1Biological Sciences, Marquette University, Milwaukee, Wisconsin; and 2Animal Sciences, Purdue University, West Lafayette, Indiana
Submitted 25 April 2005 ; accepted in final form 29 June 2005
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ABSTRACT |
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vinculin; talin; fibronectin; caveolin; stomach; ileum; colon; trachea
All of these proteins have been studied in smooth muscle in an effort to understand its regulation and contraction. Talin and vinculin are two cytoskeletal proteins associated with the AJs on the cytoplasmic side of the membrane. Talin is a 230-kDa protein (49) that has been reported to associate with actin, vinculin, and -integrins (9, 12, 51). Talin also has been reported to nucleate actin filament growth at the membrane (34, 35). Vinculin and metavinculin are splice variants of the same protein, and both are present in vertebrate smooth muscle (8, 25). In some smooth muscle tissue, metavinculin is actually present in higher concentrations than vinculin (24), but both of these proteins colocalize at dense plaques in smooth muscle cells (SMCs) (4). There is no known unique function for metavinculin, but a deficiency in the human heart can result in a cardiomyopathy (39).
Fibronectin is a ubiquitous extracellular glycoprotein that can bind to itself, collagen, integrins, and numerous other extracellular proteins (46). Fibronectin is involved in external linkages for cell-matrix interactions and may be involved in the regulation of the cell via activation of the AJs (41, 47, 52).
Caveolae are distinct plasmalemmal microdomains that are identified by their unique morphology (clusters of flask-shaped invaginations of the plasma membrane), high concentrations of receptors and channels, and the presence of the protein caveolin (1, 45). Caveolin is an integral membrane protein that acts as a scaffolding protein to localize and regulate a wide range of kinases. Caveolae are interdispersed with AJs in an alternating pattern along the plasma membrane in which interactions between the receptors and channels in the caveolae and the neighboring AJs may be important in cell regulation and function (2, 36, 42, 55, 58).
The role and function of these AJ-associated proteins are not well defined. Smooth muscle AJs (dense plaques) are known to be present at the plasma membrane, where they alternate with caveolin (16, 42, 50, 60) and have been observed in a variety of tissues (16, 17). While vinculin is a major constituent of AJs, it is not present in cytoplasmic dense bodies (21) but has been reported to be in equilibrium between cytosolic and cytoskeletal pools in chick embryo fibroblasts (38). Using tissue homogenization and fractionation, Kim et al. (37) reported that cholinergic stimulation of bovine trachea smooth muscle resulted in simultaneous increases in force and the recruitment of -actinin, talin, and metavinculin (but not vinculin) from the cytosolic fraction to the cytoskeletal fraction. Opazo Saez et al. (43), using indirect immunofluorescence, also reported that cholinergic activation of canine tracheal SMCs resulted in the translocation of vinculin, talin, paxillin, and focal adhesion kinase (FAK) from the cytoplasm to the membrane. These results suggest that vinculin, talin, and other cytoskeletal proteins demonstrate dynamic SMC distribution upon activation and relaxation of the tissue. However, both talin and vinculin are reported to dissociate readily from focal adhesions in permeabilized cells and show high solubility in the presence of nonionic detergents (6). Thus it is not clear whether these AJ-associated proteins are stably localized at the cell periphery under physiological conditions.
In smooth muscle, mechanical plasticity, a deviation of muscle force or shortening behavior from that mandated by static isometric force-length curves (57) has been reported by researchers at a number of laboratories. Pratusevich et al. (48) reported force production that is length independent in canine airway SMCs, and subsequently Seow et al. (53) suggested a series-to-parallel transition in the filament lattice to explain this observation. Others have proposed a repositioning of actin filament anchorage to dense plaques and dynamic actin filament remodeling to explain these results (22, 28, 29, 40). Hai et al. (10) also reported smooth muscle plasticity in tracheal smooth muscle that appeared to be related to a "memory" of previous strain and length. In contrast, Wingard et al. (68) found excellent evidence for the conventional length-tension curve in porcine carotid tissue. Thus numerous possible mechanisms could result in mechanical plasticity in smooth muscle.
The purpose of the present study was to determine the dynamic range of distribution of vinculin, talin, fibronectin, and caveolin in intact smooth muscle tissue under reduced-Ca2+ PSS, normal PSS, and carbachol-activating PSS conditions. The results show that in several different smooth muscle tissues, the cellular distribution of these proteins does not change when the tissue is incubated in Ca2+-free PSS, in normal PSS, or in normal PSS activated with carbachol before freezing and immunostaining. Thus immunohistochemical results from intact tissue suggest that these proteins do not translocate to and from the cytoplasm and plasma membrane in normal physiological function.
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METHODS |
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Reagents. The antibodies used were obtained from the following sources: talin (8D4), vinculin (VIN-11-5), and fibronectin (IST-3) were purchased from Sigma Chemical (St. Louis, MO); caveolin-1 was purchased from BD Biosciences/Pharmingen (San Diego, CA); Cy2 and Cy3 donkey anti-mouse or anti-rabbit secondary antibodies were purchased from Jackson ImmunoResearch (West Grove, PA); and phalloidin and DAPI were obtained from Molecular Probes (Eugene, OR). Antibody reactions were performed using standard procedures at room temperature. Frozen tissue sections (5 µm) picked up on glass slides were fixed with 2% paraformaldehyde for 10 min, permeabilized in 0.5% Triton X-100 for 10 min, and blocked with 5 mg/ml BSA for 1 h before being reacted with the primary antibody for 1 h and then the appropriate secondary antibody for 1 h. After reaction with the secondary antibody, the tissues were incubated in DAPI (0.5 µM), phalloidin (1050 nM), or DAPI-phalloidin as appropriate for staining nuclei and/or filamentous actin. Multiple washes were performed after the primary and secondary incubations. Coverslips were mounted over the tissue sections using buffered 75% glycerol with 0.2% n-propyl gallate to minimize fading. All immunoreacting solutions were made in PBS-Tween 20 (in g/l: 8.0 NaCl, 0.2 KH2PO4, 1.15 Na2HPO4, 0.2 KCl, and 0.1% Tween 20, pH 7.4) with 0.1% BSA.
Microscopy. Sections were observed using an Olympus IX70 inverted microscope with epifluorescence illumination. Digital images were obtained with a 16-bit Princeton Instruments (Princeton, NJ) charge-coupled device camera controlled through a PCI board via IPLab for Windows (version 3.6; Scanalytics, Fairfax, VA) on a personal computer. Images were obtained using either a x60 magnification/1.25 numerical aperture (NA) or x100 magnification/1.3 NA oil-immersion objective and stored on the personal computer. Montages were assembled in Adobe PhotoShop (version 6.0; Adobe Systems, San Jose, CA). Sections were viewed using a Zeiss confocal microscope (Axiovert 200 with LSM 5 Pascal software). No quantitative differences were observed in immunofluorescence distributions using these two different systems. All figures were obtained using the Olympus microscope. Histograms of fluorescence intensity (below camera saturation) of individual cells were obtained across the transverse tissue section using the IPLab software. Data from pairs of different cells (carbachol activated and relaxed in reduced Ca2+) were normalized to peak intensity and cell width using SigmaPlot software (version 8.0; Jandel Scientific, Corte Madera, CA) and replotted to allow for comparisons between cells. At least 4 animals were used for each tissue and antibody reported, with as many as 18 animals used to study the ileum and colon with some antibodies.
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RESULTS |
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To quantify the distribution of these proteins under these different physiological conditions, we studied fluorophore intensity histograms. Figure 5, A and B, shows high-magnification transverse photomicrographs of tissue sections from colon longitudinal smooth muscle after 10 µM carbachol activation (Fig. 5, Aa and Ba) or Ca2+-free PSS (Fig. 5, Ab and Bb). As in the other figures, the cytoskeletal proteins (vinculin, Fig. 5A, and talin, Fig. 5B; both green) were always localized near the periphery of the cell and alternated with caveolin (red in Fig. 5, A and B). In each image, a line is drawn across a SMC and the histogram of the intensity of the fluorophore of the respective cytoplasmic protein from a cell in the activated tissue and one in the relaxed state is plotted (Fig. 5, Ac and Bc). As shown for vinculin and talin (Fig. 5, A and B, respectively, histograms), the relative distribution did not change between the Ca2+-reduced and carbachol-activated conditions. The decay function of the decrease in fluorescence intensity with increasing distance away from the cell periphery to the center of the cell was not different between the relaxed and activated cells, and neither was the peak-to-valley ratio.
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DISCUSSION |
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The AJs are known to be a complex arrangement of transmembrane, intracellular, and extracellular proteins. Although >50 proteins have been reported to be associated with these structures (70), the exact roles of all of these proteins are not completely understood. In the literature, there are reports of numerous cytoskeletal proteins being localized to the AJs in smooth muscle. -Actinin (54), fibronectin (42), filamin (14, 56), integrins (42), metavinculin (25), paxillin (61), plectin (60, 66, 67), talin (13, 15, 63), and vinculin (21, 42, 54) are some of the commonly cited examples. These proteins are observed to be located at the cell periphery in a punctate pattern in transverse sections of smooth muscle in alternation with the caveolae (Refs. 18, 42, 54, 60; present study). In longitudinal sections of smooth muscle, the pattern of these proteins appears as stacks (60), ribs (42, 54), or staves (present study).
The morphology, stability, and regulation of the proteins associated with the AJs, as well as the entire cell, have been reported to undergo dynamic changes (see Ref. 70). These observations are generally reported in cultured cells, but remodeling of and/or translocation to and/or from the AJ also has been reported in tissues. For example, Taggart et al. (59) reported movement of receptor-coupled excitation molecules PKC, RhoA, and Rho kinase to the cell membrane in isolated SMCs with agonist stimulation, and Urban et al. (62) reported that RhoA kinase translocates to the membrane with K+ depolarization of arterial smooth muscle.
There also have been reports of cytoskeletal proteins translocating within cells as a result of stimulation. Beckerle et al. (3) reported talin redistribution from the cytoplasm to the adhesion plaque in platelets after activation. Opazo Saez et al. (43) reported the translocation of cytoskeletal proteins (vinculin, talin, and paxillin) as well as FAK to the cell membrane with agonist activation of isolated SMCs. Kim et al. (37) also reported translocation of -actinin, talin, and metavinculin (but not vinculin) in smooth muscle tissues with cholinergic stimulation.
Freshly frozen tissues from the gut and trachea showed that three cytoskeletal proteins (vinculin, talin, and fibronectin) were localized to the cell periphery at the AJ. They were located in a punctate pattern around the cell periphery in an alternating pattern with caveolin (Fig. 1) as also reported by others (42, 60). This pattern appeared to be stable and independent of tissue activation. A reduction of intracellular Ca2+ by tissue incubation in Ca2+-free PSS (no added Ca2+ and 0.5 mM EGTA) at 4°C overnight or by carbachol activation (3090 min) after 12 h at 37°C before freezing had no effect on the intracellular distribution of these proteins (Fig. 4). Fluorophore intensity histograms of vinculin and talin (Fig. 5, A and B) distribution in relaxed and activated smooth muscle also showed no differences in intracellular distribution. In both the Ca2+-free and activated conditions, these proteins were localized at the cell membrane and showed an identical decay function in fluorescence intensity from the cell periphery toward the center of the cell (Fig. 5, Ac and Bc).
To our knowledge, our present study represents the first time that these protein distributions have been analyzed in tissues after relaxation (Ca2+ reduction) and activation using immunohistochemistry. These results suggest that vinculin, talin, and fibronectin are stably associated at the AJs in gut and trachea SMCs in relaxed and activated conditions. Thus, in these intact tissues, the association of these proteins at the AJs appears to be physiologically important. While no other authors of whom we are aware have examined the localization of these proteins directly in tissues with relaxation and activation, numerous reports of the localization of these proteins in smooth muscle have been published, and these proteins are always found at the plasma membrane (see references cited at start of DISCUSSION). Immunolocalization of these proteins to the plasma membrane with specific antibodies leaves little room for alternative explanations. There is a possibility that the cold temperature in which the smooth muscle tissue was stored adversely affected the cytoskeletal system (including microtubules) and thus might have affected our results reported herein. However, other studies that we have performed did not show changes in microtubule pattern or staining intensity when freshly isolated SMCs were cold treated, and microtubule depolymerization favors Rho kinase activation, which would favor modest tissue activation, suggesting that freezing isolated SMCs did not affect our results (Swartz DR and Zhang D, unpublished results). In addition, the PSS sample served as a control for the influence of rewarming on microtubule status (both regular PSS and Ca2+-free PSS samples were treated in the same manner). Finally, in experiments in which tissues were warmed in PSS at 37°C for 12 h and then either relaxed in Ca2+-free PSS or activated with carbachol (all at 37°C) and then frozen, the AJ-associated proteins remained at the periphery, regardless of treatment.
Using high-speed centrifugation, Kim et al. (37) reported that cholinergic activation of bovine trachea strips resulted in a shift of -actinin, talin, and metavinculin (but not vinculin) from the cytoplasmic to the cytoskeletal membrane fraction. Peak force in this tissue was observed with 1 µM carbachol, and 80% of peak force was obtained with 0.1 µM carbachol. However, while the shifts from the cytoplasmic to the cytoskeletal membrane fractions were significant, they required activation with 110 µM carbachol, a concentration 12 orders of magnitude higher than that required for 80% peak force. Thus the tissue can generate
80% of peak force without significant shifts in the cellular distribution of these proteins. This leaves open the question regarding the physiological significance of movement from the soluble (cytoplasmic) to the insoluble (cytoskeletal) fraction with cholinergic activation.
Using immunohistochemistry, Opazo Saez et al. (43) reported that vinculin, paxillin, talin, and FAK were all evenly distributed throughout the cytoplasm of freshly isolated tracheal SMCs. When these cells were activated with 10 µM ACh, the distribution changed to one in which these proteins were primarily at the cell periphery. These authors concluded that this shift is physiologically relevant and critical for force generation and/or shortening. The results of the present study showing that two of these proteins (vinculin and talin) did not translocate to the membrane upon cholinergic activation are inconsistent with the results reported by Opazo Saez et al. The difference cannot be explained by tissue differences, because dog tracheal smooth muscle also was used in this study. The difference between activation with ACh by Opazo Saez et al. and the use of carbachol in the present study does not seem like a reasonable explanation. It is possible that the enzymatic and mechanical disruption of the tissue for SMC isolation used by Opazo Saez et al. (43) may have altered the AJs, leading to dissociation of these proteins. Subsequent activation of the cells could then have led to reformation and organization of the AJs.
An increasing body of literature suggests that the contractile filaments in smooth muscle are not static but assemble and disassemble with activation, relaxation, and/or changes in mechanical load. For example, if the thick and/or thin filaments within SMCs were not as stable as striated thick and thin filaments, they could assemble and disassemble more readily. This would allow the filamentous system to reorganize on the basis of overall cell length and activation history. This might explain in part the ability of smooth muscle tissue to generate high tension over a significantly wider range of muscle lengths than striated muscle. There are reports of increased birefringence in rat anococcygeus muscle with activation that was shown to correlate with a near-doubling of the myosin filament density of this tissue, of which only a portion can be explained by shrinkage or other changes (23, 26). Researchers from the same group subsequently reported that the increase in filaments was 23% in the anococcygeus but was not observed in the taenia coli, suggesting tissue differences (69). This finding is similar to the results described by Watanabe et al. (65), who reported an increase in myosin filaments in rat anococcygeus but not in guinea pig taenia coli.
The well-established length-tension relationship of skeletal muscle (27, 31, 32) may not be directly applicable to smooth muscle. While many groups (see, e.g., Ref. 68) have found evidence for the conventional length-tension curves in smooth muscle, other investigators (30) have reported a lack of a unique length-tension relationship in single SMCs. As mentioned in the introduction, this could be a result of changes in the arrangement of the contractile filaments (53), a repositioning of actin filaments relative to dense plaques (22, 28, 29, 40), or a "memory" of previous strain and length (10). Translocation of AJ-associated proteins (43) might be another mechanism that allows for mechanical plasticity in smooth muscle. The results of the present study suggest that the latter option is inconsistent with a physiologically relevant explanation for mechanical plasticity in intact smooth muscle. This study does not rule out the possibility of dynamic changes in the association of filamentous actin with the AJs, however.
In conclusion, the role of the AJ-associated proteins vinculin, talin, and fibronectin appears to be of such import that these proteins remain intact in smooth muscle tissues under relaxed and activated conditions. Changes to the cytoskeleton that would allow for the extreme range of shortening possible in smooth muscle most likely takes place elsewhere in the system, perhaps at the association of the thin filaments to the AJ-associated proteins, the arrangement of the thick and thin filaments, or changes in thin and thick filament lengths.
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GRANTS |
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ACKNOWLEDGMENTS |
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FOOTNOTES |
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The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
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