A decrease in the Triton-insoluble fraction of desmoglein in DPNTP-expressing cells is accompanied by reductions in intercellular adhesion and cell sheet integrity

As described above, DPNTP did not detectably alter the cell surface expression of classic or desmosomal cadherins. However, as IFs are uncoupled from these adhesive structures, we predicted that the loss of IF attachment might alter the amount of Dsg 2 in the IF-associated Triton-insoluble pool. To test this possibility, untreated or treated A431/SS, C2, and G4 lines were lysed in 1% Triton X-100 buffer and the insoluble pellet was analyzed by SDS-PAGE (Fig. 3 D). Both the A431/C2 and G4 lines exhibited an approximately twofold decrease in the Triton-insoluble fraction, whereas no change was detected in the solubility of E-cadherin.

We next sought to test whether this desmosomal cadherin-specific alteration in solubility translated into a functional difference in intercellular adhesion and epithelial sheet integrity. We first employed an adaptation of a previously described hanging drop aggregation assay using EDTA-dissociated single cells to generate clusters of cells not adherent to the substrate (Thoreson et al., 2000). Cells allowed to aggregate for at least 6 h in a hanging drop on the underside of a culture dish were subjected to trituration through a 200-μl Gilson pipette tip in an attempt to disrupt intercellular adhesion. The degree of dissociation was quantified by counting the particles that dissociated from the original cluster. Cells induced with Dox to express DPNTP were more susceptible to dissociation than untreated cells (Fig. 4 A) and exhibited on average an approximately fourfold increase in particle number when compared with the untreated cells after trituration (Fig. 4 B). Dox-treated and untreated A431/SS cells did not show any differences in the degree of dissociation, which was comparable to untreated C2 cells.

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Figure 4.

DPNTP weakens adhesion in cell clusters formed in hanging drop culture. (A) 1 × 10³ cells that had previously been Dox induced for at least 24 h or left untreated were seeded into hanging drop cultures without (top left) or with Dox-containing (top right) media and allowed to aggregate for 6 h. After trituration by passing the cell cluster 30 times through a 200-μl pipette tip, the degree of dissociation of the cell cluster was visualized by microscopy (A) and quantified by manual counting in a dissecting microscope (B). Note that cell clusters were imaged while suspended in a hanging drop and that dark patterns at the edge of field are caused by refraction of light. Due to cropping of the images, some of the dissociated small particles at the periphery in the bottom right panel of A are not visible but are still reflected in the graph in B. Error bars represent the standard deviation from an experiment in which each condition was tested in triplicate.

Because cell clusters grown in suspension were not well suited for analysis by immunofluorescence microscopy to monitor desmosome status, we sought to use an assay in which we could both verify formation of intercellular junctions and test adhesive strength. Confluent monolayers of Dox-induced and uninduced A431/SS and C2 cells were harvested from tissue culture dishes by incubation with dispase as previously described (Calautti et al., 1998). Monolayers were then transferred to 15-ml conical tubes containing 5 ml of Dulbecco's PBS (containing 0.9 mM calcium chloride and 0.5 mM magnesium chloride [DPBS]). After inverting the tubes 50 times on a rocker, monolayer fragments were counted (Fig. 5 A). DPNTP-expressing monolayers dissociated into numerous smaller fragments, whereas both Dox-treated and untreated control cell lines and uninduced A431 monolayers exhibited only minimal dissociation. Quantification of the number of fragments showed a 23-fold (t test, P < 0.001) and fourfold (P < 0.005) increase in fragmentation by the DPNTP-expressing A431/C2 and G4 monolayers, respectively (Fig. 5 B).

DPNTP-dependent dissociation of cell sheets is not accompanied by lactate dehydrogenase (LDH) release

To address whether the observed increase in fragmentation in DPNTP-expressing monolayers was due to increased cellular fragility and cytolysis, we assessed cellular damage by measuring the release of lactate dehydrogenase (LDH) from monolayers after application of mechanical stress. LDH release measured from DPNTP-expressing monolayers after mechanical disruption was not statistically altered compared with that released from the uninduced and control cell lines, which had minimal fragmentation (Fig. 5 C). Thus, despite a dramatically increased degree of monolayer dissociation, the DPNTP-expressing cell lines did not exhibit a corresponding increase in LDH release.

Time-lapse microscopy of living cells reveals that DPNTP-expressing cells dissociate more rapidly and with less cell distortion under stress than control cells

To examine at higher resolution the behavior of IF-uncoupled cells undergoing stress within epithelial cell sheets, time-lapse imaging of living cells was performed. Tet-On cell lines expressing DPNTP COOH-terminally tagged with EGFP were generated to confirm that DPNTP was being expressed and incorporated into cell junctions specifically in the cells under observation. These cell lines behave similarly in all respects to the DPNTP–FLAG-expressing cells. One cell line, termed A431/I3, which homogeneously expressed DPNTP–EGFP and exhibited the best morphological characteristics for imaging, was selected for this analysis. Data obtained with Dox-induced I3 cells were compared with either uninduced cells or control cell lines treated with Dox to rule out Dox-dependent effects.

The I3 cell line inducibly expressing DPNTP–EGFP was grown to confluence in the presence or absence of Dox, wounded, and then mounted in a living cell viewing chamber as described in the Materials and methods. To initiate stress-induced tearing of the cell sheet, the wounded monolayers were released from the center of the coverslip by flowing dispase through the chamber. The edges of the cell sheet remained tethered by the circumferential gasket associated with the living cell chamber; thus, cells at the edge of the wound began to stretch and tear due to contraction of the sheet as it lifted from the coverslip. Time-lapse images were captured at 4- (Fig. 6 A) or 8-s (Fig. 6 B) intervals and are shown here as selected individual still images and as four supplemental video clips (Videos 1–4, available online at http://www.jcb.org/cgi/content/full/jcb.200206098/DC1). Aspect ratios, which describe the geometry or “squareness” of a cell, were determined by dividing the diameter along the axis of stretch by the diameter orthogonal to the axis of stretch.

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Figure 6.

DPNTP expression leads to rapid dissociation of cells within an epithelial sheet with less cell distortion than control cells. (A) The A431 I3 DPNTP–EGFP line treated with Dox (top panels) or without (bottom panels) was imaged during stress-induced tearing. Expression of DPNTP was confirmed by visualizing EGFP fluorescence (top, second panel from right). Individual still images from the time-lapse record have been enlarged here to include the boxed regions and are shown with a time stamp. The entire time-lapse record for each movie can be viewed at http://www.jcb.org/cgi/content/full/jcb.200206098/DC1 as Video 1 (top panels) and Video 2 (bottom panels). (B) A Dox-induced DPNTP–EGFP line (top panels) was imaged during stress-induced tearing and compared with control cells treated with Dox (bottom panels) to rule out Dox- dependent effects. Individual still images from the time-lapse record are shown here and the complete movies can be viewed as Video 3 (top panels) and Video 4 (bottom panels). Expression of DPNTP was confirmed by visualizing GFP fluorescence (unpublished data). Quantitative information for the cells marked with the asterisk is shown in C, left. Note that in the examples in both A and B, cell sheets expressing DPNTP tear more rapidly, and cells become less distended before cell–cell dissociation occurs, reflecting that less stress is required for dissociation to occur. This can be appreciated best by watching the video clips. (C) Cell aspect ratios for the cells in B were determined as described in the Materials and methods and plotted versus time. Arrows represent the time points shown in B. Note that the DPNTP–EGFP cells dissociated more quickly with less distension along the axis of stretch. Population data summarizing the average time to breakage are provided in C, right. Error bars represent the SEM for a population of 10–15 cells.

Although the precise degree of stretch could not be controlled using this method, we observed a dramatic difference between DPNTP-expressing cells and control cells, both in the length of time it took for tearing to occur and the degree of cell distortion observed at the time of dissociation (Fig. 6, A–C). DPNTP-expressing cells dissociated much more rapidly than control cells (Fig. 6, C and D) and did so without undergoing as much distension along the axis of stretch (Fig. 6 C).

DPNTP expression does not alter E-cadherin function

As shown above, total and cell surface expression of E-cadherin were not altered by expression of DPNTP. To verify that DPNTP did not interfere with E-cadherin function, a capillary tube flow assay was employed (Yap et al., 1997; Gottardi et al., 2001). Uninduced and induced A431/C2 and G4 cells were first allowed to adhere to the inner surface of capillary tubes precoated with the extracellular domain of E-cadherin and then subjected to shear flow through the capillaries after either 10 or 45 min (Fig. 7), the latter time frame having previously been shown to allow for a strengthening phase of adhesion (Yap et al., 1997). The percentage of cells still adhering to the capillary tube after being subjected to various flow rates was recorded. CHO cells, which do not express E-cadherin, were used as a negative control, whereas transfected CHO cells expressing E-cadherin were used as a positive control. Although previous studies suggested that the β-catenin binding site is not necessary for adhesion or strengthening, when assays were performed in the presence of LtnA, E-cadherin–dependent adhesion was abrogated, supporting the importance of the actin cytoskeleton in this assay (Fig. 7). Both uninduced and induced A431/C2 and G4 cells exhibited adhesion comparable to the E-cadherin–expressing CHO cells at both time points (unpublished data). These results suggest that DPNTP expression does not impair the early phase of E-cadherin–dependent adhesion or the strengthening phase, both of which are also dependent on an intact actin cytoskeleton.

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Figure 7.

DPNTP expression does not alter the adhesive function of E-cadherin. Capillary flow assays were performed on induced (2 μg/ml Dox) and uninduced (0 μg/ml Dox) A431/C2 and G4 cells using glass capillary tubes coated with recombinant E-cadherin extracellular domain as described in the Materials and methods. Adhesion of both treated and untreated cells was similar under various flow rates. In addition, no difference was seen in the strengthening of adhesion after cells were allowed to adhere to capillary tubes for longer periods of time (45 min). Note that pretreatment with 2 μM latrunculin A abrogated E-cadherin–dependent adhesion. Note that the G4 line exhibited a somewhat greater loss of binding in response to flow than the C2 line in both the Dox-treated cultures and controls. It is possible this could be explained in part by the lower expression of E-cadherin in this line.

Contribution of microfilaments to adhesive strength in uninduced and induced cell lines

To further verify that DPNTP expression did not interfere with the cortical actin cytoskeleton–dependent strengthening of adhesion, Dox-treated and untreated A431 monolayers were incubated with LtnA before application of mechanical strain. LtnA disrupts the actin cytoskeleton by sequestering G-actin, which subsequently causes depolymerization of F-actin and disrupts adherens junctions (Cai et al., 2000). If the observed DPNTP-dependent loss of adhesive strength was solely due to interfering with adherens junction function, one would not expect a dramatic increase in stress-induced dissociation when DPNTP is expressed in the presence of LtnA.

As expected, Dox-treated A431/C2 monolayers exhibited significant dissociation (Fig. 8 A). LtnA alone also resulted in the dissociation of cell sheets. When the A431/C2 line was treated with 2 μM LtnA after Dox treatment to induce DPNTP expression, the monolayer dissociated into >500 particles. Measurement of LDH release indicated that there was no statistical difference among the samples and, in all cases, the amount of LDH released after mechanical strain was <5% of the total amount of LDH expressed.

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Figure 8.

LtnA and DPNTP act synergistically to weaken adhesion. (A) Dox-treated and untreated A431/SS and A431/C2 monolayers were treated with dispase and subjected to mechanical stress as described above for Fig. 4, with or without a 30-min treatment with 2 μM LtnA. (B) Again, LDH release was not significantly increased despite the dramatic fragmentation of LtnA-treated monolayers expressing DPNTP. Error bars represent the SEM from an experiment in which each condition was tested in triplicate.

Contributions of the desmosomal plaque and IF attachment to adhesive strength

Drawing a parallel to adherens junctions, one might envision that desmosomal strengthening of adhesion would involve at least two major steps: (1) lateral clustering and proper orientation of desmosomal cadherins through proper assembly of the plaque and (2) physical anchoring of the cadherin complex via IFs. In the case of adherens junctions, it has been suggested that lateral cadherin clustering of the extracellular domain, whether through artificial means or via p120^ctn, can increase adhesive strength (Brieher et al., 1996; Yap et al., 1997, 1998; Thoreson et al., 2000). We showed previously that the p120^ctn-related protein plakophilin 1 cooperates with plakoglobin to cluster desmosomal cadherins in the presence of DP (Bornslaeger et al., 2001). Furthermore, the NH₂-terminal 584 amino acids of DP, which comprise DPNTP, are sufficient for such clustering in L cell fibroblasts (Kowalczyk et al., 1997). It seems likely that this clustering event constitutes a necessary step in desmosomal cadherin–dependent adhesive strengthening. Consistent with this idea is the observation that the DP NH₂ terminus appeared to restore sealing of epithelial sheets and possibly even some measure of IF attachment in DP-null keratinocytes (Hatsell and Cowin, 2001; Vasioukhin et al., 2001). However, it is clear from the present work that even though DPNTP is capable of clustering desmosomal cadherins and forming junctional plaques in A431 cells, adhesive strength is severely compromised in epithelial sheets derived from these cells.

As DPNTP-dependent clustering is not sufficient to support adhesion comparable to wild-type DP, this ability must require the remaining DP domains, which include the rod and the COOH-terminal IF-binding domains. Previous work suggests that the DP rod contributes to the formation of oligomeric coassemblies with IF polypeptides that build the innermost portion of the plaque (Stappenbeck and Green, 1992; Kouklis et al., 1994). Thus, it could be envisioned that loss of these domains may compromise the ability of the plaque to withstand mechanical stress. However, the JD-1 cell experiments suggest that even small deletions of the DP COOH terminus that interfere with IF anchorage are enough to compromise adhesive strength when stress is applied to cells in vitro (Fig. 9 A) and in vivo (Norgett et al., 2000).

IF–membrane attachments may enhance adhesive strength in several ways. As has been suggested for actin, IFs may stabilize desmosomal cadherin clusters formed by the armadillo–DP NH₂-terminal complex, and thus, the proper disposition of desmosomal cadherin extracellular domains. Another possibility is that IFs anchor desmosomal cadherins within the plane of the membrane and that uncoupling of this connection results in molecular extraction of desmosomal cadherins out of the plane of the membrane. Similar molecular extraction has been reported previously for agglutinin receptors from red blood cells (Evans et al., 1991), L-selectin from neutrophils (Shao and Hochmuth, 1999), and integrins from fibroblasts moving across a substratum (Palecek et al., 1996). Another possibility is that the entire plaque might be extracted, as might be suggested by the plaque-associated membrane blebs seen in electron micrographs of the epidermis of DP-null animals (Vasioukhin et al., 2001). Whether such blebs seen in vivo would be enough to lead to cytolysis is not clear, but if this sort of extraction occurred in A431 cells, membrane resealing would have to be rapid so that LDH release would not occur.

The actin and IF cytoskeletons contribute synergistically to provide strong intercellular adhesion through classic and desmosomal cadherins

DPNTP was previously shown in stable A431 clones to lead to the codistribution of certain adherens junction and desmosome components (Bornslaeger et al., 1996), raising the possibility that DPNTP might interfere with adhesive strength solely by interfering with E-cadherin function. However, as shown here, the fluorescence patterns for E-cadherin and Dsg 2 did not detectably differ in uninduced and induced cells (Fig. 3, B and C), and the dramatic codistribution between desmosomal and adherens junction components that was previously reported in those clonal lines was not observed here. The basis for this difference is not currently known, although it may be related to DPNTP expression levels, clonal variability, or both.

Nevertheless, we sought to ensure that the observed adhesive defect was not through effects on E-cadherin. The experiments outlined here show that, in addition to the fact that neither cell surface expression nor distribution was altered, the fraction of E-cadherin present in the detergent-insoluble pool remained constant. This was in contrast to Dsg 2, which was reduced in the Triton-insoluble pool, likely due to the loss of attachment to the IF cytoskeleton. The data presented here also support the idea that E-cadherin is still functional in DPNTP-expressing cells. The ability of DPNTP-expressing cells adhering to capillary tubes coated with the E-cadherin extracellular domain to resist shear stress was no different than that of uninduced cells at early or later time points, even though adhesion was totally abrogated by LtnA. In addition, DPNTP-expressing epithelial sheets treated with LtnA exhibited a dramatically increased dissociation, which would be predicted only if IF attachments have an additional strengthening effect beyond that contributed by the cortical actin–adherens junction complex. Thus, the decreased structural integrity of DPNTP-expressing monolayers is not simply due to interference with E-cadherin function.

In fact, as the combined effect of LtnA treatment and DPNTP expression is much greater than the sum of the effects of the separate treatments, our data suggest that the actin and IF cytoskeletons function synergistically to provide strong intercellular adhesion and maintain structural integrity of cell sheets. These data are consistent with earlier work demonstrating that the microfilament and IF cytoskeletons are physically associated at the ultrastructural level in cells. Indeed, our earlier work in primary mouse keratinocytes demonstrated an intimate relationship between the cortical actin circumferential ring associated with the perpendicular bundles emanating from adherens junctions and keratin IF bundles that looped in and out of the cortical actin (Green et al., 1987). More recent work suggested that F-actin serves as a template for cytokeratin organization in vitro (Weber and Bement, 2002), and such interactions might in fact be stabilized by other members of the plakin family of cytolinkers, which may cross-link these systems laterally (Fuchs and Yang, 1999).

Observations from DP-null keratinocytes support the idea that the DP–IF network plays an important role in the maturation of the keratinocyte microfilament cytoskeleton (Vasioukhin et al., 2001). Cells cultured from these animals had severely reduced desmosome numbers and, in addition, failed to undergo the normal maturation of the cortical actin cytoskeleton, which might in turn contribute to the observed failure to form epithelial sheets in vitro and skin fragility in vivo. In the current work, the actin microfilament system did not appear to be significantly altered in DPNTP-expressing cells or JD-1 keratinocytes (unpublished data), possibly due to restoration of certain DP NH₂-terminal–dependent functions (Hatsell and Cowin, 2001). Together these previous observations and the data presented here suggest two related but independent points. First, DP, possibly through its NH₂-terminal head domain, plays an important role in the development and maturation of the actin cytoskeleton. Second, actin and IF cytoskeleton connections to cadherins function to synergistically strengthen adhesion, possibly through anchoring cadherins in the plane of the membrane, thus maintaining epithelial cell sheet integrity.

Cell culture and generation of inducible lines

Parental A431 epithelial cells were cultured as previously described (Bornslaeger et al., 1996). These cells were transfected with pTet-On (CLONTECH Laboratories, Inc.) using calcium phosphate precipitate and selected using 400 μg/ml G418 (Mediatech). Expression of the tetracycline-responsive transactivator in drug-resistant lines was determined by a luciferase assay (pTRE-Luc; CLONTECH Laboratories, Inc., and pRL-CMV; Dual-Luciferase Reporter Assay System; Promega). To establish tetracycline-inducible DPNTP–FLAG lines, a single stable line, 68Q71, was cotransfected with p801 and pSV2pacΔP. After selection with 400 μg/ml G418 and 1 μg/ml puromycin, resistant clones were screened for DPNTP–FLAG by treating with 2 μg/ml Dox for 24 h. The cells were then lysed in Laemmli sample buffer and analyzed via SDS-PAGE and immunoblotting. To establish a cell line inducibly expressing EGFP-tagged DPNTP, the 68Q71 cell line was transfected with vectors p992 and pSV2pacΔP, and puromycin-resistant clones were selected as described above.

JD-1 keratinocytes isolated from patients were immortalized with an HPV16 plasmid (pJ45216) that has early region genes driven by MoMLV-LTR as previously described (Storey et al., 1988). Cells were cultured in DME/Ham's F12 (3:1) supplemented with 10% FCS, 4 mM glutamine, 0.4 μg/ml hydrocortisone, 0.1 nM cholera toxin, 5 μg/ml insulin, and 10 ng/ml EGF. NHEK were isolated from neonatal foreskin as previously described (Halbert et al., 1992) and were cultured in media M154 (Cascade Biologics, Inc.).

Antibodies

Full-length DP and DPNTP–FLAG were detected using a rabbit polyclonal antibody, NW161, against the DP NH₂ terminus (Bornslaeger et al., 1996) or poly-FLAG (Sigma-Aldrich) against the FLAG-tagged constructs. The following rabbit polyclonal antibodies were also used: C2206 against β-catenin (Sigma-Aldrich), C2081 against α-catenin (Sigma-Aldrich), 795 against E-cadherin (provided by R. Marsh and R. Brackenbury, University of Cincinnati, Cincinnati, OH), and NW6 against the DP COOH terminus (Angst et al., 1990). 1407 is a polyclonal chicken antibody recognizing plakoglobin (Gaudry et al., 2001). Mouse monoclonal antibodies used in this study were: DP2.15 (a gift from P. Cowin, New York University, New York, NY) against the central rod domain of DP, Dg3.10 against Dsg 2 (Schmelz et al., 1986), HECD-1 against human E-cadherin (Shimoyama et al., 1989), 6D8 against Dsg 2, 11E4 against plakoglobin, 5H10 against β-catenin (gifts from M. Wheelock, University of Nebraska, Omaha, NE), and KSB17.2 against keratin 18 (Sigma-Aldrich).

Preparation of whole cell lysates and Triton-insoluble pool

To obtain whole cell lysates, cells were homogenized in Laemmli sample buffer, resolved on 7.5% SDS-PAGE gels, transferred to nitrocellulose, and immunoblotted using the following antibodies: 6D8 (1:1,000), 795 (1:1,000), C2081 (1:2,000), 5H10 (1:100), 11E4 (1:1,000), and NW161 (1:5,000). Sample loading was normalized to keratin that was visualized using KSB17.2 (1:2,000). For analysis of the Triton X-100–insoluble pool, cells were lysed in 1% Triton X-100 buffer (1% Triton X-100, 145 mM NaCl, 10 mM Tris-HCl, pH 7.4, 5 mM EDTA, 2 mM EGTA, and 1 mM PMSF) followed by centrifugation (16000 g, 30 min). The Triton X-100–insoluble pellet was solubilized in Laemmli sample buffer, resolved by SDS-PAGE, transferred to nitrocellulose, and labeled using 795 (1:1,000) and Dg3.10 (1:100).

Immunofluorescence analysis

Cells were seeded onto poly-l-lysine–coated (0.1 mg/ml, 3 h) glass coverslips. 1 d after plating, samples were induced with Dox (2 μg/ml, 24 h). Coverslips were then rinsed in DPBS and fixed in methanol (−20°C, 2 min). Indirect immunofluorescence was performed using KSB17.2 (1:200), poly-FLAG (1:50), NW6 (1:50), DP2.15 (1:100), Dg3.10 (1:10), and HECD-1 (undiluted supernatant). Primary antibodies were detected using Alexa Fluor^®488 goat anti–mouse IgG and Alexa Fluor^®568 goat anti–rabbit IgG at a dilution of 1:400 (Molecular Probes). Images were captured using a Leica DMR microscope/Orca Hamamatsu digital camera system and analyzed using OpenLab software (Improvision).

Fluorescence density at cell borders was determined by measuring fluorescent signal (OpenLab) from regions of intercellular contact in induced and uninduced A431/SS, C2, and G4 cells stained using 795 (1:1,000) and Dg3.10 (1:10) or JD-1 and NHEK cells stained using DP2.15 (1:500) and C2081 (1:2,000). Fluorescence density was calculated by dividing the fluorescent signal (pixels²) by the length (pixels) of the measured cell border. Calculations were determined from 10 representative regions of intercellular contact for each cell line.

Cell surface biotinylation

Cells treated with or without 2 μg/ml Dox were grown to confluency in 60-mm dishes. Cultures were washed twice with DPBS and labeled with cell-impermeable EZ-Link™ Sulfo-NHS-LC-Biotin (4 ml at 500 μg/ml in DPBS, 4°C, 30 min) (Pierce Chemical Co.) on a rocker. Cultures were then washed twice with DPBS and incubated in 4 ml of DPBS containing 100 mM glycine (4°C, 20 min) on a rocker. Finally, cells were washed twice with DPBS and lysed with 1 ml RIPA buffer (10 mM Tris-HCl, pH 7.5, 140 mM NaCl, 1% Triton X-100, 0.1% SDS, 0.5% sodium deoxycholate, 5 mM EDTA, 2 mM EGTA, 1mM PMSF). Biotin-labeled proteins were precipitated using 40 μl of UltraLink™ Immobilized Streptavidin (Pierce Chemical Co.) in 1.5-ml Eppendorf tubes overnight at 4°C on a rotator. The streptavidin beads were washed five times using lysis buffer and then centrifuged (10,000 g, 1 min, 4°C). Precipitated proteins were eluted using Laemmli buffer, boiled, resolved by SDS-PAGE, and immunoblotted using 795 (1:1,000), C2206 (1:4,000), Dg3.10 (1:100), and 1407 (1:5,000).

Hanging drop assay

Hanging drop cultures of aggregated cells were generated from 1 × 10³ cells that were either untreated or pretreated with Dox for 24 h and then dissociated in 1mM EDTA in DPBS without Ca²⁺ and Mg²⁺. Cells were allowed to aggregate over times ranging from 2 h to overnight on the underside of a culture dish as previously described (Thoreson et al., 2000). Resulting cell clusters were subjected to 30 rounds of pipetting through a 200-μl Gilson pipette, and the degree of dissociation was quantified by counting the particles after trituration.

Dispase-based dissociation assay

A431/SS, C2, and G4 cultures were seeded in triplicate onto 60-mm dishes. At ~50% confluency, cells were treated with or without 2 μg/ml Dox. 24 h after reaching confluency, cultures were washed twice in DPBS and then incubated in 2 ml of dispase (2.4 U/ml; Roche Diagnostics GmbH) for more than 30 min. Released monolayers were carefully washed twice with DPBS and transferred to 15-ml conical tubes. Enough DPBS was added for a final volume of 5 ml. Tubes were secured to a rocker and subjected to 50 inversion cycles. Fragments were counted using a dissecting microscope (Leica MZ6). For experiments using LtnA, cell monolayers released by dispase were further incubated for 30 min in 2 ml of dispase solution containing 2 μM LtnA. Statistical analysis was performed on the average of three separate experiments in the cases of the A431/SS and C2 cell lines and one experiment using G4 cells. Statistical significance (t test) was defined as P < 0.05. For experiments with JD-1 and NHEK, application of mechanical stress included 50 inversion cycles with either five additional inversion cycles in which the sample tubes were turned rapidly end over end or shaking at 400 rpm for 1 min in a rotary shaker.

LDH release measurements

LDH measurements using the Cytotoxicity Detection Kit (Roche Diagnostics GmbH) were performed according to the manufacturer's instructions on 200 μl of cell-free supernatant collected after monolayers were subjected to the dispase-based dissociation assay.

Videomicroscopy of dispase-treated wounded cell monolayers

Cell lines inducibly expressing DPNTP–FLAG or DPNTP–EGFP were seeded onto 40-mm diameter circular glass coverslips and allowed to grow to confluence either in the presence or absence of 2–4 μg/ml of Dox. Monolayers were then wounded at the center of the coverslips using a razor blade. Coverslips were then quickly mounted into the FCS2 closed system live-cell chamber (Bioptechs). The cell chamber was then filled with DPBS containing 48 U/ml dispase. Phase contrast and/or fluorescence time-lapse record of tearing of the monolayer was obtained at 40× magnification using a Leica DMIRBE inverted microscope fitted with an Orca Hamamatsu (C4742–95) digital camera and analyzed using Openlab software. Images were captured at 4- or 8-s intervals. Aspect ratios of measured cells were determined by dividing the diameter along the axis of stretch by the diameter orthogonal to the axis of stretch. Quicktime movies of the time-lapse record were assembled using Adobe Premier software.

Shear flow assay

Glass capillary tubes coated with 1 mg/ml protein A (Amersham Biosciences) in DPBS (4–7 h, 4°C) and blocked with 0.5% casein enzymatic hydrolysate were coated with 25 μg/ml E-cadherin extracellular domain–Fc fusion protein in HBSS plus 1 mM calcium chloride (HBSS⁺) (overnight, 4°C). Tubes were then blocked with 5% nonfat dry milk (in HBSS⁺, 2 h, 4°C) followed by a wash in HBSS⁺. A431/C2 or G4 cells treated with 2 μg/ml Dox for 36 h were dispersed with 2 mM EDTA in PBS without calcium or magnesium (37°C, 15 min) and washed in HBSS⁺ containing 0.2 mg/ml RGD peptide. Cells were loaded onto tubes and allowed to incubate for 10 or 45 min. Flow of HBSS⁺ started at 0.5 ml/min and doubled every 30 s, ending with a final flow rate of 25 ml/min. The percentage of cells bound to substrate was determined at each flow rate by video microscopy. For experiments involving LtnA, cells were incubated in 2 μM LtnA for 30 min before loading onto coated capillary tubes.

The authors would like to thank all those who have generously contributed antibodies, plasmids, and other reagents, including P. Cowin, R. Marsh, R. Brackenbury, M. Wheelock, K. Johnson, and P. Purkis. Thanks also S. Getsios and A. Kowalczyk for helpful discussion and critical reading of the manuscript.

This work is supported by National Institutes of Health grants RO1 AR43380 and PO1 DE12328 (project No. 4) to K. Green and R01 GM52717 to B. Gumbiner.