22. Friedl, P., and K. Wolf. 2003. Tumour-cell invasion and migration: diversity and escape mechanisms. Nat Rev. Cancer 3:362-374.[CrossRef][Medline]
23. Fujita, N., D. Jaye, M. Kajita, C. Geigerman, C. Moreno, and P. Wade. 2003. MTA3, a Mi-2/NuRD complex subunit, regulates an invasive growth pathway in breast cancer. Cell 113:207-219.[CrossRef][Medline]
24. Gottardi, C., E. Wong, and B. Gumbiner. 2001. E-cadherin suppresses cellular transformation by inhibiting ß-catenin signalling in an adhesion-independent manner. J. Cell Biol. 153:1049-1060.
25. Gromova, I., and. J. E. Celis. Protein detection in gels by silver staining: a procedure compatible with mass-spectrometry in cell biology. In J. E. Celis, N. Carter, T. Hunter, D. Shotton, K. Simons and J. V. Small (ed.), Cell biology, a laboratory handbook, vol. 4, 3rd ed., in press. Academic Press, San Diego, Calif.
26. Guilford, P., J. Hopkins, J. Harraway, M. McLeod, N. McLeod, P. Harawira, H. Taite, R. Scoular, A. Miller, and A. Reeve. 1998. E-cadherin germline mutations in familial gastric cancer. Nature 392:402-405.[CrossRef][Medline]
27. Hennigan, R., and P. Stambrook. 2001. Dominant negative c-jun inhibits activation of the cyclin D1 and cyclin E kinase complexes. Mol. Biol. Cell 12:2352-2363.
28. Hoschuetzky, H., H. Aberle, and R. Kemler. 1994. Beta-catenin mediates the interaction of the cadherin-catenin complex with epidermal growth factor receptor. J. Cell Biol. 127:1375-1380.
29. Huelsken, J., and W. Birchmeier. 2001. New aspects of Wnt signaling pathways in higher vertebrates. Curr. Opin. Genet. Dev. 11:547-553.[CrossRef][Medline]
30. Kovacs, E. M., R. Ali, A. McCormack, and A. Yap. 2002. E-cadherin homophilic ligation directly signals through Rac and phosphatidylinositol 3-kinase to regulate adhesive contacts. J. Biol. Chem. 277:6708-6718.
31. Levenberg, S., B.-Z. Katz, K. Yamada, and B. Geiger. 1998. Long-range and selective autoregulation of cell-cell or cell-matrix adhesions by cadherin or integrin ligands. J. Cell Sci. 111:347-357.[Abstract]
32. Lochter, A., S. Galosy, J. Muschler, N. Freedman, Z. Werb, and M. Bissel. 1997. Matrix metalloproteinase stromelysin-1 triggers a cascade of molecular alterations that leads to stable epithelial-to-mesenchymal conversion and premalignant phenotype in mammary epithelial cells. J. Cell Biol. 139:1861-1872.
33. Lozano, E., M. Betson, and V. Braga. 2003. Tumor progression: small GTPases and loss of cell-cell adhesion. Bioessays 25:452-463.[CrossRef][Medline]
34. Lu, Z., S. Ghosh, Z. Wang, and T. Hunter. 2003. Downregulation of caveolin-1 function by EGF leads to the loss of E-cadherin, increased transcriptional activity of beta-catenin, and enhanced tumor cell invasion. Cancer Cell 4:499-515.[CrossRef][Medline]
35. Malliri, A., M. Symons, R. Hennigan, A. Hurlstone, R. Lamb, T. Wheeler, and B. Ozanne. 1998. The transcription factor AP-1 is required for EGF-induced activation of Rho-like GTPases, cytoskeletal rearrangements, motility, and in vitro invasion of A431 cells. J. Cell Biol. 143:1087-1099.
36. Mann, B., M. Gelos, A. Siedow, M. Hanski, A. Gratchev, M. Ilyas, W. Bodmer, M. Moyer, E. Riecken, H. Buhr, and C. Hanski. 1999. Target genes of beta-catenin-T cell-factor/lymphoid-enhancer-factor signaling in human colorectal carcinomas. Proc. Natl. Acad. Sci. USA 96:1603-1608.
37. Matsuo, K., J. Ownes, M. Tonko, C. Elliott, T. Chambers, and E. Wagner. 2000. Fosl1 is a transcriptional target of c-Fos during osteoclast differentiation. Nat. Genet. 24:184-187.[CrossRef][Medline]
38. Nieman, M., J. B. Kim, K. R. Johnson, and M. J. Wheelock. 1999. Mechanism of extracellular domain-deleted dominant negative cadherins. J. Cell Sci. 112:1621-1632.[Abstract]
39. Noren, N., W. Arthur, and K. Burridge. 2003. Cadherin engagement inhibits RhoA via p190RhoGAP. J. Biol. Chem. 278:13615-13618.
40. O'Farrell, P. H. 1975. High resolution two-dimensional electrophoresis of proteins. J. Biol. Chem. 250:4007-4021.
41. Ohkubo, T., and M. Ozawa. 2004. The transcription factor Snail downregulates the tight junction components independently of E-cadherin downregulation. J. Cell Sci. 117:1675-1685.
42. Orsulic, S., O. Huber, H. Aberle, S. Arnold, and R. Kemler. 1999. E-cadherin binding prevents ß-catenin nuclear localisation and ß-catenin/LEF-1-mediated transactivation. J. Cell Sci. 112:1237-1245.[Abstract]
43. Pece, S., M. Chiariello, C. Murga, and J. Gutkind. 1999. Activation of the protein kinase Akt/PKB by the formation of E-cadherin-mediated cell-cell junctions. Evidence for the association of phosphatidylinositol 3-kinase with the E-cadherin adhesion complex. J. Biol. Chem. 274:19347-19351.
44. Pece, S., and J. Gutkind. 2000. Signaling from E-cadherins to the MAPK pathway by the recruitment and activation of epidermal growth factor receptors upon cell-cell contact formation. J. Biol. Chem. 275:41227-41233.
45. Peinado, H., M. Quintanilla, and A. Cano. 2003. Transforming growth factor ß-1 induces snail transcription factor in epithelial cell lines. J. Biol. Chem. 278:21113-21123.
46. Perez-Moreno, M. A., A. Locascio, I. Rodrigo, G. Dhondt, F. Portillo, M. A. Nieto, and A. Cano. 2001. A new role for E12/E47 in the repression of E-cadherin expression and epithelial-mesenchymal transitions. J. Biol. Chem. 27629:27424-27431.
47. Polakis, P. 2000. Wnt signalling and cancer. Genes Dev. 14:1837-1851.
48. Qian, X., T. Karpova, A. Sheppard, J. McNally, and D. Lowy. 2004. E-cadherin-mediated adhesion inhibits ligand-dependent activation of diverse receptor tyrosine kinases. EMBO J. 21:1739-1784.[CrossRef]
49. Rappsilber, J., Y. Ishihama, and M. Mann. 2003. Stop and go extraction tips for matrix-assisted laser desorption/ionization, nanoelectrospray, and LC/MS sample pretreatment in proteomics. Anal. Chem. 75:663-670.[Medline]
50. Sambrook, J., and D. W. Russell. 2001. Molecular cloning: a laboratory manual, 3rd ed. Cold Spring Harbor Laboratory Press, Cold Spring Harbor, N.Y.
51. Sanson, B., P. White, and J.-P. Vincent. 1996. Uncoupling cadherin-based adhesion from wingless signalling in Drosophila. Nature 383:627-630.[CrossRef][Medline]
52. Savagner, P. 2001. Leaving the neighborhood: molecular mechanisms involved during epithelial-mesenchymal transition. Bioessays 23:912-923.[CrossRef][Medline]
53. Schramek, H., E. Feifel, E. Healy, and V. Pollack. 1997. Constitutively active mutant of the mitogen-activated protein kinase kinase MEK1 induces epithelial dedifferentiation and growth inhibition in Madin-Darby canine kidney-C7 cells. J. Biol. Chem. 272:11426-11433.
54. Shapiro, L., A. M. Fannon, P. D. Kwong, A. Thompson, M. S. Lehmann, G. Grubel, J.-F. Legrand, J. Als-Neilsen, D. R. Colman, and W. A. Hendrickson. 1995. Structural basis of cell-cell adhesion by cadherins. Nature 374:327-337.[CrossRef][Medline]
55. Shevchenko, A., M. Wilm, O. Vorm, and M. Mann. 1996. Mass spectrometric sequencing of proteins silver-stained polyacrylamide gels. Anal. Chem. 68:850-858.[Medline]
56. St. Croix, B., C. Sheehan, J. Rak, V. Florenes, J. Slingerland, and R. Kerbel. 1998. E-cadherin-dependent growth suppression is mediated by the cyclin-dependent kinase inhibitor p27(KIP1). J. Cell Biol. 142:557-571.
57. Stockinger, A., A. Eger, J. Wolf, H. Beug, and R. Foisner. 2001. E-cadherin regulates cell growth by modulating proliferation-dependent beta-catenin transcriptional activity. J. Cell Biol. 154:1185-1196.
58. Takeichi, M. 1993. Cadherins in cancer: implications for invasion and metastasis. Curr. Opin. Cell Biol. 5:806-811.[CrossRef][Medline]
59. Thiery, J.-P. 2003. Epithelial-mesenchymal transitions in development and pathologies. Curr. Opin. Cell Biol. 15:740-746.[CrossRef][Medline]
60. Thiery, J.-P., and D. Chopin. 1999. Epithelial cell plasticity in development and tumor progression. Cancer Metastasis Rev. 18:31-42.[CrossRef][Medline]
61. van de Wetering, M., N. Barker, I. Harkes, M. van der Heyden, N. Dijk, A. Hollestelle, J. Klijn, H. Clevers, and M. Schutte. 2001. Mutant E-cadherin breast cancer cells do not display constitutive Wnt signaling. Cancer Res. 61:278-284.
62. Vega, S., A. Morales, O. Ocaña, F. Valdés, I. Fabregat, and M. A. Nieto. 2004. Snail blocks the cell cycle and confers resistance to cell death. Genes Dev. 118:1131-1143.
63. Vial, E., E. Sahai, and C. J. Marshall. 2003. ERK-MAPK signaling coordinately regulates activity of Rac1 and RhoA for tumor cell motility. Cancer Cell 4:67-79.[CrossRef][Medline]
64. Vleminckx, K., and R. Kemler. 1999. Cadherins and tissue formation: integrating adhesion and signalling. Bioessays 21:211-220.[CrossRef][Medline]
65. von Schlippe, M., J. Marshall, P. Perry, M. Stone, A. J. Zhu, and I. R. Hart. 2000. Functional interaction between E-cadherin and av-containing integrins in carcinoma cells. J. Cell Sci. 113:425-437.[Abstract]
66. Wang, H.-D., J. Ren, and L. Zhang. 2004. CDH1 germline mutation in hereditary gastric carcinoma. World J. Gastroenterol. 10:3088-3093.[Medline]
67. Wong, A., and B. Gumbiner. 2003. Adhesion-independent mechanism for suppression of tumor cell invasion by E-cadherin. J. Cell Biol. 161:1191-1203.
68. Zantek, N., M. Azimi, M. Fedor-Chaiken, B. Wang, R. Brackenbury, and M. Kinch. 1999. E-cadherin regulates the function of the EphA2 receptor tyrosine kinase. Cell Growth Differ. 10:629-638.
This article has been cited by other articles:
| ||||||||||||