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Published ahead of print on July 10, 2003, doi:10.1165/rcmb.2003-0188OC
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American Journal of Respiratory Cell and Molecular Biology. Vol. 30, pp. 333-341, 2004
© 2004 American Thoracic Society
DOI: 10.1165/rcmb.2003-0188OC

Effect of Eosinophil Adhesion on Intracellular Signaling in Cholinergic Nerve Cells

Marie-Thérèse Walsh, David R. Curran, Paul J. Kingham, Ross K. Morgan, Niamh Durcan, Gerald J. Gleich, W. Graham McLean and Richard W. Costello

Department of Medicine, RCSI, Beaumont Hospital, Dublin, Ireland; Department of Pharmacology and Therapeutics, University of Liverpool, Liverpool, United Kingdom; and Department of Dermatology, School of Medicine, University of Utah, Salt Lake City, Utah

Address correspondence to: Richard W. Costello, Department of Medicine, RCSI, Beaumont Hospital, Dublin 9, Ireland. E-mail: rcostello{at}rcsi.ie


   Abstract
Top
 Abstract
Introduction
 Materials and Methods
 Results
 Discussion
 References
 
Eosinophil localization to cholinergic nerves occurs in a variety of inflammatory conditions, including asthma. This localization is mediated by interactions between eosinophil integrins and neuronal vascular cell adhesion molecule-1 (VCAM-1) and intercellular adhesion molecule-1 (ICAM-1). Eosinophil–nerve cell interactions lead to generation of neuronal reactive oxygen species and release of eosinophil proteins. The effects of eosinophil adhesion on neuronal intracellular signaling pathways were investigated. Eosinophil adhesion to IMR32 cholinergic nerves led to a rapid and sustained activation of the nuclear transcription factors nuclear factor (NF)-{kappa}B and activator protein (AP)-1 in the nerve cells. Eosinophil binding to neuronal ICAM-1 led to a rapid activation of ERK1/2 in nerve cells. Inhibition of ERK1/2 prevented NF-{kappa}B activation. Eosinophil adhesion to VCAM-1 resulted in AP-1 activation, mediated partially by rapid activation of the p38 mitogen-activated protein kinase. These data show that adhesion of eosinophils induces mitogen-activated protein kinase–dependent activation of the transcription factors NF-{kappa}B and AP-1 in nerve cells, indicating that eosinophil adhesion may control nerve growth and phenotype.

Abbreviations: activator protein-1, AP-1 • electrophoretic mobility shift analysis, EMSA • extracellular signal–regulated protein kinase, ERK • intercellular adhesion molecule-1, ICAM-1 • mitogen-activated protein kinase(s), MAP kinases • nuclear factor-{kappa}B, NF-{kappa}B • vascular cell adhesion molecule-1, VCAM-1 • very late antigen-4, VLA-4


   Introduction
Top
 Abstract
 Introduction
Materials and Methods
 Results
 Discussion
 References
 
The presence of eosinophils and their degranulation products in tissues is the hallmark of a variety of diseases, including the allergic conditions asthma and rhinitis, as well as inflammatory bowel disease and eosinophilic gastroenteritis (1–4). Eosinophils localize to specific tissue structures under the coordinated influence of specific chemoattractants and through interactions with adhesion molecules at sites of inflammation (5). Eosinophil survival in these tissues is mediated in part by autocrine growth factor production. These growth factors are produced in response to ligation of integrin adhesion molecules, which leads to signaling within eosinophils, an effect termed "inside out signaling" (6, 7). Thus, tissue eosinophilia is mediated by interactions of eosinophils with adhesion molecules on cells, such as nerve cells, at inflammatory sites. However, the effects of this eosinophil adhesion on the cells to which they adhere are less well established.

In prior studies, we have shown that eosinophils specifically localize to cholinergic nerves in the airways of subjects with asthma, as well as to enteric nerves in subjects with inflammatory bowel disease (8). In vitro, we have shown that the process of localization of eosinophils to nerve cells is mediated in part via interactions between the neuronal adhesion molecules intercellular adhesion molecule-1 (ICAM-1) and vascular cell adhesion molecule-1 (VCAM-1) and eosinophil CD11/18 and VLA-4, respectively (9). Eosinophil adhesion to cholinergic nerve cells leads to eosinophil activation and degranulation via adhesion-dependent, nerve-generated reactive oxygen species (ROS) (10). The nerve-induced activation of eosinophils influences acetylcholine release from these nerves through an eosinophil-mediated loss of function of neural M2 muscarinic autoreceptors (9–11). Thus, one effect of eosinophil localization to nerve cells is that, acutely, they increase acetylcholine release.

Recent studies have shown that allergic inflammation also has long-lasting effects on neuron function and phenotype (12, 13). For these effects to occur they must be mediated by an inflammation-induced activation of transcription factors and subsequent gene transcription in neurons. In some cell models, it has been shown that cross-linking of ICAM-1 or VCAM-1 induces activation of transcription factors and mitogen-activated protein (MAP) kinases (14–16). In this study, we tested the hypothesis that eosinophil adhesion to nerves results in the activation of nuclear transcription factors nuclear factor-{kappa}B (NF-{kappa}B) and activator protein-1 (AP-1), which play a pivotal role in nerve cell survival and maintenance of nerve phenotype (17–20).


   Materials and Methods
Top
 Abstract
 Introduction
 Materials and Methods
Results
 Discussion
 References
 
Materials
Dulbecco's modified Eagle's medium (DMEM) Plus Glutamax, fetal calf serum (FCS), and penicillin/streptomycin solution were purchased from GIBCO/BRL Life Technologies (Paisley, UK). The IMR32 cell line was obtained from ECACC (Salisbury, UK) and depleted of fibroblasts using immunomagenetic antifibroblast microbeads and LD MACS separation columns purchased from Miltenyi Biotech (Bisley, UK). Gentamicin, Trypan Blue, poly (dI-dC.dI-dC):poly (dI-dC.dI-dC), CDP-Star chemiluminescent substrate solution, Igepal CA-630, phenylmethylsulfonyl flouride (PMSF), dithithreitol (DTT), and all common buffer constituents were obtained from Sigma (Poole, UK). I-Block for Western blot blocking and Nitro-Block II, chemiluminescent substrate component for alkaline phosphatase, were purchased from Tropix (Bedford, MA). Dulbecco's phosphate-buffered saline (PBS) was purchased from Invitrogen Ltd (Paisley, UK). Monoclonal mouse anti-human anti-CD11/18 antibody (clone 68–5A5, isotype IgG2a) was from Cymbus Biotechnology (Chandlers Ford, UK). Polyclonal rabbit anti-human anti-phospho-p38 antibody was obtained from Cell Signaling Technology (Beverly, MA). NF-{kappa}B binding site consensus oligonucleotide (5'-AGT TGA GGG GAC TTT CCC AGG C-3'), anti-mouse IgG alkaline phosphatase (AP) conjugate, anti-rabbit IgG AP conjugate, PD98059, and T4 polynucleotide kinase were obtained from Promega (Madison, WI). Monoclonal mouse anti-human anti–phospho-ERK antibody (E-4, isotype IgG2a), polyclonal rabbit anti-rat/human anti-ERK2 antibody (K-23), monoclonal mouse anti-human anti-p38 antibody (A-12, isotype IgG1), polyclonal rabbit anti-human anti-I{kappa}Bß antibody (S-20), polyclonal goat anti-human NF-{kappa}B p50 (C-19), polyclonal rabbit anti-human NF-{kappa}B p65 (C-20), and polyclonal rabbit anti-mouse/human NF-{kappa}B c-Rel (N), consensus (5'-CGC TTG ATG ACT CAG CCG GAA-3') AP-1 binding site oligonucleotide and mutant NF-{kappa}B binding site oligonucleotide (5'-AGT TGA GGC GAC TTT CCC AGG C-3') were all purchased from Santa Cruz Biotechnology (Santa Cruz, CA). Ficoll-Paque PLUS was purchased from Amersham Pharmacia Biotech (Little Chalfont, UK). CD16 immunomagnetic beads and VS+ VarioMacs columns were purchased from Miltenyi Biotech. Speedy-Diff was obtained from Clin-Tech Ltd (Clacton-on-Sea, UK). {gamma}-32P ATP was from NEN (Zaventem, Belgium). SB239063 was a gift from Dr. Kristen E. Belmonte (GlaxoSmithKline Pharmaceuticals, Philadelphia, PA), ZD7349 was a gift from Dr. Duncan Haworth (Astra Zeneca Pharmaceuticals, Loughborough, UK).

IMR32 Nerve Cell Culture
The human cholinergic neuroblastoma cell line IMR32 was depleted of fibroblasts by labeling with immunomagenetic antifibroblast microbeads and applying to LD MACS separation columns as recommended by the manufacturer. Fibroblast depletion was verified by Western blotting with a mouse antihuman fibroblast antibody (Serotec) and by observation of cell morphology. Fibroblast-depleted IMR32 cells were used for all experiments. They were maintained in culture in proliferation media (DMEM Plus Glutamax, 5% FCS, 100 U/ml penicillin/streptomycin, 10 µg/ml gentamicin) at 37°C in an atmosphere of 5% CO2. Upon achieving confluence, cells were plated at a density of 5 x 105/well in 6-well cell culture dishes and grown in proliferation medium for 48 h. Proliferation medium was then replaced by differentiation medium (DMEM Plus Glutamax, 2% FCS, 2 mM sodium butyrate, 100 U/ml penicillin/streptomycin, 10 µg/ml gentamicin) and cells were used for experimentation after a further 6–7 d of differentiation in culture. During this period an approximate doubling in cell number was achieved.

Eosinophil Isolation
Eosinophils were prepared from 45 ml of peripheral blood from healthy human volunteers by a negative immunomagnetic selection technique, essentially as described previously (9). Following phlebotomy, 15 ml of blood were added to 25 ml of PBS containing 100 U of heparin and 30 ml of blood/PBS were layered on to 23 ml Ficoll (1.077 ± 0.001 g/ml). Centrifugation at 500 x g for 20 min at room temperature was performed, the upper layer and monocyte layer were discarded and the resulting granulocytes, and red blood cell–containing pellet was subjected to hypotonic water lysis, for removal of red blood cells. This was performed with 18 ml ice-cold dH2O for 30 s followed by 2 ml of PIPES-buffered salt solution (250 mM PIPES, 1.1 M NaCl, 50 mM KCl, 420 mM NaOH; pH 7.47). Following hypotonic lysis, the granulocyte pellet was washed in PAG buffer (PIPES-buffered salt solution plus 55.5 mM glucose and 0.0003% [wt/vol] human serum albumin). Granulocytes were then resuspended in MACS buffer (PBS plus 2 mM EDTA and 0.5% [wt/vol] bovine serum albumin) with an equal volume of human CD16 immunomagnetic microbeads at 1 µl of beads/106 cells at 4°C for 30 min. After washing and resuspension in MACS buffer, granulocytes were applied to a VS+ VarioMacs column. Immunomagnetically labeled neutrophils were retained on the column and eluted eosinophils collected and resuspended in differentiation medium (DMEM Plus Glutamax, 2% FCS, 2 mM sodium butyrate, 100 U/ml penicillin/streptomycin, 10 µg/ml gentamicin). Cell viability was assessed by Trypan blue exclusion and eosinophil purity determined by Speedy-Diff staining. Only populations of eosinophils which were > 98% pure and > 95% viable were used in experimentation. For experimentation, 2 x 105 eosinophils/well were added to differentiated IMR32 cells plated as above in 6-well cell culture plates.

Eosinophil Membrane Preparation
Immediately upon isolation, eosinophils were resuspended in cold, sterile dH2O, incubated on ice for 15 min, then centrifuged at 1,500 x g for 10 min at 4°C. This process was repeated two more times and the resulting lysed cell membranes were resuspended in differentiation medium and added to differentiated IMR32 cells plated as described above in 6-well cell culture plates at an amount equivalent to 2 x 105 whole eosinophils/well. These membrane preparations contain no granular proteins, as shown by immunoblotting for the eosinophil protein MBP.

Nuclear and Cytoplasmic Protein Preparation
IMR32 cells (5 x 105) were differentiated for 6–7 d with sodium butyrate as described above and then incubated with 2 x 105 eosinophils for varying time periods from 2 min to 24 h. In some experiments, IMR32 cells were pretreated with inhibitors of eosinophil adhesion for 1 h, and co-culture experiments with eosinophils were performed in the presence or absence of these inhibitors. In other experiments, IMR32 cells were pretreated with inhibitors of the MAP kinases ERK1/2 (PD98059, 50 µM) or p38 (SB239063, 10 µM) for 24 h. Nuclear and cytoplasmic extracts were isolated from IMR32 cells, essentially as in (21) with some modifications. Briefly, cells were harvested in 1 ml ice-cold PBS and pelleted by centrifugation at 3,800 x g for 5 min at 4°C. Cells were resuspended in 1 ml hypotonic buffer (10 mM HEPES [pH 7.9], 1.5 mM MgCl2, 10 mM KCl, 0.5 mM PMSF, 0.5 mM DTT) and pelleted by centrifugation at 13,000 x g for 10 min at 4°C before lysis for 10 min on ice in 20 µl hypotonic buffer containing 0.1% Igepal CA-630. Lysates were centrifuged as before and the supernatant cytoplasmic extract removed to fresh tubes. Protein concentration was established by the Bradford method (22) and the cytoplasmic extract stored at –80°C. The nuclear pellet was lysed in 15 µl lysis buffer (20 mM HEPES [pH 7.9], 420 mM NaCl, 1.5 mM MgCl2, 0.2 mM EDTA, 25% vol/vol glycerol, 0.5 mM PMSF) for 15 min on ice. After centrifugation, as before, supernatant nuclear extracts were removed into 35 µl storage buffer (10 mM HEPES [pH 7.9], 50 mM KCl, 0.2 mM EDTA, 20% [vol/vol] glycerol, 0.5 mM PMSF, 0.5 mM DTT). Protein concentration was determined as before and nuclear extracts stored at –80°C.

Electrophoretic Mobility Shift Assay
Nuclear extracts (10 µg) were incubated with 1.6 kBq [{gamma}-32P]ATP (3,000 Ci/mmol) and T4 polynucleotide kinase end-labeled oligonucleotides containing NF-{kappa}B or AP-1 consensus sequence. Incubations were performed for 30 min at room temperature in binding buffer (4% [vol/vol] glycerol, 1 mM EDTA, 10 mM Tris-HCl [pH 7.5], 100 mM NaCl, 5 mM DTT, 0.1 mg/ml nuclease-free BSA), and 2 µg poly (dI-dC.dI-dC):poly (dI-dC.dI-dC). In experiments to test specificity of transcription factor activation, unlabeled wild-type or mutant oligonucleotides were added before incubation with labeled oligonucleotides. Reaction mixtures were electrophoresed on native 5% polyacrylamide gels that were subsequently dried and viewed on a Storm 820 Scanner PhosphorImager (Molecular Dynamics, Sunnyvale, CA).

Western Blotting
Cytoplasmic protein extracts (10 µg for pERK or I{kappa}Bß analysis or 30 µg for phospho-p38 analysis) were heated to 95°C in sample buffer (100 mM Tris pH 6.8, 2% [wt/vol] SDS, 0.002% [wt/vol] bromophenol blue, 20% [vol/vol] glycerol, 10% [vol/vol] ß-mercaptoethanol) and separated by SDS-PAGE on 10% polyacrylamide separating gel overlaid with 4% stacking gel at 500 V for 1 h. The separated proteins were transferred on to nitrocellulose membranes in transfer buffer (20 mM Tris, 150 mM glycine, 0.01% [wt/vol] SDS, 20% [vol/vol] methanol) at 500 V overnight. For immunodetection with mouse anti-human anti–phospho-ERK antibody, rabbit anti-rat/human anti-ERK2 antibody, mouse anti-human anti-p38 antibody, or rabbit anti-human anti-I{kappa}Bß antibody, membranes were incubated in blocking buffer (Dulbecco's PBS containing 0.2% [wt/vol] I-block and 0.1% [vol/vol] Tween-20) for 1 h at room temperature, then incubated for 2 h in blocking buffer containing the individual respective antibody (1:500 for each). After six 5-min washes in washing buffer (PBS pH 7.4, 0.1% [vol/vol] Tween-20), membranes were incubated for 1 h in blocking buffer containing the appropriate goat anti-mouse (phospho-ERK, p38) or goat anti-rabbit (ERK2, I{kappa}Bß) IgG AP conjugate (1:10,000). Membranes were then washed six times for 5 min each and exposed to CDP Star chemiluminescent substrate solution plus Nitro-Block II chemiluminescent substrate compound for AP (19:1) for 5 min at room temperature. Blots were then exposed to X-OMAT light sensitive film to obtain an image. For analysis of phospho-p38, after protein transfer membranes were incubated in blocking buffer consisting of TBST (10 mM Tris pH 7.5, 100 mM NaCl, 0.1% [vol/vol] Tween-20) containing 5% [wt/vol] nonfat dry milk for 1 h at room temperature. Membranes were washed three times for 5 min each in TBST at room temperature and then incubated at 4°C overnight in rabbit anti-human phospho-p38 MAP kinase antibody (1:1,000) in TBST containing 5% (wt/vol) BSA. After three 5-min washes in TBST at room temperature, membranes were incubated for 2 h at room temperature with goat anti-rabbit IgG alkaline phosphatase conjugate (1:10,000) diluted in blocking buffer and subjected to three further 5-min washes in TBST. Blots were then processed for chemiluminescent analysis and exposed to X-ray film as described above for all other antibodies.

Statistical Analysis
Values are expressed as mean ± SD. The statistical significance of differences between treated samples and the appropriate time point control was evaluated by an unpaired, two-tail t test; *P < 0.05, **P < 0.005.


   Results
Top
 Abstract
 Introduction
 Materials and Methods
 Results
Discussion
 References
 
Eosinophil Co-Culture Induces Activation of NF-{kappa}B and AP-1 in IMR32 Cholinergic Nerve Cells; Dependence on Adhesion
Peripheral blood eosinophils were co-incubated with differentiated IMR32 cells for various time periods. Electrophoretic mobility shift analysis (EMSA) showed that co-culture induced a strong and rapid activation of NF-{kappa}B in IMR32 cells, within 2 min of co-incubation, which remained evident over a 1-h time period, declining by 2 h of co-culture (Figures 1A and 2A). Extended co-culture at 3, 6, and 24 h revealed a further wave of NF-{kappa}B activation in IMR32 cells in response to eosinophils, reaching a maximum at 24 h co-culture (Figure 1B). In competition studies with unlabeled oligonucleotide, wild-type NF-{kappa}B consensus binding site blocked NF-{kappa}B activation, whereas mutant NF-{kappa}B binding site did not, confirming that NF-{kappa}B specifically was involved (data not shown). NF-{kappa}B activation was accompanied by a concomitant degradation of I{kappa}Bß but not I{kappa}B{alpha} in cytoplasmic fractions from IMR32 cells (Figure 1C and data not shown).



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  		Figure 1. NF-{kappa}B and AP-1 are activated in IMR32 cells in response to eosinophil co-culture. EMSA of NF-{kappa}B (A, B) or AP-1 (D) in nuclear extract, Western blot of I{kappa}Bß (C) in cytoplasmic extract, from IMR32 cells (5 x 105) differentiated for 7 d and left untreated (lane 1) or co-incubated with eosinophils (2 x 105) for the indicated time period (A–D). A–D show representative results from at least three independent experiments.

 


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  		Figure 2. NF-{kappa}B activation is mediated by eosinophil CD11/18 and neuronal ICAM-1. Graphs show percentage change in neuronal NF-{kappa}B activation following co-culture with eosinophils for the indicated times compared with control samples of IMR32 cells (5 x 105) differentiated for 7 d in the absence of eosinophils. In A, a time course for the effect of eosinophils on NF-{kappa}B activation is shown. In B, the effect on neuronal NF-{kappa}B activation of inhibiting eosinophil adhesion to nerves via CD11/18 (with 0.1 µg/ml anti-CD18/11 antibody) is shown. The lower panel is a representative Western blot showing the effect of inhibition of eosinophil adhesion via CD11/18 on neuronal I{kappa}Bß degradation. In C, the effect on neuronal NF-{kappa}B activation of inhibiting eosinophil adhesion to nerves via VLA-4 (with 10 µM ZD7349) is shown. The lower panel is a representative Western blot showing the effect of inhibition of eosinophil adhesion via VLA-4 on neuronal I{kappa}Bß degradation. In D, the effect of inhibiting both adhesion molecules on neuronal NF-{kappa}B activation is shown. The lower panel is a representative Western blot showing the effect of inhibition of eosinophil adhesion via CD11/18 plus VLA-4 on neuronal I{kappa}Bß degradation. Percentage NF-{kappa}B induction is based on a comparison of band intensities from EMSA gels of nuclear extract, calculated from the area under the curve of plots of pixel intensities generated using ImageQuant on the Storm 820 phosphoimaging system. The figure shows representative results from at least four independent experiments. Data are mean ± SD. NF-{kappa}B percentage activation at each time point of eosinophil co-culture for each inhibitor is individually compared with the percentage NF-{kappa}B activation observed at that time point in the absence of inhibitor, *P < 0.05, **P < 0.005, significantly reduced compared with eosinophil-stimulated IMR32 cells at the corresponding time point.

 
EMSA also showed that eosinophil co-culture induced a strong and rapid activation of AP-1 in IMR32 cells, over a similar time course to NF-{kappa}B (Figure 1D).

In a number of preliminary experiments, we established that the signals obtained in these Western blots and EMSA were from IMR32 cells and not eosinophils. First, the number of eosinophils used in these experiments was relatively few, and when Western blots and EMSAs were performed on stimulated nuclear and cytoplasmic extracts of this quantity of eosinophils it was not possible to observe a signal (data not shown). Furthermore, we have previously shown (9) that optimally one-third of eosinophils added adhere to the nerve cells, making their potential contribution correspondingly smaller, and in the bandshift experiments described herein typically between one-tenth and one-fifth of the total nuclear protein extract (10 µg) generated was used, again further reducing any potential contribution from adherent eosinophils. Use of paraformaldehyde-fixed eosinophils, which have no response to cell-stimulating agents, also induced I{kappa}Bß degradation (data not shown).

To determine whether NF-{kappa}B activation in IMR32 cells was dependent on adhesion of eosinophils, we treated the nerve cells with inhibitors of adhesion before and during eosinophil co-culture. NF-{kappa}B activation at each time point was expressed as a percentage of NF-{kappa}B activation seen in control IMR32 cells in the absence of eosinophils. Incubation of eosinophils with nerve cells in the absence of inhibitor led to a 2- to 3-fold activation of NF-{kappa}B starting at 2 min of co-culture and reaching a maximum at 1 h (Figure 2A). To assess the effect of adhesion inhibitors on NF-{kappa}B activation at each time point, we compared NF-{kappa}B activation in cells in the presence of adhesion inhibitors and eosinophils to that observed, at the corresponding time point, in the eosinophil only–treated cells. The concentration of each inhibitor was based on observations from two prior studies in which the effect of various concentrations of each inhibitor on eosinophil adhesion to IMR32 cholinergic nerves was investigated (9, 10). None of the inhibitors used had any significant effect on baseline NF-{kappa}B activation in the absence of eosinophils (data not shown).

When eosinophil adhesion to IMR32 nerve cells via CD11/18 was inhibited, the activation of NF-{kappa}B was significantly reduced, reaching control levels at most time points. I{kappa}Bß degradation in the cytoplasmic fraction did not occur until 2 h of co-culture (Figure 2B, bottom panel). To assess a possible role for very late antigen-4 (VLA-4)/VCAM-1–mediated adhesion in transcription factor activation, experiments were performed in the presence of ZD7349, an inhibitor of VLA-4. NF-{kappa}B activation was not significantly affected by ZD7349 (Figure 2C). Similarly, I{kappa}Bß degradation in the cytoplasmic fraction was similar to IMR32 cells treated with eosinophils only (Figure 2C, bottom panel). A combination of the two inhibitors completely inhibited NF-{kappa}B activation (Figure 2D) and delayed I{kappa}Bß degradation in the cytoplasmic fraction until 2 h of co-culture, similarly to anti-CD11/18 alone (Figure 2D, bottom panel).

Similar assessment of dependence of AP-1 activation on eosinophil adhesion revealed that, in contrast to NF-{kappa}B, inhibition of eosinophil adhesion to IMR32 nerve cells via CD11/18 had no significant effect on AP-1 activation (Figure 3B). By contrast, when experiments were performed in the presence of ZD7349, AP-1 activation in IMR32 cells in response to eosinophils was significantly inhibited (Figure 3C). A combination of anti-CD11/18 with ZD7349 did not further reduce AP-1 activation (Figure 3D).



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  		Figure 3. AP-1 activation is mediated by eosinophil VLA-4 and neural VCAM-1. Graphs show percentage change in AP-1 activation following co-culture with eosinophils compared with control samples of IMR32 cells differentiated for 7 d in the absence of eosinophils (0.5 x 106). In A, a time course for the effect of eosinophils on neuronal AP-1 activation is shown. In B, the effect on neuronal AP-1 activation of inhibiting eosinophil adhesion to nerves via CD11/18 (anti CD18/11 antibody, 0.1 µg/ml) is shown. In C, the effect on neuronal AP-1 activation of inhibiting eosinophil adhesion to nerves via VLA-4 (with ZD7349, 10 µM) is shown. In D, the effect on neuronal AP-1 activation of inhibiting both adhesion molecules is shown. Percentage induction is based on a comparison of band intensities from EMSA gels of nuclear extract, calculated from the area under the curve of plots of pixel intensities generated using ImageQuant on the Storm 820 phosphoimaging system. The figure shows representative results from at least three independent experiments. Data are mean ± SD. AP-1 percentage activation at each time point of eosinophil co-culture for each inhibitor is individually compared with the percentage AP-1 activation observed at that time point in the absence of inhibitor, *P < 0.05, **P < 0.005.

 
To determine that adhesion alone, in the absence of any other eosinophil-derived factors, could induce early transcription factor activation in IMR32 cells, eosinophil membranes were prepared and used to stimulate IMR32 cells in time courses ranging from 2 min to 2 h (Figure 4). Our preliminary data have confirmed that eosinophil membrane preparations are free of eosinophil proteins such as MBP (P. Kingham, unpublished observation).



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  		Figure 4. Eosinophil membrane preparations can induce adhesion-dependent activation of NF-{kappa}B and AP-1 in IMR32 cells. EMSA of NF-{kappa}B (A) or AP-1 (B) in nuclear extract from IMR32 cells (0.5 x 106) differentiated for 7 d and exposed to membranes from eosinophils (made from 0.2 x 106 eosinophils). In C, activation was inhibited with anti-CD11/18 antibody or with the VLA-4 inhibitor ZD7349 (D). The figure shows representative results from three independent experiments.

 
Eosinophil membranes induced NF-{kappa}B activation (Figure 4A) and AP-1 activation (Figure 4B) in IMR32 cells by 2 min of stimulation. Activation remained evident at time points up to 1 h and declined at 2 h. Furthermore, eosinophil membrane–induced NF-{kappa}B was also inhibited in the presence of anti-CD11/18 (Figure 4C), whereas AP-1 activation was inhibited by ZD7349 (Figure 4D).

Eosinophil Adhesion–Induced Activation of ERK1/2 and p38 MAP Kinases
To determine the mechanism of eosinophil adhesion–induced transcription factor activation in IMR32 cells, we examined the role of the MAP kinases extracellular signal–regulated protein kinase (ERK)1/2 and p38. Western blots of cytoplasmic protein from IMR32 cells co-cultured with eosinophils for various times from 2 min to 2 h were probed with an antibody specific to the dual phosphorylated form of ERK1/2 (Figures 5A and 6A, top panels). Rapid phosphorylation of ERK1/2 in the IMR32 cytoplasmic fraction was evident within 2 min of co-incubation, declining by 10 min. Activation of p38 was observed in the IMR32 cytoplasm after 5 min of co-culture, declining by 10 min (Figures 5B and 7A, top panels). The same blots were re-probed with anti-ERK2 antibody (Figures 5A and 6A, bottom panels) or with anti-p38 antibody (Figures 5B and 7A, bottom panels) to confirm even loading of total ERK or p38.



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  		Figure 5. ERK1/2 and p38 are phosphorylated in IMR32 cells in response to eosinophil co-culture. Western blot of cytoplasmic extract (10 µg) from IMR32 cells (0.5 x 106) differentiated for 7 d and then co-incubated with eosinophils (0.2 x 106) for the indicated time period. Blots were probed with (A) anti–phospho-ERK antibody (upper blot) then stripped and re-probed with anti-ERK2 (lower blot) or (B) anti–phospho-p38 antibody (upper blot) then stripped and re-probed with anti-p38 (lower blot). The figure shows representative results from three independent experiments.

 


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  		Figure 6. Eosinophil-induced phosphorylation of ERK1/2 in IMR32 cell is dependent on CD11/18 ICAM-1 interactions. In the figure Western blots of cytoplasmic extract (10 µg) from IMR32 cells (0.5 x 106) differentiated for 7 d and exposed to eosinophils (0.2 x 106) in the presence of eosinophil inhibitors are shown. In A, a positive control time course for eosinophils only is shown. In B, eosinophil adhesion via CD11/18 was inhibited. In C, eosinophil adhesion via VLA-4 was inhibited and D, adhesion via both adhesion molecules was inhibited. Blots were probed with anti–phospho-ERK antibody (A–D, upper panels) then stripped and re-probed with anti-ERK2 (A–D, lower panels). The figure shows representative results from three independent experiments.

 


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  		Figure 7. Eosinophil-induced phosphorylation of p38 in IMR32 cells is dependent on adhesion. Western blots of cytoplasmic extract (30 µg) from IMR32 cells (0.5 x 106) differentiated for 7 d and exposed to eosinophils (0.2 x 106) in the presence of eosinophil inhibitors are shown. In A, a positive control time course for eosinophils only is shown. In B, eosinophil adhesion via CD11/18 was inhibited. In C, eosinophil adhesion via VLA-4 was inhibited and D adhesion via both adhesion molecules was inhibited. Blots were probed with anti–phospho-p38 antibody (A–D, upper panels) then stripped and re-probed with anti-p38 (A–D, lower panels). The figure shows representative results from three independent experiments.

 
Inhibition of eosinophil adhesion to IMR32 nerve cells via CD11/18-ICAM-1 strongly inhibited the activation of ERK1/2 for time periods ranging from 2 min to 2 h (Figure 6B). However, inhibition of eosinophil adhesion to IMR32 nerve cells via VLA-4/VCAM-1, in the presence of the inhibitor ZD7349, had little effect on ERK1/2 phosphorylation (Figure 6C). A combination of anti-CD11/18 and ZD7349 completely inhibited ERK1/2 phosphorylation (Figure 6D).

Inhibition of eosinophil adhesion to IMR32 nerve cells via CD11/18-ICAM-1 using the antibody to CD11/18 (Figure 7B) had no effect on p38 activation, whereas ZD7349 delayed p38 activation to 1 h (Figure 7C) and the two inhibitors further delayed p38 activation (Figure 7D).

Dependence of Transcription Factor Activation on the MAP Kinases ERK1/2 and p38
We then determined whether MAP kinase activation was involved in activation of the transcription factors NF-{kappa}B. Pretreatment of IMR32 cells with the ERK1/2 inhibitor PD98059 (50 µM, 24 h) significantly reduced NF-{kappa}B activation in response to eosinophils (Figure 8A). Pretreatment of IMR32 cells with the specific p38 inhibitor SB239063 (10 µM, 24 h) reduced NF-{kappa}B activation at the earliest time points, but not signifcantly (Figure 8A). PD98059 pretreatment of IMR32 cells had no significant effect on AP-1 activation, whereas pretreatment of IMR32 cells with SB239063 strongly reduced AP-1 activation (Figure 8B). These pretreatments had no effect on nerve cell viability or on basal transcription factor activation levels. The doses used are consistent with those published for effective inhibition of ERK (23, 24) or p38 (25) and their downstream effects.



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  		Figure 8. Both ERK and p38 inhibitors reduce activation of transcription factors in IMR32 cells in response to eosinophil co-culture. Graphs show percentage change in NF-{kappa}B activation (A) or AP-1 activation (B) following co-culture with eosinophils for the indicated times, compared with control samples of IMR32 cells (5 x 105) differentiated for 7 d in the absence of eosinophils. Time courses for the effect of eosinophils on NF-{kappa}B (A) or AP-1 (B) activation are shown in cells incubated with eosinophils only, or pre-treated with PD98059 (50 µM) (ERK inhibitor) or SB239063 (10 µM) (p38 inhibitor) for 24 h, then stimulated with eosinophils for the indicated time periods. Percentage induction is based on a comparison of band intensities from EMSA gels of nuclear extract, calculated from the area under the curve of plots of pixel intensities generated using ImageQuant on the Storm 820 phosphoimaging system. The figure shows representative results from at least three independent experiments. Solid bars, eosinophils only; open bars, eosinophils + ERK inhibitor; bars with hatching, eosinophil + p38 inhibitor. Data are mean ± SD. NF-{kappa}B or AP-1 percentage activation at each time point of eosinophil co-culture for each inhibitor is individually compared with the percentage NF-{kappa}B or AP-1 activation observed at that time point in the absence of inhibitor, *P < 0.05, **P < 0.005.

 

   Discussion
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
References
 
In this study we have investigated the intracellular consequences for a differentiated cholinergic neuroblastoma cell line, IMR-32, following co-culture with human eosinophils. IMR-32 cells have been previously employed by us and others (9, 10, 26–28) as a convenient in vitro model for studying aspects of neurobiology at a cellular and molecular level. Ongoing studies in our laboratory have confirmed that they inducibly express genes for a battery of cholinergic genes, including muscarinic M2 receptor, choline acetyl transferase, and vesicular acetylcholine transporter. Adhesion of eosinophils to IMR-32 cells induced activation of the transcription factors NF-{kappa}B and AP-1, both of which are important in nerve survival, growth and maintenance of phenotype. Activation of both transcription factors occurred within 2–5 min of co-culture with eosinophils. Although both had returned to baseline by 2 h, a further wave of activation was detected at 3 h and this persisted over 24 h of co-culture. Inhibition of eosinophil binding via ICAM-1 inhibited NF-{kappa}B activation while inhibition of eosinophil binding via VCAM-1 had no significant effect on NF-{kappa}B. In contrast, eosinophil adhesion to VCAM-1 led to activation of AP-1. Thus, co-culture of eosinophils with IMR32 nerve cells led to activation of neuronal transcription factors by adhesion-dependent mechanisms.

To confirm that the observed changes were occurring within the nerve cells and not the eosinophils, several controls were performed. First, we used a small number of eosinophils relative to the number of nerve cells. The number of eosinophils was insufficient to give a signal using the techniques used in these studies. Second, in preliminary experiments we showed that adhesion of paraformaldehyde-fixed eosinophils which adhere to IMR32 cells also activated these pathways. Third, we used eosinophil membranes alone instead of eosinophils to study the effects of adhesion on neural signaling. These membrane preparations do not contain eosinophil-derived factors, they induced both NF-{kappa}B and AP-1 in IMR32 cells, and this was inhibited by specific inhibitors of adhesion. Thus, the observed activation of MAP kinases and nuclear transcription factors was due to membrane–membrane interactions between nerves and eosinophils.

We have previously shown that eosinophil adhesion to nerve cells leads to activation and release of eosinophil proteins; our prior studies have shown that neuron-induced eosinophil activation is first detectable at least 30 min after adhesion (10). Therefore, although it is unlikely that eosinophil degranulation products have a predominant role in transcription factor activation at the earliest time points examined, they may be important at later time points. It is possible that eosinophil-derived nerve growth factor (NGF) could influence eosinophil-induced effects on nerves; however, immunologic stimuli appear to be necessary for substantial secretion of NGF from eosinophils sufficient to induce functional changes in nerve cells (29). Furthermore, the kinetics of the responses we observe are inconsistent with an NGF-induced effect and the ability of eosinophil membranes alone to induce transcription factor activation makes an important role for NGF unlikely.

To determine the intermediates involved in the adhesion-dependent activation of the neural transcription factors, we studied the MAP kinases ERK1/2 and p38. We studied these as the cytoplasmic domain of ICAM-1 has been shown to interact with MAP kinases in other cells such as endothelial cells (15–17). Furthermore, they are integrally linked to the activation of the transcription factors, but their precise role is highly dependent on the stimulus and cellular context (30–34). In our experiments, p38 and ERK1/2 were rapidly and transiently activated in IMR32 cells in response to eosinophil adhesion. Activation of p38 was dependent on VCAM-1, because inhibition of VLA-4/VCAM-1 interaction delayed the onset of p38 activation from 5 min to 1 h of co-culture. Inhibition of p38 activation strongly inhibited AP-1 activation, whereas inhibition of ERK1/2 had little or no effect on AP-1 activation. ERK1/2 activation was primarily dependent on ICAM-1–mediated adhesion and NF-{kappa}B activation was strongly dependent on ERK1/2 phosphorylation. At the earliest time points, the p38 inhibitor SB239063 also appeared to inhibit NF-{kappa}B activation, although this was not statistically significant. Thus, adhesion of eosinophils via CD11/18/ICAM-1 and VLA-4/VCAM-1 induced the activation of a network of intracellular pathways in IMR32 cells. These are summarized in Figure 9.



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  		Figure 9. Schematic representation of the proposed mechanism of activation of neural transcription factors by adhesion of eosinophils to nerves. Eosinophil adhesion to neural ICAM-1 led to a rapid activation of ERK1/2 and to NF-{kappa}B activation. Eosinophil adhesion via neural VCAM-1 led to phosphorylation of the p38 MAP kinase and in turn to activation of AP-1.

 
Activation of the transcription factors NF-{kappa}B and AP-1 by eosinophil binding has many potential consequences for gene expression in the nerve cells and particularly for their survival. Ongoing studies in our laboratory are addressing the impact of eosinophil adhesion and consequent transcription factor activation on expression of nerve cell cholinergic genes and consequent changes in cholinergic phenotype. NF-{kappa}B has been implicated in models of neuronal survival and plasticity (35, 36); however, its effects may depend on the cell type, level of differentiation or stimulus. Likewise, reports conflict on the effects of AP-1 on neuronal cell survival (37, 38). It remains to be determined whether the NF-{kappa}B and AP-1 activation rapidly induced in IMR32 cells by eosinophil binding impacts on neuronal cell survival and whether the transcription factors co-operate or interfere with each other and other intracellular modulators in this cell context. Our ongoing studies are addressing the impact of eosinophil adhesion and consequent transcription factor activation on expression of pro- and antiapoptotic genes and the influence on nerve cell survival. Recent experiments have shown that eosinophil adhesion–induced activation of p38 leads to neurite retraction, suggesting that one effect of eosinophil adhesion to cholinergic nerves is nerve remodeling (39).

In conclusion, we have demonstrated that adhesion of eosinophils to IMR32 cholinergic nerve cells induces a network of intracellular signaling pathways. If such an impact is mirrored, in vivo, during the eosinophilic infiltration and accumulation at nerves previously observed in antigen-challenged animals and in asthma in humans, it may have an important impact on the gene expression and function of the nerve cells.


   Acknowledgments
 
This work was supported by the Wellcome Trust and the Health Research Board, Ireland.

Received in original form May 9, 2003

Received in final form July 4, 2003


   References
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 References
 

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