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© The Rockefeller University Press, 0021-9525/1998//1105 $5.00
The Journal of Cell Biology, Volume 142, Number 4, , 1998 1105-1119

Mechanisms of Epithelial Cell–Cell Adhesion and Cell Compaction Revealed by High-resolution Tracking of E-Cadherin– Green Fluorescent Protein

Department of Molecular and Cellular Physiology, Stanford University School of Medicine, Stanford, California 94305-5345

Cadherin-mediated adhesion initiates cell reorganization into tissues, but the mechanisms and dynamics of such adhesion are poorly understood. Using time-lapse imaging and photobleach recovery analyses of a fully functional E-cadherin/GFP fusion protein, we define three sequential stages in cell–cell adhesion and provide evidence for mechanisms involving E-cadherin and the actin cytoskeleton in transitions between these stages. In the first stage, membrane contacts between two cells initiate coalescence of a highly mobile, diffuse pool of cell surface E-cadherin into immobile punctate aggregates along contacting membranes. These E-cadherin aggregates are spatially coincident with membrane attachment sites for actin filaments branching off from circumferential actin cables that circumscribe each cell. In the second stage, circumferential actin cables near cell–cell contact sites separate, and the resulting two ends of the cable swing outwards to the perimeter of the contact. Concomitantly, subsets of E-cadherin puncta are also swept to the margins of the contact where they coalesce into large E-cadherin plaques. This reorganization results in the formation of a circumferential actin cable that circumscribes both cells, and is embedded into each E-cadherin plaque at the contact margin. At this stage, the two cells achieve maximum contact, a process referred to as compaction. These changes in E-cadherin and actin distributions are repeated when additional single cells adhere to large groups of cells. The third stage of adhesion occurs as additional cells are added to groups of >3 cells; circumferential actin cables linked to E-cadherin plaques on adjacent cells appear to constrict in a purse-string action, resulting in the further coalescence of individual plaques into the vertices of multicell contacts. The reorganization of E-cadherin and actin results in the condensation of cells into colonies. We propose a model to explain how, through strengthening and compaction, E-cadherin and actin cables coordinate to remodel initial cell–cell contacts to the final condensation of cells into colonies.

Key Words: cadherin • actin • cell–cell adhesion • epithelia • microscopy

Abbreviations used in this paper: CD, cytochalasin D; DIC, differential interference contrast; EcadGFP, E-cadherin fused to green fluorescent protein; synGFP, synthetic jellyfish green fluorescence protein; TIP, time, intensity, and position.

CELL–CELL adhesion is crucial for the development and survival of multicellular organisms (Townes and Holtfreter, 1955; Takeichi, 1991; Steinberg, 1996). Members of the cadherin superfamily of Ca²⁺- dependent cell–cell adhesion proteins are expressed in most organs and tissues of vertebrates and invertebrates. Different cadherins are expressed in specific tissues, cell layers, and neuronal cell types consistent with their roles in distinct cellular recognition and sorting processes (Takeichi, 1987; Nose et al., 1988; Takeichi, 1991; Steinberg and Takeichi, 1994; Fannon and Colman, 1996; Uchida et al., 1996; Martinek and Gaul, 1997). While recent studies have attempted to elucidate the molecular and mechanical properties of cadherin-mediated adhesion (Brieher et al. 1996; Yap et al., 1997), little is known about the physical or molecular dynamics of cell–cell adhesion (cell stickiness), compaction (maximization of adhesive contacts), or condensation (aggregation of large cell colonies) during tissue formation.

Cadherins are single transmembrane–spanning proteins. The extracellular domain (amino terminus) is composed of five repeats that have similar structures and contain Ca²⁺-binding motifs (Shapiro et al., 1995; Aberle et al., 1996). While homotypic binding between the extracellular domains of cadherins on adjacent cells is clearly important for cell–cell recognition (Nose et al., 1988), the affinity of binding (1 µM; Shapiro et al., 1995) may not be sufficient to promote the strong cell–cell adhesion necessary to maintain tissue integrity. X-ray diffraction studies of crystals of the amino-terminal repeat domains of E-cadherin and N-cadherin revealed the presence of dimers and higher ordered complexes (Nagar et al., 1996; Shapiro et al., 1995). Formation of higher-order complexes between extracellular domains of parallel-oriented cadherins on single cells and clustering of extracellular E-cadherin between cells might cooperatively increase the strength of adhesion (Brieher et al., 1996; Yap et al., 1997), but little is known about how, when, or where clustering of cadherins occurs during cell–cell adhesion in vivo.

The cytoplasmic domain of cadherin is also required for cell–cell adhesion. The amino acid sequences of the cytoplasmic domain of different cadherins are very similar, and contain a highly conserved binding site for a family of sequence-related cytosolic proteins: β-catenin, plakoglobin, and p120^CAS (Aberle et al., 1994; Jou et al., 1995; Ozawa et al., 1989; Reynolds et al., 1994). The cadherin/β-catenin complex binds via β-catenin to another cytosolic protein, -catenin (Herrenknecht et al., 1991; Jou et al., 1995; Ozawa and Kemler, 1992), which interacts with actin filaments either directly (Rimm et al., 1995) or through other actin-associated proteins such as -actinin (Knudsen et al., 1995). Binding of this protein complex to the actin cytoskeleton is consistent with the appearance of a pool of cadherins and catenins at cell–cell contacts that is resistant to extraction by the nonionic detergent Triton X-100 (Nagafuchi and Takeichi, 1988; Ozawa et al., 1989).

In individual motile cells, actin filaments are continually polymerizing at the free cell edge of lamellae, and depolymerizing in a transition zone between the cell body and lamellae. This transition zone is often characterized by a conspicuous ring of actin that is called a circumferential actin cable (see review, Small et al., 1996). In single motile epithelial cells, such actin cables often circumscribe the cell in the form of a continuous ring. In polarized epithelial cells, actin filaments are also organized into a much thinner, much more peripherally disposed circumferential ring at the apical surface in association with the adherens junction (Hirano et al., 1987); this actin organization is also referred to as a circumferential actin cable. While circumferential actin structures are thus characteristic of both motile and tissue forms of epithelial cells, a dramatic structural transformation in actin organization occurs during the formation and stabilization of cell–cell contacts. Recent studies have suggested that circumferential actin cables in single cells either passively become parallel to cell–cell contacts (Yonemura et al., 1995) or they actively reorganize to the cell periphery (Gloushankova et al., 1997). These studies have not provided significant insight into how actin filaments in lamellae and circumferential actin cables reorganize when two lamellae interact in forming a cell–cell contact. Thus, while the actin cytoskeleton is known to be involved in cadherin-mediated cell–cell adhesion, little is known about how actin participates in the initiation and strengthening of cell–cell adhesion, or how the organization of the actin cytoskeleton in single motile cells becomes incorporated into the actin organization observed in monolayers of polarized epithelial cells.

Even less is known about the molecular mechanisms of compaction and condensation of single cells into multicellular colonies. On the other hand, much work has been done to elucidate the mechanisms involved in the healing of wounded monolayers and tissues (Martin and Lewis, 1992; Bement et al., 1993). Small wounds in cell monolayers rapidly form circumferential actin cables at the wound perimeter, which then slowly cinch together with a purse-string action (Bement et al., 1993; Brock et al., 1996). Similarly, tissue explants with ragged edges will round up in vitro over extended periods of time, presumably by a similar purse-string action. The mechanisms that mediate these events could play a role in the genesis of multicellular monolayers, but previous studies have not established similarities among these processes.

The experimental approaches used in the studies described above have not provided much insight into the dynamic processes by which cadherins, catenins, and the actin cytoskeleton cooperate to initiate, strengthen, and compact cell–cell contacts between cells initiating adhesion or reorganizing within colonies. In previous studies, we used differential interference contrast (DIC) time-lapse imaging coupled with retrospective immunocytochemistry to examine the distributions of E-cadherin, catenins, and actin in adhering MDCK cells (McNeill et al., 1993; Adams et al., 1996). We showed that during the first hour of cell–cell adhesion, E-cadherin, β-catenin, and -catenin coaccumulated into Triton X-100–insoluble aggregates (puncta) that are associated with thin actin bundles (Adams et al., 1996). However, from these previous studies many problems related to initiation of cell–cell adhesion were unresolved. For example, we were unable to determine the relationship of Triton X-100 insolubility and clustering of E-cadherin, the source of E-cadherin in puncta, the role of the actin cytoskeleton in the spatial organization of puncta, the dynamics and fate of puncta within the contact, or the dynamics of actin and cadherin reorganization in older contacts. Furthermore, we were unable to investigate the role(s) of these E-cadherin puncta in strengthening and compacting initial adhesive contacts, or the mechanisms of reorganization of cell–cell contacts as cells condensed into colonies to form a multicell monolayer. Answers to these problems are at the core of understanding mechanisms involved in cell–cell adhesion.

In the present study, we examined the dynamics of cell– cell adhesion using a fully functional protein comprising E-cadherin fused to green fluorescent protein (EcadGFP).¹ EcadGFP was stably expressed in MDCK epithelial cells, and was examined with time-lapse imaging and photobleach-recovery analysis. We define three sequential stages in cell–cell adhesion, and show how the distributions of E-cadherin and the actin cytoskeleton are remodeled to coordinate transitions in cell–cell adhesion from initial contacts, to strengthening and compaction, to the final condensation of cells into colonies. Our results provide new detailed insights into the dynamics and mechanisms involved in regulating epithelial cell–cell adhesion in vivo.

	Materials and Methods

Top Abstract Materials and Methods Results Discussion References

 
Construction of E-cadherin–GFP Recombinant cDNA and Stable Transfection of MDCK Cells
Recombinant cDNA of a synthetic jellyfish green fluorescence protein (synGFP; Haas et al., 1996; a gift from Dr. Brian Seed) was used as the PCR template to amplify the complete synGFP coding region, with Xho1 and Not1 sites added to the 5' and 3' ends, respectively. The PCR product was subcloned into CDM8FluTag (Chen et al., 1993) through Xho1/Not1 sites that generated the plasmid HA-synGFP. A second PCR reaction using canine E-cadherin cDNA as the template amplified the complete coding region of E-cadherin (without a stop codon) with a Hind3 site added at the 5' end and a Xho1 site added at the 3' end. The PCR product was subcloned into HA-synGFP through Hind3 and Xho1 sites after restriction of HA-synGFP with Hind3 and Xho1 to release the influenza virus hemagglutin tag (Chen et al., 1993), and resulted in an in-frame fusion of E-cadherin and synGFP with the Xho1 site as the linker (U-GFP1). Most of the extracellular domain of E-cadherin–GFP fusion protein cDNA was then replaced with a Hind3/Bgl2 fragment from the cDNA of E-cadherin pCEcad1 after restricting U-GFP1 with Hind3/Bgl2 enzymes, to generate U-GFP2. Accordingly, there is no sequence difference between the coding regions for E-cadherin–GFP fusion protein in U-GFP1 and U-GFP2, but most of the extracellular domain encoded by U-GFP2 is from the E-cadherin cDNA instead of the PCR product. The E-cadherin-GFP fusion cDNA in U-GFP2 was confirmed by restriction mapping and DNA sequencing, and was used for transfection. MDCK IIG cells were transfected with U-GFP2 using Ca²⁺ phosphate (Graeve et al., 1990) with pSV2neo (Southern et al., 1982) as the selection marker. Eight positive clones expressing the E-cadherin–GFP fusion cDNA in U-GFP2 (EcadGFP) were isolated using cloning rings, and all clones gave identical patterns of EcadGFP fluorescence and Western blot profiles.

Expression of EcadGFP in Mouse L-Cells
To express EcadGFP in Ltk-cells stably, the E-cad-GFP cDNA was released from pUGFP2 by Hind3 and Not1 digestion, and was subcloned into the retroviral vector LZRS-pBMN-Z (Kinsella and Nolan, 1996) after restriction by Hind3 and Not1. Production of virus and infection of Ltk– cells was performed as previously reported (Kinsella and Nolan, 1996).

Cell-Cell Adhesion Mediated by EcadGFP
Aggregation.
Ltk-cells and Ltk-cells stably transfected with U-GFP1 were tested for their ability to aggregate in suspension culture. Approximately 0.5 x 10⁶ single cells were plated in DMEM containing 5 µM Ca²⁺ and 10% FCS that had been dialyzed extensively against PBS, or normal DMEM/FBS (1.8 mM Ca²⁺) on 35-mm tissue culture dishes coated with 2% agarose (Sigma Chemical Co., St. Louis, MO) and left in a tissue culture incubator overnight. The organization of cells in clumps were recorded 18 h after plating using a Zeiss Axiovert microscope equipped with a 10x phase-contrast objective.

Kinetics of aggregation.
Wild-type MDCK type G cells, MDCK cells expressing EcadGFP, and MDCK cells expressing EcadGFP treated for 24 h with 5 mM sodium butyrate were suspended with trypsin/EGTA treatment, counted, and adjusted to a density of 7 x 10⁶ cells/ml in imaging buffer (DMEM without phenol red, 10% FBS, 50 mM Hepes). Cells were allowed to recover for 2 h before aggregation measurements. Cells were vortexed for 30 s immediately before loading into the cuvette of the aggregometer. Cells were immediately placed in an optical aggregometer (Model 440; Chrono-log, Corp., Havertown, PA) at 37°C. 270 µl of cells were transferred to a 0.6-mm diameter cuvette, and were kept in suspension by constant stirring at 300 rpm. Spontaneous aggregation was measured by monitoring the increase in OD of the cell suspension on a strip-chart recorder for up to 6 h with cell-free buffer as a reference. Maximal aggregation was determined by the value when changes in the transmitted light were zero. The half-time was manually determined as the time at half-maximal aggregation.

Binding of Catenins to EcadGFP in HEK 293 Cells and MDCK Cells
HEK 293-EBNA cells (Invitrogen Corp., Carlsbad, CA) were transfected with the E-cadherin-GFP plasmid, U-GFP2, using lipofectamine (GIBCO BRL, Gaithersburg, MD). 24 h after transfection, cells in one 35-mm dish were labeled with 250 µCi of [³⁵S]Met/Cys (Amersham Life Science, Inc., Arlington Heights, IL) for 24 h. At the end of the labeling period (48 h after transfection), cells were lysed in Triton X-100 lysis buffer (10 mM Tris-HCl, pH 7.5, 120 mM NaCl, 25 mM KCl, 2 mM EDTA, 2 mM EGTA, 0.5% Triton X-100), and proteins were immunoprecipitated with mAb 3G8, which binds to the extracellular domain of E-cadherin. The immunoprecipitate was resolved by 7.5% SDS-PAGE, and the radioactive signals were detected by fluorography. Identical labeling and immunoprecipitation experiments were performed with MDCK IIG cells stably transfected with U-GFP2.

Delivery of EcadGFP to the Basal-lateral Membrane of Polarized MDCK Cells
Confluent monolayers of MDCK cells stably transfected with U-GFP2 were grown on Transwell^TM filters (Costar Corp., Cambridge, MA) for 7 d, and were treated with 5 mM sodium butyrate for 24 h before an experiment. Cells were first incubated with Met/Cys-free medium for 45 min, and then cells on one 24-mm diameter petri dish were labeled with 250 µCi of [³⁵S]Met/Cys for 1 h. After labeling, plasma membrane domain– specific biotinylation was performed as previously described (Wollner et al., 1992). Cells were lysed in Triton X-100 extraction buffer, and proteins in the lysate were immunoprecipitated with mAb 3G8. Biotinylated protein from the immunoprecipitates was collected with immobilized avidin (Pierce, Rockford, IL). Precipitated proteins were resolved by 7.5% SDS-PAGE. Radioactive signals were detected by fluorography.

Cell Culture
Stably transfected EcadGFP MDCK cells were maintained at low density in DMEM (Gibco Laboratories, Grand Island, NY) supplemented with 10% FBS (Summit, Ft. Collins, CO). 12–24 h before experimentation, 5 mM sodium butyrate in DMEM/FBS was added to 90% confluent cultures of cells to further induce expression of EcadGFP. Cells were removed with trypsin and plated at low density in DMEM/FBS on collagen-coated coverslips for 2–5 h. For time-lapse imaging, cells were removed from the incubator, and the media was replaced with imaging buffer (DME without phenol red, 10% FBS, 50 mM Hepes). The coverslip was placed in a custom-built imaging chamber with 500 µl of imaging buffer. Cytochalasin D (Sigma Chemical Co.) solution was diluted 1:2,000 from a stock solution in DMSO to 2 µM in the imaging buffer.

Time-lapse Imaging
The Smith Mark IV multisite laser scanning confocal microscope, designed by Stephen Smith and built by Stanford's Physiology Instrument Shop, has been described elsewhere (Adams et al., 1996). In brief, it is an optical bench design using argon (488 nm) and helium-neon (633 nm) lasers, a mirror galvanometer–based scanning unit, a GaAs photomultiplier and silicon photodiode photodetectors, and a PC-based control and data acquisition unit. It allows collection of multispectral fluorescence and transmitted light (e.g., Nomarski DIC) images in perfect spatial register as determined by a single laser scanning geometry. The multisite feature (see Cooper and Smith, 1992; McNeill et al., 1993; Adams et al., 1996; software written by Dr. Noam Ziv) provides for the acquisition of time-lapse sequences at multiple specimen areas by frequent automated motions of a motorized specimen stage combined with use of an autofocusing algorithm. For observing the maturation of cell–cell contacts, time-lapse sequences were collected at 6–12 stage-position sites with consecutive DIC and fluorescence images every 2–10 min for 2–23 h at 37°C. For tracking the dynamics of new puncta, images were collected at single sites every 0.9 s to 1 min for 3–100 min.

Image Analysis
To generate time, intensity, and position graphs (TIP scans), images were rotated in Adobe Photoshop (Adobe Systems, Inc., Mountain View, CA) so that the long axis of the contact in the last image of the time series was parallel to the horizon. The rotated images were imported into MetaMorph software (Universal Imaging Corp. West Chester, PA), and the developing contact area was boxed off by 50–200 adjacent horizontal regions, 2 pixels high by 60–200 pixels wide, depending on the relative position and length of the contact through the time sequence. MetaMorph software automatically collected maximum and integrated intensity data from contact, noncontact, and noncell (background) regions for all time points. These data were then imported into Excel (Microsoft, Redmond, WA) and were corrected for photobleaching during the imaging period (<10%). Three-dimensional graphs were generated in Excel, and the pseudocolor intensity scale was created for ease of interpretation by color-blind persons and after photocopying (Livingstone, 1988). Intensity values from regions not in the contact were set to zero (black).

To determine the extent of clustering of existing vs. accumulation of new EcadGFP to the developing plaque area, measurements of the maximum fluorescence intensity and the sum of fluorescence intensities of all pixels in an area of fixed size were monitored. For analysis, time-lapse images were passed through a 3 x 3 convolution filter. A region of 20 µm² was placed over an area where multiple small puncta were coalescing into a plaque at the edge of a developing cell–cell contact. The maximum and integrated intensities were measured using MetaMorph during the time-lapse series.

Photobleach Recovery
Method.
Photobleaching experiments were performed on the scanning laser confocal microscope described above, and with EcadGFP cells prepared as for time-lapse experiments. The Smith Mark IV microscope incorporates a digitally controlled acousto-optic shutter (NEOS, Melbourne, FL). In conjunction with appropriate interface electronics and control software, this high-speed shutter allows the scanning laser beam to be turned on or off anywhere within the imaging area on a pixel-by-pixel basis. For the present experiments, the shutter was used for controlled photobleaching of circular areas within the scanning raster pattern. Typically, circular spots of a 120 pixel radius were generated with a 380 x 240 pixel scan raster. The actual spot radius at the specimen was varied by adjusting pixel sizes within the range of 0.01–0.05 µm. Scan geometries and beam intensities were varied independently as necessary during successive baseline acquisition, photobleaching, and recovery acquisition episodes. Scanning frame rates were varied to suit the needs of individual experiments between 0.33 and 3 Hz. Photobleaching energies were adjusted by varying exposure intensities and durations to achieve depths of photobleaching ranging from 30 to 70%. Calibrations of the exact position and dimensions of the photobleached area were collected using fixed specimens and aminopropysilane coverslips (Sigma Chemical Co.) coated with fluorescein-isothiocyanate (data not shown). The region of the cell to be photobleached was controlled by adjusting the x, y stage on the microscope during low-power imaging until the desired target area on the cell was in the middle of the video monitor.

Analysis.
Digital images from the photobleaching experiments were imported into MetaMorph and passed through a 3 x 3 low-pass convolution filter. Average fluorescence intensity data from the bleached regions, two non-cell regions, and two non-bleached cell regions were collected for all frames. Data were imported into Excel and corrected for the minor photobleaching caused by the fluorescence recording process by normalization to a region outside the main photobleach pulse spot. The photobleaching recovery part of the data was then imported into Igor Pro, and the characteristic diffusion time (t_d) was calculated using equations for diffusion into a circular disk (Axelrod et al., 1976) and modified for total recovery (Soumpasis, 1983; equation 16). The diffusion coefficient (D) was calculated according to the equation

where r is the radius of the bleached region. Mobile fraction constants were calculated according to equation in Axelrod (1976; equation 9) at indicated times after recovery. Measurements of the slower photobleach recovery timecourses were subject to potential errors due to gradual redistribution motions of puncta and plaques during recovery phases. TIP scans of photobleach recoveries were generated (see Fig. 9) and used to reject data from any experiment where such errors might have been significant.

View larger version (94K): [in this window] [in a new window] [Download PPT slide]   Figure 9 Mobility of E-cadherin puncta within the cell–cell contact interface. Fig. 9 shows a TIP scan of an entire contact during a photobleach-recovery experiment. A newly developing plaque in a 1.5-h-old contact was photobleached with a 2.8-µm-diameter bleach circle (0 mins, 0 µm) on the TIP scan (arrow). Images were collected every 16 s for 24 min at 0.11 µm/pixel. The fluorescence intensity scale bar ranges from 0–255 units divided into 15 colors.

	Results

Top Abstract Materials and Methods Results Discussion References

View larger version (37K): [in this window] [in a new window] [Download PPT slide]   Figure 1 EcadGFP has properties similar to those of endogenous E-cadherin. (A and B) Catenins were coimmunoprecipitated with EcadGFP from HEK 293 cells transiently transfected with EcadGFP (A) or stably transfected MDCK cells (B). Cells were labeled with [35S]Met/Cys for 24 h before immunoprecipitation. (A) Proteins in cadherin immunoprecipitates are compared among mock-transfected HEK 293 cells (No DNA), HEK 293 cells transfected with canine E-cadherin (Ecad), and HEK 293 cells transfected with EcadGFP, and (B) between untransfected MDCK cells (No DNA) and MDCK cells stably transfected with EcadGFP. (C) EcadGFP fluorescence and β-catenin immunofluorescence from HEK 293 cells transiently transfected with EcadGFP. Bar, 30 µm. (D) Preferential delivery of newly synthesized EcadGFP to the basal-lateral plasma membrane of fully polarized MDCK cells stably transfected with EcadGFP, or of endogenous E-cadherin in untransfected cells (No DNA); A, apical membrane; B, basal-lateral membrane. (E) Phase contrast images of L-cells and L-cells expressing EcadGFP after 18 h of aggregation in suspension culture in the presence or absence of extracellular Ca2+. Bar, 60 µm.

EcadGFP Distribution Changes During Cell–Cell Adhesion
EcadGFP expressing MDCK cells were imaged for 10 h to observe the dynamics of the localization of EcadGFP during formation of contacts between single cells, and during formation of small multicell colonies (Fig. 2). Expression of EcadGFP in single cells was relatively uniform over the plasma membrane with some increased intensity in a circumferential ring at the cell periphery (Fig. 2, A and B, 0 h). During the formation of cell–cell contacts between two (Fig. 2 A) or three (Fig. 2 B) cells, or single cells and larger cell clusters (Fig. 2 C), EcadGFP fluorescence became significantly more intense at the cell–cell contact during the first 2 h. After at least 2 h, the largest and brightest regions of EcadGFP fluorescence were at the edges of cell–cell contacts; we call these structures plaques (Fig. 2, circles). The fluorescence intensity of EcadGFP plaques was 6–10 times greater than that of EcadGFP in noncontacting membranes, and 2–4 times greater than that of EcadGFP in areas of the membrane in the middle of the contact (Fig. 2, A–C; 8 h).

View larger version (47K): [in this window] [in a new window] [Download PPT slide]   Figure 2 Distribution of EcadGFP during monolayer formation. A single confocal image was collected from EcadGFP expressing cells every 10 min for 12 h at 0.12 µm/pixel at 12 sites. Five representative images from each time-lapse are shown. Elapsed time is indicated on top of each column in h (0, 2, 4, 6, and 8, respectively). The circles in A highlight the edges of a cell–cell contact that have developed large aggregates of EcadGFP plaques. The arrows in B–F, columns 0 or 2 h point to the well-separated plaques at the edges of developing cell–cell contacts that reorganize into a multicellular vertex by 8 h. Note that the 0-h designation is arbitrarily set as the first time point shown. Bar, 15 µm.

To better understand the evolution of these distinct patterns of E-cadherin, the distribution of EcadGFP during development of cell–cell contacts was examined in multisite time-lapse confocal images taken over the course of 3 h (we initially focused on the formation of puncta and plaques during the first two stages of adhesion see below). The cells were then fixed and stained with phalloidin, (which labeled F-actin) and mAb 3G8 (which recognized the extracellular domain of endogenous E-cadherin and EcadGFP), and were imaged. Fig. 3 shows representative contacts from one time-lapse recording. Column 1 of Fig. 3 A shows the formation of a contact between two cells over 71 min. During cell–cell adhesion, EcadGFP fluorescence appeared at cell–cell contacts, and then increased in intensity with time and as the contact lengthened. The distribution of EcadGFP and endogenous E-cadherin were recorded retrospectively with mAb 3G8 (Fig. 3, 1C), showing that both proteins had coincident distributions as expected (see Fig. 1). Retrospective actin staining (Fig. 3, 1B) shows that circumferential actin cables were organized parallel to the cell–cell contact interface at this time.

View larger version (26K): [in this window] [in a new window] [Download PPT slide]   Figure 3 Redistribution of EcadGFP during cell–cell contact occurs in two stages and correlates with reorganization of the actin cytoskeleton. Three 0.3-µm z-sections were collected from EcadGFP-expressing cells every 4.2 min at 0.5 micron/pixel at 6 sites. (A) Combined stacks from two sites are shown (Contact 1 and Contact 2). The age of each contact is displayed in min; the zero time point defines when a stable cell–cell contact had formed. Bar, 12 µm. The arrow points to a circumferential pattern of EcadGFP observed in single cells. (B and C) Immunofluorescence of the same cells stained, after formaldehyde fixation, with rhodamine phalloidin (actin; B) and mAb 3G8/CY5 (E-cadherin; C). (D) EcadGFP organization shown in C is plotted as a function of fluorescence intensity (y axis, arbitrary fluorescence units) vs. position in the contact (x axis, µm). The dashed gray lines running between C and D approximately register the edges of the contact in the image and graph. (E and F) Triton X-100 extracted wild-type MDCK cells (i.e., without EcadGFP) that had formed a contact for <1 h (Contact 1) or >2 h (Contact 2) stained with FITC-phalloidin and mAb 3G8/CY5, respectively. Bar, 10 µm.

Fig. 3, E and F shows staining of actin and E-cadherin, respectively, in wild-type MDCK cells (i.e., not expressing EcadGFP) at a contact that is <1 h old (Fig. 3, 1E and 1F), and at a contact that is >2 h old (Fig. 3, 2E and 2F). The distributions of actin and endogenous E-cadherin in cell– cell contacts are very similar to those of actin and Ecad-GFP shown in the contacts in Fig. 3, B and C, respectively, supporting the general conclusion that EcadGFP is fully functional. Furthermore, during cell–cell adhesion, changes in the organization/distribution of EcadGFP are very similar, if not identical, to those of endogenous E-cadherin.

EcadGFP Clusters into Puncta Upon Cell–Cell Contact
Our observation that EcadGFP initially aggregated into puncta during formation of cell–cell contacts is similar to our previous observations in which E-cadherin distributions were determined retrospectively by immunolabeling cells after time-lapse DIC imaging (Adams et al., 1996). However, in that previous study we could not determine the relationship of Triton X-100 insolubility and clustering of E-cadherin, the source of E-cadherin in puncta, the role of the actin cytoskeleton in the spatial organization of puncta, or the dynamics and fate of puncta within the contact. Using EcadGFP, we were able to address these critical problems directly.

The reorganization of EcadGFP during cell–cell adhesion was examined quantitatively by measuring EcadGFP fluorescence intensity over time after initial contact between cells. The graphs in Fig. 3 D show quantitative representations of the maximum E-cadherin fluorescence intensity vs. position in the contacts shown in Fig. 3 C. The younger contact shows multiple small peaks of fluorescence along the contact, each corresponding to a punctum (Fig. 3, column 1D). In contrast, the older contact shows two pronounced peaks of fluorescence intensity at the edge of the contact, each corresponding to an EcadGFP plaque (Fig. 3, column 2D). In addition, there were multiple smaller peaks between the plaques that represented residual puncta within the cell–cell contact.

To gain information about the genesis, lifetime, and position of EcadGFP during initiation of contact formation, a single field of EcadGFP-expressing cells was imaged rapidly at high resolution. Fig. 4 A shows representative images from one of these time-lapse recordings. An arrow follows the position of a bright EcadGFP fluorescent punctum at the cell–cell interface. To provide an objective nonbiased format for the quantitative representation of dynamic data like that illustrated in Fig. 4 A, we developed the type of representation shown in Fig. 4 B. The fluorescence intensity profiles along the length of the contact (e.g., Fig. 3 D) were color-coded and combined for each time-lapse frame to provide a color map of EcadGFP intensity distribution along the length of the contact as the contact lengthened. We term such graphs TIP scans. By providing a clear representation of time-dependent changes in EcadGFP fluorescence along the cell–cell contact interface, TIP scans make it relatively easy to discern the organization of EcadGFP during contact formation. Background fluorescence in the TIP scan is contributed by overlapping regions of plasma membrane. Areas of the contact that are brighter than the background cell fluorescence correspond to brighter clusters of EcadGFP. The TIP scan in Fig. 4 B shows that the contact in Fig. 4 A grew to a length of 12 µm in 20 min. The contact then grew more slowly to reach a length of 35 µm after 90 min.

View larger version (74K): [in this window] [in a new window] [Download PPT slide]   Figure 4 EcadGFP puncta are formed and stabilized along newly formed cell–cell contacts. (A) 16 time-lapse images of EcadGFP cells taken from a sequence recorded every 1 min for 100 min at 0.11 µm/pixel; time in min after formation of the contact is shown. Arrows follow a single punctum. (B) TIP scan of all of the time-lapse images from the experiment represented in A. The contact was divided into 129 0.22-µm sections, and the maximum fluorescence intensity at 100 time points was collected for a total of 12,900 data points. The contact originates at 0 min and 0 µm. Small fluctuations in the apparent intensity of stable puncta are near the limits of instrumental noise sources such as laser output fluctuations and noise processes in the photomultiplier tube detector. (C) Double immunofluorescence of the same contact extracted with Triton X-100, fixed with formaldehyde, and stained with rhodamine phalloidin and E-cadherin mAb 3G8/CY5. The arrows in all panels point to the same punctum. Bars: (A) 10 µm; (C) 5 µm. (B) 0–210 gray scale fluorescence intensity units divided into 15 colors.

As the contact lengthened, new EcadGFP puncta appeared sequentially such that the number of puncta remained constant with respect to the length of the contact (1 punctum per 1.5 µm contact length; see also Adams et al., 1996). These new puncta also appeared de novo and gradually increased in intensity with time (Fig. 4 B). Many of these puncta were spatially stable in the contact interface over time, while some gradually changed position towards the edges of the contact. In general, the fluorescence intensity of puncta was brightest in the oldest part of the contact near the site of initial cell–cell contact (marked as 0 µm), and dimmest at the perimeter of the lengthening.

The EcadGFP puncta evident in these time-lapse sequences were Triton X-100–insoluble. Each EcadGFP punctum observed in the final frames of the time-lapse sequences colocalized with the brightest Triton X-100–insoluble E-cadherin puncta (compare similarly oriented images in Fig. 4 A, 90'; and Fig. 4 C). EcadGFP puncta were associated with thin cables of actin filaments that emerged from circumferential actin cables oriented parallel to the contact (Fig. 4 C). In summary, while we have confirmed our early observation that cell adhesion initiates the formation of E-cadherin puncta, the data presented here demonstrate that a diffuse pool of E-cadherin clusters into puncta in response to cell–cell contact, and that those puncta organize and become stabilized around actin filaments located close to the contacting membranes.

Formation of EcadGFP Plaques at the Contact Margins and Reorganization of the Actin Cytoskeleton
Next, we asked how early E-cadherin puncta are reorganized with actin to further strengthen cell–cell adhesion, and then to cause condensation of cells into multicell colonies. Over longer times (>2 h), EcadGFP and endogenous E-cadherin became organized into large plaques at the margins of the contact (Fig. 2 and Fig. 3, column 2). Fig. 5 A shows representative images from a longer time-lapse experiment. After 1.5 h, two regions of increasing EcadGFP fluorescence appeared and migrated out with the edges of the contact at velocities of up to 0.5 µm/min (Fig. 5). These regions gradually gained up to 10x the average punctum fluorescence intensity over the course of 1 h, in contrast to a punctum that reached maximum fluorescence intensity within 30 min of formation. Approximately 2.5 h after contact nucleation, EcadGFP was heavily concentrated in discrete fluorescent plaques at the margins of the contact (Fig. 5, A and B, arrows). The region of the contact between the two plaques retained a thin line of EcadGFP intensity (compare to Fig. 3, column 2B). Comparison of the last live EcadGFP images with the retrospective immunofluorescence of E-cadherin and actin (Fig. 5 C) shows that EcadGFP plaques were resistant to extraction with Triton X-100, and were sites at which circumferential actin cables terminated.

View larger version (46K): [in this window] [in a new window] [Download PPT slide]   Figure 5 Two large plaques of EcadGFP form and move to the edges of the cell–cell contact. (A) 16 time-lapse images of EcadGFP cells taken from a sequence recorded every 2 min for 2.7 h at 0.23 µm/pixel. Arrows follow a single plaque. (B) TIP scan of all of the time-lapse images from the experiment represented in A. The contact was divided into 101, 0.46-µm sections, and the fluorescence intensity at 85 time points was collected for a total of 8,585 data points. The contact originates at 0 min and 0 µm. Note that the TIP scan at this reduced resolution shows a relatively homogeneous distribution of EcadGFP within the contact during the first hour, whereas the TIP scan at a higher resolution revealed individual punctum (see Fig. 4). (C) Double immunofluorescence of the same contact stained with rhodamine phalloidin and E-cadherin mAb 3G8/CY5. The arrows in all panels point to the same plaque. (B) 0– 151 gray scale fluorescence intensity units divided into 15 colors. Bars: (A) 10 µm and (C) 5 µm.

View larger version (26K): [in this window] [in a new window] [Download PPT slide]   Figure 6 EcadGFP puncta cluster into plaques during transition between early and late stages of adhesion. (A) Representative images of a time-lapse sequence taken at 0.8 Hz for 300 s at 0.12 µm/pixel in a region of the cell–cell contact in which a plaque is developing. Time is in min; arrows point to individual puncta; bar, 2 µm. (B) Quantitative fluorescence intensities of EcadGFP. The average (gray circles) and maximum (black diamonds) intensities in a 20-µm2 region surrounding a developing plaque area are plotted over a 40-min period. The average fluorescence intensity of the same number of EcadGFP in a fixed area is constant regardless of its distribution. However, the maximum fluorescence intensity increases as EcadGFP clusters into a smaller region within that fixed area. For details, see Materials and Methods.

View larger version (24K): [in this window] [in a new window] [Download PPT slide]   Figure 7 Cytochalasin D selectively disassembles new cell–cell contacts. Representative images of a time-lapse sequence taken at 1 frame/2 min for 2 h at 0.4 µm/pixel before and after adding 2 µM CD. (A) 14 min before CD; (B) 1 min before CD; (C) 30 min after CD; (D) 60 min after CD. Immunofluorescence of the same area is shown using rhodamine phalloidin (E) or β-catenin/ CY5 (F). The arrows in D–F point to CD-induced EcadGFP clusters; bar, 10 µm. (G) The number of cells that were in contact before the time-lapse experiment began (>60 min old), and those that made contact during the imaging experiment (<60 min old) were counted and the totals shown for three independent experiments (black bars). The number of those contacts that disassembled within 1 h after CD treatment was determined (striped bars). The percentage of cell–cell contacts disassembled by CD treatment is 14% for old contacts and 73% for new contacts.

View larger version (64K): [in this window] [in a new window] [Download PPT slide]   Figure 8 Photobleach-recovery analysis shows a highly mobile pool of EcadGFP coalesces into immobile puncta. A shows a live cell before and after photobleaching. The box indicates where the cell was photobleached. The arrow points to an area that formed a contact during the photobleach. The cells were fixed in formaldehyde and stained with phalloidin and mAb 3G8. B shows the fluorescence recovery curves of a single noncontacting cell in which half of the cell was photobleached (blue). EcadGFP fluorescence of the nonbleached region (red) and the entire cell (green) was monitored during recovery. Notice that the EcadGFP fluorescence values equalize in the photobleached and nonphotobleached areas. C shows the first 3 min of photobleach-recovery data of noncontacting membrane regions photobleached with a 5.8-µm (pink) and 3-µm (black) circle. The relative fluorescence is scaled between the fluorescence intensity just after bleaching and equilibrium. The lines show the theoretical recovery curves for each region with a diffusion coefficient of 3 x 10–10 cm2/s. Note that the smaller photobleach circle (black line) recovers more quickly. D shows images taken before, 0.1 min after, and 10 min after photobleaching a 5.8-µm-diameter circle in a region of membrane not involved in cell–cell contact (Membrane), a region of membrane in a <15-min-old contact (New contact), a region of membrane in the middle of a <60-min-old contact (Puncta), and a membrane at the edge of a >2-h-old contact (Plaque). For each experiment, 300 images were collected every 3.2 s at 0.11 µm/pixel. The circles mark the photobleach region, and the colors correspond to the recovery curves shown in C (pink) and E. E shows photobleach-recovery data for the bleached contact regions identified in D. The relative fluorescence is scaled to the pre-bleach intensity value. The mobile fraction of EcadGFP in the new contact (blue), puncta (green), and plaque (orange) is 100, 50, and <10%, respectively.

Fig. 9 shows a TIP scan from an experiment where a 2.8-µm–diameter area was bleached in a 1-h-old contact containing two very bright EcadGFP puncta (Fig. 9, arrow). It is clear that bleached puncta partially recovered their fluorescence, concomitant with a partial loss of fluorescence in nonbleached puncta. However, it is also obvious that individual puncta undergo gradual redistributions during these slow recoveries. As the EcadGFP fluorescence recovered, the bleached puncta migrated out of the bleached area, and puncta adjacent to the original bleached area slowly migrated into the bleached area (Fig. 9).

Detailed TIP scan analyses of photobleach recovery data allowed us to make the following generalizations about the dynamics of EcadGFP: (a) puncta move as individual units within the cell–cell contact interface during contact expansion; (b) puncta on the edge of a fluorescence bleach area recovered their fluorescence first; and (c) adjacent nonbleached puncta sometimes exhibited a decrease in fluorescence intensity. These data indicate that the immobilized fraction of EcadGFP is associated with the cytoskeleton, and that cytoskeletal associated EcadGFP moves and exchanges within the cell–cell contact interface.

	Discussion

Top Abstract Materials and Methods Results Discussion References

 
E-cadherin plays important roles in cell–cell recognition and adhesion. However, the dynamics of E-cadherin redistribution during the processes of initial cell–cell contact through development of a polarized monolayer are unknown. It is thought that clustering of E-cadherin (Yap et al., 1998) via extracellular homotypic binding (Nose et al., 1988) is sufficient for initial cell–cell interactions: the role of cadherin clusters during the development of older cell colonies is less understood. Furthermore, the dynamic interactions of E-cadherin with intracellular proteins, especially the cytoskeleton, has not been described. By quantitatively analyzing EcadGFP redistribution and mobility in epithelial cells during adhesion development, we provide a new dynamic view of how E-cadherin and the actin cytoskeleton establish strong cell–cell adhesion.

GFP was attached to the COOH terminus of the E-cadherin cytoplasmic domain. The cytoplasmic domain of cadherin binds to catenins, which are required for binding the cadherin/catenin complex to the actin cytoskeleton. Therefore, it was necessary to show that EcadGFP protein has functions identical to those of endogenous E-cadherin. We showed that EcadGFP is fully functional by the following criteria: EcadGFP formed a 1:1:1 stoichiometric complex with - and β-catenin; EcadGFP precisely colocalized with catenins at cell–cell contacts; EcadGFP was targeted directly from the Golgi to the basal-lateral membrane in polarized MDCK cells; EcadGFP localized to cell–cell contacts and entered a Triton X-100–insoluble pool of proteins only at cell–cell contacts; and EcadGFP induced Ca²⁺-dependent cell–cell adhesion and condensation in transfected MDCK cells and nonadherent cells (fibroblasts) with kinetics that were qualitatively and quantitatively similar to those of endogenous E-cadherin (Fig. 1). In addition, the kinetics of assembly of EcadGFP puncta in live cells was similar to that measured for the assembly of Triton X-100–insoluble E-cadherin puncta by retrospective immunocytochemistry (Adams et al., 1996), and the formation of cell–cell contacts between MDCK cells expressing EcadGFP and wild-type MDCK cells appeared very similar (compare to Adams et al., 1996). Therefore, the fact that EcadGFP has functions identical to those of endogenous E-cadherin implies that EcadGFP can substitute for endogenous E-cadherin, and that the process of cell–cell adhesion is normal even when levels of endogenous E-cadherin are reduced in the presence of EcadGFP (Fig. 1).

We present evidence for three sequential stages of cell– cell adhesion that involve specific changes in E-cadherin and actin cytoskeleton organization. These stages are: (I) clustering of mobile E-cadherin into immobile puncta along the length of the forming contact; (II) reorganization of E-cadherin puncta into plaques at the edges of the contact; and (III) coalescence of E-cadherin plaques to the vertices of contacts among three or more cells.

Stage I
In the first stage of adhesion, E-cadherin spontaneously clusters into puncta at initial sites of developing cell–cell contacts. The formation of E-cadherin puncta results in decreased E-cadherin mobility. In new areas of cell–cell contact (<15 min old), EcadGFP has a high mobile fraction (>90%) and a high diffusion coefficient (3.6 ± 1.5 x 10^–10 cm²/s). However, where EcadGFP clusters into puncta and associates with the cytoskeleton, a smaller fraction is mobile (<50% within a 26-µm area). The puncta formed by EcadGFP are very similar in organization and distribution to structures formed by endogenous E-cadherin and catenins that were previously characterized by retrospective immunocytochemistry (Adams et al., 1996; see also Fig. 3).

Weak interactions between extracellular and juxtamembrane domains of cadherins may be sufficient to initiate clustering of the protein in the membrane (Yap et al., 1998). However, interactions between E-cadherin and the actin cytoskeleton are initiated quickly upon cell–cell contact, and these interactions affect the organization of the adhesion complex. We showed that as E-cadherin puncta begin to form during this first stage, they always appear to be associated with the ends of thin actin cables that are oriented toward the contact (Fig. 4). These actin filaments branch from circumferential actin cables that are organized parallel to the forming contact and circumscribe the perimeter of single cells. We speculate that binding actin filaments to E-cadherin/catenin complexes may cause further clustering and stabilization of puncta. This type of cadherin/actin organization has been shown to provide a mechanical linkage between fibroblasts (Ragsdale et al., 1997). Quantitative measurements showed that this initial stage of adhesion coincides with an exponential increase in the strength of adhesion (Angres et al., 1996). Significantly, this strengthening stage was completely inhibited by treatment of cells with CD (Angres et al., 1996; Fig. 7). In the present study we showed that during this initial stage, CD selectively disassembled contacts and caused formation of aggregates that include cell-surface EcadGFP (this study) and probably the barbed ends of actin filaments (Verkhovsky et al., 1997). It is also interesting to note that myosin is involved in the CD-induced aggregation of the barbed ends of actin filaments (Verkhovsky et al., 1997), and that actin treadmilling ceases in areas of developing cell–cell contacts (Gloushankova et al., 1997). We speculate that E-cadherin puncta gradually sequester the barbed ends of actin filaments, and directly or indirectly anchor them to the membrane at cell–cell contacts, resulting in the gradual strengthening of cell–cell adhesion. These changes in actin organization may also set up cytoarchitectural cues for stage II of adhesion.

Stage II
The second stage of contact formation is distinguished by the gradual emergence of much larger E-cadherin clusters that we designate as plaques (Figs. 2, 5, and 6). Generally, one plaque is observed to form at either end of the developing contact where the reorganized circumferential actin cables terminate. Identical plaques were formed by endogenous E-cadherin in compacted MDCK cell–cell contacts (Fig. 3). Using EcadGFP, we showed that these plaques arose by lateral clustering of a subset of puncta that were formed during the first stage of adhesion and the continual immobilization of a mobile pool of EcadGFP (Fig. 9). Plaque formation resulted in a further decrease in the mobile fraction of E-cadherin to <10%. Using TIP scans, we found that during plaque formation, EcadGFP puncta traveled within the cell–cell contact interface to the margins of the contacts at velocities of up to 0.5 µm/min (Fig. 5), which is similar to the velocity of translocation of ConA beads along cell–cell contacts (Gloushankova et al., 1997).

During the second stage of contact formation, circumferential actin cables rearrange from a parallel to a perpendicular orientation with respect to the cell–cell contact (Fig. 5). The reorganization of actin appears to be different in MDCK cells and fibroblasts (Yonemura et al., 1995), which raises the possibility that the strength of the interactions between E-cadherin and actin might be responsible for specific differences in actin dynamics between these two cell types. A consequence of the reorganization of E-cadherin and the actin cytoskeleton is the compaction of contacting cells, which is a clear sign of the establishment of strong cell–cell adhesion and cells maximizing the contacting surfaces between them (Fig. 2). We showed that these compacted cell–cell contacts are resistant to disassembly by CD, indicating either that these contacts have become mechanically resistant to depolymerization of actin, or the barbed ends of the circumferential actin cables are firmly embedded within E-cadherin plaques and are no longer accessible to CD.

Stage III
E-cadherin plaque formation coordinates circumferential actin cables to cell–cell contact and maximizes the area of membrane involved in the contact between two cells. When more than two cells form contacts, E-cadherin plaques and actin cables continue to reorganize to form a more compact cell colony (Fig. 2). This process of condensation involves movement of E-cadherin plaques towards each other until they have coalesced to form a vertex among three or more cells (Fig. 2, C and D). We suggest that E-cadherin plaques are cinched together by contraction of the actin cables that coordinate the plaques within the multicell colony. This reorganization of E-cadherin changed the shape of cells from rather cubiodal after plaque formation to cone-shaped after cell condensation in colonies; the vertices of E-cadherin were located at the tip of the cone (Fig. 2, B–D). The formation of cone-shaped cells is redolent of the effects of purse-string contraction of circumferential actin filaments during wound healing in vitro (Bement et al., 1993) and in vivo (Martin and Lewis, 1992; Brock et al., 1996). Previous studies have shown that contraction of cell monolayers around a wound coincides with reorganization of actin cables, myosin II, tropomyosin, and other actin-associated proteins on the membrane adjacent to the wound. It is assumed that actin– myosin contraction pulls on membranes at the edge of the wound to close the opening between cells. We suggest that circumferential actin contraction initiated during cell–cell compaction continues until an equilibrium, perhaps equal tension, is reached between the circumferential actin bundles throughout the forming multicell colony. Studies are underway to test the roles of actin and actin contractility in this stage of adhesion and cell reorganization.

A Model
In summary, we suggest a model for how contacts between cells are initiated, strengthened, compacted, and condensed as cells transform from the migratory phenotype of a single cell to a sedentary phenotype of one cell in a multicell monolayer. Cell–cell adhesion is initiated by weak binding between extracellular domains of E-cadherin that are present in a highly mobile pool at the plasma membrane. At or near the same time, E-cadherin/catenin complexes attach to actin filaments that branch from actin cables that circumscribe the perimeter of migratory cells. These two processes act synergistically to assemble puncta, which, as a group, are sufficiently adhesive to hold the nascent cell–cell contact together (Fig. 10, stage I). Subsequently, there is a change in actin dynamics as actin treadmilling ceases in areas of cell–cell contact, perhaps due to sequestration of the barbed ends of actin filaments into E-cadherin puncta. We hypothesize that reduced actin treadmilling causes the dissolution of the circumferential actin cables immediately adjacent to the developing contact. It is also possible that a signaling event at the cell surface induced by cell–cell adhesion causes a change in the organization or polymerized state of the circumferential actin cables adjacent to the contact site. We suggest that stabilization of actin via the clustered cadherin/catenin complex engages the myosin II clutch (Suter et al., 1998), thereby inducing translocation of circumferential actin cables and the rest of the cell body to the cell–cell contact interface and the rapid movement of associated E-cadherin puncta into large plaques. This coordinated reorganization of E-cadherin and the actin cytoskeleton results in the establishment of strong compacted cell–cell contacts and the generation of an actin cable that circumscribes the free edges of the newly contacting cells and is embedded into either side of a E-cadherin plaque at the margins of the contact (Fig. 10, stage II).

View larger version (39K): [in this window] [in a new window] [Download PPT slide]   Figure 10 A three-stage model for cell–cell adhesion and colony formation. Stage I: multiple E-cadherin puncta form along the developing contact and loosly hold contacting cells together. A circumferential actin cable (thick red line) surrounds isolated cells. As cells adhere, E-cadherin clusters into puncta within the cell–cell contact interface (blue circle) and rapidly associates with thin actin bundles and filaments (thin red lines). As the contact lengthens, puncta continue to develop along the length of the contact at a constant average density during the first 2 h. Stage II: E-cadherin plaques develop at the edges of the contact which compact and strengthen cell–cell interactions. Stabilization of actin filaments by E-cadherin puncta within the cell–cell contact results in gradual dissolution of the circumferential actin cable behind the developing contact and insertion of the circumferential actin cables into the cell–cell contact accompanied by additional clustering of E-cadherin puncta into E-cadherin plaques (green ovals). Between cell plaques, E-cadherin is more diffusely distributed (green line) and associates with actin filaments oriented along the axis of the cell–cell contact (red line). Stage III: E-cadherin plaques cinch together to form multicellular vertices, further condensing cell colonies. In multicellular colonies, contractility within the circumferential actin cable brings E-cadherin plaques from adjacent cells together. Dynamics within the cytoskeleton result in continual rearrangement.

Additional cells systematically join a multicellular colony as described above. E-cadherin and actin in newly forming cell–cell contacts reorganize, and then cinch together adjacent plaques attached to actin cables at the free cell edges (Fig. 2, D and E). Eventually some cells will become engulfed by a multicellular colony such that there is no longer a circumferential actin cable, and the cell is surrounded by contacts on all sides. There still exist larger clusters of E-cadherin at the vertices of cells within the multicellular colony, and the cells are probably still able to exert tension on each other through these regions to maximize the contact area. The organization of actin in the multicellular colony is now distinctly different from that in a cell–cell contact that is <1 h old, and is characterized by thin actin bundles running parallel to the cell–cell contact in association with E-cadherin (see Fig. 3). These E-cadherin and actin organizations are also characteristic of the mature monolayer.

Our findings reveal a very tight coordination among cadherin binding, aggregation, adhesion events, and dramatic reorganizations of the actin cytoskeleton. These reorganizations occur in parallel with transitions from weak initial adhesion to strong adhesions associated with cell– cell compactions, condensations of cells into colonies, and formation of a belt of cadherin and actin as observed in mature monolayers. These results provide a new framework for future studies aimed at identifying the effects of other regulatory molecules (e.g., GTPases, kinases, and cytoskeletal motors) and cadherin-associated proteins (e.g., p120^CAS) on these distinct adhesive stages of cell–cell adhesion.

	Acknowledgments

 
We thank members of the Nelson and Smith labs and Chris Hazuka for their comments during the course of this work, Dr. Richard Lewis for help programming diffusion solutions in Igor Pro, Dr. Lee Rubin (University College London) for the canine E-cadherin cDNA clone, and Dr. Brian Seed (Harvard Medical School) for the generous gift of humanized codon-preference-adjusted red-shifted GFP before its publication. We also thank Dr. David Loftus for his help with the aggregation assay.

Submitted: 13 February 1998

Revised: 1 June 1998

W.J. Nelson and S.J Smith were supported by grants from the National Institutes of Health and The Mathers Charitable Foundation. Y.-T. Chen is a recipient of a postdoctoral fellowship award from National Kidney Foundation.

The current address of Cynthia L. Adams is Cytokinetics, Inc., 280 East Grand Ave., South San Francisco, CA 94040.

Address all correspondence to Stephen J Smith, Department of Molecular and Cellular Physiology, Beckman Center, Stanford University School of Medicine, Stanford, CA 94305-5345. Tel.: 650-723-6799; Fax: 650-498-5286; E-mail: sjsmith{at}leland.stanford.edu

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