1 Thoracic Surgery Research Laboratory, Toronto General Hospital, University Health Network, and Departments of 2 Anatomy and Cell Biology, 3 Surgery, and 4 Medicine, University of Toronto, Toronto, Ontario, Canada M5G 2C4
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ABSTRACT |
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Lipopolysaccharide
(LPS) polymerizes microfilaments and microtubules in macrophages and
monocytes. Disrupting microfilaments or microtubules with cytochalasin
D (CytoD) or colchicine can suppress LPS-induced tumor necrosis
factor- (TNF-
) gene expression and protein
production from these cells. We have recently demonstrated that primary
cultured rat alveolar epithelial cells can produce TNF-
on LPS
stimulation. In the present study, we found that the LPS-induced
increase in TNF-
mRNA level and protein production in alveolar
epithelial cells was not inhibited by CytoD or colchicine (1 nM to 10 µM). In fact, LPS-induced TNF-
production was further enhanced by
CytoD (1-10 µM) and inhibited by jasplakinolide, a polymerizing
agent for microfilaments. Immunofluorescent staining and confocal
microscopy showed that LPS (10 µg/ml) depolymerized microfilaments
and microtubules within 15 min, which was prolonged until 24 h for
microfilaments. These results suggest that the effects of LPS on the
cytoskeleton and the role of the cytoskeleton in mediating TNF-
production in alveolar epithelial cells are opposite to those in immune
cells. This disparity may reflect the different roles between nonimmune
and immune cells in host defense.
tumor necrosis factor-; cytokines; lipopolysaccharide; microfilament; microtubule
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INTRODUCTION |
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LIPOPOLYSACCHARIDE (LPS) is the principal
shock-inducing factor extracted from the outer membrane of
gram-negative bacteria. Proinflammatory cytokines including tumor
necrosis factor- (TNF-
) derived from LPS-stimulated cells are
responsible for the lethal effect of LPS (5). TNF-
, an
early-response proinflammatory cytokine, is produced by many types of
cells in response to LPS stimulation (4). Immune cells such as alveolar
macrophages are major sources of TNF-
in the lung (31). It has
recently been recognized, however, that nonimmune cells such as
epithelial cells may also play an important role in host defense by
producing cytokines and chemokines (15). McRitchie et al.
(22) recently found that isolated rat alveolar epithelial
cells can produce TNF-
in response to LPS stimulation. TNF-
produced in alveolar epithelial cells may function as an intermediate
signal to amplify LPS-induced cellular responses by further triggering
the production of chemokines such as macrophage inflammatory protein-2
(32). TNF-
may play a crucial role in mediating acute lung injury. Nash et al. (24) used immunohistochemistry staining to examine the
distribution of TNF-
in lung tissue from patients dying with adult
respiratory distress syndrome. They found that TNF-
protein was
located within epithelial cells resembling type II pneumocytes. There
were relatively few TNF-
-positive cells in the early stages of adult
respiratory distress syndrome, and TNF-
-positive cells were found
throughout the epithelium in the later stages (24). It has also been
demonstrated that ventilator-induced lung injury increased TNF-
mRNA
expression in lung epithelial cells with in situ hybridization (30).
The mechanisms by which LPS stimulates TNF- production from
macrophages, monocytes, and other inflammatory cells have been studied
extensively; however, the regulatory mechanisms acting in LPS
stimulation of alveolar epithelial cells are unknown. The cytoskeleton
plays an important role in mediating cytokine production by immune
cells in response to LPS stimulation. Treatment of macrophages or
monocytes with LPS increases polymerization of microfilaments (14, 28)
and microtubules (3). Cytochalasin D (CytoD) is an inhibitor of actin
polymerization, acting largely by binding to the barbed end of the
actin filament (10). Colchicine is a broadly used anti-inflammatory
drug inhibiting intracellular microtubule polymerization (1). CytoD
(27) or colchicine (19, 25) blocks LPS-induced TNF-
gene expression
and/or protein synthesis in macrophages. The purpose of the present
study was to determine the effects of polymerization status of the
cytoskeleton on LPS-induced TNF-
production in alveolar epithelial
cells. We found that neither CytoD nor colchicine inhibited TNF-
gene expression and protein production. On the contrary,
jasplakinolide, a novel microfilament-polymerizing agent (7),
attenuated LPS-induced TNF-
production from alveolar epithelial
cells but enhanced this effect from macrophages. Furthermore, LPS
induced depolymerization of both microfilaments and microtubules in
alveolar epithelial cells. These data suggest that although alveolar
epithelial cells produce TNF-
in response to LPS stimulation, the
role of the cytoskeleton in mediating LPS-induced TNF-
production is
different from that acting in immune cells such as macrophages and monocytes.
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MATERIALS AND METHODS |
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Reagents. LPS
(Escherichia coli and
Salmonella typhosa), CytoD,
colchicine, rat IgG, mouse monoclonal anti-actin antibody, and rabbit
skeletal muscle actin were purchased from Sigma (St. Louis, MO). DMEM,
fetal bovine serum (FBS), and gentamicin were purchased from GIBCO BRL
(Life Technologies, Mississauga, ON). Porcine pancreatic elastase was
purchased from Worthington Biochemical (Freehold, NJ). Pentobarbital
sodium was purchased from Bimeda-MTC Pharmaceuticals (Cambridge, ON).
Rat anti-mammalian -tubulin antibody and FITC-conjugated rabbit
anti-rat IgG were purchased from Serotec (Oxford, UK). Jasplakinolide,
rhodamine-phalloidin, and FITC-DNase 1 were purchased from Molecular
Probes (Eugene, OR). Peroxidase-linked sheep anti-mouse IgG was
purchased from Amersham Life Science (Oakville, ON).
Rat alveolar epithelial cells isolation and culture. Alveolar type II cells were obtained with the method of Dobbs (13) as previously described (20, 22). Briefly, male adult Sprague-Dawley rats (Harlan, Indianapolis, IN) weighing ~250 g were anesthetized by an 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 the cells were stained with crystal violet and trypan blue exclusion. The viability of the fresh alveolar epithelial cell suspensions was >95%.
The cells were cultured in DMEM containing 10% (vol/vol) FBS and 12.5 µg/ml of gentamicin. In most experiments, 1 ml of cell suspension
(106 cells/ml) was seeded in
24-well culture plates (Corning Glass Works, Corning, NY) and
maintained at 37°C in 5% CO2.
The cells were cultured in 6-well plates to determine the relative
amount of filamentous actin (F-actin) and to study TNF- gene
expression or in 96-well plates for 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. (22) have recently reported, this maneuver reduced the number of macrophages to undetectable levels 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 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; data not shown).
RAW 264.7 cell culture. Murine macrophage cells (RAW 264.7 cell line purchased from American Type Culture Collection, Manassas, VA) were cultured in DMEM supplemented with 4 mM L-glutamine, 4.5 g/l of glucose, 1 mM sodium pyruvate, and 10% (vol/vol) FBS. The cells were maintained in a humidified atmosphere of 5% CO2 at 37°C and split twice weekly. One day before the experiment, 1 ml of RAW 264.7 cell suspension (106 cells/ml) was seeded into 24-well culture plates (Corning) and allowed to adhere overnight at 37°C in 5% CO2.
Cytotoxicity assays. The cytotoxic effects of LPS, CytoD, jasplakinolide, and colchicine were examined by simultaneous double staining with fluorescein diacetate (FDA) and propidium iodide (PI), a rapid, convenient, and reliable method to determine cell viability (17). The cells were incubated in 96-well plates (Corning). After treatment with the agents mentioned above, the culture medium was removed, and the cells were washed twice with Dulbecco's phosphate-buffered saline (DPBS). The stock solutions of FDA (5 mg/ml in acetone) and PI (0.02 mg/ml in DPBS) were stored at 4°C in the dark. The working solution was freshly diluted and mixed with DPBS. The final solution containing 2 µg of FDA and 0.6 µg of PI was added to each well for 3 min at room temperature. The viability of cells was examined with a fluorescent microscope with 520- and 590-nm filters. Viable cells fluoresced bright green, whereas nonviable cells were bright red. The viability of cells in all groups in this study was found to be comparable to that of the control group without LPS or drugs.
Immunocytofluorescent 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 before LPS
treatment. On the day of experiment, the culture medium was replaced
with 10% FBS-DMEM with and without LPS (10 µg/ml) for 15 min or 24 h. The cells were washed three times with cold PBS and fixed in 3.7%
formaldehyde for 10 min at room temperature followed by washing 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 washing with PBS. For
localization of F-actin and globular actin (G-actin), the fixed and
permeabilized cells were stained with rhodamine-phalloidin (1:40 in
PBS) and/or FITC-DNase 1 (1:50 in PBS) for 30 min in the dark. For
localization of microtubules, the fixed and permeabilized cells were
stained with rat anti-mammalian -tubulin antibody (1:100 in
distilled water) for 30 min at room temperature. After being washed
with PBS, the slides were incubated with FITC-conjugated rabbit
anti-rat IgG (1:100 in distilled water) for 30 min at room temperature
in the dark. After unbound antibody was removed by washing with PBS,
the slides were mounted with an antifading reagent (SlowFade, Molecular
Probes). To determine the specificity of staining, the first antibody
was replaced with nonspecific rabbit IgG (Sigma) or omitted from the
staining procedure. Confocal laser scanning was performed with a
confocal laser scanning microscope (MRC-600, Bio-Rad, Mississauga, ON)
equipped with a Kr/Ar laser. In each experiment, control and
LPS-treated cells were cultured on the same chamber slide and processed
simultaneously for comparison.
Measurement of TNF-.
TNF-
concentrations in the culture medium were measured with ELISA
kits (Biosource, Camarillo, CA) following the manufacturer's
instructions. According to the manufacturer, the kit for rat TNF-
has no cross-reactivity to many human and mouse cytokines and no
cross-reactivity with rat interferon-
and macrophage inflammatory
protein-2. It has 0.15% cross-reactivity to human TNF-
and 100%
cross-reactivity to mouse TNF-
. The optical density of each well was
read at 450 nm with a NM600 microplate reader (Dynatech Laboratories,
Chantilly, VA). The final concentration was calculated by
converting the optical density readings against a standard curve.
Extraction, gel electrophoresis, and immunoblotting
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 LPS (10 µg/ml) in 10% FBS-DMEM
for 24 h. After being washed twice with ice-cold PBS, the cells were
lysed by adding 200 µl 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 to each well and holding 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
collected as the Triton-soluble fraction. 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. A fraction of the
Triton-soluble supernatants (15 µl) was mixed with 5 µl of 4×
SDS sample buffer. All samples were boiled for 10 min, and 15 µl of
each sample were subjected to SDS-PAGE (10% polyacrylamide gel) (23).
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) and by
comparison with purified actin from rabbit skeletal muscle as a
positive control. For immunoblotting, after electrophoresis, the
proteins were electrophoretically transferred onto nitrocellulose membranes. Nonspecific binding was blocked by incubation with 8%
(wt/vol) nonfat milk in PBS for 60 min at room temperature. The
blots were washed with PBS and incubated overnight at 4°C with a
mouse monoclonal anti-actin antibody (1:500 dilution). The blots were
incubated for 60 min at room temperature with peroxidase-linked sheep
anti-mouse IgG (1:10,000 dilution). After being washed, the blots were
developed with SuperSignal ULTRA Chemiluminescent Substrate (Pierce,
Rockford, IL). The actin bands were scanned with a GS-690 imaging
densitometer and quantified with a computer program, Molecular Analyst
version 1.5 (Bio-Rad, Hercules, CA).
RNA extraction and semiquantitative
RT-PCR. To study TNF- gene expression, 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 and
without various inhibitors for 2 h followed by stimulation with LPS (10 µg/ml) in 10% FBS-DMEM for 4 h with and without inhibitors. The
medium was removed, and the cells were washed twice with ice-cold PBS.
Total RNA was extracted with an RNeasy total RNA extraction kit
(Qiagen, Chatsworth, CA) following the manufacturer's
instructions. Briefly, the cells were lysed with 0.25 ml of lysis
buffer with 1% (vol/vol) freshly added
-mercaptoethanol and
homogenized with a QIAshredder column (Qiagen). The RNA concentration
was measured with a spectrophotometer (Beckman DU 640B, Fullerton, CA).
Two micrograms of RNA were used for the RT reaction with a Superscript
II kit (GIBCO BRL) following the manufacturer's instructions. RT
products from 0.2 µg of RNA were used for PCR. PCR primers for
-actin and TNF-
were synthesized by ACGT Corporation (Toronto,
ON). The sequences of primers for
-actin and TNF-
are listed in
Table 1. The PCR mixture was set up in a
total volume of 30 µl containing 3 µl of 10× PCR buffer (200 mM Tris-Cl, pH 8.4, and 500 mM KCl), 1 µl of 50 mM
MgCl2, 0.5 µl of 10 mM
deoxynucleotide triphosphate mixture, 0.5 µl of each PCR primer (10 µM), and 0.3 µl of Taq polymerase
(GIBCO BRL). PCR was performed with a programmable thermal cycler
(PTC-100, MJ Research, Watertown, MA). The optimized PCR conditions are described in Table 1. Ten microliters of PCR product were
electrophoresed on a 1% agarose gel, with ethidium bromide staining
for visualization, and the gels were photographed and quantified with a
gel-documentation system (Gel Doc 1000, Bio-Rad). To ensure
comparability, RT-PCR was performed simultaneously on all samples
collected from each experiment. PCR products were analyzed on the same
gel. The optical density of the PCR product bands was quantified with
integrated image-analysis software (Molecular Analyst version 1.5, Bio-Rad). With optimized PCR conditions, all data were collected
without saturation or missing bands. The background of the optical
density reading for each band was subtracted locally. RT-PCR was
conducted at least two times for each sample to ensure reproducibility.
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Statistical analysis. All experiments were carried out with materials collected from at least two to three separate cell cultures in duplicate or triplicate. All data are expressed as means ± SE from separate measurements and were analyzed with a personal computer with SigmaStat for Windows version 1.0 (Jandel, San Rafael, CA). Comparison of two groups was analyzed with Student's t-test. Comparison of more than two groups was carried out with one-way or two-way ANOVA followed by Student-Newman-Keuls test, with significance defined as P < 0.05.
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RESULTS |
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CytoD enhanced LPS-induced TNF-
release from alveolar epithelial cells. McRitchie et
al. (22) have recently shown that 10 µg/ml of LPS from
E. coli maximally stimulated TNF-
production by alveolar epithelial cells within 4 h. In the present
study, we found that treatment with LPS from S. typhosa (10 µg/ml) had a similar stimulatory effect
on TNF-
production in the same preparation (data not shown). LPS
from E. coli was used in subsequent
experiments except when otherwise specified. CytoD has been shown to
inhibit LPS-induced TNF-
production from macrophages, monocytes, or
neutrophils (27). We first examined whether CytoD has similar
inhibitory effects in alveolar epithelial cells. Treatment of alveolar
epithelial cells with various concentrations (1 nM to 10 µM) of CytoD
for 6 h did not change the basal levels of TNF-
in the culture
medium. The cells were pretreated with various concentrations of CytoD for 2 h, then challenged with LPS (10 µg/ml) for 4 h in the presence of CytoD. TNF-
production was not affected by CytoD at
concentrations equal to or below 0.1 µM. With higher concentrations
(1-10 µM), CytoD enhanced LPS-induced TNF-
production (Fig.
1A).
We then stimulated the cells with various concentrations of LPS (10 pg/ml to 10 µg/ml) in the presence and absence of CytoD (1 µM).
Similar to the previous observations by McRitchie et al. (22), TNF-
production increased in a dose-dependent manner with increasing doses
of LPS, which was further enhanced by CytoD treatment
(P < 0.05 as assessed by two-way
ANOVA; Fig. 1B).
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Jasplakinolide inhibited
LPS-induced TNF- release from alveolar epithelial
cells. To further clarify the role of microfilament polymerization in TNF-
production, cells were pretreated with various concentrations of jasplakinolide (1-100 nM) for 2 h and then challenged with LPS (10 µg/ml) for 4 h in the presence of jasplakinolide. A control group was handled similarly in the absence of
jasplakinolide. TNF-
production was inhibited by jasplakinolide at
all concentrations tested (Fig. 2). Treatment of
alveolar epithelial cells with jasplakinolide (1-100 nM) alone for
6 h did not change the basal levels of TNF-
in the culture
medium (data not shown).
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Colchicine had no inhibitory effect on
TNF- release in alveolar epithelial
cells. Microtubules, another major component of the
cytoskeleton, are also involved in mediating LPS-induced TNF-
production in immune cells (1, 19, 25). The effect of microtubule disruption on alveolar epithelial cells was investigated. When the
cells were incubated with various concentrations of colchicine, neither
the basal level nor the LPS stimulation-induced TNF-
production was
affected by colchicine at all concentrations tested (1 nM to 10 µM;
Fig. 3A). When the
cells were stimulated with various concentrations of LPS (1 ng/ml to 10 µg/ml), TNF-
production was not inhibited by colchicine (1 µM)
at any dose of LPS tested (Fig. 3B).
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CytoD and colchicine inhibited and jasplakinolide
enhanced LPS-induced TNF- release from macrophage
cells. To determine the efficiency of our CytoD,
jasplakinolide, and colchicine treatment protocols, we used a murine
macrophage cell line (RAW 264.7). RAW 264.7 cells were pretreated with
CytoD (1 µM), jasplakinolide (100 nM), or colchicine (1 µM) for 2 h
and then incubated with LPS and CytoD, jasplakinolide, or colchicine
for 4 h. With an optimized dose of LPS (10 ng/ml), RAW 264.7 cells
exhibited a marked increase in TNF-
release, an effect that was
significantly inhibited by CytoD or colchicine and enhanced by
jasplakinolide (Fig. 4).
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CytoD and colchicine did not block LPS-induced
TNF- mRNA expression in alveolar epithelial
cells. In macrophages, the inhibitory effects of CytoD
(28) or colchicine (19) on LPS-stimulated TNF-
production was
associated with inhibition of TNF-
mRNA expression. To determine
whether the treatment with CytoD or colchicine had any effect on
LPS-induced TNF-
gene expression by alveolar epithelial cells, the
cells were incubated with CytoD (1 µM) or colchicine (1 µM) before
and during LPS (10 µg/ml) stimulation. Total RNA was extracted, and
steady-state levels of TNF-
mRNA were determined with
semiquantitative RT-PCR. Because
-actin gene transcription was not
affected by either LPS or any inhibitors tested, the ratio of
densitometry units between TNF-
and
-actin mRNA was used to
represent the steady-state levels of TNF-
mRNA. Similar to our
previous observation, LPS induced an increase in TNF-
mRNA.
Treatment of cells with CytoD or colchicine had no effect on
LPS-induced TNF-
mRNA expression. A representative blot is shown in
Fig. 5A. Densitometry
data combined from three separate experiments are presented in Fig.
5B.
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LPS induced depolymerization of microfilaments in rat
alveolar epithelial cells. The opposite effects of
CytoD and jasplakinolide on LPS-induced TNF- production by alveolar
epithelial cells compared with that by macrophages suggest that the
effects of LPS on microfilaments of alveolar epithelial cells are
different from those on macrophages. In most cell types, actin exists
in both a G-actin and an F-actin form, which is regulated through a
dynamic polymerization and depolymerization process (8). We examined
these two kinds of actin with fluorescent staining and confocal
microscopy. The intensity of F-actin staining was decreased in primary
cultured alveolar epithelial cells by a short exposure (15 min) to LPS
(10 µg/ml; Fig. 6B)
compared with the untreated control cells (Fig.
6A). The staining of G-actin in
alveolar epithelial cells (Fig. 6C) was not affected by short periods of LPS stimulation (Fig.
6D). We also examined the effect of
LPS on microfilaments after longer periods of treatment. When alveolar
epithelial cells were incubated with LPS for 24 h, the staining of
F-actin was further reduced (Fig. 7,
A and
B), which was associated with a
clearly increased G-actin staining (Fig. 7,
C and
D). This LPS-induced
depolymerization was opposite to that described in macrophages (28) and
monocytes (14).
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To further confirm LPS-induced depolymerization of actin filaments, we
isolated F-actin from alveolar epithelial cells by Triton extraction
(23). After high-speed centrifugation, 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 (Fig.
8A). This band was
further confirmed to be actin by immunoblotting analysis (Fig.
8B). Densitometric quantification was performed, and the data were standardized by the cell number. Interestingly, the amount of actin in the Triton-insoluble fraction only accounted for a small percentage of the total actin in the cells
(Fig. 8C). LPS stimulation
significantly reduced the amount of Triton-insoluble actin to ~60%
(Fig. 8C). Similar results were obtained when the cells were stimulated with LPS from
S. typhosa (Fig.
8A).
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Effects of LPS on microtubules in rat alveolar
epithelial cells. Similarly, we investigated the effect
of LPS on microtubules in alveolar epithelial cells using
immunofluorescent staining and confocal microscopy. Unstimulated
alveolar epithelial cells demonstrated a diffuse, homogeneous pattern
of microtubules. The intensity of microtubule staining after a short
incubation period (15 min; Fig.
9A) was stronger than
that after a longer incubation period (24 h; Fig.
9B), which may be due to the change
in cell morphology during the culture. After a short period of LPS
stimulation (15 min), the intensity of microtubule staining decreased
(Fig. 9C) compared with that in
control cells (Fig. 9A). The reduced polymerization of microtubules was observed as soon as 1 min after LPS
stimulation and lasted at least 4 h (data not shown). However, no
difference in the microtubule pattern and staining intensity was
observed between control and LPS-stimulated cells after a 24-h
incubation (Fig. 9, B and
D). When the first antibody was replaced with rabbit nonimmune IgG or omitted from the staining protocol, no staining was detected (data not shown). The LPS-induced transient depolymerization of microtubules in alveolar epithelial cells was also opposite to that observed in human monocytes (3).
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DISCUSSION |
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In the present study, we observed that LPS stimulation induces TNF-
gene expression and protein production in alveolar epithelial cells,
which is consistent with the previous study by McRitchie et al. (22).
In contrast to macrophages and monocytes, LPS-induced TNF-
production by alveolar epithelial cells was enhanced by CytoD and was
not suppressed by colchicine. Jasplakinolide, a microfilament-stabilizing agent, showed opposite effects on TNF-
production between alveolar epithelial cells and macrophages. Furthermore, we found that LPS induced depolymerization of
microfilaments and microtubules in alveolar epithelial cells, which is
also opposite to that reported in macrophages and monocytes. Taken
together, these data indicate that although alveolar epithelial cells
can produce TNF-
in response to LPS stimulation, the effect of LPS on the cytoskeleton and the role of the cytoskeleton in LPS-induced TNF-
production are substantially different from those of
macrophages and monocytes.
The cytoskeleton is composed of three constituents: microfilaments,
microtubules, and intermediate filaments. Both microfilaments and
microtubules are involved in various activities of monocytes, macrophages, and neutrophils. LPS is known to affect the cytoskeleton system in immune cells (2, 3, 14, 16, 19, 27, 28). LPS induced a rapid
reorganization of F-actin assembly in macrophages (28) and increased
stiffness and F-actin assembly in monocytes (14) as well as enhanced
formyl-methionyl-leucyl-phenylalanine-induced actin polymerization in
neutrophils (16). Microfilaments are related to the trafficking of LPS
after its internalization in neutrophils (11). In mononuclear
phagocytes, LPS increased the number, length, and stability of
microtubules after either 30 min or 18 h of incubation (3). Incubation
of monocytes with LPS for 18 h increased the quantities of -tubulin,
-tubulin, and tyrosinated
-tubulin as well as
microtubule-associated protein-2 (2). Macrophages or monocytes must
change their shape to migrate to inflammatory sites in response to
stimuli such as LPS (15, 29). Increased polymerization of the
cytoskeleton may be related to these functions.
In contrast, alveolar epithelial cells play an important role in maintaining the integrity of the epithelial structure and in the recruitment of immune cells. The decrease in polymerization of microfilaments and microtubules in alveolar epithelial cells after LPS stimulation may facilitate the migration of inflammatory cells across the epithelium to enter the alveolar spaces. Both gram-negative and gram-positive bacteria can enter nonphagocytic cells such as epithelial cells (9, 18). Inhibition of microfilament (9) and microtubule (18) polymerization of host cells may reduce the invasion of bacteria. The depolymerization of the cytoskeleton induced by LPS may have a similar protective effect on alveolar epithelial cells.
In the present study, the enhancement by CytoD on TNF- production
from alveolar epithelial cells was weak, whereas the inhibitory effect
of jasplakinolide was dramatic. By contrast, the inhibitory effect of
CytoD on TNF-
production from RAW 264.7 cells was remarkable, whereas the enhanced effect of jasplakinolide was small. These results
suggest that when the effect of CytoD or jasplakinolide is opposite to
the effect of LPS on microfilament polymerization, it blocks
LPS-induced TNF-
production. On the other hand, when LPS increases
or decreases microfilament polymerization in one particular cell type,
further enhancement of this effect has less influence on TNF-
production.
LPS-induced depolymerization of microfilaments may enhance secretion
activity in alveolar epithelial cells. It has been noted that CytoD can
stimulate alveolar epithelial cells to release lung surfactant (26).
Depolymerization from F-actin to G-actin is involved in
terbutaline-induced transport and exocytosis of lamellar bodies, the
storage form of lung surfactant, from type II cells (6). Intratracheal
instillation of LPS to rats increased secretion of surfactant proteins
A and D (21). These observations suggest that a microfilament
depolymerization-dependent secretion exist in type II cells. F-actin
depolymerization induced by LPS-stimulation may facilitate TNF-
secretion through this mechanism.
The inhibitory effect of CytoD on LPS-induced TNF- production in
macrophages was observed at 0.2 µM (27), a concentration that is
incapable of affecting F-actin by itself (33). At this low
concentration, CytoD blocked LPS-induced elevation of TNF-
mRNA
(27). Suppression of LPS-stimulated TNF-
production in macrophages
by colchicine was also associated with inhibition of LPS-induced
TNF-
gene expression (1, 19, 25). LPS stimulation rapidly activated
mitogen-activated protein kinase associated with microtubules in
macrophages (12). Therefore, in macrophages, polymerization of
microfilaments and microtubules may be involved in LPS signaling
leading to TNF-
gene expression and subsequently its synthesis. In
the present study, agents affecting microfilaments and microtubules
had no effects on LPS-induced elevation in TNF-
mRNA levels. The
role of LPS-induced depolymerization of microfilaments and microtubules
in alveolar epithelial cells may be involved in LPS-induced TNF-
production at the posttranscriptional level.
Alveolar epithelial cells are continually exposed to external pathogens
and thus may play an important role in recruiting inflammatory cells by
producing various kinds of chemoattractants, including the
proinflammatory cytokines such as TNF-. The different effects of LPS
on the cytoskeleton between alveolar epithelial cells and macrophages
shown in this study may be due to their different roles in host
defense. The underlying molecular mechanisms need to be further studied.
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ACKNOWLEDGEMENTS |
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We acknowledge the technical assistance of Lu Cai, Xiao-Hui Bai, Xiao-Ming Zhang, Dr. Atsushi Baba, Dr. Young Kyoon Kim, and Dr. Michiharu Suga.
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FOOTNOTES |
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This research was supported by Medical Research Council of Canada Grants MT-13270 (to M. Liu) and MA-8558 (to A. S. Slutsky), the James H. Cummings Foundation (M. Liu), the Canadian Cystic Fibrosis Foundation (S. H. Keshavjee and M. Liu), and the Ontario Thoracic Society.
N. Isowa is a recipient of a fellowship from the Department of Surgery and Faculty of Medicine, University of Toronto (Toronto, Ontario, Canada). M. Liu is a Scholar of the Medical Research Council of Canada.
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. §1734 solely to indicate this fact.
Address for reprint requests and other correspondence: M. Liu, Thoracic Surgery Research Laboratory, Toronto General Hospital, University Health Network, Room CCRW 1-821, 200 Elizabeth St., Toronto, Ontario, Canada M5G 2C4 (E-mail: mingyao.liu{at}utoronto.ca).
Received 30 October 1998; accepted in final form 17 May 1999.
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