1Department of Pharmacology and Therapeutics, University of Liverpool, Liverpool L69 3GE, United Kingdom; 2Department of Medicine, Royal College of Surgeons in Ireland, Beaumont Hospital, Dublin 9, Ireland; 3Department of Environmental Health Sciences, School of Hygiene and Public Health, Johns Hopkins University, Baltimore, Maryland 21205; and 4Department of Dermatology, School of Medicine, University of Utah, Salt Lake City, Utah 84132
Submitted 2 April 2003 ; accepted in final form 3 June 2003
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ABSTRACT |
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inflammation; cell adhesion molecule; cholinergic nerve; major basic protein
We have also recently shown that primary cultures of parasympathetic nerves and differentiated IMR32 cholinergic neuroblastoma cells express intercellular adhesion molecule-1 (ICAM-1) and vascular cell adhesion molecule-1, to which eosinophils adhere (29). Engagement of these neural adhesion molecules by eosinophils leads to the generation of reactive oxygen species (ROS) within the nerve cells through an NADPH oxidase-dependent mechanism. Adhesion is also associated with activation and degranulation of eosinophils (18).
In animal models of inflammatory bowel disease, the associated increase in vagally mediated gut motility is followed by a period of diminished cholinergic nerve activity suggestive of a reduction in nerve function (6). We hypothesized that this may be due to the effect of eosinophil localization and adhesion to nerves. Eosinophils could influence nerve morphology or survival via the release of factors; some of these are already known to be toxic to epithelium (23) and muscle (30), whereas others, e.g., nerve growth factor (NGF), can induce cholinergic nerve growth (19). Alternatively, adhesion of eosinophils to neural cell adhesion molecules and the consequent induction of ROS could interfere with microtubule structure and thus induce neurite retraction (14).
In this study we have therefore further developed the in vitro cell culture model to investigate the consequences of eosinophil adhesion and degranulation for cholinergic nerve cell morphology.
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MATERIALS AND METHODS |
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Guinea pig eosinophil isolation and primary parasympathetic nerve cell culture. Guinea pigs were handled following the standards established by the USA Animal Welfare Acts set forth in National Institutes of Health guidelines and The Policy and Procedures Manual published by The Johns Hopkins School of Public Health Animal Care and Use Committee. Adult male Hartley guinea pigs (180-200 g) were anesthetized with xylazine and ketamine. A peritoneal lavage was performed by injecting 50 ml of PBS at 37°C through an 18-gauge catheter. Twenty milliliters of lavage fluid were layered onto 12 ml of Percoll solution (density 1.090 g/ml) and centrifuged at 400 g for 20 min. The pelleted cells were washed once with PBS and then placed in 1 ml of serum-free medium.
Parasympathetic nerves were isolated from trachealis muscles of the male Hartley guinea pigs, plated onto six-well plates, and stimulated for 48 h with IFN- (1,000 U/ml) and TNF-
(2 ng/ml) to induce the expression of cell adhesion molecules (29). Eosinophils (1 x 106/well) were then added to the nerves, and 24 h later the eosinophils were washed gently away with buffered saline, and the nerves were fixed and stained with dilute hematoxylin.
IMR32 cell culture. The IMR32 nerve cell line was maintained at 37°C in an atmosphere of 5% CO2 in proliferation medium consisting of DMEM with 10% (vol/vol) FCS, 100 U/ml penicillin, 100 µg/ml streptomycin, and 10 µg/ml gentamicin. Upon reaching confluence, cells were plated at a density of 1 x 104 cells/well in 48-well plates for subsequent measurements of neurite retraction or outgrowth and ROS and 1.5 x 105 cells/35-mm tissue culture dish for preparation of cell lysates for electrophoresis and Western blotting. After 24 h, proliferation medium was removed, and the cells were incubated for 4-6 days with differentiation medium [as for proliferation medium but with 2% (vol/vol) FCS plus 2 mM sodium butyrate] to induce neurite outgrowth and a cholinergic phenotype, before experimental treatment or addition of eosinophils.
Human eosinophil isolation. Eosinophils were prepared from the blood of healthy human volunteers as previously described (18). Briefly, 15 ml of blood were layered onto a 1.086 g/ml Percoll solution and centrifuged at 400 g for 20 min to yield a pellet containing granulocytes. Contaminating red blood cells were removed by two hypotonic lysis treatments. Granulocytes were then incubated with immunomagnetic anti-CD16 antibody-conjugated beads (1 µl of beads/106 cells) for 30 min at 4°C before depletion of CD16+ neutrophils through a magnetic separation column (MACS system, Miltenyi Biotec). Cytospins of the eluted cell population stained with Diff-Quik indicated the purity to be 98% eosinophils.
Coculture of eosinophils with differentiated IMR32 cells and measurement of neurite retraction. Eosinophils (10,000-50,000/well) were added directly to differentiated IMR32 cells in their culture medium. In some cases, the medium from coincubated eosinophils and IMR32 cells was removed after 2 h of coincubation, centrifuged to remove cells, and added to IMR32 cells. Conditioned medium was also taken from equivalent numbers of eosinophils stimulated for 2 h with 1 µM fMLP and then added to IMR32 cells. Nerves were then fixed in 4% (wt/vol) paraformaldehyde for 20 min at room temperature, stained with Coomassie blue stain [0.6% (wt/vol) Coomassie brilliant blue G in 10% (vol/vol) acetic acid, 10% (vol/vol) methanol, and 80% (vol/vol) PBS] for 10 min, and then washed once with PBS and twice with distilled H2O. Fixed and stained cells were viewed by light microscopy (Zeiss Axiovert 35M) linked by a video camera to a Kontron Vidas 2.0 image analyzer. The average length of neurites (in pixels) per cell was then calculated automatically from 200 randomly chosen cells as previously described (22). Where appropriate, drugs or enzyme inhibitors were added to IMR32 cells for various times before the addition of eosinophils or conditioned media. Experimental treatments were expressed as percentages of their appropriate control.
Coculture of eosinophils with undifferentiated IMR32 cells and measurement of neurite outgrowth. Eosinophils or their conditioned media were added to IMR32 nerve cells at the initiation of differentiation, and neurite outgrowth was subsequently measured as above. The length of neurites in cells maintained in proliferation medium (and therefore producing few neurites) was subtracted from the length of neurites in differentiating cells. The net neurite length in cells subjected to experimental treatment was then expressed as a percentage of that in control cells.
Measurement of neuronal apoptosis. The number of apoptotic IMR32 cells was assessed by Hoechst 33258 (bis-benzimide) staining. Cells were fixed in 4% (wt/vol) paraformaldehyde for 20 min at room temperature and then stained with Hoechst 33258 (5 µg/ml) in PBS for 10 min. Nuclear morphology was viewed under a fluorescence microscope (Zeiss Axiovert 35M), and apoptotic cells were defined as those that exhibited bright-condensed nuclei (17). Cells were counted in eight randomly chosen fields.
MTT assay. IMR32 nerve cells, eosinophils, or cocultures were incubated with MTT (1 mg/ml) at 37°C for 4 h. The formazan product formed was then solubilized in 20% (vol/vol) Triton X-100 for 30 min with constant agitation. Absorbance was then measured at 570 nm in a microplate spectrophotometer.
Measurement of neuronal ROS production. IMR32 nerve cells differentiated for 4-6 days were incubated with 10 µM DHR, an oxidant-sensitive probe (21), for 30 min at 37°C and then washed twice with fresh culture medium. The IMR32 cells were then pretreated with a number of inhibitors before the addition of 5 x 104 eosinophils. Alternatively, an equal number of paraformaldehyde-fixed eosinophils was used. The conversion of DHR to rhodamine 123 was detected in a microplate fluorescence reader (Bio-Tek FL600), with excitation filters at 485 ± 20 nm and emission filters at 530 ± 20 nm.
Gel electrophoresis and Western blotting. After coincubation of eosinophils with IMR32 nerve cells on 35-mm tissue culture dishes, the cultures were washed once with ice-cold PBS and lysed in 100 µl of Triton X-100 cell lysis buffer [1% (vol/vol) Triton X-100, 10 mM Tris · HCl, pH 7.4, 150 mM NaCl, 1 mM EGTA, 1 mM EDTA, 10 µg/ml leupeptin, 1 mM phenylmethylsulfonyl fluoride, 150 µM sodium orthovana-date, and 0.5 mM dithiothreitol]. Alternatively, eosinophils, nerves, or cocultures were treated with 4% paraformaldehyde for 20 min, and lysates were prepared in the same way. All lysates were then incubated at 4°C for 15 min and centrifuged at 12,000 g for 15 min at 4°C, and the supernatant was assayed for protein with the DC protein assay kit (Bio-Rad). Twenty micrograms of protein from each live nerve extract or a sample prepared from an equal number of cells for fixed cell lysates were boiled for 5 min in sample buffer [125 mM Tris, pH 6.8, 4% SDS, 10% glycerol, 0.006% (wt/vol) bromphenol blue, and 2% (vol/vol) 2-mercaptoethanol]. Proteins were separated by SDS-PAGE in a 10% polyacrylamide resolving gel overlaid with a 4% stacking gel, electrophoresed at 200 V for 45 min, and blotted onto nitrocellulose membranes in transfer buffer [25 mM Tris, 192 mM glycine, and 20% (vol/vol) methanol] at 30 V for 2 h.
The nitrocellulose membranes were then added to blocking buffer consisting of 1% (wt/vol) BSA in TBS/T [10 mM Tris, pH 7.5, 100 mM NaCl, and 0.1% (vol/vol) Tween 20] for 2 h and then incubated overnight at 4°C with mouse anti-human phosphotyrosine PY20 MAb (1:1,000) diluted in blocking buffer. After three 10-min washes in TBS/T, the membranes were incubated for 2 h at room temperature with goat anti-mouse IgG peroxidase conjugate (1:10,000) diluted in blocking buffer. Finally, the membranes were washed in TBS (3 x 10 min) and then exposed to Chemiglow West Chemiluminescent substrate for 5 min. The blots were then analyzed with an Alpha Innotech (San Leandro, CA) imaging system.
Alternatively, when the antiphospho-p38 MAP kinase antibody was used, membranes were incubated in blocking buffer consisting of TBS/T with 5% (wt/vol) nonfat dry milk for 1 h at room temperature. Membranes were then washed 3x 5 min in TBS/T at room temperature and then incubated overnight at 4°C with rabbit anti-human phospho-p38 MAP kinase antibody (1:1,000) in TBS/T containing 5% (wt/vol) BSA. After 3x 5-min washes in TBS/T, the membranes were incubated for 2 h at room temperature with goat anti-rabbit IgG alkaline phosphatase conjugate (1:10,000) diluted in blocking buffer. Membranes were then washed in TBS/T (3x 5 min) and exposed to CDP Star chemiluminescent substrate solution plus Nitro-Block II chemiluminescent substrate compound for alkaline phosphatase (19:1) for 5 min at room temperature. Blots were then exposed to X-OMAT light-sensitive film to obtain an image.
Statistical analysis. Values are expressed as means ± SE. The statistical significance of differences between control and treated samples was evaluated by one-way ANOVA; *P < 0.05, **P < 0.01, ***P < 0.001.
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RESULTS |
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The effect of eosinophils on neurite retraction was dose dependent (Fig. 3A). Under the conditions of coculture, eosinophils may mediate their effects either via contact or via the release of eosinophil products, since IMR32 nerve cells induce eosinophil degranulation (18). Therefore, we investigated whether adhesion per se or the consequent release of eosinophil products into the medium was responsible for neurite retraction. To generate conditioned medium containing eosinophil products, eosinophils were either incubated with IMR32 cells for 2 h to induce degranulation (18) or treated with 1 µM fMLP, and the medium was collected and transferred to differentiated IMR32 cells for 24 h. There was no significant change in the length of neurites in cultures treated with either set of media, suggesting that the effects of eosinophils on differentiated IMR32 nerve cells were a direct consequence of contact between the cells (Fig. 3, B and C).
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Kinase activation and ROS generation in IMR32 nerves mediate neurite retraction. It is established that the processes of neurite retraction and outgrowth can involve protein tyrosine kinase activity (2). To determine whether or not eosinophil-IMR32 cell interactions involved changes in protein tyrosine phosphorylation, we subjected cell extracts to Western blotting and probed them with an antiphosphotyrosine antibody (PY20). To ensure that any observed changes occurred in the nerves and not the eosinophils, we added paraformaldehyde-fixed eosinophils (which adhere as effectively as live eosinophils) to the IMR32 nerve cells. There was a time-dependent increase in the phosphorylation of a number of proteins in the molecular mass range 30-45 kDa (Fig. 4A). The phosphorylated protein bands were absent when lysates of paraformaldehyde-fixed eosinophils and/or paraformaldehyde-fixed nerves were electrophoresed, providing further evidence that the phosphorylation was occurring in the nerves (Fig. 4B). We also probed Western blots of cell extracts for the activated phosphorylated form of p38 MAP kinase, and this revealed a rapid, transient phosphorylation of the enzyme at 5 min (Fig. 4C).
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We have previously shown that eosinophil adhesion via CD18 leads to the production of ROS within the nerve cells, which can be prevented with the NADPH oxidase inhibitor DPI, and that IMR32 cells express the p47phox subunit of NADPH oxidase (18). Nerves were loaded with DHR, an oxidant-sensitive fluorescent probe, and cocultured with either live or paraformaldehyde-fixed eosinophils (Fig. 4D). Both samples of eosinophils induced a significant increase in fluorescence in the nerves, further indicating that the ROS were of neuronal origin and not diffusing from live eosinophils into the nerves. We also treated cocultures of nerves and eosinophils with kinase inhibitors. The increase in fluorescence associated with ROS production was significantly reduced by 72.52 ± 8.29% in the presence of genistein but not daidzein, the inactive analog of genistein (Fig. 4E). The p38 MAP kinase inhibitor SB-239063 also failed to prevent the increase in neuronal ROS induced by eosinophils. Activation of p38 MAP kinase was unaffected by DPI (data not shown).
These signaling pathways were then examined with regard to neurite retraction. Paraformaldehyde-fixed eosinophils were as effective as live eosinophils in reducing the length of neurites, providing further evidence that these effects were not mediated by released eosinophil factors and were a result of nerve signaling following eosinophil adhesion (Fig. 5A). When eosinophils were treated with MAb against CD18 (2 pg/ml) and the very late antigen (VLA)-4 peptide inhibitor ZD-7349 (10 µM) to prevent adhesion (29), there was a significant attenuation of neurite retraction (Fig. 5A). Treatment of the nerves with DPI (1 µM) also reduced the effect of eosinophils to induce neurite retraction (Fig. 5A), indicating a role for ROS. These ROS were of neuronal origin since treating eosinophils with DPI and washing out of the drug before addition of the eosinophils to the nerves did not reduce the eosinophil-mediated retraction (Fig. 5A). Both genistein and SB-239063 significantly reduced the effects of eosinophils on neurite retraction (Fig. 5B), suggesting at least two separate pathways were in operation: one involving ROS production via tyrosine kinases and the other mediated by p38 MAP kinase activation, which does not involve ROS. A possible downstream target of these pathways is Rho kinase (2). However, although the specific Rho kinase inhibitor Y-27632 (10 µM) induced neurite outgrowth on its own, it had no effect on the retraction induced by eosinophils (Fig. 5B).
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None of the kinase inhibitors used in these studies interfered with the adhesion of eosinophils to the nerves following 30-min coculture (data not shown), indicating that their effects were via downstream pathways within the nerve cells.
Eosinophil products inhibit neurite outgrowth. We also studied the process of neuronal differentiation in the presence of eosinophils or their conditioned medium by measuring the first 48 h of neurite outgrowth. Freshly plated IMR32 nerve cells were switched from proliferation medium to differentiation medium (in the presence or absence of eosinophils) or to differentiation medium that had been conditioned by eosinophils in contact with nerves or treated with fMLP. There was a significant dose-dependent reduction in the length of neurites in nerves directly in contact with eosinophils (Fig. 6A), in nerves treated with conditioned medium from nerve-stimulated eosinophils (Fig. 6B), and to a lesser extent in nerves treated with conditioned medium from fMLP-treated eosinophils (Fig. 6C). These results contrast with the failure of the same conditioned media to cause neurite retraction (Fig. 3).
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Eosinophils are known to release a number of highly charged cationic proteins such as MBP and eosinophil peroxidase that may affect nerve function (16). We treated IMR32 nerve cell cultures with both MBP and the synthetic cationic molecule poly-L-lysine and found there was a significant dose-dependent reduction in neurite outgrowth (Fig. 7, A and B) but no effect on neurite retraction (Fig. 7, C and D), which is consistent with our above observations on the effects of conditioned media from eosinophils. Neither compound was directly toxic to the IMR32 nerves (data not shown).
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DISCUSSION |
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Tissue sections taken from patients with inflammatory bowel disease and asthma show that eosinophils adhere to cholinergic nerves and degranulate in their presence (5, 9). The long-term effect of these processes on nerve function is unknown. However, studies in an animal model of inflammatory bowel disease have shown that acute inflammation causes a long-lasting reduction in the magnitude of cholinergic nerve-mediated smooth muscle contraction (6). We hypothesized that one mechanism for this inflammation-dependent reduction in cholinergic nerve function may be reflected in a change in the length of nerve processes.
Our data show that eosinophils caused a retraction of neurites of the cholinergic nerve cell line IMR32. To investigate whether this was the consequence of cell death or apoptosis (24, 26), we made measurements of MTT metabolism and cell apoptosis (using Hoechst 33258 staining). MTT metabolism was not affected by prolonged culture with eosinophils, and there was only a small increase in the number of apoptotic nerves. Therefore, eosinophils were not exerting their effect on neurite retraction via either apoptosis or necrosis; so retraction should not be considered a pathological response to eosinophils but, rather, a physiological remodeling process.
Previously, we have shown that eosinophils adhere to IMR32 nerve cells via specific integrin-cell adhesion molecule interactions and that this leads to the generation of neuronal ROS, which in turn leads to eosinophil degranulation (18). We found that blocking adhesion of eosinophils to nerves with an anti-CD18 MAb and a VLA-4 peptide inhibitor attenuated eosinophil-induced neurite retraction. Furthermore, the NADPH oxidase inhibitor DPI also attenuated neurite retraction, suggesting that adhesion-generated ROS were important. The role of ROS in neurite retraction has recently been explained by the observation that ROS induce reconfiguration of microtubules (14).
Activation of cell adhesion molecules results in their redistribution at the cell surface, an event often controlled by tyrosine phosphorylation of intracellular proteins (32). Thus we examined whether any nerve proteins were activated in this manner, by probing Western blots of extracts from nerve-eosinophil cocultures with a specific antibody directed against PY20 phosphotyrosine. In our experiments the proteins predominantly undergoing tyrosine phosphorylation in response to eosinophils were in the range 30-45 kDa, and our results indicate that the p38 MAP kinase was activated. Ligation of ICAM-1 on endothelial cells leads to the generation of ROS, in a process mediated via the p38 MAP kinase (33, 34). Activation of the p38 pathway has also been associated with the induction of neurite outgrowth (13), but conversely it can mediate ROS-induced apoptosis in neuroblastoma cells (25). Thus we investigated the role of p38 MAP kinase in eosinophil-mediated neurite retraction and ROS production. We found that, by using the p38 MAP kinase inhibitor SB-239063, we could inhibit neurite retraction but not ROS production. Equally, inhibiting ROS production had no influence on p38 MAP kinase activity. In contrast, the tyrosine kinase inhibitor genistein blocked both ROS production and neurite retraction. Together, these data suggest that two separate pathways involving ROS and p38 activation control the retraction of neurites.
The site at which genistein might be acting in our system remains uncertain. Because interaction of eosinophils with ICAM-1 on nerve cells leads to generation of ROS (18) and ICAM-1 is tyrosine phosphorylated (27), this may be where genistein is exerting its effects. Alternatively, it is possible that genistein prevents the activation of the NADPH oxidase complex by inhibiting the translocation of specific subunits such as p67phox or Rac (8, 10) and thus directly controls ROS formation. We and others have shown that nerves express subunits of the NADPH oxidase complex (18, 31). The GTPase family members Rho and Rac normally exist together in an inactive complex. After an activation signal, Rac dissociates from Rho and translocates to the membrane where it mediates the effects of the NADPH oxidase complex, leading to ROS production (7). This leaves Rho free to activate Rho kinase, which can lead to neurite retraction (1). It therefore occurred to us that genistein could act by preventing the downstream activation of Rho kinase (2). However, although the specific Rho kinase inhibitor Y-27632 induced neurite outgrowth in its own right (data not shown), it was unable to inhibit the effects of eosinophils on neurite retraction, indicating that Rho kinase was not involved.
From our experiments on differentiated nerves, we established that eosinophils induced neurite retraction through a process of cell adhesion but that the effect was independent of eosinophil degranulation, since medium from fMLP-conditioned eosinophils did not induce retraction. We have previously shown that fMLP-treated eosinophils release significant quantities of degranulation and activation products (18). However, the release of products from eosinophils did affect another aspect of nerve function, viz. neurite extension. This was demonstrated by the experiments in which we differentiated IMR32 cells in coculture with either eosinophils or their conditioned media. In both cases, there was a reduction in the ability of IMR32 cells to extend neurites during differentiation, indicating that released eosinophil products were at least partly responsible. This phenomenon may be specific to eosinophils since neutrophils had the opposite effect, promoting neurite outgrowth (data not shown). In particular, we found that eosinophil MBP could inhibit neurite outgrowth in a dose-dependent manner. The mechanism of this effect is unknown, but part of the action of MBP may be related to its cationic charge, since poly-L-lysine had a similar effect. In contrast to our observations, it has recently been shown that IgA-immune complex induces the release of NGF from eosinophils, which could promote neurite outgrowth (19). However, the study used NGF-dependent PC12 cells, and the eosinophils in our experiments were not activated in the same manner.
In conclusion, our observations have implications for the development and morphology of peripheral autonomic cholinergic nerves in conditions in which there is tissue eosinophilia (3). Although our in vitro model has identified some potential consequences of nerve-immune cell interactions, ultimately further investigation in vivo is required to determine their functional significance.
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DISCLOSURES |
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FOOTNOTES |
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The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
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REFERENCES |
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