Role of LPS-induced microfilament depolymerization in MIP-2 production from rat pneumocytes

Noritaka Isowa1 and Mingyao Liu1,2

1 Thoracic Surgery Research Laboratory, Division of Cellular and Molecular Biology, Toronto General Hospital Research Institute, University Health Network, and 2 Department of Surgery, University of Toronto, Toronto, Ontario, Canada M5G 2C4


    ABSTRACT
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ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

We have previously demonstrated that lipopolysaccharide (LPS) induces production of macrophage inflammatory protein-2 (MIP-2), a C-X-C chemokine for neutrophil recruitment and activation, in primary cultured rat lung alveolar epithelial cells. We have also demonstrated that LPS depolymerizes microfilaments in rat alveolar epithelial cells. To determine whether the polymerization status of microfilaments affects LPS-induced MIP-2 production, we treated rat alveolar epithelial cells with cytochalasin D (CytoD), a microfilament-disrupting agent, before and during LPS stimulation. A lower concentration (0.1 µM) of CytoD inhibited LPS-induced MIP-2 production without affecting microfilament polymerization. In contrast, LPS-induced MIP-2 production was enhanced by a higher concentration (10 µM) of CytoD, which disrupted the filamentous structure of actin. Jasplakinolide (1 nM to 1 µM), a polymerizing agent for microfilaments, decreased LPS-induced MIP-2 secretion. Jasplakinolide (1 µM) also blocked LPS-induced depolymerization of microfilaments. These results suggest that, in alveolar epithelial cells, LPS-induced MIP-2 production is at least partially regulated by microfilament depolymerization.

cytokines; chemokines; lipopolysaccharide; macrophage inflammatory protein-2


    INTRODUCTION
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ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

THE RESPIRATORY TRACT is an accessible portal for potentially infective microorganisms and noxious substances in the inhaled air. Thus lung defense mechanisms are crucial for the effective removal of microbes and other debris from the conducting airways and alveoli (32, 45). The alveolar epithelium is an important component in host defense. It functions as a barrier to prevent the invasion of pathogens. Type II pneumocytes produce lung surfactant that can enhance the function of immune cells in the alveoli. Surfactant proteins are also important mediators of host defense (11, 27, 52, 53). Recently, it has been found that lung alveolar epithelial cells may also function as sensors for the invasion of microorganisms and other noxious agents by producing cytokines and chemokines (46). The interaction between leukocytes and pulmonary parenchymal cells, including alveolar epithelial cells, via cytokine signaling mediates innate and acquired immunity in lung antimicrobial host defense (47, 48).

Cytokines are extracellular signaling proteins secreted by cells and have the ability to modify the behavior of other adjacent cells (26). Chemokines are chemotactic cytokines for leukocyte recruitment and activation at the sites of infection or tissue injury (2). The role of chemokines in mediating lung host defense has been the subject of several reviews (47, 48). They are also important mediators in inflammation (2-4, 15, 31, 47).

Neutrophil infiltration into the alveolar space is mainly mediated by C-X-C chemokines such as interleukin (IL)-8 and its rodent homologue macrophage inflammatory protein (MIP)-2 (3, 15, 31). MIP-2 is an important mediator in host defense (17-19, 22) and acute inflammation in the lung (20, 21, 41) by mediating recruitment and activation of neutrophils in the alveolar space (20-22, 41). Gene expression of MIP-2 has been reported from rat lung epithelial cell lines (16). Xavier et al. (54) confirmed that primary cultured rat pneumocytes are a source of MIP-2. Lipopolysaccharide (LPS) is a component of the gram-negative bacterial cell wall, which is known to induce inflammatory responses in many cell types. Isowa et al. (24) have recently found that both basal and LPS-induced MIP-2 secretion in rat pneumocytes is through the endoplasmic reticulum (ER)-Golgi pathway in a constitutive fashion. Because microtubules and associated motor proteins can facilitate the selective delivery of transport intermediates between the ER and the Golgi and the delivery of secretory vesicles from the Golgi to the plasma membrane (9, 29), Isowa et al. (25) examined the role of LPS-induced microtubule depolymerization in MIP-2 production. Further depolymerization of microtubules with colchicine or nocodazole enhanced LPS-induced MIP-2 production, whereas paclitaxel, a microtubule-stabilizing agent, partially inhibited LPS-induced MIP-2 production (24). These results suggest that the microtubule system is involved in LPS-induced MIP-2 production, but microtubule-independent mechanisms may also exist.

In addition to inducing microtubule depolymerization, LPS also reduced the polymerization of microfilaments in rat pneumocytes that is involved in LPS-induced tumor necrosis factor (TNF)-alpha production (25). Therefore, in this study, we examined the hypothesis that microfilament depolymerization may also participate in the regulation of LPS-induced MIP-2 production from rat pneumocytes.


    MATERIALS AND METHODS
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ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

Reagents. LPS (Escherichia coli), cytochalasin D (CytoD), rat IgG, and rabbit skeletal muscle actin were purchased from Sigma (St. Louis, MO). DMEM, fetal bovine serum (FBS), and gentamicin were purchased from GIBCO BRL (Mississauga, ON). Porcine pancreatic elastase was purchased from Worthington Biochemical (Freehold, NJ). Pentobarbital sodium was purchased from Bimeda-MTC Pharmaceuticals (Cambridge, ON). Jasplakinolide (Jasp) and rhodamine-phalloidin were purchased from Molecular Probes (Eugene, OR).

Rat alveolar epithelial cell isolation and culture. Alveolar type II cells were obtained with the method of Dobbs (12) as described previously (25, 35, 54). Briefly, male adult Sprague-Dawley rats (Harlan, Indianapolis, IN) weighing ~250 g were anesthetized by intraperitoneal injection of pentobarbital sodium (100 mg/kg body wt) and killed by transection of the descending aorta and inferior vena cava. Alveolar epithelial cells were separated from the alveolar basement membrane by incubation of the isolated lung tissue with porcine pancreatic elastase. Contaminating alveolar macrophages were removed by differential adherence to rat IgG-precoated petri dishes. The number and viability of fresh cell suspensions were counted after they were stained with crystal violet and trypan blue exclusion. The viability of the fresh alveolar epithelial cells was >95%.

Cells were cultured in DMEM containing 10% (vol/vol) FBS and 12.5 µg/ml of gentamicin. In most experiments, the cell suspension (106 cells · ml-1 · well-1) was seeded in 24-well culture plates (Corning Glass Works, Corning, NY) and maintained at 37°C in 5% CO2. The cells were also cultured in six-well plates to determine the relative amount of filamentous actin (F-actin) or in 96-well plates for the cytotoxicity assay. For immunofluorescent staining, the cells were seeded on four-well plastic Lab-Tek chamber slides (Nunc, Naperville, IL). To reduce the contamination of alveolar macrophages in the primary culture, the culture medium was changed daily for 2 days before LPS treatment. As McRitchie et al. (35) have reported recently, this maneuver reduced the number of macrophages to undetectable levels as confirmed by cell surface ectoenzyme-alkaline phosphatase staining or by immunofluorescent staining with a monoclonal antibody for CD45, a surface marker for macrophages and leukocytes. The purity of the alveolar epithelial cells in the culture system was confirmed with phase-contrast microscopy and immunofluorescent staining with anti-cytokeratin and anti-surfactant proprotein C antibodies (specific markers for epithelial cells and type II pneumocytes, respectively) as McRitchie et al. (35) recently described.

Cytotoxicity assays. The cytotoxic effects of LPS, CytoD, and Jasp were examined by simultaneous double staining with fluorescein diacetate and propidium iodide as previously described (25, 35, 54). The viability of the cells in all groups in this study was found to be comparable to that of the control group without LPS or drugs. No cytotoxic effect was observed under experimental conditions in this study.

Filamentous actin staining and confocal microscopy. One milliliter of freshly isolated lung cell suspension (5 × 105 cells/ml) was seeded in each well of four-well Lab-Tek chamber slides, and the culture medium was changed daily for 2 days. After treatment with various reagents, the slides were washed three times with cold PBS and fixed in 3.7% formaldehyde for 10 min at room temperature followed by a wash with PBS. The cells were permeabilized with 0.1% Triton X-100 in 100 mM PIPES buffer (pH 6.9) containing 1 mM EGTA and 4% polyethylene glycol 8000 for 3 min at room temperature followed by a wash with PBS. For the localization and structure of F-actin, the fixed and permeabilized cells were stained with rhodamine-phalloidin (1:40 in PBS) for 30 min in the dark. After a wash with PBS, the slides were mounted with an antifading reagent (SlowFade, Molecular Probes). Confocal microscopy was performed with a confocal laser-scanning microscope (MRC-600, Bio-Rad, Mississauga, ON) equipped with a krypton-argon laser. In each experiment, cells with different treatments were cultured in different wells on the same chamber slide and processed simultaneously under the same conditions for comparison. Each experiment was repeated at least three times with cells isolated from separate cultures. The laser power, magnification, and other conditions were fixed for all the slides in each experiment. Multiple fields were photographed to ensure reproducibility. The images were collected and analyzed by different people in a blinded fashion.

Measurement of MIP-2. MIP-2 concentrations in the culture medium were measured in duplicate or triplicate with ELISA kits (BioSource, Camarillo, CA) following the manufacturer's instructions. The optical density of each well was read at 450 nm with an NM600 microplate reader (Dynatech Laboratories, Chantilly, VA). The detection range of the MIP-2 kit was 10-640 pg/ml. The final concentration was calculated by comparing the optical density readings against a standard curve.

Extraction and gel electrophoresis analysis of F-actin. Cells were cultured in six-well plates (4 × 106 cells/well, 3 wells/group) in 10% FBS-DMEM. Forty-eight hours after isolation, the cells were treated with and without Jasp in 10% FBS-DMEM for 4 h. After two washes with ice-cold PBS, the cells were lysed by adding 200 µl/well of a 1% Triton X-100 solution containing 1 mM EGTA, 50 mM Tris (pH 7.2), 1 mM benzamidine, 0.1 mM Na3VO4, 250 µg/ml of leupeptin, 25 µg/ml of aprotinin, and 0.1 mM phenylmethylsulfonyl fluoride and were held on ice for 20 min. The cell lysates from each group were pooled and centrifuged at 14,000 rpm for 5 min. The supernatants (600 µl in total) were removed. The Triton-insoluble pellets were washed with cold PBS, centrifuged again, and resuspended in 40 µl of SDS sample buffer containing 60 mM Tris (pH 8.0), 5% (vol/vol) beta -mercaptoethanol, 2% (wt/vol) SDS, 0.0025% (wt/vol) bromphenol blue, and 10% (vol/vol) glycerol. All samples were boiled for 10 min, and 15 µl of each sample were subjected to SDS-PAGE (10% polyacrylamide gel) (36). The gels were stained with Coomassie blue and destained in methanol-water-acetic acid (2:7:1 by volume). Actin protein was identified by its molecular mass (43 kDa) by comparison with purified actin from rabbit skeletal muscle as a positive control and by Western blotting (25, 36).

RNA extraction and semiquantitative RT-PCR. Cells were cultured in six-well plates (4 × 106 cells/well) in 10% FBS-DMEM. Forty-eight hours after cell isolation, the cells were treated with CytoD or Jasp for 2 h followed by stimulation with LPS (10 µg/ml) in 10% FBS-DMEM for 4 h with the agents tested. The medium was removed, and the cells were washed twice with ice-cold PBS. RNA was extracted and semiquantitative RT-PCR was performed as previously described (24, 54). The forward PCR primer for beta -actin was 5'-GTGGGCCGCTCTAGGCACCAA-3', and the reverse primer was 5'-CTCTTTGATGTCACGCAGGATTTC-3'. The forward PCR primer for MIP-2 was 5'-ATGCTGTACTGGTCCTGCTCCT-3', and the reverse primer was 5'-CTTCAGGGTTGAGACAAACTTCA-3'.

Statistical analysis. All experiments were carried out with materials collected from at least three separate cell cultures in duplicate or triplicate. All data are expressed as means ± SE from separate measurements and were analyzed with SigmaStat for Windows, version 1.0 (Jandel, San Rafael, CA). Comparison of more than two groups was carried out with two-way analysis of variance followed by Student-Newman-Keuls test, with significance defined as P < 0.05.


    RESULTS
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

Biphasic effects of CytoD on LPS-induced MIP-2 release from alveolar epithelial cells. Isowa et al. (25) have previously shown that 10 µg/ml of LPS from Escherichia coli reduced microfilament polymerization, and the same dosage of LPS also maximally stimulated MIP-2 production by alveolar epithelial cells within 4 h (54). CytoD is a commonly used microfilament-disrupting agent; Isowa et al. (25) have recently shown that 1 or 10 µM of CytoD enhanced LPS-induced TNF-alpha production from rat pneumocytes. To clarify the role of the polymerization status of microfilaments in MIP-2 production, the cells were pretreated with various concentrations of CytoD (1 nM to 10 µM) for 2 h and then challenged with LPS (10 µg/ml) for 4 h in the presence of CytoD. Treatment of alveolar epithelial cells with CytoD (1 nM to 10 µM) alone for 6 h did not change the basal levels of MIP-2 in the culture medium (Fig. 1). LPS-induced MIP-2 release was inhibited by 0.1 µM CytoD but was increased by 10 µM CytoD (Fig. 1).


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Fig. 1.   Biphasic effects of cytochalasin D (CytoD) on lipopolysaccharide (LPS)-induced macrophage inflammatory protein (MIP)-2 release from alveolar epithelial cells. Cells (1 × 106/ml) were cultured with 10% fetal bovine serum (FBS)-DMEM, treated with indicated concentrations of CytoD at 37°C for 2 h, and then stimulated with and without (-) LPS for 4 h in the presence of CytoD. Results are expressed as a percentage of the group treated with 1 nM CytoD and LPS. Values are means ± SE; n = 9 samples. P < 0.0001 as analyzed by 2-way ANOVA. * P < 0.05 vs. the group treated with 1 nM CytoD and LPS as determined by Student-Newman-Keuls test.

Low concentration of CytoD (0.1 µM) did not change the polymerization of F-actin in alveolar epithelial cells. To determine whether the decrease in LPS-induced MIP-2 release from the cells treated with a low concentration (0.1 µM) of CytoD is related to the polymerization status of microfilaments, we examined the effect of 0.1 µM CytoD on microfilaments in alveolar epithelial cells with fluorescent staining and confocal microscopy. Treated with this concentration of CytoD from 15 min to 4 or 24 h, the intensity of fluorescence of F-actin did not show significant changes compared with that in nontreated cells (see Figs. 5A and 6A as examples). Fine microfilament stress fibers were clearly seen in the cells at all time points tested (Fig. 2, A, C, and E). The intensity of F-actin staining in LPS-stimulated cells (Fig. 2, B, D, and F) was decreased compared with that in cells without LPS stimulation (Fig. 2, A, C, and E) at the same time points. However, the structure of the microfilament stress fibers was retained very well in LPS-treated cells. Therefore, the inhibitory effect of a low concentration of CytoD on LPS-induced MIP-2 release seems not to be related to the status of F-actin polymerization.


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Fig. 2.   Effect of LPS and a low concentration of CytoD (0.1 µM) on filamentous (F) actin in alveolar epithelial cells. Cells were treated with CytoD at 37°C for 2 h and then incubated without (A, C, and E) and with (B, D, and F) LPS (10 µg/ml) for 15 min (A and B), 4 h (C and D), or 24 h (E and F) in the presence of CytoD. Cells were stained with rhodamine-phalloidin (for F-actin) and examined with a confocal microscope. The intensity and structure of F-actin staining was not affected by CytoD at each time point tested (A, C, and E), and LPS treatment reduced the staining intensities (B, D, and F).

A high concentration of CytoD (10 µM) enhanced the staining intensity, with a disrupted structure of F-actin filaments in alveolar epithelial cells. We also examined the effect of 10 µM CytoD on microfilaments in alveolar epithelial cells with fluorescent staining and confocal microscopy at different time points. Treated with CytoD (10 µM), the fluorescence intensity of F-actin in cells was much brighter than that in untreated cells (see Figs. 5A and 6A) or cells treated with 0.1 µM CytoD (Fig. 2) in a time-dependent manner. The intensity of F-actin staining in LPS-stimulated cells (Fig. 3, B, D, and F) was much lower compared with that in cells without LPS stimulation (Fig. 3, A, C, and E) at each time point tested. In the cells treated with 10 µM CytoD for 15 min (Fig. 3, A and B) and 4 h (Fig. 3, C and D), F-actin fibers were collapsed and aggregated. In the cells treated with CytoD for 24 h (Fig. 3, E and F), fibrous structures were slightly recovered. However, it was clear that a high concentration of CytoD disrupted the F-actin structure, whereas the major effect of LPS on F-actin was to reduce the intensity of microfilaments. CytoD may cause severing of the microfilaments and thus increase the local density of chopped actin filaments (10, 33).


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Fig. 3.   Effect of LPS and a high concentration of CytoD (10 µM) on F-actin in alveolar epithelial cells. Cells were treated with CytoD at 37°C for 2 h and then incubated without (A, C, and E) and with (B, D, and F) LPS (10 µg/ml) for 15 min (A and B), 4 h (C and D), or 24 h (E and F) in the presence of CytoD. Cells were stained with rhodamine-phalloidin and examined with a confocal microscope. Fluorescent intensity of microfilaments was increased by the high concentration of CytoD in a time-dependent manner, but the filamentous structures were disrupted. The intensity of F-actin staining was decreased in LPS-treated cells (B, D, and F) compared with that in CytoD-treated cells (A, C, and E) at each time point tested.

Jasp inhibited LPS-induced MIP-2 release from alveolar epithelial cells in a dose-dependent manner. To further determine the effect of microfilament polymerization status on LPS-induced MIP-2 production, we used a membrane-permeable cyclodepsipeptide, Jasp, to treat the cells. Jasp has been show to induce actin polymerization and stabilize preexisting actin filaments (7, 23, 28, 33). Isowa et al. (25) have shown that Jasp inhibited LPS-induced TNF-alpha release from alveolar epithelial cells (25). To determine whether Jasp has similar inhibitory effects on LPS-induced MIP-2 release in alveolar epithelial cells, the cells were treated with varying concentrations (0.1 nM to 1 µM) of Jasp for 2 h and were then challenged with LPS (10 µg/ml) for 4 h. Treatment of cells with Jasp for 6 h did not change the basal levels of MIP-2 in the culture medium (Fig. 4). LPS-induced MIP-2 in the culture medium was inhibited by Jasp in a dose-dependent manner, with a maximal inhibitory effect of ~60% (Fig. 4).


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Fig. 4.   Jasplakinolide (Jasp) inhibited LPS-stimulated MIP-2 release from alveolar epithelial cells. Cells (1 × 106/ml) were cultured with 10% FBS-DMEM, treated with indicated concentrations of Jasp at 37°C for 2 h, and then stimulated with and without LPS (10 µg/ml) for 4 h in the presence of Jasp. Results are expressed as a percentage of the group treated with 0.1 nM Jasp and LPS. Values are means ± SE; n = 6 samples. P < 0.0001 as analyzed by 2-way ANOVA. * P < 0.05 vs. the group treated with 0.1 nM Jasp and LPS as determined by Student-Newman-Keuls test.

Jasp stabilized microfilaments in rat alveolar epithelial cells in a dose-dependent manner. Jasp has been shown to polymerize microfilaments in other cell types (28, 44). We examined whether Jasp at the concentrations we used could enhance polymerization of microfilaments in alveolar epithelial cells with fluorescent staining and confocal microscopy. F-actin bundles in the cells treated with Jasp were thicker and denser in a dose-dependent manner compared with those in control cells (Fig. 5, A-C). To further confirm increased polymerization of the microfilaments in Jasp-treated cells, we isolated F-actin from alveolar epithelial cells by Triton extraction (25, 36). After high-speed centrifugation, globular actin (G-actin) can be dissolved in the Triton-soluble fraction, whereas F-actin is mainly present in the Triton-insoluble pellets. Because actin is very abundant in the cell lysates, it can be separated by gel electrophoresis and displayed by Coomassie blue staining as a single band (25). Treatment with Jasp increased insoluble F-actin in alveolar epithelial cells (Fig. 5D).


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Fig. 5.   Jasp treatment increased microfilament polymerization in rat alveolar epithelial cells. Cells were incubated with and without Jasp for 6 h. A-C: cells were stained with rhodamine-phalloidin and examined with a confocal microscope. A: control cells have fine bundles of microfilaments. After Jasp (B: 0.1 µM; C: 1 µM) treatment, thicker and more prominent bundles of microfilaments can be seen. D: Triton-insoluble fractions of alveolar epithelial cell extracts were subjected to SDS-PAGE. A 42-kDa band representing actin was intensified in a Jasp dose-dependent manner.

Jasp inhibited LPS-induced depolymerization of microfilaments in alveolar epithelial cells. To determine whether Jasp could inhibit LPS-induced depolymerization of microfilaments, the cells treated with Jasp followed by LPS stimulation were stained with rhodamine-phalloidin and examined with confocal microscopy. The intensity of F-actin staining was decreased in primary cultured alveolar epithelial cells with LPS (10 µg/ml) stimulation (Fig. 6B) compared with that in untreated control cells (Fig. 6A), which was consistent with the previous observation by Isowa et al. (25). In the cells pretreated with Jasp, LPS also reduced the intensity of microfilament bundles (Fig. 6, D vs. C). In addition, there were intensely stained clumps at the peripheral sites in the cytoplasm of the cells treated with Jasp followed by LPS stimulation (Fig. 6D). Comparison between Fig. 6, B and D, showed that Jasp treatment increased the intensity in the network of microfilament bundles in the cells.


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Fig. 6.   Jasp inhibited LPS-induced depolymerization of microfilaments in alveolar epithelial cells. Cells were cultured in 10% FBS-DMEM and treated without (A and B) and with (C and D) Jasp (1 µM) at 37°C for 2 h followed by stimulation with LPS (10 µg/ml; B and D) for 4 h. Cells were fixed, permeabilized, stained with rhodamine-phalloidin, and observed with a confocal microscope. The intensities of F-actin staining in the cells stimulated with LPS (B and D) were decreased compared with those in the cells without LPS stimulation (A and C). Treatment with Jasp increased the intensities of F-actin staining in both cells with (D) and without (C) LPS stimulation.

CytoD and Jasp did not affect LPS-induced MIP-2 gene expression in alveolar epithelial cells. We then measured mRNA levels of MIP-2 by semiquantitative RT-PCR. The cells were pretreated with CytoD (0.1 or 10 µM) or Jasp (1 µM) for 2 h and then stimulated with LPS (10 µg/ml) for 4 h. The RNA extracted was analyzed with semiquantitative RT-PCR as previously described (25). The LPS-induced increase in MIP-2 mRNA was not affected by these substances (Fig. 7).


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Fig. 7.   Polymerizing or depolymerizing microfilaments did not affect LPS-induced MIP-2 mRNA expression in pneumocytes. Cells were cultured with 10% FBS-DMEM, treated with Jasp (1 µM) or CytoD (0.1 or 10 µM) at 37°C for 2 h, and then stimulated with LPS (10 µg/ml) for 4 h in the presence of these agents. RNA was extracted, and mRNA levels were analyzed by semiquantitative RT-PCR. Arbitrary density units of MIP-2 mRNA levels were normalized against the density of beta -actin mRNA levels from the same RT reaction of each sample. Results are expressed as means ± SE; n = 3 samples. LPS significantly increased MIP-2 mRNA levels compared with those in untreated control cells, which were not affected by Jasp or CytoD. P < 0.0001 as analyzed by 2-way ANOVA. * P < 0.05 vs. other groups as determined by Student-Newman-Keuls test.


    DISCUSSION
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ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

LPS-induced MIP-2 production from rat pneumocytes is partially mediated through microfilaments. The present study was performed to determine the role of the polymerization status of microfilaments in LPS-induced MIP-2 production by alveolar epithelial cells. We found that a high concentration (10 µM) of CytoD disrupted microfilament structure and enhanced the LPS-induced MIP-2 production. In contrast, Jasp increased the intensity of fluorescent staining of the microfilaments and the amount of actin protein present in the F-actin fraction and inhibited the LPS-induced decrease in fluorescent staining of the microfilaments as well as the LPS-induced MIP-2 production in a dose-dependent manner. Although a lower concentration (0.1 µM) of CytoD inhibited LPS-induced MIP-2 secretion from alveolar epithelial cells, the inhibitory effect was not related to the polymerization status of F-actin because the microfilament structures were not affected by this dose of CytoD. Taken together, these data suggest that LPS-induced depolymerization of microfilaments is involved in the LPS-induced MIP-2 production in alveolar epithelial cells. Because these substances did not affect LPS-induced MIP-2 gene expression, we speculate that the role of microfilaments in LPS-induced MIP-2 production is mainly at the posttranscriptional levels, such as its synthesis and secretion.

Role of microfilament depolymerization in mediating cytokine productions from lung alveolar epithelial cells. Because the microfilament is an important component for cell locomotion (8), many studies have focused on immune cells with motile ability, such as neutrophils or macrophages. Those cells remodel microfilaments in response to both intracellular and extracellular signals and transfer molecules via the cytoplasm membrane (1). Treatment of macrophages (43) or monocytes (13) with LPS increases the polymerization of microfilaments by reorganization of F-actin, which may be related to the motility of these immune cells. In macrophages, CytoD blocked LPS-induced TNF-alpha gene expression and/or protein synthesis (42).

In contrast, in alveolar epithelial cells, LPS decreased the polymerization of microfilaments (25), which could be an important regulatory mechanism for secretion from these cells. It has been shown that actin depolymerization is involved in the release of pulmonary surfactant from rat alveolar epithelial cells (5, 39, 40). Stabilization of the microfilaments in rat alveolar epithelial cells prevented surfactant secretion (40). Intratracheal instillation of LPS to rats increased the secretion of surfactant proteins (34). McRitchie et al. (35) have recently demonstrated that primary cultured rat alveolar epithelial cells could produce TNF-alpha in response to LPS stimulation, which was inhibited by Jasp and enhanced by CytoD (25). In the present study, LPS-induced MIP-2 release from alveolar epithelial cells was similarly affected by Jasp and the higher concentration of CytoD. These results suggest that LPS-induced microfilament depolymerization is very important for the secretion of surfactant and cytokines from alveolar epithelial cells. Because both surfactant and cytokines are important mediators in host defense in the lung (11, 27, 52, 53), the coordinated secretion of surfactant and cytokines induced by LPS may be a very important step in the regulation of host defense.

Disassembly of actin filaments in various kinds of nonimmune cells has been found to play a significant role in secretion (33, 51). For example, the cortical actin network has been shown to create a barrier preventing exocytosis of zymogen granules in pancreatic acinar cells (38). The secretory granules of pancreatic acinar cells become "coated" with F-actin before membrane fusion in the final step of exocytosis (50). The cytoskeletal structure endows the cell with a very crowded cytoplasm, and the integrated organization of the cytoskeleton and membrane systems may provide an important barrier to the free diffusion of secretory vesicles (6, 30). In the resting condition, the actin cytoskeleton localized under the plasma membrane may prevent secretory granules from reaching their exocytotic destination. On stimulation, microfilaments may be disassembled or rearranged to allow secretory granules to reach the site of exocytosis (51). LPS-induced depolymerization of microfilaments, especially the cortical filament network, may facilitate secretion of TNF-alpha and MIP-2 as well as of other secretory proteins through similar mechanisms.

Role of cytoskeleton in LPS-induced cytokine production in rat pneumocytes. Isowa et al. (24) have found that the microtubule system is also involved in LPS-induced MIP-2 production in primary cultured rat alveolar epithelial cells. Preventing depolymerization of microtubules with paclitaxel or further enhancing microtubule depolymerization with colchicine or nocodazole inhibited or increased LPS-induced MIP-2 production, respectively (24). However, these effects were incomplete (24). In this study, we demonstrated that LPS-induced MIP-2 production was also influenced by microfilament-disrupting or -stabilizing agents. Therefore, both microtubule and microfilament systems appear to be involved in LPS-induced MIP-2 production from alveolar epithelial cells. Interestingly, both microtubule- and microfilament-regulating agents affected MIP-2 protein production but did not significantly change the steady-state mRNA levels of MIP-2. It seems that the major effects of the cytoskeletal system on LPS-induced MIP-2 production are at posttranscriptional levels. LPS-induced depolymerization of microtubules may affect intracellular transport of cytokine molecules between the ER and the Golgi or change the secretion path (24).

In addition to LPS stimulation, cytokines produced from lung cells could be regulated by other factors through the cytoskeleton. For example, mechanical ventilation is known to induce lung injury (14) and provoke proinflammatory cytokine production (49). Mourgeon et al. (37) have recently demonstrated that mechanical stretch, which simulates injurious ventilation in vitro, enhanced LPS-induced MIP-2 secretion from fetal rat lung cells in a force- and frequency-dependent manner. The direct effect of mechanical stretch on the cells is to apply physical forces to deform the cytoskeletal structures.

As mentioned earlier, the cytoskeleton is also involved in LPS-induced TNF-alpha production from lung alveolar epithelial cells (25). However, the involvement of cytoskeletal components is different from that of MIP-2 production. Both microtubule and microfilament systems are involved in LPS-induced MIP-2 production, but only microfilaments are related to TNF-alpha production (25). In addition, we did not see an inhibitory effect of the lower concentration of CytoD on TNF-alpha production (25). Therefore, the detailed mechanisms, which regulate the production of these two cytokines, merit further investigation.


    ACKNOWLEDGEMENTS

We acknowledge the technical assistance of Xiao-Hui Bai, Xiao-Ming Zhang, and Dr. Michiharu Suga.


    FOOTNOTES

This research was supported by operating grants from the Canadian Institutes of Health for Research (MT-13270 and MOP-42546) and the Ontario Thoracic Society.

N. Isowa was a recipient of a fellowship from the Department of Surgery and Faculty of Medicine (University of Toronto, Toronto, ON). M. Liu is a Scholar of the Medical Research Council of Canada and a recipient of Premier's Research Excellent Award from the Ontario Government.

Address for reprint requests and other correspondence: M. Liu, Thoracic Surgery Research Laboratory, Toronto General Hospital, Univ. Health Network, Room CCRW 1-816, 200 Elizabeth St., Toronto, Ontario, Canada M5G 2C4 (E-mail: mingyao.liu{at}utoronto.ca).

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.

Received 1 August 2000; accepted in final form 8 November 2000.


    REFERENCES
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

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