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1 KAN Research Institute, Kyoto Research Park, Shimogyo-ku, Kyoto 600-8815, Japan
2 Institute of DNA Medicine, Jikei University School of Medicine, Minato-ku, Tokyo 105-8461, Japan
3 Department of Cell Biology, Faculty of Medicine, Kyoto University, Sakyo-ku, Kyoto 606-8315, Japan
4 Solution Oriented Research for Science and Technology, Japan Science and Technology Corporation, Sakyo-ku, Kyoto 606-8501, Japan
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Abstract |
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TSV)). Although both APC proteins were similarly accumulated at the MT ends, GFP-fAPC, but not GFP-APC(TSV), was associated with the basal and lateral plasma membranes and co-localized with a PDZ protein, DLG1. Stable over-expression of GFP-fAPC enforced cell–substrate attachment and thereby enhanced cell spreading on the substratum and induced polarized extension of lamellipodia and MTs during scratch-induced migration. Truncation of the PDZ-binding motif was sufficient to abolish these effects of GFP-fAPC. Furthermore, expression of GFP-APC(TSV) disturbed the establishment of a continuous epithelial monolayer. These results suggest that APC links MTs to plasma membranes through interactions with PDZ proteins, such that the migration and morphogenesis of epithelial cells can be properly regulated. |
Introduction |
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APC is a large 310-kDa protein associated with many cytoskeletal components (Fig. 1A). It binds directly to microtubules (MTs) at its COOH-terminal basic region, and stabilizes them (Munemitsu et al. 1994; Smith et al. 1994; Zumbrunn et al. 2001). In cells, APC is transported along MTs by kinesin motor protein complexes containing KAP3 and accumulates at the MT ends, specifically at the migrating edges of cells (Näthke et al. 1996; Mimori-Kiyosue et al. 2000; Jimbo et al. 2002; Kita et al. 2006). Such molecules that specifically associate with the plus ends of MTs have been classified as MT-plus-end-tracking proteins or +TIPs, together with cytoplasmic linker proteins (CLIPs), CLIP-associated proteins (CLASPs) and the dynein/dynactin complex (reviewed in Schuyler & Pellman 2001; Galjart & Perez 2003; Mimori-Kiyosue & Tsukita 2003). These molecules are now considered to provide important clues toward understanding the molecular mechanism behind the generation of the asymmetrical organization of MTs, which is critical for directional cell migration (Vasiliev 1991; Liao et al. 1995; Gundersen 2002).
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) complexes affect the localization of APC at MT ends through local inactivation of GSK-3ß (Etienne-Manneville & Hall 2003). More recently, APC was shown to be involved in neuronal axon growth in the downstream of nerve growth factor (NGF) signaling through spatial activation of PI3K and inactivation of GSK-3ß (Zhou et al. 2004). On the other hand, since APC can activate the Rac-specific guanine nucleotide exchange factor (GEF) Asef (Kawasaki et al. 2000) and affect cancer cell motility (Kawasaki et al. 2003), it is possible that APC itself may be able to regulate the actin cytoskeleton. In mammalian and Drosophila epithelial cells, APC localizes to the basolateral plasma membrane (Miyashiro et al. 1995; Näthke et al. 1996) in an actin cytoskeleton-dependent manner (Rosin-Arbesfeld et al. 2001) and ensures the maintenance of cell–cell adhesion (Hamada & Bienz 2002). Mammalian APC is associated with a PDZ protein, DLG1 (SAP97; an ortholog of the Drosophila Discs large tumor suppressor protein), via its COOH-terminal PDZ-binding motif S/TXV (Matsumine et al. 1996; Yanai et al. 2000). Since DLG1 localizes to the basolateral plasma membrane and is essential for epithelial polarity (Muller et al. 1995; Woods et al. 1996), it is worth investigating the physiological meaning of the interaction between APC and DLG. In this context, it is interesting that the association of APC and DLG1 was recently shown to be regulated by Cdc42 and Par6-PKC at the migrating edges of cells (Etienne-Manneville et al. 2005).
Although a large amount of information has been gained from APC mutants, the normal functions of APC in vertebrate cells are poorly understood. This is attributable not only to the complexity of this multifunctional protein, but also to the apparent lack of detailed analyses of APC functions using cell biology strategies, which is mainly due to the significant difficulties in the expression of exogenous full-length APC in most cultured mammalian cells. As an example, over-expression of APC in human colorectal cancer cells resulted in a substantial diminution of cell growth due to the induction of cell death through apoptosis (Morin et al. 1996). However, by taking advantage of the A6 cell line established from a normal Xenopus laevis kidney, which can over-express exogenous full-length APC, we previously observed the behavior of GFP-fused full-length APC (GFP-fAPC) in living A6 cells (Mimori-Kiyosue et al. 2000). In the present study, we established stable A6 transfectants expressing GFP-fAPC or truncated APC and characterized these cells using a series of morphological approaches. Our results suggest that PDZ domain-based interactions of APC with plasma membranes are important for proper epithelial cell motility and morphogenesis.
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Results |
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Xenopus A6 epithelial cells were transfected with the GFP-tagged APC constructs as shown in Fig. 1A. In the GFP-APC(TSV) construct, the three COOH-terminal amino acids TSV, which represent a PDZ-binding motif for binding to PDZ proteins such as DLG1 (Matsumine et al. 1996; Yanai et al. 2000), were deleted. Several independent stable clones were isolated for each construct, and three clones were used in the present study. The expressions of the fusion proteins were analyzed by Western blotting (Fig. 1B). The expression level of GFP-fAPC was estimated to be 10- to 30-fold higher than that of endogenous APC by comparison with our previous quantitative analysis (Mimori-Kiyosue et al. 2000). In the transfected cells, the amount of endogenous ß-catenin was not affected by expression of either full-length or truncated APC (Fig. 1B), indicating that over-expression of APC in A6 cells does not affect ß-catenin signaling though modulation of ß-catenin stability, unlike cancer cell lines. All clones expressing the same construct behaved similarly in the assays described below, and statistical data were obtained by averaging the results from independent clones, unless stated otherwise.
Localization of APC proteins to the basal plasma membrane
At low cell densities, GFP-fAPC and GFP-APC(TSV) were similarly accumulated in a subset of MT ends in cell protrusions (Fig. 2A). When the MTs were depolymerized, however, GFP-fAPC showed a strikingly distinct distribution from that of GFP-APC(TSV). Without MTs, GFP-fAPC was gradually relocated from the cell edges toward the cell center, and finally became distributed along the basal plasma membrane in a strip complementary with actin stress fibers, as described previously (Mimori-Kiyosue et al. 2000). In these cells, DLG1 was clustered together with GFP-fAPC in a stripe-like pattern (Fig. 2B; see also Supplementary Fig. S1D). Endogenous APC showed a similar distribution to GFP-fAPC (data not shown). In contrast, GFP-APC(TSV) was not distributed along the basal plasma membrane with DLG1, but instead was scattered in the cytoplasm (Fig. 2B). This result clearly indicates that APC associates with the basal plasma membrane via its PDZ-binding motif. In some cells, co-localization of GFP-fAPC with DLG1 was detectable at the MT ends (Fig. 2C).
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TSV) by total internal reflection fluorescence (TIRF) microscopy, which only illuminates the area in close proximity to the coverslip (
200 nm) (Steyer & Almers 2001) (Fig. 2D). GFP-DLG1 was detectable all over the basal plasma membrane as diffuse small granular structures. The signals for GFP-fAPC covered the entire basal plasma membrane, in addition to showing strong accumulation at the MT ends. In contrast, GFP-APC(TSV) was selectively localized along MTs. These observations suggest that GFP-fAPC is predominantly associated with the basal plasma membrane via a PDZ protein such as DLG1, and is brought to the edges of the cells by kinesin-based transport along MTs. GFP-fAPC stabilizes cellular extensions
The distributions of GFP-fAPC and GFP-APC(TSV) at the MT ends appeared almost identical at first glance. However, observation of their dynamic behaviors by time-lapse imaging revealed that the dynamic properties of the GFP clusters and cell motilities differed somewhat in the cells. In many cases, GFP-fAPC accumulated at two or more sites within a cell (for example, clusters a and b in Fig. 3A; see also Supplementary Video S1). Once GFP-fAPC-positive cell protrusions had grown and elongated, they were stable for several hours. Sometimes the protrusions retracted, while retaining the APC cluster (cluster b in Fig. 3A), and the cell moved in a different direction. After retraction, the original APC cluster (b) became diffuse and was incorporated into a new cluster formed at another site. However, GFP-APC(TSV) clusters were less stable than GFP-fAPC-clusters. Even though GFP-APC(TSV) formed clusters at the cell periphery, the clusters showed repeated outgrowth and retraction over a short time and distance (Fig. 3B,D). These differences in the dynamic properties can be visualized in Supplementary Video S1.
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TSV)-positive extensions, and the retracting population was increased in GFP-APC(TSV)-expressing cells. All these observations suggest that the PDZ-binding ability of APC is important for stabilizing cellular extensions, possibly by attaching MT plus ends to specified regions of the basal plasma membrane. GFP-fAPC enhances polarized cellular extension during wound healing
The migration activities of the cells were further examined using scratch-induced migration assays. Highly polarized quiescent cell layers were scratched to induce directional migration. Shortly after wounding, we noticed that GFP-fAPC-expressing cells extended significantly larger lamellipodia into the wound than parental A6 cells, while the cells expressing GFP-APC(TSV) were retarded (Fig. 4A). When the cell extension lengths were measured on time-lapse images at 60, 120 and 240 min after scratching, it was clear that GFP-fAPC-expressing cells quickly increased the length of their lamellipodia (Fig. 4C). We also noticed that the MTs in migrating GFP-fAPC-expressing cells were largely elongated toward the wound in a polarized fashion (Fig. 4A, middle). At the leading edges of the cells, GFP-fAPC was clustered at the MT ends facing the wounded region (Fig. 4B). Although GFP-APC(TSV) was localized at the MT ends similarly to GFP-fAPC (see Fig. 2A), GFP-APC(TSV) clusters did not migrate efficiently into the wounded region, consistent with a previous observation in primary rat astrocytes expressing an NH2-teminal fragment of APC (Etienne-Manneville & Hall 2003).
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The rapid cellular extension of GFP-fAPC-expressing cells into a wound suggested a role for GFP-fAPC in cell spreading. To examine this possibility, we performed cell spreading assays. Cells seeded on coverslips were fixed and stained with TRITC-phalloidin to visualize their shape changes over time. As shown in Fig. 4D, GFP-fAPC-expressing cells began to spread at earlier time points. The cell areas were measured and plotted (Fig. 4E). The results clearly revealed that GFP-fAPC-expressing cells increased in area significantly faster than the other cells. However, after 24 h, the average size of GFP-APC(TSV)-expressing cells increased to become the same as that of GFP-fAPC-expressing cells (Fig. 4E). These results indicate that GFP-fAPC accelerates cell spreading effectively from an early stage soon after cell attachment to the substratum. Since APC has been reported to stimulate cell migration by activating the Rac-specific GEF Asef (Kawasaki et al. 2000, 2003; Watanabe et al. 2004), our results suggest that APC facilitates Rac signal transduction through associations with plasma membranes at regions where Asef and Rac are distributed.
Abnormal epithelial morphology of GFP-APC(TSV)-expressing cells
The expression of exogenous APC also affected the epithelial cell morphology. In confluent cultures (1–1.3 x 105 cells/cm2), parental A6 and GFP-fAPC-expressing cells showed cobblestone-like epithelial morphologies, whereas GFP-APC(TSV)-expressing cells were somewhat fibroblastic in appearance (Fig. 5A). These morphological differences became more pronounced as the cell density increased (4–5 x 105 cells/cm2). Specifically, parental A6 and GFP-fAPC-expressing cells established a continuous monolayer, whereas the layer of GFP-APC(TSV)-expressing cells was irregular and frequently protruded in patches (Fig. 5B). Scanning electron microscopic observation of the cell surfaces revealed that groups of GFP-APC(TSV)-expressing cells protruded from the layer, although they retained contact (Fig. 5C). In addition, GFP-fAPC-expressing cells were consistently larger in the planar dimension than the other cells, even after prolonged culture.
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TSV) expression (Fig. 5E,F). Therefore, expression of GFP-APC(TSV) disturbs the maintenance of the epithelial monolayer without affecting the assembly of cell–cell junctions. Furthermore, no nuclear accumulation of ß-catenin was observed under any of the conditions examined (Fig. 5E, F and data not shown), suggesting that all the morphological changes were independent of ß-catenin-induced transcription in the nucleus.
Ultrastructures of the cell layers
The structures of the cell layers were further examined by thin-section transmission electron microscopy. Parental A6 and GFP-fAPC-expressing cells were both organized into monolayers, although the monolayer of GFP-fAPC-expressing cells was significantly thinner (Fig. 6A,B). Together with the results of the scanning electron microscopy and immunofluorescence analyses (Fig. 5), these observations indicate that GFP-fAPC-expressing cells have a flattened morphology in the polarized cell layer.
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TSV) tended to pile up with increased intercellular space (Fig. 6C), and multiple layers developed in some areas (Fig. 6D). Magnified images of the cell–cell adhesion sites are shown in Fig. 6E–G. Similar to the parental A6 cells, both GFP-fAPC- and GFP-APC(TSV)-expressing cells retained normal junction structures near the apical cell surface (arrowheads).
Interestingly, when the substrate attachment sites were observed at higher magnification (Fig. 6H–J), the basal plasma membranes of GFP-fAPC-expressing cells were repeatedly found to be in close association with the substratum over a wider region than those of parental A6 cells (Fig. 6H,I), while only small regions of the membranes of GFP-APC(TSV)-expressing cells were attached to the substratum (Fig. 6J).
Cell–cell and cell–substrate attachments of the transfectants
The above-described morphological observations suggested that the expression of full-length or truncated APC could affect the mode of cell–cell and/or cell–substrate attachment of the cells. Therefore, we analyzed the nature of the cell–cell adhesion using cell dissociation assays. Polarized cell layers were treated with trypsin in the presence of calcium (TC treatment), which specifically disrupted only calcium-independent cadherin-based adhesion structures, followed by pipetting to mechanically dissociate the bonds (Fig. 7A). To statistically analyze the sizes of the cell clusters remaining after the pipetting, the numbers of clusters were divided by the numbers of particles remaining after trypsin-EGTA treatment (TE treatment), which disrupts all cell–cell adhesion structures (Fig. 7B). After pipetting, the layers of GFP-fAPC-expressing cells were easily broken up.
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TSV)-expressing cells were detached from the substratum as aggregates after pipetting (Fig. 7C,D). In contrast, GFP-fAPC-expressing cells remained attached to the substratum, although cell–cell attachments were lost, indicating that these cells were more strongly attached to the substratum than the other cell lines, while the cell–cell adhesion was weakened. These results seemed to be inconsistent with the electron microscopic observations shown in Fig. 6B,F, in which the intercellular space of GFP-fAPC-expressing cells was reduced due to their close contact. However, in these pictures, no obvious cell–cell adhesion structures visible as dense signals connecting neighboring cells (arrowheads in Fig. 6E–G) were observed, indicating that the alterations to the membrane architecture induced by GFP-fAPC expression did not strengthen cell–cell adhesion.
These results all indicate that, when the function of full-length APC was abolished by the loss of PDZ protein-binding ability, the truncated product caused abnormal effects on the establishment of an epithelial monolayer. Thus, the association of APC with plasma membranes through the PDZ protein is important for proper regulation of epithelial migration and morphogenesis.
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Discussion |
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Previously, to visualize the dynamic behavior of APC in living cells, we used a fAPC-GFP construct, in which GFP was fused to the COOH-terminus of APC (Mimori-Kiyosue et al. 2000). However, APC has a PDZ-binding motif at its extreme COOH-terminus, and thus fusion of the COOH-terminal with GFP may inhibit the interactions of APC with PDZ proteins. In fact, we observed that fAPC-GFP did not become distributed to plasma membranes (our unpublished data). In the present study, we generated an NH2-terminal-tagged construct, GFP-fAPC, by fusing the molecules via a longer spacer sequence. GFP-fAPC appeared to be functional, and became distributed at the ends of MTs and in the basolateral plasma membrane, similar to endogenous APC. As expected from our earlier observations, truncation of the PDZ-binding motif from GFP-fAPC (GFP-APC(TSV)) abolished the association of this molecule with plasma membranes (Fig. 2B, Supplementary Fig. S1D), indicating that interactions with PDZ proteins are important for tethering APC to plasma membranes.
However, previous reports have shown that the armadillo repeat domain of APC alone is sufficient for APC association with plasma membranes (Barth et al. 2002; Reilein & Nelson 2005). Consistent with these reports, we observed that NH2-terminal fragments of APC (e.g. nAPC-GFP shown in Supplementary Fig. S1D) were localized at the basal plasma membrane and in cell–cell adhesion sites in A6 cells (data not shown, Supplementary Fig. S1D). These observations indicate that GFP-APC(TSV) is regulated differently from nAPC-GFP that lacks the COOH-terminal region. Recently, the MT-binding region of APC was reported to interact with the armadillo repeats and inhibit the binding of KAP3 to this domain (Li & Näthke 2005). Therefore, it is possible that the COOH-terminal region regulates the ability of the armadillo repeats to associate with plasma membranes by changing the protein interactions. This hypothesis can explain our present results, as well as a previous observation that membrane-associated DLG1 interacts with MT-bound APC to attach MTs to the migrating edges of astrocytes in the downstream of Cdc42 and Par6-PKC (Etienne-Manneville et al. 2005).
Physiological meaning of the interaction of APC with PDZ proteins
At the basolateral plasma membrane, GFP-fAPC co-localized with a known binding partner, DLG1 (Matsumine et al. 1996). In addition, TIRF microscopy revealed that a portion of the GFP-fAPC was also distributed over the entire basal plasma membrane in the presence of MTs (Fig. 2D), indicating that GFP-fAPC molecules are associated with the plasma membrane and that some of them are concentrated at the edges of cells depending on active transport along MTs. Therefore, DLG1 may provide important clues for clarifying the regulation of cell functions by linking MT plus ends to the cell cortex. Recent studies have demonstrated that not only APC but also other MT-anchoring factors, such as ACF7 and CLASP, also largely contribute to the regulation of directional cell migration (Kodama et al. 2003; Mimori-Kiyosue et al. 2005; Lansbergen et al. 2006).
Next, the question arises as to what happens after MT-cortex attachment via APC. In our wound healing and cell spreading assays (Fig. 4), we observed that GFP-fAPC effectively accelerated cell spreading. Furthermore, GFP-fAPC-expressing cells showed a flattened morphology, both in low-density cultures and confluent monolayers (Figs 5 and 6). These phenomena can be explained by the fact that GFP-fAPC expression enhanced cell–substrate adhesion, rather than cell–cell attachment (Fig. 7). It is likely that the Rac-specific GEF Asef may contribute by activating the signaling cascade downstream of Rac at plasma membranes, since Asef was found to activate the migration of cancer cells expressing only truncated APC (Kawasaki et al. 2003). Under normal conditions, APC could activate Rac signaling through Asef in a spatiotemporally controlled manner, and DLG1 may help the association of APC with Asef by linking APC to plasma membranes in the downstream of signaling molecules such as Par6-PKC-Cdc42 complexes or NGF (Zhou et al. 2004; Etienne-Manneville et al. 2005). Furthermore, the MTs tethered at the cortex may help the APC-induced process, perhaps by facilitating the supply of factors required for cell spreading and cell–substratum adhesion.
However, since APC also associates with PDZ proteins other than DLG1, such as the protein tyrosine phosphatase PTP-BL (Erdmann et al. 2000), its relationships with other PDZ proteins need to be taken into account in order to fully understand the phenomena observed in A6 transfectants. An additional interesting point is that no direct relationship between APC and DLG1 in Drosophila has been described to date. Drosophila and vertebrate APC proteins share a common trait in their capacity to down-regulate the expression of ß-catenin and associate with MTs and basolateral plasma membranes, and thus the functions of mammalian APC in cell migration and morphogenesis have been estimated from studies in Drosophila. However, neither of the two Drosophila APC proteins carries a recognizable PDZ-binding motif. It is therefore necessary to clarify how APC proteins with distinct molecular architectures act differently and/or similarly in diverse cellular systems.
Functions of APC in epithelial morphogenesis
In contrast to the morphological transformation of cells following the expression of oncogenic Ras or Rho family proteins, in which the cells adopt a mesenchymal phenotype and lose E-cadherin-mediated cell–cell adhesion (reviewed in Malliri & Collard 2003), the truncated APC proteins did not inhibit cell–cell junction formation even after piling up of the cells. Interestingly, these phenotypes appear to bear some analogy to the early stages of polyp formation in the intestine of APC mutant mice (Oshima et al. 1997), as well as human sporadic colorectal adenomas (Shih et al. 2001). When both alleles of the APC gene were mutated, the cells began to develop a dysplastic epithelial tissue architecture, presenting as polyps. However, during the course of the polyp development, the cell–cell interactions remained intact, and these events preceded a transition to malignancy due to additional gene mutations (Oshima et al. 1997). Although accumulation of ß-catenin may be required for activation of Tcf/Lef response genes in the nucleus with malignant transition of APC mutated cells (Fodde et al. 2001), APC may have other tumor suppressor functions besides the degradation of ß-catenin (Kawasaki et al. 2003). For example, it has been reported that small adenomas with ß-catenin mutations are less likely to develop into larger adenomas and invasive carcinomas than adenomas with APC mutations, suggesting that APC and ß-catenin mutations are not functionally equivalent (Samowitz et al. 1999). Furthermore, loss of both APC alleles is rarely observed, and most tumors retain at least one allele that produces a roughly half-sized APC protein (Rowan et al. 2000; reviewed in Bienz 2002).
The results of the present study support the notion that loss of full-length APC or the presence of a truncated APC protein could affect the cell morphology during polyp formation and malignant transition of APC mutated cells. Therefore, it will be interesting to explore further factors and molecular mechanisms that regulate epithelial cell morphogenesis together with APC.
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Experimental procedures |
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The following primary antibodies were used: rabbit anti-GFP polyclonal antibody (pAb) (Chemicon); chicken anti-GFP pAb (Chemicon); rabbit anti-ß-catenin pAb (Sigma); mouse anti-
-tubulin monoclonal antibody (mAb) (DM1A; Sigma); mouse anti-PSD-95 family (anti-PDZ domain, Upstate Biotechnology; referred to as anti-DLG antibody in the text); anti-ZO-1 mAb (Sanko Junyaku, Japan); and rabbit anti-APC pAb (kindly provided by Dr Akiyama (Jimbo et al. 2002)). For more details regarding the anti-DLG antibody, please refer to the Supplementary Materials and methods. Tetramethyl rhodamine isothiocyanate (TRITC)-phalloidin and Alexa Fluor-594-phalloidin were purchased from Sigma and Molecular Probes, respectively. The following secondary antibodies were purchased from Jackson Immunoresearch Laboratories: Cy2-conjugated anti-mouse IgG and anti-rabbit IgG pAbs; Texas Red-conjugated anti-mouse IgG and anti-rabbit IgG pAbs; and Cy5-conjugated anti-mouse IgG and anti-rabbit IgG pAbs.
To construct the GFP-fAPC expression vector, partial fragments of APC cDNAs were cut out from pGFP-C(NheI)/APC(1-8490) or pQBI25/APC(1-8487) (Mimori-Kiyosue et al. 2000) and inserted into the SacII and XbaI sites of pEGFP-C1 to generate fAPC/pEGFP-C1. As a result, APC was fused to GFP through 24 unrelated amino acids. To construct GFP-APC(TSV), the COOH-terminal region of fAPC in pEGFP-C1 was substituted with a PCR-engineered COOH-terminal fragment lacking the coding sequence for the final three COOH-terminal amino acids (TSV). To construct GFP-DLG1/SAP97, the rat SAP97 gene inserted in pCIneo (kindly provided by Dr Y. Takai, Osaka University) was cut out with EcoRI and SalI and inserted into the EcoRI and SalI sites of pEGFP-C2 (Clontech). The Effectene reagent (Qiagen) was used for plasmid transfections, according to the manufacturer's directions. Drug-resistant clones were selected in the presence of 0.6–0.8 mg/mL G418 (Calbiochem) and screened by GFP detection under a fluorescence microscope.
Cell culture and assays
The A6 cell line was established from a normal X. laevis kidney. In culture, these cells form typical polarized epithelial monolayers, and have apical microvilli, a primary cilium and tight junctions (Perkins & Handler 1981). The A6 cells were grown at 23 °C without CO2 in Leibovitz's L-15 medium (50% L-15 medium, 40% distilled water, 10% fetal bovine serum, penicillin–streptomycin solution), routinely subcultured every 4–5 days at about 90% confluency and used for assays on day 3 after a passage. Stable transfectants were used for assays within 20 passages. To allow the cells to polarize and become quiescent, confluent cultures (1 x 105 cells/cm2) were continuously grown for 5–6 days (4–5 x 105 cells/cm2) by exchanging the medium daily.
Cell dissociation assays were performed as described previously (Nagafuchi et al. 1994), with some modifications, using 50% HCMF buffer (100% HEPES-buffered saline: 10 mM HEPES–NaOH, pH 7.4, 137 mM NaCl, 5.4 mM KCl, 0.3 mM Na2HPO4·7H2O, 5.5 mM glucose) to optimize the conditions for A6 cells. Confluent cells (4 x 105 cells/cm2) were treated with 0.02% trypsin in buffer supplemented with 1 mM CaCl2 (TC treatment) or 1 mM EGTA (TE treatment) for 15 min at room temperature, and then dissociated by pipetting 10 times. The extent of cell dissociation was represented by NTC/NTE, where NTC and NTE are the total numbers of particles after the TC and TE treatments, respectively. For low-calcium buffer (LC) treatment, the concentration of free Ca2+ was adjusted to 20.5 µM by adding 50 µM CaCl2 and 30 µM EGTA.
For scratch-induced cell migration assays, polarized quiescent cell layers were scratched with pipette tips and either subjected to time-lapse microscopy as soon as possible or fixed and stained over time.
For cell spreading assays, dissociated cells were seeded on coverslips, and then fixed and stained with TRITC-phalloidin after 15, 30, 60 and 120 min and 24 h. To observe live cells, cells were cultured on glass-bottomed dishes containing No. 1S coverslips (Iwaki) in culture medium without phenol red. To disassemble MTs, the cells were incubated with 33 µM nocodazole (Sigma) at 23 °C.
Western blotting
To analyze the expressions of APC proteins, cell lysates from various stable transfectants were separated by SDS-PAGE (2 x 105 cells/lane) using 4%–7.5% acrylamide gradient gels, and immunoblotted. For ß-catenin, lysates were separated in 7.5% acrylamide gels (4 x 104 cells/lane). Following incubation with appropriate antibodies, the bound antibodies were detected with BCIP/NBT solution (WAKO) or the ECL plus Western Blotting Detection System (Amersham).
Immunofluorescence staining
Cells cultured on coverslips were fixed in either 3.7% formaldehyde for 15–30 min or methanol at –20 °C for 2 min. After washing with PBS(-), cells were permeabilized with 0.1% or 0.5% Triton X-100 and blocked with 10% fetal bovine serum. The samples were then processed for indirect immunostaining using appropriate primary and secondary antibodies, washed several times and mounted using a Prolong Anti-fade Kit (Molecular Probes).
Fluorescence microscopy and image analysis
Images of cells were acquired with an LSM510 laser scanning confocal microscope (Ver. 2.3; Carl Zeiss) or a DeltaVision optical sectioning microscope (Ver. 2.50; Applied Precision Inc.), equipped with Axiovert (Plan Apochromat 63 x /1.40 NA or Plan Apochromat 100 x /1.40 NA oil immersion) or Olympus IX70 (UPlanApo 20 x /0.70 NA dry, PlanApo 60 x /1.40 NA oil or PlanApo 100 x /1.40 NA oil immersion) objectives, respectively. Using the DeltaVision system, images were acquired through a cooled CCD camera (Series 300 CH350; Photometrics) with appropriate ND filters, binning of pixels, exposure times and time intervals. Fluorescent signals were visualized using an Endow GFP Bandpass Emission Filter Set (41 017; Chroma) or a Sedat Quad Filter Set (86 000; Chroma). TIRF microscopy was performed as described previously (Mimori-Kiyosue et al. 2005).
Pixel positions, distances and areas were measured on the digital images using the analysis function of the DeltaVision Aquacosmos software (Hamamatsu Photonics) or MetaMorph software (Universal Imaging). Cell areas were quantified using the particle analysis function of the Aquacosmos software. The total cell area visualized with TRITC-phalloidin staining was obtained for each image, and then divided by the cell number. Statistical analyses were performed using KyPlot (Version 3.0; Kyence Inc.) or Microsoft Excel 2000 software. The statistical significance of observed differences was evaluated using the Mann–Whitney U test (P < 0.05). In all plots, the standard error (SE) is indicated.
Electron microscopy
For transmission electron microscopy, cells seeded on plastic dishes were doubly fixed with 1.2% glutaraldehyde in 0.1 M cacodylate buffer and 1% osmium tetroxide in 0.1 M cacodylate buffer, dehydrated through a graded ethanol series, and embedded in epoxy resin together with the plastic dishes to preserve cell–substrate attachments. Ultrathin sections were stained with uranyl acetate and lead citrate, and observed with an H-7500 transmission electron microscope (Hitachi).
For scanning electron microscopy, cells were doubly fixed with 1% glutaraldehyde in 0.1 M phosphate buffer and 1% osmium tetroxide in 0.1 M phosphate buffer, dehydrated through a graded ethanol series, critical point-dried, sputter-coated with gold, and observed with a JSM-5800 LV scanning electron microscope (JEOL).
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Acknowledgements |
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Footnotes |
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* Correspondence: E-mail: y-kiyosue{at}kan.gr.jp
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References |
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Received: 5 October 2006
Accepted: 8 November 2006
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