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Molecular and Cellular Biology, June 2003, p. 4283-4294, Vol. 23, No. 12
0270-7306/03/$08.00+0     DOI: 10.1128/MCB.23.12.4283-4294.2003
Copyright © 2003, American Society for Microbiology. All Rights Reserved.

Effects of Rho Kinase and Actin Stress Fibers on Sustained Extracellular Signal-Regulated Kinase Activity and Activation of G1 Phase Cyclin-Dependent Kinases

Kristin Roovers and Richard K. Assoian*

Department of Pharmacology, University of Pennsylvania School of Medicine, Philadelphia, Pennsylvania 19104-6084

Received 17 December 2002/ Returned for modification 27 January 2003/ Accepted 18 March 2003


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ABSTRACT
 
We recently reported that Rho kinase is required for sustained ERK signaling and the consequent mid-G1 phase induction of cyclin D1 in fibroblasts. The results presented here indicate that these Rho kinase effects are mediated by the formation of stress fibers and the consequent clustering of {alpha}5ß1 integrin. Mechanistically, {alpha}5ß1 signaling and stress fiber formation allowed for the sustained activation of MEK, and this effect was mediated upstream of Ras-GTP loading. Interestingly, disruption of stress fibers with ML-7 led to G1 phase arrest while comparable disruption of stress fibers with Y27632 (an inhibitor of Rho kinase) or dominant-negative Rho kinase led to a more rapid progression through G1 phase. Inhibition of either MLCK or Rho kinase blocked sustained ERK signaling, but only Rho kinase inhibition allowed for the induction of cyclin D1 and activation of cdk4 via Rac/Cdc42. The levels of cyclin E, cdk2, and their major inhibitors, p21cip1 and p27kip1, were not affected by inhibition of MLCK or Rho kinase. Overall, our results indicate that Rho kinase-dependent stress fiber formation is required for sustained activation of the MEK/ERK pathway and the mid-G1 phase induction of cyclin D1, but not for other aspects of cdk4 or cdk2 activation. They also emphasize that G1 phase cell cycle progression in fibroblasts does not require stress fibers if Rac/Cdc42 signaling is allowed to induce cyclin D1.


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INTRODUCTION
 
Much like cells deprived of growth factors or attachment to an extracellular matrix (ECM), fibroblasts cultured under conditions that preclude cell spreading become arrested in G1 phase of the cell cycle (14, 21, 26, 33, 36). The cyclin-dependent kinases (cdk's), cyclin D-cdk4 (or cdk6) and cyclin E-cdk2, are the key regulators of cell cycle progression through G1 phase, and several studies now indicate that disruption of cell spreading prevents the activation of these enzymes. Pharmacological inhibitors of actin polymerization block the induction of cyclin D1 (typically the rate-limiting step in formation of active cyclin D1-cdk4 or -cdk6 complexes), the downregulation of p21cip1 and p27kip1 (inhibitors of cyclin E-cdk2), phosphorylation of the retinoblastoma protein, and entry into S phase (10, 11, 22, 31, 32). In endothelial cells, spreading is associated with the translation of cyclin D1 mRNA, the downregulation of p27kip1, and S phase entry (31, 44). Similarly, when fibroblasts are cultured within collagen gels, the disruption of isometric tension leads to loss of actin stress fibers, the inactivation of extracellular signal-regulated kinases (ERKs), the loss of cyclin D1, the upregulation of p27kip1, and G1 phase cell cycle arrest (22, 62). Conversely, mechanical tension stimulates focal adhesion kinase (FAK) phosphorylation (69, 81), which can induce cyclin D1 and downregulate p21cip1 (84). These kinds of data have lent support to the idea that cell shape-dependent G1 phase cell cycle progression is mediated, at least in part, by stress fibers and the generation of isometric tension. However, cultured epithelial cells readily proliferate without detectable stress fibers and, except for endothelial cells and wound fibroblasts, stress fibers are not generally detected in vivo (27, 70, 78, 79). Thus, even if stress fibers have a role in G1 phase progression, there must be signaling pathways that allow G1 phase progression to proceed in the absence of stress fibers.

The Rho-Rho kinase pathway is required for stress fiber and focal adhesion formation (9, 17, 25). Rho kinase catalyzes the inhibitory phosphorylation of myosin phosphatase (38), and the resulting increase in steady-state myosin light-chain (MLC) phosphorylation by MLC kinase (MLCK) promotes both myosin filament assembly and actin-activated myosin ATPase activity (12). MLC has also been identified as a direct substrate of Rho kinase (4, 71). Independent of its effects on MLC, Rho kinase catalyzes the activating phosphorylation of LIM kinase (LIMK) (49, 72), which, in turn, phosphorylates cofilin on Ser-3. The phosphorylation of cofilin inhibits its ability to depolymerize f-actin (2, 6, 43, 80). Activation of mDia and PIP4-5 kinase by Rho and Rho kinase also contribute to stress fiber formation through their stimulatory effects on actin polymerization (37, 73).

Burridge and coworkers have proposed that RhoA promotes stress fiber and focal adhesion formation by stimulating actin-myosin contractility, which in turn, generates tensional forces that cluster integrins (16). These studies place the effect of RhoA-dependent stress fiber formation upstream of integrins. However, RhoA can also act downstream of integrins: the GTP-loading of RhoA is transiently inhibited (0 to 15 min) and then stimulated when cells are plated on fibronectin (55). Rho-GTP levels then gradually decline over a 1- to 3-h period. How this complex activation pattern affects Rho effectors such as Rho kinase remains to be explored.

Rho and Rho kinase also plays important roles in G1 phase cell cycle progression. Inhibition of Rho activity results in the upregulation of both p21cip1 and p27kip1 (1, 7, 30, 40, 51, 76). We recently reported that Rho has a dual function in regulating cyclin D1 gene expression in NIH 3T3 cells: it allows for sustained ERK activity (defined as activity occurring between 3 and 9 h after mitogenic stimulation of quiescent cells and required for mid-G1 phase induction of cyclin D1 mRNA) while suppressing an alternative Rac/Cdc42 pathway that leads to an early G1 phase induction of cyclin D1 mRNA (77). In some reports, inhibition of Rho prevents the expression of cyclin D1 altogether (19, 26), presumably because Rac/Cdc42 signaling to cyclin D1 mRNA is absent. Rac also stimulates the translation of cyclin D1 mRNA, at least in endothelial cells (44). The effects of Rho on cyclin D1 induction are largely due to activation of Rho kinase (77), whereas the Rho effector(s) that regulate p21cip1 and p27kip1 are not yet clear (35, 63, 64, 66).

We examine here the mechanism by which Rho kinase regulates G1 phase cell cycle progression. We conclude that Rho kinase-dependent stress fiber formation is required for the sustained activation of ERK and mid-G1 phase of cyclin D1 but not for the expression of cdk4, cdk2, cyclin E, p21cip1, or p27kip1. Rac/Cdc42-dependent induction of cyclin D1 is similarly independent of stress fiber formation. Our data identify the molecular event that underlies stress fiber-dependent G1 phase progression and also provide a mechanism for explaining how G1 phase progression can occur without stress fibers.


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MATERIALS AND METHODS
 
Cell culture, transient transfections, and drug treatments. NIH 3T3 cells stably expressing {alpha}5human ß1mouse chimeric integrin (h{alpha}5-3T3) have been characterized previously, and the results showed that expression of the chimeric integrin had no effect on the rate of G1 phase progression or on the cooperative signaling between growth factor receptor tyrosine kinases (RTKs) and integrins (61). Near-confluent h{alpha}5-3T3 cells were serum starved as described previously (77). The starved cells (1.5 x 106 cells) were treated with trypsin, suspended in 10 ml of defined medium (61), and pretreated in suspension (30 min at 37°C) with pharmacological inhibitors: Y27632 (10 µM; Tocris), ML-7 (10 µM; Biomol), U0126 (50 µM; Promega), or dimethyl sulfoxide (DMSO; control). The pretreated cells were then stimulated with basic fibroblast growth factor (bFGF; 10 ng/ml [final concentration]) and immediately plated on 100-mm culture dishes that had been coated with fibronectin (0.1 mg/10 ml), anti-{alpha}5ß1 (1.2 mg/10 ml), poly-L-lysine (0.25 mg/10 ml), or bovine serum albumin (BSA; 1 mg/ml) in the continued presence of the inhibitor. Established mouse embryo fibroblasts (MEFs) were similarly serum starved (see individual figure legends for times) and pretreated with the pharmacological inhibitors, except that the preincubations used 106 cells (in 10 ml of Dulbecco modified Eagle medium [DMEM]-BSA [1 mg/ml]). The pretreated cells were then stimulated with fetal bovine serum (FBS; 10% final concentration) and immediately plated on uncoated 100-mm culture dishes. MEFs stably expressing tetracycline-repressible cyclin D1 were maintained in 5% FBS with tetracycline added daily at a final concentration of 2 µg/ml. Controls with anti-phospho-MLC (gift of Fumio Matsumura) showed that 10 µM Y27632 and 10 µM ML-7 completely blocked phosphorylation of MLC (not shown). In other experiments, h{alpha}5-3T3 cells or MEFs (60 to 70% confluent) were transiently transfected, serum-starved, and plated at subconfluence in DMEM-10% FBS (MEFs) or in defined medium on fibronectin-coated dishes with 10 ng of bFGF/ml (h{alpha}5-3T3 cells) as described previously (77). Transfected plasmids encoded dominant-negative (CAT-KD) Rho kinase (3), dominant-negative (T508A) LIMK (49), dominant-negative FAK, FRNK (24, 59), p21-binding domain of PAK (PBD), constitutively active MEK1 (S218D/S222D), constitutively active Rac (Q61L), or pCDNA3 (empty vector). Transfection efficiencies were 75 to 85% as determined by immunofluorescence analysis (e.g., with antibodies to the epitope tag).

Immunoblotting. Immunoblotting was performed on extracted cell pellets as described previously (61) with antibodies specific for ERK (Transduction Lab catalog no. M12320), dually phosphorylated ERK (pERK; Cell Signaling [9101S]), FAK (Santa Cruz [sc-558]), pY397 FAK (BioSource [44-624]), MEK1 (Transduction Lab [M17020]), Ras (UBI [05-516]), Raf-1 (Santa Cruz [sc-133]), pS222 MEK (BioSource [44-452]), cyclin D1 (Santa Cruz [sc-8396]), cyclin E (Santa Cruz [sc-481]), p21cip1 (Santa Cruz [sc-6246]), p27kip1 (Transduction Lab [K25020]), cdk2 (UBI, 06-505), cdk4 (Santa Cruz [sc-260]), myc (9E10), and glutathione S-transferase (GST; a gift of Margaret Chou). Each figure panel shows results of enhanced chemiluminescence from the same filters with comparable exposure times.

In vitro Ras, Raf-1, MEK, and ERK assays. Ras activation assays were performed as suggested by the manufacturer (UBI) except that 500 µg of cell lysate in ~0.1 ml was incubated with 10 µg of the Raf-1 Ras-binding domain-bead conjugate. The beads bound to active Ras (GTP loaded) were washed twice. To assess total Ras levels, 50 µg of the lysate was fractionated on a reducing sodium dodecyl sulfate (SDS)-gel and immunoblotted with anti-Ras. Comparison of total and pull-downed Ras showed that ca. 10 to 20% of total Ras was GTP loaded.

For Raf-1 kinase assays, frozen cell pellets (~3 x 106 cells) were lysed in 0.1 ml of 50 mM Tris (pH 7.5), 150 mM NaCl, 5 mM EDTA, 1 mM dithiothreitol (DTT), 1% Triton X-100, 0.5% deoxycholate, and 0.1% SDS with protease inhibitors (10 µg of aprotinin/ml, 10 µg of leupeptin/ml, 5 mM NaF, 10 mM Na3VO4). Cell lysates (250 µg) were incubated (2 h, 4°C) with 3 µg of anti-Raf-1 and then collected (2 h at 4°C with rocking) with 50 µl of washed protein A-agarose (Invitrogen). Collected immunoprecipitates were washed once with cold lysis buffer and then four times with cold kinase reaction buffer (50 mM Tris-HCl [pH 8.0], 10 mM MgCl2). Washed immunoprecipitates were suspended in 50 µl of kinase reaction buffer containing protease inhibitors (see above), 1 mM DTT, 4 µg of GST-MEK (UBI), and 50 µM ATP and then incubated at room temperature for 30 min. Kinase reactions were stopped with an equal volume of 2x SDS sample buffer; the samples were fractionated on reducing SDS-gels and transferred to nitrocellulose membranes. Nitrocellulose membranes were immunoblotted with anti-pS222 MEK to assess in vitro Raf-1 activity. To assess total Raf-1 levels, 75 µg of the lysate was fractionated on a reducing SDS-gel and immunoblotted with anti-Raf-1. Controls demonstrated that the immunoprecipitation depleted more than 80% of the total Raf-1 and that the kinase assay was in the linear range between 100 to 500 µg of cell lysate.

For MEK kinase assays, cell pellets (~4.5 x 106 cells) were lysed in 0.15 ml of 50 mM Tris-HCl (pH 8.0), 250 mM NaCl, 2 mM EDTA, and 1% NP-40, with protease inhibitors (see above). Cell lysates (300 µg) were incubated (2 h at 4°C with rocking) in 0.3 ml of immunoprecipitation buffer (50 mM HEPES [pH 7.4], 250 mM NaCl, 2 mM MgCl2, 1 mM EDTA, 1% glycerol, 1% Triton X-100, and 1% NP-40 with protease inhibitors; see above) and 3 µg of anti-MEK1. Immune complexes were collected (2 h at 4°C with rocking) with 50 µl of washed protein A-agarose (Invitrogen). The collected immunoprecipitates were washed, and the kinase activity was determined as described for Raf-1 except that the reaction buffer contained 1 mM DTT, 6 µg of kinase-dead GST-ERK (K52R), 20 µM ATP, 20 µCi of [{delta}-32P]ATP (3,000 Ci/mmol) and protease inhibitors (see above). Nitrocellulose membranes were exposed to film to detect 32P-labeled GST-ERK fusion protein and then immunoblotted with anti-MEK1. Controls demonstrated that the immunoprecipitation depleted more than 80% of the total ERK and that the kinase assay was in the linear range of between 150 and 600 µg of cell lysate.

ERK kinase assays were performed and analyzed as described previously (61). Controls demonstrated that the immunoprecipitation depleted more than 80% of the total ERK and that the kinase assay was in the linear range between 25 and 100 µg of cell lysate.

In vitro cyclin D1-cdk4 and cyclin E-cdk2 kinase assays. Cyclin D1-cdk4 kinase assays with GST-Rb protein (Santa Cruz) as a substrate were performed as described previously (77). For cyclin E-cdk2 kinase assays, frozen cells (2 x 106 to 3 x 106 cells) were extracted in 0.1 ml of freshly prepared lysis buffer (50 mM Tris-HCl [pH 8], 250 mM NaCl, 5 mM MgCl2, and 0.1% NP-40 with protease inhibitors; see above). Equal amounts of cell lysate (250 µg in 0.1 ml of lysis buffer) were incubated with 2 µg of anti-cyclin E for 2 h on ice and then with 50 µl of washed protein A-agarose (Invitrogen) for 2 h at 4°C with rocking. Collected immunoprecipitates were washed once with cold lysis buffer and then four times with cold kinase reaction buffer (20 mM HEPES [pH 8.0], 10 mM MgCl2, 0.1 mM DTT) with protease inhibitors (see above). The washed immunoprecipitates were suspended in 50 µl of kinase reaction buffer, containing 1 µg of histone H1 (UBI), 20 µM ATP, and 20 µCi of [{delta}-32P]ATP (3,000 Ci/mmol) and incubated at room temperature for 30 min. Kinase reactions were stopped with an equal volume of 2x SDS sample buffer; the samples were fractionated on reducing SDS-gels and transferred to nitrocellulose membranes. The amount of 32P-labeled histone H1 was visualized by exposure to film. Filters were then immunoblotted with anti-cdk2. Controls demonstrated that the immunoprecipitation reaction depleted ~90% of the total cyclin E and that the kinase assay was in the linear range between 100 and 500 µg of cell lysate.

Fluorescence microscopy. Quiescent h{alpha}5-3T3 (2.5 x 105) or MEFs (2 x 105) were seeded on coverslips in 35-mm dishes with 2 ml of 10% FBS-DMEM, fixed, and permeabilized (61). Actin was stained with fluorescein-phalloidin (1 to 1.5 U/ml; 30 min). For vinculin staining, the permeabilized cells were incubated sequentially with anti-vinculin (Sigma V4505; 1:100 dilution for 2 h) and TRITC (tetramethyl rhodamine isothiocyanate)-conjugated anti-mouse immunoglobulin G (Jackson Laboratories; 1:300 dilution for 1 h). To analyze S phase entry, the stimulation with 10% FBS was performed in the presence of 3 µg of bromodeoxyuridine (BrdU; Amersham)/ml; permeabilized cells were incubated with DNase (140 U/µl) and anti-BrdU (BioDesign M20105S; diluted 500-fold; 1 h) and then fluorescein isothiocyanate-conjugated anti-sheep immunoglobulin G (Jackson Laboratories; diluted 200-fold; 1 h). Cell nuclei were stained with DAPI (4',6'-diamidino-2-phenylindole). Images were obtained by epifluorescence microscopy under oil at 40x magnification, captured by using a Hamamatsu digital charge-coupled device camera, and analyzed with Openlab Imaging System software.


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RESULTS
 
Actin stress fibers are required for sustained ERK activity and mid-G1 phase induction of cyclin D1. Many studies have shown that Rho kinase and its effectors, MLCK and LIMK, are important for the formation of actin stress fibers (see introduction). Indeed, we found that inhibition of Rho kinase with either with Y27632 (34) or dominant-negative Rho kinase blocked stress fiber formation and integrin clustering as determined by phalloidin staining and punctate vinculin staining, respectively, in both h{alpha}5-3T3 cells (Fig. 1) and MEFs (not shown). Similar results were obtained when we treated cells with ML-7 (an inhibitor of MLCK; (65) or expressed dominant-negative LIMK, except that the effect of dominant-negative LIMK was not complete (Fig. 1). Cortical f-actin persisted under these conditions (Fig. 1), a finding consistent with studies showing that Rho effectors other than Rho kinase (e.g., mDia) can mediate actin polymerization (37, 73).



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  		FIG. 1. Inhibition of Rho kinase and its effectors blocks stress fiber formation. h{alpha}5-3T3 cells were transiently transfected with a dominant-negative (dom neg) Rho kinase expression vector or dominant-negative LIMK expression vector and then serum starved or else serum starved and then pretreated with 10 µM Y27632 or 10 µM ML-7. Control cells were either transiently transfected with empty vector or pretreated with DMSO. The cells were plated at subconfluence on fibronectin-coated dishes containing coverslips and stimulated with 10 ng of bFGF/ml. Coverslips were collected at 9 h, fixed, permeabilized, and analyzed for f-actin and vinculin by fluorescence microscopy. Bar, 5 µm.

Quiescent h{alpha}5-3T3 cells pretreated with Y27632 (Fig. 2A) or ML-7 (Fig. 2B) or transfected with dominant-negative Rho kinase (Fig. 2A) or dominant-negative LIMK (Fig. 2A) were replated at subconfluence on fibronectin-coated dishes and stimulated with bFGF to examine the consequence of stress fiber disruption on the autophosphorylation of FAK at Y397 (a biochemical measure of integrin clustering) and sustained ERK activation. Each of these treatments inhibited FAK autophosphorylation, albeit not completely (Fig. 2A and B; fibronectin). In fact, residual FAK autophosphorylation was expected since some integrin clustering can occur in the absence of stress fibers (17, 45). Sustained activation of ERK (viz, ERK activity between 3 and 9 h) was also blocked by each of these treatments (Fig. 2; fibronectin). Moreover, when we plated the treated cells on anti-{alpha}5ß1, we rescued full FAK autophosphorylation and sustained ERK activity (Fig. 2, anti-{alpha}5ß1). As expected, the mid-G1 phase expression of cyclin D1 was also rescued when ML-7-treated cells were plated on anti-{alpha}5ß1 (Fig. 2B, anti-{alpha}5ß1), since that is a consequence of sustained ERK activity (reviewed in reference 60). The comparable cyclin D1 rescue experiment could not be performed with Rho kinase- or LIMK-inhibited cells, since inhibition of those Rho effectors allows for Rac/Cdc42-dependent cyclin D1 expression (Fig. 6A to C and data not shown) (77). Note that the effect of anti-{alpha}5ß1 on ERK signal duration was not blocked by cycloheximide (indicating that it was not a consequence of secreted matrix proteins) and required bFGF (indicating that the preparation of anti-{alpha}5ß1 was not mitogenic in itself) (not shown). Although use of Y27632, ML-7, dominant-negative Rho kinase, and dominant-negative LIMK may have individual effects that go beyond stress fiber formation (see, for example, reference 20), the fact that we obtained identical results for each of these treatments strongly argues that it is their common effect on stress fibers that allows for clustering of {alpha}5ß1 and sustained ERK activity (refer to Fig. 8).



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  		FIG. 2. Enforced clustering of {alpha}5ß1 integrin rescues sustained ERK activity and mid-G1 phase cyclin D1 expression. (A) h{alpha}5-3T3 cells were either transiently transfected (with empty vector, dominant-negative Rho kinase, or dominant-negative LIMK and then serum starved) or serum starved prior to treatment with Y27632. The starved cells (lanes 0) were treated with trypsin, plated on dishes coated with fibronectin or anti-{alpha}5ß1, and stimulated with 10 ng of bFGF/ml. (B) Starved h{alpha}5-3T3 cells were treated with trypsin, held in suspension for 1 h (lanes S), and pretreated with DMSO or ML-7 before being plated on dishes coated with fibronectin or anti-{alpha}5ß1. Collected cells were lysed and analyzed by immunoblotting with antibodies to pY397 FAK, FAK, cyclin D1, and cdk4 (loading control). ERK activation was determined by gel-shift and by direct detection of dually phosphorylated ERKs (pERK).



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  		FIG. 6. Rac/Cdc42-dependent cyclin D1 expression leads to activation of the G1 phase cdks in the absence of stress fibers. (A) MEFs were serum starved for 48 h; pretreated with DMSO, 10 µM ML-7, or 10 µM Y27632; plated at subconfluence; and stimulated with 10% FBS. At the times shown, collected cells were lysed and equal amounts of total protein were analyzed by immunoblotting for the levels and activation of ERK by using anti-ERK and anti-pERK, respectively. Duplicate filters were analyzed by immunoblotting for the expression of cyclin D1, cdk4, cyclin E, cdk2, p27kip1, and p21cip1. (B) MEFs transiently transfected with empty vector or the dominant-negative (dom neg) Rho kinase expression vector, serum starved for 48 h, plated at subconfluence, and stimulated with 10% FBS. Collected cells were lysed and analyzed by immunoblotting with antibodies against ERK, pERK, cyclin D1, cdk4, and myc (the epitope tag for dom neg Rho kinase). Cell lysates were also incubated with anti-cdk4, and the collected immunoprecipitates were used to assess in vitro cdk4 activity by phosphorylation of GST-Rb. The amount of immunoprecipitated (IP) cdk4 was assessed by immunoblotting (IB) by using filters from the kinase assay. (C) MEFs were transiently transfected with empty vector, serum starved for 48 h, and preincubated with DMSO or U0126 in the absence and presence of Y27632. Alternatively, MEFs were transiently transfected with the PBD expression vector, serum starved for 48 h, and preincubated with DMSO or Y27632. Cell lysates were analyzed by immunoblotting with antibodies to ERK, pERK, cyclin D1, cdk4, and GST (epitope tag for the PBD). (D) serum-starved MEFs were pretreated with DMSO, 10 µM Y27632, or 10 µM ML-7. (E) MEFs were transfected with empty vector or dominant-negative Rho kinase prior to 36 h serum starvation. For both panels D and E, the cells were plated at subconfluence and stimulated with 10% FBS. Cell lysates were incubated with anti-cyclin E, and the collected immunoprecipitates (IP) were used to assess in vitro cyclin E-cdk2 kinase activity by phosphorylation of histone H1. The levels of cyclin E-associated cdk2 was determined by immunoblotting (IB) the kinase assay filters with anti-cdk2.



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  		FIG. 8. Regulation of cyclin D1 expression by Rho kinase in fibroblasts. Rho kinase-dependent stress fiber formation allows for {alpha}5ß1 integrin clustering. Clustered {alpha}5ß1 integrin cooperates with RTK signaling upstream of Ras to sustain Ras, Raf, MEK, and ERK activities. These effects lead to mid-G1 phase cyclin D1 expression and can explain why G1 phase cell cycle progression is stress fiber dependent. Rho kinase is also involved in suppressing the early G1 phase expression of cyclin D1 by Rac/Cdc42. Rac/Cdc42 signaling induces cyclin D1 in early G1 phase, and this effect is independent of stress fiber formation. Since stress fibers do not affect the downregulation p21cip1 or p27kip1 (not depicted), G1 phase progression is independent of stress fiber formation if Rac/Cdc42 induces cyclin D1.

The results in Fig. 2 raised the possibility that stress fibers were specifically acting upstream of {alpha}5ß1 integrin to stimulate clustering and signaling. However, we found that FAK autophosphorylation remained incomplete and ERK activity was not sustained when bFGF-treated h{alpha}5-3T3 cells were incubated with anti-{alpha}5ß1-conjugated beads in suspension culture (not shown). Thus, although our data indicate that actin stress fibers are required for full FAK autophosphorylation and sustained ERK activity, they appear to act both upstream and downstream to maintain {alpha}5ß1 integrin clustering throughout G1 phase (refer to Fig. 8). One group has reported that ERK can disrupt the actin cytoskeleton by posttranscriptionally downregulating the expression of Rho kinase (52, 53). However, we found that (i) both Rho kinase isoforms (ROK{alpha}/ROCK2 and ROKß/ROCK1) are constitutively expressed even though ERK is active throughout most of G1 phase and (ii) the levels of both Rho kinase isoforms were unaffected by ERK inhibition with U0126 (not shown). Thus, in our system there was no apparent connection between ERK activity and Rho kinase expression.

Sustained ERK activity requires sustained activation of the Ras-Raf-MEK cascade. Many studies have examined the mechanisms involved in the coordinate signaling between RTKs and integrins on short-term (typically 5 to 60 min) ERK activity (reviewed in reference 60). Some of these studies indicate that integrin signaling affects the RTK itself (46-48, 67), whereas others have placed the adhesion-dependent step at different loci along the Ras-Raf-MEK-ERK signaling cascade (42, 57). To date, no studies have directly examined how integrins and RTKs cooperate to sustain an ERK signal for the several hours needed to induce cyclin D1. We asked whether adhesion and/or stress fibers regulate the duration of ERK signaling by affecting the activity of ERK phosphatases and/or MEK activity.

To assess the role of mitogen-activated protein kinase phosphatases, duplicate sets of h{alpha}5-3T3 cells were plated on fibronectin-coated dishes and stimulated with bFGF to induce sustained ERK activity. Thirty minutes prior to each collection (from 1 to 9 h), one dish of cells was treated with the MEK inhibitor, U0126. As determined by immunoblotting with phospho-ERK antibodies, activated ERK was readily detected throughout G1 phase in the cells treated with vehicle (DMSO) but absent whenever the cells were treated with U0126 (Fig. 3A). Thus, with MEK inhibited, the ERK signal is rapidly dephosphorylated, indicating that ERK phosphatases are active throughout most of the G1 phase. We then forced sustained ERK activity by ectopic expression of activated MEK so that we could compare the effect of U0126 in adherent and nonadherent cells, as well as in adherent cells treated with ML-7. ERK was rapidly dephosphorylated (reflecting the activity of ERK phosphatases) after addition of U0126 in all three culture conditions (Fig. 3B). Together, these results show that ERK phosphatases are active and continually dephosphorylating ERK throughout G1 phase, independently of cell adhesion or stress fiber formation. Thus, ERK phosphatases are not playing a major role in the regulation of ERK activity by either cell adhesion or actin stress fibers.



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  		FIG. 3. Cell adhesion and stress fibers are required for sustained MEK activity in growth factor-treated cells. (A) Quiescent h{alpha}5-3T3 cells were plated at subconflence on fibronectin (FN)-coated dishes and stimulated with 10 ng of bFGF/ml. These cells were treated with DMSO or U0126 30 min before collection, at each of the indicated times. (B) h{alpha}5-3T3 cells were transiently transfected with constitutively activated MEK, serum starved, pretreated with DMSO or ML-7, plated on fibronectin or BSA-coated dishes (suspension culture) and stimulated with 10 ng of bFGF/ml. Cells were treated with U0126 or DMSO 5 min prior to the indicated collection time (3 or 9 h). In panels A and B, cell lysates were analyzed for activation of ERK by gel-shift and by direct detection of dually phosphorylated ERKs (pERK). (C) Quiescent h{alpha}5-3T3 cells were plated on dishes coated with fibronectin or poly-L-lysine (PLL) and treated with 10 ng of bFGF/ml. At the indicated times, cells were collected and lysed. ERK activation was determined by immunoblotting as described above. Cell lysates were also incubated with anti-ERK or anti-MEK, and the collected immunoprecipitates were used to assess in vitro ERK or MEK activities by phosphorylation of myelin basic protein (MBP) or kinase-dead ERK [GST-ERK (K52R)], respectively. The amounts of ERK or MEK in the immunoprecipitates (IP) were determined by immunoblotting (IB) with anti-ERK or anti-MEK, respectively. Both kinase-dead ERK and MBP remained unphosphorylated when the lysates were incubated with a control serum (not shown). (D) Quiescent h{alpha}5-3T3 cells were pretreated with DMSO or ML-7, plated on dishes coated with either fibronectin (FN) or anti-{alpha}5ß1, and stimulated with 10 ng of bFGF/ml. Cells were collected at the indicated times, lysed, and analyzed for the activation of ERK and MEK activity as described for panel C.

Quiescent h{alpha}5-3T3 cells were then plated on fibronectin or poly-L-lysine (which mediates cell adhesion independently of integrins) and stimulated with bFGF to compare the extent and duration of MEK and ERK activities. The results showed that MEK and ERK activities were sustained (up to 9 h) when growth factor-treated cells were plated on fibronectin but transient (~1 h) when the cells were plated on poly-L-lysine (Fig. 3C). Thus, the time course of MEK activity corresponds to the time course of ERK activity. We note, however, that MEK activity, though persistent, decays somewhat faster than ERK activity. In three separate experiments, MEK activity was ~25% of maximal by 6 h, whereas ERK activity was ~50% of maximal, as determined by phosphorimager analysis. Similarly, disruption of stress fibers with ML-7 blocked both sustained MEK and ERK activities (Fig. 3D, FN), and anti-{alpha}5ß1 restored sustained MEK and ERK activities in ML-7-treated cells (Fig. 3D, anti-{alpha}5ß1). Overall, our results indicate that stress fiber-dependent clustering of {alpha}5ß1 integrin regulates the duration of ERK activity by controlling the duration of MEK activity.

To map the site of adhesion and stress fiber action within the Ras-Raf-MEK-ERK cascade, h{alpha}5-3T3 cells stimulated with bFGF were cultured on poly-L-lysine or on fibronectin in the absence or presence of ML-7 (Fig. 4A). We found that Ras-GTP loading and Raf-1 activity were sustained (up to 9 h) in the control cells plated on fibronectin and transient (~1 h) in the cells treated with ML-7 or plated on poly-L-lysine. These effects closely matched those seen for activation of MEK and ERK (Fig. 4A but also see Fig. 3C and D). Thus, the synergistic signaling between RTKs, integrins, and stress fibers that leads to sustained ERK activity maps upstream of Ras (refer to Fig. 8).



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  		FIG. 4. Integrin signaling and stress fibers act upstream of Ras to sustain ERK activity. (A) Quiescent h{alpha}5-3T3 cells were pretreated with DMSO or ML-7, plated on dishes coated with either fibronectin or poly-L-lysine (PLL), and stimulated with 10 ng of bFGF/ml. Cells were collected at the indicated times, lysed, and analyzed for total and GTP-loaded Ras. Identically treated cells were analyzed for Raf-1 activity and the level of total Raf-1 (see Materials and Methods). Total cell lysates were also used to monitor the activities of MEK and ERK by immunoblotting for phosphorylation of MEK at S222 and dual phosphorylation of pERK, respectively. (B) h{alpha}5-3T3 cells were transiently transfected with empty vector or FRNK, serum starved for 24 h (0), and then held in suspension for 1 h (lanes S), prior to stimulation on fibronectin (FN)-coated dishes with 10 ng of bFGF/ml. Collected cells were lysed and analyzed by immunoblotting with antibodies to pY397 FAK, FAK, pERK, ERK, cyclin D1, and cdk4 (loading control).

Some studies have implicated FAK phosphorylation in the synergistic activation of ERKs by RTKs and integrins (see Discussion). However, we found that expression of FRNK (a dominant-negative FAK that effectively inhibits FAK autophosphorylation at Y397; Fig. 4B) did not block sustained ERK activity or mid-G1 phase cyclin D1 expression when quiescent h{alpha}5-3T3 cells were plated on fibronectin and stimulated with bFGF. Similarly, ectopic expression of activated FAK (CD2-FAK [13]) did not rescue sustained ERK activity or cyclin D1 expression in ML-7-treated cells (not shown).

Stress fibers are not required for G1 phase cell cycle progression if Rac/Cdc42 signaling is used to induce cyclin D1. Established MEFs were used to assess the effects of stress fiber formation on G1 phase cell cycle progression. These cells behave identically to h{alpha}5-3T3 cells with regard to ERK and Rac/Cdc42-dependent cyclin D1 expression; the effects of MLCK and Rho kinase inhibition are also the same (77; the present study [see below]). However, MEFs show more consistent mitogen and adhesion-dependent regulation of p21cip1 and p27kip1 (see Fig. 6 and reference 85).

Consistent with the several studies that have implicated cellular tension in G1 phase progression (see introduction) and our data in h{alpha}5-3T3 cells (Fig. 2), the disruption of stress fibers in MEFs with ML-7 blocked entry into S phase (Fig. 5). However, equivalent disruption of stress fibers (refer to Fig. 1) and sustained ERK activity (refer to Fig. 6) with either Y27632 or dominant-negative Rho kinase led to an acceleration of S phase entry by 3 to 4 h (Fig. 5; compare DMSO or vector to Rho kinase-inhibited cells). Both flow cytometry and immunoblotting for the induction of cyclin A (a marker of S phase entry) also demonstrated a similar shortening of G1 phase upon inhibition of Rho kinase (not shown). Although DNA synthesis in the absence of stress fibers is a common property of transformed cells, S phase entry of these Rho kinase-inhibited cells remained adhesion and mitogen-dependent (Table 1), indicating that the cells were not transformed.



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  		FIG. 5. Distinct effects of Rho kinase inhibition and MLCK inhibition on progression through G1 phase. MEFs were serum starved for 36 h and then pretreated with 10 µM Y27632, 10 µM ML-7, or DMSO. Alternatively, MEFs were transiently transfected with a dominant-negative (dom neg) Rho kinase expression vector or empty vector and then serum starved for 36 h. The cells were plated in dishes containing coverslips and stimulated with 10% FBS in the presence of BrdU. Coverslips were collected and fixed at the indicated times for an analysis of S phase entry by BrdU incorporation, counting ~150 cells per sample.


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  		TABLE 1. Inhibition of Rho kinase does not phenotypically transform MEFsa

To identify the molecular basis for these distinct proliferative responses, we compared the effects of MLCK and Rho kinase inhibition on activation of the G1 phase cyclin-cdks. Although sustained ERK activity was blocked in MEFs treated with ML-7 (Fig. 6A), Y27632 (Fig. 6A), or dominant-negative Rho kinase (Fig. 6B), cyclin D1 was not expressed in the ML-7-treated cells, whereas MEFs treated with Y27632 or expressing dominant-negative Rho kinase showed the early G1 phase induction of cyclin D1 and activation of cdk4 characteristic of Rac/Cdc42 signaling. Moreover, as with our previous studies in 3T3 cells (77), the expression of cyclin D1 seen in Rho kinase-inhibited cells was blocked by transfection of the p21-binding domain of PAK (PBD; a specific Rac/Cdc42 inhibitor) and unaffected by U0126 (Fig. 6C). Conversely, the expression of cyclin D1 seen in the control cells was unaffected by the PBD and blocked by U0126 (Fig. 6C). Thus, stress fibers are required to induce cyclin D1 and activate cdk4 in response to sustained ERK signaling but not in response to Rac/Cdc42 signaling (refer to Fig. 8).

We also compared the effect of Rho kinase inhibition and MLCK inhibition on cyclin E-cdk2. The levels of cyclin E, cdk2, and the cyclin E-cdk2 inhibitory proteins, p21cip1 and p27kip1, were not affected by ML-7 (Fig. 6A), Y27632 (Fig. 6A), or dominant-negative Rho kinase (not shown). Interestingly, the activation of cyclin E-cdk2 was blocked in cells treated with ML-7 (Fig. 6D) but accelerated ~3 h in cells treated with Y27632 (Fig. 6D) or dominant-negative Rho kinase (Fig. 6E). This outcome was expected because the absence of cyclin D1-cdk4 complexes in MLCK-inhibited cells would preclude p21cip1 and p27kip1 sequestration and thereby inhibit cyclin E-cdk2, whereas the premature formation of cyclinD-cdk4 complexes in Rho kinase-inhibited cells should result in a premature sequestration of p21cip1 and p27kip1 and early activation of cyclin E-cdk2 (15, 68). Overall, these data show that cyclin E-cdk2 activation is stress fiber independent as long as cyclin D1 is induced. Moreover, the shortening of G1 phase we observed in Rho kinase-inhibited cells (refer to Fig. 5) is consistent with the early activation of both cyclin D1-cdk4 and cyclin E-cdk2.

The results described above indicate that stress fiber formation is required for S phase entry when ERK is used to express cyclin D1 but not when Rac/Cdc42 is used to express cyclin D1. We therefore reasoned that all of G1 phase progression should be unaffected by ML-7 in cells that express cyclin D1 ectopically. Quiescent MEFs expressing tetracycline-repressible cyclin D1 were stimulated with serum in the absence and presence of tetracycline. As expected, ML-7 blocked the expression of endogenous (ERK-dependent) cyclin D1 but not the expression of ectopically expressed cyclin D1 that occurred upon removal of tetracycline (Fig. 7, inset). Importantly, ectopic expression of cyclin D1 expression rescued S phase entry in the ML-7-treated cells (Fig. 7). These data show that stress fiber formation is dispensable in cells expressing cyclin D1 and argue that ERK-dependent cyclin D1 induction is the only event in the G1 phase cyclin-cdk network that requires actin stress fibers, at least in fibroblasts.



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  		FIG. 7. Ectopic expression of cyclin D1 rescues S phase entry in ML-7-treated cells. MEFs stably expressing tetracycline (tet)-repressible cyclin D1 were grown to confluence in the presence of tetracycline, washed, and serum starved for 48 h in the presence or absence of tetracycline. In the continued presence or absence of tetracycline, the starved cells were trypsinized and pretreated with 10 µM ML-7 (lanes M) or DMSO (lanes D) as described for MEFs in Materials and Methods. The pretreated cells were plated in 100-mm dishes containing coverslips and stimulated with 10% FBS in the presence of BrdU. Coverslips were collected and fixed at 6 h ({square}) and 21 h ({blacksquare}) for an analysis of S phase entry by BrdU incorporation, counting ~150 cells per sample. The results show mean ± the standard deviation. The remaining cells were collected at 21 h, lysed, and analyzed by immunoblotting with antibodies to cyclin D1 and cdk4 (inset).


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DISCUSSION
 
G1 phase cell cycle progression in the presence of Rho kinase and stress fibers. We examined here the mechanism by which Rho kinase regulates sustained ERK activity and, consequently, the mid-G1 phase expression of cyclin D1. We found that inhibition of Rho kinase, or its effectors MLCK and LIMK, blocked stress fiber formation, {alpha}5ß1 clustering and sustained ERK activity (Fig. 8). Enforced clustering of {alpha}5ß1 restored sustained ERK activity in these treated cells, and the identical results obtained with each of four independent experimental approaches (Y27632, dominant-negative Rho kinase, dominant-negative LIMK, and ML-7) strongly argues that it is their common effect on stress fiber formation that is central to sustained ERK activity. Others have already proposed that stress fiber and focal adhesion formation generate the tension required to cluster integrins (16), and our results show a functional role for this effect in G1 phase cell cycle progression.

Stress fibers and {alpha}5ß1 integrin signaling allow for sustained ERK signaling by regulating the duration of Ras/Raf/MEK activity rather than the activity of ERK phosphatases. Thus, like the studies examining RTK-integrin activation of ERK in short-term assays (reviewed in reference 60), it appears that integrin-dependent activation of the upstream kinase cascade, rather than the inhibition of ERK phosphatases, sustains the ERK signal for the 5- to 6-h period needed to induce cyclin D1. However, our data also indicate that different mechanisms underlie the short- and long-term effects of integrin signaling on RTK-dependent ERK activity. In particular, Lin et al. (42) and Renshaw et al. (57) report that cell adhesion synergizes with RTKs downstream of Ras-GTP loading, but we find that the adhesion/stress fiber-dependent locus for sustained ERK activity lies upstream of Ras-GTP loading. Other studies have also linked FAK phosphorylation to ERK activation (5, 29, 56, 84) and cyclin D1 expression (83, 84) in mitogen-treated cells, but our studies indicate that FAK phosphorylation is not required for sustained ERK activity or cyclin D1 expression when h{alpha}5-3T3 cells are plated on fibronectin and stimulated with bFGF. Barberis et al. (8) reached a similar conclusion in studies with MEFs that were unable to recruit FAK to ß1 integrins. Thus, the mechanistic relationship between FAK and ERK activation appear to be context dependent. Studies in progress are now attempting to characterize the relative roles of Shc, Src-family kinases, and ligand-independent RTK activation in regulating the duration of Ras and ERK activation. These mechanisms are thought to underlie integrin-dependent activation of ERK upstream of Ras (47, 48, 74, 75).

In contrast to the effects on cyclin D1 induction and in agreement with Sahai et al. (63, 64), we found that the expression of cdk4, cdk2, cyclin E, p21cip1, and p27kip1 were unaffected by inhibition of Rho kinase or MLCK. Others have reported that Rho is required for S phase entry and the downregulation of p21cip1 and p27kip1 (see introduction), and we have confirmed these effects of Rho (not shown) in MEFs. Thus, Rho kinase-independent effectors of Rho regulate the activation of cyclin E-cdk2, whereas Rho kinase regulates the activation of cyclin D-cdk4 via ERK-dependent cyclin D1 expression.

G1 phase cell cycle progression in the absence of Rho kinase-dependent stress fibers. In addition to describing the role of Rho kinase in regulating sustained ERK activity and mid-G1 phase expression of cyclin D1, our results also show that stress fibers will not be required for G1 phase progression if Rac/Cdc42 is used to induce cyclin D1 (Fig. 8). This bimodal view of stress fiber function is compatible with the studies that strongly implicate cellular tension in G1 phase progression but also with the facts that stress fibers are not required for G1 phase progression of many cell types in culture nor generally detected in fibroblasts in vivo. We suggest that the ultimate proliferative response to stress fiber formation and disruption will depend on the relative roles of ERK versus Rac/Cdc42 in inducing cyclin D1.

In our studies, the Rac/Cdc42 and ERK pathways to cyclin D1 are independent as well as parallel: inhibition of ERK signaling does not affect Rac/Cdc42 signaling to cyclin D1 and vice versa. However, others have reported that PAK, a Rac/Cdc42 effector, phosphorylates Raf-1 and MEK, and that these phosphorylations contribute to optimal Raf-1 and MEK activity (18, 23, 39, 41, 82). Similarly, Rac has been reported to stimulate the nuclear translocation of ERK (28). In our experiments, inhibition of endogenous Rac/Cdc42 and PAK did not have a pronounced effect on G1 phase ERK activity or ERK-stimulated cyclin D1 expression. The basis for these different results is not completely clear, but it may reflect our analysis of endogenous protein effects, our use of an optimal mitogenic stimulus, or our focus on relatively long incubation times.

Rac/Cdc42 signaling results in an early G1 phase induction of cyclin D1. Consequently, cdk4 is activated prematurely, as is cyclin E-cdk2. Although the premature activation of cdk4 and cdk2 does result in an accelerated entry into S phase, we could not detect an increased rate of cell proliferation in Rho kinase-inhibited MEFs (not shown). A proliferation assay may not be sensitive enough to distinguish a 3 to 4 h shortening in the first G1 phase. Alternatively, S phase may be lengthened to compensate for the shortened G1 phase, as has been observed when G1 phase is shortened by ectopic expression of cyclins D1 or E (50, 54, 58). We also note two studies (34, 63) reporting that the rate of S phase entry was unchanged (rather than increased) by inhibition of Rho kinase; Rac/Cdc42 signaling to cyclin D1 may be somewhat less efficient in those cells. Nevertheless, the composite data indicate that if cyclin D1 is induced by Rac/Cdc42, then activation of cdk4, activation of cyclin E-cdk2, progression through G1 phase, and cell proliferation can occur in the absence of stress fibers and the consequent imposition of cellular tension.

Our results emphasize that a subtle change in the regulation of cyclin D1 (ERK versus Rac/Cdc42-dependent induction) can have a profound effect on the growth properties of cells by allowing them to proliferate with or without cellular tension. Stress fiber formation is strongly dependent upon the ECM, and the ECM is often remodeled physiologically, e.g., during wound repair and development. Thus, the ability to cycle and proliferate in the presence or absence of stress fibers by the differential use of apparently redundant signal transduction pathways to cyclin D1 may actually allow cells to control their proliferation in response to these changing extracellular environments.


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ACKNOWLEDGMENTS
 
We thank David Boettiger, Margaret Chou, Kozo Kaibuchi, Kensaku Mizuno, Martin Schwartz, and Michael Weber for generously providing plasmids. Fumio Matsumura kindly provided antibody to phospho-MLC. Yemisi Oluwatosin generated the tet-regulated cyclin D1 MEFs.

K.R. was supported by postdoctoral fellowship DAMD17-98-1-8209 from the Department of the Army. These studies were supported by NIH grants CA72639 and GM51878 to R.K.A.


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FOOTNOTES
 
* Corresponding author. Mailing address: Department of Pharmacology, University of Pennsylvania School of Medicine, 3620 Hamilton Walk, 167 Johnson Pavilion, Philadelphia, PA 19104-6084. Phone: (215) 898-7157. Fax: (215) 573-5656. E-mail: rka{at}pharm.med.upenn.edu. Back


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Molecular and Cellular Biology, June 2003, p. 4283-4294, Vol. 23, No. 12
0270-7306/03/$08.00+0     DOI: 10.1128/MCB.23.12.4283-4294.2003
Copyright © 2003, American Society for Microbiology. All Rights Reserved.



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