1Department of Pathology, Massachusetts General Hospital, and 2Department of Pathology, Harvard Medical School, Boston 02114; 3Gastrointestinal Cell Biology, Children's Hospital Boston, Boston, Massachusetts 02115; 4Department of Bacteriology, Okayama University Graduate School of Medicine and Dentistry, Okayama 700-8530, Japan; and 5Department of Pediatrics, Harvard Medical School, and 6Harvard Digestive Diseases Center, Boston, Massachusetts 02115
Submitted 15 April 2004 ; accepted in final form 12 July 2004
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ABSTRACT |
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membrane microdomains; membrane lipids; bacterial toxins; endocytosis; intestinal mucosa
Lipid rafts and caveolae have emerged as distinct portals of entry into the cell in a variety of biological and pathological processes (reviewed in Refs. 8, 36, and 42). Caveolae are abundant in specific cell types (not including intestinal epithelial cells) and exhibit a characteristic ultrastructural morphology and a cytoplasmic coat rich in caveolin-1 (46, 56). Noncaveolar lipid rafts are membrane microdomains with no specific morphological structure or uniformly accepted biochemical definition. Lipid rafts are operationally defined on the basis of their insolubility in nonionic detergents at 4°C and their sensitivity to cholesterol depletion (8, 30). Unlike caveolae, significant heterogeneity has been reported in the structure and function of lipid rafts (2, 16, 47, 50, 52, 60, 62, 63).
The presence of actin and other cytoskeletal proteins in detergent-insoluble membrane fractions has been known since the early description of these structures, and recent proteomic analyses have documented the presence of numerous cytoskeletal proteins in these fractions (37). Several studies suggest that the actin cytoskeleton plays an essential role in structure and function of lipid rafts in a variety of cell types (4, 13, 19, 38, 43, 55, 58, 59, 61). Furthermore, cholesterol depletion, which is often regarded as a functional test for dependence on lipid rafts, is associated with loss of phosphatidylinositol 4,5-bisphosphate (PIP2) from the PM and a global reorganization of the actin cytoskeleton (21), suggesting a possible role for actin in organization and/or function of lipid rafts.
In this study, we tested for a structural and functional association between CT bound to GM1 at the cell surface and the actin cytoskeleton in intestinal epithelial cells. We show that retrograde transport of CT, as well as toxin-induced cAMP generation, is affected by disruption of the actin cytoskeleton. We also find that CT and actin are colocalized within the same detergent-insoluble lipid microdomains. Cholesterol depletion, which inhibits retrograde trafficking and CT function (64), uncouples this association. We propose that the lipid raft provides the link between CT-GM1 complex at the cell surface and the actin cytoskeleton and that this association is required for retrograde transport of the toxin from PM to the ER and the induction of toxicity.
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MATERIALS AND METHODS |
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Cell culture.
Intestinal epithelial T84 cells obtained from the American Type Culture Collection (Rockville, MD) were cultured as previously described (7, 24). All monolayers used in the experiments were polarized cultures grown on 0.4-µm pore size Transwell inserts (Corning, Acton, MA) and had a resistance of at least 500 /cm2 determined by electrophysiology as described elsewhere (1). Hanks' balanced salt solution (HBSS) buffered at pH 7.4 with 10 mM HEPES was used as base buffer in all experiments unless otherwise stated.
Disruption of actin cytoskeleton and membrane cholesterol depletion.
Actin cytoskeleton was disrupted by treatment of T84 cell monolayers with drugs that either result in breakage of F-actin filaments or interfere with actin dynamics by binding F-actin. Briefly, monolayers were preincubated in HBSS containing 20 nM cytochalasin D (Cyto-D), 1 µM jasplakinolide, or 10 µM latrunculin A for 1 h at 37°C. These experimental conditions lead to significant loss of microfilaments or their function through different mechanisms of action (14, 23, 31). To prevent a washout effect, all CT experiments were carried out in the continued presence of actin-disrupting agents in all incubation buffers. For membrane cholesterol depletion, cells were incubated in HBSS containing 0.5% methyl--cyclodextrin (M
CD) for 1 h at 37°C. This condition was previously found to deplete lipid raft cholesterol and disrupt CT function in polarized T84 cells (64).
Measurement of intracellular cAMP levels. Cells were grown to confluence for 1014 days on 0.33-cm2 Transwell inserts. CT was added to the appropriate inserts after the preincubation period with actin-disrupting agents or vehicle alone. Forskolin, a direct activator of adenylyl cyclase, was added to the basolateral well of selected inserts to measure the enzyme activity. At time 0 and after 45 and 90 min of exposure to apical or basolateral CT, monolayers were rapidly chilled by immersion in excess ice-cold HBSS buffer containing 1 mM 3-isobutyl-1-methylxanthine (a phosphodiesterase inhibitor) to prevent loss of cAMP. Cells were then lysed at 4°C and assayed for cAMP with a direct ELISA kit according to the manufacturer's instructions (Amersham, Piscataway, NJ). For each time point, background and 1 µM forskolin-induced cAMP levels were obtained from additional monolayers treated in parallel. Thus each time point was analyzed by using measurements on control inserts (vehicle alone) with and without CT, control inserts (vehicle alone) with and without forskolin, actin-disrupted inserts with and without CT, and actin-disrupted inserts with and without forskolin, all in duplicate. Samples were diluted as necessary to ensure that all measurements were in the linear region of the ELISA calibration curve.
Disruption of the cytoskeleton may have an effect on adenylyl cyclase activity unrelated to CT action (15, 17). To eliminate such potential confounding effects, cAMP data from each ELISA were converted to a standard measure of toxin potency as follows. A cAMP standard curve was constructed for each experimental condition by directly activating the cyclase in T84 monolayers with a broad range of forskolin concentrations (0100 µM). All standard curves proved to be linear in this range, thus allowing a direct (linear) conversion of absolute cAMP levels into forskolin equivalents by using the baseline and forskolin controls from each experiment as two-point internal calibration. As noted above, these two-point calibration data were obtained in parallel for each experimental condition and at every time point. Comparison between groups was made with data expressed in forskolin equivalents, thus eliminating any potential confounding effects of drug treatment on adenylyl cyclase activity unrelated to CT action.
Biochemical assays for retrograde transport to Golgi and ER. A recombinant CT holotoxin (CT-GS) was engineered to contain tyrosine sulfation and N-glycosylation motifs as previously described (11). Recombinant toxin was expressed and purified as described elsewhere (45). For in vivo sulfation experiments (Golgi transport assay), actin-disrupted and control monolayers were washed and incubated in sulfate-free HBSS for 30 min at 37°C and incubated again for an additional 1 h in fresh, sulfate-free HBSS. Monolayers were then incubated with 0.5 mCi/ml Na235SO4 in the same buffer for 30 min. CT-GS was added apically and basolaterally to a final concentration of 20 nM, incubated for indicated times at 37°C, and washed twice with ice-cold HBSS before biochemical analysis. After cell lysis and immunoprecipitation with antibodies to CTB, samples were run on 1020% denaturing gels and the signal bands were detected and analyzed with a Molecular Dynamics PhosphorImager (Sunnyvale, CA). For ER transport assay, N-glycosylation of [35S]sulfated toxin subunits was assessed by a shift in molecular mass of the B-subunit detected by SDS-PAGE and quantitative fluorography on the PhosphorImager after CTB immunoprecipitation. In control experiments described previously (11), selective digestion of sample pairs with PGNase F was used to confirm evidence for N-glycosylation. For both assays, total cell-associated CT was determined by quantitative immunoblotting of a known concentration of the lysate. Data were analyzed and compared between groups by comparing the relative fraction of sulfated or N-glycosylated CT.
CT binding and internalization assays. A CT-HRP conjugate was used to quantify binding and internalization of CT in control monolayers and in monolayers treated with actin-disrupting agents under the conditions described above. For binding measurements, T84 cells grown to confluence in 96-well plates were treated with Cyto-D (20 nM), jasplakinolide (1 µM), or vehicle alone for 1 h at 37°C. Monolayers were then chilled to 4°C and exposed to 8 nM CT-HRP in the presence of 0.25% bovine serum albumin (BSA) for various times. After the indicated incubation periods, monolayers were thoroughly washed in phosphate-buffered saline (PBS) and incubated for 20 min with 0.5 mg/ml 2,2'-azino-bis(3-ethylbenzothiazoline-6-sulfonic acid) (ABTS) in citrate buffer, pH 4.2, in the presence of 1:1,000 hydrogen peroxide. Intensity of the developed color was measured at 410 nm and calibrated against an appropriate standard curve.
To measure the quantity of internalized CT, T84 monolayers were treated with Cyto-D or jasplakinolide as described above. After actin disruption, the incubation buffer was adjusted to 8 nM CT in the presence of 0.25% BSA and monolayers were incubated for various times at 37°C. After the indicated incubation periods, cell surface CT was removed as previously described (64). Cells were washed extensively with PBS, followed by two additional 1-min washes either with PBS (control monolayers) or with pH 2.5 phosphate buffer to remove surface-bound toxin (acid-stripped monolayers). All monolayers were then solubilized in 1% Triton X-100 and 60 mM n-octyl glucoside (NOG). Lysates were incubated for 20 min with 0.5 mg/ml ABTS in citrate buffer pH 4.2 in the presence of 1:1,000 hydrogen peroxide, and intensity of the developed color was measured at 410 nm. For each time point, total internalized CT was calculated as the relative color intensity of acid-stripped monolayers.
Fractionation with lipid rafts. Confluent monolayers of T84 cells grown for 1421 days on 45-cm2 Transwell inserts were used. All reagents were kept at 4°C during the entire procedure. Monolayers were rinsed in HBSS and incubated for 1 h with 8 nM CT in 10 ml of HBSS in the apical well, the basolateral well, or both. Unbound CT was washed away in HBSS, and inserts were placed in 2 ml of 1% Triton X-100 in detergent extraction buffer (DEB; 10 mM Tris·HCl, 150 mM NaCl, pH 7.4) containing the maximum recommended quantity of EDTA-free protease inhibitor tablets (Complete tabs; Boehringer-Mannheim, Indianapolis, IN). Inserts were gently shaken for 10 min, after which the lysate was removed and saved as the starting material of the "free" rafts in the first sucrose equilibrium density centrifugation. The partially extracted inserts were rinsed in DEB and placed in an additional 2 ml of 1% Triton X-100 in DEB. Cells were gently scraped, homogenized by 15 strokes in a tight-fitting Dounce homogenizer, and saved as the starting material of the "cytoskeletal" rafts in the second sucrose equilibrium density centrifugation. This two-step procedure divides the previously described lipid raft fraction of T84 monolayers (2, 65) into two subfractions, both of which meet the accepted criteria for lipid rafts: 1) insolubility in nonionic detergents at 4°C and 2) buoyancy in a sucrose gradient. In Table 1, the protein content of lipid rafts, soluble fraction, and pellet from the first sucrose gradient are listed under subheading 1, and the corresponding fractions from the second sucrose gradient are listed under subheading 2.
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Immunoprecipitation of lipid rafts.
Lipid rafts were immunoisolated by a variation of the method that was previously shown to be able to pull down intact membrane microdomains that contain the CT-GM1 complex as well as the associated raft proteins (2). Polarized T84 monolayers grown for 1421 days on 45-cm2 Transwell inserts were incubated apically or basolaterally with 10 ml of 8 nM CT for 1 h at 4°C. Cytoskeletal rafts were prepared as described above, pelleted at 100,000 g, and resuspended in PBS, pH 7.4, supplemented with 0.5% BSA (PBS-BSA). For immunoprecipitation with anti-CTB antibodies, 2 µg of protein G purified rabbit antiserum against CTB was added to 1 ml of lipid raft suspension containing 5075 µg of total protein. The suspension was tumbled end over end for 1 h, at which time 50 µl of magnetic Dynabeads with protein A (Dynal Biotech, Lake Success, NY) in PBS-BSA were added to the mixture. The mixture was tumbled for an additional 30 min, after which the beads were magnetically separated, washed extensively in PBS-BSA, and solubilized in electrophoresis sample buffer for immunoblotting. Control experiments included 1) matching concentrations of purified rabbit IgG, 2) no primary antibody, 3) no CT, and 4) immunoprecipitation in the presence of 1% Triton X-100 and 60 mM NOG, a condition known to dissolve the lipid rafts.
Immunoblotting. Samples were resolved on denaturing Tris·HCl polyacrylamide gels (Bio-Rad, Hercules, CA) and transferred onto nitrocellulose membranes by electroblotting. Membranes were blocked with 5% nonfat milk in 10 mM Tris and 150 mM NaCl, pH 7.6, containing 0.1% Tween 20 and probed with the primary antibody. Bound primary antibody was labeled with HRP-conjugated secondary and detected with an enhanced chemiluminescence reagent (Pierce) by imaging on Image Station 440CF (Eastman Kodak, Rochester, NY). Data were quantitated and exported for printing with Kodak 1D Image Analysis software (Eastman Kodak). Occasional immunoblots in which quantitation of band intensity was not required were developed and visualized with the Opti-4CN substrate kit (Bio-Rad).
Electron microscopy.
For thin-section electron microscopy of isolated lipid rafts, membranes were pelleted at 100,000 g for 30 min at 4°C, fixed in 2% gluteraldehyde and 2.5% paraformaldehyde in 0.2 M cacodylate buffer at 4°C, dehydrated, postfixed in osmium tetroxide, and embedded in Epon 812. For thin-section electron microscopy of monolayers, cells were treated with vehicle alone, Cyto-D, or MCD and extracted with Triton X-100. Extracted monolayers were washed in cold DEB, fixed, and processed as described above for raft membranes. Ultrathin sections were cut on a Leica Ultracut R, stained with uranyl acetate and lead citrate, and examined and photographed under a Philips EM208S electron microscope.
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RESULTS |
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Although N-glycosylation can be used for quantification of CT transport to the ER (11), we are unable to document a detectable glycosylation signal after 1 h of incubation with CT in control or actin-disrupted monolayers (note lack of detectable shift in molecular mass of CT in Fig. 3A). A detectable glycosylation signal was present after 3 h of continuous incubation with CT (Fig. 3C), at which point the relative glycosylation signals for jasplakinolide and Cyto-D are identical to control (note identical ratio of sulfated to glycosylated CT-GS in Fig. 3, C and D). However, these results are limited by the long incubation time required to detect a glycosylation signal. Thus, although our data indicate that the actin cytoskeleton is not required for toxin transport from Golgi to ER, it remains possible that the actin cytoskeleton affects the efficiency of toxin transport in a manner that cannot be detected at this late time point.
Structural association between cell surface CT-GM1 complex and actin cytoskeleton.
Given that the CT cell surface receptor ganglioside GM1 is not a transmembrane molecule, the functional data presented above suggest an indirect physical association between the CT-GM1 complex and the actin cytoskeleton, presumably mediated via the lipid rafts. To test for this association, we first examined isolated lipid rafts for enrichment in both CT and actin. Detergent extraction of T84 monolayers usually has been done with mechanical disruption of cells in the presence of nonionic detergents at 4°C. In the course of these studies, however, we found that a portion of the floating detergent-insoluble T84 membranes are dissociated from the cells without the need for mechanical disruption, suggesting a possible strategy for subfractionation of lipid rafts. This floating fraction (abbreviated as free rafts) contains 0.25% of total cellular proteins and consists of a relatively uniform population of membranes ranging from 50 to 250 nm in cross-sectional diameter (Fig. 4A). This size distribution is somewhat broader than that of classic caveolae (56) but consistent with the size distribution of lipid rafts isolated from other epithelial cell types obtained without cell homogenization (33). A second floating fraction (abbreviated as cytoskeletal rafts) is obtained after a subsequent mechanical disruption (Dounce homogenization) of the T84 monolayers in nonionic detergents. This floating fraction contains
1.25% of total cellular proteins and consists of more irregular membrane fragments measuring 100500 nm in average dimension (Fig. 4B). The shape and size distribution of these rafts are comparable to those of previously reported rafts isolated after Dounce homogenization of cells in nonionic detergents (29).
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To provide further evidence for colocalization of CT-GM1 and actin on cytoskeletal rafts, we first examined for the association of cell surface-bound CT and the actin cytoskeleton in situ by electron microscopy. After removal of free rafts from T84 cell monolayers exposed to CTB conjugated to colloidal gold, particles of gold were seen localized to patches of detergent-insoluble plasma membranes that were in close proximity to cytoplasmic actin bundles (Fig. 5A). Although this proximity is not surprising, given the richness of the apical region in actin cytoskeleton, it is consistent with a physical association between the cytoskeletal rafts containing the CT-GM1 complex and the actin cytoskeleton.
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Cholesterol depletion dissociates actin from cytoskeletal rafts.
Cholesterol is a key component of lipid rafts and is required for the integrity of raft structure and function. Because depletion of membrane cholesterol in T84 cells is known to inhibit CT transport from plasma membrane to the Golgi and ER (11), and to inhibit toxin action as determined by short-circuit currents (64), we tested whether cholesterol depletion is associated with the uncoupling of CT-GM1 complex from the actin cytoskeleton. Cholesterol depletion of T84 cells does not displace the CT-GM1 complex from lipid rafts (64), and these results were confirmed and extended in the current studies (CT blots in Fig. 6A). In contrast, depletion of cholesterol by treatment of T84 monolayers with MCD caused a loss of actin in the cytoskeletal raft fractions (actin blots in Fig. 6A). Raft-associated actin, however, was not displaced by pretreatment of T84 monolayers with Cyto-D (Fig. 6A, actin blots) indicating that Cyto-D under these conditions does not result in significant loss of membrane-associated actin or the cortical cytoskeleton. These results suggest that membrane cholesterol is required for assembly of the actin cytoskeleton with cytoskeletal rafts but not for binding CT to cell surface GM1 associated with raft microdomains. Furthermore, uncoupling actin from cytoskeletal rafts correlates with a loss in CT function.
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DISCUSSION |
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The mechanisms that explain the structural and functional associations between caveolae or lipid rafts and the actin cytoskeleton are not well understood. In fibroblasts the SV40 virus enters the cell by binding GM1 in caveolae, and this process requires recruitment of actin to the caveolar membranes at the cell surface (43). However, the subsequent transport of SV40 to caveosomes en route to the ER does not depend on the formation of dynamic actin tails (43). These data suggest a role for the actin cytoskeleton in the formation of caveolae-derived endocytic vesicles, but actin-based motility does not explain the subsequent intracellular trafficking. In Madin-Darby canine kidney (MDCK) cells the actin cytoskeleton is involved in the internalization of caveolae containing CT, and in A431 cells disruption of actin filaments completely inhibits the okadaic acid-induced uptake of alkaline phosphatase by caveolae (41). Lipid raft-mediated endocytosis is dependent on RhoA and Rac1 in lymphocytes (22) and cdc42 in COS cells (49), suggesting that the actin cytoskeleton also plays a role in endocytosis and trafficking of lipid rafts in these cell types.
The endocytic intermediates involved in the physiologically relevant transport of CT from the PM to the TGN and the ER, however, are not defined. We did not detect an effect of actin-disrupting agents on the overall rate of toxin endocytosis. These results were not expected, given the emerging evidence for a critical role of actin in clathrin- and caveolae-mediated endocytosis and budding (6, 9, 18, 43). It is possible that our data are explained by the lack of pathway specificity in the endocytosis assay coupled with the fact that CT enters the cell via multiple pathways, not all of which lead to transport to the Golgi and induction of toxicity (32).
Our results demonstrate a physical association between the lipid-anchored CT-GM1 complex on the outer leaflet of the PM and actin on the cytoplasmic surface of the PM. A physical association between an outer-membrane lipid and the actin cytoskeleton requires the presence of other structural, signaling, or regulatory proteins or lipids or both. Because lipid rafts in vivo are thought to be small and highly dynamic, the idea that they are stabilized by membrane-intrinsic or membrane-associated proteins has gained acceptance (8, 35, 42). Although several different structural assemblies or signaling pathways can be envisioned, given the richness of lipid rafts in cytoskeletal and signaling proteins (10, 37), a significant body of data points to a likely physical link between actin and lipid rafts through the ezrin-radixin-moesin (ERM) family of proteins. Csk-binding protein, a broadly expressed lipid raft protein, directly binds EBP50, an ERM-binding protein that is also known as Na+/H+ exchanger regulatory factor (NHERF) (4). The ERM family of actin binding proteins contain a COOH-terminal actin-binding domain and an NH2-terminal domain that binds the membrane-associated PIP2. Furthermore, it has been shown that PIP2 fractionates with lipid rafts in a cholesterol-dependent manner (21, 44) and is involved in actin polymerization in association with lipid rafts (48). These data make PIP2 and the ERM family of actin-binding proteins possible candidates for mediating the interactions between CT lipid rafts and the actin cytoskeleton, but our current data and experimental models in T84 cells are insufficient for a specific functional analysis at this time.
A limitation of our studies is reliance on the use of detergents for biochemical analysis of CT rafts. The limitations of the use of detergents in isolation and characterization of lipid rafts are well known (53). Nevertheless, detergent insolubility has been used widely in characterization of structure and function of CT rafts, and it remains a useful technique for biochemical analysis of lipid rafts in other model systems. We also know that only raft-associated gangliosides are capable of transporting CT from the PM to the ER (11), and our functional assays confirm a link between CT trafficking and cAMP generation with the actin cytoskeleton. It is therefore likely that the specific fraction of detergent-insoluble membranes that contains both CT and actin is relevant to the biology of CT, and possibly the biology of lipid rafts in general. Further studies are needed to pinpoint the specific mechanism(s) by which actin plays a specific role in CT trafficking, as well as the specific protein and lipid components required for these interactions.
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GRANTS |
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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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