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Published ahead of print on August 21, 2003, doi:10.1165/rcmb.2003-0079OC
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American Journal of Respiratory Cell and Molecular Biology. Vol. 30, pp. 233-241, 2004
© 2004 American Thoracic Society
DOI: 10.1165/rcmb.2003-0079OC

Lipid Raft Compartmentalization of Urokinase Receptor Signaling in Human Neutrophils

Robert G. Sitrin, Douglas R. Johnson, Pauline M. Pan, Donna M. Harsh, Jibiao Huang, Howard R. Petty and R. Alexander Blackwood

Pulmonary and Critical Care Medicine Division, Department of Internal Medicine, Department of Pediatrics and Communicable Diseases, and Department of Ophthalmology, University of Michigan, Ann Arbor, Michigan

Address correspondence to: Robert G. Sitrin, M.D., 6301 MSRB III, Box 0642, 1150 West Medical Center Dr., Ann Arbor, MI 48109-0642. E-mail: rsitrin{at}umich.edu


   Abstract
Top
 Abstract
Introduction
 Materials and Methods
 Results
 Discussion
 References
 
Urokinase plasminogen activator (uPA) receptors (uPAR) can be engaged for activation signaling either by aggregation or by binding exogenous uPA. These signaling mechanisms require uPAR to associate with two distinct adhesion proteins, L-selectin and complement receptor 3 (CR3), respectively. uPAR contains a glycosylphosphatidylinositol anchor, suggesting that it is concentrated within glycosphingolipid-enriched microdomains, or "lipid rafts". This study was undertaken to determine the extent to which uPAR-mediated signaling is compartmentalized to lipid rafts. Human neutrophil uPAR was cross-linked or stimulated with uPA after pretreatment with the lipid raft–disrupting agents, methyl-ß-cyclodextrin or filipin III. Both agents suppressed increases in intracellular Ca2+ concentrations ([Ca2+]i) triggered by cross-linking, but did not affect [Ca2+ ]i in response to uPA. Neutrophil membranes were separated into lipid raft and non-raft fractions, revealing the presence of uPAR and L-selectin, but the virtual absence of CR3 {alpha} chain in lipid rafts, either constitutively or in response to uPAR aggregation. Fluorescence resonance energy transfer experiments confirmed close proximity of a lipid raft marker to both uPAR and L-selectin, but not CR3. We conclude that uPAR can engage distinct signaling pathways involving different partner proteins that are functionally and physically segregated from one another in both lipid raft and non-raft domains of the plasma membrane.

Abbreviations: intracellular calcium concentration, [Ca2+]i • complement receptor 3, CR3 • fluorescein isothiocyanate, FITC • glycosylphosphatidylinositol, GPI • high molecular weight uPA, HMW-uPA • horseradish peroxidase, HRP • methyl-ß-cyclodextrin, MßCD • tetramethylrhodamine isothiocyanate, TRITC • urokinase plasminogen activator, uPA • urokinase plasminogen activator receptor, uPAR


   Introduction
Top
 Abstract
 Introduction
Materials and Methods
 Results
 Discussion
 References
 
Urokinase plasminogen activator (uPA) participates in many aspects of inflammation and tissue remodeling through its proteolytic functions, among them converting plasminogen to plasmin and activating latent growth factors (1). More recently, attention has been drawn to the uPA receptor (uPAR, CD87) expressed on leukocytes and many types of parenchymal cells. Its originally described function as a high-affinity docking site for uPA has given way to a burgeoning array of functions in activation signaling and lateral associations with multiple partner proteins on the plasma membrane (2, 3). These associations include interactions between uPAR and ß1, ß2, and ß3 integrin adhesion proteins whereby bidirectional communication influences the adhesive and signaling functions of both participants (2, 4, 5). Other partner proteins that associate with uPAR and participate in cellular signaling include the chemotactic receptor FPRL1/LXA4R, GP130, mannose-6-phosphate/insulin-like growth factor II receptor, uPAR associated protein/Endo-180, casein kinase 2, and nucleolin (3, 6–10).

The mechanism by which uPAR is engaged strongly influences its dependency on ligand (uPA) binding for signaling and the selection of its partner protein. Cao and coworkers showed previously that stimulating human neutrophils with exogenous uPA can trigger Ca2+ signaling and augmented superoxide release, and that these events require formation of a trimeric complex comprised of uPA, uPAR, and complement receptor 3 (CR3; Mac-1; CD11b/CD18), a ß2 integrin (5). We later showed that uPAR-mediated signaling in neutrophils can be also engaged through an entirely independent pathway whereby uPAR aggregation can elicit intracellular Ca2+ mobilization and proinflammatory effector functions, including degranulation, oxidant release, and upregulated expression of CR3 (11). The physiologic relevance of uPAR aggregation is supported by observations that clusters of uPAR form at the leading edges of polarized neutrophils (12) and at cell–substratum interfaces of fibrinogen-adherent monocytes (4), although the mechanisms causing uPAR to form clusters remain to be defined. In distinct contrast to signaling mechanisms triggered by exogenous uPA, signaling through uPAR aggregation does not require receptor occupancy with uPA, and there is an obligate partner protein relationship between uPAR and another adhesion protein, L-selectin, rather than CR3 (11, 13). Fluoresence resonance energy transfer experiments corroborated this relationship by documenting a direct physical association between uPAR and L-selectin (13). The signaling partnerships formed between uPAR and L-selectin versus uPAR and CR3 appear not to be interrelated, as we observed that signaling through uPAR aggregation was unaffected by blocking anti-CR3 mAbs (13) and conversely, a blocking anti–L-selectin mAb does not affect signaling triggered by exogenous uPA (unpublished observation).

The above findings raise several questions regarding uPAR function. Because uPA is bound to the plasma membrane by a glycosylphosphatidylinositol (GPI) anchor, it has no direct access to the cytoplasmic face of the plasma membrane to engage intracellular signaling intermediates. Although the mechanisms underlying cellular signaling through uPAR and other GPI-anchored proteins are not fully understood, one commonly accepted paradigm is that the membrane-spanning proteins with which they form complexes serve as signal transduction devices. Such lateral associations on the plasma membrane are clearly dynamic, as uPAR is known to exchange its association with CR3 in favor of CR4 (CD11c/CD18) as neutrophils become polarized (14). Little is known regarding the organizational schemes that determine how uPAR is distributed among the large array of potential partner proteins. uPAR structure is clearly heterogeneous by virtue of its variable glycosylation and the existence of both intact and truncated forms, and this certainly may influence its associations with partner proteins (1). Second, not all partner proteins may be expressed concurrently, thereby regulating the availability of these proteins to uPAR on a temporal basis. Lastly, uPAR partner proteins may be physically or functionally segregated from one another within the plasma membrane. GPI-anchored proteins are often concentrated (either constitutively or upon aggregation) in "lipid raft" microdomains within the plasma membrane that are enriched with glycosphingolipids, cholesterol, and numerous lipid-substituted signaling elements. Lipid rafts are thought to provide the infrastructure for bringing certain receptors and downstream signaling intermediates into proximity, permitting the formation of competent signaling assemblies (reviewed in Ref. 15). There are many instances whereby receptor redistribution into lipid rafts is necessary for certain signaling cascades to proceed (15). In this study, we sought to determine if distribution within lipid raft versus non-raft fractions of neutrophil plasma membranes constitutes a physical basis for functionally segregating signaling pathways involving uPAR and two of its potential signaling partners, CR3 versus L-selectin.


   Materials and Methods
Top
 Abstract
 Introduction
 Materials and Methods
Results
 Discussion
 References
 
Reagents
Purified murine IgG Fc fragments and F(ab')2 fragments of goat anti-murine IgG F(ab')2 were obtained from Jackson Immunoresearch Laboratories (West Grove, PA). A hybridoma producing anti-human uPAR mAb (clone 3B10) was generously provided by Robert F. Todd III, M.D., Ph.D., University of Michigan Health System. The hybridoma cells were cultured in vitro and the IgG2a mAb purified by Protein A Sepharose (Antibody Solutions, Palo Alto CA). This mAb recognizes an epitope near the uPA binding site, and thus preferentially binds to unoccupied uPAR (16). High molecular weight uPA (HMW-uPA) was generously provided by Jack Henkin, Abbott Laboratories (North Chicago IL). Anti–L-selectin IgG1 mAb was purified from cultures of the Dreg-56 hybridoma (American Type Culture Collection, Manassas, VA). Methyl-ß-cyclodextrin (MßCD) was obtained from Sigma (St. Louis, MO), and filipin III was obtained from Cayman Chemical Co. (Ann Arbor, MI).

Purification and Stimulation of Human Neutrophils
Neutrophils were isolated from peripheral blood obtained from healthy volunteers according to provisions of the University of Michigan Institutional Review Board for Human Subject Research. Briefly, anticoagulated blood was sedimented with 6% dextran/0.9% NaCl, the erythrocytes were removed by hypotonic lysis, and neutrophils were isolated by density gradient centrifugation on a 10% ficoll-Hypaque cushion. The resulting cells (> 95% neutrophils) were washed three times in phosphate-buffered saline and transferred to the appropriate buffer for further studies. To cross-link uPAR, cells were suspended in Buffer A (145 mM NaCl, 5 mM KCl, 1 mM MgCl2, 10 mM glucose, 1 mM CaCl2, 1% wt/vol bovine serum albumin, 10 mM Hepes, pH 7.4). Cells were first incubated with murine IgG Fc fragments (150 µg/ml) at 4°C for 15 min to block binding of primary mAbs to Fc receptors. The effectiveness of this blocking step has been demonstrated previously (16). Cells were then incubated with anti-uPAR mAb (100 µg/ml) at 4°C for 30 min, washed in Buffer A, and to initiate receptor cross-linking, the F(ab')2 goat anti-murine F(ab')2 was added after warming the cells to 37°C. Essentially identical protocols for cross-linking human neutrophil uPAR with the 3B10 mAb produce visible receptor capping by fluorescence microscopy in approximately one-half of labeled cells (5). For uPAR cross-linking before membrane fractionation (see below), the cells were treated with cytochalasin B (5 µg/ml for 5 min) before adding the cross-linking Ab. Alternatively, freshly purified neutrophils were stimulated with HMW-uPA (2 µM final concentration). uPAR and HMW-uPA recognize each other in monomeric fashion, so saturating uPAR with HMW-uPA cannot directly induce uPAR aggregation (1).

Fractionation of Lipid Raft and Non-Raft Plasma Membranes
Fractionation of neutrophil membranes into lipid raft and non-raft fractions was performed using a standard method (with minor modifications) based on the low buoyant density of lipid rafts and their relative insolubility in Triton X-100 (17). Cells were treated with cytochalasin B (as above) to minimize the possibility that lipid raft constituents would be retained in higher density fractions during purification because of their association with cytoskeletal elements. Neutrophils were stimulated as described (108 cells per condition), and immediately collected by centrifugation in Buffer B (150 mM NaCl, 1 mM EDTA, 25 mM Hepes, pH 7.0, with 1.4 µg/ml pepstatin A, 1.4 µg/ml leupeptin, 1 µg/ml aprotinin, 4 mM iodoacetic acid, and 1 mM PMSF) at 4°C. The cells were then lysed in Buffer B with DFP 2.5 mM, sodium orthovanadate 1 mM, and 1% triton X-100 for 30 min at 4°C. The lysates were then mixed 1:1 vol/vol with 80% sucrose and layered under discontinuous sucrose gradients of 40–30–10%. After centrifugation at 25,000 x g for 20 h, the fractions were removed in 1-ml volumes. The lipid raft (detergent-resistant, low density) fractions were collected at the 30–10% interface, and the non-raft (high density, detergent-sensitive) fractions were removed from the 40% layer. Protein concentrations were determined with a micro-BCA assay (Pierce, Rockford IL).

Immunoblotting
Aliquots of each fraction were mixed 3:1 with 4x sample buffer (0.25 M Tris, pH 6.8, 31% glycerol, 8% SDS) and boiled for 5 min. Samples were electrophoresed on 8–16% gradient polyacrylamide gels under nonreducing conditions, transferred to PDVF membranes in standard fashion, labeled as indicated, and developed by enhanced chemiluminescence. Labeling antibodies and reagents: rabbit anti-human uPAR (American Diagnostica, Greenwich, CT), horseradish peroxidase (HRP)-conjugated donkey anti-rabbit (Jackson), sheep anti-human L-selectin (R&D Systems, Minneapolis MN), HRP-conjugated goat anti-sheep (Jackson), goat anti-human CR3 {alpha} chain (Santa Cruz Biotechnology, Santa Cruz, CA), HRP-conjugated donkey anti-goat (Santa Cruz), biotinylated anti-CD45 (Pharmingen, San Diego CA), HRP-conjugated streptavidin (Jackson), peroxidase-conjugated cholera toxin B subunit (a specific ligand of GMI ganglioside; Sigma). Densitometry measurements were obtained with a Chemi Doc Imaging System and analyzed with Quantity One Software v.4.2.1 (Bio-Rad, Hercules, CA).

Immunofluorescence Flow Cytometry
Cells were resuspended in labeling buffer (phosphate-buffered saline with 0.1% human {gamma}-globulin, 0.1% glucose, pH 7.4) and incubated with anti-uPAR (3B10) or anti–L-selectin (Dreg 56) primary mAb for 30 min, 4°C, followed by phycoerythrin (PE)-conjugated goat anti-mouse Ab (30 min, 4°C). For negative controls, cells are stained with secondary mAb alone, and with an irrelevant isotype-matched primary Ab. Fluorescence intensity was assessed as a measure of relative antigen expression using an EPICS Elite ESP flow cytometer (Coulter, Miami, FL; University of Michigan Flow Cytometry Core Facility). Mean fluorescence intensities (linear scale) were determined from >= 10,000 cells, and specific fluorescence intensities were calculated by subtracting the corresponding value of the nonspecific control. To maintain consistent results between experiments, the flow cytometer was adjusted to provide constant fluorescence intensities for Coulter Standard Brite beads.

Measurement of Intracellular Calcium Concentration
Cells were loaded (5 x 106/ml) with the Ca2+-sensitive fluorescent dye Fluo-3/AM (2 µM; Molecular Probes, Eugene, OR) at 30°C for 30 min in 145 mM NaCl, 5 mM KCl, 1 mM MgCl2, 10 mM glucose, 4 mM probenecid, 10 mM Hepes, pH 7.4. After pretreatments as indicated, 2.5 x 106 cells were suspended in 1 ml incubation buffer and prewarmed to 37°C. Fluorescence intensities were then measured with a SLM8000 spectrofluorometer equipped with SLM Spectrum Processor v3.5 software (SLM Instruments Inc., Urbana, IL), using a 1-cm light path cuvette at an excitation wavelength of 505 nm and an emission wavelength of 530 nm. Fluorescence intensities were acquired at 2-s intervals for 300 s with continuous stirring of the cell suspension. These measurements were converted to nanomolar concentrations of intracellular Ca2+ concentration ([Ca2+]i) by the calibration method of Grynkiewycz and colleagues (11, 18).

Fluorescence Resonance Energy Transfer
Cholera toxin B subunit conjugated with fluorescein isothiocyanate (FITC) or tetramethylrhodamine isothiocyanate (TRITC) were obtained from Sigma and List Biological Laboratories (Campbell, CA), respectively. FITC-labeled anti–anti-CD45 and anti–L-selectin were obtained from Biosource (Camarillo, CA). Anti-uPAR mAb (3B10) was TRITC-labeled and CR3 {alpha} chain was FITC labeled with standard labeling kits (Molecular Probes) and purified by gel chromatography. An axiovert-inverted fluorescence microscope with HBO-100 mercury illumination (Carl Zeiss, New York, NY) interfaced to a Dell 410 workstation via a Scion SG-7 video card (Vay Tek, Fairfield, IA) was used. A narrow bandpass-discriminating filter set was used with excitation at 485DF20 nm and emission of 530DF30 nm for FITC. For TRITC, an excitation of 540DF20 nm and an emission of 590DF30 nm were used (Omega Optical, Brattleboro, VT). Long-pass dichroic mirrors at 510 and 560 nm were used for FITC and rhodamine, respectively. Single-cell spectra were obtained using a imaging spectrophotometer system. Labeled cells were illuminated with an excitation filter at 485DF22 nm and a 510LP dichroic mirror for FITC emission spectra and resonance energy transfer experiments (13, 19). The emission spectra were obtained with an Acton-150 (Acton Research Corp, Acton, MA) imaging spectrophotometer fiber-optically coupled to a microscope. The exit port of the spectrophotometer was attached to a Gen-II intensifier coupled with an I-MAX-512 camera (Princeton Instruments, Trenton, NJ). Spectra collection was controlled by a high-speed Princeton ST-133 interface and a Stanford Research Systems DG-535 delay-gate generator (Sunnyvale, CA), and analyzed with Winspec software (Princeton Instruments).

Statistical Analysis
Comparisons of means were performed with t tests using a P value of 0.05 to determine significance (GraphPad Prism version 3.00 for Windows; GraphPad Software, San Diego, CA).


   Results
Top
 Abstract
 Introduction
 Materials and Methods
 Results
Discussion
 References
 
Effects of Lipid Raft–Disrupting Agents on uPAR-Mediated Signaling
Freshly purified neutrophils were stimulated either by antibody-mediated uPAR aggregation, or by stimulation with HMW-uPA (2 µM). To disrupt lipid rafts, cells were treated with MßCD, a cyclic heptasaccharide that binds cholesterol with high specificity and extracts it from plasma membranes, thereby destabilizing the highly ordered packing of glycosphingolipids and causing lipid rafts to dissipate (15, 20). Cells were pretreated with 10 mM MßCD for 5 min, added just before the cross-linking Ab or just before stimulation with HMW-uPA. As shown in Figure 1A, MßCD pretreatment completely suppressed the flux in [Ca2+]i induced by uPAR aggregation. By contrast, pretreating with MßCD under identical conditions had no significant effect on the [Ca2+]i flux triggered by exogenous HMW-uPA.



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  		Figure 1. Effects of lipid raft disruption on uPAR-mediated Ca2+ signaling. (A) Inhibition of uPAR-mediated Ca2+ signaling by MßCD. Left columns represent pooled results indicating inhibition by MßCD (10 mM, 5 min) of the flux in [Ca2+]i in response to uPAR cross-linking (uPAR XL), and the absence of an effect after stimulation with HMW-uPA. ({Delta} [Ca2+]i, the difference between baseline and peak, expressed as % inhibition relative to control cells incubated without MßCD; mean ± SEM, n = 4). Right: representative continuous tracings of [Ca2+]i over time after uPAR XL (top) or addition of HMW-uPA (bottom), with and without MßCD. Arrows indicate the time that cross-linking antibody or HMW-uPA was added. (B) Inhibition of uPAR-mediated Ca2+ signaling by filipin III. Columns represent pooled results indicating inhibition by filipin III (0.8 µg/ml, 5 min) of the flux in intracellular Ca2+ concentrations in response to uPAR cross-linking (uPAR XL), and the absence of an effect after stimulation with HMW-uPA. Data are expressed as in A.

 
To confirm that the effects of MßCD were mediated by its cholesterol-binding properties, these experiments were repeated with a chemically unrelated cholesterol chelator, filipin III (21). Filipin III (0.8 µg/ml) had an effect similar to that of MßCD, significantly reducing the Ca2+ flux triggered by uPAR cross-linking without affecting the Ca2+ flux triggered by exogenous HMW-uPA (Figure 1B). Filipin III was less effective than MßCD at suppressing the response to uPAR cross-linking, but its use was limited by toxic effects on the neutrophils observed at higher concentrations. These results suggest that the importance of lipid raft integrity is highly specific to the mechanism by which uPAR is engaged, and accordingly, may be partly related to the distinct partner proteins, L-selectin versus CR3, incorporated in these signaling pathways. These results also strongly suggest that MßCD and filipin III interrupt the assembly of specific signaling complexes within the plasma membrane and do not interfere universally with downstream signaling events leading to Ca2+ mobilization. The membrane fractionation experiments (see below) included pretreatments with cytochalasin B, so it was also confirmed that its inclusion did not interfere with Ca2+ signaling in response to HMW-uPA. The difference in [Ca2+]i between peak and baseline ({Delta}[Ca2+]i) in the presence of cyto B (5 µg/ml) was 113.9 ± 3.6% of controls without cyto B (mean ± SEM, n = 3). In response to uPAR cross-linking, the {Delta}[Ca2+]i in the presence of cyto B was 97.8 ± 18.3% of controls without cyto B (mean ± SEM, n = 8). In other preliminary experiments (not shown), it was confirmed that under identical conditions, MßCD did not affect the magnitude of the [Ca2+]i flux induced by the chemotactic peptide FMLP, further indicating that there was no nonspecific effect on Ca2+ mobilization. Furthermore, flow cytometry demonstrated that MßCD did not significantly affect plasma membrane expression of uPAR or L-selectin (Figure 2), confirming that loss of uPAR and/or L-selectin upon raft disruption does not explain the inhibitory effect of MßCD.



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  		Figure 2. MßCD (10 mM, 5 min) does not alter plasma membrane expression of uPAR or L-selectin. Data are expressed as mean fluorescence intensity (arbitrary units, background subtracted). Mean ± SEM, n = 4. (t tests; all P > 0.05).

 
Demonstration of uPAR and L-Selectin in Lipid Raft Fractions
The essential role for plasma membrane cholesterol, and presumably lipid raft integrity, for Ca2+ mobilization in response to uPAR cross-linking fosters the hypothesis that uPAR and L-selectin are present in lipid raft microdomains either constitutively or in response to its aggregation. To address this issue, neutrophils were subjected to lysis with 1% Triton X-100 and discontinuous sucrose gradient centrifugation to separate lipid raft microdomains from the dense, detergent-soluble (non-raft) membranes. The validity of the fractionation protocol is demonstrated in Figure 3, as fraction 7 was markedly enriched in the ganglioside GM1, a major constituent of lipid rafts. By contrast, CD45, a raft-excluded protein, is easily demonstrable in the non-raft membrane fraction but absent from the lipid raft fraction (22, 23). The protein content of the lipid raft fractions was considerably smaller than the non-raft membranes (29.1 ± 4.6 and 73.4 ± 18.6 mg/ml, respectively).



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  		Figure 3. Characterization of lipid raft and non-raft membrane fractions of unstimulated neutrophils. Cells were solubilized with 1% Triton X-100 and separated by density gradient centrifugation as described in MATERIALS AND METHODS. (Insert, upper right) To confirm that fraction 7 is enriched in lipid rafts relative to non-raft membranes (fractions 1–4), equal volumes were loaded onto PDVF membrane using a slot-blot apparatus and probed with peroxidase-conjugated cholera toxin B subunit, a specific ligand for the ganglioside GM1. Western blots for CD45 were performed on the same fractions. The lipid raft fraction is characterized by enrichment with GM1 and exclusion of CD45, relative to non-raft membranes. Western blots for uPAR (top panel), L-selectin (middle panel), and CR3 {alpha} chain (bottom panel) were performed on all fractions of the sucrose density gradient. The preponderance of all these proteins was found in fractions 1–4, containing non-raft membranes. Significant but smaller amounts of uPAR and L-selectin were found in fraction 7, whereas no CR3 {alpha} chain was found in this fraction. The results shown are representative of four independent experiments.

 
Immunoblots for uPAR were performed on fractions of unstimulated neutrophils (Figure 3A). To properly compare the overall distributions of specific proteins between lipid raft versus non-raft membrane fractions, all Western blots were performed by loading equal volumes of sample (representing the yields of comparable numbers of cells) per lane rather than normalizing to protein content. Most uPAR was found in fractions 1–4 (40% sucrose) containing non-raft plasma membrane, although a significant portion was also seen in fraction 7 (drawn from the 10%/30% interface) containing lipid rafts.

Given that L-selectin plays an obligate role in Ca2+ mobilization induced by uPAR aggregation (13), it follows then that L-selectin should also be found in the lipid raft fraction. The distribution of L-selectin closely paralleled the distribution of uPAR, as most was found in the non-raft membrane fractions, whereas a smaller but significant portion was detected in fraction 7 (Figure 3).

The distribution of CR3 {alpha} chain was quite distinct from uPAR and L-selectin, in that none at all was found in fraction 7 (Figure 3). The vast majority was seen in the non-raft membrane fractions, whereas smaller amounts were seen in the intermediate density fractions. Although it is possible that fractions of intermediate density contain some lipid raft material, there clearly was no demonstrable co-isolation of CR3 with uPAR outside of the non-raft membrane fractions 1–4.

To corroborate the association of uPAR and L-selectin with lipid rafts, fluorescence resonance energy transfer experiments were performed with dual-labeled, unstimulated neutrophils, using GM1 as a marker for lipid rafts (Figure 4). GM1 was labeled with cholera toxin B subunit, which binds GM1 with high specificity (20, 24). The various fluorescent-labeled antibodies were as described in MATERIALS AND METHODS. When cells were illuminated at 485 nm to excite the donor chromophore, robust acceptor emission at 590 nm (evidence of resonance energy transfer) was observed in the presence of both the donor and acceptor for both GM1/uPAR and GM1/L-selectin labeling pairs (Figures 4A and 4B). As shown, resonance energy transfer was proportionate to the donor/acceptor-labeling ratio of the cells. There was no 590 nm emission in the presence of the donor alone (not shown). Resonance energy transfer is indicative of molecular proximity between fluorochromes within a resolution limit of 7 nm, consistent with a direct "nearest neighbor" interaction, so this degree of proximity places the labeled uPAR and L-selectin within lipid rafts. By contrast, no resonance energy transfer could be detected between GM1 and CR3 {alpha} chain (Figure 4C). Likewise, no resonance energy transfer was seen in the negative control cells dual-labeled for GM1 and lipid raft-excluded CD45 (Figure 4D).



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  		Figure 4. Fluorescence resonance energy transfer. Representative emission spectra of labeled neutrophils stained with FITC-conjugated (donor) and TRITC-conjugated (acceptor) labels. GM1 ganglioside, the lipid raft marker, was labeled with cholera toxin B subunit, and the proteins were labeled with fluorescent antibodies, as described in MATERIALS AND METHODS. A shows substantial resonance energy transfer (arrows) between GM1 (the lipid raft marker) and uPAR on a dual-labeled cell, and B shows similar resonance energy transfer between GM1 and L-selectin. Each group of emission spectra demonstrates the dose–response relationship of resonance energy transfer between GM1/uPAR and GM1/L-selectin with a range of donor/acceptor-labeling ratios. C shows no demonstrable resonance energy transfer on a cell dual-labeled for GM1 and CR3 {alpha} chain, using a donor/acceptor ratio of 1:2 (solid line). The dotted line represents an emission spectrum of a cell with the FITC label alone. D shows no demonstrable resonance energy transfer between GM1 and lipid raft-excluded CD45, using a donor/acceptor ratio of 1:2 (solid line). The dotted line represents an emission spectrum of a cell with the FITC label alone. Results shown are representative of three independent experiments.

 
The amount of uPAR in the lipid raft fractions increased rapidly after uPAR cross-linking (Figure 5). Densitometry of immunoblots obtained from four separate experiments demonstrate significant increases in uPAR, peaking in the lipid raft fractions 30 s after uPAR cross-linking. This was very transient, as uPAR already returned to basal levels by 60 s. The accumulation of uPAR in the lipid rafts coincides closely with the onset of the Ca2+ flux induced by uPAR cross-linking (11). The levels of L-selectin in the lipid rafts did not increase significantly after uPAR aggregation. CR3 {alpha} chain was virtually undetectable in lipid raft fractions, with or without uPAR aggregation.



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  		Figure 5. Changes in lipid raft constituents after uPAR cross-linking. (A) Top: uPAR demonstrated in the lipid raft fractions of unstimulated cells (time 0), and at 30 and 60 s after uPAR cross-linking. Bottom: corresponding densitometries of four pooled experiments, indicating a significant increase in uPAR at 30 s. Densitometry measurements are expressed as % of the time 0 control (mean ± SEM, P values determined by t tests). (B) Levels of L-selectin in the lipid raft fraction did not change significantly after uPAR cross-linking. Data are expressed as in A. (C) Virtually no CR3 {alpha} chain was demonstrable in lipid rafts even after uPAR cross-linking. (Note: the lipid raft fractions for these experiments were concentrated, so the amount of lipid raft material per lane is 10-fold greater than in the results shown in Figure 3.)

 

   Discussion
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
References
 
Our prior work has demonstrated that uPAR can initiate two independent signaling pathways that are distinctive by the method of receptor engagement (receptor cross-linking versus ligand binding with HMW-uPA) and the corresponding obligate partner adhesion proteins (L-selectin versus CR3) (5, 11, 13, 16). In this report, the dissimilarity between these two pathways is extended to demonstrate a physical basis for their compartmentalization within the plasma membrane. The first series of experiments show that the pathway initiated by uPAR cross-linking and resulting in increased [Ca2+]i is critically dependent on plasma membrane cholesterol (Figure 1). MßCD, which extracts cholesterol from the plasma membrane, completely abrogates the resulting Ca2+ flux, whereas filipin III, a cholesterol sequestering agent, is partially effective under conditions that preserve cell viability. As cholesterol is critically required to maintain the integrity of lipid raft microdomains, these agents have been used extensively to investigate the role of lipid rafts in multiple signaling pathways (15, 20, 21, 25). It is likely that MßCD and filipin III affect the formation of a competent signaling assembly rather than inhibiting downstream Ca2+ mobilization. MßCD and filipin III are known to affect intracellular Ca2+ homeostasis very selectively. Neither affects the [Ca2+]i flux triggered by cross-linking human neutrophil Fc{gamma}RIIa, which mobilizes Ca2+ from intracellular stores, but both block Ca2+ influx through plasma membrane channels (21). We have previously shown that uPAR cross-linking primarily triggers mobilization of Ca2+ from intracellular stores, with only a secondary influx of Ca2+ from the extracellular milieu (16). In that study, depleting [Ca2+]i completely abrogated the response to uPAR cross-linking just as MßCD did, whereas reducing extracellular Ca2+ had only a modest effect. In the present study, neither MßCD nor filipin III had any effect at all on the robust Ca2+ response to exogenous HMW-uPA (Figure 1), further attesting to (i) the absence of an adverse effect of either agent on cellular viability, and (ii) the specificity of the response to uPAR cross-linking, rather than a more global effect on uPAR function or [Ca2+]i homeostasis.

The larger implication of the disparate effects of cholesterol binding on the two uPAR-mediated signaling pathways is that clustering uPAR with L-selectin as its signaling partner absolutely requires the integrity of lipid raft microdomains, whereas the uPAR/HMW-uPA/CR3 trimeric complex engages signaling mechanisms that function independently of lipid rafts. This conventional interpretation is hampered by the recent observation that lipid raft microdomains can be heterogeneous and differentially sensitive to cholesterol sequestration (26). Thus, to attribute the effects of cholesterol depletion to the function of lipid rafts, we sought further corroboration by directly examining the distribution of uPAR, L-selectin, and CR3 in lipid raft versus non-raft membranes. The results in Figures 3 and 4 clearly show that subpopulations of uPAR and L-selectin are co-localized in the lipid raft fractions of unstimulated neutrophils. After uPAR cross-linking, uPAR accumulates further during the same time frame as the corresponding Ca2+ flux, whereas L-selectin levels remain fairly constant (Figure 5). Combined with the inhibitory effects of MßCD and filipin III, this indicates that uPAR and L-selectin physically located within lipid rafts are required for a competent signaling assembly to form in response to uPAR aggregation. However, we cannot yet conclude whether the signaling pathway only requires the uPAR present constitutively, or whether accumulation of uPAR in lipid rafts is also required. It is important to recognize that the composition of lipid rafts is dynamic, and the increased levels of uPAR could represent physical redistribution into lipid rafts from non-raft membrane domains, but may also be explained by longer retention times or greater stability of these proteins within the lipid raft fractions. The presence of L-selectin in lipid rafts has not been reported previously, but the present study demonstrates that even a relatively small quantity of L-selectin in lipid rafts can be functionally significant. It is well established that L-selectin engagement can trigger neutrophil activation, and it will be important for future studies to determine if localization within lipid rafts is an important feature of any of the relevant signaling mechanisms (27).

The Ca2+ signal elicited by the uPAR/HMW-uPA/CR3 complex was unaffected by depleting or binding membrane cholesterol (Figure 1), and consistent with this observation, CR3 (as determined by its {alpha} chain) was virtually undetectable in lipid rafts, in either unstimulated or stimulated conditions (Figures 3–5). Prior studies have not established the extent to which CR3 may localize to lipid rafts. Neutrophil CR3 has been found in large detergent-resistant complexes, but it is not clear that these complexes represented lipid rafts (28). Nonopsonic phagocytosis by human neutrophils has been reported to require CR3, GPI-linked protein(s), and lipid raft integrity, but membrane fractionation studies were not performed to establish that the CR3 was actually present in lipid rafts (29). Other integrins, including LFA-1 ({alpha}Lß2), {alpha}4ß1, and {alpha}vß3, mobilize to lipid raft compartments upon activation, so it is entirely possible that CR3 will also redistribute to lipid rafts in response to other stimuli, whereas cross-linking a relatively sparse protein such as uPAR may be insufficient to redistribute CR3 (17, 22, 24).

It is well established that in parenchymal cells, uPAR can be found in or near caveolae, which are specialized structures that contain lipid rafts along with a signature protein, caveolin (7, 30). Other lipid raft constituents include signaling molecules such as Jak and src kinases that are known to be associated with uPAR-mediated signaling (7, 15, 30, 31). Given that caveolae, like lipid rafts in general, are enriched in signaling molecules, it has been presumed that they serve as signaling platforms for uPAR-mediated activation signaling (15). Wei and coworkers have shown that in 293 cells, uPAR stabilizes caveolin/ß1 integrin complexes (30, 32). However, it remains highly arguable whether neutrophils express caveolin at all (33–35). Therefore, our study raises the possibility that activation signaling through uPAR may also proceed in lipid rafts through mechanisms unrelated to caveolin. The relatively small proportion of membrane uPAR we observed in the lipid raft fractions is somewhat surprising, given the propensity for uPAR to localize to lipid rafts/caveolae in other cell types. However, this has not been addressed in leukocytes, and it is possible that extensive binding to CR3 or other partner proteins in neutrophils may limit uPAR distribution into lipid rafts. It is also possible that uPAR-bearing multiprotein complexes in lipid rafts could artifactually segregate with high density plasma membranes if they are tethered to cytoskeletal proteins (22), but all our studies were performed in the presence of cytochalasin B to minimize such associations with the cytoskeleton. In preliminary experiments, the distribution of uPAR and its partner proteins among the membrane fractions was not materially affected by including cytochalasin B (not shown). Moreover, it was demonstrated that cytochalasin B has no effect at all on Ca2+ signaling either by uPAR aggregation or stimulation with HMW-uPA. It remains to be determined if stimuli other than uPAR aggregation can cause a more dramatic accumulation of uPAR into lipid rafts.

In summary, this study demonstrates that a single GPI-linked receptor, uPAR, engages at least two distinct activation signaling pathways that are functionally and physically segregated within separate domains of the neutrophil plasma membrane. Thus, a categorical approach to defining uPAR function as being dependent or independent of lipid rafts appears to be inadequate. In this instance, the choice of adhesion proteins with which uPAR forms its signaling complexes is likewise compartmentalized, suggesting that location within the plasma membrane may be a major factor in regulating the association of uPAR among multiple candidate partner proteins and presumably, downstream intracellular signaling intermediates.


   Acknowledgments
 
This work was supported by NIH grants HL53283 (R.G.S.), AI35877 (R.A.B.), and AI 51789 (H.R.P.), and the National Multiple Sclerosis Society (H.R.P.).

Received in original form March 13, 2003

Received in final form August 12, 2003


   References
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 References
 

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