JBC

HOME HELP FEEDBACK SUBSCRIPTIONS ARCHIVE SEARCH TABLE OF CONTENTS
QUICK SEARCH:   [advanced]


     

Originally published In Press as doi:10.1074/jbc.M313414200 on February 6, 2004

J. Biol. Chem., Vol. 279, Issue 17, 17625-17633, April 23, 2004
This Article
Right arrow Abstract Freely available
Right arrow Full Text (PDF)
Right arrow All Versions of this Article:
279/17/17625    most recent
M313414200v1
Right arrow Alert me when this article is cited
Right arrow Alert me if a correction is posted
Right arrow Citation Map
Services
Right arrow Email this article to a friend
Right arrow Similar articles in this journal
Right arrow Similar articles in PubMed
Right arrow Alert me to new issues of the journal
Right arrow Download to citation manager
Right arrow reprints & permissions
Citing Articles
Right arrow Citing Articles via HighWire
Right arrow Citing Articles via Google Scholar
Google Scholar
Right arrow Articles by Frasch, S. C.
Right arrow Articles by Bratton, D. L.
Right arrow Search for Related Content
PubMed
Right arrow PubMed Citation
Right arrow Articles by Frasch, S. C.
Right arrow Articles by Bratton, D. L.
Social Bookmarking
Add to CiteULike   Add to Complore   Add to Connotea   Add to Del.icio.us   Add to Digg   Add to Reddit   Add to Technorati  
What's this?

Phospholipid Flip-Flop and Phospholipid Scramblase 1 (PLSCR1) Co-localize to Uropod Rafts in Formylated Met-Leu-Phe-stimulated Neutrophils*

S. Courtney Frasch{ddagger}, Peter M. Henson{ddagger}§, Kaz Nagaosa{ddagger}, Michael B. Fessler¶||, Niels Borregaard**, and Donna L. Bratton{ddagger}{ddagger}{ddagger}§§

From the Departments of {ddagger}Pediatrics and Medicine, Division of Cell Biology, National Jewish Medical and Research Center, Denver, Colorado 80206, the Departments of §Medicine and Pathology and {ddagger}{ddagger}Pediatrics, University of Colorado Health Sciences Center and the ||Department of Medicine, University of Colorado School of Medicine, Denver, Colorado 80262, and the **Department of Hematology, Rigshospitalet-4042, 9 Blegdamsvej, DK-2100 Copenhagen, Denmark

Received for publication, December 8, 2003 , and in revised form, February 4, 2004.


   ABSTRACT
TOP
 ABSTRACT
INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 
Movement of phosphatidylserine (PS) to the plasma membrane outer leaflet is a nearly universal marker of apoptosis and occurs during activation of many cells. Neutrophils stimulated with the chemotactic peptide formylated Met-Leu-Phe (fMLP) demonstrated transient PS exposure. Stimulated outward movement of PS was accompanied by enhanced inward movement of several phosphorylcholine lipid probes and was associated with enhanced FM 1-43 staining indicative of phospholipid packing changes. Unlike apoptosis, inward movement of exogenously added fluorescent PS did not decline, and DNA was not cleaved during fMLP stimulation. Movement of phospholipids occurred within minutes following stimulation, was independent of endocytosis/pinocytosis, and was consistent with bidirectional, transbilayer phospholipid flip-flop. While the role of phospholipid scramblase 1 (PLSCR1) is controversial in flip-flop, we sought evidence for its role in enhanced phospholipid movements during fMLP stimulation. Using antibodies to the carboxyl-terminal domain of PLSCR1, its presence in the plasma membranes of non-permeabilized neutrophils was confirmed by flow cytometry. Additionally subcellular fractionation demonstrated that PLSCR1 was also located in secretory vesicles and tertiary and secondary granules. Activation of neutrophils with fMLP, however, did not significantly alter surface labeling suggesting that stimulated phospholipid flip-flop does not require additional mobilization of PLSCR1 to the plasma membrane. As expected for palmitoylated proteins, PLSCR1 was enriched in detergent-insoluble membranes and co-localized with raft markers at the neutrophil uropod after stimulation. Of note, PS exposure, phospholipid uptake, and FM 1-43 staining also localized to the uropod following stimulation demonstrating that both PLSCR1 and phospholipid flip-flop characterize this specialized domain of polarized neutrophils.


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 
Most resting cells appear to demonstrate marked asymmetry in the transbilayer distribution of phospholipids composing the plasma membrane with the majority of sphingomyelin and phosphatidylcholine (PC)1 localizing to the membrane outer leaflet and most phosphatidylethanolamine and nearly all phosphatidylserine (PS) localizing to the membrane inner leaflet (1, 2). This phospholipid asymmetry is thought to be maintained by both an aminophospholipid translocase activity that flips (inward) PS, and to a lesser extent phosphatidylethanolamine, from the outer leaflet back to the inner leaflet and a low spontaneous rate of phospholipid flip and flop (outward) across the plasma membrane bilayer (1, 3–5).

When cells die by programmed cell death, or apoptosis, PS exposure almost universally results. Recognition of exposed PS on the apoptotic cell by a stereospecific receptor on the phagocyte signals for both phagocytic engulfment and down-regulation of inflammatory mediator production (6). As such, apoptotic cells are cleared in an efficient and non-phlogistic manner (7). PS exposure has also been reported under certain conditions during activation of inflammatory cells and on capacitated spermatozoa (8). For example, PS is exposed on platelets stimulated by thrombin and collagen, on B cells with the B cell antigen receptor capped, on mast cells with cross-linked Fc{epsilon}RI, and most recently on neutrophils treated with dimeric galectin-1 after initial stimulation with fMLP (9–13). Externalization of PS on inflammatory cells is thought to play a role in the activation of both complement and coagulation cascades and supports the binding and activity of secreted phospholipases A2 (see "Discussion").

The mechanism(s) of enhanced exposure of PS during apoptosis has been shown variously to be either caspase-dependent or -independent and to require cytoskeletal alterations, calcium flux, or oxidation (14–16). Where it has been investigated, PS exposure is thought to result, at least in part, from loss of PS flip due to declining aminophospholipid translocase activity (14, 17). The mechanisms of PS exposure during cellular activation have received much less attention. Where it has been studied, PS exposure in activation, like apoptosis, is accompanied by enhanced staining with lipophilic dyes that indicate changes in membrane lipid packing and enhanced nonspecific uptake of phospholipids bearing various head groups including PC species (12, 14, 17–21).

Notably the mechanisms of phospholipid movement and the proteins involved in either outward movement of PS or inward movement of PC have not been unequivocally identified (see "Discussion"). Members of the P-type ATPase family have been proposed as candidate proteins mediating flip of PS from the outer leaflet to the inner leaflet and are credited with establishing and maintaining the basal asymmetric distribution of PS, but reverse function resulting in PS exposure has not been noted (1, 3–5). Other proteins of the ABC family have been proposed as mediating flop of phospholipids from inner leaflet to outer leaflet (e.g. ABCA1, ABCB1, and ABCB4) (for reviews, see Refs. 1 and 22) but have not been found to enhance inward flip of phospholipids from outer leaflet to inner leaflet. Also these proteins appear to be constitutively active with little evidence that they are activated within minutes of stimulation as has been described for PS exposure in inflammatory cells. Although controversial, the PLSCR1 is hypothesized to mediate calcium-activated, bidirectional, nonspecific (with regard to head group) phospholipid movement that has been shown to accompany PS exposure in apoptosis and cellular activation (23–26) (see "Discussion"). However, where phospholipid movement has been associated with PLSCR1 expression, the precise mechanisms of movement have not been elucidated. Movement of phospholipids could involve either transbilayer flip-flop across the plasma membrane (27) and/or, given PLSCR1 localization to recycling endosomes in epithelial cells (28), could involve bilayer mixing during vesicle-plasma membrane fission and fusion.

Here we have shown that stimulated neutrophils transiently expose PS during fMLP activation in the absence of evidence for apoptosis. PS exposure was accompanied by altered plasma membrane phospholipid packing and enhanced inward movement of both alkyl- and acyl-linked choline-containing phospholipids. As these phospholipid movements were not accompanied by evidence of either actin-dependent or actin-independent endocytosis/pinocytosis, the data are strongly supportive of phospholipid movement by flip-flop across the plasma membrane. Furthermore we present evidence that phospholipid flip-flop and membrane packing changes occurred in "raft" membranes at the uropod of polarized neutrophils, and that plasma membrane PLSCR1 co-localized to these same domains.


   EXPERIMENTAL PROCEDURES
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
RESULTS
 DISCUSSION
 REFERENCES
 
Reagents—Cholera toxin B (CT-B)-Alexa 555, FM 1-43, annexin V-Alexa 488, fluorescein isothiocyanate-phalloidin, 6-((N-(7-nitrobenz-2-oxa-1,3-diazol-4-yl)amino)hexanoyl)sphingosyl phosphocholine (NBD-sphingomyelin), and Lucifer Yellow were from Molecular Probes (Eugene, OR). 1-Palmitoyl-2-[12-[(7-nitro-2,1,3-benzoxadiazol-4-yl)amino]hexanoyl[-sn-glycero-3-phosphoserine] (NBD-PS) and 1-palmitoyl-2-[12-[(7-nitro-2–1,3-benzoxadiazol-4-yl)amino]hexanoyl[-sn-glycero-3-phosphocholine] (NBD-PC) were from Avanti Polar Lipids (Alabaster, AL). Radiolabeled 1-O-[hexadecyl-1',2'-3H(N)]2-N-methylcarbamyl-platelet activating factor (cPAF) was synthesized by PerkinElmer Life Sciences. Anti-CD55 was from Santa Cruz Biotechnology (Santa Cruz, CA), and anti-CD11b was from Dako Corp. (Carpenteria, CA). Anti-CD45 clone BRA-55, fatty acid-free BSA, and fMLP were from Sigma. AEBSF, aprotinin, and leupeptin were from Calbiochem. Factor Va and anti-Factor Va were from Hematologic Technologies, Inc. (Essex Junction, VT). Rabbit PLSCR1 antiserum was raised against the 14 carboxyl-terminal amino acids of PLSCR1 (sequence CESTGSQEQKSGVW).

Cell Isolation and Culture—Using endotoxin-free reagents and plasticware, human neutrophils were isolated by the plasma Percoll method as described previously (29). Unless otherwise noted, all incubations were done in NBD buffer (137 mM NaCl, 2.7 mM KCl, 2 mM MgCl2,5mM glucose, 10 mM HEPES (pH 7.4)) or Krebs-Ringer phosphate-dextrose (KRPD) buffer (0.9% saline, 4.8 mM KCl, 0.93 mM CaCl2, 1.2 mM MgSO4, 3.1 mM NaH2PO4, 12.5 mM Na2HPO4, 5% dextrose) with designated amounts of fatty acid-free BSA. T-Rex-293 cells (Invitrogen) were cultured in Dulbecco's modified Eagle's medium supplemented with 10% fetal calf serum, L-glutamine, 1% penicillin, 1% streptomycin, 1 mM sodium pyruvate at 37 °C in a 10% CO2 humidified incubator.

cPAF Uptake—Uptake of radiolabeled cPAF was carried out as described previously (24) with the following modifications. Uptake of [3H]cPAF took place over the last 5 min of stimulation with fMLP at 37 °C. Samples were diluted with an equal volume of ice-cold 10% BSA in KRPD buffer and washed two more times with 5% BSA in KRPD buffer at 4 °C to remove outer leaflet lipid. The cell pellets were resuspended in 500 µl of 1% Triton X-100. Uptake of labeled lipid was determined by scintillation counting.

NBD-PS, NBD-PC, NBD-sphingomyelin, Annexin V, and Factor Va Staining—NBD-PC (2.5 µg), NBD-PS (2.5 µg), or NBD-sphingomyelin (5.0 µg) was dried under nitrogen and resuspended in 20 µl of NBD buffer and 180 µl of propidium iodide (50 µg/ml stock). Neutrophils were stimulated as described at a concentration of 5 x 106/ml in NBD buffer with 1 mM CaCl2 and 0.25% BSA. Following stimulation, a final concentration of 1.25 µg/ml NBD-PC or NBD-PS or 2.5 µg/ml NBD-sphingomyelin was added to 100 µl of cells at 37 °C for 5 min. Outer leaflet lipid was removed by diluting cells with an equal volume of ice-cold 10% BSA in NBD buffer followed by two additional washes at 4 °C with 5% BSA in NBD buffer. Cells were resuspended in 500 µl of ice-cold NBD buffer and analyzed by flow cytometry.

For annexin V staining, 1 x 106 cells in 100 µl of NBD buffer with 2.5 mM CaCl2, annexin V-Alexa 488 (1:50), and 5 µg/ml propidium iodide were incubated for 15 min at room temperature, diluted with 400 µl of ice-cold NBD buffer with 2.5 mM CaCl2, and analyzed by flow cytometry. For activated Factor V (FVa) staining, 1 x 106 neutrophils in 100 µl NBD buffer with 1% BSA and 1 mM CaCl2 were transferred to 100 µlof FVa and propidium iodide (10 µg/ml and 5 µg/ml, respectively) or propidium iodide alone for the last 10 min of stimulation. Cells were then fixed for 10 min at room temperature with 3% paraformaldehyde, 3% sucrose in PBS, washed once, and incubated with 100 µl of anti-FVa antibody (final concentration of 10 µg/ml) for 10 min at room temperature. Cells were washed once and incubated with 100 µl of anti-mouse IgG F(ab')2 (1:100 in NBD buffer plus 1 mM CaCl2 and 0.1% BSA) for 30 min on ice, washed once, and analyzed by flow cytometry. For fluorescence microscopy, FVa staining was performed as for flow cytometry except in the absence of propidium iodide and incubated for 15 min at room temperature with Cy3 goat anti-mouse IgG F(ab')2 diluted 1:100. Cells were mounted on coverslips with OPDA (20 mg/ml o-phenylenediamine dihydrochloride in 1 M Tris base (pH 8.5) diluted 1:1 with glycerol) and viewed with a Lieca DMRXA fluorescence microscope using a 63x oil (numerical aperture 1.32) Plan-Apo objective. Images were analyzed using Slidebook software (Intelligent Imaging Innovations, Inc., Denver, CO).

FM 1-43 and Lucifer Yellow Staining—Neutrophils at 10 x 106 cells/ml in NBD buffer with 0.05% BSA and 1 mM CaCl2 were stimulated with fMLP for the times indicated. For the last minute of stimulation, 5 x 105 cells were transferred to 450 µl of NBD buffer and FM 1-43 (final concentration of 2.0 µM) and allowed to incubate for an additional 1 min at 37 °C. Cells were centrifuged and resuspended in 500 µl of ice-cold NBD buffer and analyzed by flow cytometry. Lucifer Yellow staining was carried out as described previously (30).

Raft Membrane Isolation—Neutrophils (300 x 106)at20 x 106/ml in KRPD buffer with 0.25% BSA were stimulated with fMLP for 15 min. Following stimulation, cells were centrifuged and resuspended in 1 ml of ice-cold lysis buffer (25 mM MES (pH 6.5), 150 mM NaCl, 1% Triton X-100, 1 mM AEBSF, 1 mM NaF, 5 µg/ml leupeptin, 5 µg/ml aprotinin) and sonicated for 1 min on ice. Unbroken cells, nuclei, and debris were removed by centrifugation at 1000 x g for 10 min at 4 °C. The supernatant was added to 2 ml of 80% sucrose in MES-buffered saline (25 mM MES (pH 6.5), 150 mM NaCl), and the lysates were centrifuged through a 10–40% continuous sucrose gradient for 18–24 h at 37,000 rpm at 4 °C. Following centrifugation, 1-ml fractions were harvested from the bottom by puncturing the tube with an 18-gauge needle. Protein concentrations in each fraction were determined by the Bradford protein assay (31), and 10 µg of protein were run on 10% SDS-PAGE, transferred to nitrocellulose, and probed for PLSCR1 or CD55.

Alkaline Phosphatase Assay—Sucrose density fractions were measured for alkaline phosphatase activity to identify rafts. Fraction samples (20 µl) were added to a 96-well flat bottom plate and mixed with 200 µl of reaction buffer (5 mM p-nitrophenyl phosphate in 100 mM 2-amino-2-methyl-1-propanol (pH 10.0)). Reactions were incubated at 37 °C for at least 15 min, and the absorbance was read in a microplate reader at 405 nm.

Cloning and Transfection of PLSCR1—RNA isolated from Jurkat cells was synthesized to cDNA using a reverse transcriptase for PCR kit (Clontech Inc.) according to the manufacturer's instructions. cDNA was used as a template to amplify full-length PLSCR1 using the forward primer 5'-AAGAATTCGGCAGCCAGAGAACTGTTTTAA-3' and the reverse primer 5'-AAGAATTCGCAGTTTTTCAAAGGAAGTTTCA-3', both of which have engineered an EcoRI restriction enzyme cloning site on the 5' ends. The PCR product of the correct size was purified and cloned directly into the EcoRI sites of pcDNA4/TO (Invitrogen), which allows the expression of the cloned gene in the presence of doxycycline. T-Rex-293 cells, expressing the tetracycline repressor, were transfected with PLSCR1 in the absence or presence of doxycycline to induce expression of PLSCR1 using FuGENE transfection reagent (Roche Applied Science) according to the manufacturer's instructions. Cells were harvested 24 h after transfection into lysis buffer (50 mM Tris-HCl (pH 7.4), 150 mM NaCl, 1 mM EGTA, 0.25% deoxycholate, 1% Triton X-100). Protein concentrations were determined by the Bradford protein assay (31), and 10 µg of protein were run on 10% SDS-PAGE, blotted to nitrocellulose, and probed for PLSCR1.

PLSCR1, CD45, and Cholera Toxin B Staining—For PLSCR1 staining, neutrophils (2 x 106)at20 x 106/ml were blocked for 15 min on ice with 1% BSA in KRPD buffer and then incubated with either PLSCR1 serum (1:50) or preimmune serum (1:50) from the same rabbit on ice for an additional 30 min. Primary antibody was removed, and neutrophils (in KRPD buffer with 0.25% BSA) were stimulated with fMLP for 10 min at 37 °C, fixed in 3% paraformaldehyde, 3% sucrose in PBS at room temperature for 10 min, washed twice with PBS, and incubated with Cy3 goat anti-Rabbit F(ab')2 diluted 1:100 for 30 min on ice. Cells were washed twice more with PBS and analyzed by flow cytometry. For CD45, neutrophils were preincubated as above except with 10 µg/ml mouse anti-human CD45 or isotype control antibody, and Cy3 anti-mouse F(ab')2 diluted 1:100 was used.

For fluorescence microscopy, cells were pretreated with antibody as above or with CT-B-Alexa 555 diluted 1:200 and either settled for 20 min at room temperature on poly-L-lysine-coated coverslips (pretreated with 100 µg/ml in PBS at 4 °C overnight) followed by fMLP stimulation at 37 °C or stimulated in suspension. Following stimulation, cells were fixed as above and incubated for 15 min at room temperature with secondary antibody, washed three times with PBS, and permeabilized with 0.2% Triton X-100 for 10 min at room temperature. Cells were washed three times with PBS, stained with fluorescein isothiocyanate-phalloidin (1:250) and 5 µg/ml Hoechst in PBS for 15 min at room temperature, washed with PBS, mounted with OPDA, and viewed with a Zeiss fluorescence microscope using a 63x oil (numerical aperture 1.4) Axiovert 200M objective. Images were analyzed using Slidebook software.

Live Cell Imaging—Neutrophils (7.5 x 106 cells at 5 x 106/ml) in NBD buffer with 0.01% BSA, 1 mM EDTA, 300 µM EGTA, and either FM 1-43 or NBD-PC at a final concentration of 16 µM or 5 µg/ml, respectively, were placed in a BSA (0.01%)-coated Delta T dish (Bioptechs, Inc.) that was preheated to 37 °C. Neutrophils were stimulated with fMLP, and images were taken over 5 min with an inverted Olympus IX70 microscope equipped with a Sensicam camera connected to TILL-vision software (T.I.L.L. Photonics GmbH) every 3 s with a 500-ms exposure time for FM 1-43 and every 5 s with a 3-s exposure time for NBD-PC.

Subcellular Fractionation—Neutrophil granules were isolated by Percoll density gradients from control neutrophils as described previously (32). Plasma membrane and secretory vesicles from control neutrophils were isolated by free flow electrophoresis as described previously (33).


   RESULTS
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
DISCUSSION
 REFERENCES
 
PS Exposure on fMLP-stimulated Neutrophils—Using FVa as a sensitive probe to detect outer leaflet PS (34), we were able to demonstrate that fMLP stimulation resulted in transient PS exposure that was evident as early as 10 min and returned to base-line levels by 45 min (Fig. 1A). In contrast, this transient PS exposure was minimally detectable by the less sensitive annexin V binding (data not shown). PS exposure during fMLP stimulation was not accompanied by loss of aminophospholipid translocase activity as was seen during UV-induced apoptosis (Fig. 1B); indeed PS uptake during fMLP stimulation appeared to be enhanced. Nor was activation of caspase 3 or DNA degradation (data not shown) seen following stimulation with fMLP. These data demonstrate that transient PS exposure during neutrophil activation was not associated with apoptosis.



View larger version (36K):
[in this window]
[in a new window]
  		FIG. 1.
Stimulated neutrophils transiently expose PS on their surface and take up phospholipid probes. Neutrophils were stimulated with 10 nM fMLP, and surface exposed PS was measured by FVa binding (A). Uptake of NBD-PS was used as a measure of aminophospholipid translocase activity in fMLP-stimulated or apoptotic neutrophils and compared with unstained control (CTR Unst), fMLP-stimulated (20' Unst), or apoptotic (APO Unst) neutrophils (B). C, time course of uptake of radiolabeled cPAF in neutrophils stimulated with 10 nM fMLP. D, uptake of NBD-PC in fMLP-stimulated or apoptotic (UV-irradiated) neutrophils compared with unstained control (CTR Unst), fMLP-stimulated (20' Unst), or apoptotic (APO Unst) neutrophils. ', minutes.

 
PS Exposure Was Accompanied by Enhanced Uptake of Other Glycerophospholipids—To test whether the outward flop of endogenous PS was both specific for PS and unidirectional or alternatively was bidirectional as suggested by enhanced PS uptake observed in Fig. 1B, we tested whether exogenous phosphocholine-containing diradyl lipid probes presented to the activated neutrophil would be internalized. Using a non-hydrolyzable, alkyl-linked phosphocholine probe, c-PAF, fMLP stimulation resulted in enhanced phospholipid uptake that was rapid in onset, occurred as early as 5 min, and peaked at 15 min following stimulation (Fig. 1C). Enhanced phospholipid uptake did not result from stimulation of the PAF receptor by c-PAF as it occurred in the presence of the PAF receptor inhibitor WEB 2086 (data not shown) and was identical for the fluorescently labeled NBD-PC, an ester-linked phosphatidylcholine probe that does not stimulate the PAF receptor (Fig. 1D). As a positive control, neutrophils stimulated to undergo apoptosis with UV irradiation also showed increased phospholipid uptake of choline-containing probes (14, 17, 24). While phospholipid uptake during apoptosis was inhibited by the caspase 3 inhibitor DEVD phospholipid uptake during fMLP stimulation was not inhibited, again dissociating it from the process of apoptosis (data not shown) (24).

Phospholipid Uptake Was Independent of Endocytosis and Pinocytosis—In contrast to the enhanced uptake of choline-containing glycerophospholipids, internalization of NBD-sphingomyelin, which has been used as a lipid marker of endocytosis (35), was not enhanced during either stimulation or apoptosis (data not shown); this was despite equivalent staining for both NBD-sphingomyelin and NBD-PC of both control and unwashed samples.

To further investigate whether PC uptake was attributable to an endocytic/pinocytic pathway and whether PS exposure was the consequence of bilayer mixing during vesicle-plasma membrane budding and fusion during endosome recycling, the following experiments were conducted. First, uptake of the fluid phase marker Lucifer Yellow was determined in duplicate samples following stimulation of neutrophils with fMLP alone or fMLP with cytochalasins. Pretreatment with either cytochalasin B or D has been shown to inhibit actin assembly and actin-mediated endocytosis during fMLP stimulation but also to significantly enhance both degranulation and pinocytosis (30). As shown in Fig. 2A, fMLP stimulation alone resulted in no significant uptake of the fluid phase dye as we have published previously (30) while markedly stimulating NBD-PC uptake (Fig. 2B). Conversely pretreatment with either cytochalasin B (Fig. 2A) or D (not shown) prior to fMLP stimulation markedly enhanced uptake of Lucifer Yellow while having little effect on NBD-PC uptake over that seen with fMLP alone (Fig. 2B). Additionally inhibition of clathrin-mediated endocytosis by hyperosmolar sucrose (0.4 mM) (36, 37) had no inhibitory effect on fMLP-stimulated uptake of NBD-PC (data not shown).



View larger version (23K):
[in this window]
[in a new window]
  		FIG. 2.
Increased NBD-PC uptake and FM 1-43 staining do not result from endocytosis/pinocytosis. A, uptake of the fluid phase dye Lucifer Yellow as a measure of endocytosis/pinocytosis in neutrophils stimulated with 10 nM fMLP alone or with cytochalasin B (CB) pretreatment. B, uptake of NBD-PC in neutrophils stimulated with 10 nM fMLP alone or with cytochalasin B (CB) pretreatment compared with unstained control (Unst CTR) or cells stimulated with fMLP for 20 min (Unst fMLP). C, FM 1-43 staining in neutrophils stimulated as above for the times indicated and stained with FM 1-43 during the last minute of stimulation. ', minutes.

 
Movement of Phospholipids Was Accompanied by Altered Membrane Packing Associated with Transbilayer Flip-Flop—It is known that calcium ionophore-treated erythrocytes demonstrate bidirectional, nonspecific, transbilayer flip-flop of phospholipid probes as well as enhanced staining with FM 1-43 or the related dye MC540, which are sensitive to phospholipid packing changes (38–40). Similarly cells undergoing apoptosis and activation have shown enhanced FM 1-43 (or MC540) dye staining (12, 14, 17–21, 41). Therefore, neutrophils were stimulated with fMLP for various lengths of time and stained with FM 1-43 for the last minute of stimulation. Enhanced staining was evident as early as 2 min following fMLP addition (Fig. 2C). Furthermore FM 1-43 staining was unaffected by the presence of cytochalasin B or D. Taken together, these data demonstrate that PS exposure is accompanied by both nonspecific phospholipid uptake and membrane packing changes that are independent of both pinocytosis and actin-dependent endocytosis. These data strongly support activated, bidirectional, transbilayer flip-flop of phospholipids in the stimulated neutrophil.

Localization of PLSCR1 to Neutrophil Raft Membranes— While the role of PLSCR1 in mediating phospholipid flip-flop remains controversial (see the Introduction and "Discussion"), we sought to determine whether human neutrophils expressed PLSCR1 on the plasma membrane surface and whether its localization could be related to areas of phospholipid flip-flop and membrane packing changes. First, using an antibody raised against the 14 carboxyl-terminal amino acids of PLSCR1, a single band at 35 kDa was identified by Western immunoblot analysis of neutrophil whole cell lysates (Fig. 3A). This band was identical to PLSCR1 expressed endogenously and overexpressed in T-Rex-293 cells suggesting that this antibody does indeed recognize PLSCR1 protein. Additionally subcellular fractionation of neutrophil lysates revealed that PLSCR1 was localized mainly in the plasma membrane/secretory vesicle fraction but also to tertiary granules and to a lesser degree in secondary granules (Fig. 3B). It was absent from both the primary granules and the cytosol. Further investigation utilizing free flow electrophoresis supported the finding that PLSCR1 was localized in both the plasma membrane and, to a lesser extent, the secretory vesicles (Fig. 3B).



View larger version (32K):
[in this window]
[in a new window]
  		FIG. 3.
PLSCR1 is evident on the neutrophil plasma membrane. A, Western immunoblot of PLSCR1 in neutrophil whole cell lysates (PMN) or in T-Rex-293 cells transfected with PLSCR1 in the absence or presence of doxycycline (Dox). Doxycycline induced the expression of transfected PLSCR1 over the endogenously expressed PLSCR1 in the T-Rex-293 cells. B, top, Western immunoblot of PLSCR1 in purified granule fractions (1°, 2°, and 3°), plasma membrane/secretory vesicle (PM/SV) fraction, or cytosol (Cyt) isolated by Percoll density centrifugation of control neutrophils. Bottom, Western immunoblot of PLSCR1 from plasma membrane (PM) or secretory vesicles (SV) purified and separated by free flow electrophoresis from control neutrophils. C, surface staining of PLSCR1 in non-permeabilized control or fMLP-stimulated (10 nM for 10 min) neutrophils compared with staining of neutrophils with preimmune (PI) serum with analysis by flow cytometry. D, increased mean fluorescence of surface-stained CD11b but not PLSCR1 in neutrophils stimulated with 10 nM fMLP for 5 or 15 min. CTR, control.

 
As the protein sequence analysis predicts PLSCR1 to be a Type II protein with a single transmembrane domain near the carboxyl terminus (27), we labeled non-permeabilized control and stimulated neutrophils at 4 °C with the carboxyl-terminal domain antibody or preimmune serum from the same rabbit followed by flow cytometric analysis. As shown in Fig. 3C, we found evidence of surface staining for PLSCR1 on control neutrophils, and stimulation of cells with fMLP for 10 min resulted in little enhancement of surface staining for PLSCR1. By contrast, CD11b, also localized to secretory vesicles and secondary and tertiary granules, was increased following stimulation (Fig. 3D). These observations suggest that activated phospholipid flip-flop does not require increased PLSCR1 on the plasma membrane. Furthermore they also suggest the possibility that PLSCR1 localized to both the plasma membrane and labile secretory vesicles may be recycled between these compartments (see "Discussion") reminiscent of the recycling endosomes of epithelial cells (28).

PLSCR1, a palmitoylated protein, has been localized to raft membrane domains in epithelial carcinoma KB cells (28). Isolation of detergent-resistant membranes from both control and fMLP-stimulated neutrophils followed by Western immunoblot analysis revealed that PLSCR1 was also enriched in neutrophil rafts (Fig. 4A) as determined by co-localization with CD55, a known raft protein (Fig. 4B), and alkaline phosphatase activity (Fig. 4C). Interestingly immunoblots for PLSCR1 from membranes prepared for raft isolation showed two bands, one at the expected 35 kDa and a smaller one at 31 kDa. The presence of this smaller band, despite inclusion of protease inhibitors, remains unexplained but appeared to be related to the raft isolation procedure in that it was not apparent when whole cell lysates were prepared for Western blotting.



View larger version (22K):
[in this window]
[in a new window]
  		FIG. 4.
PLSCR1 is localized to lipid rafts. A, Western immunoblot analysis of PLSCR1 in fractions obtained from sucrose density centrifugation in control or fMLP-stimulated neutrophils. B, Western immunoblot analysis of CD55, a known raft protein, in fractions from sucrose density centrifugation. C, analysis of alkaline phosphatase activity, a marker of lipid rafts, in fractions from sucrose density centrifugation. Scr, PLSCR1; Ctr, control; Alk Phos, alkaline phosphatase.

 
Analysis of PLSCR1 surface distribution on control non-permeabilized neutrophils by epifluorescence microscopy demonstrated that the protein was localized on the plasma membrane in a patchy distribution unlike CD45, a protein often excluded from rafts, which appeared evenly distributed on the membrane surface (Fig. 5A). Stimulation of neutrophils with fMLP to produce a polarized phenotype resulted in aggregation of these patches and redistribution of PLSCR1 to the uropod as shown in cells stained with anti-PLSCR1 antibody and phalloidin to identify the leading edge (Fig. 5B). CD45 staining, on the other hand, remained evenly distributed on the membrane despite the polarized phenotype.



View larger version (17K):
[in this window]
[in a new window]
  		FIG. 5.
PLSCR1 aggregates at the uropod with a marker of lipid rafts in fMLP-polarized neutrophils. A, surface labeling of PLSCR1 (red) or CD45 (red) on control neutrophils compared with either preimmune (PI) or isotype control. B, surface PLSCR1 (red) or CD45 (red) stained prior to permeabilization for staining with phalloidin (green) to identify F-actin and Hoescht (blue) to identify the nucleus in neutrophils stimulated with 10 nM fMLP for 10 min. C, control or fMLP-stimulated neutrophils stained for surface GM1 with CT-B (red), a marker of lipid rafts, F-actin (green), and the nucleus (blue) as above. Arrows indicate the uropod of polarized cells. CTR, control.

 
It has been shown that migrating T cells exhibit a polarized phenotype with asymmetrically distributed raft domains: GM1-containing rafts localized to the uropod, and GM3-containing rafts localized to the leading edge (42). Similarly in neutrophils, it has been shown that chemoattractant receptors are redistributed to the front of the cell during migration (43) and that detergent-resistant membranes and the raft markers CD44 and CD43 localize toward the rear in polarized cells (44). Whether fMLP-polarized neutrophils asymmetrically distribute GM1-containing membrane rafts to the uropod was determined. Neutrophils were stained with CT-B to identify GM1-containing rafts and phalloidin to identify actin assembly at the leading edge. As shown in Fig. 5C, in unstimulated control cells (lacking F-actin), CT-B appeared to be patched and focally distributed similar to the pattern of staining observed with PLSCR1. Following fMLP stimulation, the leading edge was easily identified by phalloidin staining of F-actin, while GM1 stained with CT-B appeared to be excluded from the leading edge and aggregated exclusively at the uropod similar to that seen with PLSCR1. Direct co-localization could not be performed because CT-B binding appeared to interfere with PLSCR1 staining by flow cytometry (data not shown). Taken together, these data demonstrated that PLSCR1, a raft protein, associated with raft membranes at the neutrophil uropod following fMLP stimulation.

Phospholipid Flip-Flop Occurs in Neutrophil Rafts—We have shown that neutrophils exhibit enhanced nonspecific phospholipid flip-flop in response to fMLP and localized PLSCR1, a candidate protein proposed to play a role in mediating phospholipid flip-flop, to raft domains at the uropod. Therefore, we hypothesized that phospholipid flip-flop would occur in uropod raft membranes containing PLSCR1. As expected and shown in Fig. 6, staining of fMLP-polarized neutrophils with Factor Va showed bright staining of uropodia following stimulation demonstrating the uropod as the region of PS exposure. Additionally we hypothesized that the uropod was also the site of phospholipid packing changes and phospholipid uptake. Neutrophils were stimulated with fMLP in the presence of the dye FM 1-43 and visualized at 0 and 4 min. As demonstrated in Fig. 7A, FM 1-43 staining was brightest at developing uropodia in polarizing neutrophils stimulated with fMLP. Since enhanced FM 1-43 staining detects areas of altered phospholipid packing that have been associated with enhanced phospholipid flip-flop (21, 38, 39), we also stimulated neutrophils with fMLP in the presence of NBD-PC. As shown in Fig. 7B, incorporation of NBD-PC was also restricted to the uropod of the polarizing neutrophil. Taken together, the data strongly suggest that altered membrane packing, phospholipid flip, PS exposure (flop), and PLSCR1 all localize to raft membranes at the neutrophil uropod.



View larger version (88K):
[in this window]
[in a new window]
  		FIG. 6.
Surface PS exposure occurs at the uropod in fMLP-polarized neutrophils. Neutrophils were left unstimulated or stimulated with 10 nM fMLP for 10 min and stained with FVa (red) to identify surface-exposed PS and with Hoescht for DNA (blue). Nomarsky images were taken to identify the leading edge and uropod.

 



View larger version (90K):
[in this window]
[in a new window]
  		FIG. 7.
FM 1-43 and NBD-PC staining occurs at the uropod in fMLP-polarized neutrophils. A, neutrophils were stimulated with 10 nM fMLP in the presence of FM 1-43. Live cell images were taken every 3 s with a 500-ms exposure time. Images shown are at 0 and 4 min of stimulation. Arrows indicate the uropod of the polarized cells. B, neutrophils were stimulated with 10 nM fMLP in the presence of NBD-PC. Live cell images were taken every 5 s with a 3-s exposure time. Images shown are at 0 and 4 min of stimulation. Arrows indicate the uropod of polarized cells.

 

   DISCUSSION
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
REFERENCES
 
The movement of PS from inner to outer plasma membrane leaflet (flop) is demonstrated for nearly all cells undergoing apoptosis and in some cells following activation including the fMLP-stimulated neutrophil (Fig. 1). While PS exposure on apoptotic cells leads to efficient engulfment by phagocytes with anti-inflammatory consequences (6, 7), PS exposure during viable cell activation may or may not have similar consequences, likely dependent on quantity, localization, or duration of PS exposure required for recognition (10, 45) or the presence of "don't eat me signals" (6, 7). Additionally activation of both the complement and coagulation cascades are well documented consequences of PS exposure and potentially result during inflammatory cell activation (2). Finally localized domains of exposed PS on inflammatory cells also significantly enhanced the binding of secreted phospholipases A2 (46). Indeed human embryonic kidney 293 cells overexpressing PLSCR1 have been reported to have both enhanced PS exposure and sensitivity to type IIa and type V secretory phospholipases A2 (26).

Growing evidence supports that outward phospholipid flop is not specific for PS but in fact is utilized to externalize other signaling phospholipids as well. Lyso-PC and PAF have been shown to be externalized and released in both activation and apoptosis (18, 39, 47–49). In both cases, these lipid mediators are first synthesized within the cell, then moved across the plasma membrane bilayer, and released to acceptor proteins (e.g. albumin) before reaching target cells where they mediate effects through specific receptors.

Flop of phospholipids during neutrophil stimulation is accompanied by both phospholipid packing changes (Fig. 2) and inward flip of phospholipids (Fig. 1). As such, these findings are similar to observations made in apoptotic cells and in calcium ionophore-treated erythrocyte ghosts (14, 17, 39). The purpose of inward flip of phospholipids, other than the aminophospholipids in non-apoptotic cells, is not readily apparent, but compensatory movement of phospholipid between the bilayers may be required to maintain bilayer geometry (50). In contrast to activated flip-flop demonstrated here, investigation in unstimulated CHO-K1 cells suggested that NBD-PC can be internalized by endocytosis (51), and recent work in apoptosing HeLa cells suggested that while an alkyl-linked PAF-like lipid was also endocytosed, acyl-linked NBD-PC was internalized by flip-flop (52). Here, in fMLP-stimulated neutrophils, we have detected no differences in the activated uptake of sn-1 alkyl- and acyl-linked phosphorylcholine lipid species, and in both cases, phospholipid flip-flop rather than either endocytosis or pinocytosis appeared to be the route of entry (Figs. 1 and 2).

Exposed PS is described as punctate or patchy in distribution on activated cells (10, 11, 13) and has also been shown to be transiently localized to cell-cell contact areas during myotubule formation (53) and to the apical region of capacitated spermatozoa (8). Data suggest that these localized domains of outer leaflet PS represent raft membranes: PS co-localized with caveolin in macrophages (54), co-capped with IgM in raft domains following B cell stimulation (11), and co-localized with apical raft domains on capacitated spermatozoa (8, 41). In this study, we found in the fMLP-stimulated neutrophil that PS exposure and phospholipid packing changes also occurred in domains that co-localized with raft proteins as well as with PLSCR1 (Figs. 4, 5, and 6). Furthermore we found this colocalization at neutrophil uropodia, which along with the lamellipodia are known raft membrane locations in polarized neutrophils showing front-to-back segregation: while urokinase-type plasminogen activator receptor moves to lamellipodia (55), CR3, CD55, P-selectin glycoprotein ligand 1, intercellular adhesion molecule-3, CD44, and CD43 move to uropodia (44, 55, 56), and CD45 and HLA remain diffusely distributed on plasma membranes following stimulation (44). Other stimulated and polarized cells (myelocytic cells, T cells, and myeloma cells) have also shown similar raft protein segregation between lamellipodia and uropodia suggesting that these redistributions may be required for polarization and chemotaxis (42, 57–62).

Directional migration of neutrophils in response to fMLP requires phosphatidylinositol 3-kinases and protein kinases C, which recognize inner leaflet phospholipids (phosphatidyl-inositides and PS, respectively) (63). The recruitment of phosphatidylinositol 3-kinases, generation of phosphatidylinositol 3,4-bisphosphate and phosphatidylinositol 3,4,5-trisphosphate, and subsequent recruitment of pleckstrin homology domain-containing proteins (e.g. Akt/protein kinase B) and Rho GT-Pases to the leading edge are thought to transmit a shallow extracellular chemoattractant gradient into a steep intracellular gradient for the restricted assembly of F-actin at the lamellipodium (63–65). Thus spatial restriction of phosphatidylinositol 3-kinase-modified phosphatidylinositides, their catabolism (66, 67), and potentially their transbilayer movement would serve to inhibit actin assembly at locations other than the leading edge and enable directed migration. Similarly the local flop of PS to the outer leaflet may serve to "turn off" protein kinases C and cytoskeletal assembly at the uropod during chemotaxis (68). Studies in PLSCR1-overexpressing mast cells showed that enhanced PS exposure was associated with inhibition of exocytotic degranulation (69). Thus, we would also hypothesize that, in chemotaxing neutrophils, granular exocytosis needed to replenish chemotactic receptors and adhesion molecules at the leading edge would be inhibited at the uropod (60, 61).

The role of PLSCR1 in phospholipid flip-flop is controversial. Zhou et al. (27), who originally cloned the protein from erythroblastic leukemia cells, were able to show enhanced transbi-layer movement of NBD-PC in PLSCR1-reconstituted proteoliposomes. In earlier studies, we demonstrated enhanced cPAF uptake in CHO-K1 cells transfected with both PLSCR1 and protein kinase C{delta}, which was shown to phosphorylate PLSCR1 (24). Finally overexpression of PLSCR1 (with or without calcium and ionophore stimulation) enhanced annexin binding in human embryonic kidney 293 cells (26), CHO-K1 cells (23), RBL-2H3 cells (69), and Raji cells (25), although other investigations have not shown enhanced activity with overexpression of the protein (28, 70, 71). On the other hand, recent studies using platelets from PLSCR1-/- mice showed no difference in PS exposure, although direct measures of phospholipid movement using phospholipid probes were not reported, and the presence of other family member proteins raise the possibility of compensatory activity occurring in cells from knock-out mice (72). As a 35–37-kDa protein with one purported transmembrane domain, PLSCR1 would seem an unlikely candidate to directly mediate transbilayer movement of phospholipids. However, as a palmitoylated raft membrane protein, it may either self-aggregate or associate with other partners or signal via phosphotyrosine-mediated associations within the raft platforms to enhance phospholipid movement across the bilayer (28, 73).

Recent studies in the PLSCR1-/- mouse demonstrating defective myelopoiesis suggest a role for PLSCR in neutrophil differentiation as well (72). Whether this or other functions of the protein, such as binding of secretory leukocyte protease inhibitor (74), enhancement of PS biosynthesis during apoptosis (23), or signaling to downstream c-Src (73), are of importance in phospholipid flip-flop remain to be determined. While our studies here in terminally differentiated human neutrophils place PLSCR1 in the same location as phospholipid flip-flop and we hypothesize that PLSCR1 is important in polarization and chemotaxis, its role in the actual process of phospholipid flip-flop requires future studies in functionally chemotaxing and genetically mutable cells for further elucidation.


   FOOTNOTES
 
* This work was funded in part by National Institutes of Health Grants HL34303 and AI058228 [GenBank] and by the Danish Medical Research Council. The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact. Back

§§ To whom correspondence should be sent: Dept. of Pediatrics, National Jewish Medical and Research Center, Rm. D506, 1400 Jackson St., Denver, CO 80206. Tel.: 303-398-1390; Fax: 303-398-1381; E-mail: brattond{at}njc.org.

1 The abbreviations used are: PC, phosphatidylcholine; PS, phosphatidylserine; AEBSF, 4-(2-aminoethyl)benzenesulfonylfluoride, HCl; BSA, bovine serum albumin; PAF, platelet-activating factor; cPAF, 1-O-[hexadecyl-1'-,2'-3H(N)]2-N-methylcarbamyl-platelet activating factor; CT-B, cholera toxin B; fMLP, formylated Met-Leu-Phe; FVa, activated Factor V; GM1, Gal{beta}1,3GalNAc{beta}1,4(Neu5Ac{alpha}2,3)Gal{beta}1,4Glc{beta}1,1-ceramide, NH4; GM3, Neu5Ac{alpha}2,3Gal{beta}1,4Glc{beta}1,1-ceramide, NH4; KRPD, Krebs-Ringer phosphate-dextrose, MES, 2-(N-morpholino)ethanesulfonic acid; NBD, (7-nitro-2,1,3-benzoxadiazol-4-yl)amino; PBS, phosphate-buffered saline; PLSCR1, phospholipid scramblase 1; ABC, ATP-binding cassette; CHO, Chinese hamster ovary. Back


   ACKNOWLEDGMENTS
 
We thank Jenai Kailey, Cally Duncan, and Charlotte Horn for technical support and Brenda Sebern for preparation of the manuscript.



   REFERENCES
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 

  1. Daleke, D. L. (2003) J. Lipid Res. 44, 233-242[Abstract/Free Full Text]
  2. Zwaal, R. F. A., and Schroit, A. J. (1997) Blood 89, 1121-1132[Free Full Text]
  3. Tang, X., Halleck, M. S., Schlegel, R. A., and Williamson, P. (1996) Science 272, 1495-1497[Abstract]
  4. Schlegel, R. A., and Williamson, P. (2001) Cell Death Differ. 8, 551-563[CrossRef][Medline] [Order article via Infotrieve]
  5. Halleck, M. S., Lawler, J. J., Blackshaw, S., Gao, L., Nagarajan, P., Hacker, C., Pyle, S., Newman, J. T., Nakanishi, Y., Ando, H., Weinstock, D., Williamson, P., and Schlegel, R. A. (1999) Physiol. Genomics 1, 139-150[Abstract/Free Full Text]
  6. Henson, P. M., Bratton, D. L., and Fadok, V. A. (2001) Curr. Biol. 11, R795-R805[CrossRef][Medline] [Order article via Infotrieve]
  7. Henson, P. M., Bratton, D. L., and Fadok, V. A. (2001) Nat. Rev. Mol. Cell. Biol. 2, 627-633[CrossRef][Medline] [Order article via Infotrieve]
  8. De Vries, K. J., Wiedmer, T., Sims, P. J., and Gadella, B. M. (2003) Biol. Reprod. 68, 2122-2134[Abstract/Free Full Text]
  9. Bevers, E. M., Comfurius, P., and Zwaal, R. F. A. (1983) Biochim. Biophys. Acta 736, 57-66[Medline] [Order article via Infotrieve]
  10. Dias-Baruffi, M., Zhu, H., Cho, M., Karmakar, S., McEver, R. P., and Cummings, R. D. (2003) J. Biol. Chem. 278, 41282-41293[Abstract/Free Full Text]
  11. Dillon, S. R., Mancini, M., Rosen, A., and Schlissel, M. S. (2000) J. Immunol. 164, 1322-1332[Abstract/Free Full Text]
  12. Williamson, P., Bevers, E. M., Smeets, E. F., Comfurius, P., Schlegel, R. A., and Zwaal, R. F. (1995) Biochemistry 34, 10448-10455[CrossRef][Medline] [Order article via Infotrieve]
  13. Windmiller, D. A., and Backer, J. M. (2003) J. Biol. Chem. 278, 11874-11878[Abstract/Free Full Text]
  14. Bratton, D. L., Fadok, V. A., Richter, D. A., Kailey, J. M., Guthrie, L. A., and Henson, P. M. (1997) J. Biol. Chem. 272, 26159-26165[Abstract/Free Full Text]
  15. Kagan, V. E., Fabisiak, J. P., Shvedova, A. A., Tyurina, Y. Y., Tyurin, V. A., Schor, N. F., and Kawai, K. (2000) FEBS Lett. 477, 1-7[CrossRef][Medline] [Order article via Infotrieve]
  16. Kunzelmann-Marche, C., Freyssinet, J. M., and Martinez, M. C. (2001) J. Biol. Chem. 276, 5134-5139[Abstract/Free Full Text]
  17. Verhoven, B., Schlegel, R. A., and Williamson, P. (1995) J. Exp. Med. 182, 1597-1601[Abstract/Free Full Text]
  18. Bratton, D. L., Dreyer, E., Kailey, J. M., Fadok, V. A., Clay, K. L., and Henson, P. M. (1992) J. Immunol. 148, 514-523[Abstract]
  19. McEvoy, L., Schlegel, R. A., Williamson, P., and Del Buono, B. J. (1988) J. Leukoc. Biol. 44, 337-344[Abstract]
  20. Williamson, P., Christie, A., Kohlin, T., Schlegel, R. A., Comfurius, P., Harmsma, M., Zwaal, R. F., and Bevers, E. M. (2001) Biochemistry 40, 8065-8072[CrossRef][Medline] [Order article via Infotrieve]
  21. Zweifach, A. (2000) Biochem. J. 349, 255-260[CrossRef][Medline] [Order article via Infotrieve]
  22. Pomorski, T., Hrafnsdottir, S., Devaux, P. F., and van Meer, G. (2001) Semin. Cell Dev. Biol. 12, 139-148[CrossRef][Medline] [Order article via Infotrieve]
  23. Yu, A., McMaster, C. R., Byers, D. M., Ridgway, N. D., and Cook, H. W. (2003) J. Biol. Chem. 278, 9706-9714[Abstract/Free Full Text]
  24. Frasch, S. C., Henson, P. M., Kailey, J. M., Richter, D. A., Janes, M. S., Fadok, V. A., and Bratton, D. L. (2000) J. Biol. Chem. 275, 23065-23073[Abstract/Free Full Text]
  25. Zhao, J., Zhou, Q., Wiedmer, T., and Sims, P. J. (1998) J. Biol. Chem. 273, 6603-6606[Abstract/Free Full Text]
  26. Murakami, M., Kambe, T., Shimbara, S., Higashino, K., Hanasaki, K., Arita, H., Horiguchi, M., Arita, M., Arai, H., Inoue, K., and Kudo, I. (1999) J. Biol. Chem. 274, 31435-31444[Abstract/Free Full Text]
  27. Zhou, Q., Zhao, J., Stout, J. G., Luhm, R. A., Wiedmer, T., and Sims, P. J. (1997) J. Biol. Chem. 272, 18240-18244[Abstract/Free Full Text]
  28. Sun, J., Nanjundan, M., Pike, L. J., Wiedmer, T., and Sims, P. J. (2002) Biochemistry 41, 6338-6345[CrossRef][Medline] [Order article via Infotrieve]
  29. Haslett, C., Guthrie, L. A., Kopaniak, M. M., Johnston, R. B., Jr., and Henson, P. M. (1985) Am. J. Pathol. 119, 101-110[Abstract]
  30. Fittschen, C., and Henson, P. M. (1994) J. Clin. Investig. 93, 247-255[Medline] [Order article via Infotrieve]
  31. Bradford, M. M. (1976) Anal. Biochem. 72, 248-254[CrossRef][Medline] [Order article via Infotrieve]
  32. Kjeldsen, L., Sengelov, H., and Borregaard, N. (1999) J. Immunol. Methods 232, 131-143[CrossRef][Medline] [Order article via Infotrieve]
  33. Sengelov, H., and Borregaard, N. (1999) J. Immunol. Methods 232, 145-152[CrossRef][Medline] [Order article via Infotrieve]
  34. Zhai, X., Srivastava, A., Drummond, D. C., Daleke, D., and Lentz, B. R. (2002) Biochemistry 41, 5675-5684[CrossRef][Medline] [Order article via Infotrieve]
  35. Hao, M., and Maxfield, F. R. (2000) J. Biol. Chem. 275, 15279-15286[Abstract/Free Full Text]
  36. Bruhl, H., Cohen, C. D., Linder, S., Kretzler, M., Schlondorff, D., and Mack, M. (2003) Eur. J. Immunol. 33, 3028-3037[CrossRef][Medline] [Order article via Infotrieve]
  37. Heuser, J. E., and Anderson, R. G. (1989) J. Cell Biol. 108, 389-400[Abstract/Free Full Text]
  38. Williamson, P. K., Algarin, L., Bateman, J., Choe, H.-R., and Schlegel, R. A. (1985) J. Cell. Physiol. 123, 209-214[CrossRef][Medline] [Order article via Infotrieve]
  39. Bratton, D. L., Kailey, J. M., Clay, K. L., and Henson, P. M. (1991) Biochim. Biophys. Acta 1062, 24-34[Medline] [Order article via Infotrieve]
  40. Williamson, P., Mattocks, K., and Schlegel, R. A. (1983) Biochim. Biophys. Acta 732, 387-393[Medline] [Order article via Infotrieve]
  41. Flesch, F. M., Brouwers, J. F., Nievelstein, P. F., Verkleij, A. J., van Golde, L. M., Colenbrander, B., and Gadella, B. M. (2001) J. Cell Sci. 114, 3543-3555[Abstract/Free Full Text]
  42. Gomez-Mouton, C., Abad, J. L., Mira, E., Lacalle, R. A., Gallardo, E., Jimenez-Baranda, S., Illa, I., Bernad, A., Manes, S., and Martinez, A. C. (2001) Proc. Natl. Acad. Sci. U. S. A. 98, 9642-9647[Abstract/Free Full Text]
  43. Sanchez-Madrid, F., and del Pozo, M. A. (1999) EMBO J. 18, 501-511[CrossRef][Medline] [Order article via Infotrieve]
  44. Seveau, S., Eddy, R. J., Maxfield, F. R., and Pierini, L. M. (2001) Mol. Biol. Cell 12, 3550-3562[Abstract/Free Full Text]
  45. Borisenko, G. G., Matsura, T., Liu, S. X., Tyurin, V. A., Jianfei, J., Serinkan, F. B., and Kagan, V. E. (2003) Arch. Biochem. Biophys. 413, 41-52[CrossRef][Medline] [Order article via Infotrieve]
  46. Diaz, C., Leon, G., Rucavado, A., Rojas, N., Schroit, A. J., and Gutierrez, J. M. (2001) Arch. Biochem. Biophys. 391, 56-64[CrossRef][Medline] [Order article via Infotrieve]
  47. Bratton, D. L. (1993) J. Biol. Chem. 268, 3364-3373[Abstract/Free Full Text]
  48. Lauber, K., Bohn, E., Krober, S. M., Xiao, Y. J., Blumenthal, S. G., Lindemann, R. K., Marini, P., Wiedig, C., Zobywalski, A., Baksh, S., Xu, Y., Autenrieth, I. B., Schulze-Osthoff, K., Belka, C., Stuhler, G., and Wesselborg, S. (2003) Cell 113, 717-730[CrossRef][Medline] [Order article via Infotrieve]
  49. Kim, S. J., Gershov, D., Ma, X., Brot, N., and Elkon, K. B. (2002) J. Exp. Med. 196, 655-665[Abstract/Free Full Text]
  50. Mohandas, N., Greenquist, A. C., and Shohet, S. B. (1978) J. Supramol. Struct. 9, 453-458[CrossRef][Medline] [Order article via Infotrieve]
  51. Sleight, R. G., and Pagano, R. E. (1984) J. Cell Biol. 99, 742-751[Abstract/Free Full Text]
  52. Van Der Luit, A. H., Budde, M., Verheij, M., and Van Blitterswijk, W. J. (2003) Biochem. J. 374, 747-753[CrossRef][Medline] [Order article via Infotrieve]
  53. van den Eijnde, S. M., van den Hoff, M. J., Reutelingsperger, C. P., van Heerde, W. L., Henfling, M. E., Vermeij-Keers, C., Schutte, B., Borgers, M., and Ramaekers, F. C. (2001) J. Cell Sci. 114, 3631-3642[Medline] [Order article via Infotrieve]
  54. Gargalovic, P., and Dory, L. (2003) J. Lipid Res. 44, 1622-1632[Abstract/Free Full Text]
  55. Kindzelskii, A. L., Laska, Z. O., Todd, R. F., III, and Petty, H. R. (1996) J. Immunol. 156, 297-309[Abstract]
  56. Alonso-Lebrero, J. L., Serrador, J. M., Dominguez-Jimenez, C., Barreiro, O., Luque, A., del Pozo, M. A., Snapp, K., Kansas, G., Schwartz-Albiez, R., Furthmayr, H., Lozano, F., and Sanchez-Madrid, F. (2000) Blood 95, 2413-2419[Abstract/Free Full Text]
  57. del Pozo, M. A., Sanchez-Mateos, P., Nieto, M., and Sanchez-Madrid, F. (1995) J. Cell Biol. 131, 495-508[Abstract/Free Full Text]
  58. Millan, J., Montoya, M. C., Sancho, D., Sanchez-Madrid, F., and Alonso, M. A. (2002) Blood 99, 978-984[Abstract/Free Full Text]
  59. Borset, M., Hjertner, O., Yaccoby, S., Epstein, J., and Sanderson, R. D. (2000) Blood 96, 2528-2536[Abstract/Free Full Text]
  60. Pierini, L. M., Lawson, M. A., Eddy, R. J., Hendey, B., and Maxfield, F. R. (2000) Blood 95, 2471-2480[Abstract/Free Full Text]
  61. Lawson, M. A., and Maxfield, F. R. (1995) Nature 377, 75-79[CrossRef][Medline] [Order article via Infotrieve]
  62. Pierini, L. M., and Maxfield, F. R. (2001) Proc. Natl. Acad. Sci. U. S. A. 98, 9471-9473[Free Full Text]
  63. Rickert, P., Weiner, O. D., Wang, F., Bourne, H. R., and Servant, G. (2000) Trends Cell Biol. 10, 466-473[CrossRef][Medline] [Order article via Infotrieve]
  64. Manes, S., Ana Lacalle, R., Gomez-Mouton, C., and Martinez, A. C. (2003) Trends Immunol. 24, 320-326[Medline] [Order article via Infotrieve]
  65. Hannigan, M., Zhan, L., Li, Z., Ai, Y., Wu, D., and Huang, C. K. (2002) Proc. Natl. Acad. Sci. U. S. A. 99, 3603-3608[Abstract/Free Full Text]
  66. Iijima, M., and Devreotes, P. (2002) Cell 109, 599-610[CrossRef][Medline] [Order article via Infotrieve]
  67. Funamoto, S., Meili, R., Lee, S., Parry, L., and Firtel, R. A. (2002) Cell 109, 611-623[CrossRef][Medline] [Order article via Infotrieve]
  68. Manno, S., Takakuwa, Y., and Mohandas, N. (2002) Proc. Natl. Acad. Sci. U. S. A. 99, 1943-1948[Abstract/Free Full Text]
  69. Kato, N., Nakanishi, M., and Hirashima, N. (2002) Biochemistry 41, 8068-8074[CrossRef]