Regulation of RhoA GTPase and various transcription ...

MINI-REVIEW Regulation of RhoA GTPase and various transcription factors in the RhoA pathway† Running title: RhoA and tumorigenesis Jae-Gyu Kim1*, Rokibul Islam1*, Jung Yoon Cho1,2*, Hwalrim Jeong3*, Kim-Cuong Cap1, Yohan Park1, Abu Jubayer Hossain1,2, Jae-Bong Park1,2,‡ 1

Department of Biochemistry, 2Institute of Cell Differentiation and Aging, Hallym University College of Medicine, Chuncheon, Kangwon-do, 24252, 3Department of Paediatrics, Chuncheon Sacred Hospital Hallym University, Chuncheon, 24253, Kangwon-do, Republic of Korea

‡

Correspondence: Jae-Bong Park, Department of Biochemistry, Hallym University College of Medicine, Chuncheon, Kangwon-Do, Republic of Korea, 200-702; Tel, 82-33-248-2542; Fax, 82-33244-8425; E-mail, [email protected]

*Equally contributed.

†

This article has been accepted for publication and undergone full peer review but has not been through the copyediting, typesetting, pagination and proofreading process, which may lead to differences between this version and the Version of Record. Please cite this article as doi: [10.1002/jcp.26487]

Received 10 November 2017; Accepted 11 January 2018 Journal of Cellular Physiology This article is protected by copyright. All rights reserved DOI 10.1002/jcp.26487

This article is protected by copyright. All rights reserved

Abstract RhoA GTPase plays a variety of functions in regulation of cytoskeletal proteins, cellular morphology and migration along with various proliferation and transcriptional activity in cells. RhoA activity is regulated by guanine nucleotide exchange factors (GEFs), GTPase activating proteins (GAPs) and the guanine nucleotide dissociation factor (GDI). The RhoA-RhoGDI complex exists in the cytosol and the active GTP-bound form of RhoA is located to the membrane. GDI displacement factors (GDFs) including IκB kinase γ (IKKγ) dissociate the RhoA-GDI complex, allowing activation of RhoA through GEFs. In addition, modifications of Tyr42 phosphorylation and Cys16/20 oxidation in RhoA and Tyr156 phosphorylation and oxidation of RhoGDI promote the dissociation of the RhoA-RhoGDI complex. The expression of RhoA is regulated through transcriptional factors such as c-Myc, HIF-1α/2α, Stat 6 and NF-κB along with several reported microRNAs. As the role of RhoA in regulating actin-filament formation and myosin-actin interaction has been well described, in this review we focus on the transcriptional activity of RhoA and also the regulation of RhoA message itself. Of interest, in the cytosol, activated RhoA induces transcriptional changes through filamentous actin (F-actin)-dependent (“actin switch”) or -independent means. RhoA regulates the activity of several transcription regulators such as serum response factor (SRF)/MAL, AP-1, NF-κB, YAP/TAZ, β-catenin and hypoxia inducible factor (HIF)-1α. Interestingly, RhoA also itself is localized to the nucleus by an as-yet-undiscovered mechanism. This article is protected by copyright. All rights reserved

Key words: RhoA, SRF, MAL, AP-1, YAP/TAZ, β-catenin, HIF-1α/2α, transcription


Introduction The Rho GTPase subfamily is a member of Ras-related small GTP-binding family of proteins that include RhoA, Cdc42 and Rac1/2. These perform several important cellular functions including cytoskeletal rearrangement, regulation of cell morphology and motility along with transcription regulation and reactive oxygen species (ROS) production. Rho GTPases are activated by incorporation of GTP, catalyzed by guanine nucleotide exchange factors (GEFs); they are also inactivated by GTP hydrolysis, catalyzed by GTPase activating proteins (GAPs) (Bos et al., 2007; Buchsbaum, 2007; Moon and Zheng, 2003). As there are specific GEFs and GAPs for Rho GTPases, these are in turn regulated by a set of stimulants that control Rho activity for various cellular processes. Inactive Rho GTPases, which are associated with GDP, exist in the cytosol in a complex with guanine nucleotide dissociation inhibitor (RhoGDI). For activation of Rho GTPases, GEFs cannot directly act on the Rho GTPases-RhoGDI complex and the Rho GTPases first need to be dissociated from RhoGDI. This dissociation from RhoGDI is accomplished by GDI displacement factor (GDF). Once a Rho GTPase is GTP-bound, it is generally associated with the cell plasma membrane through its prenyl group (Fig. 1). Activated Rho GTPases transmit the Rho-mediated signal to downstream target proteins through binding to effector proteins such as Rho-associated coiled coil kinase (ROCK) and p21activated protein kinase (PAK). ROCK1 and ROCK2 are activated by RhoA and PAK1-6 are activated by Cdc42 and Rac1. It has been established that RhoA regulates filamentous actin formation through the axis of ROCK/LIM kinases1/2 (LIMKs1/2)/p-cofilin/F-actin and mDia/profilin/Factin with actin-myosin interaction through phosphorylation of myosin light chain kinase and myosin light chain phosphatase (Jaffe and Hall, 2005). Here in this review, we describe target proteins regulated by RhoA and other regulatory proteins in the cytoskeleton (Heasman and Ridley, 2008). A dysregulation of Rho GTPases, not surprisingly, is closely linked to a variety of diseases including various types of cancer. In particular, RhoA is involved in carcinogenesis when it is overexpressed, mutated or its activity differentially regulated. In such cases, over-expression of Rho A is


brought about transcriptionally and differential regulation of Rho activity occurs by post-translational modifications. Expression of RhoA level is regulated in the transcriptional level and several microRNAs (miRs). From these changes, RhoA in turn alters transcription of multiple target genes involved in several physiological processes, particularly those relevant to carcinogenesis. In this review, we focus on the regulatory functions of RhoA on transcriptional factors such as nuclear factor-κB (NF-κB), activator protein-1 (AP-1), serum response factor (SRF) coactivator, MAL/myocardin

related

transcription

factor

(MRTF)/megakaryoblastic

leukemia-1

(MKL1),

peroxisome proliferator-activated receptor γ (PPARγ), β-catenin and hypoxia inducible factor-1α (HIF-1α). There are two underlying mechanisms for RhoA GTPase regulating various transcriptional factors; namely, filamentous actin (F-actin)-dependent (“actin switch”) and F-actin-independent. Before discussing the transcriptional regulation changes by RhoA, regulators of RhoA activity are first described, as these factors in turn subsequently also affect RhoA targets.

Regulation of RhoA activity and expression Regulation of RhoA GTPase activity GEFs and GAPs: GEFs are the main activators of RhoA GTPase, whereas GAPs function to inactivate RhoA. There are several GEFs and GAPs for RhoA with a particular GEF or GAP acting on RhoA depending on the circumstance. In general, GEFs and GAPs have multiple domains, enabling them to interact with other proteins and lipids and for localization or becoming part of protein complex scaffolds. In some cases, GEF and GAP domains are in one protein, providing efficient means to interconnect with two signaling processes (reviewed by Bos) (Bos et al., 2007). There may also be temporal and special compartments of RhoA and GEFs/GAPs to regulate RhoA activity, formed in response to several specific stimuli. GEFs and GAPs are themselves regulated by protein-protein interactions, leading to becoming localized in specific areas or undergoing post-translational modifications. In the example of required localization to a particular region in the cell, p115RhoGEF/LARG is localized and binds to active heterotrimeric G-protein coupled receptors (GPCR) Gα12 and Gα13 in the plasma


membrane; this then leads to RhoA activation. Thus, RhoA needs to be localized near the Gα12 and Gα13, in order to be activated. In the case of post-translational modification, Vav in its active form is Tyr174 phosphorylated, which then interacts with target Rho proteins via their SH2 domains. Recently, it was shown that p-Tyr174 Vav2 binds to p-Tyr42 RhoA, resulting in RhoA activation; both of the above Tyr phosphorylations are brought about by Src tyrosine kinase (Kim et al., 2017b). In the case of GEF regulation by cellular localization, Net1, a RhoGEF, first exists in the nucleus, but in response to cell stimulation by EGF, it is acetylated at K93/95 and then localizes to the cytosol, where Net1 activates RhoA (Song et al., 2015). It should be noted that the regulation and the signaling pathway responsible for Net1 acetylation remains undefined. It is well known that p190RhoGAP inactivates RhoA, and P190RhoGAP and RhoA are key regulators of oligodendrocyte differentiation. RhoA inactivation by p190RhoGAP is critical for formation of cellular processes such as neurite outgrowth (Arthur and Burridge, 2001). In case of ARAP3, it is a Rap-dependent RhoGAP and is active against both Rho and Arf family GTPases. Interestingly, for ARAP3 recruitment to the membrane and activation by Rap, phosphatidylinositol 3,4,5-triphosphate (PIP3) is required (Krugmann et al., 2004).

GDFs as they dissociate the Rho-RhoGDI complex: Ezrin/radixin/moesin (ERM) were first identified to be GDFs for Rho. ERM directly bind to RhoGDI, reducing the activity of RhoGDI and facilitating Rho activation (Takahashi et al., 1997). Neurotrophin receptor p75NTR also directly binds to RhoGDI and initiates RhoA activation by releasing RhoA from RhoGDI. Either of the growth factors nerve growth factor (NGF) and basic fibroblast growth factor (bFGF) or cAMP induce neurite outgrowth from PC12 cells (Jeon et al., 2010a; Jeon et al., 2010b; Jeon et al., 2012). This is of interest as RhoA inactivation is closely related to neurite outgrowth and for NGF, it impairs the interaction between p75NTR and RhoGDI, leading to prevention of RhoA activation. However, Nogo and MAG (myelin-associated glycoprotein) induce p75NTR-RhoGDI complex formation, resulting in RhoA activation, and that inhibits neurite outgrowth. Thereby, p75NTR plays a role as a GDF to activate RhoA (Yamashita and Tohyama, 2003).


IκB kinase γ (IKKγ), also referred to as NEMO (NF-κB essential modulator), plays a role in the interaction with the RhoA-GDP/RhoGDI complex. In particular, RhoA binds to the N-terminal domain (amino acids 1-43) of IKKγ and RhoGDI binds to its large C-terminal domain (amino acids 100-419). As such, IKKγ significantly enhances GTP binding to RhoA via a GEF even in the presence of RhoGDI. It has been proposed that IKKγ functions as a GDF to dissociate the RhoARhoGDI complex before its GEF action on RhoA (Kim et al., 2014).

Modifications in RhoGDI regulating its affinity to Rho GTPases: Pak1 phosphorylates Ser101 and Ser174 of RhoGDI, leading to dissociation of the Rac1-RhoGDI complex, but not the RhoA-RhoGDI complex (DerMardirossian et al., 2004). Also, Src phosphorylates Tyr156 of RhoGDI, which decreases the affinity of RhoGDI to RhoA, Rac1 and Cdc42 (DerMardirossian et al., 2006). Recently, RhoGDIα was reported to be acetylated at Lys127 and Lys141, which reduces its binding to RhoA, leading to facilitation of RhoA activation (Kuhlmann et al., 2016b). It was then proposed that acetylation as well as phosphorylations of RhoGDI function as GDF events (Kuhlmann et al., 2016a). In addition, it was reported that oxidation of RhoGDI by hydrogen peroxide abolishes its complex formation with RhoA (Kim et al., 2017d), and hence, RhoGDI oxidation facilitates RhoA activation. It is of interest that oxidation of either RhoA or RhoGDI reduces one protein’s affinity towards its partner protein.

Role of phosphorylations in RhoA GTPase as part of post-translational modifications: Rho GTPases undergo post-translational modifications including phosphorylation, ubiquitination and AMPylation, which lead to changes in their function (Olson, 2016). It is well established that protein kinase A (PKA) phosphorylates RhoA Ser188 and increases RhoA’s affinity towards RhoGDI, resulting in RhoA inactivation (Lang et al., 1996). Cyclic GMP-dependent protein kinase (PKG) similarly phosphorylates RhoA Ser188 in vascular myocytes, leading to the same effect in RhoA through a direct interaction between RhoA (residue 1-44) and PKG1α (residue 1-59) 1(Kato et al., 2012; Sauzeau et al., 2000; Sawada et al., 2009). Cultured aortic smooth muscle cells from stroke-prone spontaneously hypertensive rats shows high level of activities of RhoA and ROCK, but PGK is This article is protected by copyright. All rights reserved

significantly down-regulated (Moriki et al., 2004). In addition, PKG1α mutant mice display hypertension (Michael et al., 2008), as the activation of RhoA and ROCK induces contractile vascular muscle (Loirand et al., 2006). In addition, AMP kinase α1 (AMPKα1) phosphorylates RhoA Ser188 in vascular smooth muscle cells in response to estradiol, also leading to RhoA inhibition (Gayard et al., 2011). Recently, ERK was reported to phosphorylate RhoA Ser88 and Thr100 which then upregulates the activity of RhoA upon EGF stimulation, although the signaling downstream components were not clearly established in report (Tong et al., 2016). ERK also phosphorylates another GTPase, Rac1, at Thr108, thereby reducing Rac1 activity upon EGF exposure of the cells. P-Thr108 Rac1 is localized to the nucleus, leading to Rac1 separation from its GEFs which are localized to the cytosol (Tong et al., 2013). Tyr34 and Tyr66 residues of RhoA are respectively localized to its switch I and switch II regions, and their post-translational modifications may give rise to activity changes in RhoA. In fact, phosphorylation of these two residues in RhoA crucially prevents its binding to effector proteins and negatively contributes to RhoA function (Uezu et al., 2012). Rac1 is also phosphorylated by focal adhesion kinase (FAK) and Src at its Tyr64, which corresponds to Tyr66 in RhoA; this leads to Rac1 readily binding to RhoGDI and consequently to inactivation of Rac1 (Chang et al., 2011). Similarly, in response to EGF, Cdc42 is phosphorylated by Src at its Tyr64, which also corresponds to RhoA Tyr66, enhancing the interaction of Cdc42 with RhoGDI and leading to its inactivation (Tu et al., 2003). It is noteworthy, however, that Tyr42 phosphorylation of RhoA contrary to its Tyr34 and Tyr66 phosphorylation is critical for RhoA activation in response to ROS, leading to cell proliferation and tumorigenesis, which is more described in the section below (Kim et al., 2017b).


Interaction of Prion protein with RhoA: The inactivation of RhoA is necessary for neurite outgrowth in response to NGF, bFGF, or cAMP in PC12 cells (Jeon et al., 2010a; Jeon et al., 2010b; Jeon et al., 2012). Specifically, p190RhoGAP, Rap-dependent RhoGAP (ARAP3) and RhoA phosphorylated at Ser188 by PKA lead to RhoA inactivation in response to NGF, bFGF or cAMP (Jeon et al., 2010a; Jeon et al., 2010b; Jeon et al., 2012). Recently, the cellular prion protein (PrPC) was implicated in neuritogenesis and neuronal differentiation (Kim et al., 2017a; Pietri et al., 2013) with PrPC recruiting both RhoA and p190 RhoGAP and facilitating RhoA inactivation in response to NGF stimulation in PC12. In addition, the pathogenic version of the prion protein, PrPSc, interferes with both RhoA inactivation and neurite outgrowth of PC12 cells upon cellular exposure to NGF (Kim et al., 2017a).

Mutations of RhoA gene in cancer cells RhoA gene mutants have recently been described in several cancer types including diffuse-type gastric carcinoma, angioimmunoblastic T cell lymphoma and peripheral T cell lymphoma (Porter et al., 2016). The R5Q, G17E and Y42C RhoA mutations are recurrent, and contrary to wild-type RhoA, expression of reconstituted both G17E and Y42C RhoA in SW948 colon cancer cells which previously had their RhoA knocked-down was able to rescue growth defects in the cells (Kakiuchi et al., 2014). The G17V RhoA mutation in T cell lymphoma also appears to act as a dominant negative mutation for RhoA, with RhoA losing its GTP binding and disabling the functioning of wild-type RhoA (Manso et al., 2014; Palomero et al., 2014; Sakata-Yanagimoto et al., 2014; Yoo et al., 2014). Expression of RhoA G17V increased proliferation in Jurkat cells. However, this is not congruent to the previous reports that overexpression of wild-type RhoA promoted tumorigenesis. It should be noted that it is not clearly addressed how these mutants are related to the proliferation of the cancer cell. For the Y42C mutant, we speculate that in the cancer cell, the mutation may give rise to sulfenic, sulfinic or sulfonic negative charge when Cys42 of RhoA is oxidized; this is similar to pTyr42 RhoA, the negatively charged form of Tyr42 in RhoA, which is positively correlated with tumor stage in breast cancer (Kim et al., 2017b). It is notable that mutations at Y42 reduce downstream activation of PKN but not mDia and ROCK1 (Sahai et al., 1998). This means that modification of This article is protected by copyright. All rights reserved

Tyr42 may change its effector proteins for cell behavior such as proliferation. On the other hand, RhoA G17A, which is unable to bind to GDP or GTP, plays the role of the bait for GEFs binding (Arthur et al., 2002; Waheed et al., 2012); this is similar to Ras G15A mutant, which forms a stable complex with CDC25, a GEF for Ras (Chen et al., 1994). For another RhoA mutant, RhoA G17V, we deduce that in a cancer cell, it may recruit unidentified effector binding proteins and stimulate cell proliferation.

Transcriptional regulation of RhoA gene It has been well established that there is overexpression of Rho GTPase in cancer cells. Although a detailed mechanism for this overexpression has not been established, a recent paper described the transcriptional complex for RhoA expression (Chan et al., 2010). The Myc-Skp2-Miz1p300 transcriptional complex binds to the RhoA promoter at about the minus-2 kb region for expressing RhoA. Deficiency of this complex impairs RhoA expression, cell migration, invasion and metastasis in breast cancer cells. Also, overexpression of Myc-Skp2-Miz1 complex is observed in metastatic human cancers along with increased RhoA expression. HIF-1 binds to and activates transcription of mRNA of RhoA gene at the minus-1.1 kb region, in addition to expression of proteins that in turn activate MLC and FAK, leading to increased motility of breast cancer cells (Gilkes et al., 2014). Promoter of RhoA gene also demonstrates binding to STAT6 (78-70 base upstream) and NF-κB (84-74 base upstream) in human bronchial smooth muscle cells in response to IL-13 and TNF-α (Goto et al., 2010). In addition, the CCAAT box (68-64 base upstream) in the promoter serves as a binding site for various transcription factors that include NFYA, NFYB, CEBPB and CEBPD (Nomikou et al., 2017). In addition, NO/PKG elevates RhoA level through RhoA protein stability and RhoA gene transcription. NO/PKG phosphorylates ATF-1 and subsequent binding to cAMP-response element (CRE) (upstream 118 base upstream) in RhoA promoter (Sauzeau et al., 2003) (Fig. 2).

It is noted that RhoA activation is closely linked to increased c-Myc expression through NFκB activation or increased β-catenin levels (Kim et al., 2017b; Kim et al., 2017c; Kim et al., 2017d). This article is protected by copyright. All rights reserved

Directly downstream of RhoA, ROCK1 directly interacts with and phosphorylates c-Myc at Thr58 and/or Ser62, resulting in stabilization of c-Myc and activation of its transcriptional activity (Zhang et al., 2014). Nuclear ROCK2 also phosphorylates p300 acetyltransferase, leading to an increase in overall acetyltransferase activity in the region (Tanaka et al., 2006). We, therefore, deduce a positive feedback loop of the initial RhoA/ROCK activation, resulting in c-Myc expression/p300 activation or NF-κB activation, and finally leading to increased RhoA expression in cancer cells. It is noteworthy that loss of RhoA causes a deficiency in oncogenic c-Myc levels, as c-Myc is a master transcription factor in turning on anabolic metabolism and promoting cell growth in many cancer types. It is known that the RhoA-SRF axis directly interacts with the c-Myc pathway in regulating glutaminase expression for utilization of glutamine in cancer cells (Haikala et al., 2016).

Role of microRNAs in regulating RhoA expression There are several microRNAs shown to regulate RhoA expression in response to various stimuli in given cells. In one instance, TGF-β increases miR-155 expression in normal murine mammary gland (NMuMG) epithelial cells, and elevated levels of miR-155 are frequently detected in invasive breast cancer tissues (Kong et al., 2008). miR-155 also reduces RhoA expression, which results in interference of endothelial to mesenchymal transition in response to TGF-β (Bijkerk et al., 2012). In addition, miR-155 is differentially expressed under normoxic and hypoxic conditions. Hypoxia induces miR-155 as the promoter of miR-155 has a hypoxia response element (HRE). However, miR-155 targets the HIF-1α mRNA, suggesting that miR-155 induction by hypoxia contributes to a negative-feedback loop for resolution of HIF-1α activity in cells exposed to prolonged hypoxia, leading to an oscillatory action of HIF-1α-dependent transcription (Bruning et al., 2011). Other examples of microRNAs regulating RhoA levels include miR-340 suppressing RhoA expression and overexpression of miR-340 inducing dendrite formation. UVB was shown to induce miR-340 and dendrite formation in a melanocyte cell line (Jian et al., 2014). miR-133b also promotes neurite outgrowth by targeting RhoA in PC12 cells (Lu et al., 2015). In addition, miR-146a suppresses RhoA expression in MDA-MB231 cells (Liu et al., 2016). Another microRNA, miR-31, also reduced RhoA expression in gastric cancer cells MKN-45 (Korourian et al., 2017). Loss of miRThis article is protected by copyright. All rights reserved

122 was also shown to be associated with poor prognosis in hepatocellular carcinoma progression along with cell migration and invasion and interestingly, RhoA 3’ UTR may be a binding site for miR122 (Wang et al., 2014). Indeed, the over-expression of miR-122 inhibits cytoskeletal structure, RhoA/ROCK signaling, cell adhesion, migration and invasion, along with mesenchymal-epithelial transition (MET) as it targets RhoA expression (Wang et al., 2014). In addition to RhoA, Cux1, Iggap, Mapre1, Nedd4i and Slc25a34 are also direct targets for miR-122 (Hsu et al., 2016). miR-185 also targets the 3’UTR of RhoA and Cdc42 in colorectal cancer and effectively inhibits cell proliferation (Liu et al., 2011). Expression of certain microRNAs can also lead to increased expression of RhoA as part of a network. In one study, a JNK inhibitor suppressed chondrogenic differentiation, likely through Rac1 inhibition, which results in up-regulation of miR-34a and RhoA. Blockade of miR-34a decreases expression of RhoA protein and stress fibers, along with up-regulation of Rac1 and type II collagen (Kim et al., 2012).

Transcription factors and RhoA Regulation of transcription factor MRTF and SRF with respect to RhoA The serum response element (SRE) for c-fos gene recruits and forms a ternary complex with TCF (ternary complex factor) and SRF (serum response factor). Serum, lysophosphatidic acid (LPA) and activated heterotrimeric G-proteins activate SRF through RhoA. In a pioneering research, it was demonstrated that RhoA regulated transcriptional activity (Hill et al., 1995). However, the molecular mechanism by which RhoA activates SRF was reported several years later. Upon serum stimulation, it was shown that MAL/MRTF/MKL1 binds to SRF, leading to SRF activation as a coactivator. In a resting state, MAL binds to the monomeric G-actin that keeps MAL/MRTF/MKL1 in the cytosol. Upon serum stimulation, RhoA is activated; this induces polymeric fibrous actin (F-actin), to which MAL/MRTF/MKL1 cannot bind. MAL protein is thus released by F-actin formation and it translocates to the nucleus where MAL/MRTF/MKL1 forms a complex with SRF on SRE, leading to activation of


SRF (Miralles et al., 2003). Elucidation of this RhoA/ROCK/F-actin pathway was seminal in revealing a role for actin filament formation in transcriptional regulation. MAL/MRTF/MKL1 can also be a repressor of a specific transcription factor such as peroxisome proliferator-activated receptor γ (PPARγ). Adipocyte differentiation is mainly mediated by PPARγ, a master transcriptional regulator of adipogenesis. As such, the transcriptional regulator MAL/MRTF/MKL1 has a preventive effect on adipogenic differentiation with MAL/MRTF/MKL1 and PPARγ being mutually antagonistic. As noted above, MAL/MRTF/MKL1 binds to the monomeric Gactin and not to F-actin, and G-actin prevents nuclear translocation of MAL/MRTF/MKL1. Thereby, RhoA activity needs to be diminished to a certain threshold for PPARγ activation and adipocyte differentiation (Nobusue et al., 2014) (Fig. 3).

Transcription factor YAP/TAZ and RhoA Changes in cell density and cell-matrix interactions can trigger MST1/2 – LATS1/2 – YAP/TAZ phosphorylation. Phosphorylated YAP/TAZ is a target for degradation; however, when dephosphorylated, YAP/TAZ moves to nucleus, where it mainly activates TEAD, leading to cell proliferation (Moroishi et al., 2015). RhoA activation is required for YAP/TAZ activation, although its detailed mechanism remains to be discovered. Recently, it was revealed that cancer-associated Gαq and Gα11 mutants, which are involved in RhoA activation, activate YAP through inhibiting LATS, leading to dephosphorylation of YAP (Yu et al., 2014). Inhibition of HMG-CoA reductase, as the enzymatic reaction is a limiting step in producing geranylgeranyl group and is required for RhoA prenylation, impairs YAP activity and nuclear accumulation (Sorrentino et al., 2014). Furthermore, cancer-associated Gαq and Gα11 mutants stimulate the Rho GEF, TRIO, leading to activation of RhoA and Rac1 and F-actin. The factor AMOT is available to bind to F-actin instead of YAP, resulting in the release of YAP and its nuclear translocation (Feng et al., 2014). The above pathway may address one mechanism in RhoA-mediated activation of YAP; however, it is of note that YAP was also recently reported to induce transcription of ARHGAP29 in suppressing RhoA activity, which reduces cytoskeletal rigidity and promotes metastasis of cancer cells (Qiao et al., 2017).


Transcription factor Sox9 and RhoA The dedifferentiation of primary chondrocytes into fibroblast morphology accompanies an induction of RhoA. Conversely, a loss of RhoA or expressing a dominant negative RhoA markedly enhances chondrogenesis. Also, Rho antagonist C3 transferase induces chondrocyte gene expression. Sox9 may play a role in RhoA mediated effects on chondrogenesis as RhoA-mediated modulation of actin-polymerization regulates Sox9 activity and Sox9 in turn positively regulates expression of chondrocyte markers (Kumar and Lassar, 2009).

Transcription factor AP-1 and RhoA AP-1 is a transcription factor formed from the homo- or heterodimer of c-Fos and c-Jun family proteins. SRF is activated by serum stimulation and that induces c-Fos expression with c-Jun becoming phosphorylated and activated by Jun N-terminal kinase (JNK). The AP-1 complex of cFos/p-c-Jun is required for cell cycle and transformation by several oncogenes. From the study of growth factors leading to c-Jun activation, the RING-domain-containing protein, RING domain AP-1 co-activator-1 (RACO-1), was shown as a c-Jun co-activator; RACO-1 is regulated by growth factors and it is stabilized through MEK/ERK activation, allowing stimulation of AP-1 (Davies et al., 2010). The molecular mechanism for activating AP-1 through RhoA involves RhoA/ROCK phosphorylating JNK, which then phosphorylates c-Jun and ATF2 upon LPA cell activation. The stability of c-Jun is increased by phosphorylation, leading to an increase in c-Jun levels. RhoA/ROCK also activates c-Fos through SRF/MAL attributed by F-actin action. Stimulation of cJun and c-Fos then leads to activation of AP-1, which is a complex of active c-Fos/p-c-Jun (Marinissen et al., 2004). For Cyr61, also known as CCN1, it is an angiogenic factor and is secreted and present in the extracellular matrix as a matricellular protein. Cyr61 is overexpressed in invasive and metastatic human breast cancer cells and tissues. MCF-7 breast cancer cells made to express Cyr61 become estrogen E2-independent and also anchorage-independent; as such, Cyr61 is a tumor-promoting factor; it is also shown to be a key regulator of breast cancer progression (Tsai et al., 2002). With regards to RhoA involvement in Cyr61 signaling, thrombin activates the G12/13 receptor, stimulating a This article is protected by copyright. All rights reserved

RhoGEF, which activates RhoA. Consequently, thrombin induces AP-1-mediated Cyr61 expression through RhoA, shown in 1321N1 glioblastoma cells (Walsh et al., 2008). Besides AP-1, Cyr61 expression is also regulated by a great number of transcription factors including NF-κB, MRTF-A and YAP, all of which are downstream targets of RhoA signaling (Jun and Lau, 2011).

Transcription factor β-catenin and RhoA Aberrant mutations in adenomatous polyposis coli (APC) and β-catenin are found in 90% of sporadic colon cancer cases. In addition, colorectal tumors also express elevated levels of Wnt3 (Voloshanenko et al., 2013). Activated RhoA has been shown to contribute to β-catenin accumulation in response to Wnt3A in HEK cells (Kim et al., 2017c). Downstream of RhoA, ROCK phosphorylates glycogen synthase kinase (GSK)-3β in response to Wnt3A and active ROCK has been shown to directly phosphorylate GSK-3β at Ser9 in vitro. As p-Ser9 GSK-3β is not relevant to β-catenin stabilization (McManus et al., 2005), RhoA/ROCK stabilizes β-catenin through an as yet unknown mechanism(s). The increased expression of c-Myc and cyclin D1 from Wnt3A stimulating the cell proliferation is governed by RhoA and ROCK1 (Kim et al., 2017c). RhoA/ROCK-mediated βcatenin activation from Wnt3A acting on the cell surface also induces expression of chemokines such as MIP-1α, which also leads to an increase in cell migration (Kim et al., 2017c). It should be noted that whether RhoA activation is critical for β-catenin accumulation and cancer progression remains controversial. It has been observed that inactivation of RhoA can contribute to colorectal cancer progression and metastasis through activation of Wnt/β-catenin signaling, and in colorectal cancer, RhoA plays a role as a tumor suppressor (Rodrigues et al., 2014). Therefore, the discrepancy between RhoA activity and β-catenin accumulation may be attributed to differences in cell types/tissues under study, and it warrants further study.

Transcription factors STAT3, GATA-4 and HIF-1α in relation to RhoA Active RhoA, Rac1, and Cdc42 could all mediate STAT3 Ser727/Tyr705 phosphorylation and the resultant nuclear translocation of STAT3; however, active RhoA, Rac1, and Cdc42 are not able to form a stable complex with STAT3. Moreover, STAT3 is required for the RhoA-induced NF-κB and This article is protected by copyright. All rights reserved

cyclin D1 transcription and is involved in NF-κB nuclear translocation. Thus, loss of STAT3 expression inhibits RhoA-promoted cell proliferation (Debidda et al., 2005). In relation to GATA-4, RhoA potentiates GATA-4 through p38MAPK with phosphorylation of GATA-4 activation domain, thus ensuring GATA-4 binding to target promoters as shown for cardiac promoters that control sarcomeric reorganization (Charron et al., 2001). For hypoxia-induced HIF-1α expression, Rho/ROCK signaling is for preventing HIF-1α degradation in endothelial cells (Takata et al., 2008). Moreover, C3 toxin, which inhibits RhoA, reduces HIF-1α protein accumulation during hypoxia (Hayashi et al., 2005; Turcotte et al., 2003). Hypoxia induces RhoA and β-catenin in a 2D cell culture system, but suppresses RhoA and βcatenin in a 3D one, suggesting that the microenvironment of cells is critical for determining this molecular switch (Ozturk et al., 2017). The overexpression and activation of Rho GTPases during hypoxia also depend on ROS production (Turcotte et al., 2003).

Transcription factor NF-κB relationship with RhoA RhoA has been shown to activate NF-κB in multiple reports (Kim et al., 2006); however, additional details in the various pathways need to be established. In the TGF-β1 signaling pathway, RhoAGTP activates ROCK1, which in turn directly phosphorylates IKKβ at Ser177/Ser181, leading to activation of IKKβ. Then, IKKβ phosphorylates IκB, resulting in IκB degradation. Consequently, NFκB dimeric form of p50/p65 released from IκB moves to the nucleus, where NF-κB induces expression of target genes (Kim et al., 2014). Reactive oxygen species (ROS) are important for NF-κB activation in the induction of inflammation. ROS have been reported to inhibit RhoA through p190RhoGAP activation. The mechanism involves ROS inhibiting the low-molecular weight protein tyrosine phosphatase (LMWPTP), leading to tyrosine phosphorylation and activation of p190RhoGAP. As Rac is involved in superoxide production, this indicates that Rac downregulates RhoA through p190RhoGAP activation (Nimnual et al., 2003). Indeed, we discovered that hydrogen peroxide as low as 1 μM induces Tyr42 phosphorylation and Cys16/20 oxidation of RhoA and subsequently its activation (Kim et al., 2017b; Kim et al., 2017d). Actually, Rho GTPases are regulated by the redox-state of This article is protected by copyright. All rights reserved

the cell (Hobbs et al., 2014). In particular, Cys16/20 in RhoA are oxidized to form a disulfide bond (Heo et al., 2006). RhoA-GTP respectively oxidized and phosphorylated at Cys16/20 and Tyr42 in response to hydrogen peroxide readily associates with the region of amino acids 100-200 in IKKγ. RhoA-GTP stimulates IKKβ that concurrently binds to IKKγ, juxtaposing with RhoA on IKKγ. Consequently, activated IKKβ phosphorylates IκB, leading to IκB degradation and NF-κB activation. NF-κB ensures increased expression of c-Myc and cyclin D1 and consequently increased cell proliferation (Kim et al., 2017b; Kim et al., 2017d). It is noteworthy that RhoA both oxidized at Cys16/20 and phosphorylated at Tyr42 is reduced in its affinity to RhoGDI (Kim et al., 2017b; Kim et al., 2017d). Tyr42 of RhoA is a substrate for Src tyrosine kinase with Src also phosphorylating and activating Vav2, a GEF for RhoA. Vav2 has an SH2 domain for binding specific p-Tyr-containing proteins. As such, the region of p-Tyr42 of RhoA may be a binding site for Vav2, an interaction that ensures increased incorporation GTP to RhoA. In tumor xenograft experiments, the 4T1 breast cancer cells expressing either wild-type RhoA (WT RhoA) or the phospho-mimic mutant Y42E RhoA gave rise to tumorigenesis; this did not occur with the dephospho-mimic mutant RhoA Y42F or the oxidative resistant mutant RhoA C16/20A (Kim et al., 2017b; Kim et al., 2017d). Furthermore, p-Tyr42 Rho has been detected in breast cancer tissues from patients and levels of p-Tyr42 Rho, p-Tyr416 Src and p-Ser527 p65/RelA positively correlate (Kim et al., 2017b). Thereby, we speculate that RhoA Tyr42 phosphorylation and Cys16/20 oxidation are likely to be critical for tumorigenesis in response to ROS or other cancer-promoting signals. As we utilized hydrogen peroxide as a ROS source to stimulate several cell types, it was also reported that ROS contributes to Src activation through hyperoxidation of Src Cys245 and Cys487 residues, leading to Tyr416 phosphorylation of Src (Giannoni et al., 2005). There are several sources of cellular ROS including mitochondria and certain enzymes that include NADPH oxidase (NOX). NOX has been well established as a source of ROS. In particular, Rac1/2 is involved in activation of p67PHOX, a component of NOX, leading to superoxide generation (Bokoch and Zhao, 2006). Here, we emphasize that RhoA has also been reported to regulate superoxide production through indirect activation of NOX (Kim et al., 2004; Moon et al., 2013). In This article is protected by copyright. All rights reserved

addition, Rap1 and RhoA have been reported to have complementary additive functions for superoxide production (Li et al., 2012). It is noteworthy that a co-culture of macrophages and cancer cells in vitro generates superoxides through production of soluble factor(s) but not through a physical interaction between these cell types, all leading to increased p-Tyr42 RhoA levels (Kim et al., 2017b). Conditioned media from the co-cultures stimulate superoxide production from both macrophages and cancer cells. These results remind us that tumor associated macrophages (TAMs) are critical for tumorigenesis, possibly helped through superoxide produced from both cancer cells and macrophages, leading to increased p-Tyr42 RhoA levels and subsequent cell proliferation. We speculate other factors in addition to ROS could also produce increased p-Tyr42 RhoA levels and consequently lead to NF-κB activation and cell proliferation. The superoxide produced in co-cultured cancer cells with macrophages is abolished by Rho inhibitor Tat-C3, suggesting that Rho GTPase is also involved in superoxide formation from cancer cells and tumor associated macrophages (unpublished data) (Fig. 4). Cell proliferation in response to hydrogen peroxide is associated with increased expression of cyclin D1 and c-Myc expression along with repression of p21Cip1/Waf1 levels. These responses are regulated by Rho and ROCK in that treatment with either the Rho inhibitor Tat-C3 or the ROCK inhibitor Y27632 reverses these changes (Kim et al., 2017b). Indeed, it has been well established that expression of cyclin D1 and p21Cip1/Waf1 are regulated by RhoA activity. Also, adhesion of NIH 3T3 cells to fibronectin stimulates RhoA activity along with cyclin D1 induction and p21 Cip1/Waf1 suppression in RhoA-dependent fashion (Danen et al., 2000). However, there has been a contentious report on RhoA activity and these target changes. In hematopoietic stem cells, low RhoA activity was associated with a higher proliferation rate and increases in cell cycle, most likely through decreasing p21 Cip1/Waf1 and increasing cyclin D1 levels (Ghiaur et al., 2006). Since Rac1/2 along with cyclin D1 induction is also shown to be essential for proliferation of hematopoietic progenitor cells (Gu et al., 2003) and RhoA and Rac1/2 act in an antagonistic manner, it is proposed that inactive RhoA can induce Rac1/2 activation along with increased cell proliferation. This


discrepancy in role of RhoA with respect to cell proliferation may be attributed to differences in cell types and stimuli.

Nuclear localization of RhoA Actin polymerization occurs not only in the cytoplasm but also in the nucleus. Polymerization of nuclear actin may directly regulate the amount of free MAL/MRTF/MKL1 and along with its association to SRF in the nucleus, it controls MAL/MRTF/MKL1-dependent transcription. Nucleartargeted constitutively active mutant RhoA (NLS-RhoA L63), mDia and MAL/MRTF/MKL1 induce SM22 expression, thereby inducing differentiation of smooth muscle cells (Staus et al., 2014). In that study, there were also increased levels of activated RhoA in the nucleus of the cells, following stimulation of the cells sphingosine-1-phosphate (S1P). We also observed that RhoA was localized to the nucleus in HeLa cells (Kim et al., 2014). Furthermore, several RhoGEFs including Net1, Ect2 and p115GEF/LARG are also found in nucleus (Chalamalasetty et al., 2006; Garcia-Mata et al., 2007; Schmidt and Hall, 2002; Staus et al., 2014). It is noteworthy that active Net1 activates RhoA in the nucleus when DNA undergoes damage with ionizing irradiation (Dubash et al., 2011). This result suggests that Net1/RhoA signaling pathway in nucleus of the cell is implicated in the DNA damage response. The detailed mechanism of nuclear RhoA activation, however, still remains undiscovered and it is yet not clear whether nuclear RhoA acts to induce actin filament formation or that RhoA itself performs novels functions such as transcriptional regulation.

Perspectives It is well established that tightly regulating the activity of RhoA is critically relevant to cellular processes in cells such as neuronal cells; on the other hand, overactive Rho GTPases are also required for tumorigenesis. Metabolically, increased glutamine (Gln) use is critical for various cancer types, as it replenishes the citrate cycle intermediates for ATP production and with the input of pyruvate being deficient as manifested by the high glycolytic metabolism of the Warburg effect. It is noteworthy that Rho GTPases and NF-κB activate cellular glutaminase activity converting glutamine


to glutamate, although its detailed mechanism still remains to be discovered (Wang et al., 2010). Modulation of metabolism by RhoA may be critical for tumorigenesis as RhoA mutations have also been discovered in several cancer tissues. The resultant functional changes due to many of these mutations in RhoA remain to be described and there is the added complication of the RhoA activity changes that may not be directly implicated in tumorigenesis. This suggests other mechanisms relevant to tumorigenesis such as recruitment of various factors to a given mutant RhoA and not relevant to RhoA activity changes. Besides activity changes in RhoA due to post-translational modifications, the RhoA expression changes at the transcriptional and translational levels also occur and are also critical for tumorigenesis. This is in par with RhoA playing a role as a regulator of various transcriptional factors governing cell proliferation. In addition, it is noteworthy that RhoA is also localized in the nucleus, where RhoA may perform its own unique functions in regulating cell physiology. Thereby, further studies on the transcriptional regulation of RhoA and those by RhoA need to be performed for a more detailed understanding of the tumorigenesis process.

Acknowledgements: This research was supported by the Basic Science Research Program of the National Research Foundation of Korea (NRF) (NRF-2015R1D1A1A01060393 for JBP; 2017R1D1A1B03029091 for JGK), and the Center for Women In Science, Engineering and Technology (WISET) Grant, funded by the Ministry of Science and ICT under the Program for Returners into R&D (WISET-2017-373 for JYC), and Hallym University (HRF-201303-017 for JBP).

Conflicts of interest: None.

The abbreviations used are as follows: CA, constitutively active; DMEM, Dulbecco’s modified Eagle’s medium; DN, dominant negative; GAP, GTPase activating protein; GDF, GDI displacement factor; GDI, guanine nucleotide dissociation inhibitor; GEF, guanine nucleotide exchange factor; GSH, glutathione; HIF, hypoxia inducible factor; IκB, inhibitor of NF-κB; IKK, IκB kinase; MAL/myocardin related transcription factor (MRTF)/megakaryoblastic leukemia1 (MKL1); NF-κB, nuclear factor-κB; NEMO, NF-κB essential modulator; PKA, protein kinase A; PKC, protein kinase C; Rhotekin-RBD, Rhotekin Rho-binding domain; ROCK, Rho-dependent coiled-coil kinase; ROS, reactive oxygen species; SRF, serum response factor; TAM, tumor associated macrophage; WT, wild-type


Figure Legends Fig. 1. Regulation of RhoA activity. Inactive RhoA-GDP binds to RhoGDI in the cytosol. For RhoA to be activated, GDF allows GEF to incorporate GTP to RhoA. Active RhoA-GTP is either located in the membrane or in the cytosol, where it binds to effector proteins such as ROCK. GAP activity accelerates RhoA inactivation by catalyzing hydrolysis of GTP to GDP and Pi. Effector proteins, which are activated by RhoA-GTP, then transmit the RhoA signals to downstream components, thus changing cellular functions.

Fig. 2. Regulation of RhoA transcription. Promoter of RhoA consists of several elements including the c-Myc and Max-binding region (-2kb), for HIF-1α/2α (-1.1 kb), NF-κB (-84 ~ -74 bp), STAT6 (-78 ~ -70bp) along with a CCAAT box (-68 ~ -64 bp). In particular, c-Myc forms a complex with Max, Miz1, Skp2 and p300 acetyltransferase. The RhoA mRNA regulation is also a target for several microRNAs including miR-31, miR-34a, miR-122, miR-133b, miR-146, miR-155, miR-185 and miR-340.

Fig. 3. Activation of transcription factors through RhoA and G/F-actin. Through an actin switch, the distribution of MAL in the cellular and nuclear compartments is regulated. MAL binds to G-actin, resulting it becoming sequestered in the cytosol; on the other hand, MAL cannot bind F-actin and can translocate to the nucleus. Nuclear MAL binds to SRF on SRE and activates SRF as a coactivator, leading to c-fos expression. On the other hand, when nuclear MAL binds to PPARγ, MAL acts as a co-repressor, leading to repression of PPARγ activity and consequently prevention of adipogenesis. YAP/TAZ binds to F-actin, which it can be replaced with AMOT, leading to its release and nuclear translocation. YAP/TAZ in the nucleus induces target genes to induce cell proliferation. Sox9 also binds to F-actin and it enters the nucleus when F-actin is disassembled by RhoA inactivation. In the nucleus, Sox9 induces genes related to chondrogenesis.


Fig. 4. IKKγ/NEMO is required for activation of NF-κB through RhoA GTPase. Oxidation of Cys16/20 and phosphorylation of Tyr42 of RhoA result in the dissociation of the RhoA-RhoGDI complex. Oxidized/phosphorylated RhoA can be activated by Vav2 GEF and then binds to IKKγ/NEMO, where RhoA directly activates IKKβ juxtaposed to RhoA on IKKγ/NEMO. IKKβ phosphorylates IκB, leading to IκB degradation and NF-κB activation. From this chain of events, ROS induces tumorigenesis through the signaling pathway.


References Arthur WT, Burridge K. 2001. RhoA inactivation by p190RhoGAP regulates cell spreading and migration by promoting membrane protrusion and polarity. Mol Biol Cell 12(9):2711-2720. Arthur WT, Ellerbroek SM, Der CJ, Burridge K, Wennerberg K. 2002. XPLN, a guanine nucleotide exchange factor for RhoA and RhoB, but not RhoC. J Biol Chem 277(45):42964-42972. Bijkerk R, de Bruin RG, van Solingen C, Duijs JM, Kobayashi K, van der Veer EP, ten Dijke P, Rabelink TJ, Goumans MJ, van Zonneveld AJ. 2012. MicroRNA-155 functions as a negative regulator of RhoA signaling in TGF-beta-induced endothelial to mesenchymal transition. Microrna 1(1):2-10. Bokoch GM, Zhao T. 2006. Regulation of the phagocyte NADPH oxidase by Rac GTPase. Antioxid Redox Signal 8(9-10):1533-1548. Bos JL, Rehmann H, Wittinghofer A. 2007. GEFs and GAPs: critical elements in the control of small G proteins. Cell 129(5):865-877. Bruning U, Cerone L, Neufeld Z, Fitzpatrick SF, Cheong A, Scholz CC, Simpson DA, Leonard MO, Tambuwala MM, Cummins EP, Taylor CT. 2011. MicroRNA-155 promotes resolution of hypoxia-inducible factor 1alpha activity during prolonged hypoxia. Mol Cell Biol 31(19):40874096. Buchsbaum RJ. 2007. Rho activation at a glance. J Cell Sci 120(Pt 7):1149-1152. Chalamalasetty RB, Hummer S, Nigg EA, Sillje HH. 2006. Influence of human Ect2 depletion and overexpression on cleavage furrow formation and abscission. J Cell Sci 119(Pt 14):30083019. Chan CH, Lee SW, Li CF, Wang J, Yang WL, Wu CY, Wu J, Nakayama KI, Kang HY, Huang HY, Hung MC, Pandolfi PP, Lin HK. 2010. Deciphering the transcriptional complex critical for RhoA gene expression and cancer metastasis. Nat Cell Biol 12(5):457-467. Chang F, Lemmon C, Lietha D, Eck M, Romer L. 2011. Tyrosine phosphorylation of Rac1: a role in regulation of cell spreading. PLoS One 6(12):e28587. Charron F, Tsimiklis G, Arcand M, Robitaille L, Liang Q, Molkentin JD, Meloche S, Nemer M. 2001. Tissue-specific GATA factors are transcriptional effectors of the small GTPase RhoA. Genes Dev 15(20):2702-2719. Chen SY, Huff SY, Lai CC, Der CJ, Powers S. 1994. Ras-15A protein shares highly similar dominant-negative biological properties with Ras-17N and forms a stable, guaninenucleotide resistant complex with CDC25 exchange factor. Oncogene 9(9):2691-2698. Danen EH, Sonneveld P, Sonnenberg A, Yamada KM. 2000. Dual stimulation of Ras/mitogenactivated protein kinase and RhoA by cell adhesion to fibronectin supports growth factorstimulated cell cycle progression. J Cell Biol 151(7):1413-1422. Davies CC, Chakraborty A, Cipriani F, Haigh K, Haigh JJ, Behrens A. 2010. Identification of a coactivator that links growth factor signalling to c-Jun/AP-1 activation. Nat Cell Biol 12(10):963972. Debidda M, Wang L, Zang H, Poli V, Zheng Y. 2005. A role of STAT3 in Rho GTPase-regulated cell migration and proliferation. J Biol Chem 280(17):17275-17285. DerMardirossian C, Rocklin G, Seo JY, Bokoch GM. 2006. Phosphorylation of RhoGDI by Src regulates Rho GTPase binding and cytosol-membrane cycling. Mol Biol Cell 17(11):47604768. DerMardirossian C, Schnelzer A, Bokoch GM. 2004. Phosphorylation of RhoGDI by Pak1 mediates dissociation of Rac GTPase. Mol Cell 15(1):117-127. Dubash AD, Guilluy C, Srougi MC, Boulter E, Burridge K, Garcia-Mata R. 2011. The small GTPase RhoA localizes to the nucleus and is activated by Net1 and DNA damage signals. PLoS One 6(2):e17380. Feng X, Degese MS, Iglesias-Bartolome R, Vaque JP, Molinolo AA, Rodrigues M, Zaidi MR, Ksander BR, Merlino G, Sodhi A, Chen Q, Gutkind JS. 2014. Hippo-independent activation of YAP by the GNAQ uveal melanoma oncogene through a trio-regulated rho GTPase signaling circuitry. Cancer Cell 25(6):831-845. Garcia-Mata R, Dubash AD, Sharek L, Carr HS, Frost JA, Burridge K. 2007. The nuclear RhoA This article is protected by copyright. All rights reserved

exchange factor Net1 interacts with proteins of the Dlg family, affects their localization, and influences their tumor suppressor activity. Mol Cell Biol 27(24):8683-8697. Gayard M, Guilluy C, Rousselle A, Viollet B, Henrion D, Pacaud P, Loirand G, Rolli-Derkinderen M. 2011. AMPK alpha 1-induced RhoA phosphorylation mediates vasoprotective effect of estradiol. Arterioscler Thromb Vasc Biol 31(11):2634-2642. Ghiaur G, Lee A, Bailey J, Cancelas JA, Zheng Y, Williams DA. 2006. Inhibition of RhoA GTPase activity enhances hematopoietic stem and progenitor cell proliferation and engraftment. Blood 108(6):2087-2094. Giannoni E, Buricchi F, Raugei G, Ramponi G, Chiarugi P. 2005. Intracellular reactive oxygen species activate Src tyrosine kinase during cell adhesion and anchorage-dependent cell growth. Mol Cell Biol 25(15):6391-6403. Gilkes DM, Xiang L, Lee SJ, Chaturvedi P, Hubbi ME, Wirtz D, Semenza GL. 2014. Hypoxiainducible factors mediate coordinated RhoA-ROCK1 expression and signaling in breast cancer cells. Proc Natl Acad Sci U S A 111(3):E384-393. Goto K, Chiba Y, Matsusue K, Hattori Y, Maitani Y, Sakai H, Kimura S, Misawa M. 2010. The proximal STAT6 and NF-kappaB sites are responsible for IL-13- and TNF-alpha-induced RhoA transcriptions in human bronchial smooth muscle cells. Pharmacol Res 61(5):466-472. Gu Y, Filippi MD, Cancelas JA, Siefring JE, Williams EP, Jasti AC, Harris CE, Lee AW, Prabhakar R, Atkinson SJ, Kwiatkowski DJ, Williams DA. 2003. Hematopoietic cell regulation by Rac1 and Rac2 guanosine triphosphatases. Science 302(5644):445-449. Haikala HM, Marques E, Turunen M, Klefstrom J. 2016. Myc requires RhoA/SRF to reprogram glutamine metabolism. Small GTPases:1-9. Hayashi M, Sakata M, Takeda T, Tahara M, Yamamoto T, Minekawa R, Isobe A, Tasaka K, Murata Y. 2005. Hypoxia up-regulates hypoxia-inducible factor-1alpha expression through RhoA activation in trophoblast cells. J Clin Endocrinol Metab 90(3):1712-1719. Heasman SJ, Ridley AJ. 2008. Mammalian Rho GTPases: new insights into their functions from in vivo studies. Nat Rev Mol Cell Biol 9(9):690-701. Heo J, Raines KW, Mocanu V, Campbell SL. 2006. Redox regulation of RhoA. Biochemistry 45(48):14481-14489. Hill CS, Wynne J, Treisman R. 1995. The Rho family GTPases RhoA, Rac1, and CDC42Hs regulate transcriptional activation by SRF. Cell 81(7):1159-1170. Hobbs GA, Zhou B, Cox AD, Campbell SL. 2014. Rho GTPases, oxidation, and cell redox control. Small GTPases 5:e28579. Hsu SH, Delgado ER, Otero PA, Teng KY, Kutay H, Meehan KM, Moroney JB, Monga JK, Hand NJ, Friedman JR, Ghoshal K, Duncan AW. 2016. MicroRNA-122 regulates polyploidization in the murine liver. Hepatology 64(2):599-615. Jaffe AB, Hall A. 2005. Rho GTPases: biochemistry and biology. Annu Rev Cell Dev Biol 21:247-269. Jeon CY, Kim HJ, Lee JY, Kim JB, Kim SC, Park JB. 2010a. p190RhoGAP and Rap-dependent RhoGAP (ARAP3) inactivate RhoA in response to nerve growth factor leading to neurite outgrowth from PC12 cells. Exp Mol Med 42(5):335-344. Jeon CY, Kim HJ, Morii H, Mori N, Settleman J, Lee JY, Kim J, Kim SC, Park JB. 2010b. Neurite outgrowth from PC12 cells by basic fibroblast growth factor (bFGF) is mediated by RhoA inactivation through p190RhoGAP and ARAP3. J Cell Physiol 224(3):786-794. Jeon CY, Moon MY, Kim JH, Kim HJ, Kim JG, Li Y, Jin JK, Kim PH, Kim HC, Meier KE, Kim YS, Park JB. 2012. Control of neurite outgrowth by RhoA inactivation. J Neurochem 120(5):684-698. Jian Q, An Q, Zhu D, Hui K, Liu Y, Chi S, Li C. 2014. MicroRNA 340 is involved in UVB-induced dendrite formation through the regulation of RhoA expression in melanocytes. Mol Cell Biol 34(18):3407-3420. Jun JI, Lau LF. 2011. Taking aim at the extracellular matrix: CCN proteins as emerging therapeutic targets. Nat Rev Drug Discov 10(12):945-963. Kakiuchi M, Nishizawa T, Ueda H, Gotoh K, Tanaka A, Hayashi A, Yamamoto S, Tatsuno K, Katoh H, Watanabe Y, Ichimura T, Ushiku T, Funahashi S, Tateishi K, Wada I, Shimizu N, Nomura S, Koike K, Seto Y, Fukayama M, Aburatani H, Ishikawa S. 2014. Recurrent gain-of-function mutations of RHOA in diffuse-type gastric carcinoma. Nat Genet 46(6):583-587. Kato M, Blanton R, Wang GR, Judson TJ, Abe Y, Myoishi M, Karas RH, Mendelsohn ME. 2012. This article is protected by copyright. All rights reserved

Direct binding and regulation of RhoA protein by cyclic GMP-dependent protein kinase Ialpha. J Biol Chem 287(49):41342-41351. Kim D, Song J, Kim S, Park HM, Chun CH, Sonn J, Jin EJ. 2012. MicroRNA-34a modulates cytoskeletal dynamics through regulating RhoA/Rac1 cross-talk in chondroblasts. J Biol Chem 287(15):12501-12509. Kim HJ, Choi HS, Park JH, Kim MJ, Lee HG, Petersen RB, Kim YS, Park JB, Choi EK. 2017a. Regulation of RhoA activity by the cellular prion protein. Cell Death Dis 8(3):e2668. Kim HJ, Kim JG, Moon MY, Park SH, Park JB. 2014. IkappaB kinase gamma/nuclear factor-kappaBessential modulator (IKKgamma/NEMO) facilitates RhoA GTPase activation, which, in turn, activates Rho-associated KINASE (ROCK) to phosphorylate IKKbeta in response to transforming growth factor (TGF)-beta1. J Biol Chem 289(3):1429-1440. Kim JG, Choi KC, Hong CW, Park HS, Choi EK, Kim YS, Park JB. 2017b. Tyr42 phosphorylation of RhoA GTPase promotes tumorigenesis through nuclear factor (NF)-kappaB. Free Radic Biol Med 112:69-83. Kim JG, Kim MJ, Choi WJ, Moon MY, Kim HJ, Lee JY, Kim J, Kim SC, Kang SG, Seo GY, Kim PH, Park JB. 2017c. Wnt3A Induces GSK-3beta Phosphorylation and beta-Catenin Accumulation Through RhoA/ROCK. J Cell Physiol 232(5):1104-1113. Kim JG, Kwon HJ, Wu G, Park Y, Lee JY, Kim J, Kim SC, Choe M, Kang SG, Seo GY, Kim PH, Park JB. 2017d. RhoA GTPase oxidation stimulates cell proliferation via nuclear factor-kappaB activation. Free Radic Biol Med 103:57-68. Kim JS, Diebold BA, Kim JI, Kim J, Lee JY, Park JB. 2004. Rho is involved in superoxide formation during phagocytosis of opsonized zymosans. J Biol Chem 279(20):21589-21597. Kim JS, Kim JG, Moon MY, Jeon CY, Won HY, Kim HJ, Jeon YJ, Seo JY, Kim JI, Kim J, Lee JY, Kim PH, Park JB. 2006. Transforming growth factor-beta1 regulates macrophage migration via RhoA. Blood 108(6):1821-1829. Kong W, Yang H, He L, Zhao JJ, Coppola D, Dalton WS, Cheng JQ. 2008. MicroRNA-155 is regulated by the transforming growth factor beta/Smad pathway and contributes to epithelial cell plasticity by targeting RhoA. Mol Cell Biol 28(22):6773-6784. Korourian A, Roudi R, Shariftabrizi A, Madjd Z. 2017. MicroRNA-31 inhibits RhoA-mediated tumor invasion and chemotherapy resistance in MKN-45 gastric adenocarcinoma cells. Exp Biol Med (Maywood):1535370217728460. Krugmann S, Williams R, Stephens L, Hawkins PT. 2004. ARAP3 is a PI3K- and rap-regulated GAP for RhoA. Curr Biol 14(15):1380-1384. Kuhlmann N, Wroblowski S, Knyphausen P, de Boor S, Brenig J, Zienert AY, Meyer-Teschendorf K, Praefcke GJ, Nolte H, Kruger M, Schacherl M, Baumann U, James LC, Chin JW, Lammers M. 2016a. Structural and Mechanistic Insights into the Regulation of the Fundamental Rho Regulator RhoGDIalpha by Lysine Acetylation. J Biol Chem 291(11):5484-5499. Kuhlmann N, Wroblowski S, Scislowski L, Lammers M. 2016b. RhoGDIalpha Acetylation at K127 and K141 Affects Binding toward Nonprenylated RhoA. Biochemistry 55(2):304-312. Kumar D, Lassar AB. 2009. The transcriptional activity of Sox9 in chondrocytes is regulated by RhoA signaling and actin polymerization. Mol Cell Biol 29(15):4262-4273. Lang P, Gesbert F, Delespine-Carmagnat M, Stancou R, Pouchelet M, Bertoglio J. 1996. Protein kinase A phosphorylation of RhoA mediates the morphological and functional effects of cyclic AMP in cytotoxic lymphocytes. EMBO J 15(3):510-519. Li Y, Kim JG, Kim HJ, Moon MY, Lee JY, Kim J, Kim SC, Song DK, Kim YS, Park JB. 2012. Small GTPases Rap1 and RhoA regulate superoxide formation by Rac1 GTPases activation during the phagocytosis of IgG-opsonized zymosans in macrophages. Free Radic Biol Med 52(9):1796-1805. Liu M, Lang N, Chen X, Tang Q, Liu S, Huang J, Zheng Y, Bi F. 2011. miR-185 targets RhoA and Cdc42 expression and inhibits the proliferation potential of human colorectal cells. Cancer Lett 301(2):151-160. Liu Q, Wang W, Yang X, Zhao D, Li F, Wang H. 2016. MicroRNA-146a inhibits cell migration and invasion by targeting RhoA in breast cancer. Oncol Rep 36(1):189-196. Loirand G, Guerin P, Pacaud P. 2006. Rho kinases in cardiovascular physiology and pathophysiology. Circ Res 98(3):322-334. This article is protected by copyright. All rights reserved

Lu XC, Zheng JY, Tang LJ, Huang BS, Li K, Tao Y, Yu W, Zhu RL, Li S, Li LX. 2015. MiR-133b Promotes neurite outgrowth by targeting RhoA expression. Cell Physiol Biochem 35(1):246258. Manso R, Sanchez-Beato M, Monsalvo S, Gomez S, Cereceda L, Llamas P, Rojo F, Mollejo M, Menarguez J, Alves J, Garcia-Cosio M, Piris MA, Rodriguez-Pinilla SM. 2014. The RHOA G17V gene mutation occurs frequently in peripheral T-cell lymphoma and is associated with a characteristic molecular signature. Blood 123(18):2893-2894. Marinissen MJ, Chiariello M, Tanos T, Bernard O, Narumiya S, Gutkind JS. 2004. The small GTPbinding protein RhoA regulates c-jun by a ROCK-JNK signaling axis. Mol Cell 14(1):29-41. McManus EJ, Sakamoto K, Armit LJ, Ronaldson L, Shpiro N, Marquez R, Alessi DR. 2005. Role that phosphorylation of GSK3 plays in insulin and Wnt signalling defined by knockin analysis. EMBO J 24(8):1571-1583. Michael SK, Surks HK, Wang Y, Zhu Y, Blanton R, Jamnongjit M, Aronovitz M, Baur W, Ohtani K, Wilkerson MK, Bonev AD, Nelson MT, Karas RH, Mendelsohn ME. 2008. High blood pressure arising from a defect in vascular function. Proc Natl Acad Sci U S A 105(18):67026707. Miralles F, Posern G, Zaromytidou AI, Treisman R. 2003. Actin dynamics control SRF activity by regulation of its coactivator MAL. Cell 113(3):329-342. Moon MY, Kim HJ, Li Y, Kim JG, Jeon YJ, Won HY, Kim JS, Kwon HY, Choi IG, Ro E, Joe EH, Choe M, Kwon HJ, Kim HC, Kim YS, Park JB. 2013. Involvement of small GTPase RhoA in the regulation of superoxide production in BV2 cells in response to fibrillar Abeta peptides. Cell Signal 25(9):1861-1869. Moon SY, Zheng Y. 2003. Rho GTPase-activating proteins in cell regulation. Trends Cell Biol 13(1):13-22. Moriki N, Ito M, Seko T, Kureishi Y, Okamoto R, Nakakuki T, Kongo M, Isaka N, Kaibuchi K, Nakano T. 2004. RhoA activation in vascular smooth muscle cells from stroke-prone spontaneously hypertensive rats. Hypertens Res 27(4):263-270. Moroishi T, Hansen CG, Guan KL. 2015. The emerging roles of YAP and TAZ in cancer. Nat Rev Cancer 15(2):73-79. Nimnual AS, Taylor LJ, Bar-Sagi D. 2003. Redox-dependent downregulation of Rho by Rac. Nat Cell Biol 5(3):236-241. Nobusue H, Onishi N, Shimizu T, Sugihara E, Oki Y, Sumikawa Y, Chiyoda T, Akashi K, Saya H, Kano K. 2014. Regulation of MKL1 via actin cytoskeleton dynamics drives adipocyte differentiation. Nat Commun 5:3368. Nomikou E, Stournaras C, Kardassis D. 2017. Functional analysis of the promoters of the small GTPases RhoA and RhoB in embryonic stem cells. Biochem Biophys Res Commun 491(3):754-759. Olson MF. 2016. Rho GTPases, their post-translational modifications, disease-associated mutations and pharmacological inhibitors. Small GTPases:1-13. Ozturk E, Hobiger S, Despot-Slade E, Pichler M, Zenobi-Wong M. 2017. Hypoxia regulates RhoA and Wnt/beta-catenin signaling in a context-dependent way to control re-differentiation of chondrocytes. Sci Rep 7(1):9032. Palomero T, Couronne L, Khiabanian H, Kim MY, Ambesi-Impiombato A, Perez-Garcia A, Carpenter Z, Abate F, Allegretta M, Haydu JE, Jiang X, Lossos IS, Nicolas C, Balbin M, Bastard C, Bhagat G, Piris MA, Campo E, Bernard OA, Rabadan R, Ferrando AA. 2014. Recurrent mutations in epigenetic regulators, RHOA and FYN kinase in peripheral T cell lymphomas. Nat Genet 46(2):166-170. Pietri M, Dakowski C, Hannaoui S, Alleaume-Butaux A, Hernandez-Rapp J, Ragagnin A, MouilletRichard S, Haik S, Bailly Y, Peyrin JM, Launay JM, Kellermann O, Schneider B. 2013. PDK1 decreases TACE-mediated alpha-secretase activity and promotes disease progression in prion and Alzheimer's diseases. Nat Med 19(9):1124-1131. Porter AP, Papaioannou A, Malliri A. 2016. Deregulation of Rho GTPases in cancer. Small GTPases 7(3):123-138. Qiao Y, Chen J, Lim YB, Finch-Edmondson ML, Seshachalam VP, Qin L, Jiang T, Low BC, Singh H, Lim CT, Sudol M. 2017. YAP Regulates Actin Dynamics through ARHGAP29 and Promotes This article is protected by copyright. All rights reserved

Metastasis. Cell Rep 19(8):1495-1502. Rodrigues P, Macaya I, Bazzocco S, Mazzolini R, Andretta E, Dopeso H, Mateo-Lozano S, Bilic J, Carton-Garcia F, Nieto R, Suarez-Lopez L, Afonso E, Landolfi S, Hernandez-Losa J, Kobayashi K, Ramon y Cajal S, Tabernero J, Tebbutt NC, Mariadason JM, Schwartz S, Jr., Arango D. 2014. RHOA inactivation enhances Wnt signalling and promotes colorectal cancer. Nat Commun 5:5458. Sahai E, Alberts AS, Treisman R. 1998. RhoA effector mutants reveal distinct effector pathways for cytoskeletal reorganization, SRF activation and transformation. EMBO J 17(5):1350-1361. Sakata-Yanagimoto M, Enami T, Yoshida K, Shiraishi Y, Ishii R, Miyake Y, Muto H, Tsuyama N, SatoOtsubo A, Okuno Y, Sakata S, Kamada Y, Nakamoto-Matsubara R, Tran NB, Izutsu K, Sato Y, Ohta Y, Furuta J, Shimizu S, Komeno T, Sato Y, Ito T, Noguchi M, Noguchi E, Sanada M, Chiba K, Tanaka H, Suzukawa K, Nanmoku T, Hasegawa Y, Nureki O, Miyano S, Nakamura N, Takeuchi K, Ogawa S, Chiba S. 2014. Somatic RHOA mutation in angioimmunoblastic T cell lymphoma. Nat Genet 46(2):171-175. Sauzeau V, Le Jeune H, Cario-Toumaniantz C, Smolenski A, Lohmann SM, Bertoglio J, Chardin P, Pacaud P, Loirand G. 2000. Cyclic GMP-dependent protein kinase signaling pathway inhibits RhoA-induced Ca2+ sensitization of contraction in vascular smooth muscle. J Biol Chem 275(28):21722-21729. Sauzeau V, Rolli-Derkinderen M, Marionneau C, Loirand G, Pacaud P. 2003. RhoA expression is controlled by nitric oxide through cGMP-dependent protein kinase activation. J Biol Chem 278(11):9472-9480. Sawada N, Itoh H, Miyashita K, Tsujimoto H, Sone M, Yamahara K, Arany ZP, Hofmann F, Nakao K. 2009. Cyclic GMP kinase and RhoA Ser188 phosphorylation integrate pro- and antifibrotic signals in blood vessels. Mol Cell Biol 29(22):6018-6032. Schmidt A, Hall A. 2002. The Rho exchange factor Net1 is regulated by nuclear sequestration. J Biol Chem 277(17):14581-14588. Song EH, Oh W, Ulu A, Carr HS, Zuo Y, Frost JA. 2015. Acetylation of the RhoA GEF Net1A controls its subcellular localization and activity. J Cell Sci 128(5):913-922. Sorrentino G, Ruggeri N, Specchia V, Cordenonsi M, Mano M, Dupont S, Manfrin A, Ingallina E, Sommaggio R, Piazza S, Rosato A, Piccolo S, Del Sal G. 2014. Metabolic control of YAP and TAZ by the mevalonate pathway. Nat Cell Biol 16(4):357-366. Staus DP, Weise-Cross L, Mangum KD, Medlin MD, Mangiante L, Taylor JM, Mack CP. 2014. Nuclear RhoA signaling regulates MRTF-dependent SMC-specific transcription. Am J Physiol Heart Circ Physiol 307(3):H379-390. Takahashi K, Sasaki T, Mammoto A, Takaishi K, Kameyama T, Tsukita S, Takai Y. 1997. Direct interaction of the Rho GDP dissociation inhibitor with ezrin/radixin/moesin initiates the activation of the Rho small G protein. J Biol Chem 272(37):23371-23375. Takata K, Morishige K, Takahashi T, Hashimoto K, Tsutsumi S, Yin L, Ohta T, Kawagoe J, Takahashi K, Kurachi H. 2008. Fasudil-induced hypoxia-inducible factor-1alpha degradation disrupts a hypoxia-driven vascular endothelial growth factor autocrine mechanism in endothelial cells. Mol Cancer Ther 7(6):1551-1561. Tanaka T, Nishimura D, Wu RC, Amano M, Iso T, Kedes L, Nishida H, Kaibuchi K, Hamamori Y. 2006. Nuclear Rho kinase, ROCK2, targets p300 acetyltransferase. J Biol Chem 281(22):15320-15329. Tong J, Li L, Ballermann B, Wang Z. 2013. Phosphorylation of Rac1 T108 by extracellular signalregulated kinase in response to epidermal growth factor: a novel mechanism to regulate Rac1 function. Mol Cell Biol 33(22):4538-4551. Tong J, Li L, Ballermann B, Wang Z. 2016. Phosphorylation and Activation of RhoA by ERK in Response to Epidermal Growth Factor Stimulation. PLoS One 11(1):e0147103. Tsai MS, Bogart DF, Castaneda JM, Li P, Lupu R. 2002. Cyr61 promotes breast tumorigenesis and cancer progression. Oncogene 21(53):8178-8185. Tu S, Wu WJ, Wang J, Cerione RA. 2003. Epidermal growth factor-dependent regulation of Cdc42 is mediated by the Src tyrosine kinase. J Biol Chem 278(49):49293-49300. Turcotte S, Desrosiers RR, Beliveau R. 2003. HIF-1alpha mRNA and protein upregulation involves Rho GTPase expression during hypoxia in renal cell carcinoma. J Cell Sci 116(Pt 11):2247This article is protected by copyright. All rights reserved

2260. Uezu A, Okada H, Murakoshi H, del Vescovo CD, Yasuda R, Diviani D, Soderling SH. 2012. Modified SH2 domain to phototrap and identify phosphotyrosine proteins from subcellular sites within cells. Proc Natl Acad Sci U S A 109(43):E2929-2938. Voloshanenko O, Erdmann G, Dubash TD, Augustin I, Metzig M, Moffa G, Hundsrucker C, Kerr G, Sandmann T, Anchang B, Demir K, Boehm C, Leible S, Ball CR, Glimm H, Spang R, Boutros M. 2013. Wnt secretion is required to maintain high levels of Wnt activity in colon cancer cells. Nat Commun 4:2610. Waheed F, Speight P, Dan Q, Garcia-Mata R, Szaszi K. 2012. Affinity precipitation of active RhoGEFs using a GST-tagged mutant Rho protein (GST-RhoA(G17A)) from epithelial cell lysates. J Vis Exp(61). Walsh CT, Radeff-Huang J, Matteo R, Hsiao A, Subramaniam S, Stupack D, Brown JH. 2008. Thrombin receptor and RhoA mediate cell proliferation through integrins and cysteine-rich protein 61. FASEB J 22(11):4011-4021. Wang JB, Erickson JW, Fuji R, Ramachandran S, Gao P, Dinavahi R, Wilson KF, Ambrosio AL, Dias SM, Dang CV, Cerione RA. 2010. Targeting mitochondrial glutaminase activity inhibits oncogenic transformation. Cancer Cell 18(3):207-219. Wang SC, Lin XL, Li J, Zhang TT, Wang HY, Shi JW, Yang S, Zhao WT, Xie RY, Wei F, Qin YJ, Chen L, Yang J, Yao KT, Xiao D. 2014. MicroRNA-122 triggers mesenchymal-epithelial transition and suppresses hepatocellular carcinoma cell motility and invasion by targeting RhoA. PLoS One 9(7):e101330. Yamashita T, Tohyama M. 2003. The p75 receptor acts as a displacement factor that releases Rho from Rho-GDI. Nat Neurosci 6(5):461-467. Yoo HY, Sung MK, Lee SH, Kim S, Lee H, Park S, Kim SC, Lee B, Rho K, Lee JE, Cho KH, Kim W, Ju H, Kim J, Kim SJ, Kim WS, Lee S, Ko YH. 2014. A recurrent inactivating mutation in RHOA GTPase in angioimmunoblastic T cell lymphoma. Nat Genet 46(4):371-375. Yu FX, Luo J, Mo JS, Liu G, Kim YC, Meng Z, Zhao L, Peyman G, Ouyang H, Jiang W, Zhao J, Chen X, Zhang L, Wang CY, Bastian BC, Zhang K, Guan KL. 2014. Mutant Gq/11 promote uveal melanoma tumorigenesis by activating YAP. Cancer Cell 25(6):822-830. Zhang C, Zhang S, Zhang Z, He J, Xu Y, Liu S. 2014. ROCK has a crucial role in regulating prostate tumor growth through interaction with c-Myc. Oncogene 33(49):5582-5591.


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