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Am J Physiol Lung Cell Mol Physiol 294: L178-L186, 2008. First published November 9, 2007; doi:10.1152/ajplung.00263.2007
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EDITORIAL FOCUS

Neutrophil caveolin-1 expression contributes to mechanism of lung inflammation and injury

Guochang Hu,1,3 Richard D. Ye,1 Mary C. Dinauer,4 Asrar B. Malik,1,3 and Richard D. Minshall1,2,3

Departments of 1Pharmacology and 2Anesthesiology and 3Center for Lung and Vascular Biology, University of Illinois College of Medicine, Chicago, Illinois; and 4Herman B. Wells Center for Pediatric Research, Riley Hospital for Children, Indiana University School of Medicine, Indianapolis, Indiana

Submitted 6 July 2007 ; accepted in final form 6 November 2007


   ABSTRACT
TOP
 ABSTRACT
MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Caveolin-1 present in immune cells may be involved in regulation of the inflammatory response. Here, using caveolin-1-null (Cav-1–/–) mice, we addressed the role of caveolin-1 in polymorphonuclear neutrophils (PMNs) in regulating PMN activation-mediated lung injury. In lungs of wild-type (Cav-1+/+) mice perfused at constant flow with Krebs-Henseleit solution, addition of Cav-1+/+ PMNs (4 x 106 cells) into the perfusate followed by their activation with formyl-Met-Leu-Phe (fMLP, 1.0 µM) plus platelet-activating factor (1.0 nM) increased pulmonary microvessel filtration coefficient by 150% and wet-to-dry lung weight ratio by 50% as well as PMN accumulation in lungs. These responses were markedly reduced in lungs perfused with Cav-1–/– PMNs followed by addition of the same activating agents. fMLP-stimulated adhesion of Cav-1–/– PMNs to pulmonary microvascular endothelial cells and migration of Cav-1–/– PMNs across endothelial monolayers were also impaired compared with Cav-1+/+ PMNs. Cav-1–/– PMNs showed 50–80% reduction in PMA- or fMLP-stimulated superoxide production compared with Cav-1+/+ PMNs. In addition, Cav-1–/– PMNs had decreased migratory activity (50%) and adhesion to fibrinogen (40%) in response to fMLP. Rac1 and Rac2 were activated in Cav-1+/+ PMNs after stimulation of fMLP but not in Cav-1–/– PMNs. Exogenous expression of caveolin-1 in COS-phox cells augmented the fMLP-induced Rac1 activation and superoxide production, indicating a direct role of caveolin-1 in the mechanism of superoxide production. Thus caveolin-1 expression in PMNs plays a key role in mediating PMN activation, adhesion, and transendothelial migration and in PMN activation-induced lung inflammation and vascular injury.

caveolae; permeability; edema; Rac

CAVEOLAE, THE 50- TO 100-NM "flask-shaped" invaginations of the plasma membrane, are found in nearly all types of cells, but they are especially abundant in endothelial cells, adipocytes, vascular muscle cells, and fibroblasts (35). These organelles have been reported to play a role in endocytosis, oncogenesis, and uptake of bacteria and viruses (10, 35). Caveolin-1, the 22-kDa caveola-associated protein, exists primarily as membrane-associated oligomers on the cytoplasmic face of the plasma membrane. There are three isoforms of caveolin in vertebrates: caveolin-1 and caveolin-2 are the most prominent forms coexpressed in endothelial cells, adipocytes, and fibroblasts, whereas expression of caveolin-3 is cardiac and skeletal muscle specific. Caveolin-1 is a scaffolding protein that also functions to regulate multiple signaling pathways originating in caveolae. We (33, 40, 41, 43) and others (14, 26) have identified the key role of endothelial caveolin-1 in regulating transcellular albumin permeability. Studies have also implicated involvement of caveolin-1 in oxidant generation, leukocyte migration, and inflammation-induced tissue injury (5, 13, 52). Caveolin-1 was shown to be essential for angiotensin II-induced reactive oxygen species production secondary to the activation of NADPH oxidase in vascular smooth muscle cells (52). Knockdown of endothelial caveolin-1 reduced transendothelial migration of leukocytes (5). Caveolin-1-null (Cav-1–/–) mice exhibited significantly reduced polymorphonuclear neutrophil (PMN) binding and sequestration, lung microvascular permeability, and lung edema formation after LPS challenge (13).

The presence and function of caveolin-1 and caveolae in human PMNs have been a matter of uncertainty (39, 50). Evidence suggests that caveolin-1 and caveolae are present in immune cells, where caveolin-1 expression and distribution may be dependent on the activation and maturation state of cells (18). Caveolin-1 has been detected in human (50) and rat (38) PMNs, mouse (25, 46) and rat (24) macrophages and mast cells (42), bovine (19) and mouse (31) lymphocytes, human and bovine dendritic cells (47), and human T cell leukemia cell lines (20). Caveolin-1 in immune cells may be involved in regulating apoptosis, lipid metabolism, and endocytosis (11, 12). Also, caveolin-1 may serve an immunomodulatory function in macrophages (46). B lymphocytes from Cav-1–/– mice have reduced IgG3 secretion in response to LPS exposure (31). Caveola-disrupting agents blocked Escherichia coli entry into mouse bone marrow-derived mast cells, and markers of caveolae were actively recruited to sites of bacterial invasion (42).

In the present study, we observed caveolin-1 expression in mouse peripheral blood PMNs. Using PMNs obtained from Cav-1–/– mice, we investigated the role of PMN caveolin-1 in the mechanism of inflammatory lung injury. Our results show that PMN caveolin-1 regulates PMN-induced lung vascular injury by modulating PMN function such as the production of oxidants. These results raise the possibility that inhibition of PMN caveolin-1-activated signaling in inflammation represents a novel therapeutic strategy for treatment of lung inflammation and injury such as seen with acute lung injury.


   MATERIALS AND METHODS
TOP
 ABSTRACT
 MATERIALS AND METHODS
RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Animals. Caveolin-1-null mice and wild-type B6/129SJ mice were purchased from Jackson Laboratory (Bar Harbor, ME). Mice were housed in microisolator cages under specific pathogen-free conditions, fed with autoclaved food, and used in experiments at 8–12 wk of age. All studies using mice were approved by the University of Illinois Institutional Animal Care and Use Committee.

Endothelial cell cultures. Mouse lung vascular endothelial cells (MLVECs) were isolated from 3- to 5-day-old mouse pups as described previously (8). Briefly, lung tissues were minced and digested with collagenase A (1.0 mg/ml) for 45–60 min at 37°C. The released cells were centrifuged and suspended in 5 ml of suspension buffer and filtered through 200-µm and 60-µm mesh filters. Endothelial cells were purified with an anti-platelet endothelial cell adhesion molecule (PECAM)-1 MAb magnetic bead (BD Pharmingen, San Diego, CA) separation technique, suspended in 8 ml of growth medium, and transferred to a 100-mm culture dish. The endothelial cells were allowed to grow for 3–4 days and then identified by light microscopy. Cloning rings were placed around colonies of endothelial cells, and the cells were trypsinized and transferred to culture dishes. Endothelial cells were repeatedly purified by PECAM-1 MAb-coated magnetic beads to maintain >90% endothelial purity of cell monolayers. Endothelial cell cultures between passages 2 and 4 were used for these studies.

COS-phox cell culture model system. Transgenic COS-phox cells were generated as described previously (37). The cells were maintained in high-glucose DMEM supplemented with 10% FBS, 50 U/ml of penicillin, 50 µg/ml of streptomycin, 0.2 mg/ml hygromycin (Sigma-Aldrich, St. Louis, MO), 0.8 mg/ml neomycin (Invitrogen Life Technologies, Carlsbad, CA), and 1 mg/ml puromycin (Calbiochem, La Jolla, CA) at 37°C in a humidified atmosphere of 5% CO2 in room air. Lipofectamine Plus was used to transiently transfect 1–2 µg of DNA per 60-mm dish of COS-phox cells, which were then analyzed 24 h after transfection. Transient transfection efficiency, which was assayed by flow cytometry to detect expression of the Myc-epitope tag, averaged 45–55%.

Isolation and activation of mouse PMNs. Mouse PMNs were isolated from peripheral blood by hetastarch exchange transfusion as described previously (22, 48), with minor modification. Briefly, blood was collected from the jugular vein with exchange performed by alternatively withdrawing blood and infusing hetastarch solution (total ~8 ml) in 0.2-ml increments. The collected blood was maintained at room temperature to allow erythrocyte sedimentation, and the leukocyte-rich plasma layer was then centrifuged at 500 g for 10 min at 4°C. Contaminating erythrocytes in the pellet were removed with sterile distilled water and subsequent addition of 0.6 M KCl. The leukocyte-rich suspension (2 ml) was then layered on top of 3 ml of Ficoll-Paque and centrifuged at 800 g at 4°C for 20 min. The yield of PMNs obtained by our procedure was ~2 x106 cells per mouse. The purity of isolated PMNs was >98%, and viability was >95% as evaluated by Trypan blue exclusion (22).

The selection of formyl-Met-Leu-Phe (fMLP) as the PMN activator for these studies was based on the finding that fMLP stimulates mouse PMNs to release oxygen free radicals but not detectable amounts of elastase (23). Furthermore, fMLP activates primarily the PMNs and exerts no effect on endothelial cells (36). The phorbol ester PMA was also used as an alternate stimulator to address the effects of PMN caveolin-1 on superoxide production. In isolated lung studies, platelet-activating factor (PAF) was added as a costimulator with fMLP to augment the level of PMN-induced inflammatory response (28).

Lung preparation. Mouse lungs were isolated as described previously (45, 49). Briefly, after the animals were anesthetized and ventilated, a thoracotomy was performed and the pulmonary artery was cannulated in situ for perfusion of the lungs with a modified Krebs-Henseleit solution and mounted on a perfusion apparatus. The composition of the solution was as follows (mM): 118 NaCl, 4.7 KCl, 1.0 CaCl2, 1.0 MgCl2, 5.0 HEPES, 11 glucose, and 0.025 EDTA (pH 7.35–7.45), with 5 g/100 ml BSA. The lung preparation was perfused via the pulmonary artery at constant flow (2 ml/min) and venous pressure (3 cmH2O) and pulmonary artery pressures of 8 ± 2 cmH2O. Pulmonary arterial pressure and lung weight were continuously monitored during experiments. Labtech software (Andover, MA) was used to control data acquisition and storage. At the end of experiments, lungs were weighed, dried, and reweighed. Wet-to-dry lung weight ratio was used as an index of accumulation of lung water content. After a stabilization period of 15 min, PMNs (4 x 106 cells) from either Cav-1+/+ or Cav-1–/– mice, along with fMLP (1.0 µM) and PAF (1.0 nM), were infused for 30 min via a sidearm cannula and a syringe pump.

Capillary filtration coefficient measurement. At 30 min after infusion, the capillary filtration coefficient (Kf,c) was measured to determine pulmonary microvascular permeability to fluid. The rate of lung weight gain was obtained after a step increase (+6 cmH2O) in venous pressure and then normalized by the lung dry weight and recorded step size to calculate Kf,c (in ml·min–1·cmH2O–1·g dry lung–1). The analytic procedures used for computing Kf,c from recordings of lung wet weight were as described previously (45, 49).

Lung tissue MPO activity. Lung PMN sequestration was determined by measuring MPO activity (22). At the end of experiment, lungs were immediately removed, frozen, and stored at –70°C until being assayed. Lungs were homogenized in 5% hexadecyltrimethylammonium bromide buffer, sonicated three times for 15 s on ice, and centrifuged at 10,000 rpm for 30 min at 4°C. A 10-µl sample of the supernatant was loaded into a cuvette plate. o-Dianisidine dihydrochloride with 0.0005% hydrogen peroxide in phosphate buffer (190 µl) was then added to samples. Absorbance change was measured at 460 nm for 3 min. MPO activity was expressed as change in absorbance per minute per gram of tissue.

Lung histology. Cav-1+/+ mouse lungs, perfused with either Cav-1+/+ or Cav-1–/– PMNs plus fMLP for 30 min, were fixed with 10% formalin via tracheal injection for 1 h, harvested, and resuspended in 10% formalin overnight. Formalin-fixed tissue was washed with PBS and dehydrated in 70% ethanol before paraffin embedding. The fixation procedure was performed with a fixed pressure. Fourmicrometer-thick sections were stained with hematoxylin-eosin and examined by light microscopy.

Western blot analysis. Mouse PMN lysates were prepared as described previously (38). Equal amounts of protein were loaded on 12% acrylamide gels, separated by SDS-PAGE, and transferred to nitrocellulose membranes. The membranes were washed three times with TBS-T solution (0.05% Tween 20 in Tris-buffered saline) and incubated with primary antibodies (caveolin-1, tubulin, PKC-{delta}, Rac1, or Rac2; 1:5,000) in 5% nonfat dry milk. Incubation was carried out overnight; the membranes were then washed three times for 5 min each and incubated for 60 min with goat anti-rabbit (polyclonal) or anti-mouse (monoclonal) IgG conjugated to horseradish peroxidase (1:5,000). Membranes were washed three times for 5 min each, and the protein bands were detected with enhanced chemiluminescence reagent (Pierce, Rockford, IL). Molecular mass of the proteins was determined with known marker proteins. Relative intensities of various bands were measured with Scion Image (National Institutes of Health).

Rac1 and Rac2 activity assay. Rac activity was determined with glutathione S-transferase (GST)-P21-activated kinase binding domain (GST-PBD) as described previously (32). Cell lysates were clarified by centrifugation at 14,000 g for 2 min at 4°C, and then equal volumes were incubated with GST-PBD beads (15 µg) for 1 h at 4°C. The beads were then washed three times with wash buffer (50 mM Tris, pH 7.4, 1% Triton X-100, 150 mM NaCl, 10 mM MgCl2, 10 µg/ml each of aprotinin and leupeptin, and 0.1 mM PMSF), and the bound Rac was eluted from beads by boiling each sample in Laemmli sample buffer. Eluted samples and total cell lysate were then separated on 10% SDS-PAGE gels, transferred to nitrocellulose, blocked with 5% nonfat milk, and analyzed by Western blotting using monoclonal anti-Rac1 or polyclonal anti-Rac2 antibodies.

Immunofluoresence microscopy. Freshly isolated PMNs were rinsed in HEPES-buffered HBSS, fixed with 4% paraformaldehyde in HBSS, and then permeabilized with 0.1% Triton X-100 (Sigma) in HBSS. After incubation for 30 min in 5% goat serum, the cells were stained overnight at 4°C with anti-caveolin-1 polyclonal antibody or normal mouse IgG. Caveolin-1 expression (green) was visualized with Alexa 488-conjugated secondary antibody by confocal microscopy in optical sections midway through the cell. Cell nuclei (blue) were labeled with 4',6-diamidino-2-phenylindole dihydrochloride (DAPI). Confocal images were acquired with a laser scanning confocal microscope (Zeiss LSM 510) using 364- and 488-nm excitation laser lines to detect DAPI [band pass (BP)385–470 nm emission] and Alexa 488 (BP505–550 nm emission), respectively. Optical sections had a thickness of <1 µm (pinhole set to achieve 1 Airy unit).

PMN chemotaxis assay. PMN chemotaxis in response to fMLP was measured in 24-well-format Transwell filter chambers (pore size 3.0 µm; Corning-Costar) as described previously (9). PMNs (1x 105) in HBSS (50 µl) were placed into the upper chamber, and fMLP was added into the lower chamber in HBSS. Transwells with PMNs were incubated for 45 min at 37°C in 5% CO2 to allow PMN migration. PMNs in three randomly chosen x20 fields were counted to determine the number of migrated PMNs. The checkerboard assay was used to distinguish between chemotaxis (directed movement) and chemokinesis (random movement). Varying concentrations of fMLP were placed in lower wells (to assess chemotaxis) or in both upper and lower wells (to assess chemokinesis). Using this system, we observed that fMLP-induced PMN migration into the lower well was a result of chemotaxis rather than chemokinesis, confirming previous observations (15).

PMN adhesion assay. PMNs were placed in flat-bottom 96-well-format plates coated with fibrinogen (250 g/ml) in the absence or presence of fMLP (1.0 µM) for 30 min at 37°C. After thorough washing, adherent PMNs were quantified by determining membrane acid phosphatase activity (2). Acetate buffer (0.15 M acetate, 0.2% Triton X-100, pH 5.3) was added into the wells for 5 min, followed by 10 mM p-nitrophenyl phosphate in acetate buffer. The reaction was stopped by the addition of 2 N NaOH. The p-nitrophenol produced in the reaction was measured at 405 nm. The percentage of adherent PMNs was calculated on the basis of a standard curve obtained with a known number of PMNs.

For detection of PMN adhesion to endothelium, confluent monolayers of MLVECs were prepared in 96-well plates in 250 µl of EBM-2 medium supplemented with 2% FBS (13). Peripheral blood PMNs were loaded with 2 µg/ml calcein-AM (Molecular Probes) for 30 min at room temperature and then added to MLVECs. The cells were then incubated with fMLP (1.0 µM) for 60 min. After fMLP treatment, the MLVECs were washed with PBS and the fluorescence was measured in duplicate with a spectrofluorometer (Photon Technology International) at excitation and emission wavelengths of 485 and 535 nm, respectively.

PMN transmigration assay. Transendothelial PMN migration was determined as described previously (4). MLVECs were plated onto 24-well-format Transwell filter inserts in 200 µl of 10% FBS-containing EBM-2 medium and allowed to grow to confluence. Five hundred microliters of serum-free DMEM was added to the lower chamber of each well. Immediately before the addition of PMNs, the upper chambers were washed twice with serum-free DMEM and medium in the lower chambers was replaced with 500 µl of serum-free DMEM or serum-free DMEM with 1.0 µM fMLP. Cells (2 x 105 in 200 µl of medium) were added to the upper chamber. After 3 h at 37°C in 5% CO2, nonadherent cells in the upper chamber were removed. Medium, including migrated PMNs, was collected from the lower chamber by repeatedly rinsing the lower chamber; absence of additional adherent PMNs was confirmed microscopically. The medium and all washes were pooled, pelleted, and resuspended, and PMNs were counted with a hemocytometer. All determinations were carried out in duplicate and repeated at least twice.

Superoxide production. PMNs and COS-phox cells were resuspended in BSA buffer (0.5% BSA in HBSS containing Ca2+, Mg2+, and 10 mM HEPES) at 5 x 106 cells/ml. Superoxide anion released was measured as described previously (29). Briefly, isoluminol was added to the cell suspension to a final concentration of 50 µM, and horseradish peroxidase was added to a final concentration of 40 U/ml. Cells were then seeded into 96-well-format flat-bottom tissue culture dishes (E&K Scientific, Campbell, CA). Chemiluminescence (CL) was measured every minute with a Wallac multilabel counter plate reader (Perkin Elmer, Boston, MA) starting from 5 min before and continuing for 50 min after stimulation with fMLP (1.0 µM) or PMA (400 ng/ml). Unstimulated PMNs or COS-phox cells (control) were recorded simultaneously. Relative level of superoxide production was calculated based on the integrated CL intensity (peak area) during the first 20 min (COS-phox cells) or 10 min (PMN stimulation) after agonist stimulation. In some experiments, Cav-1+/+ PMNs were pretreated with methyl-β-cyclodextrin, the cholesterol-binding agent that disassembles caveolae (51). This pretreatment consisted of incubation of PMNs with 10.0 mM methyl-β-cyclodextrin in HBSS for 30 min at 37°C, followed by two washes with HBSS.

Drugs and reagents. Chemicals and reagents used were obtained from Sigma (St. Louis, MO) unless otherwise stated. Horseradish peroxidase-conjugated goat anti-rabbit and anti-mouse IgG were obtained from Santa Cruz Biotechnology (Santa Cruz, CA). Anti-Rac1 monoclonal and anti-Rac2 polyclonal antibodies were from Upstate (Lake Placid, NY). Alexa 488 goat anti-mouse and anti-rabbit IgG, Alexa 488-albumin, and DAPI were purchased from Molecular Probes (Eugene, OR). HBSS containing NaHCO3 (4.2 mM) and HEPES (10 mM) was adjusted to pH 7.4. EBM-2 medium and FBS were obtained from Clonetics (Walkersville, MD) and Hyclone (Logan, UT), respectively.

Statistical analysis. One-way analysis of variance and Student's Newman-Keuls test for post hoc comparisons were used to determine differences between control and experimental groups. Student's t-test was performed for paired samples. Data are expressed as means ± SD. Differences were considered significant when P < 0.05.


   RESULTS
TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
DISCUSSION
 GRANTS
 REFERENCES
 
Caveolin-1 expression in peripheral mouse PMNs. A band of 22 kDa (molecular mass for caveolin-1) was detected by using a caveolin-1-specific antibody in Cav-1+/+ mouse PMN lysates but not in Cav-1–/– mouse PMNs. Compared with mouse pulmonary endothelial cells, the level of caveolin-1 expressed in PMNs was remarkably lower (Fig. 1A). To further characterize localization of caveolin-1 expressed in mouse PMNs, we incubated freshly isolated mouse PMNs with caveolin-1 antibody followed by secondary fluorescent antibody. Consistent with immunoblot results, caveolin-1 immunostaining was detected in Cav-1+/+ PMNs but not in Cav-1–/– PMNs (Fig. 1B).


Figure 1
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  		Fig. 1. Caveolin-1 (Cav-1) protein expression in peripheral mouse neutrophils. A: Cav-1 expression in wild-type (Cav-1+/+) polymorphonuclear neutrophils (PMNs) was detected by Western blot analysis. Cav-1+/+ mouse lung vascular endothelial cells (MLVECs) and Cav-1-null (Cav-1–/–) mouse PMN lysates were used as positive and negative controls; 60 µg of PMN and 10 µg of MLVEC lysates were loaded. B: confocal images show Cav-1 (green) expression in Cav-1+/+ PMNs and 4',6-diamidino-2-phenylindole dihydrochloride (DAPI) staining of the nucleus (blue) in mouse PMNs. Scale bar, 5 µm. Data in both A and B are representative of 3 experiments.

 
PMN caveolin-1 contributes to mechanism of microvascular injury and edema formation in mouse lungs. In Cav-1+/+ mouse lungs, infusion of Cav-1+/+ PMNs (4 x 106 cells) followed by fMLP (1.0 µM) plus PAF (1.0 nM) infusion for 30 min increased pulmonary capillary filtration coefficient by 150% and lung wet-to-dry weight ratio by 50% (Fig. 2, A and B). We also observed significant PMN accumulation in lung tissue (Fig. 2, C and D). These responses were abolished when Cav-1–/– PMNs followed by fMLP plus PAF were infused into Cav-1+/+ mouse lung (Fig. 2, A–C). Neither PAF nor fMLP alone, nor the combination of these mediators, affected Kf,c or wet-to-dry lung weight ratio in the absence of Cav-1+/+ PMNs.


Figure 2
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  		Fig. 2. Effect of PMN Cav-1 on pulmonary microvessel liquid permeability [lung capillary filtration coefficient (Kf,c), A] pulmonary edema formation [wet-to-dry (W/D) lung wt ratio, B], and PMN infiltration (MPO activity, C) in mouse lungs (see MATERIALS AND METHODS for details). PMNs (4 x 106 cells) from either wild-type (Cav-1+/+) or Cav-1-null (Cav-1–/–) mice along with formyl-Met-Leu-Phe (fMLP, 1.0 µM) and platelet-activating factor (PAF, 1.0 nM) were infused for 30 min (n = 6–8 for each group). *P < 0.05 vs. control (A and B) or PMN (C) group; {dagger}P < 0.05 compared with corresponding wild type. D: hematoxylin-eosin-stained lung sections. Left: infiltrating PMNs and septal thickening indicative of edema in mouse lungs perfused with Cav-1+/+ PMNs and fMLP. Right: normal alveolar septa and no obvious PMN infiltration in mouse lung perfused with Cav-1–/– PMNs + fMLP. Magnification x25.

 
We performed histological analysis at 30 min after administration of Cav-1+/+ or Cav-1–/– PMNs with fMLP. Activated Cav-1+/+ PMNs were actively recruited into lungs and caused septal thickening (edema), whereas these effects were markedly reduced when Cav-1–/– PMNs were added to the perfusate (Fig. 2D).

Caveolin-1 deletion in PMNs reduces PMN adhesion to endothelial cells. We examined static adhesion of PMNs to dishes coated with fibrinogen (Fig. 3A). Compared with the vehicle control group (22% of cells adhered), we observed a significant increase in Cav-1+/+ PMN adhesion on fMLP stimulation (47%). However, Cav-1–/– PMN adhesion to fibrinogen in response to fMLP was significantly reduced (30%) even though unstimulated Cav-1–/– PMN adhesion was not different from that of Cav-1+/+ PMNs (Fig. 3A). Next, we examined the role of PMN caveolin-1 in adhesion to resting MLVECs. Confluent monolayers of MLVECs were incubated for 30 min with PMNs in the absence or presence of fMLP. Endothelial monolayer adhesion of fMLP-activated PMNs more than doubled relative to vehicle-treated Cav-1+/+ PMNs, whereas this effect of fMLP was significantly reduced in Cav-1–/– PMNs (Fig. 3B).


Figure 3
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  		Fig. 3. Cav-1 participates in the mechanism of PMN adhesion. A: PMN adhesion to fibrinogen-coated dishes. Wild-type (Cav-1+/+) or Cav-1-null (Cav-1–/–) mouse PMNs were placed onto fibrinogen-coated dishes for 30 min with or without (Control) fMLP. Nonadherent cells were removed by gentle washing and % of adherent cells was determined (n = 8 wells for each stimulus). B: PMN adhesion to MLVECs. Fluorescently labeled Cav-1+/+ or Cav-1–/– mouse PMNs were allowed to adhere to confluent monolayers of MLVECs for 60 min in the presence or absence of fMLP. MLVECs were then washed, and fluorescence was measured (arbitrary units) in duplicate (n = 6 wells for each stimulus). Data in both A and B are representative of 3 experiments. *P < 0.05 vs. control group; {dagger}P < 0.05 compared with corresponding wild type.

 
Deletion of caveolin-1 in PMNs inhibits PMN chemotaxis and transendothelial migration. We examined agonist-induced PMN chemotaxis in vitro in the Transwell chamber. A PMN suspension in the upper chamber was separated by a filter with a pore size of 3 µm from a lower chamber containing medium with chemoattractant. PMN chemotaxis from the upper chamber into the lower chamber was assessed after 30 min by counting the number of cells in the bottom chamber attached to the filter. Cav-1+/+ PMNs showed 114% and 185% increase in migratory activity in response to 1.0 and 10 µM fMLP, respectively. The chemotactic response was significantly reduced in Cav-1–/– PMNs, which showed only 23% and 61% increase in chemotaxis in response to fMLP over vehicle control (Fig. 4A). A similar effect was observed in the presence of wild-type mouse lung endothelial monolayers: fMLP greatly increased Cav-1+/+ PMN transendothelial migration into the bottom compartment, whereas fMLP-stimulated transendothelial migration was reduced in the Cav-1–/– PMN group (Fig. 4B).


Figure 4
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  		Fig. 4. Role of Cav-1 in PMN chemotaxis and transendothelial migration. A: Transwell chemotaxis assay. Wild-type (Cav-1+/+) and Cav-1-null (Cav-1–/–) mouse PMN migration in response to gradients of fMLP was measured for 45 min in Transwell chambers (n = 8 wells for each stimulus). B: transmigration of PMNs across unactivated MLVEC monolayers. Purified mouse PMNs from Cav-1+/+ or Cav-1–/– mice were added to the top chamber, in which confluent monolayers of MLVECs were grown on a microporous filter. DMEM containing fMLP (1.0 µM) or DMEM alone was added to the bottom chamber. After 3 h at 37°C in 5% CO2, PMNs that had migrated across the monolayer were collected from the bottom chamber and counted (n = 6–8 wells for each stimulus). *P < 0.05 vs. control group; {dagger}P < 0.05 compared with corresponding wild type.

 
Deletion of caveolin-1 reduces PMN superoxide production. We measured agonist-stimulated superoxide generation by Cav-1+/+ and Cav-1–/– mouse peripheral blood PMNs. Freshly isolated PMNs from Cav-1–/– mice showed a 50–80% reduction in PMA- and fMLP-stimulated superoxide production compared with Cav-1+/+ PMNs (Fig. 5, A and B). To address whether lipid rafts/caveolae are involved in the mechanism of superoxide production, we pretreated Cav-1+/+ PMNs with methyl-β-cyclodextrin, a lipid raft/caveola-disrupting agent, as described in MATERIALS AND METHODS. Pretreatment of PMNs with methyl-β-cyclodextrin significantly reduced both fMLP- and PMA-induced superoxide production (Fig. 5C).


Figure 5
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  		Fig. 5. Superoxide production in wild-type (Cav-1+/+) and Cav-1-null (Cav-1–/–) mouse PMNs. PMNs were stimulated with PMA (400 ng/ml, A) or fMLP (1.0 µM, B). Superoxide production was measured by isoluminol-enhanced chemiluminescence assay and recorded at 37°C for 50 min. Data in both A and B are representative of 4 experiments. C: integrated chemiluminescence (CL) values showing inhibitory effects on fMLP- or PMA-induced superoxide production in Cav-1+/+ mouse PMNs pretreated with methyl-β-cyclodextrin (MβCD). *P < 0.05 vs. control (buffer) group; {dagger}P < 0.05 compared with corresponding stimulator groups. Data in C are representative of 3 experiments.

 
Caveolin-1 deletion prevents fMLP-induced Rac activation in mouse PMNs. We determined the relative level of fMLP-stimulated Rac1-GTP and Rac2-GTP in peripheral blood PMNs from Cav-1+/+ and Cav-1–/– PMNs by P21 binding domain (PBD) pull-down assay. Stimulation of PMNs with fMLP induced the formation of both Rac1- and Rac2-GTP, indicating Rac1 and Rac2 activation in Cav-1+/+ mouse PMNs. However, in Cav-1–/– mouse PMNs, fMLP did not induce the activation of Rac1, and Rac2 activation was reduced (Fig. 6).


Figure 6
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  		Fig. 6. fMLP activation of Rac1 and Rac2 in wild-type (Cav-1+/+) and Cav-1-null (Cav-1–/–) mouse PMNs. PMNs were stimulated with fMLP (1.0 µM) for 60 s, followed by solubilization and Rac activity measurement. Immunoblot from a representative experiment shows activation of Rac1 and Rac2 in peripheral mouse PMNs. Results are representative of at least 3 experiments.

 
Expression of caveolin-1 induces Rac activation and superoxide production in reconstituted COS-phox cells. We used the COS-phox cell line expressing gp91phox, p67phox, p22phox, and p47phox as a model system to investigate the role of caveolin-1 in NADPH oxidase activation by fMLP (21, 27). A previous study showed that PKC-{delta} is critical for fMLP-induced superoxide production in COS-phox cells and PMNs (21). Therefore, since COS-phox cells contain relatively little PKC-{delta} and caveolin-1, we transfected COS-phox cells with formyl peptide receptor, PKC-{delta}, and caveolin-1 cDNAs to determine the role of caveolin-1 in fMLP activation of NADPH oxidase and superoxide production. Figure 7A shows expression levels of endogenous and exogenous PKC-{delta} and caveolin-1. Because COS-phox cells only express Rac1 (21), we determined the effects of caveolin-1 overexpression on Rac1 activation as well as superoxide production. As shown in Fig. 7B, expression of exogenous PKC-{delta} and caveolin-1 did not affect the basal level of Rac1-GTP. However, in response to fMLP, expression of PKC-{delta} alone significantly increased the formation of Rac1-GTP, consistent with previous observations (21). When both caveolin-1 and PKC-{delta} were expressed in COS-phox cells, both Rac1 activation and fMLP-induced superoxide production were significantly greater than seen in PKC-{delta}-expressing COS-phox cells. Thus, in this "add-back" experiment, caveolin-1 regulated NADPH oxidase activity and superoxide production by facilitating fMLP-induced Rac1 activation.


Figure 7
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  		Fig. 7. Effects of exogenous Cav-1 expression on fMLP-induced Rac1 activation and superoxide production in COS-phox cells. COS-phox cells were transfected with formyl peptide receptor (FPR), PKC-{delta}, and/or caveolin-1 cDNA (see MATERIALS AND METHODS for details). A: endogenous and exogenous expression of PKC-{delta} and caveolin-1 were detected by Western blot analysis using PKC-{delta}- and caveolin-1-specific antibodies. Representative blots from 4 experiments are shown. B: activation of Rac1 by fMLP. Transfected COS-phox cells were stimulated with fMLP (1.0 µM) for 60 s, followed by solubilization and Rac activity measurement by glutathione S-transferase-P21-activated kinase binding domain pull-down assay and immunoblot. Representative blots from 3 experiments are shown. C: integrated chemiluminescence (Int. CL) from the first 20 min showing % difference in superoxide production with and without exogenous Cav-1. Data are from 3 experiments. *P < 0.05 vs. control (buffer) group; {dagger}P < 0.05 vs. corresponding FPR + PKC-{delta} group.

 

   DISCUSSION
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 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
GRANTS
 REFERENCES
 
Here we showed caveolin-1 expression in peripheral mouse PMNs by immunostaining and Western blot analysis. Although the level of caveolin-1 in mouse PMNs was lower than seen in endothelial cells, we found, as described below, that caveolin-1 has a profound role in the mechanism of PMN activation and PMN activation-mediated lung injury.

It is fair to say that the function of caveolin-1 in PMNs is incompletely understood. A recent study reported that caveolin-1 can regulate LPS-induced cytokine production in macrophages (46). Caveolin-1-deficient mice also showed defects in innate immunity and inflammatory immune responses (30) and exhibited reduced PMN binding and sequestration, lung microvascular permeability, and lung edema formation after LPS challenge (13). These observations suggest (but do not prove) that PMN caveolin-1 may be important in regulating the inflammatory response induced by the activation of PMN. Thus, to address this question, we infused either Cav-1+/+ or Cav-1–/– PMNs (obtained from knockout mice) along with the PMN-activating mediators fMLP and PAF into isolated, Krebs-Henseleit-perfused Cav-1+/+ lungs as described in MATERIALS AND METHODS. Activation of Cav-1+/+ PMNs in these lungs increased microvessel fluid permeability and PMN accumulation and caused lung edema formation. However, we observed that lung inflammation and injury were markedly reduced on perfusion of Cav-1+/+ mouse lungs with Cav-1–/– PMNs and the activation of these caveolin-1-deficient PMNs. These data show that caveolin-1-deficient PMNs have a reduced ability to induce PMN activation-mediated lung vascular injury and edema formation.

Superoxide production induced by activation of PMNs adherent to the vessel wall can cause endothelial injury leading to edema formation. Formation of superoxide is the first of several steps resulting in the production of other oxygen-derived reactive species. These include hydrogen peroxide and hydroxyl radical. To address the role of caveolin-1 in regulating PMN activation, we studied PMN superoxide production. Freshly isolated PMNs from Cav-1–/– mice showed a 50–80% reduction in PMA- or fMLP-stimulated superoxide production compared with Cav-1+/+ PMNs. In addition, we observed, using the cell-based NADPH oxidase reconstitution system (COS-phox cells), that exogenous expression of caveolin-1 increased the fMLP-stimulated superoxide production. Although the COS-phox cell system does not completely mimic PMNs, it is a well-accepted in vitro reconstitution system that has the advantage of titrating in signaling molecules to assess their impact on NADPH oxidase activation (21, 37). Our findings in reconstituted cells demonstrate the important role of caveolin-1 in mediating NADPH oxidase activation-induced superoxide production. Some previous findings support a role of caveolin-1 in regulating superoxide production, although our observations are the first to our knowledge to identify clearly that caveolin-1 is crucial in the mechanism of activation of phagocytic cells. In vascular smooth muscle cells, caveolin-1 was involved in mediating angiotensin II-induced reactive oxygen species production (52), and disruption of lipid rafts and caveolae prevented TNF-{alpha}-induced superoxide production in endothelial cells (51).

How does caveolin-1 regulate PMN oxidant production? The primary source of superoxide anion production in PMNs is through the NADPH oxidase enzyme complex. The phagocyte NADPH oxidase complex consists of two membrane-bound components, gp91phox and p22phox, and three cytoplasmic subunits, p47phox, p67phox, and p40phox. Rac GTPases Rac1 and Rac2 are key molecular switches regulating NADPH oxidase complex activity and thereby superoxide production (3, 7). On agonist stimulation, phosphorylated p47phox, p67phox, p40phox, and activated Rac1 are recruited to the plasma membrane to form the functional NADPH oxidase complex in association with gp91phox/p22phox (1, 21). Our findings show that caveolin-1 participates in the activation of NADPH oxidase in PMNs, but the mechanisms are not understood. Caveolin-1 could mediate the response at multiple levels to promote the assembly of NADPH oxidase as well as its localization to the plasma membrane of phagocytes. Caveolin-1-mediated activation of Rac1 may in particular be involved in NADPH oxidase assembly. Both Rac1 and NADPH oxidase subunits were shown to localize in caveolin-1-containing caveolae in various cell types (44). Rac also dynamically associates with caveolae and lipid rafts after agonist stimulation of endothelial cells (6, 16). The regulatory effect of caveolin-1 on Rac activation may be cell type specific. In bovine aortic endothelial cells, caveolin-1 knockdown enhanced both basal and agonist-induced Rac activity (16), and caveolin-1-deficient mouse embryonic fibroblasts exhibited increased Rac activity (17); thus caveolin-1 may serve as an inhibitory signal required for Rac activation. However, a study using a caveolin-1 small interfering RNA knockdown demonstrated that caveolin-1 facilitated Rac1 activation and Rac1 translocation to the membrane of vascular smooth muscle cells (52). Caveolin-1 was also shown to bind Rac1 and to keep Rac1 in its active conformation (52). Thus caveolin-1-mediated activation of Rac1 may be important in the mechanism of PMN activation. In the present study, we observed that Rac1 and Rac2 activation did not occur in response to fMLP in Cav-1–/– PMNs. Also, exogenous expression of caveolin-1 in COS-phox cells enhanced fMLP-induced Rac-1 activation as well as concomitant superoxide production.

PMN adhesion to pulmonary microvascular endothelial cells and migration into the air space are the main features in the pathogenesis of acute inflammatory lung injury, and thus we evaluated the role of caveolin-1 in the mechanism of PMN adhesion and migration. Cav-1–/– PMNs showed decreased migration and adhesion to fibrinogen in response to fMLP. Using pulmonary microvascular endothelial cell monolayers, we also showed that activated Cav-1–/– PMNs adhered and migrated less effectively than Cav-1+/+ PMNs to and across the endothelium. These findings show that caveolin-1 is essential for PMN adhesion and migration, consistent with the proposed role of caveolin-1 in mediating migration of endothelial cells (34). Thus caveolin-1 may be a generalized factor regulating cell migration, but the mechanisms are far from clear. Caveolin-1 may regulate PMN migration via its effects on cytoskeletal remodeling, signal transduction, alterations in the lipid profile of the plasma membrane, or all of the above.

The present study differs from a previous study that demonstrated marked attenuation of LPS-induced PMN sequestration and edema formation in lungs of Cav-1–/– mice (13). In the present study, fMLP was used to address specifically the role of PMN caveolin-1 in the mechanism of PMN activation-mediated lung injury. LPS activates multiple cell types including endothelial and epithelial cells (23, 36). Also, we used the approach of studying solely the effects of PMN caveolin-1; that is, PMNs isolated from Cav-1–/– mice were perfused into the isolated Cav-1+/+ mouse lung. This approach enabled us to address specifically the role of PMN caveolin-1 in the mechanism of PMN activation-induced lung inflammation and injury.

In summary, we have demonstrated that caveolin-1 expressed in peripheral mouse PMNs contributes to PMN activation-mediated acute lung inflammation and injury and does so by increasing PMN superoxide production and adhesion and migration responses. Caveolin-1 may regulate PMN activation through its ability to control Rac activity in response to PMN secretagogues such as fMLP. Thus our findings suggest that PMN-expressed caveolin-1 plays an important role in the mechanism of PMN activation-mediated lung inflammation and injury responses.


   GRANTS
TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
REFERENCES
 
This work was supported by National Heart, Lung, and Blood Institute Grants HL-71626 (R. D. Minshall) and P01-HL-60678 (A. B. Malik and R. D. Minshall) and American Heart Association Grant 0730331N (G. Hu).


   ACKNOWLEDGMENTS
 
We thank David J. Visintine and Maricela Castellon (Depts. of Pharmacology and Anesthesiology, University of Illinois College of Medicine) for technical assistance.


   FOOTNOTES
 

Address for reprint requests and other correspondence: G. Hu, Dept. of Pharmacology (m/c 868), Univ. of Illinois College of Medicine, 835 South Wolcott Ave., Chicago, IL 60612 (e-mail: gchu{at}uic.edu)

The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.


   REFERENCES
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 MATERIALS AND METHODS
 RESULTS
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 REFERENCES
 

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