Threonine 37 of the mouse RhoE cDNA in pCMV5-Flag was mutagenized to asparagine by use of a QuikChange site-directed mutagenesis kit (Stratagene). The sequence changes were verified with an ABI Prism dye terminator cycle sequencing kit (Applied Biosystems). Wild-type and mutant (N37RhoE) mouse RhoE proteins were amplified from pCMV5-Flag-RhoE vectors by PCRs with the following oligonucleotides: forward, 5′-CCCAAGCTTTAAAGGAGAGAGCCAGC-3′, and reverse, 5′-GCTCTAGATCACATCACAGTACAGCTC-3′. The amplified sequence was then digested with HindIII and XbaI, and the resulting fragment was subcloned into the pTRE-HA vector (Clontech). pTRE-HA-RhoE constructs were cotransfected with pBabepuro into S2-6 cells. After puromycin (2 μg/ml) selection, individual puromycin-resistant clones were selected, amplified, and screened by Western blotting for inducible expression (with or without tetracycline) of hemagglutinin (HA)-RhoE by use of an anti-HA antibody (Covance Research). In addition, a dish of puromycin-resistant clones were pooled to create a population with inducible expression of HA-RhoE. Individual clones and the cell population are referred to collectively as RhoE-3T3 cells. Stable cell lines for Flag-RhoAV14, Flag-cyclin D1, cyclin D1-cdk4R24C, and E1A cells were generated by transfecting the indicated plasmids together with a hygromycin-resistant vector into RhoE-3T3 cells and following the same procedure as that outlined above.

NIH 3T3 and RhoE-3T3 cells were transfected by using Lipofectamine 2000 (Invitrogen) according to the manufacturer's instructions with the following expression plasmids: cyclin D1 promoter-luciferase reporter (pGL3-1748CD1Luc; a gift of S. Cook), H-RasV12 (pcDNA3-H-RasV12; a gift of J. Downward), RafCAAX (pEF-mycRafCAAX; a gift of R. Marais), Flag-RhoE (pCMV5-Flag-RhoE [35]), Flag-RhoAV14 (pCMV5-Flag-RhoAV14; a gift of G. O. Cory), Flag-cyclin D1 (pCDNA3-Flag-cyclin D1; a gift of Carme Gallego), cyclin D1-cdk4R24C (pBabeDK; a gift of Parmjit Jat), cyclin E (pCMX-Myc-cyclin E; a gift of Roger Watson), E7 (pBabeE7; a gift of Parmjit Jat), E1A (pLPCE1A; a gift of Parmjit Jat), pEGFPC1 (Clontech), and an E2F-responsive luciferase reporter (pE2F-Luc; Clontech).

Cells were harvested in a buffer containing 50 mM Tris-HCl (pH 7.4), 150 mM NaCl, 1 mM EDTA, and 1% (vol/vol) Triton X-100 plus protease and phosphatase inhibitors (2 μg of aprotinin/ml, 10 μg of leupeptin/ml, 1 mM phenylmethylsulfonyl fluoride, 1 mM dithiothreitol, 1 mM NaF, and 0.2 mM Na₃VO₄). Protein content was measured by the Bradford procedure, with bovine serum albumin as a standard (5). Cell lysates or proteins from pull-down or immunoprecipitation experiments were electrophoresed in sodium dodecyl sulfate (SDS)-polyacrylamide gels. After electrophoresis, the proteins were transferred to Immobilon P strips for 2 h at 60 V. The sheets were preincubated in a mixture of TBS (20 mM Tris-HCl [pH 7.5], 150 mM NaCl), 0.05% Tween 20, and 5% defatted milk powder for 1 h at room temperature and then were incubated for 1 h at room temperature in a mixture of TBS, 0.05% Tween 20, 1% bovine serum albumin, and 0.5% defatted milk powder containing the following antibodies, as appropriate: rabbit anti-phospho-ERK (9101; Cell Signaling) (1:1,000), anti-phospho-Akt (9271; Cell Signaling) (1:1,000), anti-p107 (sc-318; Santa Cruz) (1:1,000), anti-cyclin A (sc-596; Santa Cruz) (1:1,000), anti-p27^kip1 (sc-528; Santa Cruz) (1:1,000), anti-p21^cip1 (sc-471; Santa Cruz) (1:1,000), anti-cdk4 (sc-260; Santa Cruz) (1:1,000), anti-RhoE (35), anti-phospho-FoxO3a (06-952; Upstate Biotechnology) (1:1,000), and anti-phospho(Y397)-FAK (44-624; Biosource International) (1:1,000), antibodies; goat anti-cdk2 antibody (sc-163-G; Santa Cruz) (1:1,000); and mouse monoclonal anti-HA (MMS-101R; Covance Research) (1:1,000), anti-Rac1 (05-383; Upstate Biotechnology) (1:1,000), anti-cyclin D1 (sc-8396; Santa Cruz) (1:1,000), anti-RhoA (sc-418; Santa Cruz) (1:100), and anti-Flag (M2; Sigma) (1:1,000) antibodies. After being washed in TBS-0.05% Tween 20, the sheets were incubated with a peroxidase-coupled secondary antibody (1/2,000 dilution; Amersham) for 1 h at room temperature. After incubation, the sheets were washed twice in TBS-0.05% Tween 20 and once in TBS. The peroxidase reaction was visualized by ECL (Amersham).

Cell cycle profiles were measured by flow cytometry using propidium iodide. Briefly, trypsinized cells were collected by centrifugation, washed in phosphate-buffered saline, and fixed for 30 min at 4°C in 70% ethanol. After being washed twice with phosphate-buffered saline, DNAs were stained with propidium iodide (50 μg/ml) in the presence of 50 μg of RNase A (Sigma)/ml. Stained cells were then processed with a Becton Dickinson FACScan instrument and analyzed with Cell Quest software. For flow cytometric analysis of transiently transfected cells, cells were transfected with green fluorescent protein alone or together with the indicated vectors, and at 48 h posttransfection, the cells were harvested and incubated for 45 min at 37°C with 20 μg of Hoechst 33342 (Sigma)/ml. Green fluorescent protein-positive cells were then analyzed for DNA content with a UV laser (325 nm) in a Becton Dickinson LSR1 instrument and were analyzed with Cell Quest software. For analyses of S-phase entry, serum-starved or serum-stimulated cells were incubated with 3 μg of BrdU/ml for 2 h and then fixed in 3.7% formaldehyde for 15 min. Fixed cells were incubated for 1 h at 37°C with an anti-BrdU antibody (Amersham) and then for 45 min at 37°C with tetramethyl rhodamine isocyanate (TRITC)-conjugated goat anti-mouse immunoglobulin G (IgG) (Jackson Immunoresearch) (1:200).

The capacities of Rho-GTP and Rac-GTP to bind to GST-rhotekin and GST-PBD (p21-activated kinase-binding domain), respectively, were used to analyze the amounts of active GTPases (31). Cells (5 × 10⁶ to 10 × 10⁶) were lysed with Rho/Rac extraction buffer (50 mM Tris-HCl [pH 7.5], 1 mM EDTA, 500 mM NaCl, 10 mM MgCl₂, 1% [vol/vol] Triton X-100, 0.5% sodium deoxycholate, 0.1% SDS, 10% [vol/vol] glycerol, 0.5% [vol/vol] 2-mercaptoethanol) plus the protease and phosphatase inhibitors described above. Cleared (10,000 × g) lysates were incubated for 45 min at 4°C with glutathione-Sepharose 4B beads coupled to GST-rhotekin or GST-PBD (Upstate Biotechnology) for Rho-GTP or Rac-GTP pull-down assays, respectively. The beads were washed four times in extraction buffer. Bound proteins were solubilized by the addition of 35 μl of Laemmli loading buffer and were separated in SDS-12.5% polyacrylamide gels. The amount of Rho or Rac in the bound fraction was detected by Western blotting.

Cells grown on coverslips were fixed in 3.7% (vol/vol) paraformaldehyde for 20 min and permeabilized in 0.2% (vol/vol) Triton X-100 for 5 min. Actin filaments were visualized by incubating the fixed cells for 45 min at 37°C with TRITC-phalloidin (Sigma) (1:500). For vinculin staining, cells were incubated for 1 h at 37°C with an antivinculin antibody (v-4505; Sigma) (1:200) followed by 1 h at 37°C with fluorescein-conjugated anti-mouse IgG (Jackson Immunoresearch) (1:200). For HA-RhoE staining, fixed cells were incubated for 1 h at 37°C with an anti-HA antibody (Covance) (1:400) followed by 1 h at 37°C with fluorescein-conjugated anti-mouse IgG (Jackson Immunoresearch) (1:200).

Total RNAs were isolated from RhoE-3T3 cells by use of an RNeasy midikit (Qiagen) according to the manufacturer's instructions. The RNAs (15 μg of each) were denatured, electrophoresed in agarose-formaldehyde gels, and transferred to nylon membranes. The membranes were prehybridized with ExpressHyb (Clontech) for 1 h at 68°C and hybridized with a ³²P-labeled cyclin D1 probe diluted in ExpressHyb solution for 2 h at 45°C. The results were quantified by scanning autoradiograms.

For luciferase reporter assays, cells were seeded the day before transfection at a density of 5 × 10⁴ cells per well in 24-well plates. NIH 3T3 cells were transfected with 0.4 to 0.7 μg of a cyclin D1 luciferase reporter construct (pGL3-1748D1Luc; a gift of S. Cook) together with 0.25 to 0.35 μg of the indicated expression plasmids. RhoE-3T3 cells were transfected with 0.6 μg of an E2F-responsive luciferase reporter plasmid (pE2F-Luc; Clontech). The cells were harvested 48 h later in passive lysis buffer (Promega), and the luciferase activity was measured with a MicroBeta counter. Levels of reporter gene induction in transiently transfected cells were calculated by normalizing the luciferase levels to total Renilla luciferase levels. Results are presented as means ± standard deviations (SD) of data from three independent experiments, each of which was done in duplicate.

Serum-starved cells grown in 10-cm-diameter dishes were stimulated with serum (10% fetal calf serum [FCS]) for 6 h in complete DMEM followed by 1 h in methionine- and cysteine-free DMEM. The cells were then labeled for 30 min with 100 μCi of [³⁵S]methionine-cysteine (Redivue Promix [³⁵S] in vitro cell labeling mix; Amersham)/ml and chased for the indicated times in complete DMEM. The cells were harvested on ice in lysis buffer (50 mM Tris-HCl [pH 7.4], 150 mM NaCl, 1 mM EDTA, and 1% [vol/vol] Triton X-100, plus the protease and phosphatase inhibitors described above) and were clarified by centrifugation (10,000 × g). The supernatants (equalized for protein concentration) were precleared by an overnight incubation at 4°C with 30 μl of protein G-Sepharose (Amersham), and radioactivity levels in the supernatants were also measured to check that the levels of incorporated ³⁵S correlated with the protein concentration. Immunoprecipitations were carried out by incubating the cleared supernatants for 2 h at 4°C with 2 μg of anti-cyclin D1 antibody (sc-8396; Santa Cruz) and then for 1 h at 4°C with 15 μl of protein G-Sepharose (Amersham). Immunoprecipitates were then washed four times in lysis buffer and resuspended in Laemmli loading buffer. Immunoprecipitated proteins were run in SDS-12% polyacrylamide gels, and the incorporated radioactivity was detected with a phosphorimager (Bio-Rad) and quantitated with Quantity One software.

To date, the only biological effects described for Rnd proteins are on the actin cytoskeleton. Transient RhoE and Rnd1 expression in different cell types induces stress fiber disassembly, a loss of focal adhesions, and cell rounding (13, 23). We therefore tested whether the induction of HA-RhoE expression in RhoE-3T3 fibroblasts would have the same effects. RhoE induced a dramatic loss of actin stress fibers within 6 h (Fig. ​(Fig.1B),1B), together with a significant reduction in the number of focal adhesions, as indicated by the diffuse vinculin staining (Fig. ​(Fig.1C).1C). However, a time course analysis of RhoE expression revealed that at longer time points, the cells recovered and reverted to normal levels of stress fibers and a spread phenotype indistinguishable from that of control cells, despite the high levels of HA-RhoE expression (Fig. ​(Fig.1D).1D). Thus, the strong effects elicited by acute RhoE expression on the actin cytoskeleton are transient and disappear within hours in cells that express RhoE for longer periods of time.

RhoE has been shown to prevent ROCK signaling, and thus stress fiber assembly, by binding directly to the N-terminal region of ROCK I (35). In addition, it has been reported that RhoE binding to p190RhoGAP stimulates its GAP activity towards RhoA, resulting in lower Rho-GTP levels and thus to a loss of stress fibers (49). We therefore investigated whether either of these mechanisms was involved in RhoE-induced cell cycle arrest. ROCK inhibition is not likely to be responsible for RhoE-mediated cell cycle arrest because cells treated with the ROCK inhibitor Y-27632 were not impaired in their proliferative ability (Fig. ​(Fig.3A),3A), consistent with previously published observations (36, 39). In contrast, Rho inhibition is associated with cell cycle inhibition (48, 50). We therefore tested whether RhoE could modulate Rho-GTP levels in RhoE-3T3 cells. Pull-down experiments using GST-rhotekin indicated that HA-RhoE expression did not affect RhoA-GTP levels, either after HA-RhoE induction in growing cells (Fig. ​(Fig.3B)3B) or in starved RhoE-expressing cells stimulated with serum to induce maximal RhoA activity (Fig. ​(Fig.3C).3C). We also investigated whether constitutively active RhoA could rescue RhoE-induced cell cycle inhibition, using stably transfected RhoAV14-expressing RhoE-3T3 cells. Upon RhoE induction, cell growth was still inhibited in these cells, despite the expression of active RhoA (Fig. ​(Fig.3D).3D). Altogether, these data suggest that neither ROCK nor RhoA inhibition, which is involved in RhoE-mediated stress fiber disassembly, accounts for RhoE-mediated cell cycle arrest.

In order to understand how RhoE inhibits G₁ phase progression, we analyzed the expression levels of different cell cycle regulators in RhoE-3T3 cells. Cyclin D1 peaked at mid-to-late G₁ (~9 h) in serum-stimulated control cells (Fig. ​(Fig.4A).4A). In sharp contrast, cyclin D1 was almost undetectable in RhoE-expressing cells. A similar pattern was observed for p21^cip1, which was almost absent from RhoE-expressing cells. p27^kip1 was efficiently downregulated as control cells progressed through G₁ phase. However, although RhoE-expressing cells showed some downregulation of p27^kip1, they did not induce complete p27^kip1 degradation, probably as a consequence of their failure to progress into S phase. cdk4 and cdk2 levels were similar in control and RhoE-3T3 cells, and as expected, HA-RhoE was only present in tetracycline-deprived cells (Fig. ​(Fig.4A).4A). Since G₁ progression ultimately leads to E2F activation via pocket protein phosphorylation, we analyzed the expression of two characterized E2F-dependent targets, p107 and cyclin A (20), as a readout of E2F activity and hence pocket protein inactivation. Whereas in control cells both proteins were induced in a cell cycle-dependent manner, their expression was almost undetectable in cells expressing RhoE at all time points analyzed (Fig. ​(Fig.4B).4B). In concordance with these results, RhoE expression inhibited E2F transcriptional activity in a reporter gene assay using a luciferase-based reporter for E2F (Fig. ​(Fig.4C).4C). Altogether, these results indicate that RhoE prevented the release of E2F from pocket proteins, an event induced by cdk phosphorylation. The lack of cyclin D1 protein in RhoE-expressing cells could therefore contribute to RhoE-induced G₁ arrest.

We next focused on which signaling pathway promoting cyclin D1 expression might be affected by the expression of HA-RhoE. We first analyzed the activation levels of the ERK and phosphatidylinositide-3-kinase (PI3K)/Akt pathways in control and RhoE-3T3 cells, as they have been clearly implicated in cyclin D1 expression (37, 46). ERKs were transiently activated shortly after serum stimulation of control cells, and a very similar pattern of activation was observed for RhoE-expressing cells (Fig. ​(Fig.5A).5A). Mid-G₁ expression of cyclin D1 has been shown to rely on sustained ERK activation, which is dependent on integrin clustering (38) and is regulated by ROCK (36). RhoE also did not affect the levels of phosphorylated focal adhesion kinase (FAK) (Fig. ​(Fig.5A),5A), which has been shown to be involved in integrin-dependent cyclin D1 expression (51). PI3K pathway activation was monitored by analyzing the phosphorylation levels of two downstream effectors of this pathway that have been implicated in the regulation of cyclin D1 expression, specifically Akt and the forkhead family member FoxO3a (10, 41). Both Akt and FoxO3a were transiently phosphorylated in response to serum stimulation in both control and RhoE-expressing cells (Fig. ​(Fig.5A).5A). Finally, we investigated the ability of RhoE-3T3 cells to induce Rac activation, since Rac has been shown to promote cyclin D1 expression (12, 16, 19). Rac was rapidly activated in both serum-stimulated control and RhoE-expressing cells, reaching a peak at 5 min and then declining back to basal levels (Fig. ​(Fig.5B).5B). Thus, neither ERK, FAK, PI3K/Akt, nor Rac activation was impaired by RhoE expression, ruling out the possibility that RhoE blocked cyclin D1 induction due to an inhibition of any of these pathways.

Since the absence of cyclin D1 probably prevents cyclin-dependent kinase activity and thus pocket protein hyperphosphorylation, we analyzed whether enforced activation of the cdk-pRb axis could rescue cell cycle progression in RhoE-expressing cells. Interestingly, cyclin D1 expression alone was insufficient to rescue S-phase entry in RhoE-expressing cells, as shown in cell growth assays using cell lines stably expressing ectopic cyclin D1 (Fig. ​(Fig.7A)7A) or by FACS analysis of transiently transfected cells (Fig. ​(Fig.7B).7B). Very similar results were obtained with cells expressing a cyclin D1 construct fused to a p16-insensitive cdk4 point mutant (cdk4R24C) (30): both cell growth assays using cells stably expressing ectopic cyclin D1-cdk4R24C (Fig. ​(Fig.7A)7A) and FACS analysis (Fig. ​(Fig.7C)7C) showed that RhoE expression was still able to induce cell cycle arrest. However, the expression of human papillomavirus E7 or adenovirus E1A, which inhibits the function of the Rb family of pocket proteins, was able to rescue cell cycle progression in RhoE-3T3 cells, as indicated by growth curves of cells stably expressing E7 or E1A (Fig. 7A and C) and by FACS analysis of transiently transfected cells (Fig. 7B and C). Furthermore, the ectopic expression of cyclin E also rescued S-phase entry in RhoE-expressing cells (Fig. ​(Fig.7B),7B), although to a lesser extent than E7 and E1A. These results indicate that RhoE requires functional pocket proteins to induce cell cycle arrest.

Since RhoE inhibits cell cycle progression in fibroblasts, we investigated whether endogenous RhoE levels changed upon stimulation with signals that affect cell proliferation. No significant changes in endogenous RhoE levels were observed when quiescent NIH 3T3 cells were stimulated with serum to induce cell cycle reentry (Fig. ​(Fig.8A).8A). We next treated NIH 3T3 cells with a number of DNA-damaging agents in order to analyze RhoE expression in response to checkpoint induction. Interestingly, the treatment of cells with cisplatin induced an increase in endogenous RhoE levels that correlated with a decrease in cyclin D1 (Fig. ​(Fig.8B),8B), suggesting that changes in RhoE levels may be relevant for cisplatin-mediated DNA damage-induced cell cycle arrest.