From the Department of Medicine and Cancer Center, University of California, San Diego, La Jolla, California 92093-0652
Received for publication, August 5, 2002, and in revised form, November 7, 2002
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ABSTRACT |
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Transformation by oncogenic Ras requires
signaling through Rho family proteins including RhoA, but the
mechanism(s) whereby oncogenic Ras regulates the activity of RhoA is
(are) unknown. We examined the effect of Ras on RhoA activity in NIH
3T3 cells either stably transfected with H-Ras(V12) under control of an inducible promoter or transiently expressing the activated H-Ras. Using
a novel method to quantitate enzymatically the GTP bound to Rho, we
found that expression of the oncogenic Ras increased Rho activity
~2-fold. Increased Rho activity was associated with increased plasma
membrane binding of RhoA and decreased activity of the
Rho/Ras-regulated p21WAF1/CIP1 promoter. RhoA
activation by oncogenic Ras could be explained by a decrease in
cytosolic p190 Rho-GAP activity and translocation of p190 Rho-GAP from
the cytosol to a detergent-insoluble cytoskeletal fraction.
Pharmacologic inhibition of the Ras/Raf/MEK/ERK pathway prevented
Ras-induced activation of RhoA and translocation of p190 Rho-GAP;
expression of constitutively active Raf-1 kinase or MEK was sufficient
to induce p190 Rho-GAP translocation. We conclude that in NIH 3T3 cells
oncogenic Ras activates RhoA through the Raf/MEK/ERK pathway by
decreasing the cytosolic activity and changing the subcellular
localization of p190 Rho-GAP.
Proteins of the Ras superfamily, including the Ras and Rho
families, cycle between active GTP- and inactive GDP-bound forms and
function as essential switches in signal transduction pathways that
regulate cell growth, differentiation, and survival (1). Activating
mutations in H-, K-, and N-Ras are found in up to 30% of all human
cancers; in cancers with wild type Ras, overexpression of growth factor
receptors frequently leads to activation of the Ras/Raf/MEK1/ERK pathway,
suggesting an important contribution of Ras functions to the
development of human cancers (1-3). Although there are no reports of
activating mutations of Rho proteins in human tumors, several Rho
proteins are overexpressed in tumors, and Rho family-activating guanine
nucleotide exchange factors (GEFs) have been isolated in screens for
transforming genes, suggesting a role of Rho proteins in tumorigenesis
(4).
Members of the Rho family regulate the actin cytoskeleton, thereby
affecting cell morphology and motility; in addition, they modulate gene
expression, cell cycle progression, and cell survival (1, 4, 5). RhoA,
B, and C and Rac1 play critical roles in cell transformation induced by
activated, oncogenic Ras, with dominant negative Rho and Rac1
constructs inhibiting Ras-induced transformation and constitutively
active constructs inducing anchorage-independent growth and other
features of the transformed phenotype (4, 6-9). The requirement of Rho
for Ras-induced transformation exists in part because Ras and Rho play
opposing roles in control of the cyclin-dependent kinase
inhibitor p21WAF1/CIP1, with Ras inducing and
Rho inhibiting p21WAF1/CIP1 transcription; thus increased
Ras activity actually blocks cell cycle progression when Rho signaling
is inhibited by C3 exoenzyme (8, 10, 11).
Activated, oncogenic Ras may regulate RhoA and Rac1 activities, but the
effects of Ras appear to be cell type-specific, vary with the Ras
subtype, and depend on the kinetics and duration of Ras activation (8,
12-18). Microinjection or transient transfection of oncogenic
H-Ras(V12) into Swiss 3T3 cells leads to acute cytoskeletal changes,
suggesting a hierarchal system with Ras activating Rac (causing
membrane ruffling) and Rac in turn activating RhoA (causing induction
of stress fibers); however, these studies were performed before direct
measures of Rac and Rho·GTP levels were available (12, 13). Ras
activation of Rac can occur through the Ras effector
phosphatidylinositol 3-kinase, with increased phosphoinositides activating a multimolecular complex including a Rac-activating GEF
(19-21). How Rac activation can lead to activation of RhoA is less
clear, but it may involve Rac activation of phospholipase A2 with subsequent arachidonic acid and leukotriene
production, at least in Swiss 3T3 cells (22).
In contrast to the acute response to oncogenic Ras, studies in
Ras-transformed cell lines have produced conflicting results, with some
studies reporting decreased Rac and increased RhoA activation compared
with nontransformed cells, and others reporting increased activation of
both GTPases or no changes (8, 14-18). In v-H-Ras-transformed MDCK
cells, decreased Rac activity appeared to be secondary to transcriptional down-regulation of the Rac-specific GEF Tiam1; the
mechanism for increased RhoA activation was not elucidated, but the
effect of oncogenic Ras on RhoA and Rac activity was mimicked by
stable transfection of constitutively active Raf (16).
H-Ras(V12)-transformed Swiss 3T3 cells also demonstrated
decreased Rac and increased Rho activity compared with untransformed
cells, but short term expression of a constitutively active Raf in
Swiss 3T3 cells did not lead to elevation of RhoA activity; elevated
RhoC·GTP levels were only seen after prolonged (>4 weeks) culture of
the active Raf-overexpressing cells, suggesting that they were a
consequence of selection rather than direct signaling (8). In HT1080
human fibrosarcoma cells containing oncogenic N-Ras, both Rac and RhoA activity were increased compared with cells lacking the mutant N-Ras;
Rac and RhoA activities were also increased when cells lacking the
mutant N-Ras were stably transfected with constitutively active Raf or
MEK (18). In K-Ras(V12)-transformed normal rat kidney cells, no
significant change of RhoA activity was observed compared with
untransformed cells (23). Several older studies reported loss of stress
fibers in Ras-transformed Rat1 cells, with restoration of stress fibers
upon transfection of constitutively active RhoA, suggesting loss of
RhoA activity in the Ras-transformed cells (7, 24); others reported
increased stress fibers in Ras-transformed breast cancer cells and NIH
3T3 cells without direct measurement of Rho activity (14, 15).
Because the mechanism of Rho regulation by Ras is not clear, we decided
to examine the effects of oncogenic Ras on the activation state of RhoA
in NIH 3T3 cells stably expressing H-Ras(V12) under control of an
inducible promoter (LTR-H-Ras(A) cells (25)); these cells have low
basal and high induced levels of H-Ras(V12) and allowed us to study
short term effects of Ras activation avoiding complex genetic changes
that may occur during long term culture of Ras-transformed cells. Using
two different methods to assess Rho activation, we found that induction
of H-Ras(V12) in LTR-H-Ras(A) cells or transient transfection of
H-Ras(V12) into wild type NIH 3T3 cells caused an approximate 2-fold
increase in Rho·GTP levels. Concomitant with Rho activation, we found
increased RhoA translocation to membranes and decreased activity of a
p21WAF1/CIP1 promoter construct. The mechanism for
increased Rho activation appeared to be decreased p190 Rho-GAP activity
because of translocation of p190 Rho-GAP from the cytosol to a
detergent-insoluble cytoskeletal fraction.
Cell Culture and Transfection Experiments
Wild type NIH 3T3 fibroblasts and LTR-H-Ras(A) NIH 3T3 cells
stably expressing activated Ras(V12) under the inducible murine mammary
tumor virus promoter were routinely cultured in Dulbecco's modified
Eagle's medium (DMEM) supplemented with 10% fetal bovine serum (FBS)
as described previously (26). To induce Ras(V12) expression, cells were
treated with 1 µM dexamethasone for 24 h. For
transient transfection experiments, cells were plated in six-well
cluster dishes and 24 h later were transfected with a total of 1.5 µg of DNA/well using LipofectAMINE PlusTM or
LipofectAMINE 2000TM (Invitrogen) as described previously
(27). The specific MEK inhibitor U0126 was from Calbiochem (28). The
following plasmids were used: pcDNA3-EE-RhoA(wt),
pcDNA3-EE-RhoA(V14), pcDNA3-EE-RhoA(63L), and pRC/CMV-BXB,
described previously (27, 29); pDCR-H-Ras(V12) from M. Wigler (30);
p Measurement of Rho Activation
The activation state of endogenous Rho was measured by two
different methods: (i) measurement of absolute amounts of GTP and GTP + GDP bound to Rho and (ii) assessment of Rho-bound GTP by Western
blotting. In both methods, the Rho binding domain (RBD) of Rhotekin was
used to isolate Rho·GTP as originally described by Ren et
al. (36). Glutathione S-transferase (GST)-tagged
Rhotekin RBD was purified from bacterial lysates; the bacterial
expression vector was provided by M. A. Schwartz (36). In the first
method, there is the potential to measure activation of RhoA, B, and C simultaneously; however, NIH 3T3 cells express mainly RhoA with low
amounts of RhoC and negligible amounts of RhoB (37). The first method
was modified to allow quantitation of GTP and GTP + GDP bound to
transfected RhoA constructs.
Measurement of Absolute Amounts of GTP and GTP + GDP Bound to
Rho--
This method is a modification of a procedure we have used
previously to measure GTP, and GTP + GDP, bound to Ras, Rap1, and Rheb
(2, 26, 38-40). Cells grown on a 100-mm plate under the conditions
indicated under "Results" and in the figure legends were extracted
quickly in situ by washing once with ice-cold Tris-buffered saline, pH 7.4, and adding lysis buffer consisting of 50 mM
Tris-HCl, pH 7.4, 1% Nonidet P-40, 1% CHAPS, 200 mM NaCl,
1 mM MgCl2, 10 µg/ml leupeptin, 10 µg/ml
aprotinin, and 1 mM phenylmethylsulfonyl fluoride. After a
1-min incubation on ice, the lysed cells were scraped with a rubber
policeman, transferred to a microcentrifuge tube, and subjected to
vortexing for 10 s. Cell extracts were centrifuged at 10,000 × g for 2 min, and a portion of the supernatant was added
to tubes containing 10 mM MgSO4 and 30 µg of
GST-tagged Rhotekin RBD bound to glutathione beads; these samples were
used for measuring GTP bound to Rho ("unloaded" samples). The
remaining supernatant was added to tubes containing 10 µM
GTP, 10 mM EDTA, and 30 µg of GST-tagged RBD on
glutathione beads, allowing the free GTP to exchange for GDP bound to
Rho, thus converting all of the Rho to the GTP-bound state
("loaded" samples). After gentle shaking for 1 h at 4 °C,
the beads with Rho·GTP bound to the Rhotekin RBD were washed four
times with 50 mM Tris-HCl, pH 7.4, 2% Nonidet P-40, 500 mM NaCl, 10 mM MgSO4, and twice
with 20 mM Tris-PO4, pH 7.4, 5 mM
MgSO4. GTP was released from Rho by heating the beads for 3 min at 100 °C in 5 mM Tris-PO4, pH 7.4, 2 mM dithiothreitol, 2 mM EDTA (TDE buffer). We
have shown previously >95% recovery of GTP under these conditions
(26).
GTP eluted from the unloaded and loaded samples was measured in
a coupled enzymatic assay by conversion to ATP in the presence of ADP
and nucleoside diphosphate kinase (26); the resulting ATP was measured
by the firefly luciferase method in a photon-counting luminometer (MGM
Instruments, Hamden, CT). This method is sensitive to 1 fmol of GTP and
is quantitative because the second reaction is irreversible, from light
generation, allowing both reactions to go to completion (26).
Assessment of Rho-bound GTP by Western Blotting--
Cells were
extracted and processed as described above for the unloaded samples
except the magnesium concentration in the initial lysis buffer was
increased to 10 mM; Rho·GTP isolated by binding to the
Rhotekin RBD-coated beads was quantitated by Western blotting using a
RhoA-specific antibody (Santa Cruz Biotechnology), as described by Ren
et al. (36).
Measurement of the Activation State of EE Epitope-tagged RhoA
Expressed in Cells--
Cells transfected with EE epitope-tagged RhoA
constructs were extracted in situ in 50 mM
Tris-HCl, pH 7.4, 1% Nonidet P-40, 500 mM NaCl, 10 mM MgCl2, 0.5% deoxycholate, 0.05% SDS, 1 mM EDTA, 1 mM phenylmethylsulfonyl fluoride, 10 µg/ml aprotinin, 10 µg/ml leupeptin (RIPA buffer). After
centrifuging the extracts, supernatants were split in half and added to
tubes containing protein G-agarose beads coated with either a mouse
monoclonal anti-EE antibody or control mouse IgG. The tubes were shaken
gently for 1 h at 4 °C, and the beads were washed four times
with RIPA buffer, and twice with 20 mM
Tris-PO4, pH 7.4, 5 mM MgSO4. GTP
and GDP were released from the immunoprecipitated Rho as described
above by heating the beads in TDE buffer. In one aliquot of the sample,
GTP was measured as described above, and in another aliquot the sum of GDP plus GTP was measured as described previously (38) by converting GDP to GTP using pyruvate kinase and phosphoenolpyruvate with the
resulting GTP (representing the sum of GDP plus GTP) measured as
described above.
Assessment of p21WAF1/CIP1
Promoter Activity
Cells were plated at 8 × 104/well on a 24-well
culture plate, and 24 h later the cells were transfected with 300 ng of DNA using PolyfectTM (Invitrogen) according to the
manufacturer's recommendation. All cells received 25 ng of p21-Luc,
and as indicated some cells additionally received 50 ng pEF-C3exo or
100 ng of pcDNA3-EE-Rho(63L). Cells were treated for 24 h with
1 µM dexamethasone, and luciferase activity was measured
in cell extracts as described previously (41). We did not include an
internal control vector because all four that were tested,
i.e. pRSV- Assessment of Rac Activation
Subconfluent cells grown on two 150-mm plates were extracted by
incubating for 2 min in cold RIPA buffer, and extracts were centrifuged
at 10,000 × g for 2 min. The p21(Rac/CDC42)-binding domain of human PAK-1 bound to glutathione-agarose beads was used to
isolate Rac·GTP, and the amount of Rac·GTP bound to the beads was
quantitated by Western blotting with a mouse monoclonal anti-Rac antibody as described previously (42), using an assay kit from Upstate Biotechnology.
Assessment of Subcellular Localization of RhoA, Rho-GAP, and
Ras-GAP
Cells grown on 150-mm plates were extracted by Dounce
homogenization in 10 mM Hepes, pH 7.5, 2 mM
EDTA, 1 mM MgCl2 (HEM buffer). The resulting
cell homogenate was centrifuged at 500 × g for 5 min
to remove nuclei and subcellular organelles, and the supernatant was
centrifuged at 37,000 × g for 30 min. The supernatant
and pellet from the second centrifugation are referred to as cytosol and membranes, respectively, with the membrane preparation washed twice
in HEM buffer to remove contaminating cytosol. Protein concentrations were determined according to Bradford (43), and equal amounts of
protein from each preparation (30 µg of homogenate, 20 µg of cytosol, and 40 µg of membranes) were subjected to SDS-PAGE/Western blotting using mouse monoclonal anti-Rho-GAP (Santa Cruz Biotechnology, 1:1,000) and anti-Ras-GAP (Sigma, 1:500) antibodies, or a rabbit polyclonal anti-RhoA antibody (Santa Cruz Biotechnology, 1:1,000).
Triton X-100-insoluble cytoskeletal fractions were prepared as
described previously (44). Cells were washed in phosphate-buffered saline and extracted in situ for 40 s at room
temperature in 50 mM Na-Hepes, pH 6.4, 3 mM
EGTA, 5 mM MgCl2, 0.5% Triton X-100. The
detergent-soluble supernatant was removed; the detergent-insoluble material was scraped off with a rubber policeman in the presence of
phosphate-buffered saline and protease inhibitor mixture, and centrifuged for 10 min at 300 × g at 4 °C. Pellets
were resuspended in SDS-sample buffer and analyzed by SDS-PAGE/Western
blotting using the monoclonal anti-Rho-GAP antibody and an
actin-specific antibody (C2, Santa Cruz Biotechnology, 1:300 dilution).
Assessment of Rho/Rho-GDI Association
Cytosolic extracts were prepared as described above and
subjected to immunoprecipitation using a rabbit polyclonal anti-Rho-GDI antibody or control rabbit IgG. Immunoprecipitates were collected on
protein G-agarose beads and analyzed by SDS-PAGE/Western blotting using
a mouse monoclonal anti-RhoA antibody and the rabbit polyclonal anti-Rho-GDI antibody (both from Santa Cruz Biotechnology, used at
1:1,000).
Measurement of Rho-GEF Activity
Cells grown on 100-mm plates were extracted by sonication in 10 mM Hepes, pH 7.4, 1 mM EDTA. Bacterially
expressed GST-RhoA was purified on glutathione-Sepharose beads and was
loaded with [3H]GDP (specific activity of 11.7 Ci/mmol)
by a 10-min incubation at 37 °C in 50 mM Hepes, pH 7.5, 5 mM EDTA, as described previously (45). The
[3H]GDP-loaded RhoA was incubated with cell extracts in
the presence of 1 mM GTP for 10 and 20 min at 37 °C, and
the reaction was stopped by adding a 40-fold excess of ice-cold RIPA
buffer. After washing the beads three times with RIPA buffer, they were
dried on filter paper, and radioactivity was measured by liquid
scintillation counting. The data are expressed as the percent increase
in GDP·GTP exchange compared with [3H]GDP-loaded RhoA
incubated in extract buffer.
Measurement of Rho-GAP Activity
Cells grown on 150-mm plates were extracted by Dounce
homogenization in HEM buffer supplemented with 1 mM
Na3VO4 and a protease inhibitor mixture, and
the cytosol was prepared as described above. After addition of 1%
Triton X-100, 250 or 500 µg of extract protein was subjected to
immunoprecipitation using protein G-agarose beads precoupled with
either mouse monoclonal anti-p190 Rho-GAP antibody or control mouse
IgG. The beads were washed three times with Triton X-100-containing
buffer and once with a reaction buffer containing 50 mM
Tris-HCl, pH 7.5, 10 mM MgCl2, 1 mM
dithiothreitol, 1 mg/ml bovine serum albumin, 1 mM GTP. The
beads were resuspended in 100 µl of reaction buffer, and 50 ng of
GST-RhoA preloaded with [ Assessment of Ras-GAP and Rho-GAP Association and of Rho-GAP
Phosphorylation
Cytosolic extracts were prepared and subjected to
immunoprecipitation with either the anti-Rho-GAP or anti-Ras-GAP
antibodies as described above. Immunoprecipitates were analyzed by
SDS-PAGE/Western immunoblotting using the same antibodies. The amount
of phosphorylated Rho-GAP was determined by probing the blots with an
anti-phosphotyrosine-specific antibody (Santa Cruz Biotechnology,
1:500).
Assessment of MAP Kinase Activation
MAP kinase activity was assessed by Western blotting using a
phospho-ERK-specific antibody that recognizes a dually phosphorylated peptide sequence corresponding to Thr183 and
Tyr185 of p42 MAP kinase, as described previously (27).
Quantitative Measurement of Rho Activation--
As part of these
studies, we developed a new quantitative method to assess Rho
activation by measuring absolute amounts of GTP and of total
nucleotides i.e. the sum of GTP plus GDP, bound to Rho. To
measure Rho·GTP, it was isolated from cell extracts according to the
method of Ren et al. (36) using glutathione-agarose beads
coated with a GST-Rhotekin RBD fusion protein; however, instead of
assessing the Rho·GTP semiquantitatively by Western blotting, we
eluted GTP from Rho and measured it in a coupled enzymatic assay as
described previously for measuring GTP bound to other Ras-related
proteins (26, 38-40). To measure total nucleotides bound to Rho, we
converted Rho·GDP to Rho·GTP by incubating a separate aliquot of
extract in the absence of magnesium and in the presence of 10 µM GTP; under these conditions, Rho·GDP is converted
rapidly to Rho·GTP (46), and the latter was measured as just described.
In extracts prepared from logarithmically growing NIH 3T3 cells, the
assay yielded a linear response over a 5-fold range of cellular protein
for both unloaded samples, i.e. those in which Rho·GTP was
measured directly (Fig. 1a),
and for loaded samples, i.e. those in which total
nucleotides bound to Rho were measured after converting Rho·GDP to
Rho·GTP (Fig. 1c). Because the graphs obtained with
unloaded and loaded samples overlap, the assay was linear over more
than a 5-fold range, and the data allow calculation of Rho activation,
i.e. [Rho·GTP/(Rho·GTP + Rho·GDP)] × 100. Thus, the
two data points marked with an asterisk and a pound
sign in Fig. 1, a and c, yielded a Rho
activation of 4.1%, and the data point marked with a double
dagger yielded a Rho activation of 3.2%. Rho activation can also
be calculated from the slopes of the lines in Fig. 1, a and
c (181 and 4,887 fmol of GTP/mg of protein, respectively),
yielding a Rho activation state of 3.7%. Similar results were found in
two other independent experiments, and experiments performed with
suspension cells (HL-60 human leukemia cells) also yielded a linear
response over a 5-fold range of cell number. For comparison, Rho·GTP
bound to the Rhotekin RBD-coated beads was also assessed by Western
blotting using a RhoA-specific antibody; within the limits of the
method, a linear response was obtained for both unloaded and loaded
samples (Fig. 1, b and d). The enzymatic method
detected Rho·GTP with slightly higher sensitivity compared with the
Western blot method; we define our limit of detection as the amount of
Rho·GTP which produces a signal greater than 2-fold above
background.
Rho Activation under Different Harvesting and Culture Conditions
and Activation Levels of Transfected Wild Type and Constitutively
Active RhoA Constructs--
Because Rho activation levels can change
in response to shear stress, changes in integrin ligation, and cell
density (36, 47, 48), we studied the effect of different cell
harvesting methods on Rho activation in NIH 3T3 cells. We found that
scraping cells with a rubber policeman from tissue culture dishes
followed by centrifugation and extraction yielded Rho activation levels 1.7 ± 0.3-fold higher than directly lysing cells on culture
plates (data are the means ± S.D. of three independent
experiments performed in duplicate, p < 0.05). These
results are consistent with the findings of others that Rho can be
activated by mechanical stimuli (47). In all subsequent experiments,
cells were extracted rapidly by lysis in situ. When we
extracted cells from subconfluent and confluent cultures, we found no
dependence of the Rho activation state on cell density, as described by
others for NIH 3T3 cells (48).
After 24 h of serum starvation, we found a Rho activation level of
4.8 ± 1.1% in wild type NIH 3T3 cells; this level increased to
14.4 ± 0.5% and 7.9 ± 2.5% after 1 and 3 min of serum
stimulation, respectively, yielding a 3-fold increase in Rho activation
at 1 min (Fig. 2a). Assessment
of Rho activation by Western blotting also yielded an approximate
three-fold increase after one minute of serum stimulation (Fig.
2b), thus confirming that the two methods provide similar
results. Previous workers reported a 2-6-fold increase in RhoA
activation during serum stimulation of Swiss 3T3 cells (36).
We modified the enzymatic assay to quantitate the activation state of
transfected RhoA constructs. We transfected NIH 3T3 cells with EE
epitope-tagged constructs of wild type RhoA, RhoA(63L), and RhoA(14V),
isolated the expressed protein from cell extracts using an anti-EE
epitope antibody, and measured GTP and total nucleotides bound to the
isolated RhoA as described under "Experimental Procedures." We
found activation states of 3.6 ± 0.8%, 64 ± 5%, and
83 ± 4% for wild type RhoA, RhoA(63L), and RhoA(14V),
respectively (Fig. 2c; compare RhoA(wt), open bar
on the far left, with RhoA(63L) and (14V), closed
bars). When we cotransfected the constitutively active
Rho Activation in NIH 3T3 LTR-H-Ras(A) Cells and NIH 3T3 Cells
Transiently Transfected with Wild Type and Activated Ras--
Rho
proteins are required for Ras-induced transformation, but there are
conflicting data concerning whether Ras signaling directly causes Rho
activation or whether some changes in Rho activity occur only during
the selection of Ras-transformed cells (5, 8, 16, 18, 23). Some of the
variation in results may be the result of cell type-specific
differences, but some may also be methodologic in origin. To address
the latter specifically, we used two different methods to assess Rho
activation, i.e. the quantitative enzyme-based method and
Western blotting, and we performed the studies under two different
conditions: (i) in NIH 3T3 cells stably expressing activated Ras under
an inducible promoter (LTR-H-Ras(A) cells) and (ii) in NIH 3T3 cells
transiently transfected with wild type Ras and Ras(12V) expression vectors.
LTR-H-Ras(A) cells express H-Ras(V12) under control of the
dexamethasone-inducible mouse mammary tumor virus long terminal repeat
and exhibit a transformed phenotype strictly dependent on the presence
of hormone in the growth medium (50). The addition of dexamethasone
leads to a significant increase in Ras expression at 8 h with peak
expression at 24 h; this is associated with a dramatic change in
cell morphology and acquisition of anchorage-independent growth (50).
Tumor formation in nude mice does not require dexamethasone treatment,
presumably because of endogenous glucocorticoid hormones in the animals
(50). When we induced oncogenic Ras expression in LTR-H-Ras(A) cells by
treating the cells with 1 µM dexamethasone for 24 h,
we found a 2-fold increase in Rho activation using the quantitative
enzymatic method to measure GTP bound to Rho; treating wild type NIH
3T3 cells with dexamethasone had no effect on Rho·GTP levels (Fig.
3a; p < 0.05 for the difference in Rho activation for LTR-H-Ras(A) cells in the
absence and presence of dexamethasone). Confirming these results,
dexamethasone induced a similar approximate 2-fold increase in
Rho·GTP in LTR-H-Ras(A) cells when assessed by the immunoblot method
(Fig. 3b, top panel, compare lanes 3 and 4), whereas no increase in Rho·GTP was observed in
wild type NIH 3T3 cells (Fig. 3b, top panel,
lanes 1 and 2). Dexamethasone had no effect on
total RhoA expression in either cell type, measured either by Western
blotting (Fig. 3b, middle panel) or by
determining total nucleotides bound to Rho (not shown).
As expected, dexamethasone increased Ras expression in LTR-H-Ras(A)
cells but had no effect on Ras levels in wild type cells (Fig.
3b, bottom panel, shows a Western blot).
Measuring total nucleotides bound to Ras (26), we found that
dexamethasone induced a 2.2-fold increase in total Ras protein in the
LTR-H-Ras(A) cells, and the activation state of Ras increased from 7.1 to 24.7% in the absence and presence of dexamethasone, respectively,
with no effect of dexamethasone on Ras activity or expression levels in
wild type NIH 3T3 cells (Table I). The
degree of Ras activation in dexamethasone-treated LTR-H-Ras(A) cells is
similar to the degree of Ras activation found in human pancreatic tumor
cells containing a mutated K-Ras
allele,2 or in HL-60 cells
containing a mutated N-Ras (51). Because the antibody Y13259 used for
immunoprecipitation of Ras recognizes all three Ras isoforms, the Ras
activation measured in Table I represents an average activation state
of H-, K-, and N-Ras, with the mutant H-Ras(V12) probably contributing
only about half of total Ras in dexamethasone-treated LTR-H-Ras(A)
cells.
To be sure that the increased Rho activity in dexamethasone-treated
LTR-H-Ras(A) cells was not influenced by clonal selection of the stably
transfected cells, we performed transient transfection experiments in
wild type NIH 3T3 cells and found a 2.1-fold increase in Rho activation
in cells expressing Ras(12V) compared with cells transfected with empty
vector (Fig. 3c; p < 0.05 for the
difference between control and Ras(12V)-expressing cells). Rho
activation required activated Ras because there was only a minimal
increase in Rho activation which did not reach statistical significance in cells transfected with wild type Ras (Fig. 3c). The
effects of the drug U0126 on Rho activation (Fig. 3, a,
b, and c) will be presented later.
Effect of Ras-induced Rho Activation on the
p21WAF1/CIP1 Promoter--
To determine whether
the observed Ras activation of Rho in NIH 3T3 cells had downstream
effects, we studied the activity of the
p21WAF1/CIP1 promoter in the absence and
presence of dexamethasone in LTR-H-Ras(A) cells. The
p21WAF1/CIP1 promoter was chosen because it is regulated
coordinately by both Ras and Rho, with Ras activating the promoter and
Rho inhibiting it (10). We transfected LTR-H-Ras(A) cells with a
p21-Luc construct and found no significant effect of dexamethasone
(Fig. 4, cells cotransfected with empty
vector). However, when these cells were cotransfected with an
expression vector for C3 exoenzyme, which ADP ribosylates and inhibits
RhoA (52), there was a 1.7-fold increase in luciferase activity in the
absence of dexamethasone and more than a 3-fold increase in the
presence of dexamethasone (Fig. 4); the difference between the absence
and presence of dexamethasone was statistically significant
(p < 0.05). The increase in p21WAF1/CIP1
promoter activity in the presence of C3 exozyme was likely secondary to
Rho inhibition, and the further increase with dexamethasone was the
effect of activated Ras on the promoter free of inhibition by Rho. We
demonstrated RhoA inhibition of the p21WAF1/CIP1 promoter
by expressing a constitutively active Rho (RhoA(63L); Fig. 4); the
RhoA(63L) effect was dominant over that of activated Ras, possibly
because of high expression levels of the transiently transfected
construct and the extremely high activation level of the mutant RhoA
(Fig. 2d). We conclude that RhoA activation by Ras is
sufficient to prevent stimulation of the p21WAF1/CIP1
promoter by Ras.
Effect of Oncogenic Ras on Rac·GTP Levels--
Rac1 is required
for Ras-induced transformation, but there are conflicting data
concerning whether Ras activation leads to an increased or decreased
Rac activation state (8, 9, 16, 18, 53). Active Rac may modulate the
activity of Rho in a cell type-specific fashion because in NIH 3T3
cells both transient and sustained Rac activation decreases Rho·GTP
levels, whereas in Swiss 3T3 fibroblasts and human endothelial cells,
microinjection of activated Rac1 appears to activate Rho (12, 22, 54,
55). We examined the effect of oncogenic Ras on Rac activity in
LTR-H-Ras(A) cells, and we found a reproducible 1.6 ± 0.2-fold
increase in Rac·GTP levels when oncogenic Ras expression was induced
by dexamethasone (Fig. 5a
shows a representative experiment, and Fig. 5c summarizes five independent experiments). The drug was without effect on Rac·GTP
levels in wild type NIH 3T3 cells (Fig. 5, b and
c). Because transfection of either activated Rac(V12) or the
Rac-GEF Tiam1 in NIH 3T3 cells leads to down-regulation of RhoA
activity (55), the dexamethasone-induced increase in Rac1 may limit the
increase in RhoA activity observed in LTR-H-Ras(A) cells.
Effect of Oncogenic Ras on the Subcellular Localization of RhoA,
RhoA Association with Rho-GDI, and on Rho-GEF Activity--
Changes in
the distribution of RhoA between membrane and cytosol have been used as
indication for Rho activation because increased membrane association of
RhoA occurs when RhoA is activated by the addition of GTP
Rho-GDI forms a cytosolic complex with Rho·GDP; Rho-GDI prevents Rho
activation by Rho-GEFs and serves to regulate membrane association/dissociation of Rho (56). We hypothesized that Ras-induced Rho activation in LTR-H-Ras(A) cells could be from a decreased Rho/Rho-GDI interaction. We assessed the amount of Rho complexed with
Rho-GDI in LTR-H-Ras(A) cells by coimmunoprecipitation and found no
change in the amount of RhoA in Rho-GDI immunoprecipitates when the
cells were treated with dexamethasone (Fig. 6b). The reciprocal experiment, i.e. assessing the amount of Rho-GDI
in RhoA immunoprecipitates, also yielded no difference in the absence or presence of dexamethasone (not shown).
Another possible mechanism for Rho activation by Ras could be through
an increase in Rho-GEF activity. Because there are ~20 different
Rho-GEFs, and any of them could potentially be activated by Ras, we
decided to assess total cellular Rho-GEF activity rather than assessing
individual Rho-GEFs. We used an assay analogous to that described for
measuring total cellular Ras-GEF activity (57, 58) and found no effect
of dexamethasone on total Rho-GEF activity in LTR-H-Ras(A) cells
(Fig. 6c, left panel). However, Rho-GEF activity
was 2.7-fold higher in extracts derived from wild type NIH 3T3 cells
transfected with the constitutively active Effect of Oncogenic Ras on Rho-GAP Activity and Subcellular
Localization--
Because we found no effect of oncogenic Ras on
Rho/Rho-GDI association and Rho-GEF activity, we hypothesized that Ras
could activate Rho through a decrease in Rho-GAP activity and/or a
change in the subcellular location of a Rho-GAP. A likely candidate for regulation by Ras is p190 Rho-GAP because this protein associates with
p120 Ras-GAP in a reversible, tyrosine
phosphorylation-dependent manner, and several observations
suggest that the p190 Rho-GAP·p120 Ras-GAP complex may function to
regulate actin cytoskeletal dynamics through RhoA (59, 60, 60-63).
We measured p190 Rho-GAP activity in immunoprecipitates prepared from
cytosolic extracts of cells by following the hydrolysis of
[
The basis for decreased cytosolic p190 Rho-GAP activity during
expression of oncogenic Ras could be a decrease in the amount of total
cellular p190 Rho-GAP, a shift in subcellular localization, or a
decrease in the specific activity of the protein, e.g.
because of decreased tyrosine phosphorylation (60, 64). We observed a
significant decrease in the amount of cytosolic p190 Rho-GAP in
dexamethasone-treated LTR-H-Ras(A) cells with no effect of dexamethasone in wild type NIH 3T3 cells (Fig. 7c,
cytosol, shows a representative experiment, and Fig.
7b summarizes the results of eight independent experiments).
We found no change in total cellular p190 Rho-GAP when LTR-H-Ras(A)
cells were treated with dexamethasone (Fig. 7c,
Homogenate), and there was only a slight increase in
membrane-associated p190 Rho-GAP in response to dexamethasone which was
not seen in wild type NIH 3T3 cells (Fig. 7c,
Membrane). To determine whether the decrease in cytosolic
Rho-GAP may be the result of a shift into a cytoskeletal compartment,
we extracted cells in Triton X-100-containing lysis buffer and examined
the detergent-insoluble cytoskeletal fraction (44). Dexamethasone treatment of LTR-H-Ras(A) cells, but not of wild type NIH 3T3 cells,
induced a significant increase in the amount of p190 Rho-GAP associated
with the Triton-insoluble cytoskeletal fraction (Fig. 7d).
We simultaneously assessed p120 Ras-GAP expression and found that total
and cytosolic p120 Ras-GAP were unaffected by dexamethasone in both
LTR-H-Ras(A) and wild type NIH 3T3 cells but that dexamethasone induced
a shift of p120 Ras-GAP to the membrane as has been described under
other conditions of Ras activation (Fig. 7c and Ref.
65).
Effect of Oncogenic Ras on the Interaction of p190 Rho-GAP with
p120 Ras-GAP and on p190 Rho-GAP Phosphorylation--
Because the
association of p190 Rho-GAP with p120 Ras-GAP is regulated by tyrosine
phosphorylation and may influence the specific Rho-GAP activity of p190
(60, 62), we examined the effect of oncogenic Ras expression on p190
tyrosine phosphorylation and association with p120. We
immunoprecipitated p190 Rho-GAP from cytosolic extracts and assessed
the immunoprecipitates for the presence of p120 Ras-GAP; we also
performed the reciprocal experiments, immunoprecipitating Ras-GAP and
looking for Rho-GAP. As would be expected from the decrease in
cytosolic p190 Rho-GAP, we found less p190 in anti-Rho-GAP
immunoprecipitates from dexamethasone-treated LTR-H-Ras(A) cells
compared with nontreated cells (Fig.
8a, compare lanes 4 and 5 with lanes 2 and 3).
Dexamethasone treatment appeared to cause a proportional decrease
in the amount of p120 Ras-GAP in the anti-Rho-GAP immunoprecipitates,
without causing any changes in cytosolic p120 Ras-GAP (Fig.
8a, compare lanes 4 and 5 with lanes 2 and 3, and lanes 8 and
9 with lanes 6 and 7). We also found
about 70% less p190 Rho-GAP in anti-Ras-GAP immunoprecipitates of
dexamethasone-treated cells, even though the amount of Ras-GAP in the
immunoprecipitates was constant in the presence and absence of
dexamethasone (Fig. 8b). In parallel experiments, we found no effect of dexamethasone on the amount of p190 Rho-GAP/p120 Ras-GAP
association in wild type NIH 3T3 cells (data not shown). We conclude
that induction of oncogenic Ras decreases the amount of p190
Rho-GAP·p120 Ras-GAP complex present in the cytosol of LTR-H-Ras(A)
cells in proportion to the decrease in cytosolic p190 Rho-GAP.
The amount of tyrosine-phosphorylated p190 Rho-GAP, both in
anti-Rho-GAP and anti-Ras-GAP immunoprecipitates, was not decreased in
dexamethasone-treated LTR-H-Ras(A) cells, suggesting that the remaining
p190 in dexamethasone-treated cells may actually be slightly more
tyrosine-phosphorylated compared with untreated cells (Fig.
8c shows anti-Rho-GAP immunoprecipitates). Thus, the decrease in the amount of p190 Rho-GAP·p120 Ras-GAP complexes appeared to be secondary to the shift in p190 Rho-GAP location and was
not associated with a detectable decrease in p190 Rho-GAP phosphotyrosine.
Effect of the Raf/MEK/MAP Kinase Pathway on
Ras-induced Rho Activation--
Multiple effector pathways contribute
to transformation by oncogenic Ras, but the function of the Raf/MEK/ERK
pathway is critical because activating mutants of Raf-1 and MEK are
necessary and sufficient for transformation of rodent fibroblasts (34,
66). We examined whether RhoA activation by oncogenic Ras required activation of the Raf/MEK/ERK pathway by using the specific MEK inhibitor U0126 (28). We found that dexamethasone treatment of
LTR-H-Ras(A) cells led to activation of the MAP kinases ERK-1/2, as
demonstrated by Western blotting using an antibody specific for the
dually phosphorylated, activated form of ERK-1/2 (Fig. 9a shows a 24 h time
point); the kinetics of ERK-1/2 activation closely follows the kinetics
of Ras(V12) induction (not shown). Dexamethasone had no effect on
ERK-1/2 activity in wild type NIH 3T3 cells (not shown). Treating
LTR-H-Ras(A) cells with 20 µM U0126 prevented
dexamethasone-induced ERK-1/2 activation (Fig. 9a). At the
same concentration, U0126 significantly inhibited Ras activation of Rho
when assessed both by the quantitative enzymatic assay (Fig.
3a) and by Western immunoblotting (Fig. 3b).
Similarly, U0126 significantly inhibited the effect of transiently
transfected Ras(12V) on Rho activity in wild type NIH 3T3 cells (Fig.
3c). The mechanism for the decrease in Rho activation by
U0126 appeared to be mediated, at least in part, by preventing the
dexamethasone-induced decrease in cytosolic p190 Rho-GAP (Fig.
9b) and increase in cytoskelton-associated p190 Rho-GAP
(Fig. 9c).
To determine whether activation of the Raf/MEK/ERK pathway is
sufficient to induce translocation of p190 Rho-GAP from the cytosol to
the cytoskeleton, we used constitutively active constructs of Raf-1
kinase (BXB) and MEK (MEK(Glu218,Asp222)) (27,
34). In wild type NIH 3T3 cells transfected with HA-tagged p190
Rho-GAP, significantly more HA-tagged Rho-GAP was found in the
detergent-insoluble cytoskeletal fraction when cells were cotransfected
with BXB than when cotransfected with empty vector (Fig.
9d). Similar results were found when endogenous p190 Rho-GAP was examined in the cytoskeletal fraction of cells transfected with BXB
or activated MEK(E218,D222); however, the
observed increases in cytoskeletal p190 Rho-GAP were less pronounced
because the transfection efficiency was only about 30% (Fig.
9e). Thus, the Ras/Raf/MEK/MAP kinase pathway appears to
regulate RhoA activity by regulating the translocation of p190 Rho-GAP
from the cytosol to a detergent-insoluble cytoskeletal fraction.
We found that increasing the amount of activated, oncogenic H-Ras
in NIH 3T3 cells increased RhoA activity. We used two different approaches, i.e. transient transfection and stable
transfection of an H-Ras(V12) construct under conditional promoter
control, and we measured Rho activation by two different assays,
i.e. Rhotekin pulldown with Western blotting for RhoA and
enzymatic quantitation of GTP bound to Rho. The mechanism of RhoA
activation by Ras involved redistribution of p190 Rho-GAP from the
cytosol to a detergent-insoluble cytoskeletal fraction with decreased
cytosolic Rho-GAP activity; the Ras-induced translocation of p190
Rho-GAP and increase in RhoA activity were prevented by pharmacologic
inhibition of the Raf/MEK/ERK pathway. The enzymatic assay for
measuring Rho activation is based on a similar assay we have used for
measuring Ras, Rap1, and Rheb activation (26, 38-40) and has several
noteworthy features: (i) it is quantitative and highly sensitive; (ii)
it provides a measure of the total amount of intracellular Rho; (iii)
it can be used for Rho expressed from transfected Rho constructs; and (iv) it can be used on human tumor samples that may have large variations in the amount of Rho present (4, 67). Using the enzymatic
assay, we have measured Ras activation in several different human
cancers (2, 68, 69), and work is in progress to measure Ras and Rho
activation in breast and prostate cancers. A potential problem is that
Rhotekin interacts with RhoA and C and may bind weakly to RhoB
(49); thus measuring GTP eluted from Rhotekin-bound Rho may provide a
mean activation of all three Rho subtypes. However, for the present
studies this was not a significant concern because NIH 3T3 cells
contain predominantly RhoA, with much less RhoB and C (37).
While the present work was in progress, several groups reported
increased Rho·GTP levels in different types of Ras-transformed cells,
although one group did not confirm this (8, 16, 18, 23). Our work
differs from these published results because we purposefully avoided
comparing Rho activities in cell lines with different clonal origins,
which may differ because of secondary genetic changes that accumulate
during selection in culture. Sahai et al. (8) stably
transfected Swiss 3T3 cells with H-Ras(V12) and found higher RhoA
activity and lower Rac activity in transformed versus
untransformed clones. The increased RhoA activity appeared to be the
result of long term selection of the Ras-transformed clones rather than
direct Ras signaling because Swiss 3T3 cells stably transfected with a
hormone-inducible, constitutively active Raf-1 construct demonstrated
an increase in Rho·GTP only after more than 4 weeks of selection in
the presence of the hormone; no change in RhoA activity after a 6-h
induction of active Raf-1 was observed, even though activation of the
Raf/MEK/ERK pathway was demonstrated (8). Zondag et al. (16)
transformed MDCK cells with v-H-Ras or constitutively active Raf and
found similarly increased RhoA and decreased Rac1 activity compared
with untransformed cells. In both Ras- and Raf-transformed Swiss 3T3
and MDCK cells, pharmacologic inhibition of MEK with U0126 or PD98059
was without effect on RhoA and Rac1 activities, further strengthening
the view that selective pressures rather than direct signaling events determined the levels of RhoA and Rac1 activity observed in these studies (8, 16). In contrast, we found that relatively short term
(24-36 h) expression of oncogenic H-Ras increased RhoA and Rac1
activity in NIH 3T3 cells, with U0126 preventing this effect at
concentrations at which it prevented activation of the Raf/MEK/ERK pathway. Our results are more compatible with the results of Gupta et al. (18), who found increased RhoA and Rac1 activities in human fibrosarcoma cells expressing mutant N-Ras and in wild type Ras-containing cells stably transfected with constitutively active Raf
or MEK compared with wild type Ras-containing cells transfected with
empty vector. These authors also found a partial decrease of RhoA and
Rac1 activity when the hyperactive Raf/MEK/ERK pathway in mutant
N-Ras-containing cells was blocked by transfection of dominant negative
Raf and MEK constructs (18). Interestingly, treatment of
v-H-Ras-transformed MDCK cells with the drug IND 12 inhibited activity
of the Raf/MEK/ERK pathway and restored the activity of RhoA and Rac1
to that found in untransformed MDCK cells (17). Taken together, these
results suggest that the increased RhoA activity found in
Ras-transformed cells is, at least in part, a consequence of increased
Raf/MEK/ERK signaling.
Although our work does not exclude the possibility that RhoA activation
by oncogenic Ras involves activation of one or multiple Rho-GEFs, we
did not find any detectable change in overall cellular Rho-GEF
activity. However, we observed a significant decrease in cytosolic p190
Rho-GAP activity which likely, at least partially, accounts for the
increased Rho·GTP levels in cells expressing H-Ras(V12). In murine
fibroblasts, p190 Rho-GAP accounts for the majority of Rho-GAP activity
in cell lysates, and inhibition of Rho-GAP activity is sufficient for
induction of RhoA-mediated actin reorganization (70). We found that
expression of oncogenic Ras in LTR-H-Ras(A) cells induced the
translocation of p190 Rho-GAP from the cytosol to a detergent-insoluble
cytoskeletal fraction. Similarly, integrin-mediated cell-substrate
interaction has been shown to result in p190 Rho-GAP recruitment to the
cytoskeleton at 1-2 h after plating NIH 3T3 cells on fibronectin, and
this change in subcellular localization of Rho-GAP occurs at a time when Rho·GTP levels are elevated (11, 36, 44). In contrast to the
stimulation of RhoA activity by prolonged integrin engagement, early
integrin signaling within 10-15 min after plating cells on fibronectin
induces Src-dependent tyrosine phosphorylation of p190
Rho-GAP, increases Rho-GAP activity, and transiently inactivates RhoA
(64). We found no definitive change in p190 Rho-GAP tyrosine phosphorylation after induction of oncogenic Ras. However, we found
that inhibition of the Raf/MEK/ERK pathway prevented the translocation
of p190 Rho-GAP from the cytosol to a cytoskeletal fraction, whereas
expression of a constitutively active Raf-1 or MEK was sufficient to
induce p190 Rho-GAP translocation. It is tempting to speculate that
redistribution of p190 Rho-GAP to the cytoskeletal fraction might be
associated with phosphorylation by ERK because p190 Rho-GAP contains
several potential ERK phosphorylation sites (35). Future work will be
to evaluate fully the p190 Rho-GAP phosphorylation status in
Ras-transformed cells.
Many Ras- or Raf-transformed cells show a loss of stress fibers
compared with their untransformed counterparts (7, 8, 18, 23, 24), and
dexamethasone treatment of LTR-H-Ras(A) cells is associated with a
dramatic loss of stress fibers in apparent contradiction to the
increased Rho·GTP levels found in these cells (71). Prolonged
activation of the Raf/MEK/ERK pathway appears to suppress the ability
of active RhoA to promote stress fiber formation because treating
Ras-transformed cells with pharmacologic MEK inhibitors or transfecting
a dominant negative MEK can restore stress fibers to basal levels found
in untransformed cells and lead to morphological reversion (8, 16, 23).
The effect of MEK inhibition on stress fibers has been explained by the
finding that MEK inhibitors prevent translocation of the RhoA effector ROCK from the cytosol to a detergent-insoluble cytoskeletal fraction in
Ras-transformed cells (8). The mechanism by which prolonged activation
of the Raf/MEK/ERK pathway leads to translocation of ROCK has not been
resolved, but it might be similar to the mechanism by which oncogenic
Ras leads to the redistribution of p190 Rho-GAP.
Our analysis of p21WAF1/CIP1 promoter activity in
LTR-H-Ras(A) cells demonstrated that Rho activation by Ras had a
physiological effect with downstream changes in gene transcription. The
opposing function of Ras and Rho in the regulation of the
p21WAF1/CIP1 promoter appears to be important for Ras
transformation because when Rho signaling is blocked, oncogenic Ras
induces cell cycle arrest in p21WAF1/CIP1-expressing, but
not in p21WAF1/CIP1-deficient cells (10). In pancreatic
carcinoma cells expressing oncogenic Ras, a dominant negative RhoA, but
not Rac1, was able to activate the p21WAF1/CIP1 promoter,
whereas a constitutively active RhoA suppressed it, similar to our
findings in LTR-H-Ras(A) cells (72).
In conclusion, our work demonstrates direct regulation of RhoA activity
by oncogenic Ras and offers an explanation for the increased RhoA
activation found in Ras-transformed cells. Further work is needed to
understand the mechanism(s) whereby the Ras/Raf/MEK pathway induces
redistribution of p190 Rho-GAP and ROCK (8) from the cytosol to a
cytoskeletal fraction.
INTRODUCTION
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
EXPERIMENTAL PROCEDURES
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
N-p115Rho-GEF from M. Hart (31); p21-Luc from X.-F. Wang
(32); pEF-C3exo from R. Treisman (33);
pMEK1(E218,D222) from S. Cowley (34),
courtesy of P. M. McDonough; and pHA-p190Rho-GAP from J. Settleman
(35).
-galactosidase, pCMV-
-galactosidase, pSV40-
-galactosidase, and pTK-
-galactosidase, demonstrated some increase in transcription when LTR-H-Ras(A) cells were treated with dexamethasone.
-32PO4]GTP
(specific activity 6,000 Ci/mmol) was added to initiate the reaction
(45). Tubes were shaken vigorously in a water bath at 20 °C, and at
the indicated times the reaction was stopped by transferring 10 µl of the reaction mixture to 1 ml of ice-cold stop buffer
containing 50 mM Tris-HCl, pH 7.5, 50 mM NaCl,
5 mM MgCl2, 1 mM dithiothreitol.
Samples were collected on nitrocellulose filters, which were washed and
dried overnight. Cerenkov radiation was measured in a scintillation
counter, and data were expressed as a percentage of the amount of
[
-32PO4]GTP bound to RhoA in the
zero time samples.
RESULTS
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
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Fig. 1.
Quantitative measurement of Rho
activation. Varying numbers of logarithmically growing NIH 3T3
cells cultured in DMEM supplemented with 10% FBS were extracted
in situ. From one aliquot of the extracts, Rho·GTP was
isolated directly on glutathione-agarose beads using GST-tagged
Rhotekin RBD (unloaded samples, a and b); to
another aliquot of the extracts 10 mM EDTA plus 10 µM GTP were added prior to adding the Rhotekin RBD
(loaded samples, c and d). GTP eluted from the
washed beads was measured in a coupled enzymatic assay as described
under "Experimental Procedures" (a and c), or
RhoA protein isolated on the beads was assessed by Western blotting
(b and d) with lanes 1-5
corresponding to the five protein concentrations in a and
c. The data in a and c are the means
of duplicates of a representative experiment, with similar results
found in two other independent assays; the asterisks,
pound sign, and double dagger indicate paired
unloaded and loaded samples.
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Fig. 2.
Serum stimulation of endogenous Rho activity
and activation levels of transfected wild type and constitutively
active RhoA constructs. a and b, NIH 3T3 cells
were serum starved for 36 h in DMEM containing 0.1% FBS and 0.1%
bovine serum albumin followed by stimulation with 10% FBS for the
indicated time. Cells were extracted in situ, and Rho·GTP
was isolated by RBD-Rhotekin pulldown as described under
"Experimental Procedures." In a, the amount of GTP and
total nucleotides bound to Rho was measured enzymatically as described
in the legend to Fig. 1. In b, the amount of RhoA·GTP
bound to the beads was quantitated by Western immunoblotting using a
mouse monoclonal anti-RhoA antibody (upper blot); to
demonstrate the input of total RhoA protein, 5% of cellular lysate was
analyzed by Western blotting using the same antibody (lower
blot). c, NIH 3T3 cells grown in six-well culture
dishes were transiently transfected with 400 ng of EE epitope-tagged
RhoA constructs (wild type RhoA (Rho wt) and mutant
RhoA(63L) or RhoA(V14)) as described under "Experimental
Procedures." Some experiments with wild type Rho included 500 ng of
an expression vector encoding constitutively active N-p115-Rho-GEF.
Cells were extracted 36 h later, and Rho proteins were isolated on
protein G-agarose beads using an anti-EE antibody. To some samples 80 µg of soluble Rhotekin RBD peptide was added immediately after
extraction. Rho activation was measured enzymatically as described in
the legend to Fig. 1. d, NIH 3T3 cells were transfected as
described in c, and whole cell extracts were analyzed by
SDS-PAGE/Western blotting using a monoclonal antibody specific for
RhoA. Lane 1, empty vector; lane 2, EE-RhoA(wt);
lane 3, EE-RhoA(63L); lane 4, EE-RhoA(V14).
Transfection efficiency was about 50%. Reprobing the blot with an
anti-EE epitope antibody confirmed that the EE-tagged RhoA proteins
migrate with a higher apparent molecular weight than endogenous RhoA
(not shown); the 63L and V14 point mutations are known to alter the
migration of RhoA differently (73). The data in a and
c are the means ± S.D. of at least three independent
experiments performed in duplicate.
N-p115-Rho-GEF with wild type RhoA, we found that Rho activation
increased to 15.1 ± 2.2% (Fig. 2c). Although this
increase in Rho activation is similar to that observed in
serum-stimulated cells (Fig. 1b), it is considerably less
than the activation state of the constitutively activated,
GTPase-deficient forms of RhoA. Because it seemed possible that the
lower activation level of wild type RhoA, compared with the
constitutively activated RhoA, was from its higher intrinsic GTPase
activity leading to a loss of GTP during immunoprecipitation, we
performed experiments in which we added soluble Rhotekin RBD to cell
extracts to protect Rho·GTP; binding of the RBD of Rhotekin to Rho
inhibits both intrinsic and GAP-enhanced Rho GTPase activity (49). We
found no effect of the peptide on Rho activation, either at basal
levels or in the presence of p115-Rho-GEF (Fig. 2c),
suggesting little loss of GTP during the assay procedure. In Fig.
2d we show the expression of the three RhoA constructs, with
wild type EE-RhoA and RhoA(14V) expressed at somewhat higher and
RhoA(63L) at somewhat lower levels, than endogenous RhoA, considering a
transfection efficiency of about 50%.
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Fig. 3.
Measurement of Rho activation in LTR-H-Ras(A)
cells and in wild type NIH 3T3 cells transiently transfected with
H-Ras(V12). Wild type NIH 3T3 cells (NIH 3T3 wt) and
stably transfected NIH 3T3 cells expressing H-Ras(V12) under control of
the inducible murine mammary tumor virus promoter (LTR-H-Ras(A) cells)
were cultured in the absence or presence of 1 µM
dexamethasone (Dex) for 24 h; some of the LTR-H-Ras(A)
cultures were also treated with the MEK inhibitor U0126 (20 µM) for 24 h as indicated. a, Rho
activation was measured by a quantitative enzymatic assay as described
in the legend to Fig. 1; open bars, without dexamethasone;
filled bars, with dexamethasone. Rho activation in the
absence of drugs was assigned a value of 1 for both cell types. For the
LTR-H-Ras(A) cells, the asterisk indicates a statistically
significant difference (p < 0.01) between the absence
and presence of dexamethasone, and the pound sign designates
a statistically significant difference (p < 0.05)
between the absence and presence of U0126 in dexamethasone-treated
cells. b, Rho·GTP (top blot), total cellular
RhoA (middle blot), and Ras (bottom blot) were
assessed by Western immunoblotting as described in Fig. 2c;
for total RhoA and Ras, 5% of the cellular lysate was used.
Lanes 1 and 2 are wild type NIH 3T3 cells
cultured in the absence and presence of dexamethasone, and lanes
3-6 are LTR-H-Ras(A) cells cultured in the absence and presence
of dexamethasone and U0126 as indicated. c, wild type NIH
3T3 cells were transiently transfected with EE epitope-tagged RhoA(wt)
as described in the legend to Fig. 2c and were cotransfected
with either 700 ng of empty vector or expression vectors for wild type
Ras (Ras wt) or Ras(12V). During the last 24 h before
harvest, 20 µM U0126 was added as indicated. Rho
activation was measured as described in Fig. 2c. The
asterisk indicates a significant difference
(p < 0.05) between the absence and presence of
U0126.
Ras activation state and Ras expression in wild type NIH 3T3 and
LTR-H-Ras(A) cells
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Fig. 4.
Effect of Ras-induced Rho activation on
the p21WAF1/CIP1
promoter. LTR-H-Ras(A) cells were cotransfected with p21-Luc
(a luciferase reporter gene under control of the
p21WAF1/CIP1 promoter) and empty
vector, expression vectors encoding C3 exozyme, or activated RhoA
(RhoA(63L)) as described under "Experimental Procedures." After a
24-h culture in DMEM containing 10% FBS in the absence (open
bars) or presence of 1 µM dexamethasone
(filled bars), luciferase activity was measured as described
under "Experimental Procedures." The luciferase activity obtained
in untreated cells transfected with empty vector was assigned a value
of 1; the data are the means ± S.D. of at least three independent
experiments performed in duplicate.
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Fig. 5.
Effect of oncogenic Ras on Rac·GTP
Levels. LTR-H-Ras(A) cells (a) and wild type NIH 3T3
cells (b) were cultured for 24 h in DMEM containing
10% FBS in the absence or presence 1 µM dexamethasone
(Dex) as indicated. Cells were extracted, and Rac·GTP was
isolated using the CDC42/Rac binding domain of Pak bound to glutathione
beads as described under "Experimental Procedures." Rac·GTP bound
to the beads (left panels) and total Rac present in 5% of
the input cell lysate (right panels) were detected by
Western blotting using a Rac-specific antibody. Ctrl refers
to cell lysates incubated with GST-loaded glutathione beads lacking the
CDC42/Rac binding domain of Pak. c, Rac·GTP was isolated
on beads and detected as described above; the Western blots for
Rac·GTP were scanned, and the band intensity obtained with untreated
cells was assigned the value of 1. The data represent the means ± S.D. of five independent experiments.
S to cell
lysates (52). We assessed the amount of RhoA associated with membrane
fractions in wild type NIH 3T3 cells and LTR-H-Ras(A) cells in the
absence and presence of dexamethasone. We found no change in the amount
of RhoA in membranes of wild type cells, but there was an approximately
2-fold increase in membrane-bound RhoA in dexamethasone-treated
LTR-H-Ras(A) cells (Fig. 6a,
upper blots). Wild type NIH 3T3 cells appeared to express
slightly higher amounts of RhoA protein; we do not know the
significance of this finding because it may simply reflect the effects
of clonal evolution in culture. The increased membrane association of
RhoA in dexamethasone-treated LTR-H-Ras(A) cells correlated well with
the 2-fold increase in Rho activity found under these conditions (Fig.
3a); because the increase in membrane-bound RhoA represented
only a small fraction of total cellular RhoA, it is not surprising that
we did not detect any significant change in the amount of RhoA in the
cytosol (Fig. 6a, lower blots; ~2.5% of the
cytosolic fraction compared with 50% of the membrane fraction was
loaded on the gels).
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Fig. 6.
Effect of oncogenic Ras on the subcellular
location of RhoA, RhoA association with Rho-GDI, and on Rho-GEF
Activity. Wild type (WT) NIH 3T3 cells (a
and c) and LTR-H-Ras(A) cells (a-c) were
cultured for 24 h in DMEM containing 10% FBS; as indicated, some
cells were treated with 1 µM dexamethasone
(Dex). a, cells were extracted in hypotonic lysis
buffer by Dounce homogenization, and cytosol and membrane fractions
were generated by differential centrifugation as described under
"Experimental Procedures." The two fractions were subjected to
SDS-PAGE/Western blotting using a mouse monoclonal anti-RhoA antibody.
b, cytosolic extracts were prepared and subjected to
immunoprecipitation using either rabbit control immunoglobulin
(C, lane 1) or a rabbit polyclonal anti-Rho-GDI
antibody (lanes 2-5) as described under "Experimental
Procedures." Immunoprecipitates were analyzed by SDS-PAGE/Western
blotting using a mouse monoclonal anti-RhoA antibody (upper
blot) and the rabbit anti-Rho-GDI antibody (lower
blot). For comparison, 5% of the cytosolic extracts were analyzed
in parallel (lanes 6-10). c, In the left
panel, LTR-H-Ras(A) cells were extracted in a buffer containing
1% Triton X-100, the extracts were centrifuged, and to the
supernatants were added 1 mM GTP and GST-tagged RhoA
preloaded with [3H]GDP that was bound to glutathione
beads. After 10 and 20 min, the reaction was stopped, and radioactivity
remaining on the washed beads was quantitated by liquid scintillation
counting as described under "Experimental Procedures." The data are
expressed as the percent increase in [3H]GDP exchange
compared with control beads incubated with extract buffer. In the
right panel, wild type NIH 3T3 cells were transfected with
either empty vector or a vector encoding constitutively active
N-p115-Rho-GEF; Rho-GEF activity was quantitated as described for
the left panel, and the activity found in cells transfected
with empty vector was assigned a value of 1. The data are the
means ± S.D. of three independent experiments performed in
duplicate.
N-p115-Rho-GEF compared
with extracts from cells transfected with empty vector (Fig.
6c, right panel). Although these studies do not
rule out a small change in an individual Rho-GEF, they suggest that
oncogenic Ras does not induce major changes in total Rho-GEF activity.
-32PO4]GTP preloaded on RhoA (62).
We found significantly less activity in extracts from
dexamethasone-treated LTR-H-Ras(A) cells compared with extracts from
nontreated cells (p < 0.05), whereas dexamethasone had
no effect on p190 Rho-GAP activity in wild type NIH 3T3 cells (Fig.
7a shows results for
LTR-H-Ras(A) cells; at 12 min, RhoA incubated with p190 Rho-GAP
immunoprecipitates from dexamethasone-treated cells had 24% more
bound GTP remaining compared with RhoA incubated with
immunoprecipitates from untreated cells).
View larger version (20K):
[in a new window]
Fig. 7.
Effect of oncogenic Ras activation on p190
Rho-GAP activity and subcellular localization. LTR-H-Ras(A) cells
(a-d) and wild type NIH 3T3 cells (WT,
b-d) were cultured for 24 h in DMEM containing 10%
FBS in the absence or presence of 1 µM dexamethasone
(Dex). a, cells were extracted in hypotonic
buffer by Dounce homogenization as described in Fig. 6a.
From the cytosolic fraction, p190 Rho-GAP was immunoprecipitated using
a mouse monoclonal antibody (filled diamonds, Rho-GAP
immunoprecipitated from nontreated cells, and filled
squares, Rho-GAP immunoprecipitated from dexamethasone-treated
cells); control immunoprecipitates were obtained using mouse control
immunoglobulin (open circles). Rho-GAP activity was
determined as described under "Experimental Procedures" by
incubating the washed immunoprecipitates with purified GST-RhoA
preloaded with [ -32PO4]GTP.
At the indicated times, the reaction was stopped by collecting proteins on nitrocellulose filters; radioactivity
bound to RhoA was measured by liquid scintillation counting. The data
are the means ± S.D. of five independent experiments performed in
duplicate and are expressed as the percent of GTP bound to RhoA
remaining at the indicated time. Note the intrinsic GTPase activity of
RhoA measured in the presence of control IgG immunoprecipitate
(open circles). b, cells were cultured in the
absence (open bars) or presence (filled bars) of
dexamethasone, and cytosolic extracts were prepared as described above.
Equal amounts of extract protein were analyzed by SDS-PAGE/Western
blotting using the anti-p190 Rho-GAP antibody, and signal intensities
were determined by scanning autoradiographs within the linear range of
exposure. The amount of p190 Rho-GAP detected in untreated cells was
assigned a value of 100%. The data represent the means ± S.D. of
six independent experiments. c, cells were extracted, and
subcellular fractions were generated as described in Fig.
6a. Equal amounts of extract protein were analyzed by
SDS-PAGE/Western blotting using mouse monoclonal antibodies specific
for p190 Rho-GAP and p120 Ras-GAP. d, cells were extracted
in situ by incubation in a Triton X-100-containing buffer;
the remaining Triton-insoluble cytoskeletal structures were collected
and solubilized in SDS-sample buffer as described under "Experimental
Procedures." The Triton-insoluble fractions from equal numbers of
cells were analyzed by Western blotting using a p190 Rho-GAP-specific
antibody (upper blot). Duplicate samples were blotted with
an actin-specific antibody (lower blot). Results similar to those shown
in c and d were obtained in three other
experiments.
View larger version (21K):
[in a new window]
Fig. 8.
Effect of oncogenic Ras on the interaction of
p190 Rho-GAP with p120 Ras-GAP and on p190 Rho-GAP
phosphorylation. LTR-H-Ras(A) cells were cultured for 24 h in
DMEM containing 10% FBS in the absence or presence of 1 µM dexamethasone (Dex). Cytosolic extracts
were prepared and subjected to immunoprecipitation using specific
antibodies for p190 Rho-GAP (a c) or p120 Ras-GAP
(b). a, Anti-Rho-GAP immunoprecipitates
(lanes 2-5) or 5% of the cytosolic input (lanes
6-9) were analyzed by SDS-PAGE/Western blotting using
anti-Rho-GAP and anti-Ras-GAP antibodies; lane 1 shows
immunoprecipitates obtained with control mouse IgG. b,
anti-Ras-GAP immunoprecipitates were analyzed by Western blotting using
the same antibodies as in a. Rho-GAP and Ras-GAP bands were
analyzed by scanning densitometry, and the ratio of Rho-GAP to Ras-GAP
found in untreated cells was assigned a value of 1. The amount of
immunoprecipitated Ras-GAP was the same in the absence and presence of
dexamethasone. The data are the means ± S.D. of two experiments
performed in duplicate and scanned at two different exposures.
c, anti-Rho-GAP immunoprecipitates were analyzed by Western
blotting using a phosphotyrosine-specific antibody (upper
panel) or an anti-Rho-GAP antibody (lower panel).
Results similar to those shown in a and c were
obtained in two other experiments.
View larger version (23K):
[in a new window]
Fig. 9.
Effect of the MEK inhibitor U0126 on
ERK phosphorylation and subcellular location of p190 Rho-GAP.
LTR-H-Ras(A) cells (a-c) and wild type NIH 3T3 cells
(d and e) were cultured as described in the
legend to Fig. 8, but some cells were treated with 20 µM
U0126 for 24 h as indicated. a, cells were extracted
in situ in SDS-sample buffer, and whole cell extracts were
analyzed by Western blotting using an anti-phospho-ERK-specific
antibody (upper blot) or an antibody that recognizes ERK-1/2
irrespective of its phosphorylation state (lower blot).
b, cytosolic extracts were prepared as described in Fig.
7c, and duplicate Western blots were analyzed using
antibodies specific for p190 Rho-GAP (upper blot) or actin
(lower blot). c, Triton X-100-insoluble
cytoskeletal fractions were isolated as described in Fig.
7d, and Western blots were generated as described above.
Similar results were obtained in two other experiments. d,
cells were transfected with HA-tagged p190 Rho-GAP and either empty
vector (lanes 1 and 2) or vector encoding the
constitutively active Raf-1 kinase BXB (lanes 3 and
4). Cytoskeletal fractions were isolated as described in
Fig. 7d, and Western blots were developed with an anti-HA
antibody (upper panel) and reprobed with an anti-actin
antibody (lower panel). e, cells were transfected
with empty vector (lanes 1 and 2), BXB
(lanes 3 and 4), or a constitutively active MEK
(E218,D222) (lanes 5 and
6). Western blots of cytoskeletal fractions were analyzed
using antibodies against endogenous p190 Rho-GAP (upper
panel) or actin (lower panel). Transfection efficiency
was about 30% by -galactosidase staining (27).
DISCUSSION
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
![]() |
ACKNOWLEDGEMENTS |
---|
We thank S. Cowley, P. M. McDonough, M. Hart, R. Treisman, M. A. Schwartz, J. Settleman, X.-F. Wang, and M. Wigler for providing valuable DNA constructs.
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FOOTNOTES |
---|
* This work was supported in part by United States Public Health Service Grants GM55586 (to R. B. P.) and CA89828 and CA90932 (to G. R. B.) and by a University of California Cancer Coordinating Committee grant (to R. B. P.).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.
Both authors contributed equally to this work.
§ To whom correspondence should be addressed: Dept. of Medicine, University of California at San Diego, 9500 Gilman Dr., Basic Science Bldg. 5080, La Jolla, CA 92093-0652. Tel.: 858-534-8805; Fax: 858-534-1421; E-mail: rpilz@ucsd.edu.
Published, JBC Papers in Press, November 11, 2002, DOI 10.1074/jbc.M207943200
2 F. C. von Lintig and G. R. Boss, unpublished observation.
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ABBREVIATIONS |
---|
The abbreviations used are:
MEK, mitogen-activated protein kinase/extracellular signal-regulated kinase
kinase;
CHAPS, 3-[(3-cholamidopropyl)dimethylammonio]-1-propanesulfonic acid;
CMV, cytomegalovirus;
DMEM, Dulbecco's modified Eagle's medium;
ERK, extracellular signal-regulated kinase;
FBS, fetal bovine serum;
GAP, GTPase-activating protein;
GDI, guanine dissociation inhibitor;
GEF, guanine nucleotide exchange factor;
GST, glutathione
S-transferase;
GTPS, guanosine
5'-O-(thiotriphosphate);
LTR, long terminal repeat;
Luc, luciferase;
MAP, mitogen-activated protein;
MDCK, Madin-Darby canine
kidney;
RBD, Rho binding domain;
RSV, Rous sarcoma virus;
TK, thymidine
kinase;
BXB, catalytic domain of Raf-1 kinase.
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