Transformation by the Rho-specific Guanine Nucleotide Exchange Factor Dbs Requires ROCK I-mediated Phosphorylation of Myosin Light Chain*

Dbs was identified in a cDNA-based expression screen for sequences that can cause malignant growth when expressed in murine fibroblasts. In previous studies we have shown that Dbs is a Rho-specific guanine nucleotide exchange factor that can activate RhoA and/or Cdc42 in a cell-specific manner. In this current study we have used a combination of genetic and pharmacological approaches to examine the relative contributions of RhoA·PRK and RhoA·ROCK signaling to Dbs transformation. Our analysis indicates that ROCK is activated in Dbs-transformed cells and that Dbs transformation is dependent upon ROCK I activity. In contrast, there appears to be no requirement for PRK activation in Dbs transformation. Dbs transformation is also associated with increased phosphorylation of myosin light chain and stress fiber formation, both of which occur in a ROCK-dependent manner. Suppression of myosin light chain expression by small interfering RNAs impairs Dbs focus formation, thus establishing a direct link between actinomyosin contraction and Rho-specific guanine nucleotide exchange factor transformation.

GEF family, historically the family has attracted the most attention for its role in transformation. Chromosomal rearrangements with breakpoints that fall within BCR and LARG occur in distinct subsets of human leukemias, and overexpression of Clg has been implicated in murine leukemias (6 -8). Additionally, a large subset of the RhoGEFs has been isolated in screens for proteins whose expression cause deregulated growth in NIH 3T3 mouse fibroblasts (3). Typically NIH 3T3 cells that are transiently transfected with oncogenic RhoGEFs fail to undergo contact inhibition when confluence is reached, forming large dense foci that are morphologically distinct. Cell lines that stably express these oncogenic RhoGEFs also exhibit transformation as measured by a variety of parameters including growth in low serum, anchorage independence, and tumorigenicity in nude mice.
Dbs is a RhoGEF family member that was isolated in a screen for cDNAs whose overexpression cause deregulated growth in NIH 3T3 mouse fibroblasts (9,10). Like most RhoGEF family members, Dbs contains a tandem DH/PH domain motif that is both necessary and sufficient for Dbs transforming activity (9). DH domains are the catalytic cores of RhoGEFs and contain virtually all of the residues that are required for substrate recognition, binding, and exchange. PH domains have multiple roles in the context of RhoGEF family members (11)(12)(13)(14). In Dbs the PH domain contributes several residues to the catalytic interface (15), is a ligand for phosphoinositides (16), and contains a docking site for activated Rac1 (17). Whereas the role of Rac1 in Dbs transformation remains unclear, mutations that block lipid binding are completely impaired in transformation (16).
Like many members of the RhoGEF family, Dbs has potent transforming activity in NIH 3T3 cells, and this activity has been mapped to the catalytic domain (9,10). Although Dbs can target RhoA and Cdc42 in vitro (10,12), substrate usage can vary in a cell type-specific manner. In NIH 3T3 cells Dbs preferentially activates RhoA (18), in 293T cells it activates Cdc42 (17), and in COS-7 cells it does not appear to activate either GTPase. 5 A recent determination of the crystal structures of Cdc42 in complex with RhoA and Cdc42 has allowed for the design of mutants with more restricted target specificities (18,19). An analysis of the biological activity of these mutants has determined that Cdc42 activation is dispensable for Dbs transformation (18). Consistent with this, we are able to detect elevated levels of activated RhoA, but not Cdc42, in stable NIH 3T3 cell lines that express activated Dbs, and competitive inhibitors of RhoA specifically block Dbs transformation. Collectively, these observations suggest that RhoA is a relevant target for Dbs transformation in this cell type.
A number of potential target kinases and structural proteins have been described that interact with RhoA and may mediate Dbs transformation. These include ROCK I (20), ROCK II (21), PRK I (22), PRK II (23), p140mDia (24), rhotekin (25), rhophilin (22), citron kinase (CRIK) (26), kinectin (27), p116RIP (28), and the myosin binding subunit of myosin phosphatase (29). However, the evidence linking particular RhoA effector proteins to the transformation phenotype is still somewhat limited. RhoA mutants that are selectively impaired in their ability to interact with specific effectors have implicated ROCK I but not PRK I in RhoA transformation (30). In support of this, a pharmacological inhibitor of ROCK I and a genetic inhibitor of its substrate ezrin, both, block transformation by the RhoGEF family members Net and Dbl (31). However, the RhoA(L-63,V-39) mutant interacts efficiently with ROCK I and is competent in transformation but fails to stimulate actin polymerization (30). This suggests that ROCK I may mediate RhoA transformation independently of its ability to remodel the actin cytoskeleton. Consistent with this possibility, a recent study suggests that ROCK I can stimulate JNK-mediated transcriptional pathways independently of its ability to regulate actin (32). Transformation by RhoA has also been linked to transcriptional activation through ROCK I-independent pathways. For example, dominant inhibitors of MKK3 and MKK6, which are responsible for PRK I-mediated activation of p38␥, also block RhoA transformation (33). Although these studies suggest that ROCK I-and PRK I-mediated signaling may be necessary for RhoA transformation, it is unclear whether these kinases are directly activated in the context of RhoA transformation or are simply cooperating with additional RhoAmediated signaling pathways in transformation.
Because both ROCK I and PRK I have been implicated in RhoA transformation, we wished to determine whether RhoA-mediated activation of either kinase contributes to Dbs transformation. Our analysis of the activation status and cellular distribution of these kinases in Dbs-transformed cells suggests a role for ROCK I, but not PRK I, in Dbs transformation. Consistent with this, pharmacological and genetic inhibitors that target ROCK I impair Dbs transformation. ROCK-mediated hyperphosphorylation of myosin light chain (MLC) was also observed in Dbstransformed cells, and siRNA targeted against MLC also impaired Dbs transformation. These observations support a model whereby ROCKmediated stimulation of actino-myosin contraction contributes directly to Dbs transformation.
Cell Culture, Transfection, and Transformation Assays-NIH 3T3 cells were cultured at 37°C in 10% CO 2 in Dulbecco's modified Eagle's medium supplemented with 10% bovine calf serum (JRH Biosciences). Primary focus formation assays were performed as described previously (37). Briefly, NIH 3T3 cells were transfected by Lipofectamine reagent (Invitrogen) according to the manufacturer's recommended protocol. Cells were maintained in growth media for 14 days, and then cells were stained with 0.5% crystal violet to visualize foci. To examine a role for ROCK I in transformation, transfected cells were split into six 6-cm dishes, three of which were treated with 10 M ROCK inhibitor (Y-27632, Calbiochem). Cell media was changed, and fresh inhibitor was added every 24 h. To generate NIH 3T3 cell lines stably transfected by pCTV3H-dbs-HA6 or its cognate vector, the transfected cells were selected in growth media supplemented with hygromycin B (200 g/ml, Invitrogen) for 10 days. Multiple drug-resistant colonies were pooled together to establish stable cell lines. Three independent cell lines were established and tested for each construct.
GTPase Activation Assay-Affinity purification assays to measure the levels of endogenous GTP-bound RhoA and Cdc42 were performed using the Rho binding domain of Rhotekin (GST-C21) and the Cdc42 binding domain of PAK (GST-PAK) as described previously (17,18). Cells were serum-starved for 16 h before harvesting the lysates.
Small Interfering RNA (siRNA) Transfections and Secondary Focus Formation Assay-Validated siRNA oligonucleotides, which were designed to suppress endogenous ROCK I (Santa Cruz, sc-36432), ROCK II (Santa Cruz, sc-36433), PRK I (Santa Cruz, sc-36262), or MLC (Santa, Cruz, sc-35940) were transfected by SiLentFect TM lipid reagent (Bio-Rad) according to the manufacturer's protocol into NIH 3T3 cells, which were stably transfected by pCTV3H-dbs-HA6 or its cognate vector. Scrambled siRNA oligonucleotides (Santa Cruz, sc-37007) were also included in the analysis as negative controls. The medium was replaced 5 h after transfection, and cells were cultured in fresh medium for 2 h before they were used to perform a secondary focus formation assay. Secondary focus formation assays were performed as described previously (12). Briefly, 1 ϫ 10 3 transfected cells were mixed with 1 ϫ 10 6 untransfected NIH 3T3 cells and then plated on 60-mm-diameter dishes. Foci were scored after 7 days. Cells from parallel transfections were cultured in fresh medium overnight and then lysed and examined by Western blot analysis. In a parallel assay for growth inhibition, 1 ϫ 10 2 transfected cells were plated on 60-mm-diameter dishes. Colonies were counted after 7 days.
Subcellular Fractionation Analysis-Cellular fractionations were performed as described previously (12). Briefly, NIH 3T3 cells that stably express Dbs-HA6 or its cognate vector were serum-starved in Dulbecco's modified Eagle's medium supplemented with 0.5% bovine calf serum for 18 h. Cells were washed by cold phosphate-buffered saline twice and incubated on ice for 30 min with saline containing 10 mM Tris, pH 7.4, 1 mM MgCl 2 , and a protease inhibitor mixture (Calbiochem). Cells were homogenized and then centrifuged at 500 ϫ g at 4°C for 10 min. The supernatant was collected, supplemented with 1 M NaCl to a final concentration of 150 mM, and centrifuged at 100,000 ϫ g at 4°C for 60 min. The pellet was resolved in saline containing 10 mM Tris, pH 7.4, 1 mM MgCl 2 , 150 mM NaCl, 0.5% Triton X-100, and protease inhibitors. The protein concentrations of total, particulate, and supernatant fractions were determined by BCA protein assay kit (Pierce). Equal amounts of protein (40 g) from each fraction were analyzed by SDS-polyacrylamide gel electrophoresis, then transferred to Hybond-P polyvinylidene difluoride membrane (Amersham Biosciences) and analyzed by Western blot.
ROCK I Kinase Assay-The in vitro ROCK I kinase assay was performed as described previously (38). Briefly, NIH 3T3 cells that stably express Dbs-HA6 or cognate vector were transiently transfected with 5 g pCAG-ROCK I. Lysates were collected at 48 h, clarified by centrifugation, and then immunoprecipitated with an agarose-conjugated anti-Myc antibody (9E10, Santa Cruz). Beads were washed 3 times with buffer containing 50 mM Tris, pH 7.5, 10 mM MgCl 2 , 50 mM NaCl, 1 mM dithiothreitol, 10% glycerol, 0.03% Brij 35, and a protease inhibitor mixture (Calbiochem). The immunoprecipitate was then split into two parts, one of which was subjected to Western blot using an anti-Myc antibody (9E10, Santa Cruz) to confirm equal expression of Myc-tagged ROCK I. The second fraction was used in an in vitro kinase assay. Immunoprecipitates were washed twice in kinase assay buffer containing 50 mM Tris, pH 7.5, 1 mM EDTA, 10 mM MgCl 2 , 50 mM NaCl, 1 mM dithiothreitol, and 0.03% Brij 35. Samples were then resuspended in 30 l of kinase assay buffer containing 10 M cold ATP, 50 ng/l histone H1 (Calbiochem) and 135 nCi/l [ 32 P]ATP (Amersham Biosciences). After a 30-min incubation at 30°C, the reaction was terminated by the addition of Laemmli buffer. Samples were resolved by 10% SDS-polyacrylamide gel electrophoresis and subjected to autoradiography. Where indicated, cells were treated with 25 ng/ml recombinant Rat tumor necrosis factor-␣ (Sigma) for 3 h before harvesting the cells. Three independent experiments were performed on each of two independent cell lines.
PRK I Kinase Assay-The in vitro PRK-I kinase assay was performed as described previously (39). NIH 3T3 cells that stably express Dbs-HA6 or cognate vector were transiently transfected with 5 g of Myc-PRK I. Lysates were collected at 48 h, cleared, and then immunoprecipitated with an agarose-conjugated anti-Myc antibody (9E10, Santa Cruz). The beads were washed twice with 1 ml of buffer B containing phosphatebuffered saline, 1% Nonidet P-40, and proteases, twice with 1 ml of buffer C containing 10 mM Tris, pH 7.5, 0.5 M LiCl, and proteases, and three times with 1 ml of buffer D containing 10 mM Tris-HCl, pH 7.5, 1 mM EDTA, 50 mM NaCl, and proteases. Immunoprecipitates were then washed twice with 1 ml of kinase assay buffer containing 20 mM Tris-HCl, pH 7.6, 5 mM MgCl 2 , 40 mM KCl, 1 mM EDTA, 1 mM EGTA, 1 mM dithiothreitol, and a protease inhibitor mixture. Samples were then resuspended in 30 l of kinase assay reaction buffer containing 10 M unlabeled ATP, 50 ng/l histone H1 (Calbiochem, 382151) and 135 nCi/l [ 32 P]ATP (Amersham Biosciences). After a 7-min incubation at 30°C, the reaction was terminated by the addition of stop solution (200 mM ATP, 500 mM EDTA, pH 8.0). Samples were resolved by 10% SDSpolyacrylamide gel electrophoresis and subjected to autoradiography. As indicated, 40 mM arachidonic acid (Sigma, A3555) was added to the kinase assay reaction mix (40).
Immunostaining-Immunostaining of transiently transfected NIH 3T3 cells was performed as described previously (17). Briefly, NIH 3T3 cells were transiently transfected with 1 g of pAX142-dbs-HA6 using Lipofectamine reagent (Invitrogen). Cells were replated on coverslips at low cell density 24 h post-transfection. Cells were serum-starved (0.5% serum) for 16 -18 h before they were fixed in 100% ice-cold methanol on ice for 10 min. Fixed cells were then permeabilized and blocked in 0.1% Triton X-100, 3% bovine serum albumin in phosphate-buffered saline for 30 min. Coverslips were then incubated with appropriate primary antibodies for 1 h in 0.1% Triton X-100 with 3% bovine serum albumin and then in secondary antibodies for 1 h in the dark in 0.1% Triton X-100 with 3% bovine serum albumin. Coverslips were washed in phosphate-buffered saline and mounted on glass slides using SlowFade Antifade kits (Molecular Probes). Cells were viewed with an Zeiss inverted microscope equipped with an ApoTome-based imaging system (Zeiss). Axial stacks were captured for each cell, and a single axial plane is shown. Images were analyzed using Axiovision software. Immunostaining of F-actin was performed as described above except that cells were fixed in 3.7% formaldehyde in phosphate-buffered saline for 10 min at room temperature. Where indicated, cells were treated with 10 M Y-27632 for an additional 6 h of serum starvation.

RESULTS
A Dbs Mutant That is Selectively Impaired in RhoA Exchange Activity Shows Reduced Transformation in NIH 3T3 Cells-Recent structural studies have identified the key residues in Dbs that are responsible for promoting exchange activity on RhoA and Cdc42 (18,19). Using this information it is possible to design Dbs mutants that are selectively attenuated in their ability to productively interact with their substrates. For example, the Dbs(L759M/L766M) mutant shows normal exchange activity for RhoA but is completely impaired in Cdc42 exchange (18). Because this mutant exhibits normal transforming activity when expressed in NIH 3T3 cells, Cdc42 activation appears to be dispensable for Dbs-mediated transformation in this cell type. A new mutant of Dbs (Dbs(K758A)) was described recently that exhibits an opposite specificity to the Dbs(L759M/L766M) mutant when examined in an in vitro exchange assay (19). That is, this mutant shows normal exchange activity for Cdc42 but is significantly impaired in its activity toward RhoA. We reasoned that if Dbs targets RhoA in NIH 3T3 cells for transformation, then this mutant should be impaired in transforming activity in these cells. To test this possibility, an equivalent substitution was placed in the context of onco-Dbs. To confirm that this substitution specifically impairs RhoA exchange activity in vivo, we first examined the mutant by GST-based affinity precipitation assays for its ability to activate Cdc42 and RhoA in 293T and NIH 3T3 cells, respectively. We have shown previously that transient expression of onco-Dbs in 293T cells causes the accumulation of Cdc42 in the activated GTP-bound form (17), whereas expression in NIH 3T3 cells causes the accumulation of activated RhoA (18). Consistent with the in vitro data, we observed that the Dbs(K758A) mutant can activate Cdc42 in 293T cells to the same levels as onco-Dbs ( Fig. 1A) but is impaired in its ability to activate RhoA in NIH 3T3 cells (Fig. 1B). To determine whether the mutant is also impaired in transformation, we compared Dbs(K758A) with onco-Dbs in an NIH 3T3 cell primary focus formation assay (Fig. 1C ). Consistent with a role for RhoA in Dbs transformation, we observed that the Dbs(K758A) mutant had a transforming activity that was only 40 -60% of onco-Dbs despite being expressed at equivalent levels. When combined with our previous observations, these results provide strong genetic evidence that RhoA is a physiologically relevant target for Dbs transformation in this cell type.
Elevated Levels of Activated ROCK, but Not PRK, in NIH 3T3 Cells That Express Onco-Dbs-ROCK I and PRK I are effector kinases for RhoA that have been previously implicated in RhoA-mediated transformation (33,42). Thus, we wondered if either of these two kinases mediates Dbs transformation in NIH 3T3 cells. Although ROCK II and PRK II have not been implicated in RhoA transformation, they were included in this analysis as additional controls. As demonstrated by Western blot, NIH 3T3 cells express detectable levels of ROCK I, ROCK II, PRK I, and PRK II, and no changes in overall level of expression of these kinases are observed in response to transient expression of onco-Dbs and activated RhoA or stable expression of onco-Dbs ( Fig. 2A). To determine whether Dbs causes activation of PRK I or PRK II, blots were stripped and rep-robed with an antibody that detects the activated, phosphorylated form of both kinases. For both PRK I and PRK II, no differences were observed between vector, onco-Dbs, and RhoA(63L)-expressing cells, suggesting that Dbs does not activate either kinase in this cell type ( Fig. 2A). In contrast, treatment of parental cells with the phosphatase inhibitor  Antibodies are as described under "Materials and Methods." B, lysates were collected from NIH 3T3 cells that were treated for 10 min with calyculin A (100 nM) or that transiently express onco-Dbs (Dbs). Lysates were then examined by Western blot as described above. C, NIH 3T3 cells that stably express either cognate vector or oncogenic Dbs were transiently transfected with an expression plasmid that encodes Myc-tagged ROCK I. As an additional control Dbs-expressing cells were treated with tumor necrosis factor-␣ (TNF-␣) as described under "Materials and Methods." Lysates were collected at 48 h and examined by Western blot for expression of either ROCK I (c-Myc) or Dbs (HA). ROCK I was immunoprecipitated using an anti-Myc monoclonal antibody, and immunoprecipitates were then used in an in vitro kinase assay using histone H1 as a substrate. The lower band indicates levels of phosphorylated histone H1 (histone), whereas the upper band (ROCK I) indicates ROCK I autophosphorylation. Relative levels of kinase activity were determined using an Odyssey IR imager, normalized against ROCK1 expression, and expressed as integrated intensity units (lower bar graph). Data shown are an average of three independent experiments and show S.D. D, NIH 3T3 cells that stably express either cognate vector or oncogenic Dbs were transiently transfected with an expression plasmid that encodes Myctagged PRK I. As a positive control, immunoprecipitates were mock-treated or treated with arachidonic acid (AA). Lysates were collected at 48 h and examined by Western blot for expression of either PRK I (c-Myc) or Dbs (HA). PRK I was immunoprecipitated using an anti-Myc monoclonal antibody, and immunoprecipitates were then used in an in vitro kinase assay as described above. Relative levels of kinase activity were determined using an Odyssey IR imager, normalized against ROCK1 expression, and expressed as integrated intensity units (lower bar graph). Data shown are an average of three independent experiments and show S.D. calyculin A as a positive control for activation resulted in a dramatic increase in the levels of phosphorylated PRK I and PRK II (Fig. 2B).
Next we wished to know whether Dbs can activate ROCK I in the same cells. Unfortunately no antibodies are currently available that can be used to detect the activated form of ROCK I. Thus, we performed an in vitro kinase assay using exogenously expressed, Myc-tagged ROCK I that was immunoprecipitated from cell lines that stably express onco-Dbs (Fig. 2C ). As a positive control for ROCK I activation, onco-Dbsexpressing cells were also treated with tumor necrosis factor-␣. When compared with vector-expressing cells, ROCK I that was immunoprecipitated from Dbs-expressing cells, consistently showed a 3-fold increase in levels of trans-and autophosphorylation, and the level of activation was further increased (5-6-fold) when cells were treated with tumor necrosis factor-␣. When we performed an equivalent in vitro kinase assay using exogenously expressed, Myc-tagged PRK I (Fig. 2D), no difference was observed between the vector and Dbs-expressing cells (compare lanes 3 and 4). In contrast, treatment with the PRK I activator, arachidonic acid (40), resulted in a 3-fold increase in activity (compare lanes 1 and 2). These results suggest that ROCK I, but not PRK I, is activated in Dbs-transformed cells.
ROCK I and PRK I Are Translocated to the Plasma Membrane in Dbs-transformed Cells-It has been shown previously that endogenous ROCK I is translocated from the cytosol to the plasma membrane in response to expression of activated RhoA (43). If oncogenic Dbs activates ROCK I in a RhoA-dependent manner in NIH 3T3 cells, we reasoned that this may be reflected in a similar translocation event. To test this possibility we stably expressed onco-Dbs in NIH 3T3 cells and then performed cell fractionations (Fig. 3). Although we were able to detect ROCK I expression in both the particulate and soluble fractions (Fig. 3A), there was a substantial increase in the amount of ROCK I in the particulate fraction in the Dbs-expressing cells relative to the vector controls (Fig. 3B). The localization of ROCK I coincided with the distribution of onco-Dbs, which was also found predominantly in the particulate fraction (Fig. 3A, lower  panel). The overall distribution of ROCK II was not substantially changed in response to Dbs expression (Fig. 3B).
To further examine the possibility that Dbs may modify PRK activity, the equivalent blots were then examined for PRK I and PRK II distribution (Fig. 3). Similar to what we observed for ROCK I, we observed that PRK I was redistributed to the particulate fraction in Dbs-expressing cells, whereas PRK II was not.
Next we wished to more precisely characterize the translocation of ROCK I and PRK I in the Dbs-transformed cells. For this analysis cells that transiently express onco-Dbs were examined by indirect immunofluorescence to determine the cellular distribution of endogenous PRK I and ROCK I (Fig. 4). ROCK II was included in this analysis as an additional control. siRNAs targeted against each of the kinases were used to demonstrate the specificity of the antibodies (lower panels). Whereas endogenous ROCK I and PRK I exhibited dispersed punctate staining throughout the cytosol in control cells, in Dbs-expressing cells a discrete fraction of the ROCK I and PRK I immunoreactivity was observed at the cell periphery. An equivalent accumulation at the plasma membrane was not observed for ROCK II. These observations are consistent with the fractionation studies and suggest that the accumulation of PRK I and ROCK I in the particulate fraction can be partially explained by their translocation onto the plasma membrane.   JUNE 9, 2006 • VOLUME 281 • NUMBER 23

Dbs Transformation in NIH 3T3 Cells Requires ROCK-mediated
Signaling-Because the results of our kinase and cellular localization studies suggest that natively expressed ROCK I may be activated in Dbs-transformed cells, we wondered whether ROCK activation is necessary for Dbs transformation. For this analysis we utilized the Y27632 inhibitor, which has been shown previously to be a selective inhibitor of ROCK I and ROCK II. Cells were transiently transfected with either cognate vector or onco-Dbs, and then primary focus formation assays were performed in the presence or absence of the inhibitor. As an additional control, the inhibitor was also tested on an unrelated oncogene (Src) that has potent focus-forming activity in this cell type. Previous studies have shown that Src transformation in NIH 3T3 cells is independent of ROCK (42). In the presence of the inhibitor, onco-Dbstransforming activity was reduced to background levels, whereas the inhibitor had no effect on transformation by Src (Fig. 5A). We then confirmed that the inhibition could not be attributed to an effect on overall expression levels of ROCK I, ROCK II, PRK I, PRK II, or onco-Dbs (Fig. 5B). The inhibition could also not be attributed to a nonspecific inhibition of PRK I and PRK II activity since the levels of phospho-PRK I and phospho-PRK II also did not change.
To confirm the dependence of Dbs transformation on ROCK I signaling, we utilized a kinase-dead dominant-inhibitory version of ROCK I. When we co-expressed this mutant with onco-Dbs in NIH 3T3 cells, we observed a dose-dependent inhibition of Dbs transformation (Fig. 5C ). In contrast, the mutant had no effect on Src transformation, again confirming that Src and Dbs transform cells through distinct mechanisms.
Additional Members of the RhoGEF Family Transform NIH 3T3 Cells through a ROCK-dependent Manner-Because many members of the RhoGEF family were isolated based on their transforming activity in NIH 3T3 cells, we wondered whether they transform this cell type through an equivalent mechanism. We have previously described a panel of transforming derivatives of RhoGEF family members that includes Ect2, Lfc, and Lsc (34). NIH 3T3 cells were transiently transfected with all members of the panel, and then focus formation assays were performed in the presence or absence of the inhibitor (Fig. 5A). Similar to what we observed for Dbs, all members of the panel exhibited greatly reduced transforming activity when tested in the presence of the inhibitor. When viewed in the context of previous studies (31,44), these results strongly suggest that there is common ROCK-dependent mechanism through which RhoGEF family members transform NIH 3T3 cells.
Dbs Transformation Does Not Require PRK-mediated Signaling-To further examine the possible contribution of PRK-mediated signaling to Dbs transformation, we also determined whether expression of dominant-inhibitory versions of MKK3 and MKK6 could block Dbs transformation (Fig. 5D). Both of these kinases are downstream of PRK, and it has been shown previously that both of these mutants can block transformation by constitutively activated RhoA. Although a dominant inhibitory version of RhoA effectively blocked Dbs transformation, no effect was observed with the dominant-inhibitory MKK3 and MKK6 despite the fact that they were expressed at significant levels (Fig. 5D, lower panels). We conclude that the PRK I-mediated signaling pathway that has been previously implicated in RhoA transformation does not contribute to Dbs transformation in NIH 3T3 cells.
siRNA Targeted against ROCK I, but Not ROCK II, Blocks Dbs Transformation-To confirm a specific role for ROCK I in Dbs transformation, siRNAs were used to target ROCK I, ROCK II, and PRK I in Dbs-transformed cells (Fig. 6, A and B). For this analysis cells were transiently transfected with siRNAs and then examined in secondary focus formation (Fig. 6C ) and clonogenicity (Fig. 6D) assays. Although siRNAs targeted against ROCK I consistently inhibited Dbs transformation by ϳ45% (Fig. 6C ), we observed that ROCK II expression was also suppressed in these cells (Fig. 6B). However, siRNAs targeted against PRK I and ROCK II were much more specific in their action and had no effect on Dbs transformation. The reduced transforming activity associated with the ROCK I siRNAs could not be attributed to nonspecific growth inhibition as demonstrated by the clonogenicity assays (Fig. 6D). These results are consistent with genetic and pharmacological inhibitors and suggest that endogenous ROCK I, but not ROCK II or PRK I, is required for Dbs transformation.
Dbs Transformation Requires ROCK-mediated Phosphorylation of MLC-Because onco-Dbs causes profound changes in the actin cytoskeleton including the formation of stress fibers, we wondered whether targets of ROCK that regulate actino-myosin contraction contribute to transformation. Initially we measured the levels of phosphorylated cofilin and MLC2 in Dbs-transformed cells (Fig. 7A). Whereas we observed a 3-fold increase in phosphorylated MLC2 in Dbs-trans- formed cells relative to vector controls, no differences in activated cofilin were noted. Treatment of Dbs-transformed cells with the ROCK inhibitor reduced the level of MLC2 phosphorylation to basal levels ( Fig.  7A) and disrupted the formation of actin stress fibers in Dbs-transformed cells (Fig. 7E). Interestingly, the accumulation or cortical actin that is associated with Dbs expression was not affected by the inhibitor, suggesting that Dbs can also modify the actin cytoskeleton through ROCK-independent events. To determine whether phosphorylation of MLC2 is required for Dbs transformation, Dbs-transformed cells were transfected with siRNAs targeted at MLC (Fig. 7B) and then assessed in a secondary focus formation assay (Fig. 7C ). A reduction in transforming activity by about 40% was observed in the MLC suppressed cells, which is consistent with what we observed for siRNAs targeted against ROCK I in the equivalent assay (see Fig. 6). Successful silencing of MLC was demonstrated by Western blot using lysates derived from parallel transfections (Fig. 7B), and the reduced transforming activity associated with the MLC2 siRNAs could not be attributed to nonspecific growth inhibition (Fig. 7D). These results suggest that Dbs causes phosphorylation of MLC in a ROCK-dependent manner and suggests that actinomyosin contraction is required to support Dbs transformation.

DISCUSSION
Although members of the RhoGEF family are well known for their transforming activity in NIH 3T3 cells, the identification of the molecular mechanisms that underlie this activity has remained elusive. In recent studies (including the current one), we have identified RhoA as a relevant physiological target for Dbs transformation in this cell type (12,18). We have shown previously that onco-Dbs mutants that cannot activate Cdc42 retain full transforming activity (18), whereas in this current study we showed that a mutant that is selectively impaired in RhoA activity shows reduced transformation. Because Dbs transformation is blocked by RhoA inhibitors and activated mutants of RhoA produce foci that are phenotypically indistinguishable from Dbs foci, a role for RhoA in Dbs transformation seems apparent. Based on these observations we have attempted to identify RhoA effector proteins that may contribute to Dbs transformation. Although many of such proteins have been described, two recent studies suggest that pathways that are regulated by ROCK I and PRK I are essential for RhoA transforming activity (33,42). Thus, we chose to focus our attention on these two kinases.
PRK I is a lipid activated protein serine/threonine kinase that was originally identified as a binding partner for activated RhoA. Although the native function of PRK I is unknown, a recent study has implicated PRK I in RhoA transformation in NIH 3T3 cells (33). PRK I was shown to activate p38␥ in these cells through an MKK3-and MKK6-dependent mechanism, and dominant-inhibitory mutants of MKK3 and MKK6 were effective in blocking RhoA transformation. However, despite the fact that we are able to detect PRK I and PRK II expression in Dbstransformed cells, we found no evidence of elevated kinase activity and no evidence that the integrity of PRK-mediated signaling pathways is required to support Dbs transformation. Thus, although Dbs transforms NIH 3T3 cells in a RhoA-dependent manner, the mechanism of transformation by onco-Dbs and the constitutively activated RhoA(63L) mutant may be mechanistically distinct. Because constitutively activated RhoA mutants have a much weaker transforming activity than onco-Dbs, these mutants may not mimic activation of the endogenous GTPase by  Dbs and, thus, may utilize a distinct roster of signaling pathways to mediate transformation.
Unlike PRK I, the evidence linking Dbs transformation to ROCK I activation seems clearer. An elevated level of ROCK I kinase activity was observed in Dbs-transformed cells, and both genetic and pharmacological inhibitors of ROCK block Dbs focus-forming activity. Previous studies that utilized the same ROCK inhibitor have implicated ROCK signaling as necessary for transformation by both RhoA (42) and several members of the RhoGEF family (Tim, Dbl, and Net1) (31,44). In these studies however, it was unclear whether RhoGEFs directly activate ROCK to support transformation or whether endogenous ROCK I simply cooperates with RhoGEFs in transformation. In this current study we have found evidence to suggest that ROCK I is activated by onco-Dbs expression. Endogenous ROCK I is redistributed to the plasma membrane in response to Dbs expression, and Dbs can strongly activate ectopically expressed ROCK I.
Interestingly, Dbs had no discernible effect on the cellular distribution of ROCK II despite the fact that it is expressed at detectable levels in these cells. In addition, siRNAs targeted at ROCK I blocked Dbs transformation, whereas those targeted at ROCK II did not. Thus, Dbs-mediated activation of RhoA appears to be associated with the recruitment of only a subset of the available RhoA effectors in this cell type. This would be consistent with previous studies which suggest that Rho effectors can also serve as scaffolding proteins that mediate the association of GTPases with specific RhoGEFs (45,46).
This study expands the roster of members of the RhoGEF family that transform NIH 3T3 cells through a ROCK-dependent mechanism and suggests that RhoA may be a common target of transformation in this cell type. Interestingly, despite the fact that the foci that are induced by RhoGEFs and Src are virtually indistinguishable, both we and others see no effect of the ROCK inhibitor on Src transformation (42). Thus, if Src and RhoA transform NIH 3T3 cells through equivalent mechanisms, this would suggest that Src acts downstream of ROCK I activity in this pathway. Because activated ROCK I has no focus-forming activity when expressed alone in these cells, it is likely that additional pathways that are commonly activated by both RhoGEFs and Src are also necessary for the full transformed phenotype. In support of this we have observed that the ability of Dbs to stimulate cortical actin or to activate the SRE reporters 6 is relatively insensitive to the effects of the ROCK inhibitor, suggesting that Dbs can activate ROCK I-independent pathways in this cell type.
Our analysis of ROCK substrates in the context of Dbs transformation suggests an important role for MLC. We observed an elevated level of phosphorylated MLC in Dbs-transformed cells that was effectively blocked by the ROCK inhibitor. Suppression of MLC by siRNAs inhibited focus formation by Dbs, which implies a direct role for actinomyosin contraction in the transformation phenotype. The phosphorylation of MLC causes the bundling of filamentous actin into stress fibers (47,48), and ROCK has been shown to stimulate actino-myosin contractility by directly phosphorylating MLC (49). We have shown previously that cells that stably express onco-Dbs are more rounded and exhibit elevated actin stress fibers and a dense ring of cortical actin (41). Interestingly, we observed in this current study that the ROCK inhibitor blocks the formation of stress fibers, which is consistent with the downregulation of MLC but has little effect on the accumulation of cortical actin. This suggests that Dbs can activate changes in the actin cytoskeleton that are independent of ROCK but that such changes are dispensable for transformation.
Although it has been reported that transformation by the RhoGEF family members Dbl and Net1 is blocked by genetic inhibitors of ezrin (31), we were unable to see any evidence of increased ezrin phosphorylation in Dbs-transformed cells 6 nor were we able to see changes in phosphorylation status of cofilin. Thus, although it is possible that Dbs transformation is dependent on the activation status or integrity of these actin regulatory proteins, they do not appear to be targets for ROCK I in the context of Dbs transformation.