Introduction

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Immunoglobulin (Ig)A is the predominant Ig isotype in secretory fluids such as gastrointestinal, respiratory, and urogenital secretions. In secretions, IgA is predominantly polymeric (p-IgA) and is synthesized by local mucosal plasma cells before it is transported through epithelial cells by their polymeric Ig receptor (pIgR) (1) and secreted into the mucosal fluids as secretory IgA (S-IgA), which consists of dimeric or p-IgA complex containing a joining J chain and secretory component. In contrast, serum IgA is largely monomeric and derived from plasma cells in the bone marrow (2). Polymeric and especially S-IgA, which constitutes the main mediator of specific humoral immunity at mucosal surfaces, neutralize bacterial toxins and viral particles (3) and inhibit adherence of bacteria to epithelial cells (4). Despite some well-recognized anti-infectious properties of IgA antibodies (Abs), the mechanisms of IgA-derived cell responses still remain poorly understood. The interaction of IgA with pIgR on epithelial cells and with IgA-specific Fc high affinity receptors (FcR [CD89]) expressed on a wide variety of blood and tissue myeloid cells (5, 6) may provide additional elements to explain the effector functions of IgA. FcR are heavily but variably glycosylated transmembrane proteins on leukocytes (5). They are expressed on the cell surface in association with the common FcR -chain homodimer (7). The FcR -chain is recognized as a signaling molecule, which, upon FcR crosslinking, is phosphorylated on its immunoreceptor tyrosine-based activation motif (ITAM) and leads to the activation of several sets of cytoplasmic protein tyrosine kinases, such as those of the Src family, and further downstream proteins such as protein kinase C (PKC) and phospholipase C (PLC) (8). Crosslinking of FcR can trigger several biologic responses, including phagocytosis of IgA immune complexes (9, 10), killing of IgA-opsonized bacteria (11), generation of reactive oxygen intermediates (ROI) (12), and release of inflammatory cytokines such as tumor necrosis factor (TNF)- and interleukin (IL)-6 (13). However, the interaction of FcR with IgA is not exclusively associated with the activation of proinflammatory processes. Other studies have shown that IgA can also downregulate the oxidative burst and the release of proinflammatory cytokines such as TNF- and IL-6 by monocytes and neutrophils (14, 15). This suggests that IgA could also to some extent control mucosal and systemic inflammatory responses, thereby protecting tissues from injury. These observations suggest that the ability of IgA to activate leukocytes through FcR does not necessarily conflict with its proposed anti-inflammatory activity, and that IgA-mediated cell responses depend not only on the type of costimulatory signals and the nature of the effector cell triggered but also on the molecular form of IgA. Even though the biology of FcR has been widely investigated in monocytes, neutrophils and eosinophils, the molecular mechanisms of IgA-mediated cellular responses still remain poorly understood.

FcR are constitutively well expressed on human alveolar macrophages (HAM) (16), an essential cell population at the interface of respiratory mucosal and systemic immunity, representing therefore the first line of defense in the lung (19). FcR of HAM bind both IgA1 and IgA2, although the IgA1 subclass predominates in lung secretions. However, upon deglycosylation, FcR of HAM are resolved to a protein core of 28 kD (18) instead of 32 kD in monocytes (5). Indeed, HAM express at their surface a protein product of an alternatively spliced FcR transcript (18). The biologic importance and the IgA-mediated signaling through these receptors, however, remain unknown.

In this study, we have investigated the effect of HAM's FcR binding by p- and S-IgA, the two main and abundant forms of IgA in mucosal secretions, on the oxidative burst and the production of the inflammatory cytokine TNF-, as well as the potential involvement of extracellular signal-related protein kinases 1 and 2 (ERK1/2) and NF-B activation in the modulation of FcR function in resting, lipopolysaccharide (LPS)-, and phorbol myristate acetate (PMA)-activated HAM. The mitogen-activated protein (MAP) kinases superfamily represents an important signaling pathway for many cell functions (20, 21). NF-B, which is held in an inactivated state in the cytoplasm of unstimulated cells through interaction with a class inhibitory proteins of IB family (22), translocates into the nucleus in activated cells upon IB phosphorylation, polyubiquitination, and subsequent degradation (23). NF-B controls the expression of several genes, especially cytokines such as TNF- and IL-8, at their level of gene transcription.

We found that the ERK1/2 pathway plays a central role in the IgA-modulated respiratory burst in both LPS- and PMA-stimulated HAM. Activated ERK kinases also control the production of TNF- in both LPS (CD14 receptor-dependent) and PMA (receptor-independent) membrane activation of HAM, but not in IgA-treated HAM. Both LPS and IgA, but not PMA, activate NF-B nuclear translocation through the classic pathway of IB phosphorylation and proteolysis. Activated NF-B plays an important role in TNF- release by LPS-activated HAM and only a minor role in that by IgA-treated HAM, suggesting that an alternative pathway distinct from ERK1/2 and NF-B is implicated in the IgA-mediated upregulation of TNF- secretion. Oxidants do not play an important role in NF-B activation and play a minor role in LPS-induced TNF- release. In contrast, PMA do not activate NF-B and activates HAM essentially through ERK1/2 MAP kinase pathway.

	Materials and Methods

Top Abstract Introduction Materials and Methods Results Discussion References

Reagents and Antibodies

Recombinant human TNF- (rh-TNF-, specific activity > 1 × 10⁸ U/mg) was purchased from Boehringer (Boehringer Mannheim GmbH, Mannheim, Germany). LPS from Salmonella typhimurium, PMA, N-acetylcysteine (NAC), glutathione (GSH), cytochrome c, and superoxide dismutase (SOD) were purchased from Sigma (Sigma Chemical Co., St. Louis, MO). PD98059, SB203580, and MG 132 were purchased from BioMol (Plymouth Meeting, PA). The mouse monoclonal antibodies (mAbs) A77, A59, and A62 specific for FcR were prepared as described previously (24). Polyclonal rabbit anti-ERK1/2 MAP Kinases, anti-phospho ERK1/2 (pERK1/2) MAP Kinases, anti-p38 and anti-phospho p38 MAP kinases, anti-IB and anti-phospho IB Abs were purchased from New England Biolabs (Beverly, MA). The mouse anti-human CD14 mAb conjugated to fluorescein isothiocyanate (FITC) was purchased from Becton Dickinson (San Jose, CA), the swine anti-rabbit (SAR) IgG conjugated to horseradish peroxidase (HRP) was purchased from Dako (Glostrup, Denmark) and the F(ab')₂ sheep anti-mouse (SAM) IgG conjugated to FITC was from Sigma. Other materials were reagent grade.

Isolation of HAM

Bronchoalveolar lavages were obtained from normal nonsmoking volunteers as described previously (16) and approved by the local ethics committee. The lavage fluid was passed through a layer of sterile gauze to remove gross mucus and then centrifuged at 500 rpm for 10 min at 4°C to separate cells from fluid. The cell pellet was resuspended and washed twice in complete culture medium: i.e., RPMI-1640 medium (Bio-Whittaker, Walkersville, MD) supplemented with 10% decomplemented (56°C, 30 min) fetal calf serum (FCS), 2 mM L-glutamine, 100 U/ml penicillin, and 100 µg/ml streptomycin. HAM were > 95% pure and contained < 1% of neutrophils and monocytes; the remaining cells were lymphocytes as determined by morphologic characteristics. HAM were cultured in complete culture medium and they were stimulated either with LPS (1 µg/ml) or PMA (0.1 µg/ml). The pharmacologic inhibitor of MEK/ERK pathways, PD98059, added 1 h before stimulation of HAM, was used at 50 µM. This concentration inhibited all detectable pERK1/2 MAP Kinases in Western blots and was not toxic to HAM as determined by trypan blue staining.

Purification of IgA

Pure polymers of monoclonal IgA1 were isolated from myeloma sera containing high levels of monoclonal IgA with a large proportion of p-IgA as described by Vaerman and colleagues (25). Polyclonal milk S-IgA was prepared as described previously (26). Purity of IgA was checked by immunoelectrophoresis and/or Ouchterlony analysis using a battery of monospecific antisera against serum proteins as described previously (27). Size distribution of IgA preparations was assessed by sucrose density gradient ultracentrifugation and by gradient polyacrylamide gel electrophoresis (PAGE) with and without sodium dodecyl sulfate (SDS) (25). Polymeric IgA was conjugated to FITC as described previously (16) and the F/P ratio was 2.5.

Anti-IgA Abs Preparations and Fluorescent Conjugates

Rabbit anti-IgA isotype (iso)-specific and idiotype (Id)-specific IgG Abs were prepared as described by Sibille and coworkers (17). To prevent the Fc portion of the anti-IgA Abs from binding to FcR, F(ab')₂ fragments were prepared by pepsin digestion of rabbit IgG for 16-20 h at 37°C with an enzyme/substrate weight ratio of 1/50 in 0.1 M Na acetate buffer, pH 4.5. F(ab')₂ fragments were separated by Ultrogel AcA 44 and their purity was checked by their lack of precipitation with a goat antiserum specific to rabbit Fc in Ouchterlony immunodiffusion and by SDS-PAGE. F(ab')₂ were conjugated with FITC as described previously (16).

Flow Cytometric Analysis

Surface expression of FcR on HAM was assayed by indirect immunofluorescence. To mask FcR, cells (3 × 10⁵) were first incubated with aggregated human IgG (10 mg/ml) in flow cytometric analysis (FCA) buffer (Hanks' balanced salt solution [HBSS] with 3% FCS and 0.05% sodium azide) for 30 min at 4°C. Cells were then washed and thereafter incubated with mouse anti-FcR A59, A62, or A77 mAbs (0.1 mg/ml) for 1 h at 4°C. After four washes, cells were exposed to FITC-labeled F(ab')₂ of SAM IgG Ab (1%) for 1 h at 4°C. Cells were then washed and fixed in 2% (vol/vol) formaldehyde in HBSS. Cell-associated fluorescence was analyzed by flow cytometry using a FACScan (Becton Dickinson, Mountainview, CA). In parallel, autofluorescence and nonspecific binding of the secondary Ab were assessed. Surface expression of CD14 LPS receptors on HAM was assessed by direct immunofluorescence using an anti-CD14 IgG mAb conjugated with FITC.

For IgA binding, assays were performed by incubating cells (3 × 10⁵) at 4°C for 1 h with p- or S-IgA at 1.5 mg/ml in FCA buffer. Cells were then washed and incubated at 4°C for 1 h with FITC-labeled F(ab')₂ of rabbit anti (id)-IgA (10 µg/ml) to reveal idiotypic p-IgA, or with FITC-labeled F(ab')₂ of rabbit anti(iso)-IgA (10 µg/ml) for detecting S-IgA. Cells were thereafter washed and cell-associated fluorescence analyzed by FACScan.

Confocal Microscopy

The internalization of surface-bound IgA was investigated by confocal microscopy. HAM seeded on coverslips were incubated on ice with FITC-labeled p-IgA or with S-IgA at the indicated concentrations in complete culture medium for 1 h in the presence of 10 µg/ml of human transferrin conjugated to Texas-Red (Molecular Probes, Eugene, OR). Transferrin was used to assess intracellular colocalization of IgA/FcR complexes within the transferrin receptor recycling endosomes. Cells were then washed with cold culture medium before incubation at 37°C, 5% CO₂ for different time intervals. At each time, cells were washed and fixed with 3.7% (vol/vol) formaldehyde. For the intracellular staining of S-IgA, cells were permeabilized for 10 min with 0.2% Triton X-100 in phosphate-buffered saline (PBS) and incubated for 1 h with 10 µg/ml of FITC-labeled F(ab')₂ anti(iso)-IgA. Coverslips were mounted in 2.5% 1,4-diacylbicyclo-(2,2,2)octane (DABCO; Sigma) in Mowiol (Calbiochem-Novabiochem International Inc, Darmstadt, Germany). Observations were made under oil immersion with a ×63 objective with an MRC1024 (Bio-Rad Laboratories, Richmond, CA) confocal microscope. Images were digitally recorded with a Focus Graphics Image Recorder and used for direct computer-assisted reproduction with an ink-jet photo printer.

Measurements of Respiratory Burst

Fluorometric assay.The formation of ROI was assessed in LPS- or PMA-activated and nonactivated HAM by measuring the oxidation of dichlorofluorescein diacetate (DCFH; Sigma) to its fluorescent analog dichlorofluorescein (DCF) by fluorometry adapted for use in microplates as described by Wan and associates (28). Briefly, treated and untreated HAM (3 × 10⁵ cells/ well) were loaded with DCFH (15 µM) for 15 min at 37°C, 5% CO₂ in the dark. The DCF-specific fluorescence resulting from DCFH oxidation via the respiratory burst was assessed by fluorometry using a fluorimetric plate reader (Packard Instruments, Downers Grove, IL) with excitation and emission wavelengths of 485 nm and 538 nm, respectively. Results are expressed as nmol of DCF (calculated from a standard curve with defined amounts of fluorescent DCF) per mg of cell proteins. Total cell protein was assessed using a BCA protein assay reagent (Pierce, Rockford, IL.). The autofluorescence of HAM was so negligible and did not contribute to these calculations. To assess the effect of antioxidants or MAP kinase inhibitor, HAM were pretreated for 1 h with NAC, GSH, or PD98059, respectively, and then treated or not for 1 h with IgA and finally stimulated or not with LPS for 2 h or with PMA for 1 h.

Measurement of superoxide anion (O₂) generation.O₂ production by HAM was measured by the superoxide dismutase (SOD)- inhibitable reduction of cytochrome c assay as previously reported (29). HAM (1 × 10⁵/well) seeded in 100 µl of complete culture medium on 96-well flat-bottom tissue culture plates were used to measure O₂ production. Cells were loaded with 160 µM cytochrome c in HBSS and the optical density (OD) at 550 nm was measured with a microplate autoreader (Labsystems, Brussels, Belgium) at different times of incubation. Plates were incubated at 37°C, 5% CO₂ between OD measurements. Control reactions were conducted with 300 U/ml of SOD.

Western Blot Analysis of Total and Phosphorylated ERK1/2 MAP Kinases and IB

Treated and untreated HAM were washed with ice-cold PBS and then lysed in lysis buffer (150 mM NaCl, 20 mM Tris-HCl pH 7.4, 5 mM EDTA, 1% Nonidet P-40, 0.1% SDS, 0.5% sodium deoxycholate) supplemented with 1 mM phenyl methyl sulfonyl fluoride (PMSF), 10 µg/ml of each protease inhibitor (leupeptin, aprotinin, pepstatin, trypsin inhibitor), 1 mM Na₃VO₄, and 50 mM NaF. Lysates were cleared by centrifugation at 13,000 rpm for 15 min at 4°C and protein concentrations were determined using a BCA protein assay reagent (Pierce). Equal amounts of total extract proteins were resuspended in 2× Laemmli buffer under reducing conditions, boiled for 5 min, subjected to 12% SDS-PAGE, and then transferred to Hybond-nitrocellulose membranes (Amersham Life Sciences, Little Chalfont, UK). Blots were blocked with 5% bovine serum albumin (BSA) in Tris-buffered saline supplemented with Tween-20 (TBST: 50 mM Tris-HCl pH 7.5, 0.15 M NaCl, and 0.1% Tween-20) for 1 h. Membranes were then washed three times with TBST and probed overnight with specific anti- phospho-ERK1/2 rabbit Ab (1/1,000) for pERK1/2, anti-phospho-IB rabbit Ab (1/1,000) for pIB, respectively, or with anti-ERK1/2 rabbit Ab (1/1,000) for total ERK1/2 or anti-IB rabbit Ab (1/1,000) for total IB proteins, respectively, in TBST with 5% BSA. After three washes, membranes were incubated with HRP-conjugated SAR IgG secondary Ab (1/2,000 in TBST-5% BSA) for 1 h. Blots were washed and immunoreactive bands were visualized with the Supersignal Enhanced Chemiluminescence detection reagents (Pierce). Phosphorylated ERK2 protein and extracts from TNF--stimulated Hela cells were used as positive controls for ERK1/2 and IB revelation, respectively.

Phospho-Specific Antibody Cell-Based ELISA

Quantitation of pERK1/2 was performed by phospho-specific antibody cell-based ELISA (PACE) assay. HAM (1 × 10⁵/well) were seeded on 96-well flat-bottom tissue culture plates. Following stimulation, HAM were fixed with 8% formaldehyde in PBS for 20 min. Cells were washed with 0.1% Triton X-100 in PBS (PBS-Tx) and then incubated for 20 min in PBS-Tx with 0.1% azide and 1% H₂O₂ to quench endogenous peroxidases. After three washes with PBS-Tx, cells were blocked with 10% FCS in PBS-Tx for 1 h and then incubated with specific anti-phospho ERK1/2 rabbit Ab (1/250) overnight at 4°C. Cells were then washed three times (5 min each) and incubated with HRP-conjugated SAR IgG secondary Ab (1/500) for 1 h. After three washes, 50 µL of 0.04% o-phenylendiamine (OPD) and 0.015% H₂O₂ in OPD-buffer (35 mM citric acid in 66 mM Na₂HPO₄, pH 5.6) were added to the wells, and plates were incubated on a shaker at 200 rpm/min for 15 min in the dark. Reactions were stopped by addition of 25 µl/well of 1 M H₂SO₄ and OD measured on microplate reader at 492 nm. Results were standardized by cell number quantification using crystal violet as described by Scragg and colleagues (30) and OD reading at 595 nm. Results are expressed as arbitrary units (OD_{492 nm}/OD _{595 nm}).

Nuclear Extracts and Electrophoretic Mobility Shift Assay

Nuclear extracts were prepared from HAM as described by Carter and coworkers (31) with minor modifications. HAM were washed with cold PBS and then resuspended in 100 µl of ice-cold electrophoretic mobility shift assay (EMSA) lysis buffer (10 mM Hepes pH 7.9, 10 mM KCl, 2 mM MgCl₂, 2 mM EDTA, 1 mM DTT, 1 mM PMSF) supplemented with protease inhibitors and incubated on ice for 15 min. Nonidet P-40 (NP-40, 10%) was added to lyse the cells and cells were vortexed and centrifuged at 4°C at 13,000 rpm. Nuclei were resuspended in extraction buffer (50 mM Hepes pH 7.9, 10% glycerol, 50 mM KCl, 300 mM NaCl, 0.1 mM EDTA, 1 mM DTT, 0.5 mM PMSF and protease inhibitors) for 20 min on ice. The nuclear suspension was then centrifuged at 13,000 rpm and supernatant stored at 70°C.

EMSA was performed using the Promega (Madison, WI) gel shift assay system. NF-B consensus oligonucleotide (5'AGTTGA GGGGACTTTCCCAGGC3') was end-labeled with [-³²P]ATP (Amersham Pharmacia Biotech) using T4 polynucleotide kinase for 10 min at 37°C in a final volume of 10 µl. The reaction was stopped by addition of 1 µl of 0.5 M EDTA and labeled oligonucleotides were purified by chromatography through chromaspin columns (Clontech Laboratories, Inc., Palo Alto, CA). EMSA were performed at room temperature for 30 min with 5 µg of nuclear extract proteins and the ³²P-labeled oligonucleotides as described in the gel shift assay system (Promega E3300 kit). Binding specificity was conducted with specific competitor used at 100-fold excess of unlabeled NF-B consensus sequence oligonucleotides added 10 min before the addition of the labeled probe. The protein-DNA complexes were separated on 5% nondenaturing polyacrylamide gel in 0.5 × Tris-EDTA-borate. The gels were dried and exposed to autoradiographic films at -70°C.

TNF- Assay

TNF- produced by HAM was measured by its cytotoxicity against the highly sensitive WEHI 164 clone 13 (WEHI) fibrosarcoma cells as described previously (32) with minor modifications. WEHI cells (5 × 10⁴/well) in complete RPMI-1640 culture medium were plated in 96-well flat-bottom plates and incubated for 3 h at 37°C, 5% CO₂. WEHI cells then received 100 µl of serial dilutions of test supernatant samples and incubated for 24 h at 37°C, 5% CO₂. After 24 h, 20 µl of 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide (MTT) at 5 mg/ml in PBS were added to each well and plates were incubated for an additional 4 h at 37°C, 5% CO₂. Cells were then lysed (100 µl/well) with SDS/HCl solution (15% SDS in 0.015 M HCl) and the plates further incubated overnight to dissolve the blue formazan crystals and bleach the phenol red color of the culture medium. The amount of MTT taken up and reduced was detected by OD reading at 550 nm. Serial dilutions of known concentrations of rh-TNF- were used to quantify HAM secreted TNF-. Results are expressed as pg of TNF- per mg of cell proteins.

Statistical Analysis

Experiments were performed in triplicate and repeated at least three times. Results are expressed as means ± SD. When appropriate, the statistical significance of the differences observed between treated groups and controls or between pertinent groups was analyzed by the Student t test with a probability value of P < 0.05 considered to be significant.

	Results

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HAM Express FcR, Bind and Internalize p- and S-IgA

Indirect immunofluorescence analysis of FcR expression on HAM was performed by staining HAM cells with anti-FcR mAbs. Figure 1A shows representative flow cytometric profiles of HAM from normal nonsmoking volunteers stained with anti-FcR A59 or A62 mAbs. Moreover, and as reported previously (18), FcR was immunoprecipitated with A77 anti-FcR mAb from metabolically labeled HAM (data not shown). Immunofluorescence assay for IgA-binding to HAM carried with purified p- and S-IgA shows that HAM efficiently bound both p- (Figure 1B) and S-IgA (Figure 1C).

View larger version (10K): [in this window] [in a new window]   Figure 1.   Analysis of FcR expression on HAM by cytofluorometry (A) and immunofluorescence analysis of p- (B) and S-IgA (C ) binding to HAM. (A) Cells (3 × 105) on 24-well flat-bottom tissue culture plates, first incubated with aggregated human IgG (10 mg/ml) to mask FcR, were incubated with anti-FcR A59 or A62 mAbs (0.1 mg/ml) and then stained with FITC-labeled F(ab')2 SAM IgG as described in MATERIALS AND METHODS. (B and C) Cells (3 × 105) were incubated with p- (B) or S-IgA (C ) at 1.5 mg/ml for 1 h at 4°C and then with FITC-labeled F(ab')2 of rabbit anti-IgA (10 µg/ml) as described in MATERIALS AND METHODS. Profiles of cells autofluorescence and nonspecific binding of the secondary FITC-labeled Abs are indicated by thin and bold lines, respectively.

Endocytosis of HAM Surface-Bound IgA

Confocal microscopy shows that at 4°C, p- and S-IgA were distributed homogeneously on the cell surface of HAM (Figure 2A). Incubation at 37°C induced rapid internalization of p- and S-IgA, which, after 1 min, were already intracellularly localized, as shown by numerous intracellular vesicles containing IgA (Figure 2A). Colocalization experiments revealed that IgA was largely found within vesicles that stained positively for transferrin. However, and as shown in Figure 2A, p-IgA is preferentially perinuclear while S-IgA is widely distributed in the cytoplasm. These intracellular distributions were maintained from 5 to 30 min after incubation at 37°C, and IgA still clearly colocalized within the recycling transferrin receptor endosomes, as illustrated by the yellow fluorescent dots due to colocalization of the two markers. Concentrations as low as 0.25 mg/ml induced a significant internalization of both p- and S-IgA and only a slight increase of IgA-endocytosis was observed with concentrations ranging from 0.5 to 2 mg/ml (Figure 2B).

View larger version (80K): [in this window] [in a new window]   Figure 2.   Endocytosis of HAM surface-bound IgA. Cells (3 × 105) on coverslips were incubated for 1 h at 4°C with FITC-labeled p-IgA or with unlabeled S-IgA at 2 mg/ml (A) or with increasing concentrations of IgA ranged from 0.25 to 1.5 mg/ml (B) in the presence of transferrin conjugated to Texas-Red (5 µg/ml) to stain the recycling transferrin receptor endosomes. Cells were then incubated at 37°C for different time periods, washed and fixed with 3.7% (vol/vol) formaldehyde in PBS. Intracellular staining of S-IgA was carried out as described in MATERIALS AND METHODS. Cells were observed under a confocal microscope and photographs taken 1, 5, and 30 min (A) or 15 min (B) after incubation at 37°C. In this system, green fluorescence corresponds to IgA staining and red fluorescence to transferrin (Tr), whereas yellow fluorescence represents the superimposition of Texas-Red over the fluorescein (Tr/IgA). Scales are in micrometers.

LPS-Induced Increase of HAM Respiratory Burst Depends on Activated ERK1/2 MAP Kinases

HAM were first analyzed for the expression of CD14, the specific binding sites of LPS/LPS binding protein. Immunofluorescence analysis of HAM directly stained with FITC- labeled anti-CD14 mAb showed that CD14 were distributed homogeneously on the surface of HAM (Figure 3A). The CD14 basal expression on HAM was, however, lower than that on monocytes (data not shown), as also reported by Hopkins and associates (33). Quantitation of HAM oxidative burst revealed that HAM constitutively exhibited substantial respiratory burst, as demonstrated by intracellular oxidation of loaded DCFH (Figure 4A). Treatment of HAM with LPS induced significant enhancement of DCFH oxidation (Figure 4A). Kinetic studies (not shown) of the LPS-increased respiratory burst revealed that 2 h treatment of HAM with LPS were necessary to induce important enhancement of oxidative burst. The 2-h period of LPS activation was then respected in all experiments with LPS. Pretreatment of HAM with PD98059, a pharmacologic inhibitor of the MEK pathway (the upstream activator of ERK), completely inhibited the LPS-induced increase of DCFH oxidation (Figure 4A). These results indicate that activation of MEK/ERK MAP kinases pathway is of critical importance in the mechanism by which LPS enhances the respiratory burst in HAM.

View larger version (19K): [in this window] [in a new window]   Figure 3.   Immunofluorescence analysis of CD14 expression on HAM. HAM (3 × 105) on coverslips or on 24-well flat-bottom tissue culture plates were incubated for 1 h at 4°C with a mouse anti-human CD14 mAb conjugated to FITC. Cells were then washed, fixed, and either examined under a confocal microscope (A) or analyzed by flow cytometry using a FACScan for basal expression of CD14 (B), and its modulation by p- (green line), and S-IgA (blue line) (C ). Autofluorescence is indicated by the black line.

View larger version (29K): [in this window] [in a new window]   Figure 4.   Effect of p- and S-IgA on the LPS-enhanced oxidative burst. Role of activated ERK1/2. Respiratory burst was assessed by the measure of the DCFH oxidation (A, B, C ), and NADPH oxidase activity/O2 release using the cytochrome c reduction assay (D, E, F ) as described in MATERIALS AND METHODS. HAM were pretreated (A, C, D, F ) or not (B, E ) for 1 h with PD98059 (50 µM), then treated (B, C, E, F ) or not (A, D) with p- or S-IgA (1.5 mg/ml) or medium () for 1 h and then stimulated (with LPS, 1 µg/ml, striped bars) or not (no LPS, solid bars) for 2 h. PD98059 was present through the assay. Cells were thereafter loaded with DCFH (15 µM) for 15 min at 37°C, 5% CO2 (A, B, C ) or analyzed for the NADPH oxidase activity/O2 release (D, E, F ). Oxidation of DCFH to DCF was evaluated by the measure of the green fluorescence of cell suspensions using a fluorometer microplate reader, and cytochrome c reduction evaluated by OD measurement. Data are shown as means ± SD (n = 3). Statistical analysis was performed by the Student t test. §P < 0.005, P < 0.001 compared with control. The horizontal square brackets show the statistical significance of the differences used in our analysis of the data (P < 0.001).

Downregulation of LPS-Increased HAM Respiratory Burst by p- and S-IgA: Role of Activated ERK1/2

Both p- and S-IgA, which had no significant effect on the spontaneous respiratory burst of HAM (Figure 4B), downregulated the increase of respiratory burst induced by LPS (Figure 4B). This effect was similar to that obtained with PD98059 treatment of HAM (Figures 4A and 4C), suggesting that p- and S-IgA could modulate the LPS-enhanced respiratory burst through the activated ERK1/2 MAP kinases.

The release of O₂ was also investigated to better evaluate effects of IgA on the respiratory burst of HAM. Activation by LPS resulted in  2.5-fold greater release of O₂ as compared with unstimulated HAM (Figure 4D). Treatment with PD98059 completely abolished this stimulatory effect of LPS (Figure 4D). Preincubation of HAM with p- or S-IgA significantly inhibited the LPS-increased O₂ release and also diminished its basal release (Figure 4E). PD98059, together with p- or S-IgA, had pronounced inhibitory effects on the LPS-induced increase of O₂ release (Figure 4F), suggesting that PD98059 might have at least an additive effect with p- and S-IgA. These data again support the idea that activated ERK1/2 play an important role in both LPS-increased and IgA-decreased HAM respiratory burst. These effects of p- and S-IgA on LPS-activated respiratory burst of HAM were not related to a downregulation of their CD14 surface expression by IgA (Figures 3B and 3C).

Downregulation of LPS-Induced ERK1/2 Activation in HAM by p- and S-IgA

To further demonstrate the role of activated ERK1/2 in the LPS-induced increase of HAM respiratory burst and its downregulation by p- and S-IgA, we analyzed the activated (dually tyrosine/threonine phosphorylated) ERK1/2 by Western blotting and PACE assay using a phospho-specific ERK1/2 Ab. In comparison with untreated HAM, treatment of HAM with LPS led to activation of ERK1/2 (Figures 5A, 5B, and 5D). HAM were stimulated with LPS for 30, 60, and 120 min to determine the kinetic of ERK1/2 phosphorylation. Figure 5A shows that high levels of ERK1/2 phosphorylation were observed at 30 min, and although it decreased after HAM stimulation longer than 30 min, there was still significant ERK1/2 phosphorylation over an extended time course and as late as 120 min. Total ERK1/2 proteins were not modified by LPS (data not shown). PD98059, an inhibitor of ERK1/2 phosphorylation, was used to evaluate the role of ERK1/2 pathway in both LPS-induced oxidative burst and TNF- release in HAM. As shown in Figure 5A, PD98059 inhibited significantly and time-dependently the phosphorylation of ERK1/2. Moreover, pretreatment of HAM with PD98059 resulted in a dose-dependent suppression of LPS-induced ERK1/2 phosphorylation (Figure 5B) and complete inhibition was observed with PD98059 at 50 µM (Figures 5A and 5J). We also checked the specificity of the inhibitory effect of PD98059 on ERK1/2 MAP kinases. As shown in Figure 5K, PD98059 had no effect on LPS-induced p38 MAP kinase phosphorylation nor on total p38 MAP kinase (Figure 5L). SB203580, a specific inhibitor of p38 MAP kinase pathway, had no effect on ERK1/2 phosphorylation (Figure 5B). These results, as reported by Carter and colleagues (21), confirmed the effectiveness and the specificity of PD98059 inhibitor. LPS- induced activation of ERK1/2 was quantitated by PACE which showed a 3.5-fold increase as compared with untreated HAM (Figure 5I). To evaluate a potential role of IgA on LPS-induced oxidative burst through ERK1/2 pathway modulation, the effect of p- and S-IgA on LPS-activated ERK1/2 in HAM was investigated. In untreated HAM, p- and S-IgA did not change the basal level of ERK1/2 activation (Figure 5C). In contrast, p- and S-IgA inhibited the LPS-induced activation of ERK1/2 (Figures 5D and 5I). The inhibitory effects observed with p- and S-IgA (Figure 5I) were similar to those obtained with PD98059 (Figure 5J). S- and p-IgA drastically inhibited the LPS-activated ERK1/2 as quantitated by the sensitive PACE assay and no phosphorylated forms were detected by Western blot (Figure 5D). Moreover, kinetic studies showed that p- and S-IgA downregulated the LPS-induced ERK1/2 phosphorylation as early as 30 min and as late as 120 min after LPS stimulation (data not shown). Total ERK levels were not affected by the LPS, IgA, or PD98059 treatments (Figures 5E-5H). As stated above, LPS activation of ERK1/2 occurred rapidly, whereas important enhancement of LPS-induced ROI production was delayed to 2 h, suggesting therefore that ERK1/2 phosphorylation is independent of oxidative burst. Our results (Figure 5M) indeed confirmed that antioxidant treatment of HAM did not alter the LPS- induced ERK1/2 activation.

View larger version (36K): [in this window] [in a new window]   Figure 5.   Downregulation of LPS-induced ERK1/2 activation in HAM by p- and S-IgA. HAM (3 × 105/well) were pretreated or not for 1 h with PD98059 (PD) at the indicated concentrations (B) or with SB203580 (SB, 10 µM), or with PD98059 at 50 µM (A), and then activated or not with LPS (1 µg/ml) for 1 h. (C-H ) HAM (3 × 105/ well) pretreated (G, H ) or not (C-F ) with PD98059 (50 µM), were treated with p- or S-IgA (1.5 mg/ml) or medium () for 1 h and then activated (striped bars) or not (solid bars) with LPS (1 µg/ml) for 2 h. Cells were then harvested and lysed, and active pERK1/2 or total ERK1/2 in equivalent amounts of total lysate proteins were analyzed by Western blot using a phospho-ERK1/2 specific Ab (A-D) or total ERK1/2 (E-H ) specific Ab, respectively. (I, J ) HAM (1 × 105/well) on 96-well flat-bottom tissue culture plates were treated (J ) or not (I ) with PD98059 (50 µM) for 1 h, treated or not with p- or S-IgA (1.5 mg/ml) or medium (), and then activated or not with LPS (1 µg/ml) for 2 h. Cells were then analyzed for the expression of activated forms of ERK1/2 (pERK1/2) by PACE assay as described in MATERIALS AND METHODS. Data are means ± SD (n = 3). (K, L) HAM were pretreated or not for 1 h with PD98059 (50 µM), activated or not for 1 h with LPS (1 µg/ml) and then analyzed by Western blot for p-p38 MAP kinase (K ) or total p38 MAP kinase (L). (M ) HAM were pretreated or not with NAC (10 mM) for 1 h, then activated or not with LPS (1 µg/ml), and cells were analyzed for the expression of activated ERK1/2 (pERK1/2) by phospho-Western blots.

Upregulation of TNF- Production in HAM by p- and S-IgA is Independent of ERK1/2 Activation

HAM released a significant amount of TNF- which was significantly enhanced upon LPS stimulation (Figure 6A). Pretreatment of HAM with PD98059 inhibited both LPS-activated and basal TNF- release to the same extent (Figure 6A, and inset) showing that activated ERK1/2 are required for basal and LPS-stimulated TNF- production. Moreover, PD98059 inhibited dose-dependently the LPS-induced TNF- production in HAM (Figure 6B). Addition of p- or S-IgA to resting HAM stimulated TNF- production (Figure 6A), and this effect was maintained in the presence of PD98059. Moreover, IgA-increased TNF- production was also observed in LPS-stimulated HAM, although IgA inhibited ERK1/2 activation by LPS (Figure 5D). Thus, IgA-stimulated TNF- release was independent of ERK1/2 activation.

View larger version (19K): [in this window] [in a new window]   Figure 6.   Modulation of TNF- release by p- and S-IgA is independent of ERK1/2 activation pathway in HAM. (A) HAM pretreated or not for 1 h with PD98059 (50 µM) were treated or not for 1 h with p- or S-IgA (1.5 mg/ml) or medium () and then activated or not with LPS (1 µg/ml) for 2 h. Supernatants were then collected and assayed for TNF- using the cytotoxicity bioassay on WEHI target cells as described in MATERIALS AND METHODS. Fold increase is defined as the ratio of the TNF- produced in treated cells to that in controls (without and with PD98059 [PD] respectively). Inset, representative effect of LPS-increased production of TNF- (pg/mg protein) by HAM. (B) HAM were pretreated or not for 1 h with increasing concentrations of PD98059, then stimulated (striped bars) or not (solid bars) with LPS (1 µg/ml) for 2 h and supernatants assayed for TNF- production. Data are means ± SD (n = 3). Statistical analysis was performed using Student t test. ¶P < 0.05, §P < 0.005, P < 0.001 compared with the corresponding control ().

NF-B Activation by LPS in HAM: Role of Oxidants

NF-B is a crucial transcription factor that regulates the expression of TNF- in macrophages. NF-B activation allows it to translocate from the cytosol into the nucleus and this occurs following signal-induced phosphorylation and subsequent degradation of its inhibitors from the IB family proteins including IB (34). Western immunoblots of cytoplasmic extracts from control and LPS-treated HAM were performed to determine changes in steady state of both phosphorylated and total IB proteins. In unstimulated HAM, no significant phosphorylation of IB could be detected (Figure 7A) and the level of total IB was higher in comparison with LPS-activated HAM (Figure 7B), indicating that NF-B is maintained under inactivated state in control cells. In contrast, LPS induced a rapid phosphorylation (Figure 7A) and subsequent degradation of IB proteins from 5 to 30 min (Figure 7B). This indicates that LPS activated NF-B in a conventional pathway requiring IB proteolysis. At 60 min after activation with LPS, IB was newly present in the cytosol and phosphorylated (Figure 7A), suggesting newly synthesized IB, which indeed is thought to be an NF-B-driven process. The phosphorylation of the newly synthesized IB decreased again at 120 min, demonstrating continuous activation of NF-B by LPS. EMSA studies were performed to assess the NF-B binding activity in HAM, because phosphorylation and subsequent degradation of IB did not always correlate with NF-B nuclear translocation. Figure 8 is a representative gel showing NF-B binding activity in HAM stimulated with LPS. This stimulation is effectively and significantly inhibited by the proteasome degradation inhibitor MG 132 (Figure 9), suggesting that NF-B activation of HAM by LPS is highly dependent upon IB degradation. MG 132 had no direct effect on NF-B binding activity in unstimulated HAM (Figure 10A, lane 1), and competition analysis performed with 100-fold excess of unlabeled NF-B consensus sequence oligonucleotide (Figure 10A, lane 2) confirmed the specificity of the retarded complexes observed in EMSA.

View larger version (22K): [in this window] [in a new window]   Figure 7.   Time course of changes in cellular levels of the inhibitory protein IB in HAM stimulated with LPS and effect of antioxidants. (A) Western immunoblots of pIB from cytosolic extract proteins of HAM pretreated or not for 1 h with NAC or GSH at 10 mM and then stimulated with LPS (1 µg/ml) for the indicated times. (B) Western immunoblots of total IB from HAM treated as in A.

View larger version (83K): [in this window] [in a new window]   Figure 8.   EMSA analysis of NF-B binding activity in nuclear extracts from HAM. EMSA demonstrating NF-B binding activity in HAM treated with LPS (1 µg/ml), p- or S-IgA (1.5 mg/ml), but not with PMA (0.1 µg/ml) for 2 h.

View larger version (57K): [in this window] [in a new window]   Figure 9.   Inhibition of NF-B binding activity in HAM by the proteasome inhibitor MG 132. MG 132 (20 µM) was added for 1 h before treatment of HAM with LPS (1 µg/ml), p- or S-IgA (1.5 mg/ml) for 2 h.

View larger version (53K): [in this window] [in a new window]   Figure 10.   (A) Specificity of EMSA analysis of NF-B. Binding activity in LPS-activated HAM was blocked by adding excess of unlabeled NF-B (NF-B*) oligonucleotide, showing specificity (lane 2). MG 132 had no effect on NF-B in unstimulated HAM (lane 1). (B) Effect of antioxidant NAC on the NF-B binding activity in HAM. HAM were treated with NAC (10 mM) for 1 h and then treated for 2 h with LPS (1 µg/ml), p- or S-IgA (1.5 mg/ml).

ROI have been proposed as activators of NF-B (35). To test a potential implication of ROI in LPS-induced NF-B activation, HAM were first pretreated with either NAC or GSH antioxidants at a concentration that suppressed the oxidative burst (Table 1) and subsequently activated with LPS. Figures 7A and 7B show that NAC had no effect on the kinetic of dual phosphorylation/degradation of IB from 5 to 30 min. However, NAC inhibited the newly synthesized IB at 60 and 120 min in comparison with LPS alone (Figure 7B). The effect of NAC is in accordance with the fact that ROI production by LPS-stimulated HAM is delayed to 60 min and increased at 120 min, whereas IB phosphorylation occurred as early as 5 min. Thus, ROI could play a minor role (but at late stages) in NF-B activation by LPS in HAM. In addition, EMSA studies showed that NAC had no effect on LPS-induced NFB binding activity (Figure 10B).

Role of NF-B and Oxidants in LPS-Induced Production of TNF- in HAM

The role of NF-B in LPS-induced TNF- release in HAM was investigated using the proteasome inhibitor MG 132, which inhibits NF-B activation by preventing IB degradation. As shown in Figure 11A, preincubation of HAM for 1 h with 20 µM of MG 132 significantly inhibited TNF- release in LPS-activated HAM. The effect of MG 132 was not generalized to all protein synthesis because total proteins from cell lysates were not affected by MG 132. The effect of MG 132 was specifically related to alteration of NF-B activation and of NF-B nuclear translocation, as illustrated in Figure 9. Moreover, MG 132 reversed the effects of LPS on IB degradation without altering IB phosphorylation (Figures 11C and 11D) and EMSA studies demonstrated that MG 132 blocked nuclear translocation of NF-B. These results also support the idea that MG 132 works specifically at the level of proteasome.

View larger version (22K): [in this window] [in a new window]   Figure 11.   Role of NF-B and oxidants in LPS-induced TNF- production in HAM. HAM were treated (striped bars) or not (solid bars) for 1 h with MG 132 (20 µM) (A) or NAC (10 mM) (B) and then activated or not with LPS (1 µg/ml) for 2 h. Supernatants were assayed for TNF- production as described in MATERIALS AND METHODS. Data are means ± SD (n = 3). Statistical analysis was performed using Student t test. P < 0.001 compared with the corresponding control (). (C, D) Inhibition of IB proteolysis by the proteasome inhibitor MG 132. HAM were pretreated or not for 1 h with MG 132 (20 µM) and then activated for 30 and 60 min with LPS (1 µg/ml) and cytosolic extracted proteins assayed for pIB (C ) and total IB (D) proteins.

We next investigated the role of ROI in LPS-induced TNF- release as LPS stimulated oxidative burst in HAM (Figures 4A and 4B). Results presented in Figure 11B show that NAC reduced by ~ 40% the LPS-induced production of TNF-. These results are in accordance with the late effects of NAC on NF-B activation (Figure 7) and with the delayed production of ROI by LPS.

Effects of IgA on NF-B and Its Role in IgA-Mediated TNF- Production in HAM

Western immunoblots of cytosolic extracts from p- and S-IgA-treated HAM show that IgA induced phosphorylation of IB with different kinetic from that observed with LPS. Substantial phosphorylation of IB was observed with IgA from 5 to 30 min followed by its depletion but as late as at 120 min (Figures 12A and 12B). NF-B nuclear translocation was also induced by treatment of HAM either with p- or S-IgA (Figures 8 and 9), and the NF-B binding activity was increased as much as observed with LPS activation. The effects of IgA on IB phosphorylation and NF-B nuclear translocation were not affected by the presence of NAC (Figures 12 and 10B, respectively). IgA, indeed, has no effect on the oxidative burst in unstimulated HAM. Both p- and S-IgA-induced NF-B binding activity was reversed by MG 132 (Figure 9). NF-B activation by IgA could explain the IgA-induced TNF- production in HAM. Pretreatment of HAM with MG 132 for 1 h before the addition of IgA reduced but did not completely inhibit IgA-increased TNF- production (Figure 13A). In contrast, NAC treatment of HAM had no effect on TNF- production by IgA-treated HAM (Figure 13B). This suggests that IgA-induced production of TNF- is partly NF-B- dependent but oxidant-independent.

View larger version (32K): [in this window] [in a new window]   Figure 12.   IgA-mediated phosphorylation of IB is not affected by antioxidant treatment of HAM. HAM were pretreated or not for 1 h with NAC (10 mM) and then treated with p- (A) or S-IgA (B) (1.5 mg/ml) for the indicated times. Cytosolic extracts were assayed for pIB by Western immunoblots.

View larger version (16K): [in this window] [in a new window]   Figure 13.   Role of NF-B and oxidants in IgA-mediated TNF- production in HAM. HAM were pretreated (striped bars) or not (solid bars) for 1 h with MG 132 (20 µM, A) or NAC (10 mM, B), then treated with IgA (1.5 mg/ml) or medium () for 2 h and supernatants were assayed for TNF- production as described in MATERIALS AND METHODS. Data are means ± SD (n = 3). Statistical analysis was performed using Student t test. §P < 0.005 compared with the corresponding control ().

Upregulation of PMA-Induced Increase of HAM O₂ Release by p- and S-IgA: Role of Activated ERK1/2

The role of ERK1/2 in the regulation by p- and S-IgA of receptor-independent activated HAM respiratory burst was investigated using PMA as stimulus. PMA-stimulated HAM released 2-fold more O₂ than unstimulated HAM (Figure 14A). PD98059 did not much alter the basal production of O₂ but completely inhibited the effect of PMA (Figure 14A). Treatment of HAM with p- or S-IgA led to a synergistic increase of O₂ release with PMA (Figure 14B) which was completely abolished by PD98059 (Figure 14B). Thus, ERK1/2 pathway also plays an important role in receptor-independent stimulated oxidative burst in HAM. Similar results were obtained when the respiratory burst was assayed through DCFH oxidation (data not shown).

View larger version (19K): [in this window] [in a new window]   Figure 14.   Effect of p- and S-IgA on the PMA-induced increase of O2 release by HAM. Role of activated ERK1/2. HAM (1 × 105) on 96-well flat-bottom tissue culture plates were pretreated or not for 1 h with PD98059 (50 µM) and then incubated (B) or not (A) with p- or S-IgA (1.5 mg/ml) or medium () for 1 h. Cells were therefore activated (with PMA, 0.1 µg/ml, striped bars) or not (no PMA, solid bars) for 1 h and then analyzed for the NADPH oxidase activity/O2 release using the cytochrome c reduction assay as described in MATERIALS AND METHODS. Data are shown as means ± SD (n = 3). Statistical analysis was performed using Student t test. §P < 0.005, P < 0.001 compared with the corresponding control ().

Upregulation of PMA-Induced ERK1/2 Activation in HAM by p- and S-IgA

HAM stimulated with PMA showed an increased amount of phosphorylated ERK1/2 in comparison with unstimulated cells as shown by phospho-Western blots (Figures 15A and 15B) and by PACE assay (Figure 15F). The PMA-induced ERK1/2 phosphorylation was detected as early as 2 min and as late as 2 h (data not shown). This effect, which is time dependent, was maximal at 30 min, but demonstrable ERK1/2 activation could still be observed as late as 60 min (Figure 15A). Preincubation of HAM with PD98059 blocked PMA-induced ERK1/2 phosphorylation, and this inhibitory effect, which is dose-dependent (Figure 15B), was maximal at 50 µM of PD98059 (Figures 15A and 15G). As shown for LPS-activated HAM, PD98059 selectively inhibited PMA-induced ERK1/2 phosphorylation without affecting that of p38 MAP kinase (Figures 15H and 15I). In PMA-activated HAM, both p- and S-IgA increased the phosphorylation of ERK1/2 (Figures 15C and 15F), and this IgA-induced upregulation of ERK1/2 was observed as early as 30 min and as late as 60 min after PMA stimulation (data not shown). These effects were also inhibited by PD98059 (Figure 15G). No effect either of PMA, IgA, or PD98059 on total ERK1/2 expression was observed (Figures 15D and 15E). In addition, ERK1/2 phosphorylation in PMA-activated HAM was independent of ROI production, as neither NAC nor GSH antioxidants altered this effect of PMA (Figure 15J).

View larger version (35K): [in this window] [in a new window]   Figure 15.   Upregulation of PMA-induced ERK1/2 activation in HAM by p- and S-IgA. (A, B) HAM (3 × 105/well) were pretreated or not for 1 h with PD98059 at 50 µM (A) or at the indicated concentrations (B) and then stimulated or not with PMA (0.1 µg/ml) for 1 h. (C-E ) HAM (3 × 105/well) on 24-well flat-bottom tissue culture plates were pretreated (E ) or not (C, D) with PD98059 (50 µM) for 1 h and treated or not with p- or S-IgA (1.5 mg/ml) or medium () and then activated with PMA (0.1 µg/ml) for 1 h. Cells were then harvested, lysed, and active pERK1/2 or total ERK1/2 in equivalent amounts of total lysate proteins were analyzed by Western blot using a phospho-ERK1/2 specific Ab (A, B, C ) or total ERK1/2 specific Ab (D, E ), respectively. The (p-ERK2)* phosphorylated in vitro by MEK1/2 kinase is used as positive control for the revealed ERK1/2 MAP kinases. (F, G ) HAM (1 × 105/well) on 96-well flat-bottom tissue culture plates were pretreated (G ) or not (F ) with PD98059 (50 µM) for 1 h, then treated or not with p- or S-IgA (1.5 mg/ml) or medium () for 1 h and then activated (striped bars) or not (solid bars) with PMA (0.1 µg/ml) for 1 h. Cells were then analyzed for the expression of activated forms of ERK1/2 (pERK1/2) by PACE assay. Data are means ± SD (n = 3). (H, I ) HAM were pretreated or not for 1 h with PD98059 (50 µM), activated or not for 1 h with PMA (0.1 µg/ml), and then analyzed by Western blot for p-p38 MAP kinase (H ) or total p38 MAP kinase (I ). (J ) HAM were pretreated or not with NAC or GSH (10 mM) for 1 h, then activated or not with PMA (0.1 µg/ml), and cells were analyzed for the expression of activated ERK1/2 (pERK1/2) by phospho-Western blots.

IgA Modulates TNF- Production in Both Unstimulated and PMA-Stimulated HAM Independently of ERK1/2 Activation

HAM stimulated with PMA released more TNF- than unstimulated HAM (Figure 16A and inset). Basal and PMA-stimulated productions of TNF- depended largely on the ERK1/2 pathway, both being inhibited by PD98059 (Figure 16A and inset). Moreover, inhibition of PMA-induced TNF- production was PD98059 dose-dependent (Figure 16B). The effect of PMA, however, did not much add to that of p- and S-IgA, both of which stimulated TNF- release in resting HAM (Figure 16A). In contrast to PMA, IgA-mediated increase of TNF- production was largely independent of the ERK1/2 pathway (Figure 16A). These results suggest that there is no complete correlation between ERK1/2 activation and TNF- release and that an alternative ERK-independent pathway exists for IgA-stimulated TNF- release in HAM.

View larger version (20K): [in this window] [in a new window]   Figure 16.   Modulation of HAM release of TNF- by p- and S-IgA is independent of ERK1/2 activation pathway. (A) HAM on 24-well flat-bottom tissue culture plates were pretreated or not with PD98059 (50 µM) for 1 h, treated with p- or S-IgA (1.5 mg/ ml) or medium () for 1 h, and then activated (striped bars) or not (solid bars) with PMA (0.1 µg/ml) for 1 h. Supernatants were then collected and assayed for TNF- using the cytotoxicity bioassay on WEHI target cells as described in MATERIALS AND METHODS. Fold increase is defined as the ratio of the TNF- in treated cells to that in controls (without and with PD98059 [PD], respectively). Inset, representative effect of PMA-increased production of TNF- (pg/mg protein) by HAM. (B) HAM were pretreated or not for 1 h with increasing concentrations of PD98059 and then activated or not with PMA (0.1 µg/ml) for 1 h before measuring TNF- production in supernatants. Data are means ± SD (n = 3). Statistical analysis was performed using Student t test. ¶P < 0.05, §P < 0.005, P < 0.001 compared with the corresponding control ().

Role of NF-B and Oxidants in PMA-Induced TNF- Production in HAM

In contrast to LPS, treatment of HAM with PMA resulted in very low phosphorylation of IB from 5 to 30 min and substantial elevation at 60 min (Figure 17A). However, and surprisingly, the level of total IB increased during PMA stimulation up to 30 min. At 60 min, a low depletion of IB was observed with a level remaining comparable to that in unstimulated HAM (Figure 17B). EMSA studies showed no nuclear translocation of NF-B in PMA-stimulated HAM (Figures 8 and 10A). Antioxidants, NAC, and GSH had no effect on IB phosphorylation/degradation in PMA-activated HAM (Figures 17A and 17B). In unstimulated HAM, the antioxidant GSH did not show any significant effect on IB proteins (Figure 17). Thus, PMA failed to activate NF-B despite the small increase in phosphorylation of IB. PMA-induced TNF- release was not only independent of NF-B, as no nuclear translocation of NF-B occurred subsequent to PMA activation and no effect of MG 132 on this production was seen (data not shown), but also was not affected by the presence of NAC (Figure 17C).

View larger version (29K): [in this window] [in a new window]   Figure 17.   Time course of changes in cellular levels of the inhibitory protein IB in HAM stimulated with PMA and effect of antioxidants. (A) Western immunoblots of pIB from cytosolic extract proteins of HAM pretreated or not for 1 h with NAC or GSH at 10 mM and then stimulated or not with PMA (0.1 µg/ml) for the indicated times. (B) Western immunoblots of total IB from HAM treated as in A. The (IB)* from TNF--stimulated Hela cells is used as positive control of the revealed IB proteins. (C) Role of oxidants in PMA-mediated TNF- production in HAM. HAM were pretreated (striped bars) or not (solid bars) for 1 h with NAC (10 mM), then treated or not with PMA (0.1 µg/ml) for 1 h, and supernatants were assayed for TNF- production as described in MATERIALS AND METHODS.

	Discussion

Top Abstract Introduction Materials and Methods Results Discussion References

HAM represent the major cellular line of defense against invading microorganisms and inhaled antigens in the lung (19). Binding sites for IgA were described on HAM (16) and more recently, the molecular nature of FcR in HAM has been characterized and revealed that HAM express an FcR derived from an alternative splicing of the primary FcR transcript and therefore different from that of neutrophils and monocytes (18). Much progress has been made regarding the biology of FcR on monocytes (5, 7, 36), neutrophils (37), and eosinophils (38, 39). However, the biology and the functional aspects of FcR on HAM remained poorly investigated. Using HAM from healthy nonsmoking volunteers, we have studied the effects and the mechanisms by which interaction of their FcR with the predominant IgA forms in mucosal respiratory secretions, namely p- and S-IgA, could modulate LPS receptor- dependent and PMA receptor-independent stimulation of the respiratory burst and TNF- release. The IgA concentrations we used are likely to be in the range of those observed in respiratory secretions. Considering that the BAL procedures induce a dilution factor of roughly 100 (40), we and others (41, 42) have reported that in vivo IgA concentration in respiratory secretions is in the range of 1 mg/ml in normal nonsmoking subjects and can increase in disease states. After first confirming FcR expression on HAM, we investigated the IgA-mediated endocytosis and FcR-mediated signal transduction in HAM. We showed that both p- and S-IgA were efficiently and rapidly internalized by HAM even without crosslinking of IgA by secondary Abs as reported for the internalization of anti-FcR Abs (43). Intracellular vesicles containing p- and S-IgA were largely colocalized with those containing the recycling transferrin receptors, suggesting the existence of a recycling pathway for HAM FcR.

HAM stimulated either with LPS or PMA show an increased respiratory burst as measured by oxidation of DCFH and O₂ release. Using two methods, DCF (which measures the production of H₂O₂), and cytochrome c (which measures the release of O₂), we observed similar increased oxidative burst with stimuli and its decrease with inhibitors. Both LPS and PMA stimuli were reported to activate ERK1/2 kinases in HAM (44). We observed that the inhibition of ERK1/2 activity by its selective inhibitor PD98059 completely abolished both LPS- and PMA-stimulated respiratory burst in HAM, suggesting that activated ERK1/2 play an important role in the respiratory burst of HAM. A role of activated ERK1/2 in the stimulation of the respiratory burst by different proinflammatory stimuli has been documented for other phagocytic cells (20, 45- 47) but, to our knowledge, this is the first report that the ERK pathway controls the respiratory burst in HAM.

Incubation of HAM with p- or S-IgA downregulated their LPS-activated respiratory burst. This effect is consistent with our finding that p- and S-IgA, as well as PD98059, downregulated the LPS-induced ERK1/2 activation. These results for the first time demonstrate an effect of IgA on ERK1/2 kinases and suggest that the ERK1/2 pathway is critical in HAM for the modulation of the respiratory burst by both p- and S-IgA. This effect of p- and S-IgA was not related to a downregulation of CD14 expression on HAM by given forms of IgA.

PMA also increased the respiratory burst of HAM as previously reported (48). We now show in addition that this effect of PMA is upregulated by both p- and S-IgA, in contrast to the observed effects of IgA on LPS-activated respiratory burst. Other contrasting effects on HAM respiratory burst were reported, such as for IL-4 that inhibited the LPS-induced respiratory burst but increased that induced by IFN- (49). HAM stimulated with PMA showed increased amounts of activated ERK1/2 kinases. This effect was further upregulated by both p- and S-IgA and accounts for the enhanced oxidative burst observed in IgA-treated PMA-activated HAM. PD98059 fully inhibited the increased oxidative burst of HAM induced by both PMA and IgA. Therefore, activated ERK1/2 are also critical for the regulation of receptor independent stimulation of respiratory burst in HAM. Altogether, our data suggest that IgA is an important modulator of the respiratory burst in HAM via different pathways depending on the nature of the stimulus.

Our data further suggest that LPS and PMA stimulated ERK1/2 activity in HAM through different pathways as IgA differentially modulates LPS- and PMA-stimulated ERK kinases. The best described pathway of ERK activation is the Ras-Raf-1-MEK/ERK kinases cascade (50). However, Raf-1-independent pathways of ERK activation exist as demonstrated for LPS- (51), TNF-- (52), and insulin (53)- regulated ERK kinase pathways. In HAM, Monick and coworkers (44) reported that LPS regulates ERK activation independently of the Raf-1 pathway. These authors showed that LPS-induced ERK activation can occur either through the phospatidylcholine-phospholipase C-ceramide-PKC -MEK pathway or the phosphatidylinositol-3 kinase-PKC -MEK pathway (44, 54). The PKC  therefore plays a central role in the LPS-stimulation of ERK1/2 pathway in HAM. Monick and associates (44) further demonstrated that PMA, a strong activator of classical PKC, activates both Raf-1 and ERK pathways, supporting the idea that PMA activates ERK through the classical Raf-1-dependent pathway. Moreover, earlier data (55) showed that in HAM, PMA did not activate PKC , as demonstrated by its negligible membrane translocation, suggesting therefore that PMA could not activate ERK through PKC -dependent pathway. Thus, PMA and LPS activate ERK kinases in HAM through distinct pathways, and independently of ROI production. The contrasting effects of p- and S-IgA on the LPS- and PMA-induced ERK activation in HAM could therefore result from a differential effect of a given form of IgA on one or several components of the LPS- and PMA-used pathways. Additional studies are needed to examine how IgA could inhibit one or several kinases implicated in the LPS-induced ERK activation and, in parallel, activate the PMA-induced Raf-1-dependent ERK pathway.

To produce a respiratory burst, phagocytes must assemble in their plasma or phagolysosomal membranes the constituents of the NADPH oxidase complex. The mechanism through which activated ERK kinases increase the respiratory burst in PMA- and LPS-stimulated HAM could result from the phosphorylation by ERK of cytosolic and/or membrane bound components of the NADPH oxidase. It has been shown that ERK as well as p38 MAP kinases phosphorylate p47^phox, a cytosolic factor of the NADPH oxidase system (56).

In our study, both LPS- and PMA-stimulated TNF- release by HAM through the ERK pathway. Inhibition of ERK activity by PD98059 completely inhibited LPS- and PMA-increased TNF- release. Carter and colleagues (21) showed that ERK but also p38 MAP kinases were required for optimal LPS-induced TNF- gene expression and protein release in HAM. In our system, the effect of LPS and PMA on TNF- release were investigated after relatively short times of stimulation, 2 h and 1 h, respectively, whereas Carter and associates (21) observed the 24-h effect of LPS. We may therefore presume that ERK activation is sufficient to regulate LPS- and PMA-increased production of TNF- for short times, whereas both ERK and p38 MAP kinases would be required for a more persistent production of TNF-. Analysis of p38 MAP kinase in our system showed that p38 MAP kinase is highly activated by LPS and IgA, and to a low extent by PMA. Thus, IgA-induced TNF- could be mediated through p38 MAP kinase. Further studies are therefore required to determine either a cooperative or antagonistic effect between ERK and p38 MAP kinases.

The different effects of LPS and PMA on NF-B activation are another demonstration that these stimulators used different pathways to activate HAM.

Our data confirm that LPS induced NF-B activation in HAM as reported earlier (31). Now, we further demonstrate that NF-B activation by LPS is highly dependent upon classical pathway phosphorylation and subsequent degradation of IB. This effect was indeed reversed by the proteasome inhibitor MG 132. Production of TNF-, like that of other cytokines such as IL-6 and IL-8, is, at least in part, controlled at the level of their gene transcription. Our data show in addition that inhibition of NF-B activation via MG 132 results in altered TNF- release in LPS-stimulated HAM. Others have shown that the proteasome inhibitor MG 132 inhibits TNF- release by LPS- activated HAM and that this effect correlated with the blockade of NF-B activation (57). Thus, both ERK1/2 and NF-B play crucial roles in LPS-increased TNF- secretion in HAM. Several studies have shown that NF-B activation may be regulated by a variety of agents, including cytokines, mitogens, and oxygen radicals (58, 59). However, other studies (60, 61) have now questioned this concept and limited the effect of oxidants on NF-B to only certain cell types such as human breast (62) and T cell lines (63), but not monocytic cell lines (64), epidermal cells (60), or epithelial cell lines (61). Our results using NAC (a precursor of GSH) and GSH demonstrate that antioxidants efficiently inhibited ROI production as reported for other macrophages (65), and that ROI play a minor and delayed role in LPS-induced NF-B activation in HAM. This effect is, in addition, consistent with the partial inhibition of TNF- release by antioxidants. Reduced TNF- production by NAC and GSH in LPS-activated HAM was reported previously (66). Thus, the intracellular glutathione redox status plays a minor role in the control of TNF- production in HAM. Paradoxically, antioxidants such as NAC can increase NF-B activation, as observed in adenocarcinoma cell lines (67). In addition, antioxidants can modulate NF-B in a manner that is independent of their scavenging activity. Although the inhibitory effects of antioxidants on NF-B but also on other transcription or activated protein factors are often controversial and inconclusive, we can speculate that these effects are limited to certain cell types. Our data also showed that both p- and S-IgA activated nuclear translocation of NF-B in HAM independently of oxidants. Activation of NF-B by IgA contributed partly to the upregulation of TNF- by IgA, indicating that other transcription or activated protein factors and/or MAP kinases play an essential role in the TNF- control by IgA. Incubation of HAM with p- or S-IgA resulted in enhanced TNF- release. In monocytes, FcR crosslinking and p-IgA binding have been shown to stimulate TNF- production (13). In our system, treatment of HAM with PD98059 did not inhibit IgA-mediated TNF- release in both resting and PMA- or LPS-stimulated HAM. Moreover, IgA-induced TNF- release was mediated only partly by NF-B. These results therefore suggest the existence of alternative pathway(s) for IgA-induced TNF- release and that the TNF- promoter may contain MAP kinase-dependent and -independent regulatory elements as reported previously (68). Our finding that both p- and S-IgA activated strongly the phosphorylation of p38 MAP kinase may suggest that p38 MAP kinase plays a critical role in regulating TNF- release in HAM. The AP-1 site, which is present in TNF- promoter, plays an important role in the transcriptional regulation of TNF-. However, AP-1 activity was reported to be loss in HAM (69), which therefore suggests that AP-1 do not play an important role in regulating TNF-, but its activation by IgA could not be ruled out.

In contrast to LPS and IgA, PMA failed to activate NF-B nuclear translocation in HAM, and even slightly increased IB phosphorylation as late as 60 min after stimulation. Other studies have demonstrated the inability of PMA to activate NF-B in macrophages (70, 71), supporting the idea that NF-B activation is independent of PKC including in HAM (31).

It is important to outline that the mechanism by which IgA increased TNF- production did not add with the ERK-dependent LPS- and PMA-stimulated TNF- release.

TNF- produced by IgA-treated HAM was also oxidant-independent because neither p- nor S-IgA modulated the oxidative burst in unstimulated HAM. In addition, IgA downregulated the oxidative burst in LPS-activated HAM through inhibition of ERK1/2 and not by acting as radical scavengers like antioxidants do.

The ERK1/2 MAP kinase and NF-B signaling systems represent two distinct but interactive signal transduction pathways. LPS activates both ERK1/2 and NF-B pathways, which play an important role in TNF- secretion. However, the two pathways appear to contribute to TNF- production competitively rather than additively. MAP kinases may act directly on the NF-B transcription activation complex or through intermediate kinases. This will be of great interest to investigate to determine if ERK1/2 or other MAP kinases could modulate NF-B activation in HAM, especially through IB kinase which is the first step in NF-B activation. Preliminary data from our laboratory have demonstrated that inhibition of NF-B activation by MG 132 did not block LPS-activated ERK1/2 MAP kinases. Examples of NF-B regulation by MEK-ERK pathway have been described, such as in lung epithelial cells where ERK regulates negatively, but JNK positively, NF-B (72).

HAM play an important role in host defense against infectious pathogens. The release of toxic ROI and proinflammatory cytokines following activation with endotoxin or contact with bacteria is of critical importance in HAM-mediated protective immunity. By contrast, these mediators may also play an important role in the pathogenesis of inflammatory processes in the lung (73). Regulation of HAM is therefore of critical importance in controlling lung inflammation, as recurrent activation of HAM could be deleterious for the alveoli and the lower lung mucosal environment. Our results show a complex HAM regulation by IgA. In LPS- but not PMA-stimulated HAM, p- and S-IgA could act as anti-inflammatory modulators by downregulating the production of ROI. This effect of IgA could protect lung tissues from injury. ROI are indeed implicated in a number of pulmonary diseases such as the adult respiratory distress syndrome (ARDS) (74). In contrast, IgA could exacerbate the noxious effect of PMA-related stimuli. The interaction of both resting and activated HAM with p- and S-IgA leads to increased production of TNF-, which could be beneficial to host defense. TNF- plays a crucial role in the development of protective responses to bacterial and viral pathogens. However, continuous interactions of FcR with IgA could also lead to elevated levels of cytokines such as TNF- that have also been correlated with lung inflammatory injury as in patients with ARDS. Although p-IgA and in particular S-IgA account for the majority of IgA in mucosal secretions, substantial amounts of monomeric IgA can be detected (~ 28% of total IgA) (41) in BAL. We are currently investigating whether the size of IgA could influence the response of HAM. Thus, IgA could act as an important and balancing modulator in the HAM-mediated host protective pro- and anti-inflammatory responses.