Trafficking of cholera toxin-ganglioside GM1 complex into Golgi and induction of toxicity depend on actin cytoskeleton

Kamran Badizadegan,1,2,3 Heidi E. Wheeler,3 Yukako Fujinaga,3,4 and Wayne I. Lencer3,5,6

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


    ABSTRACT
 TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Intestinal epithelial lipid rafts contain ganglioside GM1 that is the receptor for cholera toxin (CT). The ganglioside binds CT at the plasma membrane (PM) and carries the toxin through the trans-Golgi network (TGN) to the endoplasmic reticulum (ER). In the ER, a portion of the toxin unfolds and translocates to the cytosol to activate adenylyl cyclase. Activation of the cyclase leads to an increase in intracellular cAMP, which results in apical chloride secretion. Here, we find that an intact actin cytoskeleton is necessary for the efficient transport of CT to the Golgi and for subsequent activation of adenylyl cyclase. CT bound to GM1 on the cell membrane fractionates with a heterogeneous population of lipid rafts, a portion of which is enriched in actin and other cytoskeletal proteins. In this actin-rich fraction of lipid rafts, CT and actin colocalize on the same membrane microdomains, suggesting a possible functional association. Depolymerization or stabilization of actin filaments interferes with transport of CT from the PM to the Golgi and reduces the levels of cAMP generated in the cytosol. Depletion of membrane cholesterol, which also inhibits CT trafficking to the TGN, causes displacement of actin from the lipid rafts while CT remains stably raft associated. On the basis of these observations, we propose that the CT-GM1 complex is associated with the actin cytoskeleton via the lipid rafts and that the actin cytoskeleton plays a role in trafficking of CT from the PM to the Golgi/ER and the subsequent activation of adenylyl cyclase.

membrane microdomains; membrane lipids; bacterial toxins; endocytosis; intestinal mucosa


CHOLERA TOXIN (CT) produced by Vibrio cholerae causes a secretory diarrhea that is responsible for the morbidity and mortality associated with the disease cholera. The active holotoxin consists of a pentameric receptor-binding B-subunit (CTB) that binds ganglioside GM1 at the cell surface and an enzymatic A-subunit (CTA) that activates adenylyl cyclase by catalyzing ADP-ribosylation of GTPase Gs{alpha} (12, 54). The resulting elevation in cytosolic cAMP in intestinal epithelia leads to a chloride secretory response that is fundamental to the pathogenesis of diarrhea. Entry of CT into the cytosol of intestinal epithelial cells occurs after retrograde transport of CT from the plasma membrane (PM) to the endoplasmic reticulum (ER) via the trans-Golgi network (TGN) (3, 11, 24, 26, 39). The pathway into the cell taken by CT is a general lipid-dependent sorting pathway used by all AB5-subunit toxins and certain viruses, including polyoma and SV40 (reviewed in Ref. 28). In the ER, CTA unfolds and dissociates from CTB in a reaction facilitated by protein disulfide isomerase (57). The free, unfolded CTA retrotranslocates from the ER to the cytosol through an analog of the sec61 complex (20, 51). Partitioning of the CT-GM1 complex into lipid rafts at the cell surface is required for retrograde trafficking of CT into the ER and thus for CT action (40, 65), but specifically how the lipid raft or the cytoskeleton contributes to toxin transport in this pathway remains largely unexplained.

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.


    MATERIALS AND METHODS
 TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Reagents and antibodies. CT and CTB were purchased from Calbiochem (San Diego, CA). Rabbit polyclonal antisera to CTB were previously described (27). Rabbit polyclonal antibody to caveolin-1 was purchased from Santa Cruz Biotechnology (Santa Cruz, CA). Rabbit polyclonal antibodies to actin and {alpha}-actinin, mouse polyclonal antibodies to actin and vinculin, and all secondary antibodies labeled with horseradish peroxidase (HRP) were purchased from Sigma (St. Louis, MO). Mouse monoclonal antibodies to occludin, transferrin receptor (TfR), and ZO-1 were obtained from Zymed Laboratories (South San Francisco, CA) and to villin were obtained from BD Transduction Laboratories (San Jose, CA). Jasplakinolide and latrunculin A were purchased from Molecular Probes (Eugene, OR). All other reagents were purchased from Sigma unless otherwise stated.

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 {Omega}/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-{beta}-cyclodextrin (M{beta}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 10–14 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 (0–100 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 10–20% 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 14–21 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.


View this table:
[in this window]
[in a new window]
 
Table 1. Quantitative distribution of proteins in tandem detergent extraction of T84 monolayers and corresponding lipid raft subfractions

 
Sucrose equilibrium density centrifugation. All steps were carried out at 4°C or on ice. Nuclei and large debris were pelleted and discarded by centrifugation of each detergent-lysate at 1,000 g for 5 min. Approximately 2 ml of the 1,000 g supernatant were mixed with an equal volume of 80% sucrose in DEB, placed in a 12-ml ultracentrifuge tube, and layered with a step gradient consisting of 4 ml of 30% and 4 ml of 5% sucrose. Gradients were centrifuged for 3 h at 39,000 rpm in a SW-41 swinging bucket rotor (Beckman Instruments, Palo Alto, CA), after which a floating membrane fraction was visible at the 30%-5% sucrose interface. Floating membranes were collected in a volume of 0.5 ml from the gradients, pelleted for 30 min at 100,000 g, and either resuspended or solubilized in an appropriate buffer for subsequent experiments. A 0.5-ml sample of the soluble fraction was also removed from the bottom of each gradient without disrupting the other layers and saved for subsequent analysis. The remaining gradient was discarded, except for the pellet at the bottom of each tube, which was resuspended or solubilized in an appropriate buffer for subsequent analysis. When needed, protein concentrations were determined by the bicinchoninic acid method according to the manufacturer's instructions (Pierce, Rockford, IL) with BSA standards.

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 14–21 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 50–75 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 M{beta}CD 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.


    RESULTS
 TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Disruption of actin cytoskeleton inhibits CT function. In initial studies, we used Cyto-D (20 nM), latrunculin A (10 µM), or jasplakinolide (1 µM) to test whether CT activity depends on an intact and dynamic actin cytoskeleton. Polarized T84 monolayers preincubated with one of these actin-disrupting drugs or with vehicle alone were treated with CT applied to either the apical or the basolateral cell surface. Toxin entry into the cell and induction of toxicity was measured as an increase in intracellular cAMP assessed at 45 or 90 min after toxin application. These time points correspond to the induction of partial (45 min) or maximal (90 min) toxicity in T84 cells as assessed by the induction of a cAMP-dependent chloride secretory response (Refs. 25 and 65 and our unpublished studies). Compared with control monolayers that were treated with vehicle alone, toxin-induced cAMP in monolayers treated with actin-disrupting compounds was decreased by 50–80% at both time points after toxin application (Fig. 1). Each of the three actin-disrupting compounds tested caused similar levels of inhibition, and both apically and basolaterally applied CT were affected. These results show a dependence on the actin cytoskeleton for CT function.



View larger version (29K):
[in this window]
[in a new window]
 
Fig. 1. Effect of actin disruption on cholera toxin (CT)-induced cAMP generation. cAMP levels were measured in T84 monolayers exposed to apical (A–C) or basolateral CT (D–F). Measurements were made in the presence and absence of actin-disrupting agents cytochalasin D (Cyto-D; A and D), jasplakinolide (B and E), and latrunculin A (C and F). For each experimental condition, vehicle control (gray bars) and treatment (open bars) groups are compared at 45 and 90 min after the addition of CT. Toxin activity at each time point is normalized to forskolin equivalent units as described in MATERIALS AND METHODS. Under all experimental conditions, disruption of the actin cytoskeleton was associated with a reduction in CT-induced cAMP generation. Each bar represents data calculated from 6 independent monolayers assayed in duplicate for cAMP content. An additional 8 monolayers served as baseline controls for calculations.

 
Disruption of actin cytoskeleton affects CT trafficking. Toxin binding to the cell surface GM1 was not inhibited by treatment with actin-disrupting agents (Fig. 2A), and there was no detectable effect on the overall rate of CT internalization under these conditions (Fig. 2B). However, CT is internalized by a variety of pathways, not all of which result in functional trafficking to the Golgi and subsequent induction of toxicity (32). We therefore tested for a direct effect of disruption of the actin cytoskeleton on trafficking of the CT-GM1 complex from the PM to the Golgi/ER, which we believe to be a functionally active trafficking pathway (Refs. 28 and 57; Fig. 3). For this transport assay, we used a recombinant CT engineered to contain tyrosine sulfation and N-glycosylation consensus motifs on the COOH terminus of each B-subunit monomer (CT-GS). The functional activity of CT-GS is similar to that of the wild-type CT in T84 monolayers (11). Transport of CT-GS to the Golgi was assessed as the fraction of total cell-associated toxin that is modified to contain [35S]sulfate (Fig. 3, A and B). Transport to the ER was assessed as the fraction of toxin that is modified to contain N-linked oligosaccharides as documented by a shift in molecular mass of [35S]sulfated CT (Fig. 3C). Efficiency of transport from Golgi to ER was assessed as the ratio of sulfated CT-GS to glycosylated CT-GS (Fig. 3D).



View larger version (22K):
[in this window]
[in a new window]
 
Fig. 2. Effect of actin disruption on CT binding and internalization. Binding of CT to the cell surface and internalization of CT were colorimetrically assayed with CT-horseradish peroxidase (HRP) as the ligand. Compared with control monolayers with intact cytoskeleton, there was no detectable difference in the rate of binding of CT to the cell surface at 4°C after disruption of F-actin with Cyto-D or stabilization of F-actin with jasplakinolide (A). Similarly, there was no significant difference in continuous uptake of CT at 37°C between control monolayers and monolayers treated with Cyto-D or jasplakinolide (B).

 


View larger version (34K):
[in this window]
[in a new window]
 
Fig. 3. Effect of actin disruption on CT trafficking to Golgi/endoplasmic reticulum (ER). Transport of recombinant CT holotoxin (CT-GS) to the Golgi is assessed as the fraction of total cell-associated toxin that is modified to contain [35S]sulfate. A: representative data for CT transport from plasma membrane (PM) to Golgi at 1 h after toxin application. Top, [35S]sulfated CT B-subunit (CTB) after immunoprecipitation and SDS-PAGE. Bottom, immunoblot for CTB in the same samples to quantify the total toxin available for sulfation in each monolayer. B: fraction of sulfated CT-GS for 3 independent experiments are normalized and combined. A significant reduction in the fraction of toxin transported from PM to Golgi and sulfated is demonstrated for monolayers treated with either jasplakinolide or Cyto-D (*ANOVA: P = 0.03, multiple comparison procedures: P = 0.02–0.05). Control studies (not shown) show no effect of Cyto-D or jasplakinolide on the overall Golgi sulfotransferase activity. C and D: results of a similar experiment after 3 h of incubation with CT-GS in which both sulfation and glycosylation signals are detected as a shift in molecular mass of 35S-labeled toxin (C, top, double bands). D: band intensities are quantified and normalized with respect to total CT in each sample to express sulfation and glycosylation as % of control. Treatment with Cyto-D and jasplakinolide results in a reduction in sulfation (dots) similar to that shown in B. However, the ratio of sulfation (dots) to glycosylation (bars) is identical between groups, suggesting no effect of actin disruption on transport from Golgi to ER.

 
When measured after ~1 h of exposure to CT, only the sulfated B-subunit was detected (Fig. 3A). The fraction of sulfated toxin was quantified as the ratio of sulfated toxin to total toxin and combined for three independent experiments. At 1 h of incubation, there was an ~50% reduction in the sulfation of CT-GS caused by either jasplakinolide or Cyto-D (Fig. 3B; ANOVA: P = 0.03, multiple comparison procedures: P = 0.02–0.05). Control studies (not shown) showed no effect of Cyto-D or jasplakinolide on the total sulfotransferase activity. These data suggest an inhibition in toxin transport from PM to the Golgi caused by disruption of the actin cytoskeleton.

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 100–500 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).



View larger version (122K):
[in this window]
[in a new window]
 
Fig. 4. Structural and biochemical comparison of "free" and "cytoskeletal" raft fractions. By transmission electron microscopy, free rafts are a relatively uniform population of membranes (A) whereas cytoskeletal rafts consist of larger and more irregular membrane fragments (B). Immunoblotting data show free rafts to be enriched in caveolin-1 (C, lane 1), whereas actin and cytoskeleton-associated raft proteins are enriched in the cytoskeletal rafts (C, lane 2). Relative enrichment of CT in cytoskeletal rafts is shown in D, where nearly all of the basolateral raft-associated CT and more than half of the apical raft-associated CT are blotted in the cytoskeletal rafts. See Table 1 for quantitative data. EGFR, EGF receptor; TfR, transferrin receptor.

 
Free rafts and cytoskeletal rafts also show a distinct protein composition as assessed by immunoblotting for selected proteins. The three major components of each sucrose gradient (floating fraction, soluble fraction, and detergent-insoluble pellet) were reconstituted in electrophoresis sample buffer and immunoblotted simultaneously for selected raft-associated and cytoskeletal proteins (Fig. 4, C and D). Band intensities were quantitatively measured in a linear range, and the relative abundance of each protein was determined (Table 1). As shown in Table 1 and qualitatively presented in Fig. 4C, free rafts (Fig. 4C, lane 1) are enriched in caveolin-1 but show little to no detectable cytoskeleton-associated proteins. Cytoskeletal rafts (Fig. 4C, lane 2), on the other hand, contain less caveolin-1 but are enriched in actin, as well as actin-binding and regulatory proteins such as villin and EGF receptor (EGFR). Other cytoskeletal and trafficking proteins, such as ezrin, actinin, and Arf6, and TfR are predominantly found in the detergent-soluble fractions (Table 1) and do not significantly associate with lipid rafts under these conditions. When CT is applied to the apical membranes of T84 cell monolayers, the CT-GM1 complex is found in both free and cytoskeletal rafts (Fig. 4D and Table 1). In contrast, basolaterally applied CT is predominantly found in the cytoskeletal rafts. This polarized pattern of distribution is likely related to structural differences between apical and basolateral rafts in T84 cells and may explain in part why basolateral exposure to CT results in more efficient signaling in T84 cells as measured by the chloride secretory response (Ref. 25 and data not shown).

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.



View larger version (46K):
[in this window]
[in a new window]
 
Fig. 5. Membrane-bound CT is linked to actin via lipid rafts. Transmission electron microscopy of detergent-extracted T84 monolayers (A) shows that cell surface CTB-gold (arrowheads) is present on detergent-insoluble membranes in immediate proximity to cytoplasmic actin bundles (arrows). These detergent-insoluble membranes are the source of cytoskeletal rafts. Immunoprecipitation of cytoskeletal rafts isolated after sucrose equilibrium density centrifugation (B) shows that antibodies against CT pull down membranes that are enriched in both CT (lane 1, CT blot) and actin (lane 1, actin blot). No CT or actin is present in identical preparations immunoprecipitated with nonspecific rabbit IgG or without a primary antibody (lanes 2 and 3). Specificity for CT is shown in lane 4, in which cytoskeletal rafts are isolated from cell monolayers that were not exposed to CT. No CT or actin is precipitated under these conditions. Specificity for lipid rafts is shown in lane 5, in which cytoskeletal rafts from monolayers exposed to CT were solubilized in 60 mM n-octyl glucoside (NOG) before immunoprecipitation. CT but not actin is precipitated under these conditions, indicating the requirement for lipid rafts in associations between CT and actin. Representative data from 4 independent experiments are shown.

 
To test for a specific association between cytoskeletal CT rafts and actin on the same membrane microdomain, we tested for the coprecipitation of actin in immunoisolated cytoskeletal CT rafts (Fig. 5B). In these studies, cytoskeletal rafts were isolated from T84 cells that were preincubated with CT at 4°C to allow toxin binding to the cell surface before fractionation. Isolated cytoskeletal rafts were resuspended in buffer and immunoprecipitated with magnetic beads coated with rabbit antibodies against CTB. Incubations with anti-CTB-coated beads immunoprecipitated a fraction of lipid raft membranes that contained both CT (on the external surface of the raft membrane) and actin (on the cytoplasmic surface of the raft membrane) (Fig. 5B, lane 1). There was no immunoprecipitation of CT or actin by beads coated with nonspecific antibodies or by beads alone (Fig. 5B, lanes 2 and 3). Furthermore, neither actin nor CT was immunoprecipitated from cytoskeletal rafts that were isolated from cell monolayers not exposed to CT, proving that the assay is specific for CT rafts (Fig. 5B, lane 4). Finally, when cytoskeletal rafts were fully solubilized in buffer containing both Triton X-100 and 60 mM NOG, only CT and not actin was immunoprecipitated by antibodies to CT, proving a requirement for intact lipid rafts for coprecipitation of CT and actin (Fig. 5B, lane 5). These studies indicate that there is a physical association mediated by the cytoskeletal rafts between the CT-GM1 complex, which is anchored to the membrane in the extracellular leaflet only, and actin, which is associated only with the cytoplasmic surface of the raft microdomain.

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 M{beta}CD 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.



View larger version (39K):
[in this window]
[in a new window]
 
Fig. 6. Membrane cholesterol depletion dissociates actin from cytoskeletal rafts. Immunoblotting for CT shows no significant loss of the toxin from free or cytoskeletal rafts in cells treated with methyl-{beta}-cyclodextrin (M{beta}CD) or Cyto-D (A, CT blots). In contrast, when monolayers are treated with M{beta}CD there is a significant loss of actin from the cytoskeletal rafts (A, actin blots, arrowhead) whereas treatment with Cyto-D does not displace membrane-associated actin from cytoskeletal rafts. Transmission electron micrographs of detergent-extracted T84 monolayers (B–D) show that treatment with Cyto-D results in the loss of microfilaments [arrows, C vs. B (control)] but does not alter the morphology of detergent-insoluble membranes [asterisks, C vs. B (control)]. In contrast, cholesterol depletion results in the formation of extended stretches of somewhat thickened and disrupted detergent-insoluble bilayers [asterisks, D vs. B (control)] but has no detectable effect on the number and size of the microfilaments [arrows, D vs. B (control)].

 
To strengthen this idea, we examined T84 cells depleted in cholesterol for defects in cytoskeletal raft ultrastructure (Fig. 6, B–D). Although Cyto-D treatment resulted in its well-known effect of loss of microfilaments (Fig. 6C; compare microfilaments highlighted by arrows with those in control cells in Fig. 6B), detergent-insoluble membranes of Cyto-D-treated cells showed no appreciable abnormality in size or bilayer structure (Fig. 6C; compare membranes marked with asterisks with those in Fig. 6B). In contrast, M{beta}CD treatment resulted in a change in the ultrastructure of detergent-insoluble membranes (Fig. 6D; compare membranes marked with asterisks with those in Fig. 6, B and C). Detergent-insoluble membranes of M{beta}CD-treated cells are qualitatively larger and thicker than controls or Cyto-D-treated cells, and the bilayer structure is not readily resolved. Unlike Cyto-D treatment, however, M{beta}CD did not result in a significant loss of microfilaments (Fig. 6D, arrows). Collectively, these data suggest that the loss of CT function in M{beta}CD-treated T84 monolayers is associated with a structural defect in lipid rafts that results in loss of association with actin.


    DISCUSSION
 TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
These studies show a functional association between CT and the actin cytoskeleton in model intestinal epithelia. Disruption of actin dynamics inhibits CT trafficking from the PM to the Golgi and the induction of toxicity. We find evidence for a structural association between the lipid-anchored toxin and the actin cytoskeleton that may explain how the actin cytoskeleton affects the biology of the toxin. The association is disrupted by cholesterol depletion, and this correlates with a loss in CT function. On the basis of these results, we propose that the cell surface CT-GM1 complex links to the actin cytoskeleton via the lipid rafts (Fig. 7A). We propose that disruption of the actin cytoskeleton by drugs such as Cyto-D leaves the lipid rafts intact but results in loss of cytoskeleton-dependent lipid raft functions such as retrograde CT trafficking (Fig. 7B). In contrast, depletion of membrane cholesterol with drugs such as M{beta}CD results in loss of lipid raft function through biochemical changes in raft structure that result in disruption of the raft association with the membrane-bound cortical cytoskeleton (Fig. 7C). The exact nature of molecular complexes that mediate this complex interaction among gangliosides, lipids, and membrane and cytoplasmic proteins is currently unknown.



View larger version (27K):
[in this window]
[in a new window]
 
Fig. 7. Schematic representation of the association between CT and actin cytoskeleton. In intact cytoskeletal rafts (A), there is a physical association between CT-GM1 on the external surfaces of the plasma membrane and the actin cytoskeleton in the contralateral cytoplasmic compartment through the lipid rafts. Disruption of actin by agents such as Cyto-D (B) does not result in a detectable change in the raft structure but results in loss of raft function due to the loss of actin filaments and/or disruption of actin dynamics. Depletion of membrane cholesterol with agents such as M{beta}CD (C) does not directly alter the actin filaments but results in loss of structural/functional association between lipid rafts and the cytoskeleton (and secondary changes in the cytoskeletal architecture) due to structural changes in the lipid bilayer that result in the loss of membrane-cytoskeleton associations.

 
Similar roles for the actin cytoskeleton in the biology of lipid rafts have been implicated in other systems, most notably in T-cell signaling. Disruption of lipid rafts by cholesterol depletion in T cells inhibits the inducible tyrosine phosphorylation of the T-cell receptor (TCR)-{zeta} chain and prevents its association with actin (34). Actin reorganization in T cells is required for stable lipid raft clustering induced either by antigen or by CT-mediated cross-linking of GM1 (59). The idea that actin plays a central role in T-cell signaling via the lipid rafts is further supported by the observation that activation of the TCR induces reorganization of actin in a cholesterol-dependent manner (58). In addition to T cells, functionally important interactions between the high-affinity Fc{epsilon} receptor in activated mast cells and lipid rafts are shown to be dependent on the actin cytoskeleton (13), and in ciliary neurons cholesterol depletion results in redistribution of the actin cytoskeleton over the cell surface, as well as causing a substantial decrease in average size of membrane-associated F-actin patches (5).

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.


    GRANTS
 TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
This work was supported in part by a Charles Hood Foundation grant (to K. Badizadegan) and National Institutes of Health Grants DK-02907 (to K. Badizadegan), DK-48106 and DK-/AI-53056 (to W. I. Lencer), and DK-34854 (to the Harvard Digestive Diseases Center).


    FOOTNOTES
 

Address for reprint requests and other correspondence: K. Badizadegan, Dept. of Pathology, Massachusetts General Hospital, 55 Fruit St., WRN219, Boston, MA 02114 (E-mail: kbadizadegan{at}partners.org)

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.


    REFERENCES
 TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
1. Badizadegan K, Collier RJ, and Lencer WI. Membrane translocation by bacterial AB toxins. Methods Microbiol 31: 277–296, 2002.[ISI]

2. Badizadegan K, Dickinson BL, Wheeler HE, Blumberg RS, Holmes RK, and Lencer WI. Heterogeneity of detergent-insoluble membranes from human intestine containing caveolin-1 and ganglioside GM1. Am J Physiol Gastrointest Liver Physiol 278: G895–G914, 2000.[Abstract/Free Full Text]

3. Bastiaens PI, Majoul IV, Verveer PJ, Soling HD, and Jovin TM. Imaging the intracellular trafficking and state of the AB5 quaternary structure of cholera toxin. EMBO J 15: 4246–4253, 1996.[Abstract]

4. Brdickova N, Brdicka T, Andera L, Spicka J, Angelisova P, Milgram SL, and Horejsi V. Interaction between two adapter proteins, PAG and EBP50: a possible link between membrane rafts and actin cytoskeleton. FEBS Lett 507: 133–136, 2001.[CrossRef][ISI][Medline]

5. Bruses JL, Chauvet N, and Rutishauser U. Membrane lipid rafts are necessary for the maintenance of the {alpha}7 nicotinic acetylcholine receptor in somatic spines of ciliary neurons. J Neurosci 21: 504–512, 2001.[Abstract/Free Full Text]

6. Carreno S, Engqvist-Goldstein AE, Zhang CX, McDonald KL, and Drubin DG. Actin dynamics coupled to clathrin-coated vesicle formation at the trans-Golgi network. J Cell Biol 165: 781–788, 2004.[Abstract/Free Full Text]

7. Dharmsathaphorn K and Madara JL. Established intestinal cell lines as model systems for electrolyte transport studies. Methods Enzymol 192: 354–389, 1990.[Medline]

8. Edidin M. The state of lipid rafts: from model membranes to cells. Annu Rev Biophys Biomol Struct 32: 257–283, 2003.[CrossRef][ISI]

9. Engqvist-Goldstein AE and Drubin DG. Actin assembly and endocytosis: from yeast to mammals. Annu Rev Cell Dev Biol 19: 287–332, 2003.[ISI]

10. Foster LJ, De Hoog CL, and Mann M. Unbiased quantitative proteomics of lipid rafts reveals high specificity for signaling factors. Proc Natl Acad Sci USA 100: 5813–5818, 2003.[Abstract/Free Full Text]

11. Fujinaga Y, Wolf AA, Rodigherio C, Wheeler H, Tsai B, Allen L, Jobling M, Rapoport TA, Holmes RK, and Lencer WI. Gangliosides that associate with lipid rafts mediate transport of cholera and related toxins from the plasma membrane to endoplasmic reticulum. Mol Biol Cell 14: 4783–4793, 2003.[Abstract/Free Full Text]

12. Holmes RK, Jobling MG, and Connell TD. Cholera toxin and related enterotoxins of gram-negative bacteria. In: Bacterial Toxins and Virulence Factors in Disease, edited by Moss J, Iglewski B, Vaughan M, and Tu AT. New York: Dekker, 1995, p. 225–255.

13. Holowka D, Sheets ED, and Baird B. Interactions between Fc{epsilon}RI and lipid raft components are regulated by the actin cytoskeleton. J Cell Sci 113: 1009–1019, 2000.[Abstract/Free Full Text]

14. Holzinger A. Jasplakinolide. An actin-specific reagent that promotes actin polymerization. Methods Mol Biol 161: 109–120, 2001.[Medline]

15. Insel PA and Koachman AM. Cytochalasin B enhances hormone and cholera toxin-stimulated cyclic AMP accumulation in S49 lymphoma cells. J Biol Chem 257: 9717–9723, 1982.[Free Full Text]

16. Iwabuchi K, Handa K, and Hakomori S. Separation of "glycosphingolipid signaling domain" from caveolin-containing membrane fraction in mouse melanoma B16 cells and its role in cell adhesion coupled with signaling. J Biol Chem 273: 33766–33773, 1998.[Abstract/Free Full Text]

17. Jasper JR, Post SR, Desai KH, Insel PA, and Bernstein D. Colchicine and cytochalasin B enhance cyclic AMP accumulation via postreceptor actions. J Pharmacol Exp Ther 274: 937–942, 1995.[Abstract]

18. Kaksonen M, Sun Y, and Drubin DG. A pathway for association of receptors, adaptors, and actin during endocytic internalization. Cell 115: 475–487, 2003.[ISI][Medline]

19. Kanzaki M and Pessin JE. Caveolin-associated filamentous actin (Cav-actin) defines a novel F-actin structure in adipocytes. J Biol Chem 277: 25867–25869, 2002.[Abstract/Free Full Text]

20. Koopmann JO, Albring J, Huter E, Bulbuc N, Spee P, Neefjes J, Hammerling GJ, and Momburg F. Export of antigenic peptides from the endoplasmic reticulum intersects with retrograde protein translocation through the Sec61p channel. Immunity 13: 117–127, 2000.[CrossRef][ISI][Medline]

21. Kwik J, Boyle S, Fooksman D, Margolis L, Sheetz MP, and Edidin M. Membrane cholesterol, lateral mobility, and the phosphatidylinositol 4,5-bisphosphate-dependent organization of cell actin. Proc Natl Acad Sci USA 100: 13964–13969, 2003.[Abstract/Free Full Text]

22. Lamaze C, Dujeancourt A, Baba T, Lo CG, Benmerah A, and Dautry-Varsat A. Interleukin 2 receptors and detergent-resistant membrane domains define a clathrin-independent endocytic pathway. Mol Cell 7: 661–671, 2001.[ISI][Medline]

23. Lamaze C, Fujimoto LM, Yin HL, and Schmid SL. The actin cytoskeleton is required for receptor-mediated endocytosis in mammalian cells. J Biol Chem 272: 20332–20335, 1997.[Abstract/Free Full Text]

24. Lencer WI, Constable C, Moe S, Jobling MG, Webb HM, Ruston S, Madara JL, Hirst TR, and Holmes RK. Targeting of cholera toxin and Escherichia coli heat labile toxin in polarized epithelia: role of COOH-terminal KDEL. J Cell Biol 131: 951–962, 1995.[Abstract]

25. Lencer WI, Constable C, Moe S, Rufo PA, Wolf A, Jobling MG, Ruston SP, Madara JL, Holmes RK, and Hirst TR. Proteolytic activation of cholera toxin and Escherichia coli labile toxin by entry into host epithelial cells. Signal transduction by a protease-resistant toxin variant. J Biol Chem 272: 15562–15568, 1997.[Abstract/Free Full Text]

26. Lencer WI, Delp C, Neutra MR, and Madara JL. Mechanism of cholera toxin action on a polarized human intestinal epithelial cell line: role of vesicular traffic. J Cell Biol 117: 1197–1209, 1992.[Abstract]

27. Lencer WI, Moe S, Rufo PA, and Madara JL. Transcytosis of cholera toxin subunits across model human intestinal epithelia. Proc Natl Acad Sci USA 92: 10094–10098, 1995.[Abstract]

28. Lencer WI and Tsai B. The intracellular voyage of cholera toxin: going retro. Trends Biochem Sci 28: 639–645, 2003.[CrossRef][ISI][Medline]

29. Lisanti MP, Scherer PE, Vidugiriene J, Tang Z, Hermanowski-Vosatka A, Tu YH, Cook RF, and Sargiacomo M. Characterization of caveolin-rich membrane domains isolated from an endothelial-rich source: implications for human disease. J Cell Biol 126: 111–126, 1994.[Abstract]

30. London E and Brown DA. Insolubility of lipids in triton X-100: physical origin and relationship to sphingolipid/cholesterol membrane domains (rafts). Biochim Biophys Acta 1508: 182–195, 2000.[ISI][Medline]

31. Madara JL, Stafford J, Barenberg D, and Carlson S. Functional coupling of tight junctions and microfilaments in T84 monolayers. Am J Physiol Gastrointest Liver Physiol 254: G416–G423, 1988.[Abstract/Free Full Text]

32. Massol RH, Larsen JE, Fujinaga Y, Lencer WI, and Kirchhausen T. Cholera toxin toxicity does not require functional Arf6- and dynamin-dependent endocytic pathways. Mol Biol Cell 15: 3631–3641, 2004.[Abstract/Free Full Text]

33. Mirre C, Monlauzeur L, Garcia M, Delgrossi MH, and Le Bivic A. Detergent-resistant membrane microdomains from Caco-2 cells do not contain caveolin. Am J Physiol Cell Physiol 271: C887–C894, 1996.[Abstract/Free Full Text]

34. Moran M and Miceli MC. Engagement of GPI-linked CD48 contributes to TCR signals and cytoskeletal reorganization: a role for lipid rafts in T cell activation. Immunity 9: 787–796, 1998.[ISI][Medline]

35. Munro S. Lipid rafts: elusive or illusive? Cell 115: 377–388, 2003.[ISI][Medline]

36. Nabi IR and Le PU. Caveolae/raft-dependent endocytosis. J Cell Biol 161: 673–677, 2003.[Abstract/Free Full Text]

37. Nebl T, Pestonjamasp KN, Leszyk JD, Crowley JL, Oh SW, and Luna EJ. Proteomic analysis of a detergent-resistant membrane skeleton from neutrophil plasma membranes. J Biol Chem 277: 43399–43409, 2002.[Abstract/Free Full Text]

38. Oliferenko S, Paiha K, Harder T, Gerke V, Schwarzler C, Schwarz H, Beug H, Gunthert U, and Huber LA. Analysis of CD44-containing lipid rafts: recruitment of annexin II and stabilization by the actin cytoskeleton. J Cell Biol 146: 843–854, 1999.[Abstract/Free Full Text]

39. Orlandi PA. Protein-disulfide isomerase-mediated reduction of the A subunit of cholera toxin in a human intestinal cell line. J Biol Chem 272: 4591–4599, 1997.[Abstract/Free Full Text]

40. Orlandi PA and Fishman PH. Filipin-dependent inhibition of cholera toxin: evidence for toxin internalization and activation through caveolae-like domains. J Cell Biol 141: 905–915, 1998.[Abstract/Free Full Text]

41. Parton RG, Joggerst B, and Simons K. Regulated internalization of caveolae. J Cell Biol 127: 1199–1215, 1994.[Abstract]

42. Parton RG and Richards AA. Lipid rafts and caveolae as portals for endocytosis: new insights and common mechanisms. Traffic 4: 724–738, 2003.[CrossRef][ISI][Medline]

43. Pelkmans L, Puntener D, and Helenius A. Local actin polymerization and dynamin recruitment in SV40-induced internalization of caveolae. Science 296: 535–539, 2002.[Abstract/Free Full Text]

44. Pike LJ and Miller JM. Cholesterol depletion delocalizes phosphatidylinositol bisphosphate and inhibits hormone-stimulated phosphatidylinositol turnover. J Biol Chem 273: 22298–22304, 1998.[Abstract/Free Full Text]

45. Rodighiero C, Fujinaga Y, Hirst TR, and Lencer WI. A cholera toxin B-subunit variant that binds ganglioside GM1 but fails to induce toxicity. J Biol Chem 276: 36939–36945, 2001.[Abstract/Free Full Text]

46. Rothberg KG, Heuser JE, Donzell WC, Ying YS, Glenney JR, and Anderson RG. Caveolin, a protein component of caveolae membrane coats. Cell 68: 673–682, 1992.[ISI][Medline]

47. Roy S, Luetterforst R, Harding A, Apolloni A, Etheridge M, Stang E, Rolls B, Hancock JF, and Parton RG. Dominant-negative caveolin inhibits H-ras function by disrupting cholesterol-rich plasma membrane domains. Nat Cell Biol 1: 98–105, 1999.[CrossRef][ISI][Medline]

48. Rozelle AL, Machesky LM, Yamamoto M, Driessens MH, Insall RH, Roth MG, Luby-Phelps K, Marriott G, Hall A, and Yin HL. Phosphatidylinositol 4,5-bisphosphate induces actin-based movement of raft-enriched vesicles through WASP-Arp2/3. Curr Biol 10: 311–320, 2000.[CrossRef][ISI][Medline]

49. Sabharanjak S, Sharma P, Parton RG, and Mayor S. GPI-anchored proteins are delivered to recycling endosomes via a distinct cdc42-regulated, clathrin-independent pinocytic pathway. Dev Cell 2: 411–423, 2002.[ISI][Medline]

50. Schade AE and Levine AD. Lipid raft heterogeneity in human peripheral blood T lymphoblasts: a mechanism for regulating the initiation of TCR signal transduction. J Immunol 168: 2233–2239, 2002.[Abstract/Free Full Text]

51. Schmitz A, Herrgen H, Winkeler A, and Herzog V. Cholera toxin is exported from microsomes by the Sec61p complex. J Cell Biol 148: 1203–1212, 2000.[Abstract/Free Full Text]

52. Schnitzer JE, McIntosh DP, Dvorak AM, Liu J, and Oh P. Separation of caveolae from associated microdomains of GPI-anchored proteins. Science 269: 1435–1439, 1995.[ISI][Medline]

53. Shogomori H and Brown DA. Use of detergents to study membrane rafts: the good, the bad, and the ugly. Biol Chem 384: 1259–1263, 2003.[ISI][Medline]

54. Spangler BD. Structure and function of cholera toxin and the related Escherichia coli heat-labile enterotoxin. Microbiol Rev 56: 622–647, 1992.[ISI][Medline]

55. Stahlhut M and van Deurs B. Identification of filamin as a novel ligand for caveolin-1: evidence for the organization of caveolin-1-associated membrane domains by the actin cytoskeleton. Mol Biol Cell 11: 325–337, 2000.[Abstract/Free Full Text]

56. Stan RV, Roberts WG, Predescu D, Ihida K, Saucan L, Ghitescu L, and Palade GE. Immunoisolation and partial characterization of endothelial plasmalemmal vesicles (caveolae). Mol Biol Cell 8: 595–605, 1997.[Abstract]

57. Tsai B, Rodighiero C, Lencer WI, and Rapoport TA. Protein disulfide isomerase acts as a redox-dependent chaperone to unfold cholera toxin. Cell 104: 937–948, 2001.[ISI][Medline]

58. Valensin S, Paccani SR, Ulivieri C, Mercati D, Pacini S, Patrussi L, Hirst T, Lupetti P, and Baldari CT. F-actin dynamics control segregation of the TCR signaling cascade to clustered lipid rafts. Eur J Immunol 32: 435–446, 2002.[CrossRef][ISI][Medline]

59. Villalba M, Bi K, Rodriguez F, Tanaka Y, Schoenberger S, and Altman A. Vav1/Rac-dependent actin cytoskeleton reorganization is required for lipid raft clustering in T cells. J Cell Biol 155: 331–338, 2001.[Abstract/Free Full Text]

60. Vyas KA, Patel HV, Vyas AA, and Schnaar RL. Segregation of gangliosides GM1 and GD3 on cell membranes, isolated membrane rafts, and defined supported lipid monolayers. Biol Chem 382: 241–250, 2001.[ISI][Medline]

61. Watzl C and Long EO. Natural killer cell inhibitory receptors block actin cytoskeleton-dependent recruitment of 2B4 (CD244) to lipid rafts. J Exp Med 197: 77–85, 2003.[Abstract/Free Full Text]

62. Waugh MG, Lawson D, and Hsuan JJ. Epidermal growth factor receptor activation is localized within low-buoyant density, non-caveolar membrane domains. Biochem J 337: 591–597, 1999.[CrossRef][ISI][Medline]

63. Waugh MG, Lawson D, Tan SK, and Hsuan JJ. Phosphatidylinositol 4-phosphate synthesis in immunoisolated caveolae-like vesicles and low buoyant density non-caveolar membranes. J Biol Chem 273: 17115–17121, 1998.[Abstract/Free Full Text]

64. Wolf AA, Fujinaga Y, and Lencer WI. Uncoupling of the cholera toxin-GM1 ganglioside receptor complex from endocytosis, retrograde Golgi trafficking, and downstream signal transduction by depletion of membrane cholesterol. J Biol Chem 277: 16249–16256, 2002.[Abstract/Free Full Text]

65. Wolf AA, Jobling MG, Wimer-Mackin S, Ferguson-Maltzman M, Madara JL, Holmes RK, and Lencer WI. Ganglioside structure dictates signal transduction by cholera toxin and association with caveolae-like membrane domains in polarized epithelia. J Cell Biol 141: 917–927, 1998.[Abstract/Free Full Text]





This Article
Abstract
Full Text (PDF)
All Versions of this Article:
287/5/C1453    most recent
00189.2004v1
Alert me when this article is cited
Alert me if a correction is posted
Citation Map
Services
Email this article to a friend
Similar articles in this journal
Similar articles in ISI Web of Science
Similar articles in PubMed
Alert me to new issues of the journal
Download to citation manager
Search for citing articles in:
ISI Web of Science (1)
Google Scholar
Articles by Badizadegan, K.
Articles by Lencer, W. I.
Articles citing this Article
PubMed
PubMed Citation
Articles by Badizadegan, K.
Articles by Lencer, W. I.


HOME HELP FEEDBACK SUBSCRIPTIONS ARCHIVE SEARCH TABLE OF CONTENTS
Visit Other APS Journals Online
Copyright © 2004 by the American Physiological Society.