Article

Ras^V12 and mutant p53 cooperatively induce cell motility

To explore mechanistic aspects of multistep carcinogenesis with a focus on cancer cell progression to malignancy, activated H-Ras^V12 and mutant p53^175H, a pair of strongly cooperating oncogenes⁷ with characteristic point mutations frequently associated with cancers, were introduced into polyclonal populations of early-passage young adult mouse colon cells^{38, 39}. Four cell populations were derived in parallel via infection with pairs of recombinant retroviruses (shown in parentheses): control (Bleo/Neo), mp53 (p53^175H/Neo), Ras (Bleo/Ras^V12) and mp53/Ras (p53^175H/Ras^V12) (see Methods for details). Confirming previous studies performed in other cell types, Ras^V12 and p53^175H cooperated to transform the murine colon cells. Only cells carrying both Ras^V12 and p53^175H were able to form colonies in soft agar in the absence of anchorage to substratum and induced invasive tumors when injected subcutaneously into nude mice (data not shown).

Enhanced cell motility and invasiveness arose as a consequence of cooperation between Ras^V12 and p53^175H, as determined by analysis of the above cell populations in three distinct assay systems (Fig. 1). In the so-called 'wounding assay', swaths of cells were physically removed from a confluent monolayer and the ability of cells to fill the available space was quantified after 16 h using phase-contrast microscopy. Whereas control cells did not migrate, mp53/Ras cells had filled 95% of the wound by this time. Ras or mp53 cells, however, had covered only approximately 30% of the wound area (Fig. 1a,c). To eliminate any contribution of cell division to this process, the experiments were carried out in the presence of mitomycin C. Time-lapse video microscopy to monitor cell motility confirmed that cell division was blocked during the 'wound closure' process (data not shown). Transwell migration assays using Boyden chambers yielded comparable results (Fig. 1b,d). In addition, when placed into three-dimensional matrigel cultures, only mp53/Ras cells were able to generate lattice structures. In contrast, control, Ras and mp53 cells were unable to form such lattices (Fig. 1e). Thus, Ras^V12 and p53^175H cooperate to induce cell motility in two- and three-dimensional cultures with both transwell migration and lattice formation in matrigel, phenotypes that strongly correlate with cancer cell invasiveness^{40, 41}.

Figure 1: Activated Ras and mutant p53 cooperatively induce cell motility in mouse colon cells.

(a) Phase-contrast micrographs depicting the migratory capacity of the retrovirally infected control (Ctrl), Ras, mp53 and mp53/Ras cells in the wound-closure assay 16 h after wounding. (b) Representative phase-contrast micrographs of cell populations after migration through matrigel-coated transwell membrane. (c) Quantification of cell motility in wound-closure assay from three independent experiments, as described in Methods. Error bars indicate s.d. (d) Quantification of cell invasiveness in transwell assay. Experiments were performed at least three times; a representative experiment is shown. Error bars indicate s.d. (e) Cells indicated were plated into three-dimensional matrigel culture and were kept in serum-free medium for 7 d. Cells were visualized by phase-contrast microscopy. (f) Activated H-Ras^V12, total H-Ras and mutant p53 protein expression. Mutant p53 was precipitated with Pab240, which selectively recognizes mutant p53, and visualized by immunoblotting with p53 antibody (FL-393). H-Ras^V12 was immunoprecipitated with monoclonal antibody Y13-259 and visualized by western blotting using a Ras^V12-specific antibody (top gel). Total H-Ras protein was visualized by western blotting with H-Ras antibody (C-20). In Ras and mp53/Ras lanes, total H-Ras bands are unresolved doublets of endogenous WT H-Ras and exogenous H-Ras^V12 proteins. -tubulin served as a loading control (bottom gel). (g) Total p53 protein abundance, phosphorylation status on Ser15 (S15) and acetylation status on Lys382 (K382) of p53 were analyzed by western blotting. -tubulin served as a loading control.

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In our experiments, Ras^V12 proteins were expressed at levels similar to endogenous Ras proteins (Fig. 1f, bottom) and thus are not highly overexpressed. Moreover, both exogenous Ras^V12 and p53^175H are expressed at comparable levels in Ras and mp53/Ras cells and in mp53 and mp53/Ras cells, respectively (Fig. 1f, top), indicating that the two mutant proteins do not affect each other's expression. In addition, Ras^V12 does not detectably alter either endogenous p53 expression or the level of p53 Ser15 phosphorylation or Lys382 acetylation in YAMC cells. Thus, regulation of covalent p53 modifications by Ras was not detected in our model system. Conversely, p53 phosphorylation and acetylation are suppressed in presence of p53^175H, indicating that expression of p53^175H has a dominant-negative effect on endogenous p53 (Fig. 1g).

Synergistic activation of RhoA by Ras^V12 and p53^175H

One class of proteins that might mediate altered motility of mp53/Ras cells in the assays described above are the Rho family GTPases, Rho, Rac and Cdc42. Analysis of GTP loading of the three Rho family members in the four cell populations revealed that the abundance of GTP-bound, active RhoA was greatly increased in mp53/Ras cells, whereas the overall RhoA protein abundance remained unaltered (Fig. 2a). In contrast, Rac1-GTP loading was decreased in cells expressing activated Ras (Fig. 2b), and levels of GTP-bound Cdc42 remained constant across the four cell lines (Fig. 2c). GTP loading of RhoA, Rac1 and Cdc42 in control, Ras, mp53 and mp53/Ras cell populations was measured by pull-down assays from cell lysates incubated with target protein domains that specifically interact with Cdc42, Rac and Rho (PAK for Rac1 and Cdc42, rhotekin for RhoA). These target domains selectively bind, and thus allow capture of, the GTP-bound, active form of the relevant GTPases^{42, 43}. We conclude that activated Ras^V12 and p53^175H cooperate to specifically activate RhoA in our colon cell model. This is consistent with the idea that high Rho activity and low Rac activity in combination facilitate cell motility²⁸.

Figure 2: Cooperative activation of RhoA by Ras and mutant p53.

(a–c) The endogenous abundances of the GTP-bound forms of RhoA (a), Rac1 (b) and Cdc42 (c) were measured by Rho-family protein activity pull-down assays. Whole-cell lysates were also immunoblotted for total RhoA, Rac1 and Cdc42 as loading controls.

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The idea that activated Ras and p53 inhibition collaborate to increase motility via RhoA activation is supported by three independent lines of experimental evidence. First, a chimeric membrane-penetrating Rho inhibitor, Tat-C3 (ref. 44), decreased RhoA-GTP loading and inhibited cell migration of mp53/Ras cells (Fig. 3a,b). This was associated with a tail-retraction defect, typical of Rho inhibition, without a detectable effect on the formation of protrusions at the leading edge of the cell (data not shown). Second, we partially inhibited RhoA activity by expressing RhoN19, a dominant-negative RhoA mutant. RhoN19 counteracts endogenous Rho activation by titrating guanine nucleotide–exchange factors (GEFs)⁴⁵, consistent with inhibition of endogenous Rho activation. We found that the motility of mp53/Ras cells decreased with increasing levels of RhoN19 expression and a concomitant decrease in RhoA-GTP loading (Fig. 3c,d). Under these conditions, the relative RhoA-GTP abundance in RhoN19-expressing mp53/Ras cells was consistently higher than in Ras cells, and the increase of RhoA-GTP abundance in mp53/Ras cells, compared with Ras cells, was directly proportional to the observed increase in cell motility. Finally, RhoN19 expression in mp53/Ras cells also suppressed lattice formation in three-dimensional matrigel culture, whereas the proliferation capacity of the RhoN19-expressing cells in standard cultures was similar to controls, suggesting that Rho activity is also required for motility in matrigel (Fig. 3e,f).

Figure 3: Oncogene-dependent RhoA activation contributes to increased cell motility in mp53/Ras cells.

(a) The motility of mp53/Ras cells pretreated with 10 g ml^-1 Tat-C3 for 24 h and untreated control cells in wound-closure assays. Error bars indicate s.d. from three independent experiments. (b) RhoA-GTP loading was measured by Rho activity pull-down (middle blot). RhoA protein abundances were determined by western blotting (bottom blot). The uptake of Myc-tagged Tat-C3 in cell lysates was detected by immunoblotting with anti-Myc (9E10; upper blot). (c) mp53/Ras cells infected with retroviruses expressing RhoN19, concentrated RhoN19 (pBpRhoN19-C) or vector pBp alone. RhoA activity and the expression of Myc-tagged RhoN19 were analyzed by Rho activity pull-down assay and western blotting with anti-Myc (9E10), respectively. (d) Cell motility of Ras and mp53/Ras cells in c was determined by wound-closure assay. Error bars indicate s.d. from three independent experiments. The differences in motility of RhoN19 or RhoN19C cells from vector pBp or Ras cells were statistically significant, as determined by Student's t-test (^***, P < 0.001). (e) Phase-contrast micrographs of mp53/Ras cells infected with either RhoN19-C or vector pBp alone in three-dimensional matrigel culture after 1 week incubation. (f) Growth curve of drug-selected mp53/Ras cells infected with RhoN19-C or control vector. Cells were maintained in standard conditions at 39 °C.

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To test whether the RhoA activation observed in mp53/Ras cells might be sufficient to mediate the enhanced cell motility, we expressed a constitutively active, GTP-binding RhoA mutant with a single amino acid substitution (RhoA^V14) in cells expressing Ras or mp53 alone. Such stably transfected Ras/RhoA^V14 or mp53/RhoA^V14 cells had migration speeds similar to those of Ras or mp53 cells transfected with an empty vector in our cell-motility assays (data not shown), indicating that RhoA activation is necessary but not sufficient for induction of cell motility by cooperation of Ras^V12 and p53^175H.

Positive and negative regulation of RhoA by Ras^V12

To explore the mechanistic basis for the synergistic activation of RhoA by Ras^V12 and p53^175H, we tested a variety of parameters with the potential to affect RhoA regulation. As activated RhoA localizes to the cytoplasmic face of the cell membrane⁴⁶, we examined the cellular distribution of RhoA in the four cell populations.

In both Ras and mp53/Ras cells, considerable fractions of RhoA protein had translocated to the membrane, whereas in mp53 and control cells RhoA remained localized in the cytoplasm (Fig. 4a). In contrast to Ras cells, the membrane-associated RhoA protein in mp53/Ras cells was found predominantly in the GTP-bound and thus activated state (Fig. 4b), suggesting the involvement of multiple factors in the regulation of RhoA activity in Ras cells (Fig. 2a and Fig. 4b). Rho GDP dissociation inhibitor (RhoGDI), a protein that binds and retains RhoA in the cytoplasm, was expressed at similar levels in each of the four cell populations and is thus unlikely to have a role in the Ras-dependent redistribution of RhoA. As expected, RhoGDI was found only in the cytosolic fraction. Conversely, 1 integrin partitioned exclusively to the membrane fraction, so that both RhoGDI and 1 integrin served as quality controls for cell fractionation (Fig. 4a,b). Together, these results suggest that, in the presence of functional p53, activated Ras promotes RhoA membrane localization without stimulating GTP binding.

Figure 4: Ras positively and negatively regulates RhoA activation by increasing RhoA membrane localization and stimulating p190 RhoGAP activity, respectively.

(a) Subcellular distribution of RhoA protein in the indicated cell populations. Membrane and cytosolic fractions were isolated and immunoblotted using RhoA antibody. RhoGDI and integrin 1 are markers for cytosolic and membrane fractions, respectively. (b) RhoA GTP loading in membrane and cytosolic fractions from Ras and mp53/Ras cells. RhoA-GTP loading was determined by Rho activity pull-down assay in both fractions. The abundances of RhoA protein were monitored by immunoblotting with RhoA antibody. (c) p190 RhoGAP phosphorylation on tyrosine and its association with p120 RasGAP in the indicated cell populations. Endogenous p190 RhoGAP was immunoprecipitated with an antibody to p190 RhoGAP. Phosphotyrosine content and association of p190 RhoGAP with p120 RasGAP in the immunoprecipitates were detected by immunoblotting with anti-phosphotyrosine and anti–p120 RasGAP antibodies, respectively. Immunoprecipitated p190 RhoGAP was visualized with anti–p190 RhoGAP, serving as input control (upper blot). The relative expression levels of p190 RhoGAP and p120 RasGAP proteins in the indicated cell populations were monitored by western blotting using equivalent amounts of cell lysates (lower blot). (d) Activity of endogenous p190 RhoGAP in immunoprecipitates from the cell populations indicated. Immunoprecipitates were incubated with RhoA-GTP from HeLa cell lysates at room temperature, as indicated. RhoA-GTP remaining unhydrolyzed was collected by GST-RBD pull-down and detected using anti-RhoA. To indicate the RhoA-GTP input, an equivalent amount of HeLa cell extract was incubated for 10 min at room temperature in the absence of p190 RhoGAP immunocomplexes before RhoA pull-down (top). Total HeLa RhoA in each sample was monitored by western blotting after incubation at room temperature (middle). The relative abundances of p190 RhoGAP in immunoprecipitates were visualized by immunoblotting with anti–p190 RhoGAP (bottom).

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GTPase-activating proteins (GAPs) constitute a group of important regulators that inhibit Rho family proteins by stimulating their rate of GTP hydrolysis. A Rho-specific GAP, p190 RhoGAP, has been shown to mediate inactivation of Rho in a variety of cell-signaling contexts. This inactivation correlates with p190 RhoGAP Tyr1105 phosphorylation, followed by increased association with p120 RasGAP and enhanced p190 RhoGAP activity^{29, 47, 48, 49, 50}.

To our surprise, we detected considerable activation of p190 RhoGAP specifically in Ras^V12-expressing cells, which may explain the finding that Ras cells contain membrane-bound but mostly inactive RhoA. We observed that tyrosine phosphorylation of p190 RhoGAP and its association with p120 RasGAP were strongly increased in Ras cells but not in mp53/Ras, mp53 or control cells. However, overall abundances of both p190 RhoGAP and p120 RasGAP remained unaltered, suggesting that changes in abundances of these proteins do not account for the inactivity of RhoA in Ras cells (Fig. 4c). In addition, direct measurement of RhoGTPase activity in p190 RhoGAP immunoprecipitates revealed detectable RhoA-GTP hydrolysis in association with p190 RhoGAP from Ras cells, but not in any of the other cell populations (Fig. 4d). The detected RhoGAP activity was specific to p190 RhoGAP from Ras cells, as control experiments using nonspecific antibodies did not promote hydrolysis of Rho-bound GTP (data not shown).

On the basis of the data described above, we conclude that Ras induces both membrane localization of RhoA and the Rho-inhibitory p190 RhoGAP activity. This implies that Ras controls RhoA activity through simultaneous engagement of two opposing signals with RhoA activity, reflecting the ratio of these two signaling inputs (see also Discussion below).

We also observed that p190 RhoGAP activation by Ras^V12 is abrogated in the presence of p53^175H, whereas RhoA membrane localization remains unaffected. This suggests that both Ras and p53 function are required for p190 RhoGAP activation, whereas RhoA membrane localization is dependent on Ras activity alone. We therefore predict that p53, presumably acting through p190 RhoGAP, prevents RhoA activation and thus cell mobilization by regulating the ratio of the two opposing Ras signals converging on RhoA activity control.

Ras-dependent inhibition of RhoA mediated by p190 RhoGAP

The model proposed above predicts that p190 RhoGAP has a causal role in the control of RhoA activity by Ras^V12 and p53^175H. We therefore targeted p190 RhoGAP messenger RNA for knockdown through stable expression of two independent retrovirus-based short hairpin RNA (shRNA) constructs in Ras cells. In both of the resulting polyclonal cell populations, p190 RhoGAP protein expression was considerably reduced, whereas GTP loading of RhoA was dramatically increased, as compared with vector controls (Fig. 5a). In contrast, p190 RhoGAP knockdown in mp53 or mp53/Ras cells had no effect on the respective RhoA activities in these cell populations (Fig. 5b), presumably because p190 RhoGAP is relatively inactive in both mp53 and mp53/Ras cells, as compared with Ras cells (Fig. 4d). This demonstrates that the activation of RhoA in mp53/Ras cells relies, at least in part, on the inhibition of p190 RhoGAP activity after expression of p53^175H. As with expression of activated RhoA^V14 (see above), p190 RhoGAP knockdown is not sufficient to induce cell motility of Ras cells (data not shown).

Figure 5: Negative regulation of RhoA by activated Ras mediated by p190 RhoGAP.

(a) shRNA-mediated knockdown of p190 RhoGAP induces RhoA-GTP loading in Ras cells. Knockdown efficiency of Ras cells stably infected with retroviruses expressing p190 RhoGAP–specific shRNAs p190i1, p190i2 or vector (pSR, pSuperRetro) was assessed by western blotting with p190 RhoGAP antibodies; p120 RasGAP served as a loading control (top blot). RhoA-GTP loading was measured by Rho pull-down assay. RhoA-GTP loading in mp53/Ras cells is shown for comparison (bottom blot). (b) ShRNA-mediated knockdown of p190 RhoGAP does not alter RhoA activity in mp53 and mp53/Ras cells. ShRNAs p190i1 and p190i2 were used to knock down p190 RhoGAP in mp53 and mp53/Ras cells as described in a. The knockdown efficiency was assessed by western blotting with p190 RhoGAP antibodies; p120 RasGAP served as a loading control (top blot). RhoA-GTP loading was measured by Rho pull-down assay (bottom blot). (c) Expression of shRNA-resistant p190 RhoGAP reverts p190 shRNA-mediated RhoA-GTP loading, whereas tyrosine substitution mutants Y1105F and Y1087F Y1105F do not. Ras-p190i1 and Ras-p190i2 cells were infected with retroviruses expressing shRNA-resistant HA-tagged WT p190 RhoGAP, p190 RhoGAP(Y1105F), p190 RhoGAP(Y1087F Y1105F) or empty vector (pBabeHygro). The resulting cell populations were subjected to RhoA activity pull-down as described in Methods. Expression of exogenous p190 RhoGAP proteins was verified by blotting with anti-HA.

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Reduced p190 RhoGAP activity in mp53/Ras cells relative to Ras cells probably involves a decrease in p190 RhoGAP tyrosine phosphorylation (see Fig. 4c). We directly tested this idea in a genetic rescue experiment in which wild-type (WT) p190 RhoGAP and its tyrosine phosphorylation–site mutants Y1105F or Y1105F Y1087F⁵⁰, lacking the p190 shRNA targeting sequences, were expressed in the p190 RhoGAP–knockdown Ras cells described above. Expression of WT p190 RhoGAP reduced the activity of endogenous RhoA to levels similar to those found in Ras cells in which expression of endogenous p190 RhoGAP had not been altered. This indicated that p190 RhoGAP mediates the Ras-induced RhoA-inhibitory signal and reconfirmed the specificity of the p190 RhoGAP shRNA constructs used. In contrast, both p190 RhoGAP mutant proteins, Y1105F and Y1105F Y1087F, did not suppress RhoA-GTP loading, indicating that Tyr1105 is essential for p190 RhoGAP activation (Fig. 5c). This strongly supports a causative role for tyrosine phosphorylation in Ras-mediated p190 RhoGAP activation.

In agreement with the idea that Ras controls RhoA activity through two opposing output signals, we determined that p190 RhoGAP regulation and RhoA membrane localization are independent events in Ras cells, as p190 RhoGAP knockdown does not alter the distribution of RhoA between membrane and cytosol induced by activated Ras (Supplementary Fig. 1 online). This is similar to the condition found in mp53/Ras cells, where RhoA is predominantly localized at the membrane and p190 RhoGAP activity is low.

Ras activation of p190 RhoGAP dependent on p53 function

The use of the p53 mutant protein p53^175H in our experiments does not allow a distinction to be made between dominant-negative and potential gain-of-function effects of p53^175H, which have previously been observed for some p53 mutations in the absence of WT p53 (refs. 51,52). To directly test whether the effects of p53^175H on Rho activity are a consequence of endogenous p53 inhibition, we reduced wild-type p53 protein abundance through stable expression of two independent retrovirus-based shRNA constructs targeting p53 mRNA in Ras cells. In both cases, this resulted in a large reduction in p190 RhoGAP activity, as indicated by decreased tyrosine phosphorylation and p120 RasGAP association. Moreover, RhoA GTP binding was strongly increased. As expected, expression of p53^175H in the same Ras cell population had effects similar to p53 knockdown (Fig. 6a). These results indicate that the p53^175H-mediated changes in cell signaling observed in our experiments are not due to gain-of-function effects, instead suggesting that p53^175H acts in a dominant-negative fashion. An shRNA construct directed against human p53 (ref. 53) served as a specificity control in the murine cells used here and had no effect. Moreover, expression of either human p53– or mouse p53–specific shRNA constructs did not affect the level of p190 RhoGAP expression, further confirming the specificity of the p190 RhoGAP knockdown described above.

Figure 6: Loss of p53 function induces reduction of p190 RhoGAP tyrosine phosphorylation and accumulation of RhoA-GTP as well as motility in cancer cells expressing activated Ras.

(a) Ras cells were infected with retroviruses expressing mutant p53 (p53^175H), mouse-specific p53 shRNAs mp53i1 or mp53i2, human-specific p53 shRNA hp53i or vector alone (pSR). Infected cells were drug-selected, pooled and assayed for p53 protein expression by western blotting (top blot). Tyrosine phosphorylation of p190 RhoGAP and p120 RasGAP binding to p190 RhoGAP were monitored by immunoprecipitation followed by western blotting (middle blot), and RhoA-GTP loading was assayed by Rho pull-down assay (bottom blot). (b) RhoA-GTP loading (middle blot) and p190 RhoGAP tyrosine phosphorylation and association with p120 RasGAP (bottom blot) were measured in human colon carcinoma cells HCT116 (p53^+/+), HCT116 (p53^-/-) and HCT116 (p53^+/+) transfected with p53^175H or vector pBabePuro (pBp). Levels of p53 protein expression (top blot) were monitored by immunoblotting with anti-p53 (FL-393). (c) Invasiveness of HCT116 (p53^+/+), HCT116 (p53^-/-) and HCT116 (p53^175H) cells was measured by transwell assay as described in Methods, in absence (Control) or presence of the Rho inhibitor TatC3 (10 g ml^-1). Experiments were carried out at least three times; a representative experiment is shown, and error bars represent s.d. TatC3 at 10 g ml^-1 is not lethal to the cells in this assay, as determined by propidium iodide (PI) and trypan blue exclusion (data not shown). (d) RhoA-GTP loading and p190 RhoGAP tyrosine phosphorylation (pTyr)/association with p120 RasGAP in human colon carcinoma cells SW480, SW620, HT29 and HCT116 (p53^+/+). (e) Invasiveness of SW480, SW620 and HT29 cells as measured by transwell assay in absence (Control) or presence of TatC3 (10 g ml^-1) as described in c.

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RhoA-GTP loading, p190 RhoGAP phosphorylation and cell motility are influenced by p53 status during malignant transformation of mouse colon cells. We therefore tested whether human colon cancer cells show similar effects. Human HCT116 cells contain a ras gene mutation and wild-type p53 (ref. 54). In these cells, both RhoA activity and cell motility are low (Fig. 6b,c), although p14^ARF is defective⁵⁵. In contrast, in HCT116 cells in which p53 has been inactivated via homologous recombination⁵⁶, or that exogenously express mutant p53^175H, we found that GTP binding of RhoA and cell motility are greatly induced, whereas p190 RhoGAP activity, as measured by tyrosine phosphorylation and p120 RasGAP binding, is greatly decreased (Fig. 6b,c). Notably, loss of p14^ARF is not sufficient for RhoA activation, even though loss of p14^ARF promotes many other aspects of tumorigenesis dependent on loss of p53 function. In addition, human colon cancer cells such as SW480, SW620 and HT29 that carry mutations in the genes for both ras or B-raf (HT29) and p53 contain high RhoA and low p190 RhoGAP activities (Fig. 6d). Moreover, nonlethal concentrations of the Rho inhibitor Tat-C3 effectively inhibited the motility of each of the human colon carcinoma cells in which p53 function is defective (Fig. 6c,e) and RhoA activity is high (Fig. 6d). Notably, these data show that p53-dependent restraint of Ras-induced cell motility through activation of p190 RhoGAP and consequent suppression of RhoA activity is a mechanism conserved between murine and human cancer cells. A causal role for Rho activity in human cancer cell motility is also confirmed^{33, 34}.

Termination of Ras signal propagation by p53

Our data demonstrate a previously unrecognized mechanism of tumor suppression in which p53 prevents Rho activation mediated by activated Ras and thus suppresses cancer cell motility, a feature associated with disease progression to malignancy. This involves a signaling circuit in which Ras^V12 controls RhoA activity by simultaneously engaging both stimulatory and inhibitory signals (Fig. 7). More specifically, Ras^V12 promotes RhoA membrane translocation necessary for activation by GTP loading, but concurrently stimulates hydrolysis of RhoA-bound GTP via phosphorylation and activation of the Rho GTPase–activating protein p190 RhoGAP. This presumably increases turnover of RhoA-bound GTP, causing overall RhoA activity to remain low. Thus, while having no apparent net effect on RhoA activity, Ras^V12 converts RhoA to a state poised for activation by sensing changes in the ratio of the two opposing signals. Notably, our data also show that p53 can specifically affect this ratio, as functional p53 is essential for Ras^V12 to mount the Rho-inhibitory signal mediated by p190 RhoGAP phosphorylation. Conversely, p53 does not affect Ras^V12-dependent membrane localization. This suggests that activation and repression of RhoA by Ras^V12 are regulated by independent mechanisms. Consistent with this idea, inhibition of p190 RhoGAP through shRNA-dependent knockdown in Ras cells does not affect Ras-induced RhoA membrane localization and is sufficient to induce RhoA-GTP loading. Upon loss of p53 function, RhoA-GTP loading and cell motility thus greatly increase in the presence of activated Ras, because both p190 RhoGAP tyrosine phosphorylation and activity are reduced. Src and/or Abl-family kinases^{48, 57} and low–molecular weight protein-tyrosine phosphatase (LMW-PTP)⁵⁸ have been implicated in the regulation of p190 RhoGAP tyrosine phosphorylation. The mechanisms by which activated Ras and p53 loss of function may influence p190 RhoGAP activity will be interesting to explore. Together, our data reveal that p53 can function as a tumor suppressor by blocking Ras-mediated RhoA activation in a manner that prevents signaling through a specific Ras effector pathway, thus altering Ras signaling specificity. Mechanisms that downmodulate signaling activity in cells are as important as the processes that initiate and transmit signals. Here we present an example of a signaling mechanism that can prevent activation of inappropriate effectors through a circuit that integrates multiple inputs.

Figure 7: Tumor suppressor p53 inhibits activation of RhoA and cell motility by Ras^V12 via 'AND' logic circuit.

(a,b) Schematic representation of the distinct functional states (a) and diagram (b) of the signaling circuit that suppresses RhoA activation and cell motility by activated Ras in a p53-dependent manner, demonstrating a previously unrecognized role of p53 in tumor suppression. In the presence of functional p53, Ras^V12 simultaneously stimulates both positive and negative signals controlling RhoA activity, by promoting RhoA membrane localization and increasing the activity of the Rho-inhibitory protein p190 RhoGAP via phosphorylation. With both GDP-GTP exchange rates and GTP hydrolysis presumed high, rapid cycling between the GTP- and GDP-bound forms of RhoA prevents accumulation of active RhoA-GTP. Activated Ras induces RhoA activation only upon loss of p53 function, as loss of p53 function disables Ras-induced p190 RhoGAP activation but not RhoA membrane localization. Thus, p53 functions to restrict RhoA activation by activated Ras. (c) Table summarizing experimentally verified settings predicted by the RhoA signaling circuit, reminiscent of a multiplicative 'AND' logic controller. Left column indicates cell populations analyzed (Ctrl, control). Membrane localization, RhoA membrane localization status; p190 GAP-P, phosphorylation status of p190 RhoGAP; p190 GAP activity, GTPase-activating protein activity of p190 RhoGAP; RhoA-GTP, activation status of RhoA; plus or minus, qualitative bias of localization or activation state; ND, not determined.

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Signal ratios specify cell behavior

The advantages offered by signaling circuitry like that described above may not be intuitive at first glance. The key feature of the cellular signaling circuit by which activated Ras and p53 regulate RhoA activity is its capacity to regulate RhoA activation and cell motility by modulating the ratio of stimulatory and inhibitory RhoA input signals, while preventing any response to either Ras or p53-mediated signaling alone. Moreover, the circuit stimulates RhoA activation strongly in response to a specific configuration of the two input variables—that is, when Ras activity is high and p53 activity is low and thus RhoA is localized at the cell membrane and p190 RhoGAP activity is low. Such a circuit design resembles a Boolean 'AND' logic controller capable of generating digital signal output in response to specific signal input combinations. Moreover, circuits of this type tend to increase the robustness of signaling-network function, as response generation requires integration of multiple inputs. Notably, such a scenario provides a mechanistic rationale for why conversion of normal cells to a malignant phenotype requires multiple mutations.

Current and previous evidence indicates that cooperating oncogenic mutations can affect multiple cancer cell traits, such as cell-cycle progression, survival and motility. It is thus possible that several features of cancer cells can emerge simultaneously owing to cooperation of a few genetic lesions involving mechanisms such as those described above. This is consistent with the observation that a gene expression signature specific to metastatic cancer cells can already be detected in a subset of precancerous lesions with high probability of progression to malignancy^{59, 60}. Understanding of the underlying principles of cancer gene cooperativity may thus yield new approaches to specifically manipulate malignant cancer cell behavior.

Antibodies, compounds and plasmids.

The following antibodies were used: monoclonal antibody to p53 (Pab240, Santa Cruz), rabbit polyclonal antibodies to p53 (FL-393, Santa Cruz) and p53-pSer15 and p53-Ac382 (Cell Signaling), rat monoclonal Y13-259 (anti–v-H-Ras, Oncogene Research), monoclonal anti–Pan-Ras^V12 (Oncogene Research), rabbit polyclonal anti–H-Ras (C20, Santa Cruz), monoclonal anti-RhoA (26C4, Santa Cruz), monoclonal anti-Rac1 (BD Transduction), rabbit polyclonal anti-Cdc42 (P1, Santa Cruz), monoclonal anti–c-Myc (9E10, see Acknowledgments), monoclonal anti-phosphotyrosine (4G10, Upstate), rabbit polyclonal anti-RhoGDI (A20, Santa Cruz), monoclonal anti–p190 RhoGAP (BD Transduction), monoclonal anti-p120 RasGAP (Santa Cruz), monoclonal anti-HA11 (Covance), rabbit anti–-tubulin (H-235, Santa Cruz). Mitomycin C was obtained from Sigma.

Plasmids pGEX3X-GST-C21, containing the Rho-binding domain from Rho effector Rhotekin⁴³, and pGEX2TK-GST-PAKCD, containing the Rac- and Cdc42-binding region from human PAK1B⁴³, were gifts (see Acknowledgments). A DNA fragment encoding Myc-tagged human RhoN19 (ref. 27) (see Acknowledgments) was subcloned into the BamHI and XhoI sites of the retroviral vector pBabePuro2 (ref. 61). hemagglutinin (HA)-tagged p190 RhoGAP expression constructs WT HAp190A, HAp190A(Y1105F) and HAp190A(Y1087F Y1105F) (see Acknowledgments) lack the p190 RhoGAP shRNA targeting sequences located in the 3'-untranslated region of the mRNA described below and were cloned via EcoRI and SalI sites into pBabeHygro. pGEX-KG Tat-C3 was a gift (see Acknowledgments), and Tat-C3 was prepared as described⁴⁴. Two mouse p190 RhoGAP shRNA targeting sequences mapping to the 3' untranslated region, p190i1 (5'-GCTGTGAGCCCTCGCCTTG-3') and p190i2 (5'-GAGGACATTTCCCAGTTTG-3') and two mouse-specific p53 shRNA targeting sequences, p53i1 (5'-GACTCCAGTGGGAACCTTC-3') and p53i2 (5'-GAAGTCACAGCACATGACG-3'), and a human-specific p53 shRNA targeting sequence⁵³, hp53i, were cloned into the retrovirus vector pSUPER.retro.puro⁶².

Cell culture, retroviral infection and DNA transfection.

Early-passage polyclonal populations of young adult mouse colon (YAMC) cells (see Acknowledgments) from the Immorto mouse (also called H-2K^b/tsA58 transgenic mouse) expressing temperature-sensitive simian virus 40 large T (tsA58) under the control of a interferon–inducible promoter³⁹ were used as recipients for oncogene cooperation experiments. The cells were maintained at the permissive temperature (33 °C) for large T in the presence of interferon to support conditional immortalization. This permits expansion of the cells in tissue culture. In contrast, exposure of YAMC cells to the nonpermissive temperature for large T (39 °C) in the absence of interferon leads to growth arrest followed by cell death^{38, 39}, indicating the absence of immortalizing mutations in the cell population. Four polyclonal cell populations, control (Bleo/Neo), mp53 (p53^175H/Neo), Ras (Bleo/Ras^V12) and mp53/Ras (p53^175H/Ras^V12) were derived by retroviral infection of low-passage YAMC cells via cocultivation with mitomycin C–treated producer cells secreting pairs of retroviruses⁶¹ carrying the bleomycin resistance gene, alone or together with a complementary DNA for p53^175H, and the neomycin resistance gene, alone or together with a cDNA encoding H-Ras^V12. The infected cells were maintained in selective medium containing zeocin and geneticin (500 g ml^-1 each). After 2 weeks, drug-resistant cells were pooled. YAMC cells and all their derivatives—control, Ras, mp53 and mp53/Ras cells—were expanded in a humidified incubator with 5% CO₂ at 33 °C. All experiments testing the roles of Ras^V12 and p53^175H were carried out at the nonpermissive temperature (39 °C) for large T function and in the absence of interferon. Cells were switched to these conditions at least 1 d before the starting time of each experiment. The cells were cultured on Collagen IV–coated dishes (Sigma; 1 g cm^-2 for 2 h at room temperature) in RPMI 1640 medium (Invitrogen) containing 10% (v/v) FBS (Hyclone), 1 insulin, transferrin and selenium mix (ITS-A, Invitrogen), 2.5 g ml^-1 gentamycin (Invitrogen), and 5 U ml^-1 interferon (R&D Systems). Cells were infected with recombinant retroviruses produced as described⁶¹. For lipid-based DNA transfection, YAMC cells were seeded at 33 °C to achieve 80% confluence at the day of transfection and were treated with Lipofectamine Plus (Invitrogen) according to manufacturer's instructions.

Human cancer cell lines SW480, SW620 and HT29 were obtained from American Type Culture Collection. Human colorectal cancer cell line HCT116, and p53-deficient HCT116 cells⁵⁶ were a gift (see Acknowledgments). All of the above human colon cancer cells and the Phoenix ecotropic retroviral producer cells (see Acknowledgments) were maintained at 37 °C in DMEM (Invitrogen) with 10% (v/v) FBS, 100 g ml^-1 kanamycin (Sigma) and 2 g ml^-1 gentamycin (Invitrogen).

In vitro wound-closure assay.

For the wound-closure assay⁶³, cells were seeded in 24-well plates coated with collagen I and incubated at 39 °C overnight to generate confluent culture. Cells were then serum-starved for 8 h and treated with Mitomycin C 10 g ml^-1 for 2 h before wounding. The cell layers were scraped with a plastic pipette tip and washed three times with serum-free medium. The remaining cell culture was incubated 16 h to allow cells to migrate into the cleared space. To quantify cell migration, phase-contrast images of identical locations in each wound were taken at 0 h and 16 h after wounding. The rate of cell migration was then calculated as the average percentage of wound closure from at least three independent experiments. Error bars indicate s.d. of the mean.

Transwell migration and invasion assay.

Transwell migration and invasion assays were performed using 12-well transwell Boyden chambers (8 m PET membrane, Becton Dickinson) as described⁴⁰. The filters were coated with 0.4 mg ml^-1 matrigel (basement membrane matrix, BD Biosciences) in PBS at room temperature for 3 h. Mouse colon cells (10⁵) or human colon carcinoma cells (2 10⁵) were added to the top well resuspended in serum-free medium containing 0.1% (w/v) BSA, and the lower well was filled with medium containing 10% (v/v) FBS and 0.1% BSA. Where applicable, TAT-C3 (10 g ml^-1) was added to the top well together with the cells. After a 20-h incubation, cells remaining on the upper side of the filter were removed with cotton swabs, and the cells that had migrated to the bottom surface of the filter were fixed in methanol and stained with crystal violet. Transwell migration was evenly distributed over the filters and was quantified by counting at least three randomly chosen separate fields per filter under a bright-field microscope.

Three-dimensional matrigel culture.

Matrigel culture⁴¹ was performed using 12-well cell-culture plates. The matrigel (BD Biosciences) was thawed at 4 °C over night and then mixed with cells in ice-cold serum-free medium at a 1:3 dilution with a final cell density of 1.5 10⁵ ml^-1. The gel was placed over an agarose layer containing 0.6% (w/v) agarose, phenol red–free RPMI 1640 (Cellgro) at pH 7.4, 1 ITS-A and 2.5 g ml^-1 gentamycin. The plates were incubated at 39 °C for 1 week and inspected by phase-contrast microscopy.

Immunoprecipitation and immunoblotting.

Cell pellets were lysed for 20 min at 4 °C with rotation in RIPA buffer (50 mM Tris-HCl (pH 7.4), 150 mM NaCl, 1% (v/v) Nonidet P-40, 1 mM sodium orthovanadate, 10 mM NaF, 5 mM EDTA, 0.1% (w/v) SDS, 0.5% (v/v) deoxycholic acid, protease inhibitor cocktail tablet). Lysates were clarified by centrifugation at 13,000g for 10 min at 4 °C and subsequently quantified using Bradford (Bio-Rad) protein assay. Cell lysate (500–1,000 g) was then incubated with prebound antibody–protein G–Sepharose beads (Sigma-Aldrich) at 4 °C for 1–3 h. Immune complexes were then washed five times in RIPA lysis buffer, denatured in 2 Laemmli buffer (100 mM Tris-HCl (pH 6.8), 4% (w/v) SDS, 20% (v/v) glycerol, 0.1% (w/v) bromophenol blue, 200 mM DTT) at 95 °C for 5 min and resolved on SDS-polyacrylamide gels. For western blot analysis, 50–100 g of protein lysate was separated by SDS-PAGE and transferred to PVDF membrane (Millipore). Immunoblots were blocked in 5% (w/v) nonfat dry milk in PBS-T for 1 h at room temperature, probed first with specific primary antibodies and then with secondary antibodies conjugated with horseradish peroxidase. The immunoreactive bands were visualized using enhanced chemiluminescence kit (Amersham).

Rho activity pull-down assay.

For expression and purification of glutathione S-transferase (GST)-Rho–binding domain (RBD) polypeptides (Rhotekin for RhoA; Pak for Rac1 and Cdc42)^{42, 43}, bacteria expressing GST-RBDs (50 l of glycerol stock) were inoculated into 50 ml of LB with ampicillin and grown overnight. The culture was diluted 1:10 and then grown at 37 °C until an A₆₀₀ of 0.8 was reached. Protein expression was then induced with 0.3 mM IPTG for 2 h at 30 °C. The bacteria were pelleted and resuspended in 10 ml cold lysis buffer (20% (w/v) sucrose, 10% (v/v) glycerol, 50 mM Tris-HCl (pH 8.0), 2 mM MgCl₂, 2 mM DTT, protease inhibitor cocktail tablet). The bacterial suspension was sonicated on ice with a microtip at setting no. 3 eight times, for 15 s each. The lysate was clarified by centrifugation at 13,000g for 20 min at 4 °C and then mixed with 1 ml 50% (w/v) glutathione-Sepharose beads to incubate for 60 min at 4 °C. The beads were washed three times with lysis buffer and resuspended to 50% slurry with GST-fish buffer (50 mM Tris-HCl (pH 7.4), 150 mM NaCl, 10 mM MgCl₂, 10 mM NaF, 1 mM sodium orthovanadate, 10% (v/v) glycerol, 1% (v/v) Nonidet P-40, protease inhibitor cocktail tablet) and frozen at -80 °C in small aliquots.

To capture Rho-GTP, cells were lysed in cold GST-fish buffer and the cell lysate was clarified by centrifugation at 13,000g for 5–10 min at 4 °C. Equal amounts of cell lysates were incubated with glutathione Sepharose–conjugated GST-fusion protein for 1 h at 4 °C. The proteins bound to the Sepharose-conjugated GST-RBD were washed, eluted with sample buffer, separated on SDS-polyacrylamide gels and analyzed by immunoblotting with RhoA, Rac1 or Cdc42 antibodies.

Subcellular fractionation.

Cells were grown at 39 °C to confluence and serum-starved for 24 h. Adherent cells were then scraped into ice-cold hypotonic lysis buffer (10 mM Tris-HCl (pH 7.4), 1.5 mM MgCl₂, 5 mM KCl, 1 mM DTT, 0.2 mM sodium orthovanadate, protease inhibitor cocktail tablet) and cell lysates were homogenized with 30 strokes of a Dounce homogenizer. Homogenates were cleared of nuclei and unbroken cells by spinning at 700g for 5 min at 4 °C. The supernatants were then spun at 40,000g at 4 °C for 45 min to pellet membrane fractions. The cytosol-containing supernatant was removed and the crude membrane pellet was dissolved in detergent-containing lysis buffer. Membrane and cytosol fractions were measured for total protein and equal amounts were analyzed by SDS-PAGE and western blotting.

RhoGAP assay.

Endogenous RhoGAP activity was measured essentially as described⁶⁴. Protein G–bound p190 RhoGAP immunocomplexes precipitated from control, Ras, mp53 and mp53/Ras cell lysates (1.7 mg protein each) were incubated for 5 or 10 min at room temperature with HeLa cell extract (2 mg protein in 180 l GST-fish buffer) containing RhoA-GTP. After incubation, protein G beads and HeLa cell extract were separated by low-speed centrifugation and a 50-g aliquot of each sample was removed to monitor total RhoA expression. GTP-bound RhoA was then collected by GST-RBD pull-down from 1 mg cell extract per sample, as described above. The protein G–bound p190 RhoGAP immunocomplexes were washed in lysis buffer and the relative abundance of immunoprecipitated p190 RhoGAP in each of the samples was determined by western blotting.