Membrane Lipid Organization Is Critical for Human Neutrophil Polarization
2003; Elsevier BV; Volume: 278; Issue: 12 Linguagem: Inglês
10.1074/jbc.m212386200
ISSN1083-351X
AutoresLynda M. Pierini, Robert J. Eddy, Michele Fuortes, Stéphanie Seveau, Carlo Casulo, Frederick R. Maxfield,
Tópico(s)S100 Proteins and Annexins
ResumoIn response to chemoattractants neutrophils extend an actin-rich pseudopod, which imparts morphological polarity and is required for migration. Even when stimulated by an isotropic bath of chemoattractant, neutrophils exhibit persistent polarization and continued lamellipod formation at the front, suggesting that the cells establish an internal polarity. In this report, we show that perturbing lipid organization by depleting plasma membrane cholesterol levels reversibly inhibits cell polarization and migration. Among other receptor-mediated responses, β2 integrin up-regulation was unaffected, and initial calcium mobilization was only partially reduced by cholesterol depletion, indicating that this treatment did not abrogate initial receptor-mediated signal transduction. Interestingly, cholesterol depletion did not prevent initial activation of the GTPase Rac or an initial burst of actin polymerization, but rather it inhibited prolonged activation of Rac and sustained actin polymerization. Collectively, these findings support a model in which the plasma membrane is organized into domains that aid in amplifying the chemoattractant gradient and maintaining cell polarization. In response to chemoattractants neutrophils extend an actin-rich pseudopod, which imparts morphological polarity and is required for migration. Even when stimulated by an isotropic bath of chemoattractant, neutrophils exhibit persistent polarization and continued lamellipod formation at the front, suggesting that the cells establish an internal polarity. In this report, we show that perturbing lipid organization by depleting plasma membrane cholesterol levels reversibly inhibits cell polarization and migration. Among other receptor-mediated responses, β2 integrin up-regulation was unaffected, and initial calcium mobilization was only partially reduced by cholesterol depletion, indicating that this treatment did not abrogate initial receptor-mediated signal transduction. Interestingly, cholesterol depletion did not prevent initial activation of the GTPase Rac or an initial burst of actin polymerization, but rather it inhibited prolonged activation of Rac and sustained actin polymerization. Collectively, these findings support a model in which the plasma membrane is organized into domains that aid in amplifying the chemoattractant gradient and maintaining cell polarization. methyl-β-cyclodextrin formyl Met-Leu-Phe 4-morpholineethanesulfonic acid phosphatidylinositol bisphosphate polymorphonuclear neutrophil monoclonal antibody tetramethylrhodamine isothiocyanate Over the past decade, compelling evidence has emerged that challenges the notion of the plasma membrane lipid bilayer as a homogeneous passive entity that merely provides a scaffold for protein-mediated signaling (1Pierini L. Maxfield F. Proc. Natl. Acad. Sci. U. S. A. 2001; 98: 9471-9473Crossref PubMed Scopus (55) Google Scholar, 2Maxfield F.R. Curr. Opin. Cell Biol. 2002; 14: 483-487Crossref PubMed Scopus (244) Google Scholar). This evidence suggests that certain lipids preferentially associate and form lateral heterogeneities in the membrane (3Brown D. London E. Annu. Rev. Cell Dev. 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These lipid domains have been postulated to serve as centers for some signal transduction processes by virtue of their copurification with signaling molecules following cell lysis with cold nonionic detergents and floatation on sucrose density gradients (3Brown D. London E. Annu. Rev. Cell Dev. Biol. 1998; 14: 111-136Crossref PubMed Scopus (2546) Google Scholar). Because the direct visualization of lipid domains has been elusive, it is thought that they typically exist as dynamic submicron-sized regions within the plasma membrane (6Kurzchalia T. Parton R. Curr. Opin. Cell Biol. 1999; 11: 424-431Crossref PubMed Scopus (512) Google Scholar, 8Varma R. Mayor S. Nature. 1998; 394: 798-801Crossref PubMed Scopus (1021) Google Scholar, 9Friedrichson T. Kurzchalia T. Nature. 1998; 394: 802-805Crossref PubMed Scopus (477) Google Scholar, 10Harder T. Scheiffele P. Verkade P. Simons K. J. Cell Biol. 1998; 141: 929-942Crossref PubMed Scopus (1044) Google Scholar). However, most of the surface area of cells such as fibroblasts and neutrophils is resistant to extraction by cold Triton X-100 (11Seveau S. Eddy R. Maxfield F. Pierini L. Mol. Biol. Cell. 2001; 12: 3550-3562Crossref PubMed Scopus (106) Google Scholar, 12Mayor S. Maxfield F. Mol. Biol. Cell. 1995; 6: 929-944Crossref PubMed Scopus (249) Google Scholar, 13Hao M. Mukherjee S. Maxfield F. Proc. Natl. Acad. Sci. U. S. A. 2001; 98: 13072-13077Crossref PubMed Scopus (250) Google Scholar), suggesting that lipid organization may be more complex than depicted in a simple version of the raft model. In fact, several studies now indicate that multiple types of membrane microdomains co-exist within the plasma membranes of cells (14Roper K. Corbeil D. Huttner W. Nat. Cell Biol. 2000; 2: 582-592Crossref PubMed Scopus (482) Google Scholar, 15Gomez-Mouton C. Abad J. Mira E. Lacalle R. Gallardo E. Jimenez-Baranda S. Illa I. Bernad A. Manes S. Martinez-A C. Proc. Natl. Acad. Sci. U. S. A. 2001; 98: 9642-9647Crossref PubMed Scopus (430) Google Scholar, 16Madore N. Smith K. Graham C. Jen A. Brady K. Hall S. Morris R. EMBO J. 1999; 18: 6917-6926Crossref PubMed Scopus (332) Google Scholar) and may coalesce into dense assemblies to form larger domain structures (1Pierini L. Maxfield F. Proc. Natl. Acad. Sci. U. S. A. 2001; 98: 9471-9473Crossref PubMed Scopus (55) Google Scholar,2Maxfield F.R. Curr. Opin. Cell Biol. 2002; 14: 483-487Crossref PubMed Scopus (244) Google Scholar, 11Seveau S. Eddy R. Maxfield F. Pierini L. Mol. Biol. Cell. 2001; 12: 3550-3562Crossref PubMed Scopus (106) Google Scholar, 15Gomez-Mouton C. Abad J. Mira E. Lacalle R. Gallardo E. Jimenez-Baranda S. Illa I. Bernad A. Manes S. Martinez-A C. Proc. Natl. Acad. Sci. U. S. A. 2001; 98: 9642-9647Crossref PubMed Scopus (430) Google Scholar). Cholesterol is the most abundant lipid component of the plasma membrane, and it plays an important role in lipid organization. Extraction of cholesterol from the plasma membrane using the synthetic cholesterol chelator methyl-β-cyclodextrin (MβCD)1 causes reorganization of lipids with preferences for both ordered and disordered domains (13Hao M. Mukherjee S. Maxfield F. Proc. Natl. Acad. Sci. U. S. A. 2001; 98: 13072-13077Crossref PubMed Scopus (250) Google Scholar). While the effects of cholesterol depletion on ordered, raft-like domains have been emphasized, additional effects on the overall organization of lipids also occur (13Hao M. Mukherjee S. Maxfield F. Proc. Natl. Acad. Sci. U. S. A. 2001; 98: 13072-13077Crossref PubMed Scopus (250) Google Scholar). Cholesterol depletion has been used to examine the role of lipid domains in a multitude of cellular processes, including cell migration (15Gomez-Mouton C. Abad J. Mira E. Lacalle R. Gallardo E. Jimenez-Baranda S. Illa I. Bernad A. Manes S. Martinez-A C. Proc. Natl. Acad. Sci. U. S. A. 2001; 98: 9642-9647Crossref PubMed Scopus (430) Google Scholar, 17Manes S. Mira E. Gomez-Mouton C. Lacalle R. Keller P. Labrador J. Martinez-A C. EMBO J. 1999; 18: 6211-6220Crossref PubMed Scopus (276) Google Scholar) and phagocytosis (18Shin J. Gao Z. Abraham S. Science. 2000; 289: 785-788Crossref PubMed Scopus (272) Google Scholar, 19Peyron P. Bordier C. N′Diaye E. Maridonneau-Parini I. J. Immunol. 2000; 165: 5186-5191Crossref PubMed Scopus (185) Google Scholar, 20Gatfield J. Pieters J. Science. 2000; 288: 1647-1650Crossref PubMed Scopus (472) Google Scholar). Disruption of lipid domain organization by MβCD treatment resulted in inhibition of motility of breast cancer-derived cells (17Manes S. Mira E. Gomez-Mouton C. Lacalle R. Keller P. Labrador J. Martinez-A C. EMBO J. 1999; 18: 6211-6220Crossref PubMed Scopus (276) Google Scholar) and T cells (15Gomez-Mouton C. Abad J. Mira E. Lacalle R. Gallardo E. Jimenez-Baranda S. Illa I. Bernad A. Manes S. Martinez-A C. Proc. Natl. Acad. Sci. U. S. A. 2001; 98: 9642-9647Crossref PubMed Scopus (430) Google Scholar), and abrogation of the phagocytosis of Escherichia coli by mast cells (18Shin J. Gao Z. Abraham S. Science. 2000; 289: 785-788Crossref PubMed Scopus (272) Google Scholar) and mycobacteria by macrophages (20Gatfield J. Pieters J. Science. 2000; 288: 1647-1650Crossref PubMed Scopus (472) Google Scholar) and neutrophils (19Peyron P. Bordier C. N′Diaye E. Maridonneau-Parini I. J. Immunol. 2000; 165: 5186-5191Crossref PubMed Scopus (185) Google Scholar). Because cell motility and phagocytosis are mechanistically related, with both processes involving directional extension of actin-rich membranes, it seems likely that their inhibition following domain disruption may have the same underlying molecular basis. Migration of neutrophils and other immune cells is contingent upon their ability to adopt a polarized morphology in response to chemotactic stimulation. In the presence of chemoattractants, neutrophils rapidly transform from roughly spherical resting cells to migratory ones with distinctive leading and trailing edges. Actin polymerization occurs almost exclusively at the leading edge, resulting in a dramatic accumulation of F-actin at just one end of the cells (21Fechheimer M. Zigmond S. Cell Motil. 1983; 3: 349-361Crossref PubMed Scopus (94) Google Scholar). Notably, this remarkable asymmetry occurs even when the external chemotactic signal is uniform, suggesting that at least one signaling step leads to an internal polarization of the cell. Given that chemoattractant receptors are typically distributed uniformly across the cell surface even after polarization (22Servant G. Weiner O. Herzmark P. Balla T. Sedat J. Bourne H. Science. 2000; 287: 1037-1040Crossref PubMed Scopus (729) Google Scholar), this internal signal occurs somewhere between receptor occupancy and actin polymerization. The exact point at which polarization is induced is not known. However, the possibilities have been narrowed by an elegant study in neutrophil-differentiated HL-60 cells using the recruitment of a GFP-tagged plekstrin homology domain to the plasma membrane as a readout for polarization (22Servant G. Weiner O. Herzmark P. Balla T. Sedat J. Bourne H. Science. 2000; 287: 1037-1040Crossref PubMed Scopus (729) Google Scholar). This study showed that amplification of polarization signals depends on one or more of the Rho GTPases, which anchor to the plasma membrane via lipid tails, and is downstream of the lipid products of phosphatidylinositol 3-kinase (22Servant G. Weiner O. Herzmark P. Balla T. Sedat J. Bourne H. Science. 2000; 287: 1037-1040Crossref PubMed Scopus (729) Google Scholar). In other words, amplification of polarization occurs at or near the plasma membrane. This, together with our finding that the plasma membrane of neutrophils segregates into distinct lipid domains that comprise either pole of migrating neutrophils (11Seveau S. Eddy R. Maxfield F. Pierini L. Mol. Biol. Cell. 2001; 12: 3550-3562Crossref PubMed Scopus (106) Google Scholar), led us to hypothesize that signal amplification is dependent on plasma membrane domains. To test this, we used MβCD to disrupt membrane domain organization in human neutrophils and studied its effects on polarization and signaling. Polymorphonuclear neutrophils (PMNs) were isolated from whole blood donated by healthy volunteers by centrifugation through Polymorphprep (Invitrogen). After lysis of contaminating erythrocytes by a 30-s hypotonic shock, cells were washed with phosphate-buffered saline (pH 7.4), and then resuspended in incubation buffer (150 mmNaCl, 5 mm KCl, 1 mm CaCl2, 1 mm MgCl2, 10 mm glucose, 20 mm HEPES, pH 7.4). For all experiments, PMNs were plated onto fibronectin-coated coverslip dishes for 5 min at 37 °C, and then stimulated with 10 nm formyl-Met-Leu-Phe (fMLF) for the indicated times. To visualize membrane domains, PMNs were plated and stimulated as above, and then extracted with ice-cold detergent. Specifically, cells were bathed in ice-cold cytoskeleton stabilizing buffer (CSB; 138 mm KCl, 3 mm MgCl2, 2 mm EGTA, 0.32m sucrose, 10 mm MES, pH 6) containing 0.5% Triton X-100 and 1× protease inhibitor mixture (BD PharMingen) for 30 min, and then washed with ice-cold CSB. Samples were fixed with 3.3% paraformaldehyde and 0.05% glutaraldehyde in phosphate-buffered saline for 10 min on ice, and then incubated sequentially with a monoclonal anti-CD44 (clone HERMES-3, American Type Culture Collection, Manassas, VA) and AlexaFluor 488-goat anti-mouse secondary antibody (Molecular Probes, Eugene, OR). Anti-CD44-labeled cells were imaged immediately after completion of the labeling protocol. Images were acquired with a Zeiss LSM510 laser scanning confocal microscope (Jena, Germany) or a Leica DMIRB widefield microscope, and then analyzed with MetaMorph image analysis software (Universal Imaging Corporation, Downington, PA). Plasma membrane cholesterol was depleted by incubating PMNs (1.2 × 107/ml) in incubation buffer containing 10 mmMβCD (Sigma) for 15 min at 37 °C. At the end of this 15-min treatment, cells were diluted to 2.4 × 106/ml with incubation buffer (depleted cells). The extent of cholesterol depletion was determined by an enzymatic method using a commercially available kit (Free cholesterol C; Wako Biochemicals, Osaka). The cholesterol content of MβCD-treated cells was measured to be 20.9 ± 3.6% (mean ± s.d., n = 5) less than that of control cells. To replete membrane cholesterol, cholesterol-depleted cells were incubated with 5 mm Chol-MβCD for 2.5 min at 37 °C (repleted cells). After stimulation with fMLF, cells were fixed with 3.3% paraformaldehyde in the presence of 0.25 mg/ml saponin for 5 min at room temperature. When it was important to preserve the labile pools of F-actin, excess fluorescently conjugated phalloidin (Molecular Probes, Eugene, OR) was included during fixation and throughout the remaining labeling steps. Rac was visualized with a mouse monoclonal antibody against Rac1 and Rac2 (clone 23A8, Upstate Biotechnology Inc., Lake Placid, NY), followed by an AlexaFluor 546-conjugated goat anti-mouse secondary antibody (Molecular Probes). Cells labeled for Rac were imaged by either confocal or wide-field fluorescence microscopy. Wide-field fluorescence images of all samples within one experiment were acquired under identical conditions and quantified using Metamorph image analysis software. Following background correction, the average fluorescence intensity per cell was measured for more than 100 cells per condition, and these measurements were normalized to the level of the unstimulated control cells. Cells were fixed in 2% buffered glutaraldehyde, washed with 0.1 m sodium cacodylate buffer pH 7.3, then postfixed in 1% osmium tetroxide in sodium cacodylate buffer and washed. Samples were then dehydrated through a graded ethanol series, freeze-dried, and sputter-coated with gold-palladium. Samples were examined at 20 kV using a JEOL 100 CX-II electron microscope fitted with an ASID-scanning unit, and photographs were recorded on Polaroid Type 55 P/N film. PMNs in suspension were stimulated with fMLF (10 nm) for 5 min at 37 °C, then simultaneously fixed, permeabilized, and labeled for F-actin by the addition of an equal volume of phosphate-buffered saline containing 6.6% paraformaldehyde, 0.1% glutaraldehyde, 0.5 mg/ml saponin, and 2 units/ml AlexaFluor 488 phalloidin. Cell-associated fluorescence was measured by flow cytometry (Beckman-Coulter XL, Fullerton, CA) for 2,000 cells per condition and from three separate experiments. Aliquots from each sample were removed and imaged by confocal microscopy. To verify the results in adherent cells, PMNs were plated, stimulated, fixed, and labeled with TRITC-conjugated phalloidin as described above. Wide-field images of fluorescent-labeled cells were obtained using a Leica DMIRB equipped with a 63 × 1.32 numerical aperture objective and a Princeton Instruments (Princeton, NJ) cooled CCD camera driven by MetaMorph Imaging System software. Following background correction, the average fluorescence intensity per cell was measured for over 150 cells per condition from three experiments on different days. The average fluorescence intensity per cell for cholesterol-depleted cells was 46 ± 8% (S.D.) of the value for control cells. Motility assays were performed as described (23Pierini L. Lawson M. Eddy R. Hendey B. Maxfield F. Blood. 2000; 95: 2471-2480Crossref PubMed Google Scholar). Briefly, motility of plated and stimulated PMNs was monitored for 4 min using a Leica DMIRB (Leica Mikroscopie und Systeme GmbH, Germany) set up for differential interference contrast microscopy. Time-lapse images were acquired with a cooled CCD camera driven by MetaMorph Imaging System software. Migrating PMNs were defined as those whose tail and leading lamella moved at least 7 μm from their initial starting position within 240 s. Separate dishes were used for each treatment, and at least three dishes were monitored for each treatment on at least three different days. To monitor changes in intracellular free calcium levels ([Ca2+]i), PMNs were loaded with the ratiometric fluorescent indicator, fura-2 as described (24Pierini L.M. Maxfield F.R. Guan J.-L. Signaling Through Cell Adhesion Molecules, CRC Methods in Signal Transduction Series. CRC Press, Boca Raton1999: 279-301Google Scholar). Briefly, cells were incubated in a 5 μm solution of acetoxymethyl ester-derivatized fura-2 (Fura-2/AM, Molecular Probes) for 40 min at room temperature. Cells were then washed and incubated for an additional 10 min at room temperature to allow cleavage of the ester. Cells were treated with MβCD or not, and then placed on ice until use. 2-ml suspensions of fura-2-loaded PMNs (1–1.5 × 106 cells/ml) were maintained at 37 °C in cuvettes with continual stirring throughout each experiment. [Ca2+]i was monitored with a SLM 8000C spectrofluorometer (Aminco, Urbana, IL) operated in dual excitation mode (excitation 340 and 380 nm, emission 510 nm). In the absence of stimulation, there was a negligible upward drift of the signal due to leakage of dye from the cells (not shown). To quantify and compare peak responses, fractional responses (F) for each sample were calculated as the change in the ratio value induced by addition of fMLF (i.e. the difference between the peak ratio value attained and the basal ratio value: I peak −I basal) divided by the maximum range of ratio values (25Grynkiewicz G. Poenie M. Tsien R. J. Biol. Chem. 1985; 260: 3440-3450Abstract Full Text PDF PubMed Scopus (80) Google Scholar). The maximum range of ratio values was determined for each sample by addition of 0.5% Triton X-100 at the end of each experiment to obtain the maximum possible ratio value (I max), followed by addition of 4 mmEGTA to obtain the minimum ratio value (I min). In summary, fractional responses were calculated using the formula in Equation 1. F=(Ipeak−Ibasal)/(Imax−Imin)Equation 1 To verify the ability of PMNs to upregulate their integrins in response to fMLF, PMNs were incubated at 37 °C with Fab fragments of an anti-β2 integrin antibody (IB4) that was directly conjugated to AlexaFluor 488 in the presence or absence of fMLF, as indicated. At various time points after the start of the incubation period, aliquots of cells were removed and diluted into ice-cold incubation buffer to stop exocytosis. Cell-associated fluorescence was monitored using a Beckman-Coulter XL analytical flow cytometer. To account for any nonspecific binding of the IB4 Fab, fluorescence intensity of PMNs incubated with an irrelevant control AlexaFluor 488-conjugated Fab was set to an arbitrary value, and all other samples were measured relative to this value. Each data point represents measurements from 2,000 cells, and the data as a whole are representative of experiments from three different days. During plating, neutrophils were incubated with 10 μm myosin light chain kinase inhibitor, ML-7 (26Eddy R. Pierini L. Matsumura F. Maxfield F. J. Cell Sci. 2000; 113: 1287-1298Crossref PubMed Google Scholar). Cells were then stimulated in the continued presence of ML-7 for the indicated times. In response to the chemoattractant, fMLF neutrophils undergo polarized morphological changes with actin-driven protrusion of a lamellipod at just one end of the cell. Because of the dramatic morphological and functional asymmetry exhibited by migrating cells, it has been suggested that the front and back of polarized cells may manifest different lipid environments (11Seveau S. Eddy R. Maxfield F. Pierini L. Mol. Biol. Cell. 2001; 12: 3550-3562Crossref PubMed Scopus (106) Google Scholar, 15Gomez-Mouton C. Abad J. Mira E. Lacalle R. Gallardo E. Jimenez-Baranda S. Illa I. Bernad A. Manes S. Martinez-A C. Proc. Natl. Acad. Sci. U. S. 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These findings, coupled with those from other laboratories (15Gomez-Mouton C. Abad J. Mira E. Lacalle R. Gallardo E. Jimenez-Baranda S. Illa I. Bernad A. Manes S. Martinez-A C. Proc. Natl. Acad. Sci. U. S. A. 2001; 98: 9642-9647Crossref PubMed Scopus (430) Google Scholar, 17Manes S. Mira E. Gomez-Mouton C. Lacalle R. Keller P. Labrador J. Martinez-A C. EMBO J. 1999; 18: 6211-6220Crossref PubMed Scopus (276) Google Scholar), provide compelling evidence that the lipid environments at the front and back of polarized PMNs are dissimilar and may represent micron-scale membrane domains. MβCD, a chelator of cholesterol, has been used to selectively deplete plasma membrane cholesterol and thereby alter lipid organization in several cell types (8Varma R. Mayor S. Nature. 1998; 394: 798-801Crossref PubMed Scopus (1021) Google Scholar, 13Hao M. Mukherjee S. Maxfield F. Proc. Natl. Acad. Sci. U. S. A. 2001; 98: 13072-13077Crossref PubMed Scopus (250) Google Scholar, 17Manes S. Mira E. 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To avoid the non-cholesterol related effects of long-term cholesterol depleting strategies, we used MβCD to acutely deplete cholesterol from PMNs. To determine if cholesterol depletion by MβCD would also disrupt the large-scale lipid organization found in PMNs, we visualized domain organization in control (Fig. 1, a–c) and MβCD-treated (Fig. 1, d–f) cells. Domain organization was revealed by extracting cells with cold Triton X-100 and then labeling the cells by indirect immunofluorescence for the transmembrane protein, CD44, which localizes to raft-like domains in PMNs (11Seveau S. Eddy R. Maxfield F. Pierini L. Mol. Biol. Cell. 2001; 12: 3550-3562Crossref PubMed Scopus (106) Google Scholar). Fig. 1 (a′–c′) shows the progression of domain organization as control cells polarize in response to fMLF. As we reported previously, rafts are evenly distributed around the periphery of unstimulated cells (Fig. 1 a′), form larger patches after 15–30 s of stimulation (Fig. 1 b′), and finally coalesce into a cap toward the cell rear (Fig. 1 c′) (11Seveau S. Eddy R. Maxfield F. Pierini L. Mol. Biol. Cell. 2001; 12: 3550-3562Crossref PubMed Scopus (106) Google Scholar). After the plasma membranes of cells have been cholesterol-depleted with MβCD (Fig. 1, d–f), the raft component CD44 is less well retained following detergent extraction; the average fluorescence intensity per cholesterol-depleted cell was decreased by 40.8% ± 3.2% (mean ± S.D.) compared with control cells. (The intensities of panels d′–f′ have been enhanced relative to panels a′–c′ to allow visualization of the CD44 distribution). In contrast to control cells, fMLF stimulation does not induce capping of CD44 in cholesterol-depleted cells; rather, the CD44 that is retained remains relatively uniformly distributed around the cell periphery (Fig. 1,d′–f′). That is, cholesterol depletion inhibits the redistribution of detergent-resistant raft components into a cap at the cell rear, indicating that MβCD treatment inhibits large scale lipid organization in addition to disrupting microdomains. Acute depletion of cholesterol by treatment with MβCD has dramatic effects on neutrophil polarization and migration (Fig.2). When cellular cholesterol is depleted by just ∼21% (see "Materials and Methods"), PMN migration is inhibited by >90% (Fig. 2 a), consistent with the report by Manes et al. (17Manes S. Mira E. Gomez-Mouton C. Lacalle R. Keller P. Labrador J. Martinez-A C. EMBO J. 1999; 18: 6211-6220Crossref PubMed Scopus (276) Google Scholar) that depletion of membrane cholesterol inhibits migration of MCF-7 cells. However, in contrast to the interpretation reported in Manes et al., we find that cholesterol depletion abolishes lamellipod formation (Fig.2 b, compare center to left panel). Inhibition of migration and polarization can be attributed to effects of cholesterol modulation as opposed to nonspecific effects of MβCD treatment since the ability to polarize and migrate is restored to previously cholesterol-depleted cells upon replenishing membrane cholesterol with cholesterol-chelated MβCD (chol-MβCD, Fig. 2,a and b, repleted). Because it has been reported, based on observation of cells by light microscopy, that cholesterol depletion does not affect membrane extension and ruffling (17Manes S. Mira E. Gomez-Mouton C. Lacalle R. Keller P. Labrador J. Martinez-A C. EMBO J. 1999; 18: 6211-6220Crossref PubMed Scopus (276) Google Scholar, 20Gatfield J. Pieters J. Science. 2000; 288: 1647-1650Crossref PubMed Scopus (472) Google Scholar), we further investigated whether lamellipod extension in PMNs was affected by changes in cholesterol levels. Scanning electron microscopy reveals that cholesterol-depleted cells are unable to form membrane extensions or ruffles in response to fMLF (compare Fig. 3, b toa). The lack of membrane ruffling in cholesterol-depleted cells is accompanied by an inhibition of stimulated actin polymerization. Following cholesterol depletion and stimulation with fMLF, cells were simultaneously fixed, permeabilized, and stained for F-actin with fluorescently conjugated phalloidin. Projections of confocal images show that control cells exhibited dramatic F-actin-rich ruffles within their lamellae (Fig. 3 c), whereas cholesterol-depleted cells had no F-actin-rich projections (Fig.3 d). Using flow cytometry, we quantified the fluorescent phalloidin binding to F-actin in parallel samples of suspended cholesterol-depleted and control cells, which had been stimulated with fMLF, or left unstimulated, for 5 min at 37 °C. Typically in r
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