Department of Medicine, University of Iowa College of Medicine, and Veterans Administration Medical Center, Iowa City, Iowa 52242
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ABSTRACT |
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Exposure of macrophages to
endotoxin [lipopolysaccharide (LPS)] results in a cascade of events
resulting in the release of multiple inflammatory and anti-inflammatory
mediators. The Toll-like receptor (TLR) 4 complex is the major receptor
that mediates LPS signaling. However, there is evidence that other
surface molecules may play a complementary role in the TLR-induced
events. Integrin receptors are one class of receptors that have been
linked to LPS signaling. This study investigates the role of macrophage integrin receptors in the activation of mitogen-activated protein (MAP)
kinases by LPS. In conditions where macrophages were not permitted to
adhere to matrix or a tissue culture surface, we found a decrease in
LPS signaling as documented by a marked reduction in tyrosine
phosphorylation of whole cell proteins. This was accompanied by a
significant decrease in extracellular signal-regulated kinase and c-Jun
NH2-terminal kinase MAP kinase activation. Inhibition of
integrin signaling, with EDTA or RGD peptides, decreased LPS-induced MAP kinase activity. The functional consequence of blocking integrin signaling was demonstrated by decreased LPS-induced tumor necrosis factor- production. These observations demonstrate that, in addition to the TLR receptor complex, optimal LPS signaling requires
complementary signals from integrin receptors.
Toll-like receptor; tumor necrosis factor; integrins; lipopolysaccharide; mitogen-activated protein
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INTRODUCTION |
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LIPOPOLYSACCHARIDE
(LPS) plays a pivotal role in the innate immune response to
gram-negative bacteria (31, 42). LPS signaling is
initiated by an interaction between LPS and LPS-binding protein, allowing binding to CD14 and association with another cell membrane receptor, which contains an intracellular signaling domain. This other
receptor has recently been determined to be Toll-like receptor (TLR) 4, and significant work has been done to define this complex (for review
see Refs. 1, 17, 22). In
addition to the TLR complex, a number of other membrane receptors have
been proposed to play a role in LPS signaling. These include
2-integrins, a purinergic receptor (P2X7),
moesin, triggering receptor expressed on myeloid cells-1, a recently
described complex that includes heat shock proteins 90 and 70, complement receptor 4, and bone morphogenic protein (20, 23, 28,
40, 41). Several previous studies have suggested a role for the
integrin receptors in LPS signaling (12, 23, 29, 36).
Recently, Perera et al. (29) showed a decreased LPS
response [some cytokines, e.g., cyclooxygenase-2 and interleukin
(IL)-12 p35, and a slight decrease in p38 activation] in peritoneal
macrophages from CD11b (a major integrin chain in macrophages)-knockout
mice (29). This study evaluates the link between adherence
and LPS-induced mitogen-activated protein (MAP) kinase activation and
tumor necrosis factor (TNF)-
production in a murine macrophage line.
Integrins are a family of cell surface glycoproteins (for review see
Refs. 2, 3, 18). Each integrin
is a heterodimer that contains an - and a
-subunit. Each subunit
has a large extracellular domain, a single membrane-spanning region,
and a short cytoplasmic domain that contains binding sites for other proteins but does not itself have signaling capabilities. There are 16 known
-subunits and
8
-subunits, which can make up a multitude
of integrins. In macrophages, the main adhesion molecules are integrins
of the
1- or
2-class (39,
45). The
2-integrin (
L
2,
M
2,
X
2, and
D
2)
family has been especially linked to macrophage and leukocyte cell
adhesion and spreading (14, 15, 36). In particular,
M
2-integrin has been linked to LPS signaling by evaluation of cells from an
M-knockout
mouse (29).
Integrins link the extracellular matrix (ECM) to the actin cytoskeleton
and transmit biochemical signals across the plasma membrane
(7). As well as transmitting signals from the ECM, integrins can also be activated from signals generated intracellularly. For example, several studies have documented the intracellular activation of integrins after G protein-coupled receptor activation (25, 37, 38). This phenomenon has been described as
"inside-out" signaling. This contrasts with the signal generated
when an integrin binds to the ECM and signals are generated:
"outside-in" signaling (18). It is our hypothesis that
LPS/TLR signaling activates integrins, increasing adhesion and
generating an outside-in signal that amplifies LPS activation
of the extracellular signal-regulated kinase (ERK) and c-Jun
NH2-terminal kinase (JNK) MAP kinases. This results in
increased production of TNF-, suggesting a role for integrin
signaling in macrophage inflammatory responses (8, 26).
Integrin signaling has been linked to LPS-induced macrophage activation
(12, 27, 29). In other systems, it has been shown that
integrins and growth factor-dependent signals converge on the MAP
kinases (11). The MAP kinase family includes the ERK MAP
kinase, JNK, and p38. ERK is regulated by a variety of agents,
including integrins, growth factors, cytokines, and LPS. MAP kinase
activation follows a cascade of kinase activation [MAP kinase kinase
(MEK) kinase (MEKK)-MEK-ERK] and can be demonstrated by
phosphorylation of the threonine-tyrosine consensus motif
(46). It has not been demonstrated that maximal LPS MAP
kinase signaling depends on recruitment of integrin receptors and that
integrin-dependent signals (secondary to the TLR signal) amplify MAP
kinase activity and ultimately affect maximal TNF- release.
In this study we have evaluated the contribution of adherence-dependent MAP kinase activation compared with adherence-independent MAP kinase activation in the setting of LPS-exposed macrophages. We found a consistent but smaller LPS-induced ERK and JNK activation when cells were kept in suspension. This is linked to the production of inflammatory mediators, because inhibition of ERK or JNK by chemical inhibitors decreased LPS-induced TNF production. In addition, two blockers of integrin signaling, EDTA and soluble RGD peptides (36, 49), blocked optimal ERK and JNK activation. This finding is also linked to TNF production, because RGD peptides and EDTA decreased LPS-induced TNF production. The link with MAP kinase activation is confirmed by the decrease in LPS-induced TNF production with inhibition of ERK or JNK. From these observations, we conclude that LPS signaling to MAP kinases is decreased without the added contribution of integrin signaling. In addition, optimal cytokine production requires integrin signaling and the increased MAP kinase activation that accompanies adherence. We found also that adherence alone did not activate ERK, suggesting a possible inside-out signal from the TLR-4 complex to integrins. This inside-out signal would increase integrin activation and macrophage adherence. The added adherence would, in turn, provide an outside-in signal that could augment LPS-induced ERK or JNK activation. These observations lead us to propose that, in macrophages, integrins provide a complementary receptor to the already described TLR-4 complex and that both contributions are necessary for optimal signaling.
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MATERIALS AND METHODS |
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Materials. Chemicals were obtained from Sigma Chemical (St. Louis, MO), LPS from List Biological Laboratories (Campbell, CA), the MEK inhibitor U-0126 and the JNK inhibitor SP-600125 from Calbiochem (San Diego, CA), protease inhibitors from Boehringer Mannheim (St. Louis, MO), nitrocellulose and ECL Plus from Amersham (Arlington Heights, IL), and SuperSignal West Femto from Pierce (Rockford, IL). Antibodies were obtained from various sources: antibodies to ERK, p38, JNK, focal adhesion kinase (FAK), and phosphotyrosine from Santa Cruz Biotechnology (Santa Cruz, CA), phosphorylation-specific antibodies to p38 from Sigma Chemical, and all other phosphorylation-specific antibodies, including ERK, JNK, c-Jun, and phosphotyrosine, from Cell Signaling (Beverly, MA). Developing antibodies (horseradish peroxidase-conjugated anti-rabbit or anti-mouse Ig) were obtained from Santa Cruz Biotechnology. Special adherence-free plates (Ultra Low Attachment) were obtained from Corning (Corning, NY) and Teflon tubes from VWR (Batavia, IL). RGDS (RGD) peptide (Arg-Gly-Asp-Ser) was obtained from Bachem (Torrance, CA).
Cell culture. RAW 264.7 (RAW) cells (TIB-71, American Type Culture Collection) were maintained in DMEM with 10% fetal bovine serum and gentamicin (40 µg/ml). Cells were subcultured every 2-3 days. Experiments were run in six-well Costar tissue culture plates, low-adherence plates, or Teflon. For some experiments, cells were cultured in plates coated with fibronectin or collagen IV (BD Biocoat, Becton Dickinson Labware).
Isolation of whole cell extracts.
RAW cells were cultured in various conditions. Whole cell protein was
obtained by lysing the cells on ice for 20 min in 500 µl of lysis
buffer (0.05 M Tris, pH 7.4, 0.15 M NaCl, 1% Nonidet P-40, 0.5 M
phenylmethylsulfonyl fluoride, 50 µg/ml aprotinin, 10 µg/ml
leupeptin, 50 µg/ml pepstatin, 0.4 mM sodium orthovanadate, 10 mM
sodium fluoride, and 10 mM sodium pyrophosphate; all from Boehringer
Mannheim). The lysates were then sonicated for 20 s, kept at 4°C
for 30 min, and spun at 15,000 g for 10 min, and the supernatant was saved. Protein was determined using a protein measurement kit (protein assay kit 500-0006, Bio-Rad, Hercules, CA).
Cell lysates were stored at 70°C until use.
Immunoprecipitation of proteins. RAW cells were cultured in various conditions, and whole cell proteins were obtained. Aliquots (300 µg) from each sample were placed in a new tube for immunoprecipitation. The volume was brought up to 500 µl with lysis buffer with added sodium orthovanadate (2 mM). Rabbit anti-FAK antibody (20 µl/sample) was added, and the samples were rotated at 4°C overnight. On the next day, protein A-agarose (20 µl/sample; Santa Cruz Biotechnology) was added to each sample, and the tubes were rotated at 4°C for 1 h. The beads were subsequently washed four times with lysis buffer with added sodium orthovanadate (2 mM). The immunoprecipitated complexes were released with 2× sample buffer for Western analysis.
Western analysis.
Western analysis for the presence of particular proteins or for
phosphorylated forms of proteins was performed on whole cell proteins
from RAW cell experiments. Protein (30-80 µg) was mixed 1:1 with
2× sample buffer (20% glycerol, 4% SDS, 10% -mercaptoethanol, 0.05% bromphenol blue, and 1.25 M Tris, pH 6.8; all from Sigma Chemical) heated to 95°C for 5 min, loaded onto a 10%
SDS-polyacrylamide gel, and run at 100 V for 90 min. Cell proteins were
transferred to nitrocellulose (ECL, Amersham) by semidry transfer
(Bio-Rad) at 25 V for 45 min. Equal loading of the protein groups on
the blots was evaluated by using Ponceau S, a staining solution
designed for staining proteins on nitrocellulose membranes (Sigma
Chemical) or, in the case of phosphorylation-specific blots, by
stripping and reprobing with antibodies to the total protein. The
nitrocellulose was blocked with 5% milk in Tris-buffered saline with
0.1% Tween 20 for 1 h, washed, and then incubated with the
primary antibody overnight. The blots were washed four times with
Tris-buffered saline with Tween 20 and incubated for 1 h with
horseradish peroxidase-conjugated anti-rabbit or anti-mouse IgG
antibody. Immunoreactive bands were developed using a chemiluminescent
substrate (ECL Plus or SuperSignal West Femto). An autoradiograph was
obtained, with exposure times of 10 s-2 min.
Adherence assay. RAW cells were cultured in adherent or nonadherent conditions in special 24-well plates (106 cells/well, triplicate wells per group); some cells were kept aside for use in a standard curve. After the culture period, the wells were washed three times with prewarmed PBS with calcium and magnesium (0.5 ml/well; wide-bore pipette tips were used to add PBS), and the plate was blotted on thick blotting paper. PBS (0.5 ml) was added to each well along with a standard curve of cells in 0.5 ml PBS/well (106-105 cells/well). Calcein-AM (0.5 ml, 2 µM final concentration; Molecular Probes) was added to each well. The plate was incubated at 37°C for 30 min, and fluorescence was measured using a 485 ± 8 nm excitation filter and a 535 ± 10 nm emission filter on a Victor2 (EG & G Wallac, Gaithersburg, MD) microplate reader. Adherence was defined as the percentage of cells remaining in the plate after the PBS washes. Cell numbers were determined by comparing sample values with the standard curve generated by the known amount of cells.
Cytokine release.
RAW cells were cultured in standard medium for 6 h in adherent and
nonadherent conditions with and without LPS. After the culture period,
the supernatants were harvested and stored at 70°C until assayed.
The amount of TNF-
in the supernatant was measured by ELISA (R & D
Systems, Minneapolis, MN).
Isolation of RNA.
Total RNA was isolated using the Absolutely RNA RT-PCR Miniprep kit
(Stratagene, La Jolla, CA) according to the manufacturer's instructions. RNA was quantitated using the RiboGreen kit (Molecular Probes). RNA samples were stored at 70°C.
RT-PCR detection of TNF- mRNA.
Total RNA (1 µg) was reversed transcribed to cDNA using the
RETROscript RT-PCR kit (Ambion, Austin, TX). The resulting cDNA was
subjected to PCR as follows. In a 0.2-ml PCR tube (Bio-Rad), 2 µl of
cDNA were added to 48 µl of PCR mixture containing dNTP (Life
Technologies, Grand Island, NY) at 2 mM each, 1.5 mM MgCl2 (Life Technologies), 1:15,000 SYBR Green I DNA dye (Molecular Probes),
sense and antisense primers (Research Genetics, Huntsville, AL) at 0.2 µM each, and 2.5 units of platinum Taq DNA (Life
Technologies). Amplification was then performed in an iCycler iQ
Fluorescence thermocyler (Bio-Rad) as follows: 3 min at 95°C,
followed by 45 cycles of 20 s at 95°C, 20 s at 59°C,
20 s at 72°C, and 10 s at 81°C. Fluorescence data were
captured during the dwell at 81°C. Data were collected and recorded
by iCycler iQ software (Bio-Rad) and expressed as a function of
threshold cycle (Ct), the cycle at which the fluorescence
intensity in a given reaction tube rises above background. Primers for
murine TNF-
and hypoxanthine phosphoribosyltransferase (HPRT) genes
are as follows (5' to 3'): AACTTCGGGGTGATCGGTCC (sense) and
CAAATCGGCTGACGGTGTGGG (antisense) for TNF-
and CCTCATGGACTGATTATGGAC (sense) and CAGATTCAACTTGCGCTCATC (antisense) for HPRT. Primers were
selected on the basis of nucleotide sequences downloaded from the
National Center for Biotechnology Information data bank and designed
with software by Steve Rozen and Helen J. Skaletsky (1998 Primer3; code
available at
http://www-genome.wi.mit.edu/genome_software/other/primer3.html). PCR conditions and data collection dwell temperature are based on
melting curve analysis of each amplimer generated by the primers listed
above. Data were captured at 3°C below the lowest melting temperature
among all amplimers assayed to ensure that primer-dimers were not
contributing to the fluorescence signal generated with SYBR Green I DNA
dye. Specificity of the amplification was confirmed using melting curve analysis.
Quantitation of TNF- mRNA.
Relative quantitative gene expression was calculated as follows. For
each sample assayed, the Ct values for reactions amplifying TNF-
and HPRT were determined. The TNF-
Ct for each
sample was corrected by subtracting the Ct for HPRT
(
Ct). Untreated controls were chosen as the reference
samples, and the
Ct values for all LPS-treated
experimental samples were subtracted from the
Ct values
for the control samples (
Ct). Finally, LPS-treated
TNF-
mRNA abundance relative to control TNF-
mRNA abundance was
calculated as follows:
and HPRT genes. With the use of this set of
template mixtures, the amplification efficiencies for TNF-
and HPRT
amplimers were found to be identical.
Statistical analysis. Statistical analysis was performed on densitometry data, ELISA results, and real-time PCR data. Significance was determined by Student's t-test.
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RESULTS |
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LPS requires an adherence-induced signal for maximal TNF-
production.
We initially validated our low-adherence model by a specific
adherence assay. Using RAW cells, we cultured cells for 30 min in a
standard tissue culture plate (adherence) or in a special low-adherence
plate with and without LPS (1 µg/ml). Photomicrographs were obtained,
and an adherence assay was performed (see MATERIALS AND
METHODS). Figure
1A shows
RAW cells in the adherence and low-adherence conditions. Under
low-adherence conditions, the cells remain rounded and distributed in a
single-cell manner. In contrast, the adherence cells were clumped
together and beginning to stick to the plate. The degree of adherence
in the two plates after 30 min is quantified and shown in Fig.
1A. We also evaluated total tyrosine phosphorylation of
whole cell proteins and found that low adherence decreased the
LPS-induced tyrosine phosphorylation in a number of bands (data not
shown). This suggests a role for integrin signaling in LPS macrophage
responses. Exposure of macrophages to LPS starts a chain of events
culminating in the release of multiple inflammatory and
anti-inflammatory mediators. Central to macrophage immune function is
the release of TNF-
(13, 33). Because of the importance
of TNF-
, we wanted to evaluate whether the decreased signaling
exemplified by the lowered tyrosine phosphorylation in low-adherence
conditions had an effect on TNF-
production. RAW cells were cultured
in low-adherence and adherence conditions. LPS was added for 3 h
(RNA) or 6 h (protein). Relative TNF-
RNA levels were
determined using real-time RT-PCR (see MATERIALS AND METHODS), and protein release was determined by ELISA.
Figure 1B demonstrates that preventing adherence during LPS
exposure results in decreased production of the proinflammatory
cytokine TNF-
.
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LPS-induced MAP kinase activation is decreased in low-adherence
conditions.
Activation of multiple MAP kinases has been linked to LPS-induced
TNF- production (8, 26). We next evaluated whether lack
of adherence correlated with a decrease in MAP kinase activation by
LPS. RAW cells were cultured in adherence vs. low-adherence conditions
with and without LPS. Whole cell lysates were obtained at various times
after LPS and analyzed for MAP kinase activation using antibodies
specific for the phosphorylated form of ERK, JNK, and p38.
Phosphorylation at the tyrosine-threonine motif in these MAP kinases is
essential for activation. Figure 2 shows a significant decrease in the activation of all three MAP kinases when
LPS exposure occurred in the low-adherence conditions. The smallest
decrease was observed in the p38 blot. In addition, over multiple
experiments the observation that low adherence decreased LPS-induced
p38 activation was not consistent (data not shown). This suggests a
tenuous link between integrin signaling and LPS-induced p38 activation.
Because of the solid link between adherence and optimal ERK and JNK
activation by LPS, we concentrated on these kinases in the remaining
experiments. The data demonstrate that, over an extended time course,
low adherence decreases the degree of MAP kinase activation in RAW
cells after LPS exposure.
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ERK activation is linked to LPS-induced TNF production.
To determine that ERK activation was relevant to the decreases in TNF
production in low-adherence cells, we evaluated TNF production after
inhibition of ERK activity with the inhibitor U-0126. U-0126 prevents
ERK activation downstream of MEK (43) and is a more
complete blocker of ERK activation than the commonly used PD-98059. We
found that U-0126 almost completely blocked LPS-induced TNF in
adherence and low-adherence conditions (Fig. 3). Low adherence does not completely
block ERK activation, which explains the difference in degree of TNF
inhibition between U-0126 and low adherence alone. As controls, we also
demonstrated that U-0126 does block ERK activation and does not block
JNK activation or the tyrosine phosphorylation of FAK, an
integrin-linked kinase.
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JNK activation is linked to LPS-induced TNF production.
To determine that JNK activation was relevant to the decreases in TNF
production seen in low-adherence cells, we evaluated TNF production
after inhibition of JNK activity with the inhibitor SP-600125, which
prevents JNK activity (6). We found that SP-600125 decreases LPS-induced TNF (Fig. 4). As
controls, we also demonstrated that SP-600125 does not block ERK
activation and does block JNK activation using phosphorylation of the
JNK substrate c-Jun as a readout. SP-600125 inhibits the catalytic
activity of JNK and not the activating phosphorylations. For this
reason, the phosphorylation-specific JNK antibody cannot be used to
measure JNK activity after SP-600125. c-Jun is phosphorylated on
Ser73 by JNK and can be used to monitor JNK activity.
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Increasing the length of adherence before LPS exposure does not
activate ERK.
RAW cells were cultured in adherence vs. low-adherence conditions for
various time periods before the addition of LPS. After exposure to LPS
for 15 min, whole cell lysates were obtained, and ERK phosphorylation
was evaluated. Figure 5 demonstrates that adherence alone does not activate ERK in RAW cells. LPS is needed for
ERK activation, and LPS plus adherence is needed for optimal ERK
activation.
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LPS activation of ERK does not change on different ECM.
We next asked the question: Is LPS-induced ERK activation dependent on
a particular substrate? RAW cells were placed in culture for 30 min in
adherence plates, low-adherence plates, fibronectin-coated plates, or
collagen IV-coated plates. The cells were treated with LPS for 15 min,
whole cell protein was obtained, and ERK phosphorylation was evaluated.
Figure 6 shows an increase in ERK
phosphorylation (activity) in the adherence plates similar to that in
plates coated with fibronectin or collagen IV. This study suggests that
binding matrices other than the plastic tissue culture dish
(fibronectin and collagen IV) can also play a positive role in
LPS-induced ERK activation.
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LPS-induced tyrosine phosphorylation of FAK depends on adherence.
In other systems, integrin involvement in MAP kinase activation is via
FAK (4, 35). To determine the effect of adherence on FAK
tyrosine phosphorylation (a marker of activity) in RAW cells, we
treated cells with LPS for various periods of time and immunoprecipitated FAK. The immunoprecipitated FAK was then evaluated for tyrosine phosphorylation with an antibody that recognizes phosphotyrosine (4G10). Figure 7
demonstrates that LPS increases tyrosine phosphorylation of FAK and
that the low-adherence conditions prevented even the baseline FAK
phosphorylation. The blot was stripped and reprobed for total FAK.
There is a decrease in the amount of total FAK immunoprecipitated in
the adherence conditions because of the difficulty of isolating
proteins that are part of the detergent-resistant membrane (activated
FAK). However, this difference only makes the lack of tyrosine
phosphorylation in the low-adherence conditions more impressive. The
data suggest a link between LPS signaling and integrin activation and
also suggest that FAK activity might contribute to LPS-induced MAP kinase activation.
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Adherence-dependent LPS activation of ERK and JNK is blocked by
added EDTA.
Integrin signaling requires Ca2+, and in experimental
models of integrin signaling the addition of EDTA to the extracellular medium has been used as an integrin inhibitor (49). To
confirm that the decrease in signaling in the low-adherence conditions is mediated by integrin, we treated RAW cells with EDTA for 15 min
before adding LPS. Figure 8A
demonstrates that the increased ERK and JNK phosphorylation in
adherence plates was blocked by the preaddition of EDTA. There was also
a small decrease in ERK activation in the low-adherence samples,
perhaps because of a role of extracelluar Ca2+ in TLR-4
signaling or a small amount of integrin signaling generated by
cell-to-cell contact in the low-adherence plates. To link the integrin-dependent signals to LPS-induced TNF production, we evaluated LPS-induced TNF production in cells pretreated with EDTA. Figure 8B shows that EDTA decreases LPS-induced TNF production in
mouse macrophages.
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Adherence-dependent LPS activation of ERK and JNK is blocked by
added RGD peptides.
Cell attachment via integrins is mediated through a recognition site on
the extracellular portion of the integrin heterodimer that recognizes a
tripeptide, RGD, in target proteins of the ECM (32).
Soluble peptides containing the RGD motif can be used to inhibit cell
attachment (9). To further link optimal LPS signaling to
integrin engagement, we treated RAW cells with synthetic RGD peptides
15 min before the cells were plated on adherence plates. The cells were
cultured for 30 min, and then LPS was added for 15 min. Figure
9A shows that the RGD peptides
keep the cells separate, in contrast to the LPS-treated cells, which
demonstrate increased clumping and sticking. Figure 9B
quantifies this using an adherence assay. This assay demonstrates that
the RGD-treated cells do not demonstrate the increase in adherence that
is found in the LPS-treated cells. Figure 9C shows that
pretreatment with RGD peptides decreases ERK and JNK phosphorylation.
This correlates with the cells' ability to produce TNF-, because
addition of the RGD peptides significantly decreases the amount of
TNF-
produced by LPS-treated cells (Fig. 9D). The data
shown here, when considered in conjunction with the previous experiment
utilizing EDTA, demonstrate a strong link between increased ERK and
JNK, TNF-
production, LPS-induced activation, and integrin
signaling.
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DISCUSSION |
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The premise of this investigation was that LPS signaling to MAP
kinases would be significantly enhanced by cooperation between the
TLR-4 complex and integrin receptors. To investigate this, we utilized
RAW cells, a murine macrophage line, to study the effect of
integrin-mediated signal on LPS activation of MAP kinases. We found
that maintaining the cells in suspension (low adherence) significantly
decreased the degree of LPS activation of macrophages. We initially
correlated lack of cell adherence with a decrease in the amount of
TNF- produced by LPS-treated cells. Because of the decrease, we then
evaluated the effect of low adherence on MAP kinase activation and
found that all three MAP kinases exhibited decreased activation in low
adherence. This was especially the case for ERK and JNK, and the
remaining experiments evaluated these MAP kinases. The increase in MAP
kinase activity was mirrored by an increase in FAK tyrosine
phosphorylation that was completely absent in the low-adherence
conditions. We linked the adherence-dependent FAK phosphorylation to
LPS-induced ERK activation by showing that cytochalasin D disrupted
LPS-induced ERK activation. Two blockers of integrin signaling, EDTA
and soluble RGD peptides, blocked the increased MAP kinase activation
demonstrated in adherence plates. RGD peptide and EDTA decreases in
adherence translated into less LPS-induced TNF-
. In addition, we
found that ERK and JNK activity was linked to optimal TNF production
after LPS exposure. As an aggregate, the data suggest that, in order
for LPS to induce a maximal response in macrophages, there must be
cooperation between the TLR complex and integrin signaling (Fig.
10).
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Integrin activation of ERK has been documented in other systems (14). ERK is activated when serum-starved fibroblasts are placed on fibronectin (19). Epidermal growth factor activation of ERK requires adherence (30). How integrins are linked to activation of MEK and then ERK is not clear. Integrins are known to activate a major tyrosine kinase pathway initiated with activation of FAK (5). FAK undergoes autophosphorylation at Tyr397, which results in the recruitment of Src. This results in the phosphorylation of paxillin and p130CAS, which serve as adaptors for the recruitment of other signaling molecules (35). One event that occurs during this cascade is phosphorylation of FAK on Tyr925, creating a binding site for the adaptor protein Grb2. Grb2 recruits Sos and activates the classic Ras-Raf-MEK-ERK cascade (4, 35). However, alternative studies have demonstrated FAK-independent integrin activation of ERK (24, 44) as well as Ras-independent integrin activation of ERK (10). Our data do not address the specific pathway activated by integrin signaling, but they do suggest a role for integrins and FAK in LPS-mediated signaling. Further studies will determine the exact nature of the downstream effectors involved in LPS/integrin/ERK signaling.
Cross talk between integrins and other receptors has been documented in
a number of cases. In neutrophils, the CD11b/CD18 integrin receptor
cooperates with the Fc RIII to generate a respiratory burst
(48). Using carcinoma cells, Yebra et al. (47) found that
v
5-integrin-directed cell migration
required activation of the urokinase-type plasminogen activator
receptor. In this study, cell migration on vitronectin was blocked by
blocking antibodies to the urokinase-type plasminogen activator or the integrin receptor
v
5. This suggests a
required cross talk between the two receptors. In a T cell model of
migration through endothelium, Hwang et al. (21) found
that signaling via the glycosylation-dependent cell adhesion molecule-1
receptor (an L-selectin receptor) increased the avidity of
2-integrins, allowing for T cell recruitment into peripheral lymphoid tissue. This model is similar to our hypothesis for
TLR-4 and integrin signaling: LPS/TLR-4 provides a signal that
increases integrin affinity, resulting in adherence to ECM, further
activating the integrins, which then provide an enhanced signal to MAP kinases.
It is our hypothesis that the sequence of events leading to optimal ERK
and JNK activation in macrophages is as follows: LPS activation of the
TLR-4 complex inside-out activation of an integrin receptor
increased adherence
integrin signaling outside-in to MAP kinases. A
recent study by Schmidt et al. (36) supports the
inside-out part of this story. They found that, in J774.A1 macrophages,
LPS activated the
2-chain of the integrin family via a
sequential activation of TLR-4-myeloid differentiation factor-88-IL-1 receptor-associated kinase-p38-Rap1 GTPase. This LPS-induced pathway results in cell spreading. In support of the outside-in signaling, Perera et al. (29) showed that macrophages from
mice deficient in CD11b/CD18 (or
M
2)
produced decreased amounts of some cytokines (cyclooxygenase-2 and
IL-12 p35) in response to LPS or taxol. However, they found no
difference in the expression of TNF-
. The fact that the knockout in
question was to the CD11b chain leaves the possibility of other
2-heterodimers contributing to the TNF-
-inducing
signal. One important difference between our study and that of Perera
et al. is that their system blocks only one of the integrin
-chains,
while our system blocks all integrin signaling. Their study does
support the role of integrins in LPS signaling, inasmuch as they found
decreased NF-
B translocation, decreased p38, and decreased cytokines
in the CD11b knockouts. The differences they described were
substantially greater in CD14- and TLR-4-knockout macrophages,
suggesting that the TLR-4 complex is the primary receptor complex.
The regulation of multiple cellular events after macrophage LPS
exposure is complex. Because of the extreme bioactivity of many of the
molecules produced by the macrophage (i.e., TNF-), multiple
checkpoints are in place. One possible checkpoint is the interaction of
multiple receptors. Our data (lack of MAP kinase activation with
adherence but augmented LPS-induced MAP kinase activation with
adherence) suggest that the TLR-4 complex interacts with integrin
receptors. This suggests that optimal LPS responses are regulated at
many levels, including that of a multiplicity of receptors.
This study demonstrates that optimal LPS signaling, to important signaling molecules (MAP kinases) and ultimately to the generation of inflammatory mediators (TNF), requires the cooperation of integrin receptors. Although a role for integrin receptors in LPS responses has been suggested, this study establishes a unique link between integrins and MAP kinase activation in the context of LPS stimulation. We believe that this is an important addition to the study of inflammation.
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ACKNOWLEDGEMENTS |
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The authors thank David Fultz for graphics assistance.
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FOOTNOTES |
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This study was supported by a Veterans Administration Merit Review grant, National Institutes of Health Grants HL-60316 and ES-09607, and Environmental Protection Agency Grant R826711 to G. W. Hunninghake.
Address for reprint requests and other correspondence: M. M. Monick, Div. of Pulmonary, Critical Care, and Occupational Medicine, Rm. 100, EMRB, University of Iowa Hospitals and Clinic, Iowa City, IA 52242 (E-mail: martha-monick{at}uiowa.edu).
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.
March 1, 2002;10.1152/ajplung.00437.2001
Received 9 November 2001; accepted in final form 28 February 2002.
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