Actin-dependent membrane association of the APC tumour suppressor in polarized mammalian epithelial cells
2001; Springer Nature; Volume: 20; Issue: 21 Linguagem: Inglês
10.1093/emboj/20.21.5929
ISSN1460-2075
Autores Tópico(s)Genetic factors in colorectal cancer
ResumoArticle1 November 2001free access Actin-dependent membrane association of the APC tumour suppressor in polarized mammalian epithelial cells Rina Rosin-Arbesfeld Rina Rosin-Arbesfeld MRC Laboratory of Molecular Biology, Hills Road, Cambridge, CB2 2QH UK Search for more papers by this author Gudrun Ihrke Gudrun Ihrke The Wellcome Trust Centre for Molecular Mechanisms in Disease and Department of Clinical Biochemistry, University of Cambridge, Cambridge, CB2 2XY UK Search for more papers by this author Mariann Bienz Corresponding Author Mariann Bienz MRC Laboratory of Molecular Biology, Hills Road, Cambridge, CB2 2QH UK Search for more papers by this author Rina Rosin-Arbesfeld Rina Rosin-Arbesfeld MRC Laboratory of Molecular Biology, Hills Road, Cambridge, CB2 2QH UK Search for more papers by this author Gudrun Ihrke Gudrun Ihrke The Wellcome Trust Centre for Molecular Mechanisms in Disease and Department of Clinical Biochemistry, University of Cambridge, Cambridge, CB2 2XY UK Search for more papers by this author Mariann Bienz Corresponding Author Mariann Bienz MRC Laboratory of Molecular Biology, Hills Road, Cambridge, CB2 2QH UK Search for more papers by this author Author Information Rina Rosin-Arbesfeld1, Gudrun Ihrke2 and Mariann Bienz 1 1MRC Laboratory of Molecular Biology, Hills Road, Cambridge, CB2 2QH UK 2The Wellcome Trust Centre for Molecular Mechanisms in Disease and Department of Clinical Biochemistry, University of Cambridge, Cambridge, CB2 2XY UK *Corresponding author. E-mail: [email protected] The EMBO Journal (2001)20:5929-5939https://doi.org/10.1093/emboj/20.21.5929 PDFDownload PDF of article text and main figures. ToolsAdd to favoritesDownload CitationsTrack CitationsPermissions ShareFacebookTwitterLinked InMendeleyWechatReddit Figures & Info Adenomatous polyposis coli (APC) is mutated in most colorectal cancers. APC downregulates nuclear β-catenin, which is thought to be critical for its tumour suppressor function. However, APC may have additional and separate functions at the cell periphery. Here, we examine polarized MDCK and WIF-B hepatoma cells and find that APC is associated with their lateral plasma membranes. This depends on the actin cytoskeleton but not on microtubules, and drug wash-out experiments suggest that APC is delivered continuously to the plasma membrane by a dynamic actin-dependent process. In polarized MDCK cells, APC also clusters at microtubule tips in their basal-most regions. Microtubule depolymerization causes APC to relocalize from these tips to the plasma membrane, indicating two distinct peripheral APC pools that are in equilibrium with each other in these cells. Truncations of APC such as those found in APC mutant cancer cells can neither associate with the plasma membrane nor with microtubule tips. The ability of APC to reach the cell periphery may thus contribute to its tumour suppressor function in the intestinal epithelium. Introduction Mutations in the adenomatous polyposis coli (APC) tumour suppressor gene are common in tumours of the human colorectum, and APC loss is an early, if not initiating, event during tumorigenesis in this tissue (Kinzler and Vogelstein, 1996). In the mouse, carriers of heterozygous Apc mutations inevitably develop multiple polyps in the intestinal tract, each of which shows inactivation of both Apc alleles (Su et al., 1992; Oshima et al., 1995, 1997). Interestingly, the earliest detectable consequence of Apc loss in the intestinal epithelium is an abnormal tissue architecture rather than an increase in cell proliferation (Oshima et al., 1995, 1997). Early adenoma cells appear to lag behind in the proliferative crypt stem cell compartment rather than leaving it at the normal rate to become differentiated, which may reflect a mis-specification of cell fate (Korinek et al., 1998), or a failure in cell migration or adhesion (Näthke et al., 1996; Barth et al., 1997; Pollack et al., 1997; reviewed by Bienz and Clevers, 2000). APC proteins promote the destabilization of cytoplasmic β-catenin, an effector of the Wnt pathway, by binding to the Axin complex. This complex earmarks β-catenin for degradation by the ubiquitin pathway (Polakis, 2000). Furthermore, APC harbours a nuclear export function that is deleted by most APC colorectal tumour mutations (Rosin-Arbesfeld et al., 2000). As a consequence, colorectal cancer cells show high levels of nuclear β-catenin that, together with the transcription factor TCF (T cell factor), activate the transcription of Wnt target genes (Korinek et al., 1997). The ability of APC to exit from the nucleus and to bind to the cytoplasmic Axin complex is thought to be critical for its tumour suppressor function (Smits et al., 1999; Rosin-Arbesfeld et al., 2000; von Kries et al., 2000). APC proteins are also concentrated at peripheral subcellular sites. Most strikingly, APC accumulates at the distal tips of microtubules in cellular protrusions of motile cells (Näthke et al., 1996). This clustering of APC at microtubule tips results from the ability of APC to track along microtubules (Mimori-Kiyosue et al., 2000). Evidence suggests that these APC clusters may be functionally relevant for cellular migration and adhesion (Barth et al., 1997; Pollack et al., 1997). Furthermore, in mitotic cells, the association of APC with microtubule plus ends appears to affect the fidelity of chromosomal segregation (Fodde et al., 2001; Kaplan et al., 2001). The Drosophila homologue E-APC/dAPC2 is associated with the apicolateral adherens junctions of embryonic and larval epithelia. The junctional association of E-APC is critical for its function to destabilize Armadillo, the Drosophila homologue of β-catenin (McCartney et al., 1999; Yu et al., 1999). This association may thus reflect E-APC's binding to the Axin complex which appears to be localized underneath the plasma membrane (Zeng et al., 1997; Bienz, 1999; Fagotto et al., 1999). It could also reflect a separate function of E-APC in the maintenance of junctional integrity (Townsley and Bienz, 2000). Little is known about the subcellular distribution of the human APC tumour suppressor in polarized epithelial cells. We chose two polarized cell models to examine this: Madin–Darby canine kidney (MDCK) cells, a widely used cell line that readily polarizes under appropriate culture conditions (Simons and Fuller, 1985; Rodriguez-Boulan and Nelson, 1989), and WIF-B cells, a hybrid cell line derived from rat hepatoma cells and human fibroblasts that exhibits a hepatic morphology and attains a high degree of polarization upon prolonged culture (Ihrke et al., 1993; Decaens et al., 1996). We found that endogenous APC is associated with the lateral plasma membrane of both polarized cell types. This association depends on the actin cytoskeleton but not on microtubules. In addition, we also observed microtubule-dependent clusters of APC in the basal-most regions of polarized MDCK cells. Our evidence indicates that these cells contain two distinct peripheral pools of APC that are in equilibrium with each other: actin-dependent membrane-associated APC and microtubule-dependent APC clusters. Finally, we show that truncations of APC such as those typically found in APC-mutant cancer cells have lost the ability to reach either of these peripheral locations. Results Two distinct peripheral pools of APC in MDCK cells To examine the subcellular distribution of endogenous APC in MDCK cells, we stained subconfluent cells with an antibody raised against a central fragment of human APC (Näthke et al., 1996). Subconfluent cells show some cytoplasmic staining, characteristically grainy. This graininess is not due to the secondary antibody (not shown), and is absent from APC-mutant cancer cells that do not express detectable APC protein (see Supplementary data available at The EMBO Journal Online). Grainy staining is also observed in other mammalian cells (see below), as well as in Drosophila cells stained for E-APC (Yu et al., 1999). In addition to the grainy cytoplasmic staining, subconfluent MDCK cells show numerous bright clusters of APC associated with microtubule tips at the cell periphery (Näthke et al., 1996) (Figure 1A, arrows). However, these clusters not only seemed to coincide with microtubule tips, but also with phalloidin staining (Figure 1A), an observation that was confirmed in COS cells transfected with APC tagged with green fluorescent protein (GFP): these large spread-out cells allowed direct observation of microtubule-associated APC–GFP clusters coinciding with tips of actin filaments (not shown). We thus re-examined the dependence of the peripheral APC clusters on the cytoskeleton, using depolymerizing drugs. Figure 1.Two distinct peripheral locations of APC in subconfluent MDCK cells. Confocal sections through subconfluent MDCK cells after paraformaldehyde fixation, co-stained as indicated above the panels, drug-treated as indicated on the right (D, recovery after latrunculin A treatment). (A) Basal-most plane, providing optimal visualization of microtubule tip clusters (arrows); these disappear after treatment with microtubule- (B) or actin-depolymerizing drugs (C), but are restored after drug removal (D, arrow; not shown). Arrowheads point to membrane-associated APC staining (optimal visualization ∼2.5 μm above basal-most plane, B–D) which disappears after actin depolymerization (C), reappears 15 min after drug removal (D), and which is considerably increased after microtubule depolymerization (B). Scale bar, 10 μm (in this and all subsequent figures). Download figure Download PowerPoint As previously shown (Näthke et al., 1996), the peripheral APC clusters disappeared completely after treatment with nocodazole (Figure 1B). This drug caused thorough depolymerization of microtubules (not shown), but had no major effects on phalloidin or E-cadherin staining. Interestingly, in cell cultures that were nearly confluent, nocodazole treatment also resulted in a striking association of APC with the plasma membrane (Figure 1B, arrowheads). Some membrane-associated APC was also observed in untreated cells, especially at plasma membrane stretches that stained strongly for E-cadherin (Figure 1A, arrowhead). Nevertheless, the microtubule depolymerization caused a considerable redistribution of APC from microtubule tips to the plasma membrane. It seems that this alternative peripheral destination of APC revealed by the drug experiment is normally overshadowed by a preference of APC to cluster at microtubule tips in these cells. Treatment of MDCK cells with cytochalasin D did not significantly affect the peripheral APC clusters (Näthke et al., 1996 and data not shown). We thus applied latrunculin A, whose effect on actin depolymerization is more potent than that of cytochalasin D (Ayscough, 1998). This revealed a considerable effect on the APC clusters, most of which disappeared after latrunculin A treatment (Figure 1C). Also, the membrane-associated APC staining was no longer detectable, paralleling the disappearance of E-cadherin staining from the plasma membrane (Figure 1C). Concomitantly, the levels of cytoplasmic APC were increased in the drug-treated cells (Figure 1C, compare with A). We asked whether these effects of latrunculin A were reversible. We thus conducted drug wash-out experiments, replacing the drug solution with fresh medium and fixing cells at various intervals of recovery. This revealed that APC began to re-associate with the plasma membrane after several minutes, along with actin repolymerization and re-appearance of membrane-associated E-cadherin staining. APC staining along the plasma membrane became maximal after 15 min of recovery, and tended to be more pronounced than in cells that had not been treated with the drug (Figure 1D, arrowheads, compare with A). Microtubule tip clusters of APC also recovered, albeit at a slower rate (Figure 1D, arrow). We conclude that there are two distinct peripheral locations of APC in MDCK cells: membrane-associated APC and microtubule tip clusters of APC. The former depends on the actin cytoskeleton but not on microtubules. The clusters depend on microtubules and only partly on the actin cytoskeleton. This partial dependence indicates that the actin cytoskeleton may contribute less directly than microtubules to the formation of the peripheral APC clusters. Finally, the drug experiments indicate that APC can shift from one peripheral location to the other. Membrane-associated APC and microtubule-dependent APC clusters in polarized MDCK cells MDCK cells can be induced to polarize and form an epithelial-like monolayer when grown on micropore filters (Simons and Fuller, 1985; Rodriguez-Boulan and Nelson, 1989). We thus examined polarized MDCK cells to see whether the process of polarization affected the subcellular distribution of APC. We grew MDCK cells on filters for 4–6 days (see Materials and methods) after which time they formed a tight monolayer, with a height of cells ranging from 8–15 μm. In these monolayer cells, E-cadherin staining was predominantly in the basolateral plasma membranes, with very little staining along the apical plasma membrane (Figure 2A–C), as previously shown (Le Bivic et al., 1990; Shore and Nelson, 1991), indicating their polarization. Polarized cells showed grainy cytoplasmic staining of APC, as well as some APC staining along the lateral plasma membranes (Figure 2A, arrowheads). This membrane-associated staining was very prominent in the basal-most zones of the lateral plasma membranes (Figure 2B, arrowheads). In addition, these confocal sections through the basal zones of cells showed numerous clusters of APC staining, typically in the vertices of cells (Figure 2B and C, arrows). These basal clusters seemed to coincide with the tips of actin stress fibres, but we did not observe any significant co-localization with the focal adhesion protein vinculin (not shown). There was no detectable concentration of APC along the apical plasma membrane. Figure 2.Peripheral locations of APC in polarized MDCK cells. Confocal sections through polarized MDCK cells after paraformaldehyde fixation, co-stained as indicated above the panels, drug-treated as indicated on the right (E, recovery after latrunculin A treatment); (A, D–F) ∼8 μm above the basal-most plane; (B and G) basal-most plane, showing actin stress fibres and basal APC clusters (arrows) which disappear after microtubule depolymerization; (C) z-section along the line indicated in (A), revealing E-cadherin staining predominantly along the basolateral plasma membrane (arrow indicates basal APC cluster). Association of APC with lateral plasma membranes (A, B, arrowheads) disappears after actin depolymerization (D), reappears after drug removal (E), and increases somewhat after microtubule depolymerization (F, G). Download figure Download PowerPoint We exposed polarized MDCK cells to latrunculin A to determine whether the membrane association of APC requires the actin cytoskeleton. This was the case, although the drug-treated cells still showed phalloidin staining in the cellular cortex, presumably reflecting residual cortical actin filaments. Also there was no longer any APC staining associated with the lateral plasma membrane and the levels of cytoplasmic APC staining seemed to have increased (Figure 2D). This drug treatment also reduced the levels of E-cadherin staining of the lateral plasma membranes (Figure 2D). However, after washing out the drug, the membrane-associated APC staining recovered in a large fraction of the cells (Figure 2E, arrowheads). At the same time, the cytoplasmic APC levels became reduced, and recovery of membrane-associated E-cadherin staining was observed (Figure 2E). This demonstrated that the drug-induced dissociation of APC from the plasma membrane is reversible. We noticed that the numbers of basal APC clusters were also significantly reduced in latrunculin-treated cells; complete restoration of these clusters was slow, requiring at least 16 h post drug removal (not shown). Exposure of polarized MDCK cells to nocodazole caused thorough depolymerization of the microtubules (not shown), but this treatment neither affected significantly the E-cadherin nor the phalloidin staining (Figure 2F, compare with A). However, the basal APC clusters completely disappeared after the drug treatment (Figure 2G, compare with B, arrows), while the membrane-associated APC staining was somewhat increased under these conditions (Figure 2F and G compare with A and B, arrowheads), indicating a re-localization of APC from the former to the latter. Prolonged recovery from drug treatment led to the restoration of the basal APC clusters (not shown). We conclude that MDCK cells retain two interconnected pools of peripheral APC protein after polarization: an actin-dependent APC pool associated with the lateral plasma membrane and basal APC clusters that depend predominantly on microtubules. Furthermore, given that the membrane association of APC is more pronounced in polarized MDCK cells compared with unpolarized cells, this indicates that the process of cell polarization may cause an equilibrium shift of the peripheral APC from microtubule tips to the lateral plasma membrane. Actin-dependent membrane association of APC in polarized WIF-B cells To determine whether our findings in MDCK cells also applied to other epithelial cells, we turned to WIF-B cells. These hepatic cells undergo two successive steps of polarization, attaining first a simple epithelial polarization, similar to that of polarized MDCK cells, followed by a secondary hepatic polarization that recapitulates some aspects of bile canaliculi differentiation (Ihrke et al., 1993; Decaens et al., 1996). After culture for 10–15 days, WIF-B cells form closed ovoid-shaped extracellular spaces between them that resemble bile canaliculi. Apical markers (such as HA4) segregate into the plasma membranes enclosing these ‘ovoids’, whereas basolateral markers are excluded from them (Figure 3A). Furthermore, the ovoids are lined with a belt of ZO-1 (Ihrke et al., 1993) (Figure 3A and B), a constituent of tight junctions that separate the apical from the basolateral plasma membranes in polarized cells. Figure 3.Actin-dependent membrane association of APC in polarized WIF-B cells. (A) Schematic representation of two adjacent polarized WIF-B cells forming a bile canaliculus-like structure between them, i.e. an ovoid of extracellular space enclosed by apical plasma membranes (HA4, apical marker); a belt of ZO-1 demarcates the junction between apical and basolateral plasma membranes (n, nucleus; from Ihrke et al., 1993, reproduced from The Journal of Cell Biology, 1993, 123, pp. 1761–1775 by copyright permission of The Rockerfeller University Press). (B–F) Confocal sections through polarized WIF-B cells after methanol fixation, ∼5 μm from the basal-most level, co-stained as indicated within the panels (merges show APC in green, ZO-1, β-catenin or α-tubulin in red), drug-treated as indicated on the right. Arrows point to membrane-associated APC staining, arrowheads to apical APC puncta [co-localizing with β-catenin puncta, as shown in the higher magnification images in (C) in which an ovoid is grazed tangentially]. Membrane-associated APC and apical APC puncta disappear after actin depolymerization (D; arrow marks an ovoid with residual apical APC puncta, as occasionally observed); both are unaffected by microtubule depolymerization (F). Note also that neither drug affects the integrity of the ovoids [as revealed by normal ZO-1 staining (D) and not shown]. Download figure Download PowerPoint In polarized WIF-B cells, we observed grainy cytoplasmic APC staining as well as a considerable level of APC staining along the lateral plasma membranes (Figure 3B, arrowheads). In addition, there were discrete APC puncta in the apical cortex (Figure 3B, arrows). These cortical APC puncta coincided with similar puncta of β-catenin staining that were associated with the ovoids (Figure 3C, arrows) although there was no E-cadherin staining along the ovoids (Decaens et al., 1996; data not shown). Exposure of polarized WIF-B cells to latrunculin A resulted in thorough depolymerization of actin filaments as revealed by phalloidin staining (not shown), but did not affect the frequency nor the shapes of the ovoids, as judged by the normal ZO-1 staining pattern (Figure 3D). However, APC was largely detached from the ovoids and from the lateral plasma membranes after this drug treatment (Figure 3D), with only residual APC puncta remaining associated with occasional ovoids (Figure 3D, arrow). Essentially the same was observed after exposure to cytochalasin D (not shown). In contrast, exposure of cells to nocodazole neither affected the cortical APC puncta nor the association of APC with the lateral plasma membrane, despite causing extensive disruption of the microtubules as revealed by α-tubulin staining (Figure 3F, compare with E). Thus, intact actin filaments are required for association of APC with the plasma membrane in WIF-B cells, whereas microtubules are dispensable for this association. We asked whether the dissociation of APC from the plasma membrane in drug-treated cells was reversible. Indeed, APC staining began to re-associate rapidly with lateral plasma membranes and ovoids after drug removal, and membrane-associated APC staining was restored to nearly normal after 15 min (Figure 4C, compare with A and B). These experiments confirmed that the association of APC with the plasma membrane depends on a continuous function of intact actin filaments. Figure 4.Reversible membrane association of APC in polarized WIF-B cells. Merged images of polarized WIF-B cells, controls (A) or treated with latrunculin A before (B) or 6–7 min after drug removal (C), co-stained with antibodies against APC (green) and ZO-1 (red). Note the restoration of apical APC puncta (arrows) and of membrane-associated APC staining (arrowheads) after recovery from drug treatment (C). Download figure Download PowerPoint APC truncations cannot reach peripheral locations In an attempt to identify the domains that target APC to the plasma membrane or to microtubule tips, we generated GFP-tagged versions of full-length APC and of three APC fragments. One of these spans the central third of the protein (MAPC–GFP) which is capable of complementing APC's nuclear export function in APC-mutant cancer cells (Rosin-Arbesfeld et al., 2000). The second one spanned the C-terminus (CAPC–GFP), a construct similar to one made by Mimori-Kiyosue et al. (2000) which is capable of decorating microtubules. The third one spanned the N-terminal half of the protein (NAPC–GFP), and mimics truncations that are typically found in APC-mutant cancer cells (Rowan et al., 2000). Since transfection of WIF-B cells was highly inefficient, we tested these constructs in transiently transfected MDCK cells before and after polarization. We found that CAPC–GFP and MAPC–GFP were expressed at high levels in transiently transfected MDCK cells, as revealed by western blot analysis of total cellular protein (Figure 5A, lanes 5 and 6, respectively). In contrast, NAPC–GFP was readily detectable only at early time points (e.g. at 18 h after transfection; Figure 5A, lane 4); however, 24 h after transfection, this fusion protein became undetectable by western blot analysis (see Materials and methods). This was due to gradual rounding off, detachment and, ultimately, lysis of transfected cells (not shown), indicating a noxious effect of this construct on cells. Similarly, full-length APC–GFP protein was only expressed at low levels, and was readily detectable by western blot analysis early on (18 h after transfection; Figure 5A, lane 3), fading away later, reflecting a gradual loss of transfected cells from the culture. This parallels earlier observations by Morin et al. (1996) who found that inducible overexpression of full-length APC in colorectal cancer cells caused the transfected cells to undergo apoptosis. Consistent with the western blot analysis, the frequency of transfected MDCK cells was high in the case of MAPC–GFP and CAPC–GFP, even at later time points, but was much lower in the case of APC–GFP and NAPC–GFP, and many of the cells transfected with the latter constructs looked somewhat unhealthy, especially after polarization (Figure 5D and E). Figure 5.Expression of APC–GFP fusion proteins in transfected MDCK cells. (A) Western blot analysis of APC fusion proteins transiently expressed in transfected subconfluent MDCK cells (lane 3, GFP–APC; lane 4, NAPC–GFP; lane 5, CAPC–GFP; lane 6, MAPC–GFP), probed with anti-GFP antibody; arrowheads indicate bands of the correct sizes expected for the different GFP fusion proteins (for comparison, endogenous full-length APC protein in HCT116 cells colorectal cancer cells, lane 1, and truncated APC protein in DLD-1 cells, lane 2, probed with anti-M-APC). (B–E) Confocal sections through subconfluent MDCK cells (B and C; basal-most plane) or confluent MDCK monolayer cells (D and E; ∼8 μm above basal-most plane), transfected with APC–GFP or NAPC–GFP (B and C, fluorescence; D and E, anti-GFP antibody), fixed with paraformaldehyde and co-stained as indicated above panels; representative examples are shown. Arrow in (B) points to a microtubule tip cluster formed by APC–GFP; arrowheads in panels B and D indicate membrane-associated APC–GFP. Neither of these is ever observed with NAPC–GFP, which shows an even cytoplasmic distribution (C and E). Download figure Download PowerPoint To assess the subcellular distributions of these GFP fusion proteins, we restricted our analysis to relatively healthy looking cells with low or moderate levels of expression. We found that, in transfected subconfluent MDCK cells, the green fluorescence due to full-length APC–GFP was very similar to the endogenous APC staining pattern: in many cells, we saw conspicuous clusters of APC–GFP at the cell periphery (Figure 5B, arrow; see also Supplementary data, Figure 3A), apparently associated with microtubule tips (not shown). Furthermore, we also observed cells in which some green fluorescence was associated with the plasma membrane (Figure 5B, arrowhead) in addition to variable levels of cytoplasmic APC–GFP. As mentioned above, a similar subcellular distribution of APC–GFP was observed in transiently transfected COS cells in which conspicuous APC–GFP clusters at peripheral microtubule tips were observed. This contrasted with the subcellular distribution of the GFP-tagged APC fragments which looked virtually indistinguishable from each other in transfected MDCK cells. Namely, none of the APC fragments showed any microtubule tip clusters nor any green fluorescence along the plasma membrane. Instead, each fragment produced a green fluorescence that was spread throughout the cytoplasm (Figure 5C, and data not shown). This was true even after nocodazole treatment which emphasizes the membrane-associated APC (see above): we never observed any lining of the plasma membrane with any of the APC fragments, whereas a significant proportion of the cells transfected with full-length APC–GFP showed membrane-associated green fluorescence after the drug treatment (not shown). Similarly, in transfected COS cells, we never observed any microtubule tip clusters with any of these GFP-tagged APC fragments (not shown). However, we did observe decoration of microtubules by CAPC–GFP, albeit not by MAPC–GFP nor by NAPC–GFP, in transfected COS cells, in agreement with the results of Mimori-Kiyosue et al. (2000) who showed that a C-terminal fragment of APC exhibited microtubule decoration in transfected Xenopus kidney epithelial cells. Note that the microtubule network is much more conspicuous, and readily detectable by immunofluorescence, in COS cells compared with MDCK cells, which may explain why the fluorescence pattern of CAPC–GFP was not detectably different from that of the other APC fragments in the latter. Essentially the same was found in transfected MDCK cells that were transferred to filters to allow monolayer formation and polarization. Namely, a significant fraction of cells transfected with full-length APC–GFP showed cytoplasmic GFP staining as well as distinct lining of the plasma membrane (Figure 5D, arrowheads), while none of the APC fragments produced GFP staining along the plasma membranes (Figure 5E). Furthermore, we observed occasional basal clusters in cell vertices only with full-length APC–GFP, but never with any of the APC fragments (not shown). These results indicate that neither the microtubule-dependent clusters of APC nor its membrane association is mediated by a single domain of APC. Furthermore, they imply that the APC truncations typically found in colorectal cancer cells would not be capable of reaching either of these peripheral locations. To test this, we examined the subcellular distribution of endogenous APC in a number of colorectal cancer cell lines, wild-type or mutant for APC. Indeed, cells that express wild-type APC (HCT116, LS174T; Figure 5A, lane 1) showed relatively low levels of cytoplasmic APC staining, characteristically grainy, but significant association of this staining with the plasma membrane, coinciding with β-catenin staining (Figure 6A) (see also Rosin-Arbesfeld et al., 2000). In contrast, APC-mutant cells that express truncated APC, for example DLD-1 or LOVO cells (Figure 5A, lane 2), showed moderately high levels of cytoplasmic as well as nuclear APC staining, but never any staining associated with the plasma membrane (Figure 6B; see also Supplementary data). Essentially the same was found in other APC mutant cells, e.g. in SW480 cells (Rosin-Arbesfel
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