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Published ahead of print on April 28, 2005, doi:10.1165/rcmb.2005-0056OC
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American Journal of Respiratory Cell and Molecular Biology. Vol. 33, pp. 169-177, 2005
© 2005 American Thoracic Society
DOI: 10.1165/rcmb.2005-0056OC

Diverse Effects of Eosinophil Cationic Granule Proteins on IMR-32 Nerve Cell Signaling and Survival

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

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

Correspondence and requests for reprints should be addressed to Richard 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
 
Activated eosinophils release potentially toxic cationic granular proteins, including the major basic proteins (MBP) and eosinophil-derived neurotoxin (EDN). However, in inflammatory conditions including asthma and inflammatory bowel disease, localization of eosinophils to nerves is associated with nerve plasticity, specifically remodeling. In previous in vitro studies, we have shown that eosinophil adhesion to IMR-32 nerve cells, via nerve cell intercellular adhesion molecule-1, results in an adhesion-dependent release of granule proteins. We hypothesized that released eosinophil granule proteins may affect nerve cell signaling and survival, leading to nerve cell remodeling. Culture in serum-deprived media induced apoptosis in IMR-32 cells that was dose-dependently abolished by inclusion of MBP1 but not by EDN. Both MBP1 and EDN induced phosphorylation of Akt, but with divergent time courses and intensities, and survival was independent of Akt. MBP1 induced activation of neural nuclear factor (NF)-{kappa}B, from 10 min to 12 h, declining by 24 h, whereas EDN induced a short-lived activation of NF-{kappa}B. MBP1-induced protection was dependent on phosphorylation of ERK 1/2 and was related to a phospho-ERK-dependent upregulation of the NF-{kappa}B–activated anti-apoptotic gene, Bfl-1. This signaling pathway was not activated by EDN. Thus, MBP1 released from eosinophils at inflammatory sites may regulate peripheral nerve plasticity by inhibiting apoptosis.

Key Words: plasticity • apoptosis • MAP kinases


   Introduction
Top
 Abstract
 Introduction
MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 References
 
Recruitment of eosinophils and the release of their granule proteins in tissues is the hallmark of a variety of common clinical conditions, including asthma and a number of gastrointestinal diseases. The eosinophil cationic granule proteins are the major basic proteins (MBP1 and 2), eosinophil peroxidase (EPO), and the closely related eosinophil RNases, eosinophil-derived neurotoxin (EDN) and eosinophil cationic protein (ECP). The numbers and levels of eosinophils and granule proteins, respectively, in tissue, correlate with disease severity, suggesting that they play a pathogenic role in these conditions (1–3). The cationic granule proteins have toxic effects on tissue-invasive parasites and bacteria in vitro and eosinophils are thus considered to be important in host defense (4). However, the proteins also have toxic and degenerative effects on mammalian cells (3, 5), suggesting that eosinophil granule proteins may contribute to the pathogenesis of disease via tissue toxicity.

A central role for eosinophils in asthma was called into question when an antibody to interleukin (IL)-5, a major eosinophil survival factor, had disappointing results as a potential asthma therapy (6). However, this antibody reduced airway tissue eosinophil accumulation by only ~ 50% and had no effect on MBP1 deposition (7). Furthermore, in a recently developed transgenic mouse model congenitally deficient in eosinophils, allergen challenge did not result in pulmonary mucus accumulation and the airway hyperresponsiveness associated with asthma (8). Recent studies suggest another role of eosinophils in repair and remodeling after injury, which may also contribute to the apparent paradox between the correlation of eosinophils with disease severity and the failure of specific inhibitory therapies to improve acute clinical symptoms (6, 9).

Under the influence of inflammatory signals nerves may also undergo remodeling, termed neural plasticity, an event that can render the nerves hyperresponsive (10). We have previously shown that eosinophils accumulate at cholinergic nerves in a variety of diseases, including at airway nerves of individuals with asthma and at enteric nerves of subjects with inflammatory bowel disease (11). In in vitro studies we have employed the neuroblastoma cell line IMR-32 as a useful model of cholinergic nerve cell function. Differentiated IMR-32 cells express neuronal muscarinic M2 receptors and release acetylcholine in response to eosinophil adhesion in an M2-dependent manner (12, 13). Differentiated IMR-32 cells also express other cholinergic genes and their protein products, including choline acetyltransferase (N. Durcan and R. Costello, unpublished results). Our in vitro studies have shown that eosinophils adhere to both primary parasympathetic nerves and to IMR-32 cells via interactions between nerve intercellular adhesion molecule-1 (ICAM-1) and vascular cell adhesion molecule-1 (VCAM-1) and the respective eosinophil counterligands (12–14). This adhesion results in both the release of eosinophil granule proteins and the generation of signaling events within the nerve cells (12–15). We have elucidated some of the intracellular signaling pathways consequent to eosinophil adhesion to IMR-32 cells; for example, eosinophil adhesion via ICAM-1 results in a phospho-ERK-dependent activation of the transcription factor nuclear factor (NF)-{kappa}B, while adhesion via VCAM-1 results in activation of transcription factor activator protein-1, which is partially dependent on the MAP kinase, p38 (15). We also showed that eosinophil adhesion induces neurite retraction of IMR-32 cells via a dual mechanism comprising both a p38-dependent element and tyrosine kinase–dependent generation of reactive oxygen species (ROS) (16). Furthermore, eosinophil adhesion to nerves promotes the release of acetylcholine and a cholinergic phenotype due to factors released from eosinophils (16). Thus, both eosinophil adhesion and the eosinophil products released as a consequence of such adhesion lead to nerve remodeling.

Recently, we have shown that eosinophil adhesion to IMR-32 cells confers protection on nerve cells from apoptosis induced either by proinflammatory cytokines (tumor necrosis factor, IL-1ß, and interferon-{gamma}) or by serum deprivation (17). This is consequent to eosinophil adhesion to IMR-32 cells because eosinophil membrane preparations, which lacked eosinophil granule products, conferred similar protection to that provided by whole eosinophils. Protection was dependent on adhesion via both ICAM-1 and VCAM-1 and on ERK 1/2 but not p38 activation (17). Having identified the role of surface adhesion per se, we have now attempted to identify the actions of the products released from the eosinophils. Given that eosinophils and their granule proteins are involved in remodeling after injury, we hypothesized that the eosinophil granule proteins MBP1 and EDN would influence nerve plasticity by protecting nerves from injury.


   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 IMR-32 cell line was obtained from ECACC (Salisbury, UK) and depleted of fibroblasts with immunomagenetic antifibroblast microbeads and LD MACS separation columns purchased from Miltenyi Biotech (Bisley, UK). TRI reagent, Gentamicin, Trypan Blue, poly (dI-dC.dI-dC):poly (dI-dC.dI-dC), LY294002, wortmannin, Igepal CA-630, phenylmethylsulfonyl flouride (PMSF), dithiothreitol (DTT), and all common buffer constituents were obtained from Sigma (Poole, UK). I-Block for Western blot blocking was purchased from Tropix (Bedford, MA). Dulbecco's phosphate-buffered serum (PBS) was purchased from Invitrogen Ltd (Paisley, UK). Polyclonal rabbit anti-human anti–phospho-p38 antibody and LumiGlo Reagent A and Peroxide Reagent B for HRP detection were 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 (HRP) conjugate, anti-rabbit IgG HRP conjugate, PD98059, and T4 polynucleotide kinase were obtained from Promega (Madison, WI). Monoclonal mouse anti-human phospho-ERK antibody (E-4, isotype IgG2a), polyclonal rabbit anti-rat/human ERK 2 antibody (K-23), monoclonal mouse anti-human p38 antibody (A-12, isotype IgG1), polyclonal goat anti-human Akt 1 antibody (C-20), polyclonal rabbit anti-human phospho-Akt 1/2/3 (Ser473), and rabbit anti-goat IgG HRP conjugate were all purchased from Santa Cruz Biotechnology (Santa Cruz, CA). [{gamma}-32P] ATP was from NEN (Zaventem, Belgium). TACS Annexin V–FITC apoptosis detection kit was purchased from R&D Systems (Minneapolis, MN). First-strand cDNA synthesis kit was purchased from Roche Diagnostics (Mannheim, Germany).

Isolation of Eosinophil MBP1 and EDN
The human eosinophil cationic proteins MBP1 and EDN were isolated from eosinophil granule proteins as previously described (18, 19).

IMR-32 Nerve Cell Culture
The human cholinergic neuroblastoma cell line IMR-32 was depleted of fibroblasts by labeling with immunomagenetic anti-fibroblast microbeads and applying to LD MACS separation columns as recommended by the manufacturer. Fibroblast depletion was verified by Western blotting with a mouse anti-human fibroblast antibody (Serotec) and by observation of cell morphology. Fibroblast-depleted IMR-32 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.

Nuclear and Cytoplasmic Protein Preparation
IMR-32 cells (5 x 105) were differentiated for 6–7 d with sodium butyrate as described above and then incubated with the eosinophil proteins MBP1 or EDN for various time periods from 10 min to 24 h. In some experiments, IMR-32 cells were pretreated with inhibitors of the MAP kinase ERK 1/2 (PD98059, 50 µM) and/or the PI3K/Akt inhibitors wortmannin (100 nM) or LY294002 (10 µM) for 2 h. Nuclear and cytoplasmic extracts were isolated from IMR-32 cells, essentially as in Ref. 15. 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 (20) 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 oligonucleotides containing NF-{kappa}B or AP-1 consensus sequence, which were end-labeled with 1.6 kBq [{gamma}-32P]ATP (3,000 Ci/mmol) and T4 polynucleotide kinase. 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). 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 or total protein extracts (10 µg for pERK, Akt 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 phospho-ERK antibody, rabbit anti-rat/human ERK2 antibody, goat anti-human Akt 1, rabbit anti-human phospho-Akt 1/2/3 (Ser473), or mouse anti-human p38 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 (ERK 2, phospho-Akt 1/2/3) IgG HRP conjugate (1:2,000) or rabbit anti-goat (Akt 1) IgG HRP conjugate (1:10,000). Membranes were then washed six times for 5 min each and exposed to LumiGlo substrate solution (Cell Signaling Technology) for 1 min at room temperature according to the manufacturer's instructions. 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 HRP conjugate (1:2,000) 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.

Annexin V Binding Apoptosis Assay
Differentiated cholinergic IMR-32 cells were grown to confluence in six-well plates for 6–7 d. Cells were then untreated or subjected to the apoptotic stimulus of culturing in serum-free medium, previously shown to induce apoptosis (17) in the presence or absence of eosinophil granule proteins MBP1 or EDN for various times. In some experiments, IMR-32 cells were pretreated with inhibitors of the MAP kinase ERK 1/2 (PD98059, 50 µM) or the PI3K/Akt inhibitors wortmannin (100 nM) or LY294002 (10–100 µM) for 2 h.

Medium was removed and cells harvested in PBS and washed twice by centrifugation at 500 x g. Cells were then incubated with annexin V–FITC and propidium iodide (PI) according to manufacturer's guidelines. The cells were subsequently analyzed by Flow Cytometry (Coulter Epics XL; Beckman Coulter, High Wycombe, UK). Data from 10,000 events were collected in logarithmic mode. Dual staining with Annexin V–FITC and PI allowed for the differentiation between early apoptotic cells (Annexin V–FITC positive), late apoptotic, and/or necrotic cells (Annexin V–FITC and PI positive) and viable cells (unstained).

RNA and Total Protein Extraction and RT-PCR
For detection of message for pro- and antiapoptotic genes, cells were untreated or subjected to apoptotic stimuli as above in the presence or absence of eosinophil granule proteins. After removal of medium, IMR-32 nerve cells were harvested and washed in warm PBS, lysed at room temperature in TRI reagent and RNA, and total protein extracted according to the manufacturer's guidelines. One microgram of RNA was reverse transcribed with AMV reverse transcriptase, random hexamer primers and a 1-st strand cDNA synthesis kit. RT-PCR analysis of cDNA preparations was performed in 50-µl reactions with Taq-DNA polymerase and the following primers sets: Bfl-1: 5'- TTA CAG GCT GGC TCA GGA CT-3' (forward) and 5'-CCC AGT TAA TGA TGC CGT CT-3' (reverse); ß-actin: 5'-TCC TGT GGC ATC CAC GAA ACT-3' (forward) and 5'-GAA GCA TTT GCG GTG GAC GAT-3' (reverse). PCR conditions were: 94°C, 3 min (1 cycle); 94°C, 1 min, 54°C, 1 min, 72°C, 2 min (20–30 cycles); 72°C, 10 min (1 cycle). PCR products were separated by 1.5% agarose gel electrophoresis and photographed under ultraviolet illumination. Results presented are for 30 cycles of PCR (Bfl-1), or 25 cycles (ß-actin). Band intensities were quantified by densitometry scanning. The results were expressed as a ratio of the band intensity relative to the corresponding ß-actin band obtained by amplification of the same template cDNA.

Statistical Analysis
Values are expressed as mean ± SD. The statistical significance of differences between treated samples and the appropriate time point control or between treated and untreated samples was evaluated by ANOVA followed by Tukey-Kramer pairwise multiple comparisons. A P value of 0.05 or less was taken as being significant.


   RESULTS
Top
 Abstract
 Introduction
 MATERIALS AND METHODS
 RESULTS
DISCUSSION
 References
 
MBP1 Confers Protection from Apoptosis on IMR-32 Nerve Cells
We studied the effect of eosinophil granule proteins on nerve cell survival in a pro-apoptotic model, culture of IMR-32 cells in serum-free medium for 24 h. This serum deprivation induced IMR-32 cell apoptosis, confirming prior observations (17) (Figure 1A). Serum deprivation reduced the percentage of viable cells from 82.4 ± 9.5% to 66.9 ± 8.2% (P < 0.05; n = 4) and increased the percentage of apoptotic cells from 8.0 ± 4.7% to 22.4 ± 4.3% (P < 0.05; n = 4). Neither MBP1 nor EDN alone induced apoptosis of IMR-32 cells in the absence of serum deprivation (Figure 1A). Inclusion of MBP1 (1 µg/ml; ~ 7 x 10–8 M) during serum deprivation conferred protection from apoptosis on IMR-32 cells (Figure 1A), increasing the percentage of viable cells to non–serum-deprived levels of 85.8 ± 2.2% and decreasing the percentage of apoptotic cells to 11.6 ± 4.0% (n = 4; P < 0.05 versus serum-deprived cells). Dose–response experiments confirmed that MBP1, the predominant eosinophil granular product in terms of molar concentration, significantly abrogated apoptosis of IMR-32 nerve cells induced by serum deprivation at concentrations from 0.1–10 µg/ml (Figure 1B). By contrast, EDN (1 µg/ml; ~ 6 x 10–8 M), while not by itself stimulating IMR-32 cell apoptosis, also did not confer protection on IMR-32 cells subjected to serum starvation (Figure 1A), and the amount of apoptosis in serum-deprived cells in the presence of EDN was significantly greater than that observed in the presence of MBP (P = 0.01).



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  		Figure 1. MBP1 but not EDN protects IMR-32 cells from serum deprivation–induced apoptosis. IMR-32 cells (0.5 x 106) were differentiated for 7 d before being subjected to serum deprivation for 24 h. (A) Serum deprivation took place in the presence or absence of 1 µg/ml MBP1 or 1 µg/ml EDN. As a control, non–serum-deprived cells were also incubated in the presence of 1 µg/ml MBP1 or 1 µg/ml EDN. IMR-32 cells were then stained for Annexin V and propidium iodide (PI) to allow for the discrimination of necrotic (PI staining only), apoptotic (Annexin V only), and viable cells (no staining) and subjected to FACS analysis. The graph shows the fold increase in apoptosis over non–serum-starved control cells. Data are mean ± SD from three to five independent experiments. ***P < 0.001 compared with serum-deprived cells. (B) Dose response for protection of serum-deprived IMR-32 cells by MBP1 is shown. Serum deprivation took place in the presence or absence of MBP1 at the indicated concentrations. The graph shows the fold increase in apoptosis over non–serum-deprived control cells. Data are mean ± SD from three independent experiments. *P < 0.05 compared with serum-deprived cells.

 
We sought to delineate the mechanism of MBP1 effects on nerve cell survival in the serum deprivation model. Therefore, we determined whether ERK 1/2, which is involved in adhesion-mediated eosinophil protection of IMR-32 cells (17), and/or Akt, which is involved in apoptosis inhibition in other cells (21), was phosphorylated/activated or inhibited in nerve cells in response to MBP1 or EDN.

Western blot analysis confirmed that MBP1, in the presence of serum deprivation for 24 h, induced both ERK 1/2 and Akt phosphorylation compared with serum deprivation only (Figure 2). Pretreatment with the MEK/ERK inhibitor PD98059 inhibited ERK phosphorylation but not Akt phosphorylation (Figures 2A and 2B), while pretreatment with the PI3K/Akt inhibitor LY249002 inhibited Akt but not ERK 1/2 phosphorylation (Figures 2A and 2B), confirming that ERK and Akt are activated by MBP1 via independent pathways in IMR-32 cells.



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  		Figure 2. MBP1 induces phosphorylation of ERK and Akt in serum-deprived IMR-32 cells by independent pathways. Serum-deprived (24 h) differentiated IMR-32 cells (0.5 x 106) were incubated in the presence or absence of 1 µg/ml MBP1 (A and B). Cells were either not pretreated or were pretreated for 2 h with the MEK/ERK inhibitor PD98059 (50 µM) and/or with the PI3K/Akt inhibitor LY249002 (10 µM). Representative Western blots (A) of phospho-ERK (top panel) or total ERK 2 (bottom panel) and (B) of phospho-Akt (top panel) or total Akt 1 (bottom panel) in total protein extract (10 µg) from serum-deprived IMR-32 cells in the presence or absence of MBP1 (1 µg/ml) and the presence or absence of PD98059 and/or LY249002 pretreatment. Blots are representative of three similar experiments.

 
Time Course of Activation of ERK and Akt but Not p38 MAP Kinase by MBP1
We determined the pattern of activation of ERK and Akt by MBP1. Western blots of cytoplasmic protein from IMR-32 cells treated with MBP1 (1 µg/ml; Figure 3A) for various times from 10 min to 24 h were probed with an antibody specific to the dual phosphorylated form of ERK 1/2 and re-probed with anti-ERK 2 antibody to allow quantification of phospho-ERK. The intensity of the signal for the dual phosphorylated form of ERK 1/2 was quantified against that for ERK 2 at each time point (Figure 3A). MBP1 induced a 2- to 3-fold sustained increase in phosphorylation of ERK 1/2 over untreated levels in IMR-32 cells. This was evident from 10 min and lasted for at least 24 h of treatment (P < 0.05 for all time points; Figure 3A). By contrast, when Western blots were probed with an antibody to phosphorylated p38, no activation of this MAP kinase was evident (data not shown). Western blots of total protein from IMR-32 cells treated with MBP1 (1 µg/ml; Figure 3B) were probed with an antibody specific to the phosphorylated/activated form of Akt and subsequently stripped and re-probed with antibody to Akt1 to allow quantification of phospho-Akt. MBP1 induced an early increase in the relative expression of phosphorylated Akt above control, reaching a 2.5- to 3-fold increase after 10–20 min of treatment. This too was sustained for 24 h (Figure 3B).



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  		Figure 3. Eosinophil major basic protein (MBP1) induces sustained ERK and Akt phosphorylation in IMR32 cells. IMR32 cells (0.5 x 106) were incubated in the presence of 1 µg/ml MBP1 (A and B) for the indicated times. A shows ratio of pERK compared with total ERK 2 following IMR-32 cell treatment with 1 µg/ml MBP1 compared with untreated cells for the indicated times, whereas B shows ratio of phospho-Akt compared with total Akt 1 following IMR-32 cell treatment with 1 µg/ml MBP1 for the indicated times. Data are mean ± SD for three to four independent experiments. *P < 0.05 compared with untreated control cells.

 
MBP1 Induces Activation of NF-{kappa}B Transcription Factor in IMR-32 Nerve Cells
We have previously shown that eosinophils induce a rapid and sustained activation of NF-{kappa}B in IMR-32 cells (15). Thus, we examined whether eosinophil proteins could activate NF-{kappa}B in IMR-32 cells. IMR-32 cells were incubated with MBP1 (1 µg/ml) for various time periods. Electrophoretic mobility shift assay (EMSA) showed that MBP1 induced a strong and sustained activation of NF-{kappa}B in IMR-32 cells within 10 min of treatment, which lasted over 12 h but had declined by 24 h (Figure 4A). To determine whether NF-{kappa}B activation by MBP1 was dependent on ERK 1/2 and/or Akt activation, IMR-32 cells were pretreated with inhibitors of ERK 1/2 and/or Akt phosphorylation, before 10 min or 4 h incubation with MBP1. NF-{kappa}B activation in response to MBP1 was dependent on both ERK and Akt at 10 min (Figure 4B). However, by 4 h, NF-{kappa}B activation was dependent on ERK only, no longer on Akt (Figure 4C).



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  		Figure 4. Eosinophil major basic protein induces NF-{kappa}B activation in IMR32 cells. IMR32 cells (0.5 x 106) were incubated in the presence of 1 µg/ml MBP1 for the indicated times. (A) Representative EMSA and quantification of NF-{kappa}B in nuclear extract (10 µg) from IMR32 cells left untreated (0) or treated with 1 µg/ml MBP1 for the indicated times. Fold increase shown in graph 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. Data are mean ± SD from at least four independent experiments. *P < 0.05, significantly increased compared with untreated control cells. B and C show fold change in NF-{kappa}B activation following co-culture with MBP1 for 10 min or 4 h in cells either not pretreated (Control) or pretreated for 2 h with the MEK/ERK inhibitor PD98059 (50 µM) and/or with the PI3K/Akt inhibitor LY249002 (10 µM). *P < 0.05, pretreatment induces significant reduction compared with nonpretreated MBP1-stimulated cells.

 
Effect of EDN on Nerve Cell Signaling
As EDN did not effect protection of IMR-32 cells subjected to serum deprivation (Figure 1), the signaling effects of EDN were compared with those of MBP1. In terms of ERK 1/2 phosphorylation, although EDN (1 µg/ml) treatment induced some early phosphorylation of ERK 1/2 (Figure 5A), this did not become significantly increased over untreated levels of ERK 1/2 until 4 and 12 h of treatment, and declined to control levels by 24 h (Figure 5A), in contrast to the prolonged activation induced by MBP1 (Figure 3A). Also in contrast to MBP1, EDN induced a significant reduction of phospho-Akt by 30 min, with values returning to approximately baseline levels thereafter (Figure 5B). EDN did not induce phosphorylation of the p38 MAP kinase (data not shown). Thus MBP1 and EDN induced different levels and time courses of both Akt and ERK 1/2 activation. Furthermore, the pattern of NF-{kappa}B activation also differed between MBP1 and EDN. EDN-induced activation was maximal at 10 min, but declined by 1 h and did not rise significantly thereafter (Figure 5C). EDN-induced NF-{kappa}B activation was dependent on neither ERK nor Akt at 10 min of treatment (Figure 5D).



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  		Figure 5. Eosinophil-derived neurotoxin (EDN) signaling in IMR32 cells. IMR32 cells (0.5 x 106) were incubated in the presence of 1 µg/ml EDN for the indicated times. A shows quantification of ratio of phospho-ERK to total ERK. Data are mean ± SD for three to four independent experiments. *P < 0.05, significantly increased compared with untreated control cells. Insert shows representative Western blots of phospho-ERK (top panel) to total ERK (bottom panel). B shows quantification of ratio of phospho-Akt to total Akt. Data are mean ± SD for three to four independent experiments. *P < 0.05, significantly decreased compared with untreated control cells. Insert shows representative Western blots of phospho-Akt (top panel) to total Akt (bottom panel). C shows a representative EMSA and quantification for the effect of EDN on NF-{kappa}B activation. Data in graph are mean ± SD for three independent experiments. *P < 0.05, significantly increased compared with untreated control cells. D shows fold change in NF-{kappa}B activation following co-culture with EDN for 10 min or 4 h in cells either not pretreated (Control) or pretreated for 2 h with the MEK/ERK inhibitor PD98059 (50 µM) and/or with the PI3K/Akt inhibitor LY249002 (10 µM). Data are mean ± SD for three independent experiments.

 
MBP1-Induced Protection of IMR-32 Nerve Cells from Apoptosis Is Dependent on ERK 1/2 Activation
To determine whether the intracellular signaling pathways activated by MBP1 in IMR-32 cells were important in MBP1-induced protection of nerve cells from apoptosis, IMR-32 cells were pretreated or not with the MEK/ERK inhibitor PD98059 and/or the PI3K/Akt inhibitor LY249002. IMR-32 cells were then subjected to serum deprivation in the presence or absence of MBP1. MBP1-induced protection of IMR-32 cells from serum deprivation–induced apoptosis was dependent on ERK 1/2 activation, as pretreatment of the IMR-32 cells with PD98059 significantly reduced the protective effect of MBP1 in serum-deprived cells by ~ 70% (Figure 6). By contrast, pretreatment with the PI3K/Akt inhibitor LY249002 had no significant effect on MBP1-induced IMR-32 cell protection (Figure 6). Concomitant pretreatment with PD98059 and LY249002 resulted in a reduction in protection comparable to that caused by ERK inhibition alone (Figure 6).



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  		Figure 6. MBP1-induced protection from apoptosis of serum-deprived IMR-32 cells is dependent on ERK but not Akt activation. Serum-deprived (24 h) IMR-32 cells (0.5 x 106) were incubated in the presence or absence of 1 µg/ml MBP1. Cells were either not pretreated or were pretreated for 2 h with the MEK/ERK inhibitor PD98059 (50 µM) and/or with the PI3K/Akt inhibitor LY249002 (10 µM). Cells were subjected to staining for Annexin V and PI and FACS analysis. The graph shows the fold increase in apoptosis over non–serum-starved control cells. Data are mean ± SD from three independent experiments. *P < 0.05, **P < 0.01 compared with serum-deprived cells or to MBP1-stimulated serum-deprived cells as indicated by the bars.

 
MBP1 but Not EDN Induces Transcriptional Upregulation of the Antiapoptotic NF-{kappa}B–Activated Gene, Bfl-1
We have previously shown that eosinophils and eosinophil membranes induce adhesion-dependent protection of IMR-32 cells via upregulation of transcription of the NF-{kappa}B–mediated anti-apoptotic gene, bfl-1 (17). In this study we showed that MBP1 both protected IMR-32 cells from apoptotic stimuli and induced prolonged activation of the transcription factor NF-{kappa}B (Figures 1 and 4). Therefore, we examined the effect of stimulation of IMR-32 cells with MBP1 on expression of bfl-1 and compared it to the effects of EDN, which we did not observe to be protective. We observed that MBP1 induced transcriptional upregulation of Bfl-1 expression in serum-deprived cells by ~ 1.5-fold (Figures 7A and 7B). This upregulation was abolished by pretreatment of the IMR-32 cells with the MEK/ERK inhibitor PD98059, but not by the PI3K/Akt inhibitor LY249002 (Figures 7A and 7B). By contrast, EDN did not induce any upregulation of Bfl-1 transcription (Figure 7C).



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  		Figure 7. MBP1 induces transcriptional upregulation of the antiapoptotic gene Bfl-1 in serum-deprived IMR-32 cells: dependence on ERK phosphorylation. Serum-deprived (24 h) differentiated IMR-32 cells (0.5 x 106) were incubated in the presence or absence of 1 µg/ml MBP1 (A and B). Cells were either not pretreated or were pretreated for 2 h with the MEK/ERK inhibitor PD98059 (50 µM) and/or with the PI3K/Akt inhibitor LY249002 (10 µM). RNA was then extracted from cells and converted to cDNA, which was subjected to RT-PCR using primers specific to Bfl-1 or ß-actin. A shows fold increase in the ratio of Bfl-1 to ß-actin (top graph) induced by MBP1 in serum-deprived IMR-32 cells in the presence or absence of PD98059 and /or LY249002 as indicated. Data are mean ± SD from three independent experiments. **P < 0.01, pretreatment induces significant reduction compared with MBP1-stimulated serum-deprived cells. B shows representative ethidium bromide stained agarose gels of RT-PVR products for Bfl-1 and ß-actin. C shows no increase in the ratio of Bfl-1 to ß-actin induced by EDN in serum-deprived IMR-32 cells. Data are mean ± SD from three independent experiments.

 

   DISCUSSION
Top
 Abstract
 Introduction
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
References
 
In this article we have demonstrated that the eosinophil granule protein MBP1, but not EDN, at physiologically relevant concentrations (22, 23) protected IMR-32 cholinergic nerves from an apoptotic stimulus and that this protection was due to differences in the intracellular signaling induced by these proteins. Both MBP1 and EDN induced the MAP kinase ERK 1/2 and Akt, but over different time courses. MBP1 induced a prolonged activation of NF-{kappa}B, extending from 10 min to 12 h. Early MBP1-induced NF-{kappa}B activation was dependent on both ERK 1/2 and Akt, whereas later NF-{kappa}B activation was dependent on ERK 1/2 only. EDN-induced NF-{kappa}B activation was short-lived and independent of either ERK 1/2 or Akt. MBP1-induced ERK 1/2 activation mediated nerve cell protection from apoptosis induced by serum deprivation. Protection was associated with an ERK-dependent transcriptional upregulation of the NF-{kappa}B–regulated, antiapoptotic gene Bfl-1, which was not induced by the nonprotective EDN.

A protective effect of an eosinophil cationic granule protein on nerve cells is unexpected, as previous studies have primarily identified the cytotoxic properties of these proteins (4, 24, 25). However, not all studies have indicated that eosinophil granule proteins are toxic in vitro; for example, MBP1 and EPO inhibit mucin release from primary hamster tracheal surface epithelial cell cultures, they are not cytotoxic at concentrations up to 1 µM (26), and MBP1 activates a survival signal via PI3K/Akt in neutrophils (27). Other cationic proteins have been noted to induce apoptosis and/or necrosis in various cell types—for example, poly-L-arginine on airway epithelial cells (28). In the present study, we have shown that MBP1 increases expression of the antiapoptotic gene Bfl-1 in a nerve cell line at the transcriptional level. Thus, it is possible that the effect of eosinophil cationic granule proteins, in terms of toxicity, differ according to the cell type studied.

The observation that MBP1 is protective toward IMR-32 cells is consistent with our previous observations (17) in which we demonstrated an adhesion-dependent protection of IMR-32 cells co-cultured with whole eosinophils, which was mediated via activation of ERK. Here we extend these observations to show that MBP1, an eosinophil granule protein, can independently confer further protection on nerve cells, also via ERK. However, in this study, inhibition of ERK confers an ~ 70% reduction in protection of IMR-32 cells by MBP1 rather than completely abolishing protection. This implies that other pathways, which we have not identified here, are also important in MBP1-conferred nerve cell resistance to apoptosis. EDN, which is nonprotective, also induces ERK activation, but over a different, less sustained time course than MBP1. The whole eosinophil protective effect (17) is likely to be a combination of the adhesion-induced effects demonstrated by the use of eosinophil membranes (17) and of MBP1 effects demonstrated herein. We have previously shown that adhesion of eosinophils to nerve cells induces degranulation of the eosinophils (12). Adhesion of eosinophils to IMR-32 cells in itself induces a protective effect (17), and the consequent release of MBP1, the predominant eosinophil granular product in terms of molar concentration, from the degranulating eosinophils induces further protective effects.

NF-{kappa}B has been previously implicated in models of nerve cell survival and plasticity (21, 29). Our previous studies showed that NF-{kappa}B is rapidly and consistently activated in IMR-32 cells in response to eosinophil adhesion (15). In this study, we have demonstrated that NF-{kappa}B is activated both by MBP1 and by EDN over different time courses. MBP1 induces a very sustained activation of NF-{kappa}B in IMR-32 cells, lasting from ~ 10 min and declining by 24 h. Early activation of NF-{kappa}B is dependent on ERK and Akt activation, whereas later activation is only dependent on ERK. We further observed that MBP1 treatment of IMR-32 cells induces transcriptional upregulation of the NF-{kappa}B–activated antiapoptotic gene Bfl-1.

In contrast to MBP1, the nonprotective EDN induced no upregulation of Bfl-1. EDN induced an early but unsustained activation of NF-{kappa}B, but this fell to basal levels by 1 h. The brief period of EDN-induced NF-{kappa}B activation is observed before detection of EDN-induced ERK activation, and therefore is not reduced by ERK inhibition. Thus, as well as occurring over a different time course, the EDN-induced NF-{kappa}B activation is mediated by a different mechanism than that observed for MBP1. The level of NF-{kappa}B activation induced by EDN may be insufficient to support upregulation of Bfl-1 transcription. In addition, other inhibitory transcription factors or other intracellular signaling molecules may be activated by EDN.

In addition to the differences in ERK and NF-{kappa}B activation, EDN and MBP1 also have different effects on Akt phosphorylation in IMR-32 cells. MBP1 induces a prolonged phosphorylation of Akt from 10 min to 24 h, whereas EDN induces a reduction in Akt phosphorylation up to 30 min of treatment and thereafter has no effect on Akt activation. Perhaps unexpectedly, however, Akt does not appear to play a role in MBP1-induced activation of Bfl-1 or in protection of serum deprivation–induced apoptosis in IMR-32 cells. This may be related to the finding that although Akt is important in early activation of NF-{kappa}B by MBP1 in IMR-32 cells, it does not play a role in later NF-{kappa}B activation.

The mechanism of how MBP1 and EDN interact with nerve cells in seemingly different ways to induce intracellular signaling is uncertain. Various mechanisms can be considered, including interaction with specific nerve cell membrane receptors. For example, the neutrophil cathelicidin cationic antimicrobial granule protein LL-37 binds to the G protein–coupled formyl peptide-like receptor 1 (FPRL-1) to induce chemotaxis of monocytes, neutrophils, and subsets of T cells (30). However, LL-37 can also activate the MAP kinases ERK and p38 in monocytes independently of FPRL-1 (31), indicating that cationic granule proteins can exert their effects via multiple mechanisms. The eosinophil granule proteins could also, for example, transactivate tyrosine kinase receptors, analogously to LL-37–induced ERK-dependent transactivation of the epidermal growth factor receptor (EGFR) (32). Alternatively, one or all of the eosinophil proteins could become internalized in the nerve cells, in a manner reminiscent of polycation gene delivery systems currently under intensive development (reviewed in Ref. 33). In this way, the proteins could, over time, exert direct intracellular effects on, for example, the nucleus, mitochondria, or endoplasmic reticulum. Elucidation of these mechanisms and any potential applications of the antiapoptotic effects of MBP1 or other eosinophil granule proteins, for example in models of neurodegeneration, is one focus of our future investigations.

Peripheral nerves are often damaged in eosinophil-associated inflammatory diseases (34). Yet these diseases are also associated with remodeling of nerves, so that under the action of inflammatory substances such as neurotrophins (e.g., nerve growth factor) they adapt to become hyperresponsive (35). Remodeling of nerves occurs in many inflammatory diseases and is a mechanism of protection and adaptation (36). The results of this study suggest that eosinophils may play a role in neural remodeling, not alone by providing neurotrophins but also via MBP1 by protecting from harmful apoptotic stimuli. However, it is also possible that under pathologic conditions, neural remodeling and plasticity may not always be an advantage. These results should further our understanding of the complex interactions that occur between inflammatory cells such as eosinophils and nerve cells in inflammatory diseases.


   Footnotes
 
This work was supported by the Health Research Board of Ireland and The Wellcome Trust.

Conflict of Interest Statement: None of the authors have a financial relationship with a commercial entity that has an interest in the subject of this manuscript.

Received in original form February 8, 2005

Received in final form April 1, 2005


   References
Top
 Abstract
 Introduction
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 References
 

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