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[Cancer Research 65, 8836-8845, October 1, 2005]
© 2005 American Association for Cancer Research

Cell and Tumor Biology

E-Cadherin Regulates the Association between ß-Catenin and Actinin-4

Yasuharu Hayashida1,2, Kazufumi Honda1, Masashi Idogawa1, Yoshinori Ino1, Masaya Ono1, Akihiko Tsuchida2, Tatsuya Aoki2, Setsuo Hirohashi1 and Tesshi Yamada1

1 Chemotherapy Division and Cancer Proteomics Project, National Cancer Center Research Institute and 2 Third Department of Surgery, Tokyo Medical University, Tokyo, Japan

Requests for reprints: Tesshi Yamada, Chemotherapy Division, National Cancer Center Research Institute, 5-1-1 Tsukiji, Chuo-ku, Tokyo 104-0045, Japan. Phone: 81-3-3542-2511; Fax: 81-3-3547-6045; E-mail: tyamada{at}ncc.go.jp.


   Abstract
Top
 Abstract
Introduction
 Materials and Methods
 Results
 Discussion
 References
 
The E-cadherin/catenin system acts as an invasion suppressor of epithelial malignancies. This invasion suppressive activity seems be mediated not only by the cell adhesive activity of E-cadherin but by other undetermined signaling pathways elicited by ß-catenin. In fact, cancer cells that have infiltrated the stroma reduce the expression of E-cadherin and accumulate ß-catenin. We attempted to identify the alternative partner proteins that make complexes with ß-catenin in the absence of E-cadherin. An ~100-kDa protein was constantly coimmunoprecipitated with ß-catenin from SW480 colorectal cancer cells, which lack the expression of E-cadherin, and was identified as actinin-4 by mass spectrometry. Transfection of E-cadherin cDNA suppressed the association between ß-catenin and actinin-4. Inhibition of E-cadherin by RNA interference transferred the ß-catenin and actinin-4 proteins into the membrane protrusions of DLD-1 cells. Immunofluorescence histochemistry of clinical colorectal cancer specimens showed that the ß-catenin and actinin-4 proteins were colocalized in colorectal cancer cells infiltrating the stroma. We reported previously that overexpression of actinin-4 induces cell motility and specifically promotes lymph node metastasis by colorectal cancer. The association between ß-catenin and actinin-4 and its regulation by E-cadherin may represent a novel molecular link connecting cell adhesion and motility. Shutting down the signals mediating this association may be worth considering as a therapeutic approach to cancer invasion and metastasis.


   Introduction
Top
 Abstract
 Introduction
Materials and Methods
 Results
 Discussion
 References
 
Accumulating bodies of evidence indicate that E-cadherin acts as an invasion suppressor of various epithelial malignancies (1–3). Nontransformed Madin-Darby canine kidney cells acquire invasive properties following the addition of anti-E-cadherin antibody (4). Transfection of E-cadherin cDNA suppresses the invasive migration of L-cells (5). In the process of invasion, cancer cells are dissociated from tumor nests and infiltrate the stroma in a scattered manner (6). The primary cause of cancer invasion is thus believed to be a disturbance of the integrity of intercellular adhesion. However, recent experimental results have implied that adhesion-independent mechanisms are also involved in the suppression of tumor cell invasion by E-cadherin (7, 8). E-cadherin has been implicated not only in cell adhesion but also in the signaling of cell migration, proliferation, and survival (5, 9, 10). E-cadherin and its cytoplasmic binding proteins, catenins and p120ctn, are involved in tyrosine kinase and Rho signaling pathways (11–13). The involvement of ß-catenin-mediated pathway(s) in the invasion suppressor activity of E-cadherin was proposed recently. The activity of E-cadherin is mediated through its ß-catenin-binding domain but not through its p120ctn-binding domain (14). Accumulation of ß-catenin is frequently observed in the invasive front of clinical colorectal cancer (6, 15). However, transcriptional activity of the ß-catenin and T-cell factor (TCF)/lymphoid enhancer factor (LEF) complex is not required for the invasion-suppressing activity of E-cadherin (14). Based on these findings, we hypothesized that dysfunction of E-cadherin liberates ß-catenin protein from the cadherin/catenin complexes and elicits an alternative ß-catenin-mediated pathway that makes cancer cells more motile and invasive.

ß-Catenin is a multifunctional protein whose functional role and subcellular localization are determined by partner proteins that form complexes with it. We attempted to identify alternative partner proteins that form complexes with ß-catenin in the absence of E-cadherin, and we found that ß-catenin associates with actinin-4. Actinin-4 is a cell motility–associated actin-binding protein that was first identified in our laboratory (16–18). The cytoplasmic localization of actinin-4 is closely associated with the invasive phenotype of breast cancer and is a predictor of the outcome of breast cancer patients (17). A cDNA microarray analysis identified actinin-4 as a significant prognostic predictor in non–small cell lung cancer patients (19). We reported recently that increased expression of actinin-4 significantly increases cell motility and mediates invasive growth and lymph node metastasis by colorectal cancer (20). In this article, we report an association between ß-catenin and actinin-4 and regulation of the association between them by E-cadherin. The dynamic shift of the ß-catenin protein from the cell adhesion complex into a complex containing actinin-4 may evoke cell movement and mediate cancer invasion and metastasis.


   Materials and Methods
Top
 Abstract
 Introduction
 Materials and Methods
Results
 Discussion
 References
 
Antibodies. Anti-hemagglutinin (HA) rat monoclonal antibody (clone 3F10) was purchased from Roche Diagnostics GmbH (Mannheim, Germany). Anti-ß-catenin mouse monoclonal antibody (clone 14) was purchased from BD Transduction Laboratories (Palo Alto, CA). Anti-ß-catenin goat polyclonal (C-18) and anti-{alpha}-catenin rabbit polyclonal (SC-7894) antibodies were purchased from Santa Cruz Biotechnology (Santa Cruz, CA). Anti-ß-catenin (9562) and poly(ADP-ribose) polymerase-1 (PARP-1; 9542) rabbit polyclonal antibodies were purchased from Cell Signaling Technology, Inc. (Beverly, MA; ref. 21). Anti-ß-actin mouse monoclonal (AC-15) antibody was obtained from Abcam Ltd. (Cambridgeshire, United Kingdom). Anti-human E-cadherin mouse monoclonal (HECD-1) and anti-actinin-4 rabbit polyclonal (Ab-1 and Ab-2) antibodies were produced as described previously (20, 22).

Cell culture. Human colorectal cancer cell lines SW480 and LS411N and pancreatic cancer cell line BxPC-3 were purchased from the American Type Culture Collection (Rockville, MD). The human colorectal cancer cell line DLD-1 and the human embryonal kidney epithelial cell line HEK 293 were obtained from the Health Science Research Resources Bank (Osaka, Japan). The human colorectal cancer cell line COLO-320 was obtained from the Riken Cell Bank (Tsukuba, Japan). The human colorectal cancer cell line NCC-CO31 was established in our laboratory (23). A colorectal cancer cell line capable of inducing the actinin-4 protein (DLD-1 Tet-off ACTN4) and its control (DLD-1 Tet-off Control) were established using the tetracycline-regulatory promoter system as described previously (20). Doxycycline (Sigma-Aldrich, St. Louis, MO) was added to the culture medium to final concentrations of 0.01 µg/mL for maintenance and 0.1 µg/mL for suppression of induction.

Poly-D-lysine (control)– and poly-D-lysine/laminin–coated culture dishes and glass coverslips were obtained from BD Biosciences (Franklin Lakes, NJ).

Immunoblot analysis. Cells were extracted on ice for 30 minutes with lysis buffer [10 mmol/L HEPES (pH 7.4), 150 mmol/L NaCl, 1 mmol/L EDTA, 1% Triton X-100, 1% NP40, 1 mg/mL NaN3] containing a protease inhibitor cocktail (Sigma-Aldrich) before centrifugation at 12,000 x g for 30 minutes. Nuclear extracts were prepared with a CelLytic nuclear extraction kit (Sigma-Aldrich). Protein samples were separated by SDS-PAGE and transferred to Immobilon-P membranes (Millipore, Bedford, MA). After incubation with primary antibodies at 4°C overnight, the blots were detected with horseradish peroxidase–conjugated anti-mouse, anti-rabbit, and anti-goat IgG antibodies and enhanced chemiluminescence Western blotting detection reagents (Amersham Biosciences, Little Chalfont, United Kingdom) in accordance with the manufacturer's instructions.

Immunoprecipitation and mass spectrometry. Cell lysates were incubated with anti-ß-catenin polyclonal (sc-1496), anti-actinin-4 rabbit polyclonal antibody, normal goat IgG, or normal rabbit IgG overnight at 4°C and precipitated with Dynabeads Protein G (Dynal Biotech, Oslo, Norway). Immunoprecipitated proteins were separated by SDS-PAGE and detected by silver staining or immunoblotting. Proteins in gels were digested with modified trypsin (Promega, Madison, WI) and extracted as described previously (24). Matrix-assisted laser desorption/ionization was done with {alpha}-cyano-4-hydroxycinnamic acid as the matrix (Sigma-Aldrich). The masses of tryptic peptides were measured with a Voyager DE time-of-flight mass spectrometer (Applied Biosystems, Foster City, CA). Comparison of the mass values against the Swiss-Prot database was done using the Mascot search engine (Matrix Science, London, United Kingdom).

Immunofluorescence cytochemistry. Cells cultured on poly-L-lysine–coated (Asahi Technoglass Corp., Tokyo, Japan), poly-D-lysine/laminin–coated, or poly-D-lysine–coated glass coverslips were fixed with 4% paraformaldehyde for 10 minutes at room temperature and made permeable with 0.2% Triton X-100. After blocking with 10% normal swine serum (Vector Laboratories, Inc., Burlingame, CA) for 30 minutes at room temperature, the cells were incubated with various primary antibodies at 4°C overnight. After incubation with Alexa Fluor 488 anti-rat, anti-rabbit, anti-mouse, or anti-goat antibody, Alexa Fluor 594 anti-mouse or anti-rabbit IgG antibody, and Alexa Fluor 594-phalloidin (Molecular Probes, Inc., Eugene, OR), the specimens were observed under a laser scanning microscope (Radiance 2000 MP, Bio-Rad Laboratories, Hercules, CA).

Quantitative immunofluorescence histochemistry. Thin sections (5 µm) of formalin-fixed and paraffin-embedded specimens of colorectal cancer (26 cases) were used for the immunofluorescence histochemical analysis. After incubation with primary antibodies at 4°C overnight, each protein was detected with Alexa Fluor 594 anti-rabbit IgG and Alexa Fluor 488 anti-mouse IgG (Molecular Probes). Fluorescence intensity was evaluated using the surface plotting and line profiling functions of LaserPix image analysis software (Bio-Rad Laboratories) as described previously (20).

Inhibition of E-cadherin and actinin-4 expression by RNA interference. Two small interfering RNAs (siRNA) targeting E-cadherin (Genbank accession no. NM_004360) were generated: siRNAE-cad-1 5'-GGGUUAAGCACAACAGCAA-3' and siRNAE-cad-2 5'-CAGACAAAGACCAGGACUA-3'. Three siRNAs targeting actinin-4 (Genbank accession no. NM_004924) were generated: siRNAACTN4-1 5'-ACAAAGCGCUGGACUUUA-3', siRNAACTN4-2 5'-GUUCAUCGUCCAUACCAUC-3', and siRNAACTN4-3 5'-AAAGCCCUCAUUCGCAAGCAC-3'. Four control siRNAs were used: siRNANC08 (Nonspecific Control Duplex VIII, 52% GC content), siRNANC09 (Nonspecific Control Duplex IX, 47% GC content), siRNANC11 (Nonspecific Control Duplex XI, 36% GC content), and siRNACy3 (Cy3-conjugated). The siRNAs were synthesized by Dharmacon, Inc. (Lafayette, CO).

Plasmid construction and two-hybrid assay. The expression construct carrying mouse E-cadherin cDNA was generously provided by Dr. Masatoshi Takeichi (Kyoto University, Kyoto, Japan). SW480 cells were transiently transfected with LipofectAMINE 2000 reagent (Invitrogen, Carlsbad, CA).

Physical interaction between the actinin-4 and the ß-catenin proteins was assessed with a CheckMate Mammalian Two-Hybrid System (Promega) as instructed by the supplier. Serial cDNA constructs encoding partial amino acid sequences of the actinin-4 (BAA24447) and ß-catenin (NP_001895) proteins were amplified by PCR and subcloned into pBIND and pACT expression vectors, respectively. The composition of the constructs was confirmed by sequencing. Details of the procedures used for plasmid construction are available on request. Cells were transiently transfected in triplicate with FuGENE 6 reagent (Roche Diagnostics), and 48 hours after the transfection, luciferase activity was measured with the Dual-Luciferase Reporter Assay System (Promega) using Renilla reniformis luciferase activity as an internal control.

Cell migration assay. siRNAs were transfected into BxPC-3 cells with LipofectAMINE 2000, and 96 hours after transfection, 8 x 105 cells were seeded into each insert of a 24-well Biocoat Matrigel Invasion Chamber (BD Biosciences Discovery Labware) in triplicate. Fetal bovine serum was added to a final concentration of 10% in the lower chambers to induce cell migration. After incubation for 18 hours, the cells remaining above the insert membranes were carefully removed with cotton swabs, and cells that had migrated into the other sides of the membranes were stained with Diff-Quik kit (Sysmex Corp., Japan; ref. 25). The total numbers of migrated cells in five random microscopic fields were counted. The experiment was done twice.


   Results
Top
 Abstract
 Introduction
 Materials and Methods
 Results
Discussion
 References
 
Expression of E-cadherin and ß-catenin in cancer cells infiltrating the stroma. Expression of the E-cadherin and ß-catenin proteins was examined in 26 cases of colorectal carcinoma using quantitative immunofluorescence histochemistry (ref. 16; Fig. 1A-J). Decreased levels of E-cadherin expression (Fig. 1A and G) and increased levels of ß-catenin (Fig. 1C-F and H) were observed in cancer cells infiltrating the stroma. We confirmed the increased expression of ß-catenin by measuring the fluorescence intensity along lines of the same length placed on randomly selected cancer cells that formed a polarized glandular structure (Fig. 1D and F, red) and those infiltrating the stroma (blue). ß-Catenin protein had accumulated in the cytoplasm and nuclei of these cancer cells infiltrating the stroma (Fig. 1H and I).



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  		Figure 1. Expression of E-cadherin and ß-catenin in cancer cells infiltrating the stroma. A-D and G-I, immunofluorescence histochemical analysis of human colorectal cancer cells infiltrating the stroma using anti-E-cadherin mouse monoclonal antibody (A and G and green in C and I) and anti-ß-catenin rabbit polyclonal antibody (B, D, and H and red in C and I). E, three-dimensional visualization of the spatial distribution and expression levels of the ß-catenin protein in (D). F, representative histogram of the relative fluorescence intensity along a line placed on an invasive cancer nest (blue in D) and a randomly selected line placed on a cancerous gland exhibiting the polarized structure (red in D). J, immunoblot analysis of E-cadherin, {alpha}-catenin, ß-catenin (monoclonal antibody), actinin-4, and ß-actin proteins in colorectal cancer cell lines SW480 (lane 1), DLD-1 (lane 2), NCC-CO31 (lane 3), LS411N (lane 4), and COLO-320 (lane 5). Bar, 100 µm.

 
Immunoblot analysis revealed that two colon cancer cell lines, SW480 and COLO-320, lacked expression of E-cadherin (Fig. 1J). Because SW480 cells contained higher amounts of the ß-catenin protein than the other cell lines, we selected this cell line for identification of proteins associated with ß-catenin. N-cadherin was not expressed in any of the colorectal cancer cell lines used in this study (data not shown).

Identification of a novel association between ß-catenin and actinin-4. An ~100-kDa protein and a few other proteins were constantly coimmunoprecipitated with anti-ß-catenin antibody but not with normal control IgG (Fig. 2A). Peptide mass fingerprinting (Fig. 2B) and a protein database search revealed that the protein was actinin-4 (Fig. 2C).



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  		Figure 2. Identification of a novel association between ß-catenin and actinin-4. A, SDS-PAGE analysis of the immunoprecipitates from SW480 cells with anti-ß-catenin goat polyclonal antibody or control goat IgG. Arrowhead 1, 100-kDa protein; arrowhead 2, ß-catenin protein. B, mass spectrogram of the 100-kDa protein digested with modified trypsin. C, amino acid sequence of actinin-4. Bold letters, peptides that correspond to peaks identified by mass spectrometry. D, immunoblot analysis of the immunoprecipitates of SW480 cells with anti-ß-catenin goat polyclonal antibody, control goat IgG, anti-actinin-4 rabbit polyclonal antibody, and control rabbit IgG. The immunoprecipitates were blotted with anti-actinin-4 (top) and anti-ß-catenin (bottom) antibodies. E-G, immunofluorescence microscopy showing the subcellular localization of actinin-4 (E) and ß-catenin (F). G, merged images of (E and F). Bar, 10 µm. H, two-hybrid assay indicating interaction between serially constructed actinin-4 and ß-catenin mutants. HEK 293 cells were transiently transfected with pBIND-actinin-4 and pACT-ß-catenin constructs encoding the indicated amino acid sequences. RLU, relative luciferase unit.

 
Immunoblotting with anti-actinin-4 antibody confirmed the presence of actinin-4 protein in the immunoprecipitate with anti-ß-catenin antibody but not with control normal goat IgG (Fig. 2D, left). Conversely, the ß-catenin protein was detected in the immunoprecipitate with anti-actinin-4-antibody but not with control normal rabbit IgG (Fig. 2D, right), confirming that the ß-catenin and actinin-4 proteins formed a native complex in SW480 cells. Similar results were obtained with the other cell line COLO-320, which lacks expression of E-cadherin (Fig. 1J; data not shown).

Double immunofluorescence cytochemistry (Fig. 2E-G) showed that the ß-catenin (Fig. 2F) and actinin-4 (Fig. 2E) proteins were colocalized in bleb-like protrusions formed on the free surfaces of SW480 cell clusters (Fig. 2G, arrows). A two-hybrid assay revealed that the NH2-terminal amino acids 1 to 161 of actinin-4 and the NH2-terminal amino acids 1 to 249 of ß-catenin were responsible for the interaction (Fig. 2H).

E-cadherin regulates the association between ß-catenin and actinin-4. We investigated whether the interaction and spatial distribution of ß-catenin and actinin-4 are affected by expression of E-cadherin (Fig. 3). Transient transfection of E-cadherin cDNA into SW480 cells resulted in the formation of condensed cell colonies (Fig. 3A and B), a phenomenon that had been previously called "compaction" (26). Although the overall expression level of actinin-4 was unaffected by E-cadherin transfection (Fig. 3C, left), the amount of actinin-4 protein present in the immunoprecipitants with anti-ß-catenin antibody was significantly reduced (Fig. 3C, right, asterisk), indicating that the recovery of E-cadherin-mediated cell adhesion restrained the aberrant interaction between actinin-4 and ß-catenin in SW480 cells. Immunofluorescence microscopy revealed no colocalization of these two proteins was observed in SW480 cells transfected with E-cadherin cDNA (Fig. 3D-F).



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  		Figure 3. E-cadherin regulates the association between ß-catenin and actinin-4. A and B, phase-contrast images of SW480 cells transfected with E-cadherin cDNA (A) and control plasmid (B). Bar, 100 µm. C, whole-cell lysates and immunoprecipitants with anti-ß-catenin polyclonal antibody of SW480 cells transfected with E-cadherin cDNA or control plasmid were blotted with anti-E-cadherin, anti-actinin-4, anti-ß-actin, or anti-ß-catenin antibody. D-F, subcellular localization of actinin-4 (D and red in F) and ß-catenin (E and green in F) in SW480 cells transfected with E-cadherin cDNA. G, immunoblot analysis of E-cadherin and ß-actin expression in DLD-1 cells transfected with siRNAE-cad-1 (lane 1), siRNAE-cad-2 (lane 2), siRNANC08 (lane 3), siRNANC09 (lane 4), or siRNACy3 (lane 5). H-M, subcellular localization of actinin-4 (H and K and red in J and M) and ß-catenin (I and L and green in J and M) in DLD-1 cells transfected with siRNAE-cad-1 (H-J) or siRNANC08 (K-M). Bar, 100 µm. N, immunoblot analysis of actinin-4 and ß-actin expression in SW480 cells transfected with siRNAACTN4-1 (lane 1), siRNAACTN4-2 (lane 2), siRNAACTN4-3 (lane 3), siRNANC09 (lane 4), siRNANC11 (lane 5), or siRNACy3 (lane 6). O-T, subcellular localization of actinin-4 and (red in O, Q, R, and T) and ß-catenin (green in P, Q, S, and T) in SW480 cells transfected with siRNAACTN4-2 (O-Q) or siRNANC11 (R-T).

 
We then targeted E-cadherin using RNA interference (Fig. 3G-M). Transfection with either siRNAE-cad-1 or siRNAE-cad-2, but not with the relevant controls, significantly reduced the protein level of E-cadherin in DLD-1 cells (Fig. 3G) and intercellular adhesion (data not shown). The expression level of ß-catenin was significantly elevated by the inhibition of E-cadherin expression (Fig. 3I and L), and ß-catenin and actinin-4 proteins jointly distributed in the bleb-like protrusions formed at the free cell surfaces (Fig. 3J, arrowheads), the same as in SW480 cells lacking expression of E-cadherin (Fig. 2G). In the control transfection, ß-catenin and actinin-4 proteins overlapped mainly in the nucleus, and hardly any association between ß-catenin and actinin-4 was detected in the cell membranes (Fig. 3K-M). Similar results were obtained by disrupting the intercellular adhesion of DLD-1 cells with a functional monoclonal antibody that recognizes the ectodomain of E-cadherin, HECD-1 (ref. 22; data not shown).

Transfection with siRNAACTN4-1, siRNAACTN4-2, or siRNAACTN4-3, but not with the relevant control siRNAs, significantly reduced the protein level of actinin-4 in SW480 cells (Fig. 3N). Bleb-like protrusions disappeared from SW480 cells when actinin-4 was knocked down, and ß-catenin protein became distributed diffusely in the cytoplasm (Fig. 3O-Q), indicating that expression of actinin-4 is essential for the formation of bleb-like protrusions. Control transfection did not affect the colocalization of actinin-4 and ß-catenin in the bleb-like protrusions (Fig. 3R-T, arrowheads) or in the nucleus.

Actinin-4 recruits ß-catenin into actin-rich structures. We established previously a colorectal cancer cell clone capable of inducing actinin-4 under control of the tetracycline-regulatory promoter system (designated DLD-1 Tet-off ACTN-4). The removal of doxycycline from the culture medium increased the expression level of HA-tagged and overall actinin-4 proteins in DLD-1 Tet-off ACTN4 (Fig. 4A, left). Removal of doxycycline had no effects on the expression of endogenous ß-catenin in the DLD-1 Tet-off Control (Fig. 4A, right). On the induction of actinin-4, DLD-1 Tet-off ACTN-4 cells became scattered, spread protrusions (Fig. 4B and C), and significantly increased their motility (20). Immunofluorescence microscopy revealed that the ß-catenin and actinin-4 proteins were colocalized at the leading edges of these wavy protrusions (Fig. 4D-I), where the actin bundles were highly concentrated (Fig. 4J and L, red), and E-cadherin (Fig. 4K and L, green) or {alpha}-catenin (data not shown) protein was not colocalized with them.



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  		Figure 4. Actinin-4 recruits ß-catenin into actin-rich structures. A, immunoblot analysis of DLD-1 Tet-off ACTN4 (ACTN4) and control (Control) cells cultured in the presence [(+)] or absence [(–)] of 0.1 µg/mL doxycycline (Dox) for 72 hours with anti-HA, anti-actinin-4, anti-ß-catenin, and anti-actin antibodies. B and C, phase-contrast microscopy of DLD-1 Tet-off ACTN4 cells cultured in the presence [Dox (+); B] or absence [Dox (–); C] of 0.1 µg/mL doxycycline for 72 hours. D-L, immunofluorescence cytochemistry of DLD-1 Tet-off ACTN4 cells cultured in the absence of doxycycline for 72 hours with anti-HA (D and E and green in H and I), anti-ß-catenin (F and G and red in H and I), and E-cadherin (K and green in L) antibodies. Filamentous actin polymers were detected by Alexa Fluor 594-phalloidin (J and red in L). Squares in (D, F, and H) are enlarged as shown in (E, G, and I). Bar, 100 µm (B, C, H, and L) and 10 µm (I). M, immunoblot analysis of actinin-4 and ß-actin (control) expression in BxPC-3 cells transfected with siRNAACTN4-2 (lane 1), siRNAACTN4-3 (lane 2), siRNANC09 (lane 3), or siRNACy3 (lane 4). N, migration capacity of BxPC-3 cells transfected with siRNAACTN4-2 (lane 1), siRNAACTN4-3 (lane 2), siRNANC09 (lane 3), or siRNACy3 (lane 4). Columns, mean number of cells that invaded Matrigel insert membranes; bars, SE.

 
IEC6 Tet-off ß-catenin {Delta}N89 is capable of inducing stabilized ß-catenin protein on the removal of doxycycline from the culture medium (27). When cultured without doxycycline for 72 to 96 hours, the cell clusters of IEC6 Tet-off ß-catenin {Delta}N89 became dispersed, and individual cells spread filopodia and lamellipodia (28). The ß-catenin and actinin-4 proteins were preferentially colocalized at the edges of these podia and membrane ruffles (data not shown).

The knockdown of actinin-4 expression by siRNAs (Fig. 4M) significantly reduced the cell motility of a highly motile pancreatic cancer cell line, BxPC-3 (ref. 25; Fig. 4N).

Colocalization of ß-catenin and actinin-4 in colorectal cancer cells infiltrating the stroma. We then examined clinical colorectal cancer tissues by immunofluorescence histochemistry (Fig. 5). In cancer nests with polarized glandular structures (Fig. 5E and F, arrowheads), the actinin-4 protein lined the apical membranes of the glands (Fig. 5A and B), consistent with its association with Na+/H+ exchanger 3 (29). In these polarized glands, the ß-catenin protein was distributed mainly in the lateral membranes (Fig. 5C and D), and the locations of actinin-4 and ß-catenin seemed mostly apart from each other. In cancer cells that were infiltrating fibrous stroma and did not form glandular structures (Fig. 5E, arrows) and in cancer cells budding from the glandular structures (Fig. 5F, arrows), the distribution of actinin-4 and ß-catenin appeared to overlap.



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  		Figure 5. Colocalization of ß-catenin and actinin-4 in colorectal cancer cells infiltrating the stroma. Immunofluorescence histochemistry of colorectal cancer with anti-actinin-4 rabbit polyclonal (A and B and red in E and F) and anti-ß-catenin mouse monoclonal (C and D and red in E and F) antibodies. Bar, 100 µm.

 
Transition of ß-catenin and actinin-4 into filopodia and the nucleus by culturing cells on laminin. DLD-1 colorectal cancer cells maintained their expression of E-cadherin (Fig. 1J) and grew in cell clusters with tight cell adhesion (Fig. 6A). Immunofluorescence microscopy detected the ß-catenin protein principally in the cell borders and barely in the nucleus (Fig. 6C and E, green) and actinin-4 mainly in the nucleus (Fig. 6D and E, red). The ß-catenin and actinin-4 proteins partly overlapped (Fig. 6E). When transferred onto laminin-coated surfaces, the DLD-1 cells became scattered and extended sharp protrusions (Fig. 6B). The expression level of actinin-4 was elevated significantly on laminin (Fig. 6G and L), and the ß-catenin and actinin-4 proteins were jointly distributed in the filopodia-like protrusions at the cell periphery (Fig. 6I and J, arrowheads) and in the nucleus (Fig. 6F-K).



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  		Figure 6. Transition of ß-catenin and actinin-4 into the filopodia and the nucleus of cells cultured on laminin. A and B, phase-contrast microscopy of DLD-1 cells cultured on poly-D-lysine– (Control; A) or poly-D-lysine/laminin–coated (Laminin; B) culture dishes for 48 hours. C-K, immunofluorescence cytochemical analysis of a human colorectal cancer cell line, DLD-1, cultured on poly-D-lysine– (Control; C-E) or poly-D-lysine/laminin–coated (Laminin; F-K) glass coverslips for 48 hours using anti-ß-catenin monoclonal (C, F, and I and green in E, H, and K) and anti-actinin-4 rabbit polyclonal (D, G, and J and green in E, H, and K) antibodies. Squares in (C and H) are enlarged as shown in (E and I-K), respectively. L, immunoblot analyses of the amounts of actinin-4, ß-actin, and PARP-1 proteins present in whole lysates (left) and nuclear extracts (right) of DLD-1 cells cultured on laminin-coated dishes [(+)] or controls [(–)]. Bar, 100 µm (A-C and F) and 10 µm (I).

 

   Discussion
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
References
 
Overcoming several technical difficulties, we used a proteomic approach to search for partner proteins that formed native complexes with endogenous ß-catenin protein, because we wanted to avoid the discovery of artificial interactions, such as those often seen in transfection experiments. {alpha}-Catenin, a known ß-catenin-binding protein, was present in the immunoprecipitate with anti-ß-catenin antibody (data not shown), confirming the authenticity of the experimental procedures. We found that the ß-catenin protein formed a complex with actinin-4 (Fig. 2A-C) in an E-cadherin-deficient colorectal cancer cell line (Fig. 1J); this finding was confirmed by coimmunoprecipitation and immunoblot assays (Fig. 2D). In addition, formation of this complex was dynamically suppressed by expression of E-cadherin (Fig. 3C-F), and it was induced by disruption of the E-cadherin-mediated intercellular connection (Fig. 3G-M). Immunofluorescence microscopy revealed that the actinin-4 and ß-catenin proteins were colocalized in bleb-like membrane protrusions on the free surfaces of colorectal cancer cells (Figs. 2E-G and 3H-J and R-T). Because the binding domain of ß-catenin for actinin-4 (Fig. 2H) overlaps with its binding domain for E-cadherin (30), actinin-4 and E-cadherin may compete for the same binding domain of ß-catenin.

Mutations in the actinin-4 (ACTN4) gene have been identified as causative of familial focal segmental glomerulosclerosis (FSGS) syndrome (31), and failure of foot process extension by glomerular podocytes is considered the main cause of FSGS. The actinin-4 in cultured podocytes is localized in the sharply extended cell processes, where filamentous actin bundles are concentrated (32). Podosomes and invadopodia are morphologically similar to each other (33), and formation of the protrusions in colorectal cancer cells can be seen as analogous to the process of foot extension in glomerular podocytes.

Thus far, ß-catenin has been described to exist in at least three different subcellular locations: the adherens junction, free cytoplasmic pool, and nucleus. ß-Catenin was first described as a cell adhesion molecule that makes complexes with the cytoplasmic domains of cadherins (34). ß-Catenin binds to {alpha}-catenin and connects the adherens junction to the actin cytoskeleton (35, 36). In the cytoplasm, ß-catenin makes complexes with the adenomatous polyposis coli (APC) gene product, glycogen synthase kinase-3ß (GSK-3ß), and axin/axil (30). ß-Catenin is phosphorylated by GSK-3ß, and phosphorylated ß-catenin is specifically bound by ß-TrCP, a subunit of the E3 ubiquitin ligase complex, which ubiquitylates ß-catenin and thereby earmarks it for rapid proteosomal degradation (37). In the nucleus, ß-catenin makes complexes with TCF/LEF transcriptional factors (38, 39). The ß-catenin and TCF/LEF complexes transactivate genes for c-myc, cyclin D1, fibronectin, matrix metalloproteinase-7, CD44, TCF1, MDR1, etc. (40). However, the subcellular localization of ß-catenin described in this study (Figs. 2F, 3I and S, 4G, and 6I) appears different from all the above.

Actinin-4 is an actin-binding protein that is preferentially localized in moving structures, such as dorsal ruffles, lamellipodia, and filopodia (16, 17, 41), but not in the adherens junction (17). The expression level of actinin-4 was significantly increased in cells exhibiting enhanced motility, and the increased expression of actinin-4 dramatically changed the morphology and motility of colorectal cancer cells (ref. 20; Fig. 4B and C). Actinin-4 was up-regulated by Rac1 and Cdc42 but not by RhoA (42). Rac and Cdc42, but not Rho, are involved in the formation of filopodia and lamellipodia (43). The Rab5-specific GTPase-activating protein RN-Tre is critical to the formation of circular ruffles and has been reported to interact with actinin-4 (41). Meanwhile, ß-catenin has been implicated in cell motility and in the epithelial-mesenchymal transition. We observed previously that retroviral expression of stabilized ß-catenin induces epithelial-mesenchymal transition of intestinal epithelial cells (28). ß-Catenin has been reported to accumulate at the leading edges of migrating astrocytes (44). ß-Catenin and actinin-4 proteins were accumulated and colocalized in colorectal cancer cells infiltrating the stroma (Fig. 5). Dynamic regulation of the actin cytoskeleton by the various classes of actin-binding proteins plays a crucial role in cell movement (33). Knockdown of actinin-4 expression by siRNAs significantly reduced cell motility (Fig. 4M and N). Based on the above observations, we speculate that the complex containing ß-catenin and actinin-4 is involved in cell motility and cancer invasion.

Cancer invasion causes interactions between epithelial cancer cells and the extracellular matrix. The behavior of cancer cells infiltrating the stroma may also be affected by interaction with the extracellular matrix (45). We found laminin-induced scattering and podia extension of DLD-1 colorectal cancer cells (Fig. 6B). Laminin significantly increased the expression level of actinin-4 (Fig. 6L) and induced the transition of actinin-4 and ß-catenin to the nucleus and filamentous podia formed along the cell membrane (Fig. 6K). Actinin-4 is reported to interact with a hemidesmosome component, bullous pemphigoid antigen 2 (BP180; ref. 46). Hemidesmosomes are multimeric protein complexes that attach epithelial cells to their underlying extracellular matrices. Direct interaction between the intracellular domains of BP180 and integrin ß4 has been reported (47). Integrin {alpha}6ß4 is a receptor for laminin and is involved in the migration of cancer cells. The engagement of laminin by integrin {alpha}6ß4 can stabilize actin-rich protrusions and mediate the traction forces necessary for cell movement (48). Integrin {alpha}6ß4 mediates the phosphatidylinositol 3-kinase (PI3K) and Rho signaling pathways (49). Activation of the PI3K pathway by integrin {alpha}6ß4 enhances the formation of actin-rich protrusions (48). The interaction with extracellular matrix laminin seems to elicit a certain signaling pathway and translocate actinin-4 and ß-catenin to the actin-rich protrusions.

The colorectal cancer cell line DLD-1 (Figs. 3G-M, 4A-L, and 6) has a mutation in the APC gene but has retained the expression of E-cadherin (Figs. 1J and 3G). Consequently, ß-catenin accumulated modestly in the nuclei of DLD-1 cells (Fig. 3L). However, the nuclear expression of ß-catenin significantly increased after disruption of E-cadherin-mediated cell adhesion (Fig. 3I) and after attachment to the laminin substrate (Fig. 6F). A recent study has indicated that transcriptional activity of the ß-catenin and TCF/LEF complex is not required for the invasion-suppressing activity of E-cadherin (14). We observed that actinin-4 was colocalized with ß-catenin in the nucleus (Fig. 6K) but did not significantly enhance TCF/LEF transcriptional activity (data not shown). Nuclear translocation of actinin-4 seems to be regulated by the PI3K pathway (17). PI3K transduces several receptor tyrosine kinase signaling pathways and regulates cell growth, motility, and apoptosis (50, 51). Actinin-4 interacts with DNase Y in the nucleus and is involved in the regulation of apoptosis (52). We reported previously that actinin-4 existed in the nucleus of certain populations of less invasive breast cancer and several cell lines (17). The binding of ß-catenin to actinin-4 in the nucleus may induce cancer invasion through mechanisms other than the regulation of gene transcription.

In conclusion, we showed that the ß-catenin and actinin-4 proteins were associated in vitro and in vivo. The ß-catenin and actinin-4 complex was highly concentrated in actin-rich protrusions at the peripheries of cell clusters. This association seems to be dynamically regulated by signals evoked through cell-to-cell and cell-to-substrate adhesion. Shutting down signals mediating the association between ß-catenin and actinin-4 may be worth consideration as a novel therapeutic approach against cancer invasion and metastasis.


   Acknowledgments
 
Grant support: "Third Term Comprehensive Control Research for Cancer" from the Ministry of Health, Labour, and Welfare of Japan and "Medical Frontier Project (MF-1)" from the Pharmaceuticals and Medical Devices Agency of Japan. Dr. Y. Hayashida is a recipient of the research resident fellowship from the Foundation for the Promotion of Cancer Research.

The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked advertisement in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

We thank Dr. Masatoshi Takeichi for providing the E-cadherin cDNA.

Received 3/ 2/05. Revised 6/23/05. Accepted 7/22/05.


   References
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 References
 

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