HOME HELP FEEDBACK SUBSCRIPTIONS ARCHIVE SEARCH TABLE OF CONTENTS		QUICK SEARCH:   [advanced] Author: Keyword(s): Year:  Vol:  Page:

J. Biol. Chem., Vol. 280, Issue 44, 37021-37032, November 4, 2005

This Article Abstract Full Text (PDF) Supplemental Data All Versions of this Article: 280/44/37021    most recent M506594200v1 Submit a Letter to Editor Alert me when this article is cited Alert me when eLetters are posted Alert me if a correction is posted Citation Map Services Email this article to a friend Similar articles in this journal Similar articles in PubMed Alert me to new issues of the journal Download to citation manager Request Permissions Citing Articles Citing Articles via HighWire Citing Articles via Google Scholar Google Scholar Articles by Guichard, C. Articles by Elbim, C. Search for Related Content PubMed PubMed Citation Articles by Guichard, C. Articles by Elbim, C. Social Bookmarking                      What's this?

Interleukin-8-induced Priming of Neutrophil Oxidative Burst Requires Sequential Recruitment of NADPH Oxidase Components into Lipid Rafts^*

Cécile Guichard1, Eric Pedruzzi, Cédric Dewas, Michèle Fay, Cécile Pouzet, Marcelle Bens¶, Alain Vandewalle¶, Eric Ogier-Denis, Marie-Anne Gougerot-Pocidalo, and Carole Elbim

From the Unité INSERM 683, ^¶Unité INSERM 478, IFR02, BP 416, Facultédemédecine Xavier BICHAT, BP 416, 75870 Paris Cedex 18, France

Received for publication, June 17, 2005 , and in revised form, August 12, 2005.

	ABSTRACT

TOP ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

 
The superoxide-producing phagocyte NADPH oxidase consists of a membrane-bound flavocytochrome b₅₅₈, the cytosol factors p47^phox, p67^phox, p40^phox, and the small GTPase Rac2, which translocate to the membrane to assemble the active complex following neutrophil activation. Interleukin-8 (IL-8) does not activate NADPH oxidase, but potentiates the oxidative burst induced by stimuli such as formyl-methionyl-leucyl-phenylalanine (fMLP) via a priming mechanism. The effect of IL-8 on the components of NADPH oxidase during the priming process has never been investigated in human neutrophils. Here we showed that within 3 min, IL-8 treatment enhanced the Btk- and ERK1/2-dependent phosphorylation of p47^phox, as well as the recruitment of flavocytochrome b₅₅₈, p47^phox, and Rac2 into cholesterol-enriched detergent-resistant microdomains (or lipid rafts). Conversely, IL-8 treatment lasting 15 min failed to recruit flavocytochrome b₅₅₈, p47^phox, or Rac2, but did enhance the Btk- and p38 MAPK-dependent phosphorylation and the translocation of p67^phox into detergent-resistant microdomains. Moreover, methyl--cyclodextrin, which disrupts lipid rafts, inhibited IL-8-induced priming in response to fMLP. Our findings indicate that IL-8-induced priming of the oxidative burst in response to fMLP involves a sequential assembly of the NADPH oxidase components in the lipid rafts of neutrophils.

	INTRODUCTION

TOP ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

 
Neutrophils are key components of the early innate response against microbial pathogens. Their activation triggers microbiocidal mechanisms, including the rapid production of reactive oxygen species (ROS)² known as the oxidative burst, that play a key role in bacterial killing. However, in an excessive and/or inappropriate response, ROS can induce vascular and tissue lesions. Generation of ROS by neutrophils results from the activation of NADPH oxidase, a multicomponent enzyme system that catalyzes the NADPH-dependent reduction of oxygen to the superoxide anion (), which is the precursor of the other ROS. In resting cells, this multicomponent enzyme system is inactive, and its components are dispersed between the cytosol and the membranes. The flavocytochrome b₅₅₈ component, which is composed of two subunits, gp91^phox and p22^phox, is located in the plasma membrane and in specific granules. The other components of the NADPH complex (p47^phox, p67^phox, p40^phox, and small G protein Rac2) are cytosol proteins. The activation of neutrophils by various stimuli, such as bacterial N-formyl peptides (fMLP) and phorbol myristate acetate, triggers the phosphorylation of the p47^phox, p67^phox, and p40^phox cytosolic components and their translocation to the plasma membrane, where they interact with flavocytochrome b₅₅₈ (1-4). Concomitantly, Rac2 is dissociated from its inhibitor, RhoGDP dissociation inhibitors, and then interacts with flavocytochrome b₅₅₈ to form a binding partner for p67^phox (5). The. complete assembly of NADPH oxidase components is crucial for production (6, 7).

The activation of NADPH oxidase is tightly regulated at infectious and inflammatory sites to avoid tissue and vascular lesions. Proinflammatory cytokines, such as interleukin-8 (IL-8), granulocyte-macrophage colony-stimulating factor, and tumor necrosis factor , are known to modulate NADPH oxidase activity by means of a priming phenomenon that strengthens the bactericidal capacity of the neutrophils (8-12). These cytokines do not stimulate oxidative burst activity on their own, but strongly enhance ROS production in response to exposure to a secondary applied stimulus, such as fMLP. Partial p47^phox phosphorylation has been shown to be induced by tumor necrosis factor (13), granulocyte-macrophage colony-stimulating factor (14), and bacterial lipopolysaccharide (15) during the priming of the oxidative burst. Other priming mechanisms have also been described previously, including increased membrane expression of flavocytochrome b₅₅₈ (15, 16), activation of heterotrimeric G proteins (17), and some other mechanisms (18, 19). Several mechanisms have been reported to be involved in the priming effect of IL-8. They include increased expression and cycling of fMLP receptors (fMLP-R) (20), increased intracellular-free Ca²⁺ (21), and increased phospholipase A₂ activation (22). However, the effects of IL-8 on the transductional pathways that determine the kinetics and assembly of the various components of the NADPH oxidase complex, and which could contribute to the IL-8-induced priming of the oxidative burst, have yet to be investigated.

It has recently been shown that cholesterol-enriched detergent-resistant membrane microdomains (DRMs), or lipid rafts, play a key role in the fMLP-, phorbol myristate acetate-, and Fc-dependent receptor activation of NADPH oxidase by triggering the recruitment and/or spatial orientation of cytosolic phox proteins relative to flavocytochrome b₅₅₈ (23, 24). The involvement of lipid rafts in regulating neutrophil functions is suggested by the fact that the neutrophil oxidative burst can be severely curtailed by depleting the level of cholesterol in the plasma membranes using methyl--cyclodextrin (MCD) (23, 25). It remains to be determined whether lipid raft microdomains play a regulatory role in the IL-8-induced priming of the oxidative burst in human neutrophils.

In this study, we analyzed the effects of IL-8, alone and when combined with fMLP, on the phosphorylation and/or translocation of p47^phox, p67^phox, and Rac2, the segregation of flavocytochrome b₅₅₈ into DRMs, and the transduction pathways involved in these processes. The main findings of this study reveal that IL-8 induced in a sequential manner the phosphorylation of cytosol phox components and the recruitment of the NADPH oxidase components (including flavocytochrome b₅₅₈) into lipid rafts, suggesting that this pre-assembly could play at least some part in the priming effect of IL-8 on the neutrophil oxidative burst triggered in response to N-formyl peptides.

	MATERIALS AND METHODS

TOP ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

 
Reagents—Human recombinant interleukin-8 (rhIL-8), rabbit polyclonal anti-p22^phox (FL-195), anti-ERK1 (C-16), and anti-ERK2 (K-23) were from Tebu-Bio (Le Perray en Yvelines, France). Phorbol myristate acetate, fMLP, mouse monoclonal anti-phosphotyrosine, and anti--actin, cytochrome c, cytochalasin B, protease and phosphatase inhibitor mixture, and fluorescein-conjugated cholera toxin B subunit were purchased from Sigma. Endotoxin-free buffers and salt solutions were from Invitrogen (Cergy, France). Genistein, bisindolylmaleimide I (GFX109203X), SB203580, PD98059, LFM-A13, LFM-A11, and wortmannin were from Calbiochem (La Jolla, CA). Rabbit polyclonal anti-phospho-ERK1/2, anti-phospho-p38 mitogen-activated protein kinase (MAPK), anti-p38 MAPK, anti-phospho-Akt/PKB. and anti-Akt/PKB antibodies were from Ozyme (St.-Quentin en Yvelines, France). The rabbit polyclonal anti-gp91^phox antibody was from Euromedex (Mundolsheim, France). [³²P]Orthophosphoric acid was from PerkinElmer Life Sciences. Isotype control, mouse monoclonal anti-flotillin-2/ESA, anti-Rac1, anti-Btk, and anti-fMLP receptor were purchased from BD Biosciences. The rabbit polyclonal anti-p47^phox (a gift of B. M. Babior, The Scripps Research Institute, La Jolla, CA) was raised against the 10 COOH-terminal residues of p47^phox and purified, as previously described (26, 27). Rabbit polyclonal antibodies against the COOH-terminal sequence of p67^phox were prepared as previously described (28). Alexa Fluor 546 goat anti-rabbit IgG (H+L) and reagents for SDS-PAGE were from Invitrogen. Horseradish peroxidase-conjugated secondary antibody and Hyperfilm^TM were from Amersham Biosciences. ECL reagents were from Perbio (Brebières, France).

Neutrophil Preparation—Human neutrophils were obtained in lipopolysaccharide-free conditions by means of dextran sedimentation and Ficoll centrifugation as previously described (13).

Superoxide Anion Production Assay—Superoxide anion () production was continuously recorded for 5 min by monitoring the superoxide dismutase-inhibitable reduction of ferricytochrome c, using an UVIKON 860 spectrophotometer equipped with a thermostatted (37 °C) cuvette holder, as previously described (29). Superoxide production by 10⁶ cells suspended in 1 ml of HBSS was triggered by adding the stimulus (fMLP M). In addition, the priming effect of IL-8 on production in response to fMLP was tested by pre-treating neutrophils (10⁶ cells) at 37 °C with IL-8 at various concentrations (0.01-50 ng/ml) and for various incubation times (1-60 min). production in response to fMLP ( M) was then measured for 5 min. In some experiments, samples were preincubated with MCD (0.2 mg/ml, 30 min) before IL-8 and fMLP stimulation.

³²P Labeling, Stimulation, and Fractionation of Neutrophils—Neutrophils were incubated in phosphate-free buffer (20 mM HEPES, pH 7.4, 140 mM NaCl, 5.7 mM KCl, 0.8 mM MgCl₂, and 0.025% bovine serum albumin) (14) containing 0.5 mCi of [³²P]orthophosphoric acid, 10⁸ cells/ml for 60 min at 30 °C. The neutrophils were then incubated at 37 °C for 5 min before being exposed to IL-8 for 1-20 min. In some experiments the neutrophils were preincubated for 15 min with various kinase inhibitors before being treated with IL-8. In all cases, the reaction was stopped by adding ice-cold buffer and centrifugation at 400 x g for 7 min at 4 °C. The cells were lysed in lysis buffer (150 mM NaCl, 1 mg/ml NaF, 1 mg/ml Na₃VO₄, 0.5 mg/ml -glycerophosphate, 0.5 mg/ml P-nitrophenyl phosphate, 0.2 mg/ml levamisol, 8% sucrose, 0.02 M Tris, pH 7.4, 0.5% Triton X-100, 2.5 mM EGTA, 5 mM EDTA, 10 µg/ml leupeptin, 10 µg/ml pepstatin, 10 µg/ml aprotinin, 2 mM phenylmethylsulfonyl fluoride, 1 mg/ml DNase I, 2.5 mM diisopropyl fluorophosphate). The resuspension was sonicated on ice (3 times for 10 s). The lysate was centrifuged at 100,000 x g for 30 min at 4 °C in a TL100 ultracentrifuge (Beckman, Fullerton, CA), and cleared supernatants were used for immunoprecipitation studies.

Immunoprecipitation of p47^phox and p67^phox—The cleared supernatant was incubated overnight with anti-p47^phox (1/200) or anti-p67^phox (1/200) antibodies; immune complexes were then immunoprecipitated using -bind G-Sepharose beads (Amersham Biosciences), and washed four times as described (14).

Neutrophil Membrane Preparation—Neutrophils were preincubated with 5 µM cytochalasin B for 5 min before being incubated with IL-8 (50 ng/ml, 3-20 min), with fMLP (10^-7 M for 3 min), or with cytochalasin B, used as control. Membranes were then prepared as previously described (14). Briefly, 1 x 10⁸ neutrophils/ml were sonicated for three 10-s periods on ice in 1 ml of relaxation buffer (10 mM Pipes, pH 7.3, 3 mM MgCl₂, 100 mM KCl, 5 mM NaCl) supplemented with 0.5 mM phenylmethylsulfonyl fluoride, 1 mM EGTA, 10 µg/ml eupeptic, and 10 µg/ml pepstatin. Cytosol and membrane fractions were separated by centrifuging at 200,000 x g on 15-50% (w/w) sucrose gradient for 45 min at 4 °C.

Preparation of Lipid Rafts—Lipid rafts were isolated as previously described (23). Neutrophils were incubated at 4 °C in buffer A (20 mM HEPES, pH 7.0, 150 mM NaCl, 5 mM EDTA, 1% Triton X-100) for 30 min, homogenized by 50 up and down strokes in a Dounce homogenizer, after which 80% sucrose stock in buffer A was added to the cell lysate to give a final concentration of 40% sucrose. The mixture was then layered under a sucrose step gradient (30 and 5% in buffer A) and centrifuged at 150,000 x g for 18 h at 4 °C. Detergent-insoluble, cholesterolrich membrane fractions, representing membrane rafts, floated to the interphase between 5 and 30% sucrose layers, whereas solubilized proteins or cytoskeleton-associated, detergent-insoluble proteins remained in the pellet of the gradient. After spinning, 10 fractions were collected from the top of the tubes, and the pellet was solubilized by sonication.

Electrophoresis and Blotting—The samples were subjected to SDS-PAGE in 10% polyacrylamide gels or NuPAGE® Novex 4-12% BisTris gels, using standard techniques and Invitrogen techniques, respectively. The separated proteins were transferred to nitrocellulose. After blocking in 1% milk, the membrane was probed with the anti-p47^phox (1/5000 dilution), -p67^phox (1/1000 dilution), -Rac2 (1/1000 dilution), -gp91^phox (1/1000 dilution), -p22^phox (1/100 dilution), -P-ERK1/2 (1/1000 dilution), or -P-p38 MAPK (1/1000 dilution) antibodies, and then with species-specific horseradish peroxidase-labeled goat anti-rabbit or mouse antibodies, and developed with an ECL kit, using light-sensitive film (Amersham Biosciences).

Densitometric analysis was performed using the Scion Image analysis program from National Institutes of Health. Results from DRM experiments were expressed as the percentage of NADPH oxidase components in the DRMs (fractions 2-5).

View larger version (24K): [in this window] [in a new window]   FIGURE 1. Priming effect of IL-8 on fMLP-stimulated neutrophil oxidative burst. A, effect of IL-8 on the kinetics of fMLP-stimulated production. Neutrophils were incubated. with IL-8 (1-50 ng/ml) and/or 10-7 M fMLP, as indicated. production was measured for 5 min using the cytochrome c reduction assay, as described under "Materials and Methods." B, dose response of IL-8 priming. Neutrophils were preincubated with various concentrations of IL-8 (0.01-50 ng/ml) for 20 min, and then stimulated with 10-7 M fMLP. production was measured after fMLP stimulation during 5 min; the values are mean ± S.E. (n = 3). *, p < 0.05 versus fMLP alone (white bar). Results were expressed as nanomole of cells/5 min. C, time course of priming by IL-8. Neutrophils were preincubated with 50 ng/ml IL-8 for 1-20 min, and then treated with 10-7 M fMLP. production was measured after fMLP stimulation lasting 5 min; the values reported are mean ± S.E. (n = 3). *, p < 0.05 versus time 0 value (white bar). Results were expressed as nanomole of cells/5 min.

Confocal Laser Scanning Microscopy Analysis—After stimulation, the neutrophils (2 x 10⁶/ml) were pooled and stained with fluorescein isothiocyanate-conjugated cholera toxin B subunit (10 µg/ml) and/or anti-rabbit polyclonal p22^phox antibody raised against extracellular epitopes of the p22^phox protein (1/10 dilution) for 30 min at 4 °C. After rinsing with phosphate-buffered saline, the samples were incubated with an Alexa 546-conjugated affinity purified goat anti-rabbit IgG (1/250) for 30 min at 4 °C. After extensive rinsing, neutrophils were then fixed in 1% paraformaldehyde and cytospun. Glass coverslides were mounted and examined by confocal laser scanning microscopy (CLSM-510-META, Zeiss, Germany) equipped with epifluorescent optics (x63 NA 1.3 oil-immersion objective). Simultaneous two-channel recording was performed with a pinhole size of 90 µm using excitation wavelengths of 488/588 nm, a 510/580 double dichroic mirror, and a 515-545 bandpass fluorescein isothiocyanate filter together with a 590-nm long pass filter. Double-labeled cells were analyzed separately to avoid leakage from one channel to another. As controls, the absence of staining was confirmed when the primary antibody was omitted.

Statistical Analysis—All results are expressed as mean ± S.E. Significant differences were identified using Student's t test. p < 0.05 was taken to be significant.

	RESULTS

TOP ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

 
A Brief Incubation with IL-8 Was Sufficient to Prime the Neutrophil Oxidative Burst in Response to Formyl Peptides—The enzyme kinetics of production in response to fMLP and/or IL-8 monitored by ferricytochrome c reduction assay are presented in Fig. 1A. Incubation of isolated neutrophils with 10^-7 M of fMLP induced the rapid and transient production of , whereas incubation with 50 ng/ml of IL-8 did not. The latter result was obtained regardless of the concentration of IL-8 (0.01-50 ng/ml) or the incubation time (2-45 min) used (not shown). In contrast, pre-exposure of the neutrophils to IL-8 (50 ng/ml) for 3 or 15 min enhanced both the initial. and maximum rates of production subsequently induced by 10^-7 M fMLP, whereas exposure to 1 ng/ml IL-8 did not. As previously reported (21), these findings suggest that IL-8 has a dose- and time-dependent priming effect on the production induced by fMLP. Pre-exposure of neutrophils for 20 min with increasing concentrations of IL-8 (0.01-50 ng/ml) enhanced the total production of induced by fMLP in a dose-dependent manner, up to a maximum response at 50 ng/ml IL-8 (Fig. 1B). The results from the kinetics study showed that the priming effect of IL-8 (50 ng/ml) on production first appeared after 2 min, and then progressively increased to plateau after incubating for 10 min (Fig. 1C). . production was inhibited by superoxide dismutase and diphenyleneiodonium, an inhibitor of NADPH oxidase (not shown). Incubating neutrophils with neutralizing anti-IL-8 monoclonal antibody before adding the IL-8 and fMLP abolished ROS production (not shown). This finding rules out the possibility that endotoxin contamination was responsible for the priming effect of IL-8.

View larger version (17K): [in this window] [in a new window]   FIGURE 2. Effect of IL-8 on p47phox and p67phox phosphorylations in human neutrophils. 32P-Labeled neutrophils were incubated without or with 50 ng/ml IL-8 for 1-15 min. p47phox (A) and p67phox (B) were then immunoprecipitated, submitted to SDS-PAGE transfer, and detected by autoradiography (32P-p47phox and 32P-p67phox). Western blot analysis using anti-p47phox and anti-p67phox antibodies showed that the same amounts of p47phox and p67phox were immunoprecipitated from each sample. Phosphorylated p47phox or p67phox was quantified by PhosphorImager analysis. Results are expressed as the percentage of phosphorylation levels from resting neutrophils. Values are mean ± S.E. (n = 5). *, p < 0.05 versus time 0 value (white bars).

Transductional Pathways Involved in IL-8-mediated Phosphorylation of p47^phox and p67^phox—We then tested the effects of various kinase inhibitors after ³²P labeling of neutrophils and immunoprecipitation of p47^phox and p67^phox to investigate the signaling pathways leading to their phosphorylation under IL-8 priming conditions. Genistein, a broad-specific tyrosine kinase inhibitor, and wortmannin, a PI3K inhibitor, reduced the IL-8-induced phosphorylation of both p47^phox and p67^phox (Fig. 4, A and B). Among the tyrosine kinases, the Tec kinases, and especially Btk, has been shown to contribute to fMLP-induced neutrophil responses by modulating the MAPK and PI3K/Akt/PKB pathways (31). The role of Btk during the IL-8 priming process was previously unknown. Here we have shown that IL-8 treatment induced a rapid but transient (3-5 min) membrane translocation of Btk (Fig. 4C). In addition, leflunomide metabolite analog LFM-A13, a Btk inhibitor, significantly reduced the IL-8-induced phosphorylation of p47^phox and p67^phox (Fig. 4, A and B), whereas the inactive analog (LFM-A11) had no such inhibitory effect. We next analyzed the effect of IL-8 on PKC, Akt/PKB, ERK1/2, and p38 MAPK, which are involved in the p47^phox and/or p67^phox phosphorylation in response to various stimuli (32, 33). Preincubation with GF109203X, a PKC inhibitor, significantly reduced the level of early (3 min) IL-8-induced p47^phox phosphorylation (Fig. 4A), but had no effect on the level of later (15 min) IL-8-induced p67^phox phosphorylation (Fig. 4B). Furthermore, ERK1/2 and Akt/PKB phosphorylation transiently peaked after exposure to IL-8 for 3 min (Fig. 4C). The data showed that PD98059, a MEK1/2 inhibitor, reduced the level of the early (3 min) IL-8-induced phosphorylation of p47^phox. Conversely, no inhibitory effect of PD98059 was detected on the later (15 min) IL-8-induced phosphorylation of p67^phox (Fig. 4B). After 5 min, IL-8 triggered the activation of p38 MAPK, which slowly declined during the following 20-min incubation period (Fig. 4C). Consistently with these results, SB203580, a p38 MAPK inhibitor, significantly reduced the level of late (15 min) IL-8-induced p67^phox phosphorylation (Fig. 4B). Taken together, these findings strongly suggest that ERK1/2 is involved in the early IL-8-induced p47^phox phosphorylation, and that p38 MAPK is involved in the late IL-8-induced p67^phox phosphorylation. Furthermore, LFM-A13, a Btk inhibitor, significantly reduced the IL-8-induced early activation of ERK1/2 and Akt/PKB, and the IL-8-induced late activation of p38 MAPK (Fig. 4D). Thus, Btk seems to be involved in the upstream transduction pathways leading to the phosphorylation of p47^phox and p67^phox.

IL-8 Induced the Sequential Translocation of p47^phox, p67^phox, and Rac2 to the Plasma Membrane in a Time-dependent Manner—The phosphorylation of the cytosol components p47^phox and p67^phox induced by stimuli such as phorbol myristate acetate or fMLP has been implicated in their translocation to the plasma membrane, and their subsequent assembly at flavocytochrome b₅₅₈ (28, 34). The membrane translocation-dependent activation of Rac2 has also been shown to control NADPH oxidase activity by means of its direct interaction with p67^phox and flavocytochrome b₅₅₈ (5, 35-37). We therefore investigated the time dependence of the effect of IL-8 treatment of neutrophils on the membrane translocation of p47^phox, p67^phox, and Rac2. IL-8 induced the rapid translocation of p47^phox and Rac2 into the neutrophil plasma membranes within 3 min (2.1 ± 0.2- and 2.6 ± 0.2-fold increases, respectively, as compared with time 0 value) (mean ± S.E., n = 3), whereas p67^phox was only detected in the membranes after incubating the neutrophils with IL-8 for 5 min. The level of p47^phox and Rac2 translocation declined as the incubation times with IL-8 were increased from 3 to 15 min, but they remained detectable in plasma membranes for incubation times of up to 15 min. p67^phox remained at the same level after incubation times with IL-8 of 5-15 min (2.4 ± 0.2-fold increase as compared with time 0 value) (mean ± S.E., n = 3) (Fig. 5). Inhibitors of signaling pathways involved in p47^phox and p67^phox phosphorylation (see Fig. 4, A and B) abolished the IL-8-induced translocation of these two components (not shown).

View larger version (19K): [in this window] [in a new window]   FIGURE 3. IL-8 potentiates the phosphorylations of p47phox and p67phox induced by fMLP in human neutrophils. 32P-Labeled neutrophils were incubated without or with 50 ng/ml IL-8 for 3 or 15 min and then with or without 10-7 M fMLP for 3 min. 32P-p47phox (A) and 32P-p67phox (B) were analyzed as described in the legend to Fig. 2. Results are expressed as the percentage of phosphorylation levels from resting neutrophils. Values are mean ± S.E. (n = 3) *, p < 0.05 versus fMLP alone (white bars).

IL-8 Induced the Sequential Translocation of Cytosol p47^phox, p67^phox, and Rac2 into Lipid Rafts in a Time-dependent Manner—DRM microdomains, or lipid rafts, are thought to play important roles in signal transduction, and are believed to act as scaffolds for the assembly of cytosol factors required for holoenzyme assembly. Recently, Shao et al. (24) have reported that the cytosol NADPH oxidase components are recruited into lipid rafts in response to Fc receptor activation and fMLP stimulation. These findings led us to analyze the effect of IL-8 on the segregation of cytosol NADPH oxidase components to form lipid rafts. Membrane preparations of untreated and IL-8-treated neutrophils were fractionated by flotation centrifugation and then subjected to Western blot analyses. The low-density membrane fractions (fractions 2-5 from the top of the layer) extracted at 4 °C contained the DRM marker flotillin-2 (Fig. 7A). As expected (38, 39), flotillin-2 was no longer detected in membranes that had been treated with two cholesterol-affecting drugs: 0.2 mg/ml MCD for 30 min at 37 °C or 0.4% saponin for 60 min at 4 °C (Fig. 7A). Very small amounts of the cytosol components p47^phox, p67^phox, and Rac2 were detected in DRMs from resting neutrophils (Fig. 7B) (1.4 ± 0.1, 2.6 ± 0.1, and 0.4% ± 0.1 of the total amount of p47^phox, p67^phox, and Rac2, respectively) (mean ± S.E., n = 3). A short incubation (3 min) with IL-8 increased the amounts of p47^phox and Rac2 detected in DRMs, but had no effect on the translocation of p67^phox (Fig. 7B). Conversely, a longer incubation time (15 min) with IL-8 reduced the relative amount of p47^phox and Rac2, and increased the level of p67^phox in DRMs (Fig. 7B). Densitometric analysis of the Western blots indicated that the percentage of p47^phox and Rac2 in low density fractions 2-5 were 12.1 ± 0.2 and 20.2% ± 0.2 of the total amount of p47^phox and Rac2, respectively (mean ± S.E., n = 3). The percentage of p67^phox was 19.9% ± 0.2 (mean ± S.E., n = 3). In addition, preincubating neutrophils with IL-8 potentiated the translocation of p47^phox, p67^phox, and Rac2 into DRMs induced by fMLP (Fig. 7C) (by a factor of 1.5 ± 0.1, 2.0 ± 0.1, and 2.2 ± 0.1 for 3 min incubation and 3.2 ± 0.2, 6 ± 0.2, and 4.2 ± 0.1 for 15 min incubation, respectively) (mean ± S.E., n = 3) as compared with fMLP alone. This effect was observed whatever the incubation time (3 or 15 min). These results confirmed the data from translocation experiments (Fig. 5).

View larger version (37K): [in this window] [in a new window]   FIGURE 4. Signaling pathways involved in p47phox and p67phox phosphorylation induced by IL-8 in human neutrophils. A and B, 32P-labeled neutrophils were incubated in the presence of phosphate-buffered saline (-), genistein (Gen, 100 µM), wortmannin (Wt, 250 nM), PD98059 (PD, 50 µM), SB203580 (SB, 25 µM), GFX109203X (GFX, 5 µM), LFM-A13 (A13, 25 µM), or LFM-A11 (A11, 25 µM) for 20 min, and then stimulated with IL-8 (50 ng/ml) for 3 (A) or 15 (B) min. p47phox (A) and p67phox (B) phosphorylations were analyzed as described in the legend to Fig. 2. C, effect of IL-8 on Btk activation at the neutrophil surface and on Akt and MAPK activation in human neutrophils. Western blotting was performed using anti-phospho-ERK1/2 (P-ERK1/2), -phospho-Akt/PKB (P-Akt), -phospho-p38 MAPK (P38 MAPK), and -Btk antibodies. Antibodies raised against non-phosphorylated forms of each protein were used to show that the same amounts of protein had been loaded, and the Btk translocation was analyzed using subcellular fractionation. D, effect of LFM-A13 (A13, 25 µM) or LFM-A11 (A11, 25 µM) (20 min) on IL-8 mediated activation of MAPK and Akt. Western blotting was performed as described in C. Values are mean ± S.E. (n = 3). *, p < 0.05 versus IL-8 alone (white bars).

To further define the polarized distributions of GM1 and p22^phox after incubating with IL-8 for 15 min, we used confocal light scanning microscopy to analyze (Fig. 8C) serial slices (1 µm thick) taken from the top to the bottom of the same un-permeabilized neutrophils. After incubating with IL-8 for 15 min, raft marker GM1 (green) was clearly localized at the top of the neutrophils, while p22^phox (red) localized at the bottom with little overlap of these two markers, as shown in Fig. 8A. This polarized distribution was not detected in resting neutrophils, or in neutrophils that had been exposed to IL-8 for 3 min.

The IL-8-induced segregation of gp91^phox and p22^phox in DRMs was greatly amplified following fMLP stimulation as compared with the effect of fMLP alone (Fig. 9A). Adding fMLP also increased the co-localization of both GM1 and p22^phox in the plasma membranes of unpermeabilized neutrophils following initial priming with IL-8 for 3 or 15 min (Fig. 9B).

View larger version (26K): [in this window] [in a new window]   FIGURE 5. Effect of IL-8 on p47phox, p67phox, and Rac2 translocation to the neutrophil membrane. Neutrophils were incubated with 50 ng/ml IL-8 for 1-15 min. Neutrophil fractions were prepared as described under "Materials and Methods." Western blotting of the membrane and cytosol fractions was performed with antibodies directed against p47phox and p67phox. Equal amounts of protein were loaded, as confirmed by Coomassie Blue staining. Results of densitometric analysis are expressed as arbitrary units. Values are mean ± S.E. (n = 3). *, p < 0.05 versus time 0 value (white bars).

View larger version (30K): [in this window] [in a new window]   FIGURE 6. Effect of IL-8 on fMLP-induced p47phox, p67phox, and Rac2 translocation to the neutrophil membrane. Neutrophils were incubated with or without 50 ng/ml IL-8 for 3 or 15 min, and then with or without 10-7 M fMLP for 3 min. Western blotting was performed as described in the legend of Fig. 5. Results of densitometric analysis are expressed as arbitrary units. Data are mean ± S.E. (n = 3). *, p < 0.05 versus fMLP alone (white bars).

View larger version (38K): [in this window] [in a new window]   FIGURE 7. Recruitment of NADPH oxidase cytosol components into lipid rafts. Neutrophils were lysed in Triton X-100, in some cases with prior MCD- or saponin-facilitated cholesterol extraction (MCD, 0.2 mg/ml, 30 min at 37 °C, and Sap, 0.4%, 1 h at 4°C) (A), and the lysate was centrifuged to equilibrium in a 40-5% sucrose step gradient. Recovered fractions (number 10 is the bottom, high-density fraction of the gradient) and the solubilized pellet (P) were analyzed by Western blotting, using antibodies against the cytosol components of oxidase and flotillin-2 (Flo-2), which is a raft marker, as indicated. Lipid raft fractions were obtained from: B, resting neutrophils or IL-8-stimulated neutrophils (IL-8 50 ng/ml for 3 or 15 min at 37 °C); or C, neutrophils stimulated with 10-7 M fMLP for 3 min with or without prior exposure to 50 ng/ml IL-8 (3 or 15 min at 37 °C) as described under "Materials and Methods." One representative experiment of three is shown.

	DISCUSSION

TOP ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

The lack of significant NADPH oxidase activity in neutrophils treated by IL-8 alone may suggest that the cytosol components of the oxidase are not totally mobilized in response to IL-8 treatment. Previous studies have shown that the assembly of an active NADPH oxidase at the plasma membrane depends on the phosphorylation of several sites of p47^phox (34, 40), and of more restricted sites of p67^phox (41, 42). Here we have shown that IL-8 exerted a dual effect on the phosphorylation of p47^phox and p67^phox by enhancing rapid phosphorylation of p47^phox and later phosphorylation of p67^phox. These sequential phosphorylation kinetics of p47^phox and p67^phox correlated with the induction by IL-8 of early and transient activation of ERK1/2, followed by the later activation of p38 MAPK in neutrophils (43, 44). Furthermore, PD98059 and SB203580 (inhibitors of the ERK1/2 and p38 MAPK pathways respectively) blocked the p47^phox phosphorylation induced by a short incubation time (3 min) with IL-8, and the p67^phox phosphorylation induced by a longer (15 min) incubation with IL-8, respectively, as well as their subsequent translocation.

Thus, IL-8 induced the phosphorylation of p47^phox and p67^phox with different kinetic patterns that diverge from the accepted model required for the full activation of NADPH oxidase. Moreover, the mechanisms of IL-8-induced priming differ from those previously reported for tumor necrosis factor and granulocyte-macrophage colony-stimulating factor, which have been shown to involve the partial phosphorylation of p47^phox and the phosphorylation of a subpopulation of p67^phox without any translocation of these components to the plasma membrane; in this model this partial phosphorylation may facilitate the fMLP-induced subsequent phosphorylation of the cytosol components, and therefore their translocation to the membrane (32). In contrast, fMLP and IL-8 (both of which are chemotactic agents binding to the G protein-coupled receptors) seem to share similar signaling pathways leading to similar kinetics of p47^phox phosphorylation. fMLP-induced production is associated with both the upstream/downstream kinase-dependent phosphorylation, and with the translocation of p47^phox and p67^phox to the plasma membrane (6). p47^phox phosphorylation in response to fMLP stimulation has been shown to be directly triggered by certain PKC isoforms, MAPK and Akt/PKB (32, 45-48). Among the MAPKs, ERK1/2 have been shown to mediate the fMLP-stimulated phosphorylation of p47^phox, whereas inhibition of p38 MAPK had little impact on p47^phox phosphorylation (32, 33). The combined phosphorylation of p47^phox by ERK1/2, Akt/PKB, and PKC leads to the translocation of p47^phox to the plasma membrane, and to the induction of the adapter function, responsible for the assembly of NADPH oxidase. In addition, direct p38 MAPK-dependent phosphorylation and the subsequent translocation of p67^phox is observed in response to fMLP stimulation of neutrophils (49). However, the rapid activation of p38 MAPK in response to fMLP stimulation appeared to be delayed in time when the neutrophils had been exposed to IL-8. Despite the fact that p67^phox can be phosphorylated by both ERK1/2 and p38 MAPK at several selective sites in response to fMLP stimulation (41), we could not detect any early ERK-dependent phosphorylation of p67^phox in response to IL-8 stimulation. Tyrosine kinases and PI3K have also been shown to control downstream signaling pathways, leading to a successful phosphorylation of p47^phox and p67^phox by fMLP. On the basis of the effects of the inhibitors tested (genistein and wortmannin), we showed that the IL-8-induced phosphorylation of both p47^phox and p67^phox depended on the upstream activation of tyrosine kinase and PI3K. Among the tyrosine kinases, the Tec kinases, and especially Btk has recently been shown to be involved in fMLP signaling, thus providing a direct link between G protein-coupled receptors and activation of the tyrosine kinases (31). Activation of the Tec kinases is mediated in part by the high affinity of their pleckstrin homology domains for phosphatidylinositol 3,4,5-P₃ (50), which is generated following the activation of PI3K. We have shown that IL-8 treatment induces membrane translocation of Btk. Furthermore, after adding LFM-A13, an inhibitor of Btk, we observed the inhibition of the IL-8-induced activation of the MAPK and Akt/PKB pathways reminiscent of previous results obtained after fMLP stimulation (31). LFM-A13 also inhibited both p47^phox and p67^phox phosphorylation induced by IL-8 treatment. Thus, Btk appears to be critically involved in initiating the tyrosine phosphorylation-dependent signaling cascades leading to the phosphorylation of both p47^phox and p67^phox following IL-8 treatment of neutrophils, in a similar manner to fMLP.

View larger version (72K): [in this window] [in a new window]   FIGURE 8. Segregation of gp91phox and p22phox into lipid rafts after IL-8 treatment. Neutrophils were untreated or treated with 50 ng/ml IL-8 for different times (3 or 15 min at 37 °C). Lipid raft fractions were analyzed by Western blotting using anti-gp91phox and anti-p22phox antibodies, as described under "Materials and Methods" (A). Confocal microscopy was performed with un-permeabilized neutrophils, treated with antibody raised against extracellular epitopes of the p22phox protein; ganglioside GM1 was used as a raft marker (panel B). 1-µm slices were obtained from the neutrophil (top to bottom) seen in contrast phase (CP) and analyzed by confocal microscopy (C). One representative experiment of three is shown.

View larger version (32K): [in this window] [in a new window]   FIGURE 9. Segregation of gp91phox and p22phox into lipid rafts after IL-8 and fMLP treatment and effect of MCD on IL-8-mediated priming of neutrophil oxidative burst. Neutrophils were pretreated with 50 ng/ml IL-8 or phosphate-buffered saline for different times (3 or 15 min at 37 °C) and stimulated with 10-7 M fMLP (2 min, 37 °C). Western blotting (A) and confocal microscopy (B) were performed as described in the legend to Fig. 8. One representative experiment of three is shown. C, effect of MCD on IL-8-mediated priming of neutrophil oxidative burst. Neutrophils were preincubated with 0.2 mg/ml MCD for 30 min at 37 °C, 50 ng/ml IL-8 for 3 or 15 min, and/or treated with 10-7 M fMLP. production was measured after stimulation lasting 5 min; the values reported are mean ± S.E. (n = 3). *, p < 0.05 versus IL-8 (3 min)/fMLP value (white bar). **, p < 0.05 versus IL-8 (15 min)/fMLP value (gray bar). Results were expressed as nanomole of cells/5 min.

The contribution of lipid rafts to the efficient activation of NADPH oxidase has been investigated in neutrophils, and suggests that the assembly of the enzyme complex depends on multiple, cooperative binding and signaling interactions that are greatly facilitated in a raft environment (24, 52). Here, we have provided the first evidence that the IL-8-induced intracellular signaling events may trigger the redistribution of the NADPH oxidase components into raft microdomains. In fact, we observed that short-term IL-8 treatment enhanced the insertion of the membrane-spanning flavocytochrome b₅₅₈ into DRMs in parallel with the rapid translocation of p47^phox and Rac2. Conversely, long-term IL-8 treatment generally failed to recruit the flavocytochrome b₅₅₈ into DRMs, but did enhance the translocation of p67^phox into the lipid microdomains. This suggests that once flavocytochrome b₅₅₈ has been incorporated into DRMs it is able to move laterally into non-detergent-resistant microdomains. Moreover, previous data have demonstrated that IL-8 induces significant degranulation of neutrophils after exposure for only 10 min, thus increasing the level of flavocytochrome b₅₅₈ at the neutrophil surface (53). This pool of flavocytochrome b₅₅₈ might be mobilized into DRMs in response to fMLP stimulation, which would constitute another potential mechanism for the IL-8-induced long-term priming. These various mechanisms play critical roles in the IL-8-induced priming of the oxidative burst in response to fMLP, as demonstrated by the inhibition of IL-8-mediated priming in response to fMLP by MCD. Moreover, lipid rafts have been shown to be involved in modulating a number of intracellular signaling pathways and functional responses in neutrophils. Disruption of the integrity of the DRMs by cholesterol-affecting agents, such as MCD, is known to alter intracellular signaling events and functional responses in neutrophils. MCD interferes with the membrane-dependent initiation of tyrosine phosphorylation (25), inhibits the phosphorylation of ERK1/2 induced by IL-8 (52), increases the phosphorylation of p38 MAPK, and inhibits NADPH oxidase activity. These findings thus suggest that lipid microdomains may mediate the efficiency of the oxidase by coupling NADPH oxidase components to certain intracellular signaling pathways in neutrophils.

Our study indicates that, in accordance with the widely accepted role of lipid rafts as signaling platforms, IL-8 induced the sequential preassembly and recruitment of the NADPH oxidase components in DRMs. It also suggested that lipid rafts are essential for the signaling events triggered by IL-8. This implies that lipid rafts must have important regulatory functions in chemoattractant-mediated responses in neutrophils. A better understanding of the role of raft microdomains in the priming of the neutrophils oxidative burst should provide valuable information about the regulation of NADPH oxidase activity and about the signaling pathways involved in inflammatory diseases.

	FOOTNOTES

 
^* This work was supported by the Institut National de la Santé et de la Recherche Médicale (INSERM) and a Ministère de l'enseignement et de la recherche grant (to C. G.). 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.

The on-line version of this article (available at http://www.jbc.org) contains supplemental Fig. 1.

¹ To whom correspondence should be addressed. Tel.: 33-1-44-85-62-09; Fax: 33-1-44-85-62-07; E-mail: guichard{at}bichat.inserm.fr.

² The abbreviations used are: ROS, reactive oxygen species; fMLP, formyl-methionyl-leucyl-phenylalanine; ERK, extracellular signal-regulated kinase; IL, interleukin; DRM, detergent-resistant microdomain; MCB, methyl--cyclodextrin; PKB, protein kinase B; PKC, protein kinase C; Pipes, 1,4-piperazinediethanesulfonic acid; BisTris, N,N-bis(2-hydroxyethyl)glycine; PI3K, phosphatidylinositol 3-kinase; MAPK, mitogen-activated protein kinase; MEK1/2, mitogen-activated protein kinase/extracellular signal-regulated kinase kinase.

	ACKNOWLEDGMENTS

 
We thank F. Adam for performing the Ca²⁺ analyses, P. Nicole for performing the fluorescence studies, and J. J. Lacapere for helpful suggestions.

	REFERENCES

TOP ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

 

1. Ago, T., Nunoi, H., Ito, T., and Sumimoto, H. (1999) J. Biol. Chem. 274, 33644-33653[Abstract/Free Full Text]
2. Babior, B. M. (1999) Blood 93, 1464-1476[Free Full Text]
3. Chanock, S. J., el Benna, J., Smith, R. M., and Babior, B. M. (1994) J. Biol. Chem. 269, 24519-24522[Free Full Text]
4. Park, J. W., and Babior, B. M. (1992) J. Biol. Chem. 267, 19901-19906[Abstract/Free Full Text]
5. Bokoch, G. M., and Diebold, B. A. (2002) Blood 100, 2692-2696[Abstract/Free Full Text]
6. Babior, B. M. (2004) Curr. Opin. Immunol. 16, 42-47[CrossRef][Medline] [Order article via Infotrieve]
7. Nauseef, W. M. (2004) Histochem. Cell Biol. 122, 277-291[CrossRef][Medline] [Order article via Infotrieve]
8. Yuo, A., Kitagawa, S., Ohsaka, A., Saito, M., and Takaku, F. (1990) Biochem. Biophys. Res. Commun. 171, 491-497[CrossRef][Medline] [Order article via Infotrieve]
9. Yuo, A., Kitagawa, S., Suzuki, I., Urabe, A., Okabe, T., Saito, M., and Takaku, F. (1989) J. Immunol. 142, 1678-1684[Abstract]
10. Yuo, A., Kitagawa, S., Kasahara, T., Matsushima, K., Saito, M., and Takaku, F. (1991) Blood 78, 2708-2714[Abstract/Free Full Text]
11. Weisbart, R. H., Golde, D. W., Clark, S. C., Wong, G. G., and Gasson, J. C. (1985) Nature 314, 361-363[CrossRef][Medline] [Order article via Infotrieve]
12. Kitagawa, S., Yuo, A., Souza, L. M., Saito, M., Miura, Y., and Takaku, F. (1987) Biochem. Biophys. Res. Commun. 144, 1143-1146[CrossRef][Medline] [Order article via Infotrieve]
13. Dewas, C., Dang, P. M., Gougerot-Pocidalo, M. A., and El Benna, J. (2003) J. Immunol. 171, 4392-4398[Abstract/Free Full Text]
14. Dang, P. M., Dewas, C., Gaudry, M., Fay, M., Pedruzzi, E., Gougerot-Pocidalo, M. A., and El Benna, J. (1999) J. Biol. Chem. 274, 20704-20708[Abstract/Free Full Text]
15. DeLeo, F. R., Renee, J., McCormick, S., Nakamura, M., Apicella, M., Weiss, J. P., and Nauseef, W. M. (1998) J. Clin. Investig. 101, 455-463[Medline] [Order article via Infotrieve]
16. Ward, R. A., Nakamura, M., and McLeish, K. R. (2000) J. Biol. Chem. 275, 36713-36719[Abstract/Free Full Text]
17. McColl, S. R., Beauseigle, D., Gilbert, C., and Naccache, P. H. (1990) J. Immunol. 145, 3047-3053[Abstract]
18. Berkow, R. L., and Dodson, M. R. (1988) J. Leukocyte Biol. 44, 345-352[Abstract]
19. Hallett, M. B., and Lloyds, D. (1995) Immunol. Today 16, 264-268[CrossRef][Medline] [Order article via Infotrieve]
20. Metzner, B., Barbisch, M., Parlow, F., Kownatzki, E., Schraufstatter, I., and Norgauer, J. (1995) J. Investig. Dermatol. 104, 789-791[CrossRef][Medline] [Order article via Infotrieve]
21. Wozniak, A., Betts, W. H., Murphy, G. A., and Rokicinski, M. (1993) Immunology 79, 608-615[Medline] [Order article via Infotrieve]
22. Daniels, R. H., Finnen, M. J., Hill, M. E., and Lackie, J. M. (1992) Immunology 75, 157-163[Medline] [Order article via Infotrieve]
23. Vilhardt, F., and Van Deurs, B. (2004) EMBO J. 23, 739-748[CrossRef][Medline] [Order article via Infotrieve]
24. Shao, D., Segal, A. W., and Dekker, L. V. (2003) FEBS Lett. 550, 101-106[CrossRef][Medline] [Order article via Infotrieve]
25. Katsumata, O., Hara-Yokoyama, M., Sautes-Fridman, C., Nagatsuka, Y., Katada, T., Hirabayashi, Y., Shimizu, K., Fujita-Yoshigaki, J., Sugiya, H., and Furuyama, S. (2001) J. Immunol. 167, 5814-5823[Abstract/Free Full Text]
26. Park, J. W., Benna, J. E., Scott, K. E., Christensen, B. L., Chanock, S. J., and Babior, B. M. (1994) Biochemistry 33, 2907-2911[CrossRef][Medline] [Order article via Infotrieve]
27. Woodman, R. C., Ruedi, J. M., Jesaitis, A. J., Okamura, N., Quinn, M. T., Smith, R. M., Curnutte, J. T., and Babior, B. M. (1991) J. Clin. Investig. 87, 1345-1351[Medline] [Order article via Infotrieve]
28. Benna, J. E., Dang, P. M., Gaudry, M., Fay, M., Morel, F., Hakim, J., and Gougerot-Pocidalo, M. A. (1997) J. Biol. Chem. 272, 17204-17208[Abstract/Free Full Text]
29. Amar, M., Amit, N., Huu, T. P., Chollet-Martin, S., Labro, M. T., Gougerot-Pocidalo, M. A., and Hakim, J. (1990) J. Immunol. 144, 4749-4756[Abstract]
30. Brown, G. E., Stewart, M. Q., Bissonnette, S. A., Elia, A. E., Wilker, E., and Yaffe, M. B. (2004) J. Biol. Chem. 279, 27059-27068[Abstract/Free Full Text]
31. Gilbert, C., Levasseur, S., Desaulniers, P., Dusseault, A. A., Thibault, N., Bourgoin, S. G., and Naccache, P. H. (2003) J. Immunol. 170, 5235-5243[Abstract/Free Full Text]
32. Dewas, C., Fay, M., Gougerot-Pocidalo, M. A., and El-Benna, J. (2000) J. Immunol. 165, 5238-5244[Abstract/Free Full Text]
33. Dang, P. M., Cross, A. R., and Babior, B. M. (2001) Proc. Natl. Acad. Sci. U. S. A. 98, 3001-3005[Abstract/Free Full Text]
34. Rotrosen, D., Kleinberg, M. E., Nunoi, H., Leto, T., Gallin, J. I., and Malech, H. L. (1990) J. Biol. Chem. 265, 8745-8750[Abstract/Free Full Text]
35. Nisimoto, Y., Motalebi, S., Han, C. H., and Lambeth, J. D. (1999) J. Biol. Chem. 274, 22999-23005[Abstract/Free Full Text]
36. Karathanassis, D., Stahelin, R. V., Bravo, J., Perisic, O., Pacold, C. M., Cho, W., and Williams, R. L. (2002) EMBO J. 21, 5057-5068[CrossRef][Medline] [Order article via Infotrieve]
37. Stahelin, R. V., Burian, A., Bruzik, K. S., Murray, D., and Cho, W. (2003) J. Biol. Chem. 278, 14469-14479[Abstract/Free Full Text]
38. Rietveld, A., and Simons, K. (1998) Biochim. Biophys. Acta 1376, 467-479[Medline] [Order article via Infotrieve]
39. Brown, D. A., and London, E. (1998) J. Membr. Biol. 164, 103-114[CrossRef][Medline] [Order article via Infotrieve]
40. Okamura, N., Curnutte, J. T., Roberts, R. L., and Babior, B. M. (1988) J. Biol. Chem. 263, 6777-6782[Abstract/Free Full Text]
41. Dang, P. M., Morel, F., Gougerot-Pocidalo, M. A., and Benna, J. E. (2003) Biochemistry 42, 4520-4526[CrossRef][Medline] [Order article via Infotrieve]
42. Forbes, L. V., Moss, S. J., and Segal, A. W. (1999) FEBS Lett. 449, 225-229[CrossRef][Medline] [Order article via Infotrieve]
43. Van Lint, J., Van Damme, J., Billiau, A., Merlevede, W., and Vandenheede, J. R. (1993) Mol. Cell Biochem. 127-128, 171-177
44. Knall, C., Worthen, G. S., and Johnson, G. L. (1997) Proc. Natl. Acad. Sci. U. S. A. 94, 3052-3057[Abstract/Free Full Text]
45. El Benna, J., Faust, R. P., Johnson, J. L., and Babior, B. M. (1996) J. Biol. Chem. 271, 6374-6378[Abstract/Free Full Text]
46. Waite, K. A., Wallin, R., Qualliotine-Mann, D., and McPhail, L. C. (1997) J. Biol. Chem. 272, 15569-15578[Abstract/Free Full Text]
47. Park, H. S., Lee, S. M., Lee, J. H., Kim, Y. S., Bae, Y. S., and Park, J. W. (2001) Biochem. J. 358, 783-790[CrossRef][Medline] [Order article via Infotrieve]
48. Chen, Q., Powell, D. W., Rane, M. J., Singh, S., Butt, W., Klein, J. B., and McLeish, K. R. (2003) J. Immunol. 170, 5302-5308[Abstract/Free Full Text]
49. Lian, J. P., Huang, R., Robinson, D., and Badwey, J. A. (1999) J. Immunol. 163, 4527-4536[Abstract/Free Full Text]
50. Varnai, P., Rother, K. I., and Balla, T. (1999) J. Biol. Chem. 274, 10983-10989[Abstract/Free Full Text]
51. van Bruggen, R., Anthony, E., Fernandez-Borja, M., and Roos, D. (2004) J. Biol. Chem. 279, 9097-9102[Abstract/Free Full Text]
52. Tuluc, F., Meshki, J., and Kunapuli, S. P. (2003) Int. Immunopharmacol. 3, 1775-1790[CrossRef][Medline] [Order article via Infotrieve]
53. Peveri, P., Walz, A., Dewald, B., and Baggiolini, M. (1988) J. Exp. Med. 167, 1547-1559[Abstract/Free Full Text]

CiteULike

Complore

Connotea

Del.icio.us

Digg

Reddit

Technorati

This article has been cited by other articles:

				R. D. Ye, F. Boulay, J. M. Wang, C. Dahlgren, C. Gerard, M. Parmentier, C. N. Serhan, and P. M. Murphy International Union of Basic and Clinical Pharmacology. LXXIII. Nomenclature for the Formyl Peptide Receptor (FPR) Family Pharmacol. Rev., June 1, 2009; 61(2): 119 - 161. [Abstract] [Full Text] [PDF]

				H. Li, W. Han, V. A. M. Villar, L. B. Keever, Q. Lu, U. Hopfer, M. T. Quinn, R. A. Felder, P. A. Jose, and P. Yu D1-Like Receptors Regulate NADPH Oxidase Activity and Subunit Expression in Lipid Raft Microdomains of Renal Proximal Tubule Cells Hypertension, June 1, 2009; 53(6): 1054 - 1061. [Abstract] [Full Text] [PDF]

				N. Semiramoth, A. Gleizes, I. Turbica, C. Sandre, R. Gorges, I. Kansau, A. Servin, and S. Chollet-Martin Escherichia coli type 1 pili trigger late IL-8 production by neutrophil-like differentiated PLB-985 cells through a Src family kinase- and MAPK-dependent mechanism J. Leukoc. Biol., February 1, 2009; 85(2): 310 - 321. [Abstract] [Full Text] [PDF]

				K. Omori, T. Ohira, Y. Uchida, S. Ayilavarapu, E. L. Batista Jr., M. Yagi, T. Iwata, H. Liu, H. Hasturk, A. Kantarci, et al. Priming of neutrophil oxidative burst in diabetes requires preassembly of the NADPH oxidase J. Leukoc. Biol., July 1, 2008; 84(1): 292 - 301. [Abstract] [Full Text] [PDF]

				W. Han, H. Li, V. A. M. Villar, A. M. Pascua, M. I. Dajani, X. Wang, A. Natarajan, M. T. Quinn, R. A. Felder, P. A. Jose, et al. Lipid Rafts Keep NADPH Oxidase in the Inactive State in Human Renal Proximal Tubule Cells Hypertension, February 1, 2008; 51(2): 481 - 487. [Abstract] [Full Text] [PDF]

				I. H. K. Dias, L. Marshall, P. A. Lambert, I. L. C. Chapple, J. B. Matthews, and H. R. Griffiths Gingipains from Porphyromonas gingivalis Increase the Chemotactic and Respiratory Burst-Priming Properties of the 77-Amino-Acid Interleukin-8 Variant Infect. Immun., January 1, 2008; 76(1): 317 - 323. [Abstract] [Full Text] [PDF]

				G. M. Fuhler, N. R. Blom, P. J. Coffer, A. L. Drayer, and E. Vellenga The reduced GM-CSF priming of ROS production in granulocytes from patients with myelodysplasia is associated with an impaired lipid raft formation J. Leukoc. Biol., February 1, 2007; 81(2): 449 - 457. [Abstract] [Full Text] [PDF]

				C. D. Ellson, K. Davidson, G. J. Ferguson, R. O'Connor, L. R. Stephens, and P. T. Hawkins Neutrophils from p40phox-/- mice exhibit severe defects in NADPH oxidase regulation and oxidant-dependent bacterial killing J. Exp. Med., August 7, 2006; 203(8): 1927 - 1937. [Abstract] [Full Text] [PDF]

				H. Fu, J. Karlsson, J. Bylund, C. Movitz, A. Karlsson, and C. Dahlgren Ligand recognition and activation of formyl peptide receptors in neutrophils J. Leukoc. Biol., February 1, 2006; 79(2): 247 - 256. [Full Text] [PDF]

This Article Abstract Full Text (PDF) Supplemental Data All Versions of this Article: 280/44/37021    most recent M506594200v1 Submit a Letter to Editor Alert me when this article is cited Alert me when eLetters are posted Alert me if a correction is posted Citation Map Services Email this article to a friend Similar articles in this journal Similar articles in PubMed Alert me to new issues of the journal Download to citation manager Request Permissions Citing Articles Citing Articles via HighWire Citing Articles via Google Scholar Google Scholar Articles by Guichard, C. Articles by Elbim, C. Search for Related Content PubMed PubMed Citation Articles by Guichard, C. Articles by Elbim, C. Social Bookmarking                      What's this?