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Cardiovascular Research 2006 71(2):236-246; doi:10.1016/j.cardiores.2006.05.003
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Copyright © 2006, European Society of Cardiology

Reactive oxygen species in vascular endothelial cell motility. Roles of NAD(P)H oxidase and Rac1

Leni Moldovana, Karthikeyan Mythreyeb, Pascal J. Goldschmidt-Clermontc and Lisa L. Satterwhiteb,*

aDavis Heart and Lung Research Institute, The Ohio State University, Columbus, OH 43210, United States
bDepartment of Medicine, Duke University Medical Center, Bell Research Building, Room 345, Durham, NC 27710, United States
cLeonard M. Miller School of Medicine at the University of Miami, P. O. Box 016099, Miami, FL 33101, United States

* Corresponding author. Tel.: +1 919 681 0381. Email address: lisa.satterwhite{at}duke.edu

Received 28 January 2006; revised 29 April 2006; accepted 3 May 2006


   Abstract
Top
 Abstract
1. Introduction
 2. The non-phagocytic NAD(P)H...
 3. ROS and endothelial...
 4. Direct interaction between...
 5. ROS and regulators...
 6. Conclusions and future...
 Appendix A. Supplementary data
 References
 
Reactive oxygen species (ROS) are acknowledged generally to be multi-faceted regulators of cellular functions that trigger various pathological states when present chronically or transiently at non-physiologically high levels. Here we focus on the physiological involvement of ROS in cellular motility, with special emphasis on endothelial cells (EC). An important source of ROS within EC is the non-phagocytic NAD(P)H oxidase, and the small GTPase Rac1 plays a central role in activating this complex. Rac1 is one of the three Rho-family molecules (Rac, Rho and Cdc42) involved in the control of the actin cytoskeleton in response to various signals. In this review we examine the evidence linking ROS production, Rac1 activation and actin organization to EC motility, considering mechanisms for direct interaction of ROS and actin and the effects of ROS on proteins that regulate the actin cytoskeleton. The accumulated evidence suggests that ROS are important regulators of the actin cytoskeletal dynamics and cellular motility, and more in-depth studies are needed to understand the underlying mechanisms.

KEYWORDS Redox signalling; Endothelial function; NADPH oxidase; Oxygen radicals; Cytoskeleton


   1. Introduction
Top
 Abstract
 1. Introduction
2. The non-phagocytic NAD(P)H...
 3. ROS and endothelial...
 4. Direct interaction between...
 5. ROS and regulators...
 6. Conclusions and future...
 Appendix A. Supplementary data
 References
 
Chronic exposure to reactive oxygen species (ROS) and the resulting oxidative stress are considered central components of the onset of atheroma formation [1], diabetic complications [2,3], cell transformation and proliferation in cancer [4], and degenerative disorders and senescence [5]. Oxidative stress, viewed as the overproduction of ROS, the failure of the antioxidant defense of the organism, or both, plays a major role in human pathology [6]. However, in healthy cells ROS are not just a by-product of oxygen metabolism, but have physiologically specific roles, serving to regulate key signaling pathways [7,8]. Hydrogen peroxide (H2O2), superoxide (O2Formula), and in some instances even the hydroxyl radical (HO•) appear to be required for cellular processes as fundamental as proliferation [9], intracellular signaling [10], and gene expression [11,12]. In this review, we first describe the mechanism by which ROS are generated within non-phagocytic cells by the NAD(P)H oxidase complex and the role of the small GTPase Rac1 for its activation. Then, we turn to how the actin cytoskeleton responds to ROS with enhanced motility and how inhibitors of Rac1 influence both ROS levels and actin-based motility. Finally, we examine the possible molecular mechanisms by which ROS may control functional dynamics of the actin cytoskeleton.


   2. The non-phagocytic NAD(P)H oxidases
Top
 Abstract
 1. Introduction
 2. The non-phagocytic NAD(P)H...
3. ROS and endothelial...
 4. Direct interaction between...
 5. ROS and regulators...
 6. Conclusions and future...
 Appendix A. Supplementary data
 References
 
NADPH oxidases are a group of multimeric enzymes whose activity results in the production of O2Formula. The best characterized is the phagocytic enzyme that mediates the oxidative burst of activated neutrophils [13]. This is a multi-component transmembrane complex, with flavocytochrome b558 in its catalytic core, which consists of a large glycoprotein gp91phox/NOX2 (known as "beta subunit") and a small protein p22phox ("alpha subunit") in complex with Rap1A [14]. Rap1A is a substrate for protein kinase A and its interaction with NADPH oxidase is abrogated upon phosphorylation [15]. The other components of the phagocytic oxidase are water-soluble proteins of cytosolic origin, namely p67phox, p47phox, p40phox and the small GTP-binding protein Rac1 in monocytes/macrophages [16] or Rac2 in neutrophils [17]. Rac proteins are associated with Rho GTPase dissociation inhibitors (RhoGDI), known to prevent the exchange of GTP for GDP [18]. Upon stimulation, cytosolic components assemble with the membrane-bound flavocytochrome b558, become activated and generate O2Formula [19]. The central role of Rac1 in activation of the phagocytic oxidase complex consists in its interaction with, and activation of, p67phox and the tethering of the complex of cytosolic proteins to the membrane [20,21].

In non-phagocytic cells, homologues of the NADPH oxidase (Fig. 1) were found in vascular endothelial cells (EC) [22–28] and smooth muscle cells [29] (for a review of vascular NAD(P)H oxidases, see [30]), as well as in other normal [31–33] or transformed cells such as colon cancer [34] or melanoma [35]. Several homologues of the gp91phox have been described and designated by the common name NOX (for NADPH oxidase): NOX1, mainly found in colon and the vascular wall, NOX2, which is the phagocytic gp91phox, NOX3, found in fetal kidney, NOX4, with widespread occurrence, and NOX5, found in spleen, sperm, mammary glands and cerebrum [36]. The cytosolic components of the non-phagocytic NAD(P)H oxidase include homologues of p47phox (or NOXO1, NOX organizing protein 1) and p67phox (or NOXA1, NOX activating protein 1) [37–39]. As opposed to the oxidative burst of phagocytosis, the amount of O2Formula produced by the non-phagocytic enzyme is much less [40] and a large portion of the ROS output occurs intracellularly rather than extracellularly. Another difference from the phagocytic oxidase is that p47phox contains an auto inhibitory region, which is absent in NOXO1 [37]. As a consequence, NOXO1 is thought to be constitutively active, and indeed in non-phagocytic cells O2Formula production is present at reduced levels even in non-stimulated cells [30]. Moreover, the oxidase components exist in preassembled intracellular complexes of gp91phox, p22phox, p47phox, p67phox and p40phox that associate with the cytoskeleton [27] and are rapidly brought together and modulated by a variety of physiological, pathological and mechanical stimuli [41]. Another difference is that while the phagocytic oxidase reduces molecular oxygen to O2Formula using NADPH as the electron donor, the non-phagocytic enzyme uses either NADH or NADPH as electron donor, hence the term NAD(P)H oxidase.


Figure 1
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  		Fig. 1 NAD(P)H oxidase of non-phagocytic cells. Upon stimulation by signaling molecules or mechanical factors, Rac1 dissociates from its inhibitor and is activated by releasing GDP and binding GTP which exposes the isoprenyl moiety. Consequently, it brings together the cytosolic components of the NAD(P)H oxidase NOXO1 and NOXA1 (homologues of the phagocytic p47phox and p67phox, respectively). NOXA1 binds Rac-GTP via the N-terminal domain composed of four TPR regions, and NOXO1 in a head-to-tail interaction [149]. The complex is recruited to the plasma membrane (PM), where Rac1 is anchored through the isoprenyl moiety, NOXO1 interacts via its SH3 domain with the PRR domain of the C-terminal cytoplasmic tail of p22phox, and NOXA1 activates the NOX1-p21phox complex. Most O2Formula is intracellular in non-phagocytes. The key signaling molecule is thought to be H2O2, generated through the catalytic activity of Cu, Zn-superoxide dismutase (SOD).

 
As is the case with the phagocytic enzyme, activation of non-phagocytic NAD(P)H oxidase is also dependent on Rac1 interaction with p67phox/NOXA1 [42] (Fig. 1), and ROS production is inhibited by a Rac1 dominant negative allele [43]. Furthermore, enhanced activation and anchoring of Rac1 to the plasma membrane appears to underlie ROS stimulation in response to statin withdrawal [44]. Most importantly, the O2Formula produced by non-phagocytic cells constitutes a key signal in the control of a variety of cellular processes, such as cell growth and motility, angiogenesis, immune function, hypoxic response, and oxidative modification of extracellular matrix proteins (for a review also see Ref. [36]).

Within the redox network, NAD(P)H oxidases are situated downstream to many important modulators of EC physiology, such as vascular endothelial growth factor (VEGF) [45], angiotensin II [46,47], TGFβ [48], endothelin-1 [49] and mechanical stimuli [50–52]; the induction pathways have been surveyed in good detail in several instances. However, the events downstream of enzyme activation and ROS formation are much less understood, one reason being that few direct targets of O2Formula and of its metabolite H2O2, which is considered to be the actual effector of most ROS-mediated processes, are known. Moreover, many of the components of the NAD(P)H oxidase have various other intracellular roles than the generation of O2Formula. For instance, p47phox has a role in the activation of RelA transcription factor in EC [53] and also interacts with TRAF4, being involved in TNF{alpha}-induced oxidative activation of c-Jun N-terminal kinase [54]. In another report, p47phox was found to co-localize with WAVE1, Rac1 and PAK1 in EC membrane ruffles induced by VEGF, and the authors suggested that NAD(P)H oxidase-derived oxidants appear to mediate ruffle formation, in addition to JNK activation [55]. This finding is consistent with our work showing that oxidants-induced fluorescence was highest within membrane ruffles of actively migrating EC ([56] and Fig. 2). In vascular smooth muscle cells, the NAD(P)H oxidase subunit p47phox is involved in redox-mediated control of gene expression [57], while in melanoma cells ROS produced by an NAD(P)H oxidase were shown to act in an autocrine manner as signaling molecules in transcription and growth regulation [58].


Figure 2
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  		Fig. 2 In an in vitro wound model, ruffles are filled with ROS. DIC image of a cell at the wound margin shows high ruffling activity (a). CM-DCF-DA fluorescence was detected within these ruffles (b, arrowheads). The image was taken from a time-lapse sequence (supplementary material) (from [56], with permission).

 
In phagocytes, a complex network of interactions between the cytosolic phox proteins and cytoskeletal proteins has been gradually uncovered: p40phox and p67phox bind coronin and accumulate around the phagocytic vacuole, possibly playing a regulatory role in the reorganization of the cytoskeleton that accompanies superoxide generation [59]; p47phox binds actin through its c-terminal region [60]; p47phox and p40phox bind moesin through their pleckstrin (PX) domains in a phosphoinositide-dependent manner [61]; Rac1, p47phox and p67phox interact with the Ca2+-binding protein S100A8/A9, which acts as a scaffold protein for the cytosolic phox proteins [62]. All the above-mentioned interactions control the activation of the phagocytic NADPH oxidase, but comparable associations of NAD(P)H oxidase components with the cytoskeleton were found in EC as well [27].


   3. ROS and endothelial cell motility
Top
 Abstract
 1. Introduction
 2. The non-phagocytic NAD(P)H...
 3. ROS and endothelial...
4. Direct interaction between...
 5. ROS and regulators...
 6. Conclusions and future...
 Appendix A. Supplementary data
 References
 
Actin-based cellular motility, essential for animal cells, depends on rapid reorganization of the actin network in response to numerous signals, such as growth factors [63], integrin engagement by their ligands [64] or signals for chemotaxis [65]. Actin network remodeling, in turn, modulates cell shape and motility, cell–cell contacts and cell contact with adhesion molecules in the substratum. ROS have emerged as key regulators of these processes. A first suggestion that O2Formula was involved in actin regulation in vivo came from the observation that in post-hypoxic EC, a condition known to induce xanthine oxidase-mediated production of O2Formula [66], the pool of filamentous actin increased, and this effect was reversed by over-expression of Cu, Zn-superoxide dismutase (SOD) [67]. Furthermore, the membrane ruffling and increase in F-actin induced by the over-expression of the constitutively activated form of Rac1, RacV12 in EC, was observed to be concomitant with increased ROS generation, and were counteracted by over-expression of SOD and by antioxidants. This again suggests that in EC O2Formula is a mediator which acts downstream of Rac1 in the pathway of actin cytoskeleton remodeling [68].

These findings suggested that NAD(P)H oxidase activation and the ensuing O2Formula production could regulate cell migration. We tested this hypothesis in an in vitro EC wounding model [56]. We and others [69] have found that migrating cells at the wound margin produced enhanced amounts of ROS, which were often localized within membrane ruffles (Fig. 2). The process coincided with increased incorporation of monomeric actin into fibers. Furthermore, preventing the formation of O2Formula with the NAD(P)H oxidase inhibitor diphenylene iodonium (DPI), or scavenging O2Formula with the SOD mimetic manganese (III) tetrakis(1-methyl-4-pyridyl)porphyrin (MnTMPyP), consistently reduced both the speed and directionality of EC migration into the wound [56]. This finding might be explained by recent reports that Rho GTPases, together with phosphatidylinositol 3-kinase, through a polarized distribution, activate signaling cascades to the cytoskeleton–both actin and microtubules–that lead to polarized cell morphology and directional cell migration [70]. Thus, Rac1 and Cdc42 recruit the IQGAP1/APC complex, where IQGAP1 links APC to actin filaments for cell polarization and directional migration [71]. IQGAP1 and Rac1, in addition to Cdc42 and APC, are necessary to reorient the microtubule organizing center (MTOC) in migrating fibroblasts [71]. Similarly, shear stress activates Rac1 in a polarized manner and its activity has to be spatially restricted in order for cells to align and to migrate in the direction of flow [72].

The Rho family of small GTPases are key regulators of actin dynamics, together with the related proteins Rho and Cdc42, coordinating formation of stress fibers, focal adhesions, lamelipodia and filopodia and thus regulating overall cellular movement [73–76]. In addition, the same molecules also control polymerization dynamics of the microtubule cytoskeleton [77–79]. In particular Rac1 is an extremely versatile molecule that interacts specifically with numerous upstream regulators and downstream effectors [42] (Fig. 3). Rac1 regulates at least four distinct effector systems via multiple domains carrying specific functions, while individual domains interact with more than one effector, thus generating complex outcomes [80–85]. However, although both roles of Rac in cytoskeletal reorganization and NAD(P)H oxidase activation have been known for more than a decade and intensely studied, the two processes were regarded until recently to be independent [7].


Figure 3
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  		Fig. 3 The Rac1 molecule, interacting partners and pathways. Rac proteins interact specifically with many modulators and effectors, thereby controlling various pathways. The molecular regions/residues involved in these interactions are known in some cases (circled regions). Some of the important downstream effectors of Rac are PAK [150], arfaptin/POR [151], IQGAP [152,153], WASP and WAVE [154,155]. Three families of modulator proteins are known, each containing several members, with different consequences upon interaction with Rac proteins: GTPase-activating proteins (GAP), guanine nucleotide dissociation inhibitors (GDI) and guanine nucleotide exchange factors (GEF) [156]. The model was built with DeepVew/Swiss-Pdb Viewer v3.7, using the coordinates from Rac–RhoGDI complex (PDB 1DS6) [18] and ribbon rendering, with the {alpha}-helices in green and β-sheets in red. GDP is rendered in blue and the magnesium ion in purple.

 
In several other instances intracellular ROS were demonstrated to have a role in EC motility. VEGF is a major signaling molecule in the vascular system. Upon interaction with its receptor, VEGF triggers a cascade of events that results in ROS generation and signaling. VEGF induces migration of EC [86–88] and of smooth muscle cells [89], and in both cases inhibition of ROS formation also prevented VEGF-induced motility. Furthermore, in vivo wound healing is accelerated by hyperbaric oxygen treatments, a finding likely related to the fact that wound-related cells possess specialized enzymes that generate ROS, and defects in these enzymes are associated with impaired healing [90]. ROS were demonstrated to be involved in various in vitro angiogenesis assays, such as Matrigel invasion by EC induced by hypoxia/reoxygenation [91] and ethanol [92]. In this latter case, the mediator of actin filament reorganization and increased formation of lamelipodia and filopodia in SVEC4-10 cells was H2O2, downstream of Cdc42.

A circumstance where EC motility is essential is angiogenesis–the formation of new blood vessels. A much-studied, but still not well understood process, angiogenesis involves targeted EC motility in response to specific signals. Among the signaling molecules involved in the regulation of angiogenesis one finds again the small GTPases from the Rho family, as well as Pak1, the downstream effector of Rac1 and Cdc42 [93]. A link between Rac1 and NAD(P)H oxidase activation, ROS generation, and angiogenesis has been recently recognized [45]. Indeed, a role of NAD(P)H oxidase in VEGF-induced angiogenesis was shown in vitro [86], and confirmed in vivo in a model of ischemic hind limb neovascularization [94]. Other signaling molecules were also found to use ROS to mediate angiogenic effects: the small GTPase ARF6 was shown to be involved in the temporal–spatial organization of caveolae or lipid rafts and in ROS-dependent VEGF signaling in EC, as well as in angiogenesis in vivo [95]. The interaction of angiopoietin-1 with its receptor tie-2 on EC triggered the production of ROS through activation of NAD(P)H oxidase, thus promoting EC migration while negatively regulating Erk1/2 phosphorylation [96]. Angiotensin II induced the cell surface matrix metalloproteinase MMP-2 (required for matrix penetration by mobilized EC) in a p47phox-dependent manner [97]. Thus, cellular processes needed for angiogenesis, whether physiologically- or pathologically-induced, require ROS generation through the activation of endothelial NAD(P)H oxidase [45].


   4. Direct interaction between ROS and actin
Top
 Abstract
 1. Introduction
 2. The non-phagocytic NAD(P)H...
 3. ROS and endothelial...
 4. Direct interaction between...
5. ROS and regulators...
 6. Conclusions and future...
 Appendix A. Supplementary data
 References
 
ROS were demonstrated to be necessary in other actin-based activities in various cell types: human sperm acrosomal reaction [98], low-density lipoprotein-induced stress fiber formation [99], EC mitogenesis and apoptosis [100], VCAM-1 signaling within EC upon engagement during lymphocyte diapedesis [101], integrin engagement during cell adhesion [102], and migration of human fibrosarcoma cells [103]. Thus, a wealth of evidence has accumulated connecting ROS generation within cells to the control of actin reorganization and cellular activities driven by this process, such as migration and shape changes. However, the mechanism of this control is complex, cell- and environment-dependent, and only recently some of the molecules involved have been described. To date, no interaction between any ROS and actin or actin-binding proteins has been directly demonstrated in vivo. Therefore a mechanism of ROS involvement in cellular motility cannot be described; instead, we will focus on the evidence that oxidants affect directly actin function.

The actin cytoskeleton is affected profoundly by the redox environment. For decades, the formation of actin filaments has been reconstituted in vitro in privileged redox conditions, in which a large excess of antioxidants, such as dithiothreitol, is added to protein preparations [104]. The reason for this redox bias is that in the absence of reducing agents, monomeric actin preparations are too unstable for studies of protein–protein interaction [105]. However, until recently this redox sensitivity was not investigated in terms of actin biology, but in terms of what damage ROS, and especially H2O2, may inflict upon of actin [106]. Most models of actin regulation are drawn from studies using purified or enriched proteins that do not address the effect of ROS on actin biology. Yet recent work suggests that cytosolic redox conditions not only alter actin polymerization, but control it in a physiologically meaningful way [7].

ROS can affect the activity of signaling pathways by direct modification of regulatory proteins via disulphide bond formation, nitrosylation, carbonylation or glutathionylation [107]. Indeed, glutathionylated actin is present in vivo in various conditions of oxidative stress [108,109]. More importantly, actin glutathionylation, which inhibits actin polymerization, is reversible, suggesting that actin S-thiolation may prevent irreversible organization of microfilaments within cells by excessive actin polymerization and cross-linking under conditions of oxidative stress [110]. Finally, components of NAD(P)H oxidase are associated with the actin cytoskeleton. In both phagocytic and non-phagocytic cells, NAD(P)H oxidase was found associated with actin or with the F-actin binding protein moesin [27,55,60,61,111]. Moreover, this association is not a permanent one, but inducible, as suggested by the fact that VEGF stimulation of EC triggers translocation of p47phox, together with WAVE1, Rac1 and Pak1, to the membrane ruffles, where the most active actin reorganization takes place [55]. The topographical correlation of ROS formation and dynamical actin network suggests a reciprocal regulation of the two molecular systems.

To understand the mechanism by which O2Formula influences actin polymerization in vitro, we have studied the effect of ROS inhibitors DPI or MnTmPyP on cytochalasin D (CyD) inhibition of actin polymerization at actin filament barbed ends [56]. We found that treatment with DPI or MnTMPyP significantly reduced the elongation of actin filaments at the barbed (fast growing) ends and decreased the number of exposed barbed ends. These results support the idea that ROS regulate actin cytoskeletal dynamics by favoring the uncovering of new barbed ends, thus dramatically increasing the speed of actin polymerization [56]. Interestingly, the simultaneous treatment with CyD and MnTMPyP significantly increased the actin polymerization as compared to CyD alone or DPI+ CyD-treated cells (Fig. 4). This unexpected finding suggested that O2Formula dismutation by the SOD mimetic, which results in the secondary generation of H2O2, might enhance actin polymerization at the pointed ends, while inhibiting polymerization at barbed ends. This concept is consistent with the increase in the number and/or size of stress fibers which we had observed in the presence of MnTMPyP [56], and with previous reports indicating that H2O2 induced actin polymerization in vitro [112] and in cultured EC [113].


Figure 4
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  		Fig. 4 Effects of CyD and antioxidants on actin incorporation into filaments. Alexa488-labeled actin monomer incorporation into filaments was detected by flow cytometry during the initial 3-min interval after addition of labeled monomers in saponin-permeabilized cells, in the presence of CyD, as described in [56]. Note that while CyD strongly inhibits actin polymerization in control and DPI-treated cells, in the presence of MnTMPyP there still is a detectable level of actin incorporation within filaments. Data from this figure were used in calculating the results in Fig. 3 from [56]. *P<0.05 vs control.

 

   5. ROS and regulators of the actin cytoskeleton
Top
 Abstract
 1. Introduction
 2. The non-phagocytic NAD(P)H...
 3. ROS and endothelial...
 4. Direct interaction between...
 5. ROS and regulators...
6. Conclusions and future...
 Appendix A. Supplementary data
 References
 
ROS also appear to modulate the activity of actin-binding and actin-regulatory proteins. Rac1-mediated ROS production resulted in down-regulation of Rho activity and concomitant formation of membrane ruffles and integrin-mediated cell spreading [114]. The mechanism involved inhibition of the low-molecular-weight protein tyrosine phosphatase (LMW-PTP) and a consequent increase in the tyrosine phosphorylation and activation of its targets, including p190Rho-GAP. This latter protein is a Rho inhibitor, which means that Rac1 down-regulates Rho and stress fiber formation in a redox-dependent manner, thus allowing remodeling of the actin network. The fact that Rac1 and Rho play opposing roles in the control of EC actin cytoskeleton was more recently demonstrated in hypoxia/reoxygenation [115]: the increase in EC permeability in this pathologic situation resulted from coordinated actions of Rac1 and RhoA, that led to remodeling of stress fibers and adherens junctions. This report underscores the propensity of both GTPases to be regulated by the cellular redox environment; indeed NAD(P)H oxidase-dependent formation of ROS may act either upstream or downstream of Rac1 on this pathway [115]. Interestingly, in several other instances ROS were found to modulate intercellular adhesion, with effects on the barrier function of EC layer. For example, H2O2 induced alterations in tight junction proteins as well as actin in bovine brain EC [116]. In another study, the constitutively active form of Rac (Tat-RacV12) caused a rapid loss of VE-cadherin-mediated cell–cell adhesion in EC from primary human umbilical veins [117]. The process was dependent upon the intracellular production of ROS and facilitated EC migration, which required a transient reduction of cell–cell adhesion. Another physiologically relevant situation where cell–cell adhesion is modulated by NAD(P)H oxidase-derived ROS is the engagement of VCAM-1 by adhering lymphocytes, prior to diapedesis [118].

In other work, the human immunodeficiency virus (HIV) type 1 Tat induced alterations in the actin cytoskeleton which were mediated by Pak1 and downstream from the activation of NAD(P)H oxidase [119]. Pak1 is a serine/threonine kinase and a Rac1-interacting protein, and although the cited report did not focus on Rac, the cytoskeletal changes observed in the presence of Tat are consistent with those reported in response to Rac1 activation [68,73,119]. Another protein involved in the control of EC migration and proliferation by ROS is IQGAP1, an effector of Rac1 and Cdc42 [71,120]. IQGAP1 binds directly to F-actin and cross-links actin filaments into irregular, interconnected bundles with gel-like properties. Upon VEGF interaction with its receptor type 2 (VEGFR2), IQGAP1 directly interacted with the active VEGFR2, which in turn rapidly promoted recruitment of Rac1 to IQGAP [88]. By inhibiting IQGAP1 by siRNA, the authors showed that IQGAP1 was involved in VEGF-stimulated ROS production, Akt phosphorylation, EC migration and proliferation. In a follow-up study using the in vitro EC wound assay, IQGAP1 appeared to link NOX2 to actin at the leading edge, thereby facilitating ROS production at the site of injury [69]. These in turn contributed to EC migration toward the injury site.

ROS may directly modulate the activity of some well-known actin-binding proteins. An example is cofilin (or actin depolymerizing factor, ADF), which dramatically increases the rate of actin depolymerization as a result of either severing them or increasing the off-rate for actin subunits at the pointed end of the actin filament [121]. Cofilin forms reversible disulfide-linked dimers and oligomers, with reduced severing activity and increased actin bundling capacity [121]. Another actin-binding protein, gelsolin, regulates actin filament dynamics by severing filaments and capping barbed ends [122]. Gelsolin is a downstream effector of Rac1, which triggers uncapping of actin filaments [123], a process important for the exposure of free barbed ends at ruffling sites [124]. Interestingly, there are two forms of gelsolin, one cytosolic and one found in blood plasma: they differ by the presence of a 25-amino acid peptide at the N-terminus in the secreted form, but more importantly by the presence of a disulphide bond in plasma gelsolin, that is absent in the intracellular one [125]. Moreover, reduced plasma gelsolin severed actin filaments more slowly than non-reduced gelsolin, and the rate of actin monomer binding was decreased [125,126]. To our knowledge there are no studies on the control of cytosolic gelsolin by its microenvironment redox status. However, it is tempting to speculate that ROS may increase actin severing activity of gelsolin and exposure of barbed ends through induction of disulfide bond formation, resulting in enhanced actin polymerization. Filamin is an actin cross-linking protein [127], and is induced by exogenously added H2O2 to translocate from the submembrane cytoskeletal networks to the cytosol, thus mediating the formation of inter-endothelial junctional gaps (enlargement of inter-endothelial space that leads to increased permeability) [128,129]. Focal adhesion kinase (FAK) is another crucial signaling component, involved in actin stress fibers control, EC-matrix adhesion and cellular movement. FAK is activated by numerous stimuli and functions as a biosensor or integrator to control cell motility [130]. In numerous instances the activation of this protein by phosphorylation was also found to be regulated by ROS [131–134].

Finally, nitric oxide (NO) metabolism tightly accompanies that of ROS in endothelial and other cell systems. For example, endothelial-type NO synthase (eNOS) expression is up-regulated in migrating EC in vitro as well as in vivo after exposure to shear stress [135]. O2Formula readily interacts with NO, a reaction that controls NO bioavailability and also results in the formation of peoxynitrite (ONOO–) [136]. In turn, ONOO– is a nitrating agent for tyrosyl residues. The complex O2Formula/NO system also leads to S-nitrosation of proteins, and both these reactions control protein functions [136]. Several cytoskeletal proteins were found to be nitrated: actin in EC [137] and in other cellular systems [138,139], profilin [140,141] and alpha-actinin [142]. Functionally, the angiogenic properties of EC in vitro could be modulated via ROS-dependent nitrosylation of actin [143].


   6. Conclusions and future directions
Top
 Abstract
 1. Introduction
 2. The non-phagocytic NAD(P)H...
 3. ROS and endothelial...
 4. Direct interaction between...
 5. ROS and regulators...
 6. Conclusions and future...
Appendix A. Supplementary data
 References
 
ROS are important modifiers in the physiological control of actin cytoskeleton dynamics by the 30+ families of ABPs [144], and more evidence is accumulating that links ROS generation with cellular motility (Fig. 5). Perhaps an adaptation to the relatively strong pro-oxidant environment to which EC from the arterial side of the vasculature are exposed (oxygenated blood pO2 is on the average 100 mm Hg, as opposed to ca. 40 mm Hg within tissues), it is recognized that EC need physiological amounts of ROS in order to function properly. More work is needed to understand the mechanisms by which ROS control actin reorganization and cellular movement. Furthermore, intriguing new research addresses the mechanisms by which the vascular endothelium is replenished and repaired, apparently in some cases by endogenous vascular precursor cells derived from stem cells of the adult bone marrow [145–147]. Adult stem cells of the mammalian epidermis, engineered to carry a targeted deletion of Rac1, induce exit from the stem cell niche in this adult tissue, resulting in proliferation and differentiation of the epidermal stem cells [148]. Given the established roles for both ROS and Rac1 in proliferation and apoptosis, and the ongoing development of animal models to study the repair of the vascular endothelium, it will be interesting to investigate the role of ROS stem cell mediated repair of the vascular endothelium.


Figure 5
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  		Fig. 5 ROS involvement in actin reorganization and cellular motility. ROS may interact directly with actin (circled1neg), as such (O2Formula, H2O2) or through the control of cellular redox homeostasis and of other redox couples, such as GSH/GSSG. As a result, polymerization parameters are altered at barbed ends, pointed ends, or both. ROS also control indirectly the cytoskeleton (circled2neg). They may interact with a number of actin-binding proteins (circled3neg) that control barbed end exposure (gelsolin), actin filament length (cofilin), association of filaments (filamin, IQGAP), and stress fibers formation (FAK), all of which control the temporal and spatial turnover of actin. Another indirect mechanism is through interaction with NO (circled4neg). The metabolites resulting from these reactions may react with actin (circled5neg) or with actin-binding proteins (circled6neg), such as {alpha}-actinin and profilin. It is also plausible that the monomer binding protein profilin modifies the impact of ROS on actin.

 

   Appendix A. Supplementary data
Top
 Abstract
 1. Introduction
 2. The non-phagocytic NAD(P)H...
 3. ROS and endothelial...
 4. Direct interaction between...
 5. ROS and regulators...
 6. Conclusions and future...
 Appendix A. Supplementary data
References
 
Supplementary data associated with this article can be found, in the online version, at doi:10.1016/j.cardiores.2006.05.003. This work was supported by NIH grant HL71536 (PJ.G.-C.).


   Acknowledgement
 
L.M. thanks Dr. Tim Eubank for help with the artwork.


   Notes
 
Time for primary review 23 days


   References
Top
 Abstract
 1. Introduction
 2. The non-phagocytic NAD(P)H...
 3. ROS and endothelial...
 4. Direct interaction between...
 5. ROS and regulators...
 6. Conclusions and future...
 Appendix A. Supplementary data
 References
 

  1. Sorescu D., Weiss D., Lassegue B., Clempus R.E., Szocs K., Sorescu G.P., et al. Superoxide production and expression of nox family proteins in human atherosclerosis. Circulation (2002) 105:1429–1435.[Abstract/Free Full Text]
  2. Matsunaga T., Nakajima T., Miyazaki T., Koyama I., Hokari S., Inoue I., et al. Glycated high-density lipoprotein regulates reactive oxygen species and reactive nitrogen species in endothelial cells. Metabolism (2003) 52:42–49.[CrossRef][ISI][Medline]
  3. Rahimi R., Nikfar S., Larijani B., Abdollahi M. A review on the role of antioxidants in the management of diabetes and its complications. Biomed Pharmacother (2005) 59:365–373.[CrossRef][Medline]
  4. Storz P. Reactive oxygen species in tumor progression. Front Biosci (2005) 10:1881–1896.[ISI][Medline]
  5. Nystrom T. Role of oxidative carbonylation in protein quality control and senescence. EMBO J (2005) 24:1311–1317.[CrossRef][ISI][Medline]
  6. Galli F., Piroddi M., Annetti C., Aisa C., Floridi E., Floridi A. Oxidative stress and reactive oxygen species. Contrib Nephrol (2005) 149:240–260.[ISI][Medline]
  7. Goldschmidt-Clermont P.J., Moldovan L. Stress, superoxide, and signal transduction. Gene Expr (1999) 7:255–260.[ISI][Medline]
  8. Moldovan L., Moldovan N.I. Oxygen free radicals and redox biology of organelles. Histochem Cell Biol (2004) 122:395–412.[CrossRef][ISI][Medline]
  9. Burdon R.H. Control of cell proliferation by reactive oxygen species. Biochem Soc Trans (1996) 24:1028–1032.[ISI][Medline]
  10. Finkel T. Reactive oxygen species and signal transduction. IUBMB Life (2001) 52:3–6.[ISI][Medline]
  11. Brandes R.P. Role of NADPH oxidases in the control of vascular gene expression. Antioxid Redox Signal (2003) 5:803–811.[CrossRef][ISI][Medline]
  12. Jackson M.J., Papa S., Bolanos J., Bruckdorfer R., Carlsen H., Elliott R.M., et al. Antioxidants, reactive oxygen and nitrogen species, gene induction and mitochondrial function. Mol Aspects Med (2002) 23:209–285.[CrossRef][Medline]
  13. Babior B.M. NADPH oxidase: an update. Blood (1999) 93:1464–1476.[Free Full Text]
  14. Quinn M.T., Mullen M.L., Jesaitis A.J., Linner J.G. Subcellular distribution of the Rap1A protein in human neutrophils: colocalization and cotranslocation with cytochrome b559. Blood (1992) 79:1563–1573.[Abstract/Free Full Text]
  15. Leusen J.H., Verhoeven A.J., Roos D. Interactions between the components of the human NADPH oxidase: a review about the intrigues in the phox family. Front Biosci (1996) 1:d72–d90.[Medline]
  16. Abo A., Pick E., Hall A., Totty N., Teahan C.G., Segal A.W. Activation of the NADPH oxidase involves the small GTP-binding protein p21rac1. Nature (1991) 353:668–670.[CrossRef][Medline]
  17. Knaus U.G., Heyworth P.G., Evans T., Curnutte J.T., Bokoch G.M. Regulation of phagocyte oxygen radical production by the GTP-binding protein Rac 2. Science (1991) 254:1512–1515.[Abstract/Free Full Text]
  18. Scheffzek K., Stephan I., Jensen O.N., Illenberger D., Gierschik P. The Rac–RhoGDI complex and the structural basis for the regulation of Rho proteins by RhoGDI. Nat Struct Biol (2000) 7:122–126.[CrossRef][ISI][Medline]
  19. Vignais P.V. The superoxide-generating NADPH oxidase: structural aspects and activation mechanism. Cell Mol Life Sci (2002) 59:1428–1459.[CrossRef][ISI][Medline]
  20. Diekmann D., Abo A., Johnston C., Segal A.W., Hall A. Interaction of Rac with p67phox and regulation of phagocytic NADPH oxidase activity. Science (1994) 265:531–533.[Abstract/Free Full Text]
  21. Sarfstein R., Gorzalczany Y., Mizrahi A., Berdichevsky Y., Molshanski-Mor S., Weinbaum C., et al. Dual role of Rac in the assembly of NADPH oxidase, tethering to the membrane and activation of p67phox: a study based on mutagenesis of p67phox-Rac1 chimeras. J Biol Chem (2004) 279:16007–16016.[Abstract/Free Full Text]
  22. Bayraktutan U., Draper N., Lang D., Shah A.M. Expression of functional neutrophil-type NADPH oxidase in cultured rat coronary microvascular endothelial cells. Cardiovasc Res (1998) 38:256–262.[Abstract/Free Full Text]
  23. Al Mehdi A.B., Zhao G., Dodia C., Tozawa K., Costa K., Muzykantov V., et al. Endothelial NADPH oxidase as the source of oxidants in lungs exposed to ischemia or high K+. Circ Res (1998) 83:730–737.[Abstract/Free Full Text]
  24. Meyer J.W., Holland J.A., Ziegler L.M., Chang M.M., Beebe G., Schmitt M.E. Identification of a functional leukocyte-type NADPH oxidase in human endothelial cells :a potential atherogenic source of reactive oxygen species. Endothelium (1999) 7:11–22.[ISI][Medline]
  25. Bayraktutan U., Blayney L., Shah A.M. Molecular characterization and localization of the NAD(P)H oxidase components gp91-phox and p22-phox in endothelial cells. Arterioscler Thromb Vasc Biol (2000) 20:1903–1911.[Abstract/Free Full Text]
  26. Hohler B., Holzapfel B., Kummer W. NADPH oxidase subunits and superoxide production in porcine pulmonary artery endothelial cells. Histochem Cell Biol (2000) 114:29–37.[ISI][Medline]
  27. Li J.M., Shah A.M. Intracellular localization and preassembly of the NADPH oxidase complex in cultured endothelial cells. J Biol Chem (2002) 277:19952–19960.[Abstract/Free Full Text]
  28. Parinandi N.L., Kleinberg M.A., Usatyuk P.V., Cummings R.J., Pennathur A., Cardounel A.J. Hyperoxia-induced NAD(P)H oxidase activation and regulation by MAP kinases in human lung endothelial cells. Am J Physiol Lung Cell Mol Physiol (2003) 284:L26–L38.[Abstract/Free Full Text]
  29. Suh Y.A., Arnold R.S., Lassegue B., Shi J., Xu X., Sorescu D., et al. Cell transformation by the superoxide-generating oxidase Mox1. Nature (1999) 401:79–82.[CrossRef][Medline]
  30. Lassegue B., Clempus R.E. Vascular NAD(P)H oxidases: specific features, expression, and regulation. Am J Physiol Regul Integr Comp Physiol (2003) 285:R277–R297.[Abstract/Free Full Text]
  31. Kikuchi H., Hikage M., Miyashita H., Fukumoto M. NADPH oxidase subunit, gp91(phox) homologue, preferentially expressed in human colon epithelial cells. Gene (2000) 254:237–243.[CrossRef][ISI][Medline]
  32. Geiszt M., Kopp J.B., Varnai P., Leto T.L. Identification of renox, an NAD(P)H oxidase in kidney. Proc Natl Acad Sci U S A (2000) 97:8010–8014.[Abstract/Free Full Text]
  33. Cheng G., Cao Z., Xu X., Van Meir E.G., Lambeth J.D. Homologs of gp91phox: cloning and tissue expression of Nox3, Nox4, and Nox5. Gene (2001) 269:131–140.[CrossRef][ISI][Medline]
  34. Geiszt M., Lekstrom K., Witta J., Leto T.L. Proteins homologous to p47phox and p67phox support superoxide production by NAD(P)H oxidase 1 in colon epithelial cells. J Biol Chem (2003) 278:20006–20012.[Abstract/Free Full Text]
  35. Brar S.S., Corbin Z., Kennedy T.P., Hemendinger R., Thornton L., Bommarius B., et al. NOX5 NAD(P)H oxidase regulates growth and apoptosis in DU 145 prostate cancer cells. Am J Physiol Cell Physiol (2003) 285:C353–C369.[Abstract/Free Full Text]
  36. Lambeth J.D. NOX enzymes and the biology of reactive oxygen. Nat Rev Immunol (2004) 4:181–189.[CrossRef][ISI][Medline]
  37. Banfi B., Clark R.A., Steger K., Krause K.H. Two novel proteins activate superoxide generation by the NADPH oxidase NOX1. J Biol Chem (2003) 278:3510–3513.[Abstract/Free Full Text]
  38. Geiszt M., Lekstrom K., Brenner S., Hewitt S.M., Dana R., Malech H.L., et al. NAD(P)H oxidase 1, a product of differentiated colon epithelial cells, can partially replace glycoprotein 91phox in the regulated production of superoxide by phagocytes. J Immunol (2003) 171:299–306.[Abstract/Free Full Text]
  39. Takeya R., Ueno N., Kami K., Taura M., Kohjima M., Izaki T., et al. Novel human homologues of p47phox and p67phox participate in activation of superoxide-producing NADPH oxidases. J Biol Chem (2003) 278:25234–25246.[Abstract/Free Full Text]
  40. Souza H.P., Liu X., Samouilov A., Kuppusamy P., Laurindo F.R., Zweier J.L. Quantitation of superoxide generation and substrate utilization by vascular NAD(P)H oxidase. Am J Physiol Heart Circ Physiol (2002) 282:H466–H474.[Abstract/Free Full Text]
  41. Li J.M., Shah A.M. Endothelial cell superoxide generation: regulation and relevance for cardiovascular pathophysiology. Am J Physiol Regul Integr Comp Physiol (2004) 287:R1014–R1030.[Abstract/Free Full Text]
  42. Gregg D., Rauscher F.M., Goldschmidt-Clermont P.J. Rac regulates cardiovascular superoxide through diverse molecular interactions: more than a binary GTP switch. Am J Physiol Cell Physiol (2003) 285:C723–C734.[Abstract/Free Full Text]
  43. Hu Q., Zheng G., Zweier J.L., Deshpande S., Irani K., Ziegelstein R.C. NADPH oxidase activation increases the sensitivity of intracellular Ca2+ stores to inositol 1,4,5-trisphosphate in human endothelial cells. J Biol Chem (2000) 275:15749–15757.[Abstract/Free Full Text]
  44. Vecchione C., Brandes R.P. Withdrawal of 3-hydroxy-3-methylglutaryl coenzyme A reductase inhibitors elicits oxidative stress and induces endothelial dysfunction in mice. Circ Res (2002) 91:173–179.[Abstract/Free Full Text]
  45. Ushio-Fukai M., Alexander R.W. Reactive oxygen species as mediators of angiogenesis signaling: role of NAD(P)H oxidase. Mol Cell Biochem (2004) 264:85–97.[CrossRef][ISI][Medline]
  46. Desideri G., Bravi M.C., Tucci M., Croce G., Marinucci M.C., Santucci A., et al. Angiotensin II inhibits endothelial cell motility through an AT1-dependent oxidant-sensitive decrement of nitric oxide availability. Arterioscler Thromb Vasc Biol (2003) 23:1218–1223.[Abstract/Free Full Text]
  47. Li J.M., Shah A.M. Mechanism of endothelial cell NADPH oxidase activation by angiotensin II. Role of the p47phox subunit. J Biol Chem (2003) 278:12094–12100.[Abstract/Free Full Text]
  48. Hu T., Ramachandrarao S.P., Siva S., Valancius C., Zhu Y., Mahadev K., et al. Reactive oxygen species production via NADPH oxidase mediates TGF-beta-induced cytoskeletal alterations in endothelial cells. Am J Physiol Renal Physiol (2005) 289:F816–F825.[Abstract/Free Full Text]
  49. Duerrschmidt N., Wippich N., Goettsch W., Broemme H.J., Morawietz H. Endothelin-1 induces NAD(P)H oxidase in human endothelial cells. Biochem Biophys Res Commun (2000) 269:713–717.[CrossRef][ISI][Medline]
  50. Hwang J., Ing M.H., Salazar A., Lassegue B., Griendling K., Navab M., et al. Pulsatile versus oscillatory shear stress regulates NADPH oxidase subunit expression: implication for native LDL oxidation. Circ Res (2003) 93:1225–1232.[Abstract/Free Full Text]
  51. Hwang J., Saha A., Boo Y.C., Sorescu G.P., McNally J.S., Holland S.M., et al. Oscillatory shear stress stimulates endothelial production of O2– from p47phox-dependent NAD(P)H oxidases, leading to monocyte adhesion. J Biol Chem (2003) 278:47291–47298.[Abstract/Free Full Text]
  52. Mata-Greenwood E., Grobe A., Kumar S., Noskina Y., Black S.M. Cyclic stretch increases VEGF expression in pulmonary arterial smooth muscle cells via TGF-beta1 and reactive oxygen species: a requirement for NAD(P)H oxidase. Am J Physiol Lung Cell Mol Physiol (2005) 289:L288–L289.[Abstract/Free Full Text]
  53. Gu Y., Xu Y.C., Wu R.F., Nwariaku F.E., Souza R.F., Flores S.C., et al. p47phox participates in activation of RelA in endothelial cells. J Biol Chem (2003) 278:17210–17217.[Abstract/Free Full Text]
  54. Xu Y.C., Wu R.F., Gu Y., Yang Y.S., Yang M.C., Nwariaku F.E., et al. Involvement of TRAF4 in oxidative activation of c-Jun N-terminal kinase. J Biol Chem (2002) 277:28051–28057.[Abstract/Free Full Text]
  55. Wu R.F., Gu Y., Xu Y.C., Nwariaku F.E., Terada L.S. Vascular endothelial growth factor causes translocation of p47phox to membrane ruffles through WAVE1. J Biol Chem (2003) 278:36830–36840.[Abstract/Free Full Text]
  56. Moldovan L., Moldovan N.I., Sohn R.H., Parikh S.A., Goldschmidt-Clermont P.J. Redox changes of cultured endothelial cells and actin dynamics. Circ Res (2000) 86:549–557.[Abstract/Free Full Text]
  57. Brandes R.P., Miller F.J., Beer S., Haendeler J., Hoffmann J., Ha T., et al. The vascular NADPH oxidase subunit p47phox is involved in redox-mediated gene expression. Free Radic Biol Med (2002) 32:1116–1122.[CrossRef][ISI][Medline]
  58. Brar S.S., Kennedy T.P., Sturrock A.B., Huecksteadt T.P., Quinn M.T., Whorton A.R., et al. An NAD(P)H oxidase regulates growth and transcription in melanoma cells. Am J Physiol Cell Physiol (2002) 282:C1212–C1224.[Abstract/Free Full Text]
  59. Grogan A., Reeves E., Keep N., Wientjes F., Totty N.F., Burlingame A.L., et al. Cytosolic phox proteins interact with and regulate the assembly of coronin in neutrophils. J Cell Sci (1997) 110:3071–3081.[Abstract]
  60. Tamura M., Kai T., Tsunawaki S., Lambeth J.D., Kameda K. Direct interaction of actin with p47(phox) of neutrophil NADPH oxidase. Biochem Biophys Res Commun (2000) 276:1186–1190.[CrossRef][ISI][Medline]
  61. Wientjes F.B., Reeves E.P., Soskic V., Furthmayr H., Segal A.W. The NADPH oxidase components p47(phox) and p40(phox) bind to moesin through their PX domain. Biochem Biophys Res Commun (2001) 289:382–388.[CrossRef][ISI][Medline]
  62. Doussiere J., Bouzidi F., Vignais P.V. The S100A8/A9 protein as a partner for the cytosolic factors of NADPH oxidase activation in neutrophils. Eur J Biochem (2002) 269:3246–3255.[ISI][Medline]
  63. Disanza A., Steffen A., Hertzog M., Frittoli E., Rottner K., Scita G. Actin polymerization machinery: the finish line of signaling networks, the starting point of cellular movement. Cell Mol Life Sci (2005) 62:955–970.[CrossRef][ISI][Medline]
  64. Wiesner S., Legate K.R., Fassler R. Integrin–actin interactions. Cell Mol Life Sci (2005) 62:1081–1099.[CrossRef][ISI][Medline]
  65. Postma M., Bosgraaf L., Loovers H.M., Van Haastert P.J. Chemotaxis: signalling modules join hands at front and tail. EMBO Rep (2004) 5:35–40.[CrossRef][ISI][Medline]
  66. Zweier J.L., Kuppusamy P., Thompson-Gorman S., Klunk D., Lutty G.A. Measurement and characterization of free radical generation in reoxygenated human endothelial cells. J Am Physiol (1994) 266:C700–C708.
  67. Crawford L.E., Milliken E.E., Irani K., Zweier J.L., Becker L.C., Johnson T.M., et al. Superoxide-mediated actin response in post-hypoxic endothelial cells. J Biol Chem (1996) 271:26863–26867.[Abstract/Free Full Text]
  68. Moldovan L., Irani K., Moldovan N.I., Finkel T., Goldschmidt-Clermont P.J. The actin cytoskeleton reorganization induced by Rac1 requires the production of superoxide. Antioxid Redox Signal (1999) 1:29–43.[Medline]
  69. Ikeda S., Yamaoka-Tojo M., Hilenski L., Patrushev N.A., Anwar G.M., Quinn M.T., et al. IQGAP1 regulates reactive oxygen species-dependent endothelial cell migration through interacting with Nox2. Arterioscler Thromb Vasc Biol (2005) 25:2295–2300.[Abstract/Free Full Text]
  70. Fukata M., Nakagawa M., Kaibuchi K. Roles of Rho-family GTPases in cell polarisation and directional migration. Curr Opin Cell Biol (2003) 15:590–597.[CrossRef][ISI][Medline]
  71. Watanabe T., Wang S., Noritake J., Sato K., Fukata M., Takefuji M., et al. Interaction with IQGAP1 links APC to Rac1, Cdc42, and actin filaments during cell polarization and migration. Dev Cell (2004) 7:871–883.[CrossRef][ISI][Medline]
  72. Tzima E. Role of small GTPases in endothelial cytoskeletal dynamics and the shear stress response. Circ Res (2006) 98:176–185.[Abstract/Free Full Text]
  73. Ridley A.J., Paterson H.F., Johnston C.L., Diekmann D., Hall A. The small GTP-binding protein rac regulates growth factor-induced membrane ruffling. Cell (1992) 70:401–410.[CrossRef][ISI][Medline]
  74. Ridley A.J., Hall A. The small GTP-binding protein rho regulates the assembly of focal adhesions and actin stress fibers in response to growth factors. Cell (1992) 70:389–399.[CrossRef][ISI][Medline]
  75. Nobes C.D., Hall A. Rho, rac, and cdc42 GTPases regulate the assembly of multimolecular focal complexes associated with actin stress fibers, lamellipodia, and filopodia. Cell (1995) 81:53–62.[CrossRef][ISI][Medline]
  76. Hall A. Rho GTPases and the control of cell behaviour. Biochem Soc Trans (2005) 33:891–895.[CrossRef][ISI][Medline]
  77. Grigoriev I., Borisy G., Vorobjev I. Regulation of microtubule dynamics in 3T3 fibroblasts by Rho family GTPases. Cell Motil Cytoskeleton (2006) 63:29–40.[CrossRef][ISI][Medline]
  78. Noritake J., Watanabe T., Sato K., Wang S., Kaibuchi K. IQGAP1: a key regulator of adhesion and migration. J Cell Sci (2005) 118:2085–2092.[Abstract/Free Full Text]
  79. Wittmann T., Waterman-Storer C.M. Spatial regulation of CLASP affinity for microtubules by Rac1 and GSK3beta in migrating epithelial cells. J Cell Biol (2005) 169:929–939.[Abstract/Free Full Text]
  80. Bagrodia S., Derijard B., Davis R.J., Cerione R.A. Cdc42 and PAK-mediated signaling leads to Jun kinase and p38 mitogen-activated protein kinase activation. J Biol Chem (1995) 270:27995–27998.[Abstract/Free Full Text]
  81. Bagrodia S., Taylor S.J., Creasy C.L., Chernoff J., Cerione R.A. Identification of a mouse p21Cdc42/Rac activated kinase. J Biol Chem (1995) 270:22731–22737.[Abstract/Free Full Text]
  82. Joneson T., McDonough M., Bar-Sagi D., Van Aelst L. RAC regulation of actin polymerization and proliferation by a pathway distinct from Jun kinase. Science (1996) 274:1374–1376.[Abstract/Free Full Text]
  83. Westwick J.K., Lambert Q.T., Clark G.J., Symons M., Van Aelst L., Pestell R.G., et al. Rac regulation of transformation, gene expression, and actin organization by multiple, PAK-independent pathways. Mol Cell Biol (1997) 17:1324–1335.[Abstract]
  84. Joneson T., Bar-Sagi D. A Rac1 effector site controlling mitogenesis through superoxide production. J Biol Chem (1998) 273:17991–17994.[Abstract/Free Full Text]
  85. Zhao Z.S., Manser E., Chen X.Q., Chong C., Leung T., Lim L. A conserved negative regulatory region in alphaPAK: inhibition of PAK kinases reveals their morphological roles downstream of Cdc42 and Rac1. Mol Cell Biol (1998) 18:2153–2163.[Abstract/Free Full Text]
  86. Ushio-Fukai M., Tang Y., Fukai T., Dikalov S.I., Ma Y., Fujimoto M., et al. Novel role of gp91(phox)-containing NAD(P)H oxidase in vascular endothelial growth factor-induced signaling and angiogenesis. Circ Res (2002) 91:1160–1167.[Abstract/Free Full Text]
  87. Abid M.R., Kachra Z., Spokes K.C., Aird W.C. NADPH oxidase activity is required for endothelial cell proliferation and migration. FEBS Lett (2000) 486:252–256.[CrossRef][ISI][Medline]
  88. Yamaoka-Tojo M., Ushio-Fukai M., Hilenski L., Dikalov S.I., Chen Y.E., Tojo T., et al. IQGAP1, a novel vascular endothelial growth factor receptor binding protein, is involved in reactive oxygen