Cell Culture

T84 intestinal epithelial cells (American Type Culture Collection, Manassas, VA) were cultured in a 1:1 mixture of DMEM and Ham's F-12 medium supplemented with 10 mM HEPES, 14 mM NaHCO₃, 40 μg/ml penicillin, 100 μg/ml streptomycin, 5% newborn calf serum, and adjusted to pH 7.4 (further designated as T84 complete medium). For all experiments, T84 cells were grown for 8-14 d on collagen-coated, permeable polycarbonate filters, 0.4-μm pore size (Costar, Cambridge, MA). Filters with a surface area of 0.33 and 5 cm² were used for immunocytochemical and biochemical experiments, respectively.

Calcium Depletion and Pharmacological Modulation of Apical Junction Disassembly

To deplete extracellular Ca²⁺, confluent T84 monolayers were washed twice with calcium-free Eagle's minimum essential medium for suspension culture (Sigma) supplemented with 2 mM EGTA, 10 mM HEPES, 14 mM NaHCO₃, and 5% dialyzed newborn calf serum (designated here as S-MEM) and incubated in S-MEM for indicated times at 37°C. For pharmacological modulation of cytoskeleton, T84 cells were preincubated for 60 min with inhibitors in complete T84 medium followed by 60-min incubation in S-MEM containing the same concentration of inhibitor. Stock solutions of water-insoluble inhibitors were prepared in DMSO and diluted in cell culture media immediately before each experiment. The final concentration of DMSO was 0.1%; the same concentration of the vehicle was included in appropriate controls.

Immunofluorescence Labeling

Calcium-depleted T84 monolayers were rinsed twice with ice-cold calcium- and magnesium-free HBSS containing 10 mM HEPES (HBSS^-), whereas control monolayers were rinsed with HEPES-buffered HBSS containing calcium and magnesium (HBSS⁺). Cells were fixed/permeabilized in absolute methanol for 20 min at -20°C followed by blocking in HBSS⁺ containing 1% bovine serum albumin (blocking buffer) for 60 min at room temperature and incubation for 60 min with primary antibodies in blocking buffer. Cell monolayers were then washed, incubated for 60 min with Alexa dyeconjugated secondary antibodies followed by rinsing and mounting on slides with ProLong Antifade medium (Molecular Probes). For double labeling of junctional proteins and myosin II with F-actin, monolayers were fixed in 3.7% paraformaldehyde (PFA), permeabilized with 0.5% Triton X-100 (TX-100), and sequentially stained with primary and green Alexa dye-conjugated secondary antibodies, whereas F-actin was labeled with rhodamine-phalloidin. Stained monolayers were examined using a Zeiss LSM510 laser scanning confocal microscope (Zeiss Microimaging, Thornwood, NY) coupled to a Zeiss 100M axiovert and 63× or 100× Pan-Apochromat oil lenses. Fluorescent dyes were imaged sequentially in frame-interlace mode to eliminate cross-talk between channels. Images shown are representative of at least three experiments, with multiple images taken per slide.

Transmission Electron Microscopy

Confluent T84 monolayers were fixed in 4% buffered glutaraldehyde and cut into 1-μm strips using a microtome. The strips were postfixed in 1% osmium tetraoxide, sequentially dehydrated through graded alcohols and propylene oxide, and then infiltrated with Embed-812 (Electron Microscopy Sciences, Ft. Washington, PA). Strips were embedded for cross-section orientation. Semithin (0.5 mm) sections were cut, stained with toluidine blue, and examined for adequacy. Ultrathin section (900 Å) were cut with a diamond knife, stained with uranyl acetate and lead citrate, and examined with a Philips EM201 electron microscope (Philips Electronics, Mahwah, NJ).

Immunoblotting

Cells were homogenized in a lysis buffer (20 mM Tris, 50 mM NaCl, 2 mM EDTA, 2 mM EGTA, 1% sodium deoxycholate, 1% TX-100, and 0.1% SDS, pH 7.4), containing a proteinase inhibitor cocktail (1:100, Sigma) and phosphatase inhibitor cocktails 1 and 2 (both at 1:200, Sigma). Lysates were then cleared by centrifugation (20 min at 14,000 × g) and immediately boiled in SDS sample buffer. PAGE and immunoblotting were conducted by standard methods with 10-20 μg protein per lane. The results shown are representative immunoblots of at least three independent experiments. Quantification of protein expression was performed by densitometric analysis of Western blot images on the UN-SCAN-IT automated digitizing system (Silk Scientific, Orem, UT).

Fractionation of G- and F-actin and Incorporation of Exogenous G-actin

Quantification of G- and F-actin was performed by TX-100 fractionation of intracellular actin as previously described (Cramer et al., 2002). Briefly, control and calcium-depleted monolayers were washed with HBSS⁺ and HBSS^-, respectively, and G-actin was extracted by gentle shaking for 5 min at room temperature in HBSS⁺ containing 1% TX-100, the proteinase inhibitor cocktail, and 1 μg/ml phalloidin to prevent filament disassembly. This extraction method has been shown to remove ~95% of monomeric actin (Cramer et al., 2002). The TX-100-soluble G-actin fraction was mixed with an equal volume of SDS sample buffer and boiled. Filters were then briefly washed with HBSS⁺ and the TX-100-insoluble F-actin fraction collected by scraping cells in 2 volumes of SDS sample buffer, and boiled. The amount of actin in each fraction was determined by gel electrophoresis and Western blotting as described above. A similar procedure was also used to determine association of junctional proteins and myosin II with F-actin, except that the TX-100 extraction was performed for 15 min on ice.

Introduction of exogenous G-actin into permeabilized cells was performed as previously described (Symons and Mitchison, 1991). Briefly, T84 monolayers were calcium-depleted for 15 min and washed twice in a rinsing buffer (20 mM HEPES, 138 mM KCl, 4 mM MgCl₂, 3 mM EGTA, 1 mM ATP, pH 7.4). Cells were then incubated for 5 min at room temperature in a permeabilizing buffer (0.2 mg/ml saponin in the rising buffer) containing 1 μM of fluorescently labeled G-actin. Thereafter, cells were washed with HBSS^-, fixed with PFA, permeabilized with 0.5% TX-100, and counterstained for F-actin with rhodamine-phalloidin.

Transepithelial Resistance Measurement

Effect of calcium depletion on transepithelial electrical resistance (TEER) was measured using an EVOMX voltohmmeter (World Precision Instruments, Sarasota, FL). The resistance of cell-free collagen-coated filters was subtracted from each experimental point.

Formation of Contractile F-actin Rings Involves Turnover of Actin Filaments Associated with Activation of Cofilin-1/ADF and Recruitment of the Arp2/3 Complex

Given the observation that formation of contractile F-actin rings appears to be a driving force for disassembly of the AJC in calcium-depleted epithelial cells, we next investigated mechanisms involved in formation and contraction of these rings. Two potential mechanisms were investigated: a turnover (depolymerization/repolymerization) of cortical actin filaments and a myosin-dependent movement of stable actin filaments.

We first investigated whether depolymerization of pre-existing actin filaments is required for creation of F-actin rings in calcium-depleted cells. Actin depolymerization was inhibited using jasplakinolide, a cell-permeable agent that promotes polymerization and stabilizes actin filaments (Bubb et al., 1994, 2000). Because jasplakinolide has been shown to block phalloidin binding to F-actin (Bubb et al., 1994, 2000), we visualized total actin with polyclonal antibody. In calcium-depleted cells, this antibody detected the F-actin rings similar to those visualized by rhodamine-phalloidin (Figure 5). Incubation of T84 cells with jasplakinolide (2 μM) drastically attenuated ring formation. Furthermore, stabilization of F-actin with jasplakinolide prevented disassembly of the AJC and translocation of E-cadherin and occludin from areas of cell-cell contacts (Figure 5), thus suggesting a critical role for F-actin depolymerization in these processes.

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Figure 5.

Stabilization of actin filaments attenuates F-actin reorganization and disassembly of the AJC in calcium-depleted cells. T84 cells were preincubated for 60 min with jasplakinolide (2 μM) or vehicle followed by 60 min incubation in S-MEM containing the same concentrations of jasplakinolide. Localization of total actin, E-cadherin, and occludin was determined by immunolabeling and confocal microscopy. Jasplakinolide, which prevents depolymerization of actin filaments, substantially attenuates formation of contractile actin rings and disassembly of AJs and TJs in calcium-depleted cells. Bar, 10 μm.

We next analyzed whether calcium depletion induces activation of members of the actin depolymerizing factor (ADF)/cofilin protein family. These experiments were designed because ADF/cofilin proteins are known to greatly accelerate depolymerization of filamentous actin in living cells (Lappalainen and Drubin, 1997). It is generally believed that ADF/cofilin proteins possess much stronger F-actin-depolymerizing activity when they exist in the nonphosphorylated state (Moon and Drubin, 1995; Bamburg, 1999; Ono, 2003). Therefore, we used commercially available antibodies recognizing either total or phosphorylated (Ser3) cofilin-1 to analyze the phosphorylation status and thus activity of this protein. As shown in Figure 6, calcium depletion caused a progressive decrease in the amount of phosphorylated cofilin-1 in T84 total cell lysates without a significant effect on the total amounts of cofilin-1. Densitometry analysis of Western blots revealed that after 60 min of calcium depletion, the average amounts of phosphorylated and total cofilin-1 were, respectively, 11 and 76% of control levels (Figure 6B). Similar results (our unpublished data) were obtained using a different set of antibodies recognizing either total cofilin-1 and its homologue ADF (Morgan et al., 1993) or identically phosphorylated forms of these two proteins (Meberg et al., 1998). We also used the latter antibodies to analyze the effect of calcium depletion on intracellular localization of cofilin-1/ADF. Double labeling showed colocalization of total cofilin-1/ADF in areas of F-actin rings in calcium-depleted cells (Figure 6B, arrows). In contrast, no substantial colocalization of F-actin and phosphorylated cofilin-1/ADF was found, indicating that the nonphosphorylated, active forms of these proteins colocalize with F-actin rings. Taken together, these data suggest that calcium depletion is accompanied by activation (dephosphorylation) of cofilin-1 and its translocation to F-actin rings.

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Figure 6.

Calcium depletion is accompanied by activation of cofilin-1 and its accumulation in contractile F-actin rings. T84 cells were incubated in S-MEM for indicated times and amount of total and phosphorylated (P) cofilin-1 in cell lysates was determined by Western blotting. Representative blot (A) and quantification results from three independent experiments (B) shows that the amount of phosphorylated (inactive) but not total cofilin-1 drastically decreases during calcium depletion. (C) Calcium-depleted (30 min) cells were double-labeled for total cofilin-1/ADF and phosphorylated cofilin-1/ADF (green) with F-actin (red). Total cofilin-1/ADF colocalizes with F-actin rings (arrows); however, no such colocalization is seen for phosphorylated (inactive) cofilin-1/ADF. Bar, 5 μm.

Next we asked whether actin depolymerization is involved in the initial step of F-actin ring formation or whether biogenesis of such rings requires continuous depolymerization and de novo polymerization of actin. If the latter mechanism is valid, then sequestration of monomeric actin would be expected to induce disassembly of F-actin rings. To test this, we allowed the rings to develop during first 30 min of calcium depletion and then treated cells for another 30 min with either a high concentration of the G-actin-binding drug latrunculin A (50 μM) or vehicle. Figure 7A demonstrates that treatment with vehicle did not affect integrity of the F-actin rings, whereas sequestration of monomeric actin by latrunculin A caused rapid (within 30 min) ring disassembly. As a complementary approach, we investigated incorporation of exogenous G-actin into cytoskeletal structures of gently permeabilized, calcium-depleted T84 cells. We preincubated cells without calcium for 15 min with subsequent introduction of the fluorescently labeled G-actin in cell-permeabilizing buffer. Figure 7B shows that within 5 min of such introduction, a substantial amount of exogenous G-actin is incorporated into F-actin rings, especially at the ring edge (arrows). Interestingly, the rapid turnover of F-actin rings in calcium-depleted cells was not accompanied by significant changes in a balance between G and F-actin. Thus, the calculated G/F actin ratio was 0.96 ± 0.16 and 1.08 ± 0.3 (n = 3; p > 0.05) in T84 cells cultured in normal concentration of calcium and in calcium-free medium for 60 min, respectively.

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Figure 7.

G-actin is essential for stability of the contractile F-actin rings, and it is incorporated into rings during calcium depletion. (A). T84 cells were preincubated in S-MEM for 30 min followed by another 30 min incubation in S-MEM containing either G-actin-binding agent, latrunculin A (50 μM), or vehicle. Sequestration of G-actin with latrunculin A, but not vehicle treatment results in rapid disassembly of preformed F-actin rings. Bar, 10 μm. (B). Fluorescently labeled exogenous G-actin (green) was introduced for 5 min into saponin-permeabilized calcium-depleted T84 cells, and its intracellular localization together with topography of F-actin (red) was determined by confocal microscopy. As can be seen, exogenous G-actin is incorporated into F-actin rings in calcium-depleted cells (arrows). Bar, 5 μm.

Because nucleation of new actin filaments is regulated by the actin related protein (Arp) 2/3 complex (Suetsugu et al., 2002; Welch and Mullins, 2002), one could expect accumulation of the Arp2/3 complex in contractile F-actin rings if de novo actin polymerization occurs during their biogenesis. Indeed, double-labeling experiments revealed substantial colocalization of Arp3 protein with F-actin rings in calcium-depleted cells (Figure 8A, arrows). Furthermore, these rings were also enriched in cortactin (Figure 8A, arrows), a protein that recruits the Arp2/3 complex to actin filaments (Weed et al., 2000; Urino et al., 2001). Another argument supporting the role of Arp2/3-dependent actin polymerization in the formation of contractile F-actin rings was obtained by using 2,3-butanedione monoxime (BDM), a low-molecular-weight agent that has been reported to prevent Arp2/3 binding to F-actin (Yarrow et al., 2003). As shown in Figure 8B, BDM treatment (30 mM) attenuated the formation of F-actin rings and translocation of E-cadherin and occludin from intercellular junctions after 30 min of calcium depletion (compare images on Figure 8B to the time-matched controls presented in Figures ​Figures22 and ​and4A).4A). The Arp2/3-related data reinforces the idea that de novo actin polymerization is important for biogenesis of contractile F-actin rings and disassembly of the AJC in calcium-depleted epithelial cells.

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Figure 8.

Arp2/3 complex is involved in formation of F-actin rings and disassembly of the AJC. (A). T84 cells were depleted in calcium for 30 min and double-labeled for Arp3 and cortactin (green) with F-actin (red). As can be seen, both components of the active Arp2/3 complex colocalize with F-actin rings (arrows). Bar, 5 μm. (B). T84 cells were preincubated for 60 min with BDM (30 mM) followed by 30-min incubation in S-MEM containing the same concentrations of BDM. Localization of F-actin, E-cadherin, and occludin was determined by fluorescence labeling and confocal microscopy. BDM, which dislocates the Arp2/3 complex from actin filaments, substantially attenuates formation of contractile F-actin rings and disassembly of AJs and TJs in calcium-depleted cells (compare with time-matched images of untreated calcium-depleted cells on Figures ​Figures22 and ​and4A).4A). Bar, 10 μm.

Role for Actin Filament Turnover in Biogenesis of F-actin Rings

Contractile F-actin rings play critical roles in variety of biological processes including cell division (Field et al., 1999; Noguchi et al., 2001), wound closure (Bement et al., 1993, 1999; Mandato and Bement, 2001; Florian et al., 2002), endocytosis (Araki et al., 2002; Sokac et al., 2003), and extrusion of apoptotic cells from epithelial monolayers (Rosenblatt et al., 2001). Two different mechanisms account for generation of force required for ring contraction. One mechanism is due to dynamic depolymerization/repolymerization of actin filaments (Mandato and Bement, 2001; Henson et al., 2002; Pelham and Chang, 2002; Sokac et al., 2003). Another requires production of force by the motor protein, myosin II, (Bement et al., 1993, 1999; Mandato and Bement, 2001; Araki et al., 2002; Florian et al., 2002). Because both mechanisms have been implicated in the biogenesis of F-actin rings during cytokinesis and wound healing (Mandato and Bement, 2001; Noguchi et al., 2001), we analyzed whether they are involved in reorganization of apical F-actin in calcium-depleted epithelial cells.

Actin creates motile force via directed filament growth (Pantaloni et al., 2001; Pollard and Borisy, 2003). In this process, known as treadmilling, monomeric units are added to so-called “plus or barbed” ends of filaments and are removed from the opposite, “minus or pointed” ends (Pantaloni et al., 2001; Pollard and Borisy, 2003), thus rationalizing requirement of both polymerization and depolymerization for filament movement. In the present study, we have demonstrated that actin depolymerization is important for formation of contractile rings, because stabilization of F-actin with jasplakinolide drastically attenuated this process as well as disassembly of the AJC during calcium depletion (Figure 5). In a search for mechanisms involved in such actin depolymerization, we analyzed functional state of members of the ADF/cofilin family that accelerate disassembly of actin filaments in vivo (Moon and Drubin, 1995; Bamburg, 1999; Ono, 2003). Two members of this protein family, ADF and cofilin-1 are expressed in epithelial cells where they exist in nonphosphorylated and phosphorylated states (Bamburg, 1999). The nonphosphorylated state has much higher actin-depolymerizing activity (Bamburg, 1999; Ono, 2003). We found rapid dephosphorylation of cofilin-1 in calcium-depleted T84 cells (Figure 6, A and B), indicating activation of this protein. Furthermore, active, nonphosphorylated cofilin-1/ADF accumulated in contractile F-actin rings (Figure 6C). Similar decrease in phosphorylation and translocation of cofilin/ADF to areas of apical F-actin disassembly has been found in energy-depleted renal epithelial cells (Ashworth et al., 2001, 2003), where functional importance of these events was confirmed by expression of constitutive active and inactive forms of cofilin/ADF (Ashworth et al., 2003). We believe that activation of cofilin-1/ADF in calcium-depleted cells also has functional consequences by mediating disassembly of the apical F-actin.

In agreement with the idea that rapid turnover of actin filaments is involved in biogenesis of contractile F-actin rings in calcium-depleted cells, we demonstrated that de novo actin polymerization is essential for the rings formation and stability. Selective sequestration of G-actin with latrunculin A prevented rings formation (Figure 1) and resulted in rapid disassembly of intact ones (Figure 7A), indicating their dependence on a constant supply of monomeric actin. Furthermore, exogenous G-actin introduced into permeabilized calcium-depleted cells was rapidly incorporated into contractile F-actin rings (Figure 7B). Another group of evidence suggests a tentative mechanism for such de novo actin polymerization. Thus, we found that Arp3, the major component of the Arp2/3 complex that nucleates new actin filaments (Suetsugu et al., 2002; Welch and Mullins, 2002), colocalizes with the F-actin rings (Figure 8A). Similar colocalization was found for cortactin, which activates the Arp2/3 complex and targets it to actin filaments (Weed et al., 2000; Urino et al., 2001). Furthermore, treatment with BDM, that has been shown to displace the Arp2/3 complex from F-actin (Yarrow et al., 2003), attenuated formation of contractile F-actin rings and disassembly of apical junctions (Figure 8B). Taken together, these data suggest that the Arp2/3-assisted de novo actin polymerization is essential for biogenesis of contractile F-actin rings during calcium depletion.

The authors thank Drs. R. Adelstein, J. Bamburg, and M. Welch for generous donation of antibodies, S. Voss and R. Santoianni for the excellent technical assistance, D. Hunt for help in the manuscript preparation, and Drs. S. Ono, A. Hopkins, and K. Mandell for valuable comments on the manuscript. This work was supported by National Institute of Health Grants DK 61379 (to C.A.P.), DK 55679, and DK 59888 (to A.N.), a Digestive Diseases Minicenter grant DK 64399, and a Senior Investigator Award from the Crohns and Colitis Foundation (to A.N.).