Aberrant Cell–Cell Junctions in the Absence of αE-Catenin
DLD-1 is a colon carcinoma line with typical epithelial organization. DLD-1/R2/7, a subclone of DLD-1, showed a normal level of E-cadherin and β-catenin, but had no αE-catenin protein expression (Fig. A); the genetic background of this defect will be published elsewhere. The DLD-1/R2/7 line, abbreviated to R2/7, formed epithelioid clusters, but their intercellular associations were apparently loose (Fig. C) compared with those of the parent DLD-1 colonies (Fig. B). When R2/7 cells were transfected with αE-catenin cDNA, the original tightly associated epithelial morphology was restored (Fig. D). As these αE-catenin transfectants were indistinguishable from the parent DLD-1, one of their clones, R2/7-αE, was used as a representative line of the normal cells throughout subsequent experiments.
Figure 1 Junctional organization in DLD-1, R2/7, and R2/7-αE cells. (A) Immunoblot analysis for E-cadherin (lanes 1 and 2), β-catenin (lanes 3 and 4), and αE-catenin (lanes 5 and 6) in DLD-1 (lanes 1, 3, and 5) or in DLD-1/R2/7 (lanes (more ...)
To determine how the loss of αE-catenin affects junctional organization, we compared R2/7 and R2/7-αE for their expression of various junctional proteins. Double- immunostaining for E-cadherin and ZO-1, a component of the tight junction (TJ), showed that in R2/7-αE, these molecules were localized at cell–cell boundaries in the pattern typical for many simple epithelia: ZO-1 was confined at the most apical portion of cell–cell contacts, displaying a honeycomb-like distribution pattern (Fig. E). E-cadherin was more broadly distributed at the lateral cell–cell boundaries, and its apical edge coincided with the ZO-1 staining (Fig. H). On the other hand, in R2/7, ZO-1 was accumulated in a circular or patchy pattern on the top surface, never showing the honeycomb pattern (Fig. F); and it was rarely localized in the lateral membranes (Fig. G). E-cadherin was detected in two separate positions, one along the ZO-1-positive rings (Fig. I) and the other at portions of the lateral membrane (Fig. J). The latter E-cadherin signals were condensed at cell-cell contact sites, being little associated with ZO-1 (Fig. , G and J).
The ZO-1–positive rings on the top surface of R2/7 cells were also delineated by staining for actin (Fig. , P and Q), occludin (Fig. R), and α-actinin (Fig. S). By contrast, vinculin was never localized along the ZO-1 rings (Fig. T) nor at the lateral cell–cell contact sites in the R2/7 cells (Fig. D). In R2/7-αE, on the other hand, all these junctional proteins were accumulated together at the apical cell–cell boundaries (Fig. K–O), coinciding with the apical signals of E-cadherin. In addition, E-cadherin was always colocalized with β-catenin at cell–cell boundaries, and also coimmunoprecipitated with this catenin from both cell lines (data not shown), supporting the widely accepted view that they form a direct molecular complex.
Figure 4 Colocalization of vinculin or α-actinin with E-cadherin. (A and D) R2/7; (B and E) R2/7-αE; (C and F; H and K) R2/7-αE(1–509). (G and J; I and L) R2/7-αE(1–325/510–890). Double immunofluorescence (more ...)
The above observations suggested that TJs were disorganized by the loss of αE-catenin. To further examine this point, the above cells were analyzed by a freeze-fracture replica method. In R2/7-αE, typical TJ strands arranged into a mesh-like pattern were detected (Fig. U). Unexpectedly, even in R2/7 cells similar structures were found, although they were discontinuous and patchy (Fig. V), suggesting that local TJ formation occurs in these cells. As described later, R2/7 cells are connected with neighbors through filopodial processes at marginal areas of their apical surface (Fig. I); these contact points could serve as sites for such discontinuous TJ formation. These findings indicate that the loss of αE-catenin did not block TJ formation per se, but abolished its network-type organization.
Figure 7 Association pattern of cells expressing various αE-catenin constructs. (A–J) Vertical EM sections of cells in monolayer cultures seen at a low magnification (A–C, G and H), and their close-up views at apical cell–cell (more ...)
Amino Acid Residues 326–509 of αE-Catenin are Necessary for Organization of the ZO-1 Network
To determine the domains of αE-catenin necessary for TJ assembly, we generated a series of its deletion mutants, tagged with T7 at their COOH terminus (Fig. A
). These were introduced into R2/7 cells, and subsequently multiple transfectant clones were isolated for each construct, among which a representative clone was chosen for the following experiments. Each clone expressed a protein with the expected size (Fig. B
). From lysates of these clones, antibodies to the T7 tag coimmunoprecipitated E-cadherin only when the αE-catenin constructs contained the NH2
-terminus (Fig. C
), consistent with the finding that the NH2
-terminal region of α-catenin is required for its association with β-catenin (Bullions et al., 1997
; Nieset et al., 1997
; Obama and Ozawa, 1997
; Sehgal et al., 1997
Figure 2 Deletion constructs of αE-catenin and their characterization. (A) Schematic drawing of the mutant αE-catenin constructs, designated as 1–8, introduced into R2/7 cells. Stable transfectants were isolated for each construct. ( (more ...)
Cells expressing the above mutant αE-catenin molecules were stained for ZO-1 to assess their TJ organization. αE(1–509) in which the COOH-terminal 397 residues had been truncated induced the formation of the typical honeycomb ZO-1 network (Fig. D). However, αE(1–325) with a longer COOH-terminal deletion failed to reorganize the ZO-1 distribution (Fig. E). These observations suggest that residues 326–509 are important for ZO-1 organization. Consistently, cells expressing αE(1–325/510–890), in which the 326–509 portion was deleted but the other regions were left intact, could not redistribute ZO-1 (Fig. F), although ZO-1 tended to be more frequently condensed at cell–cell contact sites in these transfectants than in R2/7 cells. E-cadherin was redistributed into a normal pattern with αE(1–509), but not with αE(1–325) or αE(1–325/510–890; Fig. , G–I). By phase-contrast microscopy, αE(1–509) cells showed tighter cell–cell associations than the other transfectants (Fig. , A–C). The mutant αE-catenins lacking the β-catenin–binding domain had no effect on ZO-1 assembly. The staining pattern of occludin was identical to that of ZO-1 in all the above cell lines (data not shown).
Figure 3 Only αE(1–509) can rescue R2/7 cells to organize the TJ. (A, D, and G) R2/7-αE(1–509). (B, E, and H) R2/7-αE(1–325). (C, F, and I) R2/7-αE(1–325/510–890). (A–C) Phase-contrast (more ...)
Interaction of Vinculin with aE-Catenin at the 326–509 Domain
As a step to investigate the role of the 326–509 domain, we examined the localization of various proteins in the above series of αE-catenin transfectants that are known to be associated with the junctional complex. As mentioned already, vinculin was completely absent from the top to lateral surfaces in R2/7 cells, not colocalizing with E-cadherin (Fig. T
and Fig. , A
), although it was normally located at focal adhesion sites to the substratum (data not shown). However, when the cells expressed the full-length αE-catenin, vinculin was redistributed to cell–cell boundaries to colocalize with the apical-most signals of E-cadherin (Fig. , B
); the same colocalization was induced by expression of αE(1–509) (Fig. , C
). The vinculin distribution was sharp, and was restricted to the apical level of cell–cell contacts close to the ZO-1 localization (data not shown), as observed for many normal epithelial cells (e.g., Yonemura et al., 1995
). All other constructs lacking the 326–509 domain, including αE(1–325/ 510–890), never led vinculin to colocalize with E-cadherin (Fig. , G
). These results suggest that the 326–509 region of αE-catenin plays a role in attracting vinculin to the E-cadherin–positive cell–cell contacts. Vinculin localization at the focal adhesion sites appeared to be normal in all the transfectant lines. α-Actinin also codistributed with E-cadherin in the αE(1–509) cells (Fig. , H
). However, this protein sporadically colocalized with E-cadherin, even in cells expressing the αE-catenin without the 326– 509 domain, such as αE(1–325/510–890) (Fig. , I
), indicating no consistent correlation between the α-actinin/ E-cadherin codistribution and the presence of the 326–509 domain. As far as examined, vinculin was the only protein that responded to the presence or absence of the 326–509 domain in its redistribution to the apical cell–cell contact sites, although other unidentified proteins could show similar behavior.
For more direct identification of molecules that can associate with the 326–509 domain, we generated GST-αE-catenin fusion proteins having the 1–325 or 1–509 domain (Fig. A
), and incubated them with chick gizzard lysates as a source of cytoskeletal proteins. SDS-PAGE analysis of the bound materials revealed that the 1–509 construct precipitated two major proteins that migrated to the 130–120 kD region, but the other construct did not (Fig. B
). In Western blotting, both bands reacted with antibodies to vinculin (Fig. B
; middle, arrowheads
), suggesting that they are two variant forms of vinculin: metavinculin, and vinculin (Molony and Burridge, 1985
). As controls, we found that β-catenin was copurified with both constructs as expected, but α-actinin and spectrin were not associated with either construct (Fig. B
). These findings indicate that the 326– 509 domain is involved in the association of αE-catenin with vinculin, consistent with the above immunostaining results. No other proteins were copurified specifically with this domain; at least as major components, as judged by silver staining of the above electrophoretic samples.
Figure 5 In vitro binding of vinculin with αE-catenin. (A) Schematic drawing of GST-αE-catenin fusion proteins (1–7), and of MBP-vinculin fusion proteins (MBP-vinHead and MBP-vinTail). (B) Detection of proteins bound to the GST-αE-catenin (more ...)
To test if the association of αE-catenin with vinculin is mediated by their direct binding, we constructed other GST-αE–catenin mutant molecules (Fig. A
), including a molecule consisting of only the 326–509 domain, and subjected them to assays of their ability to bind the head domain or tail domain of vinculin constructed as MBP-fusion proteins (Fig. A
). The vinculin head domain (MBP-vinHead) bound not only the intact αE-catenin, but also to αE(1–509) and αE(326–509) (Fig. C
); it did not bind any other constructs lacking the 325–509 domain of αE-catenin. On the other hand, the tail domain of vinculin (MBP-vinTail) reacted with none of the αE-catenin constructs (Fig. C
). All these results indicate that αE-catenin can directly bind to the head domain of vinculin at the 326–509 domain. We also examined if E-cadherin could be copurified with vinculin by immunoprecipitation, but the results were negative as previously observed (Knudsen et al., 1995
; Tsukatani et al., 1997
; Hazan et al., 1997
). It is possible that the E-cadherin/αE-catenin/vinculin complex was not soluble to our detergent solutions, as we found that their colocalization was still detected in the cells extracted with nonionic detergents. Alternatively, their association might have been disrupted during the detergent extraction; it is of note that the above in vitro binding assays were carried out under detergent-free conditions.
αE-Catenin-Vinculin Chimeric Proteins Can Induce the ZO-1 Network
As found above, the activity of αE-catenin to organize the ZO-1 network was correlated with its ability to bind vinculin. This finding suggests that the ZO-1–organizing activity of αE-catenin could be mediated by vinculin itself. To test this idea, we constructed cDNAs encoding the 1–325 αE-catenin fragment whose COOH terminus was fused with the head (αE/vinHead) or tail (αE/vinTail) domain of vinculin (Fig. A
), and these constructs were introduced into R2/7 cells to isolate transfectants. The 1–325 αE-catenin fragment by itself cannot redistribute ZO-1, as mentioned above. Both chimeric proteins were found to bind the E-cadherin/β-catenin complex (data not shown). When αE/vinTail was expressed, ZO-1 was redistributed to exhibit a honeycomb pattern (Fig. B
), colocalizing with E-cadherin or tag signals (Fig. D
). On the other hand, αE/vinHead expression could not reorganize the ZO-1 distribution (Fig. , C
). These results suggest that the tail domain of vinculin, associated with αE-catenin, plays a role in ZO-1 organization. Incidentally, in the αE/vinTail-expressing cells, α-actinin did colocalize with E-cadherin or αE/vinTail (Fig. , F
) despite the absence of the identified α-actinin–binding sites in this chimeric protein, which are amino acids 325–394 in αE-catenin (Nieset et al., 1997
) and amino acids 1–107 in vinculin (Kroemker et al., 1994
). Therefore, the α-actinin/E-cadherin colocalization often observed during this study does not necessarily suggest that they are forming a molecular complex.
Figure 6 An αE-catenin/vinculin chimeric protein can redistribute ZO-1. (A) Schematic drawing of αE-catenin/vinculin chimeric proteins. The NH2-terminal 1–325 amino acid domain was fused with the NH2-terminal 1–823 or COOH-terminal (more ...)
Morphological Assessment of Functions of Different αE-Catenin Domains
To gain further insight into the role of different domains of αE-catenin, we analyzed the junctional structures of cells expressing various αE-catenin constructs by EM. R2/ 7-αE cells formed monolayer sheets with a smooth surface in which individual cells were essentially of cuboidal shape and closely associated with each other at their lateral membranes (Fig. A). At the apical-most portion of their lateral contacts, the junctional complexes were observed (Fig. D). The ZA was not always clearly detectable as a domain separable from the TJ, suggesting that these two junctions might be fused to each other. Similar apical junctional structures were observed in cells expressing αE(1–509) (Fig. E) and αE/vinTail (Fig. F). Despite such relatively normal organization of the apical junctional complexes, these cells differed from R2/7-αE cells in their overall association patterns. Cells with αE(1–509) tend to overlap each other at their basolateral regions (Fig. B), suggesting that their adhesive interaction was not perfectly controlled at the nonapical lateral membranes. In the case of αE/vinTail-expressing cells, their intercellular contacts were limited to the areas with the apical junctional complex and desmosomes; large intercellular spaces were formed at the sites where these junctional structures were absent (Fig. , C and F).
On the other hand, in R2/7 cells no typical junctional complex was observed (Fig. I); their plasma membranes were simply apposed to each other with an occasional presence of desmosomes, and in some cells the apical-most portion of the lateral contacts was occupied by a desmosome instead of the junctional complex (Fig. I, insert). Interestingly, these cells extended filopodia from marginal areas of their apical surface, and formed spotty contacts with neighboring cells at similar areas. At these contact sites, electron-dense junction-like structures were observed (Fig. I, arrowheads). Also, in cells expressing αE(1–325), αE(1–184), αE(1–326/509–890) (Fig. J), and αE/vinHead, no apical junctional complexes were found. As an overall view, these cells that are unable to form the apical junctional complex exhibited very irregular patterns of intercellular association (Fig. , G and H).
We then attempted to estimate the E-cadherin–mediated aggregating activity of cells expressing different αE-catenin constructs. An ideal experiment to this end is to measure the rate of reaggregation of cells that have been dissociated with preservation of their cadherins by the trypsin-Ca2+
-treatment method (Takeichi, 1977
; Takeichi, 1988
). However, DLD-1 cells were difficult to dissociate into single cells by this method. We therefore carried out the following assay: cells were dissociated into single cells by a trypsin–EDTA treatment, and were incubated overnight to induce cell reaggregation in the absence or presence of E-cadherin–blocking antibodies. Without the antibodies, R2/7-αE cells formed compact aggregates in which cells were tightly packed (Fig. K
), whereas with the blocking antibodies, the cells remained single or formed only loose clusters (Fig. P
). In contrast to R2/7-αE ones, R2/7 cells loosely aggregated, and their aggregates were never compacted (Fig. L
), although this aggregation was still sensitive to the anti–E-cadherin antibodies, as the cells were further dispersed by the antibodies (Fig. Q
). Aggregates of cells with αE(1–509) (Fig. M
) or αE/vinTail (Fig. O
) exhibited a morphology intermediate to that of the above cases; they were partly compacted, although the latter aggregates tended to be less compacted. The aggregation profile of the cells expressing αE(1–325), αE(1– 184), αE(1–325/510–890), and αE/vinHead was essentially identical to that of R2/7, although those with αE(1–325/ 510–890) were slightly more adhesive than the others (Fig. N
). To summarize, the activity of αE-catenin to induce compacted aggregates was expressed in the following order: intact αE-catenin > αE(1–509) ≥ αE/vinTail > αE(1– 325/510–890) ≥ αE(1–325) = αE(1–184) = αE/vinHead = no αE-catenin.
Apical Junctional Organization in Vinculin-deficient Cells
Finally, to confirm the importance of vinculin in cell–cell junction formation, we examined junctional organization in two independent sublines of F9 embryonal carcinoma cells—γ227 and γ229—whose vinculin genes had been disrupted (Coll et al., 1995
). Normal F9 cells, cultured at low densities, appeared flat and epithelioid. Immunostaining of these cells for ZO-1 showed that this protein was discontinuously distributed at cell–cell boundaries (Fig. , A
) as typically seen in fibroblasts; nevertheless, the ZO-1 signals were closely associated with vinculin (Fig. B
) as well as with E-cadherin at its apical borders (Fig. F
), as observed in epithelial cells. In the vinculin-null F9 cells that were less flat than the normal F9 cells, ZO-1 and E-cadherin were distributed in a pattern similar to that found in the normal cells (Fig. , C
) despite the absence of vinculin (Fig. D
), although the ZO-1 pattern appeared more complicated in the mutant cells. As these cells grew and became more packed, we noted a dramatic difference in the ZO-1 distribution between the normal and vinculin-null cells. In normal cells, ZO-1 tended to be reorganized into honeycomb- or web-like networks on their top surfaces (Fig. , I
). These ZO-1 signals colocalized with vinculin (Fig. J
), and also with the apical-most E-cadherin signals (Fig. N
). In contrast, the vinculin-null F9 cell layers rarely formed such closed web-like networks of ZO-1, maintaining the discontinuous ZO-1 pattern (Fig. , K
), although they sometimes displayed partially closed organization of this molecule. The same results were obtained with both γ227 and γ229 cells. To confirm whether this mutant phenotype was directly brought about by the loss of vinculin, we further examined two sublines of γ229—R3 and R15—in which vinculin expression had been restored by virtue of vinculin cDNA transfection (Xu et al., 1998b
). In both of the rescued lines, the web-like distribution of ZO-1, colocalizing with vinculin, was normally generated (Fig. , Q
). The above ZO-1 networks were observed only at the top focal plane of the cell layers. These findings support the idea that vinculin is involved in organization of the apical-most junctions.
Figure 8 Junctional organization in normal and vinculin-null F9 cells. (A–H) Low cell density. Cells (1 × 105) were inoculated on a collagen-coated coverslip placed in each 3.5-cm dish, and were cultured for 2 d. (I–R) High cell density. (more ...)
When we induced differentiation in the F9 lines by means of aggregation cultures with retinoic acid as described (Stephens et al., 1993
), some of the resultant differentiated cells, likely visceral endoderm cells, exhibited a honeycomb-like organization of ZO-1, even in the absence of vinculin (data not shown). This finding suggests that vinculin may not be absolutely required for junctional organization in certain cell types or under certain physiological conditions.