1 Department of Bioengineering and The Whitaker Institute of Biomedical
Engineering, University of California, San Diego, La Jolla CA 92093-0427,
USA
2 Department of Vascular Biology VB-1, The Scripps Research Institute, La Jolla,
CA 92037, USA
* Present address: Department of Bioengineering, University of California,
Berkeley, Berkeley, CA, USA
Present address: Department of Biomedical Sciences, University of California,
Riverside, Riverside, CA, USA
Present address: Lawrence Berkeley National Laboratory, One Cyclotron Road, MS
74-157, Berkeley, CA, USA
¶ Author for correspondence (e-mail: shuchien{at}ucsd.edu )
Accepted 25 February 2002
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Summary |
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Key words: Integrin, Extracellular domain, Small GTPase, Signal transduction
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Introduction |
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Rho family GTPases (Rho, Rac and Cdc42), which function as binary switches
that cycle between an active GTP-bound form and an inactive GDP-bound form,
have distinct functions in regulating the actin-based cytoskeletal structure
(Ridley and Hall 1992;
Ridley et al., 1992
;
Van Aelst and D'Souza-Schorey
1997
). Rho increases cell contractility, focal adhesions and actin
stress fibers (Ridley and Hall
1992
; Hotchin and Hall
1995
), and Rac induces lamellipodia formation and membrane
ruffles, as well as wavy, loose bundles of actin filaments in the periphery
(Ridley and Hall 1992
;
Ridley et al., 1992
;
Rottner et al., 1999
;
Small et al., 1999
). Recent
studies have suggested that integrins can activate Cdc42, Rac and Rho
(Hotchin and Hall 1995
;
Clark et al., 1998
;
Price et al., 1998
;
Ren et al., 1999
). Fibroblasts
plated on fibronectin exhibit an early activation of Cdc42 and Rac and a
delayed activation of Rho (Price et al.,
1998
; Ren et al.,
1999
), suggesting that Cdc42/Rac and Rho may be regulated through
different pathways. However, the molecular basis of the differential
activation of Rho family GTPases is not clearly understood.
Here we showed that the overexpression of ß3 integrin enhanced Rho activity and stress fiber formation, whereas the overexpression of ß1 integrin increased Rac activity and lamellipodia formation. The overexpresion of a mutant ß1-3-1 integrin, in which the extracellular I-domain-like sequence of ß1 integrin has been replaced by the corresponding sequence of ß3 integrin, also enhanced Rho activity and stress fiber formation. These results suggest that the extracellular domains of ß1 and ß3 integrins play differential roles in transducing the extracellular stimuli to result in the specific regulation of Rac and Rho activities.
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Materials and Methods |
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The methods of transfection of CHO cells to generate single transfectants
expressing human ß1, ß3 and ß1-3-1 integrins have been
previously described (Takagi et al.,
1997). The parent CHO cells did not express endogenous ß3
integrin (Ylanne et al.,
1993
). Human ß1, ß3 or ß1-3-1 integrin cDNA in
pBJ-1 vector was co-transfected with pFneo into CHO cells. ß1-3-1
integrin is a mutant in which the sequence CTSEQNC (residues 187-193) in the
extracellular I-domain-like structure of ß1 integrin is replaced with the
corresponding CYDMKTTC sequence of ß3 integrin. The transfected cells
were then selected with G418. After selection, ß1, ß3 and
ß1-3-1 cells were cultured in the presence of G418 at 100 µg/ml in CHO
cell culture medium. The expressed ß integrins were detected at focal
adhesions in each cell line (Takagi et
al., 1997
). The expression levels of human ß1 integrin on
ß1 cells and ß1-3-1 integrin on ß1-3-1 cells were analyzed by
flow cytometry using an anti-human ß1 integrin antibody (AIIB2). ß1
and ß1-3-1 cell lines with equivalent expression level of ß1 or
ß1-3-1 integrins were used in our experiments. The expression level of
ß3 integrin was not directly comparable with that of ß1 integrin,
owing to the different affinities of antibodies against ß1 and ß3
integrins.
For all experiments, culture dishes or slides were coated with fibronectin (2.5 µg/cm2) or fibrinogen (5 µg/cm2) for 2 hours, and the non-specific binding sites were blocked with 0.2% bovine serum albumin (BSA) (Calbiochem).
Actin staining and confocal microscopy
Cells were plated on coverslips that had been coated with either 2.5
µg/cm2 fibronectin or 5.0 µg/cm2 fibrinogen for 4
hours in serum-free DMEM. The plated cells were fixed in 3% paraformaldehyde
in phosphate buffer saline (PBS) for 30 minutes and permeabilized with 0.5%
Triton X-100 in PBS for 15 minutes. The specimens were then incubated with
FITC-conjugated phalloidin (5 U/ml, Molecular Probes, Eugene, OR) for 1 hour,
and the resulting actin staining was observed under a confocal microscopy
system (MRC-1024, Bio-Rad, Hercules, CA).
Rho and Rac activities assay
The activities of Rho and Rac were measured by the recently developed
affinity-precipitation assays on the basis of the specific interaction of the
activated Rho family GTPases with their downstream effectors
(Ren et al., 1999;
del Pozo et al., 2000
).
Briefly, after the cells had been passed onto culture dishes coated with
fibronectin for different durations in the absence of serum, the Rho-binding
domain (RBD) from Rho effector Rhotekin and the p21-binding domain (PBD) from
PAK were purified as GST fusion proteins. For the Rho activity assay, the
cells were lysed with a high salt RIPA buffer (50 mM Tris-HCl, pH 7.2, 1%
Triton X-100, 0.5% sodium deoxycholate, 0.1% SDS, 500 mM NaCl, 10 mM
MgCl2, 10 µg/ml leupeptin, 1 mM orthovanadate and 1 mM PMSF) and
centrifuged at 13,000 g at 4°C for 10 minutes. The cell
lysates were incubated with 20 µg GST-RBD beads for 45 minutes at 4°C.
The activated Rho was detected by immunoblotting using a monoclonal anti-RhoA
antibody (Santa Cruz Biotechnology, Santa Cruz, CA). For Rac activity assay,
the cells were lysed in a buffer containing 50 mM Tris-HCl pH 7.0, 0.5% NP-40,
500 mM NaCl, 5 mM MgCl2, 5% glycerol, 10 µg/ml leupeptin, 1 mM
orthovanadate and 1 mM PMSF. GST-PBD beads were used for affinity
precipitation assays, and the activated Rac was detected by immunoblotting
using an anti-Rac1 antibody (Transduction Laboratory). In the inhibition
experiment, the PI 3-kinase inhibitor LY294002 (Sigma) was added to ß1
cells in suspension for 30 minutes before seeding the cells on
fibronectin.
PI 3-kinase activity assay
Cells were lysed in a buffer containing 50 mM HEPEs, pH 7.5, 1% NP-40, 10%
glycerol, 150 mM NaCl, 1.5 mM MgCl2, 1.0 mM EDTA, 10 mM
Na4P2O7, 100 mM NaF, 1 mM PMSF, 1 mM
Na3VO4, 10ug/ml leupeptin and 10 µg/ml aprotinin. The
supernatant containing 500 µg protein was immunoprecipitated by using an
anti-PI 3-kinase p85 polyclonal antibody (Santa Cruz Biotechnology). After
washing, the beads were resuspended in 50 µl of the kinase buffer
containing 0.2 mg/ml of phosphatidylinositol, 20 µM ATP, 20 µCi of
[-32P] ATP, and 20 mM MgCl2. The sample was
extracted once with 160 µl methanol-chloroform (1:1) and twice with 200
µl chloroform. The combined organic phase was washed once with 200 µl of
1M HCl-methanol (1:1). Phosphatidylinositol was recovered from the organic
phase by evaporation, suspended in 15 µl chloroform and analyzed by thin
layer chromatography (TLC) on a Silica gel 60 TLC plate (VWR, Willard, OH)
(Whitman et al., 1988
). The
products were visualized by autoradiography. To detect the amounts of
immunoprecipitated PI 3-kinase in each sample, 200 µg of protein lysates
were immunoprecipitated by using an anti-PI 3-kinase p85 polyclonal antibody,
and the immunoprecipitated complexes were separated by SDS-polyacrylamide gel
electrophoresis (PAGE). The amount of PI 3-kinase p85 was detected by the
anti-PI 3-kinase p85 polyclonal antibody.
JNK kinase activity assay
Cells were lysed in a kinase lysis buffer containing 25 mM HEPES, pH 7.4,
0.5 M NaCl, 5 mM EDTA, 50 mM NaF, 1 mM Na3VO4, 1 mM PMSF
and 2 mM BGP. JNK was immunoprecipitated by using a polyclonal anti-JNK1
antibody (Santa Cruz Biotechnology) and protein A-Sepharose beads (Sigma). The
immunocomplexes were incubated in 30 µl kinase buffer containing 25 mM
HEPES, pH 7.4, 20 mM MgCl2, 1 mM PMSF, 10 µg/ml leupeptin, 20 mM
ß-glycerophosphate, 1 mM Na3VO4 and 2 mM DTT, 2
µg of glutathione S-transferase (GST-c-Jun-1-79) fusion protein, 10 µCi
[-32P] ATP and 25 µM ATP at 30°C for 20 minutes. The
reactions were stopped by the addition of a SDS sample buffer. The
phosphorylated proteins were separated by SDS-PAGE and visualized by
autoradiography.
Adhesion and detachment assays
The adhesion assay was performed as previously described
(Takagi et al., 1997).
Briefly, 96-well Immulon-2 microtiter plates (Dynatech Laboratories,
Chantilly, VA) were coated with 2.5 µg/cm2 fibronectin in 100
µl of PBS and incubated for 1 hour at 37°C. The remaining protein
binding sites were blocked by incubating with 0.2% BSA for 1 hour at room
temperature. Cells (2.5x104 cells/well) in 100 µl of DMEM
were added to the wells and incubated at 37°C for 5 minutes. After
removing the non-bound cells by rinsing the wells with the same buffer, the
bound cells were quantified by measuring the endogenous phosphatase activity
(Prater et al., 1991
).
The detachment assay was used to detect the strength of cell-substrate
adhesion, which was quantified by calculating the percentage of cells removed
by well-defined shear flow. Cells were seeded on a fibronectin-coated (2.5
µg/cm2) glass slide and incubated at 37°C for 4 hours to
allow the establishment of cell-substrate adhesion. The glass slide was then
assembled into a parallel plate flow chamber
(Hochmuth et al., 1973), and
the cells were subjected to a dislodging shear stress of 250
dyn/cm2 for 10 minutes. The numbers of cells before and after the
application of the shear flow was counted in the same field of view from
videotaped images. The results were expressed as the percentage of cells
detached after the application of the shear flow.
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Results |
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|
To further test the roles of ß1 and ß3 subunits in the formation
of lamellipodia and stress fibers, CHO, ß1, ß 3 and ß1-3-1
cells were plated on coverslips coated with fibrinogen, which binds only to
ß3 but not to ß1 integrin. As expected, CHO and ß1 cells did
not attach to the fibrinogen-coated coverslips because CHO cells did not
express endogenous ß3 subunit (Ylanne
et al., 1993). ß3 (Fig.
1E) and ß1-3-1 (Fig.
1F) cells plated on fibrinogen had a comparable level of stress
fiber formation to when these cells were plated on fibronectin.
ß1 and ß3 integrins differentially regulate Rho and Rac
activities to cause the different actin organization
To investigate whether ß1 and ß3 integrins differentially
regulate Rho and Rac activities to cause the different actin organization,
suspended CHO, ß1, ß3 and ß1-3-1 cells were plated on
fibronectin-coated culture dishes in serum-free DMEM for various lengths of
time. RhoA activity was detected by RBD of Rhotekin and pull-down assays
(Ren et al., 1999). As shown
in Fig. 2, the level of RhoA
activity in the CHO (Fig. 2A)
and ß1 (Fig. 2B) cells
plated on fibronectin was lower than that in suspension, and this decrease
remained significant for at least 4 hours (P<0.05). In contrast,
the level of RhoA activity in ß3 (Fig.
2C) and ß1-3-1 cells (Fig.
2D), following an initial decrease in the early phase of adhesion,
rose to become not significantly different from the level in suspension after
1 hour of adhesion (P>0.30). We did not detect any difference in
Rho activity among CHO, ß1, ß3 and ß1-3-1 cells in suspension
(Fig. 2E). These results
suggest that ß3 integrin preferentially promoted the increase of Rho
activity of adhered cells to the level seen in suspended cells and that the
extracellular I-domain-like structure of ß3 integrin is important for
this effect on Rho activity.
|
To examine whether expression of ß3 integrin and overexpression of
ß1 integrin affect Rac activity, we used PBD and pull down assays
(del Pozo et al., 2000). In
contrast to Rho activity, the level of Rac1 activity
(Fig. 3A) was higher in CHO and
ß1 cells after 4 hours of adhesion, suggesting that ß1 integrin
preferentially increases Rac activity. Again, there is no significant
difference in Rac activity among different types of cells in suspension
(Fig. 3B).
|
Overexpression of ß1 integrin causes adhesion induction of PI
3-kinase and JNK activities
Rac is known to be activated by PI 3-kinase and to stimulate a kinase
cascade leading to the subsequent activation of JNK
(Coso et al., 1995;
Minden et al., 1995
;
Kiyono et al., 1999
). To test
whether overexpression of ß1 integrin can enhance the activation of PI
3-kinase and JNK, we examined PI 3-kinase and JNK activities after plating
cells on fibronectin for 4 hours. The activation of PI 3-kinase
(Fig. 4A) was higher in ß1
cells than that in CHO, ß3 and ß1-3-1 cells. To test the
relationship between PI 3-kinase and Rac in this system, the specific PI
3-kinase inhibitor LY294002 was added to a ß1 cell suspension 30 minutes
before seeding the cells on fibronectin. Inhibition of PI 3-kinase with
LY294002 blocked Rac activity in ß1 cells
(Fig. 4B), suggesting that PI
3-kinase is involved in the activation of Rac1 in this system. Consistent with
the activation of PI 3-kinase and Rac1, JNK kinase activity in ß1 cells
was higher than that in CHO, ß1 and ß1-3-1 cells
(Fig. 4C). These results
suggest that ß1 integrin activates JNK through the activation of PI
3-kinase and Rac and that this signaling process is regulated by the
extracellular I-domain-like structure of the ß1 subunit.
|
Adhesion and detachment assays
Two approaches were used to quantify cell adhesion. First, we studied the
efficiency of cell attachment to a fibronectin-coated substrate. This adhesion
assay provides a very gentle removal of non-adherent cells. We detected no
significant differences in the number of bound cells in different cell lines
(Fig. 5A). In the second
approach, we applied a dislodging shear stress (250 dyn/cm2) to the
cells, which had established matured cell-substrate adhesion to test the
strength of adhesion. As shown in Fig.
5B, we did not detect any significant difference in the number of
detached cells in different cell lines. These results suggest that the
different phenotypes of ß1 and ß3 cells were not caused by the
differences in cell adhesion.
|
![]() |
Discussion |
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The I-domain-like structure within the ß1, ß2 and ß3
subunits has been reported to have components critical for ligand binding and
regulation (Loftus et al.,
1994; Bajt et al.,
1995
; Puzon-McLaughlin and
Takada 1996
; Tozer et al.,
1996
; Takada et al.,
1997
). The CHO cells that express human ß1-3-1 integrin
provided a cell system that was ideally suited to evaluate the role of the
extracellular I-domain-like structure of the integrin ß subunits in
integrin-mediated changes in cell morphology and Rho family GTPase activity.
Expression of a mutant ß1-3-1 integrin enhanced Rho activity and stress
fibers so that there was higher activity than there was in ß1 cells, that
is, this resulted in the same phenotype as ß3 rather than being like
ß1 cells. Since both the expression level of ß1 and ß1-3-1
integrins and the adhesion force of ß1 and ß1-3-1 cells were
similar, our findings indicate that the extracellular I-domain-like structures
of the integrin ß subunits play an important role in ß1- and
ß3-integrin-mediated increases of Rho and Rac activities. The molecular
mechanisms involved in the differential regulation of Rho and Rac by ß1
and ß3 integrins are not known. One possibility is that different ligand
binding on the extracellular domains of ß subunits may change the
conformation of the integrins, thus affecting the recruitment of signaling
molecules at focal adhesions and the anchorage of the cytoskeleton.
There are many possible mediators for the differential modulation of Rho
and Rac activities by integrins. Tyrosine phosphorylation has been shown to
regulate Rho and Rac activities (for a review, see
Kjoller and Hall 1999), but we
did not detect significant difference in the level of tyrosine phosphorylation
in CHO, ß1, ß3 and ß1-3-1 cells (H.M. and S.C., unpublished).
Our data indicate that PI 3-kinase mediates the enhancement of Rac activity
induced by the overexpression of ß1 integrin
(Fig. 4). PI 3-kinase could
stimulate Rac activity through its lipid product phosphatidylinositol
3,4,5-trisphosphate, which binds to the PH domain of guanine nucleotide
exchange factors (GEFs), such as Tiam1, Sos and Vav
(Rameh et al., 1997
;
Han et al., 1998
). It is likely
that the differential regulation of Rho and Rac by integrins is mediated by
the differential regulation of GEFs, GTPase-activating proteins (GAPs) and
guanine nucleotide dissociation inhibitors (GDIs). Indeed, some of the GEFs
and GAPs are specific regulators of particular Rho family members. For
example, GEF Lbc and GAP p122 specifically regulate Rho activity
(Homma and Emori 1995
;
Zheng et al., 1995
), whereas
GEF Tiam and GAP chemerin specifically regulate Rac activity
(Diekmann et al., 1991
;
Habets et al., 1994
). It is
possible that different integrin ß subunits modulate the functions of one
or more GEFs, GAPs and GDIs, thus changing the overall balance of these
regulators. Such differential regulation may go through different signaling
molecules and their combinations and/or different through feedback regulation
at focal adhesions.
The coordinated regulation of Rho GTPases is complicated. In fibroblasts,
Cdc42 activates Rac, which in turn activates Rho to regulate the actin
cytoskeleton (Nobes and Hall
1995), suggesting a hierarchical cascade for Rho GTPases. However,
the differential regulation of Rho and Rac has been shown under some
circumstances. For example, the growth of microtubules can lead to the
activation of Rac (Waterman-Storer et al.,
1999
), whereas disruption of microtubules leads to the activation
of Rho (Ren et al., 1999
).
Downstream in the activation pathway, Rac can counter the effect of Rho by
decreasing the phosphorylation of myosin light chain
(Sanders et al., 1999
). Our
data indicate that ß1 integrin increases Rac activity, whereas ß3
integrin enhances Rho activity. Since Rac and Rho regulate different
downstream effectors in signal transduction and actin cytoskeleton
organization, the signals from ß1 and ß3 integrins may complement
each other in the regulation of actin filament assembly and signaling
events.
In summary, we have shown that ß1 and ß3 integrins differentially regulate Rac and Rho activities and actin organization. Therefore, ß1 and ß3 integrins could play differential as well as coordinated roles in modulating cell functions such as migration and proliferation.
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Acknowledgments |
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