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
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ABSTRACT |
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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
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INTRODUCTION |
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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)- 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.
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MATERIALS AND METHODS |
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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 · mlCytotoxicity 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) -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 -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.
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RESULTS |
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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- 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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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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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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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- 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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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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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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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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DISCUSSION |
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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- gene expression and/or protein synthesis
(42).
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- ![]() |
ACKNOWLEDGEMENTS |
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We acknowledge the technical assistance of Xiao-Hui Bai, Xiao-Ming Zhang, and Dr. Michiharu Suga.
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FOOTNOTES |
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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.
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