Effect of the Rho-Kinase/ROCK Signaling Pathway on Cytoskeleton Components

Abstract

The mechanical properties of cells are important in tissue homeostasis and enable cell growth, division, migration and the epithelial-mesenchymal transition. Mechanical properties are determined to a large extent by the cytoskeleton. The cytoskeleton is a complex and dynamic network composed of microfilaments, intermediate filaments and microtubules. These cellular structures confer both cell shape and mechanical properties. The architecture of the networks formed by the cytoskeleton is regulated by several pathways, a key one being the Rho-kinase/ROCK signaling pathway. This review describes the role of ROCK (Rho-associated coiled-coil forming kinase) and how it mediates effects on the key components of the cytoskeleton that are critical for cell behaviour.

1. Introduction

The biomechanical properties of cells are inextricably linked to their intracellular structural components, such as the cytoskeleton [1]. Internal and external mechanical forces both act through the cytoskeleton to affect cellular mechanical properties, behaviour, cell spreading, movement, polarity and cytokinesis [2,3,4,5,6,7]. The cytoskeleton also plays essential biomechanical roles in connecting the plasma membrane of the cell, as well as internal membranes such as the endoplasmic reticulum, to the rest of the cell. The cytoskeletal network also regulates the diffusion of intracellular polymers that are larger than the network mesh size and possibly controls the permeation of water and small solutes through the cell [8]. Rho-associated coiled-coil forming kinase (ROCK) is considered to be a key regulator of the cytoskeleton and affects various important cellular functions such as cell shape, motility, secretion, proliferation and gene expression [9,10,11,12]. For example, various neural processes, including cell migration, axonal guidance, dendritic spine architecture, axonal regeneration and cell survival, are dependent on the ROCK signaling pathway [13]. The ROCK signaling pathway plays an important role in the reorganization of both microtubules and actin during the process formation of podocytes [14]. ROCK is associated with cancer progression and ROCK protein expression is higher in a number of cancer types such as primary stomach carcinoma, colon and bladder cancers and malignant melanoma [15]. A high level of ROCK protein expression correlates with poor overall survival in osteosarcoma and more aggressive behaviour in hepatocellular carcinomas [16,17,18]. This review is focused on the current knowledge of the regulation of the Rho-kinase/ROCK signaling pathway and the effects of ROCK on cytoskeletal components.

2. The Cytoskeleton

The main components of the cytoskeleton include microfilaments (specifically actin filaments), intermediate filaments and microtubules, which connect to each other, cellular organelles and the cell/nuclear membrane. They are biopolymers and organized into a three-dimensional network that resists deformation but can rearrange in response to internal or external forces and are thus responsible for maintaining the integrity of the cellular components. In general, there are four main functions of the cytoskeleton: organization of cellular contents; connection of cellular constituents to the external environment; mechanical resistance to deformation and generation of forces to move the cell or change its shape (Table 1) [2,19].

The three types of cytoskeletal components differ from each other, both chemically and physically. Generally, the most significant difference is their rigidity, which can be represented by the persistence length (l_P) [7]. The persistence length is a mechanical property quantifying the bending stiffness of a polymer, which is defined as the distance over which the filament is bent by thermal forces and is proportional to the stiffness of the polymer [66]. If a polymer is considered to be a uniform cylinder, l_P = B_S/k_BT, where B_S is bending stiffness, k_B is the Boltzmann constant and T is the absolute temperature [67]. Filament stiffness, length, and geometry of cross-linking together determine the mechanical properties of cytoskeletal networks. The most common difference among the cytoskeletal filament types is their bending stiffness. The l_P of microfilaments is 10–17 μm, the l_P of intermediate filaments is less than 0.3–1.0 μm, and the l_P of microtubules is more than 1 mm [68]. This shows that microtubules are not only larger but also stiffer than microfilaments and intermediate filament.

3. Microfilaments

Microfilaments, or more specifically actin filaments, are semiflexible (l_P ~ 10–17 μm) polymers of actin (~375 amino acids with a molecular mass of ~42-kDa) folded into a U-shaped double helix structure. Each actin monomer has four subdomains and a bound adenine nucleotide, either ATP, ADP-Pi, or ADP, and a bound divalent cation, Mg²⁺ [67,69,70]. Microfilaments are ~7 nm in diameter and up to several micrometres in length [71]. Bending, torsional, and twist-bend coupling elasticities influence the microfilament’s mechanical properties [70]. The properties of microfilaments are determined by the strength and distribution of inter-subunit contacts [72]. Microfilaments can assemble into different structures, such as networks and bundles that are closely packed and crosslinked by trans-membrane adhesion proteins.

The mechanical properties of microfilament networks are determined by the physical properties of the individual actin molecules, the connections/interactions between actin monomers, and the three-dimensional geometry of the actin arrangement [8]. Microfilament networks appear to be the primary mechanical component of the cytoskeleton and are particularly abundant and dense near the inner surface of the plasma membrane, where they form a network with a variety of capping, binding, branching, and severing proteins. G-actin (individual globular actin) is capable of polymerisation and can bind either ATP or ADP. The pool of unpolymerized actin in normal cells is nearly exclusively made up of ATP-G-actin, with little to no ADP-G-actin in the pool [73]. Polymerization is not an energy-consuming process [74]. ATP is hydrolysed a long time after polymerization occurs [75]. F-actin is formed by polymerization and the addition of G-actin at the growing (+) end of the microfilament. The process of adding a G-actin unit at one end and removing it at the other (-) end is called treadmilling [76,77]. The actin cytoskeleton network is constantly assembling and disassembling by treadmilling, which can drive the formation of protrusive structures such as filopodia and lamellipodia. As a result, the network not only provides mechanical support, determines cell shape, but also drives cell locomotion by the extension of pseudopods, thereby enabling cellular migration, division, and particle engulfment [20,21,22]. The connection between the microfilaments and the trans-membrane adhesion proteins also facilitates communication between the intracellular and extracellular mechanical signals, which allows cells to detect and respond to both chemical and mechanical signals from their extracellular environment [78]. Microfilament networks are viscoelastic as they both store and dissipate mechanical energy [79]. The elasticity of the networks allows them to resist deformation like a simple spring. This resistance to deformation and ability to recover was found to be dependent on rho-kinase mediated contractility [80].

4. Intermediate Filaments

Intermediate filaments play an important role in cell mechanics, signaling and homeostasis. They belong to a superfamily of highly conserved, α-helix-rich fibrous proteins that are encoded by approximately 65–70 different genes [81,82,83,84]. They are more flexible biopolymers (l_P = 0.3–1.0 μm) than microfilaments and microtubules. Their diameter is from 8 to 12 nm, intermediate in size between microfilaments and microtubules [7]. Thus far, six major types of intermediate filament have been described [85], and their abundance varies between cell types. Examples of the different intermediate filaments are: acidic keratins (type I), basic keratins (type II), vimentin, glial fibrillary acidic protein (GFAP), peripherin and desmin (type III), neurofilaments (type IV), lamin (type V), and filensin and phakinin (type VI) [86]. Type I and II keratins are highly expressed in epithelial cells [29,32,87,88]. The type III protein vimentin is present mainly in cells of mesenchymal origin [41], whereas desmin, GFAP and peripherin are mainly found in muscle, glial cells, and neurons of the peripheral system, respectively [36,89]. Type IV proteins are found in many types of mature neurons [42,85]. Type V proteins are the nuclear lamins which form a filamentous support inside the inner nuclear membrane [36,47]. The type VI proteins, phakinin and filensin, are lens fibre cell specific intermediate filaments [52,56,90].

Intermediate filaments are vital determinants of intracellular organelle organization and form a delicate network in the cytoplasm, encapsulating the nucleus and radiating toward the plasma membrane [19]. They are the main proteins providing mechanical resistance to the cells, because of their flexibility, elasticity and extensibility [85]. Both in vitro and in vivo studies have shown that intermediate filaments are important contributors to the elasticity and tensile strength of cells [91,92,93]. Indeed, intermediate filaments have been shown to determine cell stiffness in keratinocytes [93]. Indirect perturbation of cytoplasmic intermediate filaments has detrimental effects on cell stiffness. Research has shown that lipids such as sphingosylphosphorylcholine, induce perinuclear rearrangement of intermediate filaments leading to a significant increase in cellular elasticity, which could facilitate metastasis [94].

5. Microtubules

Microtubules are cylindrical polymers of α and β-tubulin dimers. There are multiple genes encoding tubulin protein isotypes [95,96]. According to The HUGO Gene Nomenclature Committee, there are 24 tubulin genes (10 alpha tubulins, 10 beta tubulins, 1 delta, 1 epsilon and 2 gamma tubulins) and 3 pseudogenes [97]. These tubulin proteins polymerize into protofilaments (α–β)n, and arrange in a spiral to form a hollow tube with a 25 nm diameter [67,98]. In addition, γ-tubulin which is located at the centrosome, has an important role in initiating microtubule assembly. Microtubules have an l_P > 1 mm and are stiffer than microfilaments and intermediate filaments. They are relatively brittle and can be fractured more easily than intermediate filaments [68]. The published flexural rigidity ranges from 1 × 10⁻²⁴ to 32 × 10⁻²⁴ Nm² [99]. Microtubules form a dynamic scaffold for the cells and are involved in a variety of functions, including the transportation of intracellular vesicles and organelles throughout the cell during normal physiological processes, force generation (by polymerization, depolymerization, or interactions with motor proteins), formation of intracellular transport platforms (for anchoring, signaling and force-coupling roles), segregation of chromosomes, assembly of the mitotic spindle in dividing cells, axon extension in neurons, directional cell migration and differentiation, maintaining cell stiffness/shape, and regulation of cell morphology and cell mechanics [57,100,101]. Microtubules undergo constant cycles of polymerization and depolymerization utilizing a guanosine triphosphate (GTP) dependent process (dynamic instability). The polymerization of microtubules begins with the formation of 13 linear protofilaments which are composed of head to tail arrays of tubulin dimers. Similar to actin, microtubules have two distinct ends (plus and minus ends). The unprotected minus ends are unstable and often require stabilizer proteins (calmodulin-regulated spectrin-associated proteins or Partonins) to prevent depolymerization [102]. On the other hand, microtubules elongate by the addition of tubulin subunits to the plus end [103]. They grow by αβ-tubulin heterodimers bound to GTP binding to the growing end of the microtubule. The GTP bound to the αβ-tubulin heterodimers can be hydrolyzed to GDP during or immediately after polymerization [104]. This hydrolysis decreases the binding affinity of tubulin polymers for adjacent molecules, thereby favouring depolymerization and resulting in the dynamic behaviour of microtubules (alternation between cycles of growth and shrinkage) [105]. Microtubules can therefore undergo periods of growth and disassembly, which means that microtubules have half-lives of only several minutes within the cell [106]. This is particularly important for the remodelling of the cytoskeleton that occurs during mitosis, and for force generation and signaling [107]. In order to control this dynamic instability, many organisms have developed proteins that perturb microtubule dynamics, several of which are in use as cancer chemotherapeutics and anti-inflammatory drugs [105]. For example, the anti-cancer drug paclitaxel (Taxol) stabilizes microtubules and prevents their disassembly, promoting mitotic arrest and cell death [108]. In epithelial cells, microtubules are responsible for the spatial organization of the secretory and endocytic apparatuses, the facilitation of exocytic and post-endocytic protein transportation, participation in protein sorting and apico-basolateral polarity [109].

In addition to these three main cytoskeleton components, many other proteins are associated with the cytoskeleton, such as spectrin, kinesin, plectin, emerin, Sad1 and UNC-84 proteins, Klarsicht, ANC-1, and Syne homology proteins, and they are important for cellular mechanics. [19]. Together, the cellular mechanical properties are determined by complex interactions involving cytoskeletal proteins and polymers. There are several pathways that lead to major rearrangements of the cytoskeletal structures, namely: Rho-kinase/ROCK; PI3K/AKT; β-Catenin-independent Wnt (Wnt ligands stabilize β-catenin; β-catenin helps link cadherin adhesion molecules to cytoskeleton); glycogen synthase kinase-3 beta (Gsk-3β); Mitogen-activated protein kinase (MAPK); and cyclic GMP-dependent protein kinase [110,111,112,113,114,115,116,117,118,119] (Table 2).

Other kinases, such as PI4P and protein kinase C, are also important in regulating interactions of the cell membrane proteins and protein scaffolds involved in vesicle budding, and cytoskeletal organization [197,198]. In particular, ROCK has been considered as a key regulator of the cytoskeleton as it mediates various important cellular functions such as cell shape, motility, secretion, proliferation, and gene expression [199].

6. Rho-Associated Kinase/ROCK Pathway and Associated Genes

6.1. Rho GTPases

Rho GTPases are key regulators of several cellular processes, including cytoskeletal shape, gene expression, cell cycle progression, cell polarity and cell migration [200,201,202]. Rho GTPases are a sub-family within the Ras superfamily of G-proteins and comprise at least 20 members, including Rho (A, B, C isoforms), Rac (1, 2, 3 isoforms), Cdc42 (Cdc42Hs, G25K isoforms), Rnd1/Rho6, Rnd2/Rho7, Rnd3/RhoE, RhoD, RhoG, TC10 and TTF [203,204,205]. In particular, RhoA, RhoB and RhoC, have similar effectors and modes of action, which impact many cellular processes [206]. They serve as gate control molecules by cycling between a GTP-bound active and GDP-bound inactive states-facilitated by Rho GTPase-specific guanine nucleotide exchange factors (GEFs) [207,208,209]. Subsequently, Rho proteins regulate downstream pathways which affect cell migration, adhesion, proliferation, apoptosis and the cell cycle via induction of contractile fibre bundles such as microfilaments, modulating microtubules and regulating intermediate filament turnover [203,204,210,211]. Rho GTPases interact with at least 30 effector proteins and initiate downstream signaling [212,213]. The effectors of three members of the family (RhoA, Rac1 and Cdc42) have been well studied. RhoA regulates the contraction of moving cells, whereas Rac1 and Cdc42 mediate the formation of lamellipodia and filopodia, respectively [214]. Rho GTPase effectors can be classified into different groups according to their functions, such as scaffold proteins, kinases, actin-binding proteins, formin-like molecules, and phospholipases [215]. For example, ROCK1,2 (serine/threonine kinases) and Dia1,2 (scaffold proteins) are important RhoA effectors that regulate cytoskeleton nucleation and polymerization [206]. Wiskott–Aldrich syndrome protein (WASP; a scaffold protein) and neural-WASP (scaffold protein N-Wasp) are Cdc42 effectors. N-Wasp mediates actin polymerization via the Arp2/3 complex [216]. Formin-homology-domain-containing protein (formin-like molecule Fhod1) is one of the Rac1 effectors that regulates actin cytoskeleton organization and gene transcription [217]. p21-activated (serine/threonine) kinase is the effector for both Rac1 and Cdc42 [218], it regulates cell shape and polarity through phosphorylation of multiple cytoskeletal proteins.

6.2. ROCK

Rho-associated kinase ROCK is one of the most important effectors downstream of Rho GTPase [219] (Figure 1). Human ROCKs consist of two isoforms, ROCK 1 and ROCK 2. ROCK 1 is located on chromosome 18 (18q11.1, human), and its mRNA is highly expressed in bone marrow and adipose cells (https://www.ncbi.nlm.nih.gov/gene/6093#gene-expression, accessed on 21 December 2022) whereas ROCK2 is located on chromosome 2 (2p25.1, human) and its mRNA is expressed abundantly in adipose tissue and the colon (https://www.ncbi.nlm.nih.gov/gene/9475, accessed on 21 December 2022) [220,221]. ROCK proteins are activated by binding Rho-GTPase which then, directly and indirectly, regulate the cytoskeleton. ROCK proteins induce a wide range of cellular responses that involve microfilaments, intermediate filaments and microtubules (Figure 2). Their downstream targets are membrane distal, including LIM kinases, myosin phosphatase target subunit of myosin light chain phosphatase, myosin light chain (MLC), collapsing response mediator protein, and ezrin-radixin-moesin (ERM) proteins. For example, ROCK controls the organization and stabilization of actin filaments by phosphorylating a number of proteins, such as MLC [222], LIM1/2 [223], myosin phosphatase target subunit 1 (MYPT1) [224], ERM [225], adducin [226], calponin [227], myristoylated alanine-rich C kinase substrate (MARCKS) [228], elongation factor-1 alpha (EF1α) [229], troponin I/T [230] and profilin [231]. ROCK phosphorylates MLC either directly or inactivates MLC phosphatase, resulting in the induction of actin-myosin contractility [232]. ROCK also activates LIM-Kinase by phosphorylation and LIM-Kinase in turn phosphorylates cofilin, which inhibits cofilin’s actin depolymerization activity, resulting in the stabilization of the actin cytoskeleton [223] (Figure 2). Both the ROCK/MYPT1/MLC and ROCK/LIM kinases/cofilin pathways are key elements in stress fibre assembly and cell adhesion [233]. Moreover, although ROCK1 and 2 have high sequence identity, they differ functionally, especially in relation to actin regulation (Figure 1). ROCK 1 is thought to be involved in destabilizing actin via regulating MLC and actin-myosin contraction, whereas ROCK 2 acts via cofilin to stabilize the actin cytoskeleton [233]. However, Wang et al., found that ROCK 1 phosphorylates LIM-kinase to inactivate cofilin, so regulating cell adhesion and invasion, but not ROCK 2 does not [234]. Also, Rochelle et al., demonstrated that ROCK 1 mediated amoeboid motility via destrin but not via cofilin [235].

ROCK proteins regulate different types of intermediate filaments via phosphorylation. For example, ROCK proteins can breakdown and separate the intermediate filaments via phosphorylation. It also upregulates the cell stiffness by interacting with keratin intermediate filaments [130,236,237,238,239]. Interestingly, activation of ROCK induces the deformation of intermediate filaments (Figure 2) with an instant release of inactive ROCK. The released ROCK is transported to the periphery of the cell and reactivated by Rho-GTP, it can then accelerate the phosphorylation and disassembly of intermediate filaments [240]. In addition, ROCK regulates microtubules by phosphorylation of several microtubule-associated proteins such as MAP2/Tau [241], collapsin response mediator protein 2 (CRMP2) [242] and Doublecortin [243] (Figure 2). ROCK-induced phosphorylation of MAP2/Tau leads to destabilization of the microtubules [12,244]. CRMPs stabilize microtubules however this is inhibited by phosphorylation via ROCK [245]. Doublecortin is a microtubule-associated protein. Phosphorylation of Doublecortin via ROCK inhibits microtubule bundling [243]. In addition, ROCK regulates microtubule acetylation via phosphorylation of the tubulin polymerization promoting protein 1 (TPPP1/p25), resulting in a decreased cellular level of acetylated tubulin and increased cell migration [246].

There are six genes that either enhance (disabled 2 (Dab2), synaptopodin 2 (Synpo2) and thymus cell antigen 1, theta (Thy1)) or inhibit (cysteine and histidine-rich domain (CHORD)-containing, zinc-binding protein 1 (Chordc1), heart of glass (Heg1), and Ras interacting protein 1 (Rasip1)) ROCK activity. For example, Dab2 (on chromosome 15), which plays an important role in cell proliferation and differentiation, is capable of upregulating RhoA/ROCK signaling [247]. SYNPO2 (known as myopodin), encodes an actin-binding protein, and has been characterized as a tumour suppressor which positively regulates ROCK [248]. It may promote cell migration through activation of the ROCK signaling pathway via increasing levels of Rho-GTP [249]. Thy1 (on chromosome 9) triggers the actin cytoskeleton remodelling via the Thy-1-CBP-Csk-Src-RhoA-ROCK axis [250]. However, overexpressed Morgana/chp-1, encoded by the CHORDC1 gene (on chromosome 9), negatively regulates Rho, and binds then inhibits ROCK 1 and 2 [251]. Rasip1 (on chromosome 7) increases the activity of Cdc42 and suppresses actomyosin contractility via inhibition of RhoA [252]. Silencing HEG1 (on chromosome 16) expression by siRNA or shRNA increases phosphorylation of MLC and formation of stress fibres indicating increased ROCK signaling [253]. Together with these regulating genes, the ROCK signaling pathway plays an important role in cytoskeleton and cellular mechanics.

ROCK inhibitors act to suppress the ROCK pathway through multiple mechanisms. They can relax the trabecular meshwork through inhibiting the actin cytoskeleton contractile tone. They can induce beta-catenin nuclear translocation and inhibit cell migration or block ATP-dependent phosphorylation therefore inhibiting ROCK1 and ROCK2 and can inhibit Rho GTPases from binding to ROCK [233,254,255]. More than 170 chemicals are capable of inhibiting ROCK, either selectively or non-selectively [256]. Most of them are ATP-competitive kinase inhibitors [257,258]. Structurally, they can be classified into isoquinoline/isoquinolinone, indazole, pyridine, pyrimidine, pyrrolopyridine, pyrazole, benzimidazole, benzothiazole, benzathiophene, benzamide, phthalazinone, aminofurazan, quinazoline, and boron derivatives [256]. Several ROCK inhibitors have been used in clinical trials for many therapeutic indications, such as ophthalmology, cardiovascular disease, anti-erectile dysfunction, and cancers. For example, both Ripasudil and Netarsudil have been proven to be of benefit in the treatment of glaucoma [259]. Fasudil, which is a potent vasodilator, has been approved for the treatment of cerebral vasospasm in Japan and China since 1995 [260]. Y-27632 improved ventricular hypertrophy, fibrosis, and function in a rat model [261,262]. Daily administration of Y-27632 also improved erectile responses in a rat model [263,264]. The use of ROCK inhibitors in cancer, such as gastric, bone, lung, breast, renal, liver and prostate cancers, has been widely characterised [265,266,267,268,269,270,271]. For example, the selective ROCK inhibitor RKI-1447 was shown to have significant anti-invasive and anti-tumour effects [272]. AT13148, an inhibitor of ROCK and AKT kinases, has been found to have antimetastatic and antiproliferative activity [273]. Despite the potential of ROCK inhibitors, the number of clinical trials for human cancer is still limited.

Although ROCK 1 and ROCK 2 are highly homologous kinases and are both involved in the Rho/ROCK signaling pathway, their target substrates and physiological/pathological activities differ. Unfortunately, most ROCK inhibitors cannot selectively target one or other of the two isoforms; such inhibitors would be very useful. Dimerization is required for the RhoE activation of ROCK 1 [274], and although the homology between the kinase domains of ROCK 1 and ROCK 2 is ~90%, the homology between the N-terminal dimerization domains [275] of the two proteins is only ~60% [276]. Therefore dimerization-disrupting peptides could be developed and used to selectively target the dimerization domains of the two isoforms [276]. Autophosphorylation of ROCK 1 at Ser1333 and ROCK 2 at Ser1366 is required for activation of the kinases [277]. It has been shown that the interaction between Thr405 in the hydrophobic motif of ROCK 2 and Asp39 in the N-terminal extension was essential for both kinase activation and dimerization [278]. This provides further support for the development of peptides that prevent dimerization and kinase activity of ROCK isoforms.

7. Conclusions

The cytoskeleton gives cells their shape, structure and mechanical support and plays an essential role in cellular mechanics. ROCK is an effector of the small Rho GTPase and when activated influences many cellular functions and processes by regulating different elements of the cytoskeleton. Many cellular and physiological functions are mediated by ROCK and its activity is often elevated in some disorders, making them good targets for the development of new drugs. ROCK inhibitions have been found to effectively manage several diseases in humans and animal models. Progress has been made towards understanding how non-selective ROCK inhibitors work for several diseases and conditions and efforts should be made to develop ROCK inhibitors with improved specificity and sensitivity.

Competing interests

Conflicts of InterestThe authors declare no conflict of interest.

References

1. Guan G., He Y., Mei L.. Atomic force microscopy: A nanobiotechnology for cellular research. Nano TransMed.. 2022 doi:10.26599/NTM.2022.9130004
2. Fletcher D.A., Mullins R.D.. Cell mechanics and the cytoskeleton. Nature. 2010 doi:10.1038/nature08908 PubMed
3. Zhu C., Bao G., Wang N.. Cell mechanics: Mechanical response, cell adhesion, and molecular deformation. Annu. Rev. Biomed. Eng.. 2000 doi:10.1146/annurev.bioeng.2.1.189 PubMed
4. Walker M., Rizzuto P., Godin M., Pelling A.E.. Structural and mechanical remodeling of the cytoskeleton maintains tensional homeostasis in 3D microtissues under acute dynamic stretch. Sci. Rep.. 2020 doi:10.1038/s41598-020-64725-7 PubMed
5. Reichl E.M., Effler J.C., Robinson D.N.. The stress and strain of cytokinesis. Trends. Cell Biol.. 2005 doi:10.1016/j.tcb.2005.02.004 PubMed
6. Dalous J., Burghardt E., Müller-Taubenberger A., Bruckert F., Gerisch G., Bretschneider T.. Reversal of cell polarity and actin-myosin cytoskeleton reorganization under mechanical and chemical stimulation. Biophys. J.. 2008 doi:10.1529/biophysj.107.114702 PubMed
7. Ananthakrishnan R., Ehrlicher A.. The forces behind cell movement. Int. J. Biol. Sci.. 2007 doi:10.7150/ijbs.3.303 PubMed
8. Pegoraro A.F., Janmey P., Weitz D.A.. Mechanical Properties of the Cytoskeleton and Cells. Cold Spring Harb. Perspect. Biol.. 2017 doi:10.1101/cshperspect.a022038 PubMed
9. Riento K., Ridley A.J.. ROCKs: Multifunctional kinases in cell behaviour. Nat. Rev. Mol. Cell Biol.. 2003 doi:10.1038/nrm1128 PubMed
10. McBeath R., Pirone D.M., Nelson C.M., Bhadriraju K., Chen C.S.. Cell shape, cytoskeletal tension, and RhoA regulate stem cell lineage commitment. Dev. Cell. 2004 doi:10.1016/S1534-5807(04)00075-9 PubMed
11. Maekawa M., Ishizaki T., Boku S., Watanabe N., Fujita A., Iwamatsu A. et al. Signaling from Rho to the actin cytoskeleton through protein kinases ROCK and LIM-kinase. Science. 1999 doi:10.1126/science.285.5429.895 PubMed
12. Amano M., Nakayama M., Kaibuchi K.. Rho-kinase/ROCK: A key regulator of the cytoskeleton and cell polarity. Cytoskeleton. 2010 doi:10.1002/cm.20472 PubMed
13. Schmandke A., Schmandke A., Strittmatter S.M.. ROCK and Rho: Biochemistry and neuronal functions of Rho-associated protein kinases. Neuroscientist. 2007 doi:10.1177/1073858407303611 PubMed
14. Gao S.Y., Li C.Y., Chen J., Pan L., Saito S., Terashita T. et al. Rho-ROCK signal pathway regulates microtubule-based process formation of cultured podocytes - Inhibition of ROCK promoted process elongation. Nephron Exp. Nephrol.. 2004 doi:10.1159/000078406 PubMed
15. Morgan-Fisher M., Wewer U.M., Yoneda A.. Regulation of ROCK activity in cancer. J. Histochem. Cytochem.. 2013 doi:10.1369/0022155412470834 PubMed
16. Lane J., Martin T.A., Watkins G., Mansel R.E., Jiang W.G.. The expression and prognostic value of ROCK I and ROCK II and their role in human breast cancer. Int. J. Oncol.. 2008 doi:10.3892/ijo_00000044 PubMed
17. Liu X., Choy E., Hornicek F.J., Yang S., Yang C., Harmon D. et al. ROCK1 as a potential therapeutic target in osteosarcoma. J. Orthop. Res.. 2011 doi:10.1002/jor.21403 PubMed
18. Wong C.C., Wong C.M., Tung E.K., Man K., Ng I.O.. Rho-kinase 2 is frequently overexpressed in hepatocellular carcinoma and involved in tumor invasion. Hepatology. 2009 doi:10.1002/hep.22836 PubMed
19. Darling E.M., Di Carlo D.. High-Throughput Assessment of Cellular Mechanical Properties. Annu. Rev. Biomed. Eng.. 2015 doi:10.1146/annurev-bioeng-071114-040545 PubMed
20. Jiang X., Qin Y., Kun L., Zhou Y.. The Significant Role of the Microfilament System in Tumors. Front. Oncol.. 2021 doi:10.3389/fonc.2021.620390 PubMed
21. Falahzadeh K., Banaei-Esfahani A., Shahhoseini M.. The potential roles of actin in the nucleus. Cell Journal. 2015 doi:10.22074/cellj.2015.507 PubMed
22. Gordón-Alonso M., Veiga E., Sánchez-Madrid F.. Actin dynamics at the immunological synapse. Cell Health Cytoskelet.. 2010
23. Blessing C.A., Ugrinova G.T., Goodson H.V.. Actin and ARPs: Action in the nucleus. Trends Cell Biol.. 2004 doi:10.1016/j.tcb.2004.07.009 PubMed
24. Halpain S.. Actin in a supporting role. Nat. Neurosci.. 2003 doi:10.1038/nn0203-101 PubMed
25. Lanier L.M., Gertler F.B.. Actin cytoskeleton: Thinking globally, actin’ locally. Curr. Biol.. 2000 doi:10.1016/S0960-9822(00)00685-0 PubMed
26. Pollard T.D., Blanchoin L., Mullins R.D.. Molecular mechanisms controlling actin filament dynamics in nonmuscle cells. Annu. Rev. Biophys. Biomol. Struct.. 2000 doi:10.1146/annurev.biophys.29.1.545 PubMed
27. Schmidt A., Hall M.N.. Signaling to the actin cytoskeleton. Annu. Rev. Cell Dev. Biol.. 1998 doi:10.1146/annurev.cellbio.14.1.305 PubMed
28. Bretscher A.. Microfilaments and membranes. Curr. Opin. Cell Biol.. 1993 doi:10.1016/0955-0674(93)90136-E PubMed
29. Gül D., Habtemichael N., Dietrich D., Dietrich J., Gößwein D., Khamis A. et al. Identification of cytokeratin24 as a tumor suppressor for the management of head and neck cancer. Biol. Chem.. 2022 doi:10.1515/hsz-2021-0287 PubMed
30. Jacob J.T., Coulombe P.A., Kwan R., Omary M.B.. Types I and II Keratin Intermediate Filaments. Cold Spring Harb. Perspect. Biol.. 2018 doi:10.1101/cshperspect.a018275 PubMed
31. Saitoh T., Sato K., Tonogi M., Tanaka Y., Yamane G.Y.. Expression of Cytokeratin 13, 14, 17, and 19 in 4-nitroquinoline-1-oxide-induced Oral Carcinogenesis in Rat. Bull. Tokyo Dent. Coll.. 2016 doi:10.2209/tdcpublication.2010-0029 PubMed
32. Schweizer J., Bowden P.E., Coulombe P.A., Langbein L., Lane E.B., Magin T.M. et al. New consensus nomenclature for mammalian keratins. J. Cell Biol.. 2006 doi:10.1083/jcb.200603161 PubMed
33. Honda Y., Koike K., Kubo Y., Masuko S., Arakawa Y., Ando S.. In vitro assembly properties of human type I and II hair keratins. Cell Struct. Funct.. 2014 doi:10.1247/csf.13021 PubMed
34. Infante C., Ponce M., Asensio E., Zerolo R., Manchado M.. Molecular characterization of a novel type II keratin gene (sseKer3) in the Senegalese sole (Solea senegalensis): Differential expression of keratin genes by salinity. Comp. Biochem. Physiol. B Biochem. Mol. Biol.. 2011 doi:10.1016/j.cbpb.2011.04.004 PubMed
35. Moll R., Divo M., Langbein L.. The human keratins: Biology and pathology. Histochem. Cell Biol.. 2008 doi:10.1007/s00418-008-0435-6 PubMed
36. Bowden P.E., Quinlan R.A., Breitkreutz D., Fusenig N.E.. Proteolytic modification of acidic and basic keratins during terminal differentiation of mouse and human epidermis. Eur. J. Biochem.. 1984 doi:10.1111/j.1432-1033.1984.tb08246.x PubMed
37. Ridge K.M., Eriksson J.E., Pekny M., Goldman R.D.. Roles of vimentin in health and disease. Genes Dev.. 2022 doi:10.1101/gad.349358.122 PubMed
38. Yue Q., Feng L., Cao B., Liu M., Zhang D., Wu W. et al. Proteomic analysis revealed the important role of vimentin in human cervical carcinoma HeLa cells treated with gambogic acid. Mol. Cell. Proteom.. 2016 doi:10.1074/mcp.M115.053272 PubMed
39. Kidd M.E., Shumaker D.K., Ridge K.M.. The role of Vimentin intermediate filaments in the progression of lung cancer. Am. J. Respir. Cell Mol. Biol.. 2014 doi:10.1165/rcmb.2013-0314TR PubMed
40. Menko A.S., Bleaken B.M., Libowitz A.A., Zhang L., Stepp M.A., Walker J.L.. A central role for vimentin in regulating repair function during healing of the lens epithelium. Mol. Biol. Cell. 2014 doi:10.1091/mbc.e12-12-0900 PubMed
41. Herrmann H., Bär H., Kreplak L., Strelkov S.V., Aebi U.. Intermediate filaments: From cell architecture to nanomechanics. Nat. Rev. Mol. Cell Biol.. 2007 doi:10.1038/nrm2197 PubMed
42. Wang R., Li Q., Tang D.D.. Role of vimentin in smooth muscle force development. Am. J. Physiol. Cell Physiol.. 2006 doi:10.1152/ajpcell.00097.2006 PubMed
43. Didonna A., Opal P.. The role of neurofilament aggregation in neurodegeneration: Lessons from rare inherited neurological disorders. Mol. Neurodegener.. 2019 doi:10.1186/s13024-019-0318-4 PubMed
44. Yuan A., Nixon R.A.. Specialized roles of neurofilament proteins in synapses: Relevance to neuropsychiatric disorders. Brain Res. Bull.. 2016 doi:10.1016/j.brainresbull.2016.09.002 PubMed
45. Yuan A., Rao M.V., Veeranna, Nixon R.A.. A. Neurofilaments at a glance. J. Cell Sci.. 2012 doi:10.1242/jcs.104729 PubMed
46. Lin H., Schlaepfer W.W.. Role of neurofilament aggregation in motor neuron disease. Ann. Neurol.. 2006 doi:10.1002/ana.20965 PubMed
47. Hoffman P.N.. Distinct roles of neurofilament and tubulin gene expression in axonal growth. Ciba Found. Symp.. 1988 doi:10.1002/9780470513675.ch12 PubMed
48. Khadija S.G., Chen F., Hadden T., Commissaris R.L., Kowluru A.. Biology and regulatory roles of nuclear lamins in cellular function and dysfunction. Recent Pat. Endocr. Metab. Immune Drug Discov.. 2015 doi:10.2174/1872214809666151009120402 PubMed
49. Carmosino M., Torretta S., Procino G., Gerbino A., Forleo C., Favale S. et al. Role of nuclear Lamin A/C in cardiomyocyte functions. Biol. Cell. 2014 doi:10.1111/boc.201400033 PubMed
50. Burke B., Stewart C.L.. The nuclear lamins: Flexibility in function. Nat. Rev. Mol. Cell Biol.. 2013 doi:10.1038/nrm3488 PubMed
51. Lopez-Soler R.I., Moir R.D., Spann T.P., Stick R., Goldman R.D.. A role for nuclear lamins in nuclear envelope assembly. J. Cell Biol.. 2001 doi:10.1083/jcb.200101025 PubMed
52. Parnaik V.K.. Role of Nuclear Lamins in Nuclear Organization, Cellular Signaling, and Inherited Diseases. Int. Rev. Cell Mol. Biol.. 2008 doi:10.1016/S1937-6448(07)66004-3 PubMed
53. Oka M., Kudo H., Sugama N., Asami Y., Takehana M.. The function of filensin and phakinin in lens transparency. Mol. Vis.. 2008 PubMed
54. Pittenger J.T., Hess J.F., FitzGerald P.G.. Identifying the role of specific motifs in the lens fiber cell-specific intermediate filament phakosin. Invest. Ophthalmol. Vis. Sci.. 2007 doi:10.1167/iovs.07-0647 PubMed
55. Blankenship T.N., Hess J.F., FitzGerald P.G.. Development- and differentiation-dependent reorganization of intermediate filaments in fiber cells. Investig. Ophthalmol. Vis. Sci.. 2001 PubMed
56. Goulielmos G., Remington S., Schwesinger F., Georgatos S.D., Gounari F.. Contributions of the structural domains of filensin in polymer formation and filament distribution. J. Cell Sci.. 1996 doi:10.1242/jcs.109.2.447 PubMed
57. Georgatos S.D., Gounari F., Remington S.. The beaded intermediate filaments and their potential functions in eye lens. Bioessays. 1994 doi:10.1002/bies.950160609 PubMed
58. Hlavaty D., Lechler T.. Roles for microtubules in the proliferative and differentiated cells of stratified epithelia. Curr. Opin. Cell Biol.. 2021 doi:10.1016/j.ceb.2020.10.002 PubMed
59. Chang W., Gu J.G.. Role of microtubules in Piezo2 mechanotransduction of mouse Merkel cells. J. Neurophysiol.. 2020 doi:10.1152/jn.00502.2020 PubMed
60. Matis M.. The Mechanical Role of Microtubules in Tissue Remodeling. Bioessays. 2020 doi:10.1002/bies.201900244 PubMed
61. Logan C.M., Menko A.S.. Microtubules: Evolving roles and critical cellular interactions. Exp. Biol. Med.. 2019 doi:10.1177/1535370219867296 PubMed
62. Cirillo L., Gotta M., Meraldi P.. The elephant in the room: The role of microtubules in cancer. Adv. Exp. Med. Biol.. 2017 doi:10.1007/978-3-319-57127-0_5 PubMed
63. Ganguly A., Yang H., Sharma R., Patel K.D., Cabral F.. The role of microtubules and their dynamics in cell migration. J. Biol. Chem.. 2012 doi:10.1074/jbc.M112.423905 PubMed
64. van der Vaart B., Akhmanova A., Straube A.. Regulation of microtubule dynamic instability. Biochem. Soc. Trans.. 2009 doi:10.1042/BST0371007 PubMed
65. Kulić I.M., Brown A.E.X., Kim H., Kural C., Blehm B., Selvin P.R. et al. The role of microtubule movement in bidirectional organelle transport. Proc. Natl. Acad. Sci. USA. 2008 doi:10.1073/pnas.0800031105 PubMed
66. Morse D.C.. Viscoelasticity of Concentrated Isotropic Solutions of Semiflexible Polymers. 1. Model and Stress Tensor. Macromolecules. 1998 doi:10.1021/ma9803032
67. Wen Q., Janmey P.A.. Polymer physics of the cytoskeleton. Curr. Opin. Solid. State Mater. Sci.. 2011 doi:10.1016/j.cossms.2011.05.002 PubMed
68. Wen Q., Janmey P.A.. Effects of non-linearity on cell-ECM interactions. Exp. Cell Res.. 2013 doi:10.1016/j.yexcr.2013.05.017 PubMed
69. Gittes F., Mickey B., Nettleton J., Howard J.. Flexural rigidity of microtubules and actin filaments measured from thermal fluctuations in shape. J. Cell. Biol.. 1993 doi:10.1083/jcb.120.4.923 PubMed
70. De La Cruz E.M., Gardel M.L.. Actin Mechanics and Fragmentation. J. Biol. Chem.. 2015 doi:10.1074/jbc.R115.636472 PubMed
71. Cooper G.M.. The Cell: A Molecular Approach. 2013
72. De La Cruz E.M., Roland J., McCullough B.R., Blanchoin L., Martiel J.L.. Origin of twist-bend coupling in actin filaments. Biophys. J.. 2010 doi:10.1016/j.bpj.2010.07.009 PubMed
73. Atkinson S.J., Hosford M.A., Molitoris B.A.. Mechanism of Actin Polymerization in Cellular ATP Depletion*. J. Biol. Chem.. 2004 doi:10.1074/jbc.M306973200 PubMed
74. Kudryashov D.S., Reisler E.. ATP and ADP actin states. Biopolymers. 2013 doi:10.1002/bip.22155 PubMed
75. Janmey P.A., Hvidt S., Oster G.F., Lamb J., Stossel T.P., Hartwig J.H.. Effect of ATP on actin filament stiffness. Nature. 1990 doi:10.1038/347095a0 PubMed
76. Brieher W.. Mechanisms of actin disassembly. Mol. Biol. Cell. 2013 doi:10.1091/mbc.e12-09-0694 PubMed
77. Wegner A.. Treadmilling of actin at physiological salt concentrations: An analysis of the critical concentrations of actin filaments. J. Mol. Biol.. 1982 doi:10.1016/0022-2836(82)90411-9 PubMed
78. Janmey P.A., Miller R.T.. Mechanisms of mechanical signaling in development and disease. J. Cell Sci.. 2011 doi:10.1242/jcs.071001 PubMed
79. Xu J., Schwarz W.H., Käs J.A., Stossel T.P., Janmey P.A., Pollard T.D.. Mechanical Properties of Actin Filament Networks Depend on Preparation, Polymerization Conditions, and Storage of Actin Monomers. Biophys. J.. 1998 doi:10.1016/S0006-3495(98)77979-2 PubMed
80. Haase K., Pelling A.E.. The role of the actin cortex in maintaining cell shape. Commun. Integr. Biol.. 2013 doi:10.4161/cib.26714 PubMed
81. Qin Z., Kreplak L., Buehler M.J.. Hierarchical structure controls nanomechanical properties of vimentin intermediate filaments. PLoS ONE. 2009 doi:10.1371/journal.pone.0007294 PubMed
82. Qin Z., Buehler M.J.. Flaw tolerance of nuclear intermediate filament lamina under extreme mechanical deformation. ACS Nano. 2011 doi:10.1021/nn200107u PubMed
83. Hesse M., Magin T.M., Weber K.. Genes for intermediate filament proteins and the draft sequence of the human genome: Novel keratin genes and a surprisingly high number of pseudogenes related to keratin genes 8 and 18. J. Cell Sci.. 2001 doi:10.1242/jcs.114.14.2569 PubMed
84. Strnad P., Stumptner C., Zatloukal K., Denk H.. Intermediate filament cytoskeleton of the liver in health and disease. Histochem. Cell Biol.. 2008 doi:10.1007/s00418-008-0431-x PubMed
85. Sanghvi-Shah R., Weber G.F.. Intermediate Filaments at the Junction of Mechanotransduction, Migration, and Development. Front. Cell Dev. Biol.. 2017 doi:10.3389/fcell.2017.00081 PubMed
86. Oshima R.G.. Intermediate filaments: A historical perspective. Exp. Cell Res.. 2007 doi:10.1016/j.yexcr.2007.04.007 PubMed
87. Karantza V.. Keratins in health and cancer: More than mere epithelial cell markers. Oncogene. 2011 doi:10.1038/onc.2010.456 PubMed
88. Huang T.L., Chou C.C.. Effect of mutations on the hydrophobic interactions of the hierarchical molecular structure and mechanical properties of epithelial keratin 1/10. Int. J. Biol. Macromol.. 2022 doi:10.1016/j.ijbiomac.2022.05.160 PubMed
89. Lowery J., Kuczmarski E.R., Herrmann H., Goldman R.D.. Intermediate Filaments Play a Pivotal Role in Regulating Cell Architecture and Function. J. Biol. Chem.. 2015 doi:10.1074/jbc.R115.640359 PubMed
90. FitzGerald P., Sun N., Shibata B., Hess J.F.. Expression of the type VI intermediate filament proteins CP49 and filensin in the mouse lens epithelium. Mol. Vis.. 2016 PubMed
91. Fudge D., Russell D., Beriault D., Moore W., Lane E.B., Vogl A.W.. The intermediate filament network in cultured human keratinocytes is remarkably extensible and resilient. PLoS ONE. 2008 doi:10.1371/journal.pone.0002327 PubMed
92. Nolting J.-F., Möbius W., Köster S.. Mechanics of individual keratin bundles in living cells. Biophys. J.. 2014 doi:10.1016/j.bpj.2014.10.039 PubMed
93. Ma L., Xu J., Coulombe P.A., Wirtz D.. Keratin Filament Suspensions Show Unique Micromechanical Properties *. J. Biol. Chem.. 1999 doi:10.1074/jbc.274.27.19145 PubMed
94. Beil M., Micoulet A., von Wichert G., Paschke S., Walther P., Omary M.B. et al. Sphingosylphosphorylcholine regulates keratin network architecture and visco-elastic properties of human cancer cells. Nat. Cell Biol.. 2003 doi:10.1038/ncb1037 PubMed
95. Janke C., Magiera M.M.. The tubulin code and its role in controlling microtubule properties and functions. Nat. Rev. Mol. Cell Biol.. 2020 doi:10.1038/s41580-020-0214-3 PubMed
96. Roll-Mecak A.. The Tubulin Code in Microtubule Dynamics and Information Encoding. Dev. Cell. 2020 doi:10.1016/j.devcel.2020.06.008 PubMed
97. HGNC Database, HUGO Gene Nomenclature Committee (HGNC), European Molecular Biology Laboratory, European Bi-oinformatics Institute (EMBL-EBI), Wellcome Genome Campus, Hinxton, Cambridge CB10 1SD, United Kingdom. 2022
98. Sept D., MacKintosh F.C.. Microtubule elasticity: Connecting all-atom simulations with continuum mechanics. Phys. Rev. Lett.. 2010 doi:10.1103/PhysRevLett.104.018101 PubMed
99. Schaap I.A.T., Carrasco C., de Pablo P.J., MacKintosh F.C., Schmidt C.F.. Elastic Response, Buckling, and Instability of Microtubules under Radial Indentation. Biophys. J.. 2006 doi:10.1529/biophysj.105.077826 PubMed
100. Avila J.. Microtubule functions. Life Sci.. 1992 doi:10.1016/0024-3205(92)90433-P PubMed
101. Kent I.A., Lele T.P.. Microtubule-based force generation. Wiley Interdiscip. Rev. Nanomed. Nanobiotechnol.. 2017 doi:10.1002/wnan.1428 PubMed
102. Akhmanova A., Steinmetz M.O.. Control of microtubule organization and dynamics: Two ends in the limelight. Nat. Rev. Mol. Cell Biol.. 2015 doi:10.1038/nrm4084 PubMed
103. van Haren J., Wittmann T.. Microtubule Plus End Dynamics—Do We Know How Microtubules Grow?: Cells boost microtubule growth by promoting distinct structural transitions at growing microtubule ends. Bioessays. 2019 doi:10.1002/bies.201800194 PubMed
104. MacRae T.H.. Towards an understanding of microtubule function and cell organization: An overview. Biochem. Cell Biol.. 1992 doi:10.1139/o92-131 PubMed
105. Zwetsloot A.J., Tut G., Straube A.. Measuring microtubule dynamics. Essays Biochem.. 2018 doi:10.1042/ebc20180035 PubMed
106. Sept D., Baker N.A., McCammon J.A.. The physical basis of microtubule structure and stability. Protein Sci.. 2003 doi:10.1110/ps.03187503 PubMed
107. McIntosh J.R., Grishchuk E.L., West R.R.. Chromosome-microtubule interactions during mitosis. Annu. Rev. Cell Dev. Biol.. 2002 doi:10.1146/annurev.cellbio.18.032002.132412 PubMed
108. Xiao H., Verdier-Pinard P., Fernandez-Fuentes N., Burd B., Angeletti R., Fiser A. et al. Insights into the mechanism of microtubule stabilization by Taxol. Proc. Natl. Acad. Sci. USA. 2006 doi:10.1073/pnas.0603704103 PubMed
109. Müsch A.. Microtubule Organization and Function in Epithelial Cells. Traffic. 2004 doi:10.1111/j.1600-0854.2003.00149.x PubMed
110. Lai S.-L., Chien A.J., Moon R.T.. Wnt/Fz signaling and the cytoskeleton: Potential roles in tumorigenesis. Cell Res.. 2009 doi:10.1038/cr.2009.41 PubMed
111. Hoffman L., Jensen C.C., Yoshigi M., Beckerle M.. Mechanical signals activate p38 MAPK pathway-dependent reinforcement of actin via mechanosensitive HspB1. Mol. Biol. Cell. 2017 doi:10.1091/mbc.e17-02-0087 PubMed
112. Sauzeau V., Le Jeune H., Cario-Toumaniantz C., Smolenski A., Lohmann S.M., Bertoglio J. et al. Cyclic GMP-dependent Protein Kinase Signaling Pathway Inhibits RhoA-induced Ca2+ Sensitization of Contraction in Vascular Smooth Muscle*. J. Biol. Chem.. 2000 doi:10.1074/jbc.M000753200 PubMed
113. Zou L., Zhang J., Han J., Li W., Su F., Xu X. et al. cGMP interacts with tropomyosin and downregulates actin-tropomyosin-myosin complex interaction. Respir. Res.. 2018 doi:10.1186/s12931-018-0903-z PubMed
114. James R.G., Conrad W.H., Moon R.T.. Beta-catenin-independent Wnt pathways: Signals, core proteins, and effectors. Methods Mol. Biol.. 2008 doi:10.1007/978-1-59745-249-6_10 PubMed
115. Roarty K., Pfefferle A.D., Creighton C.J., Perou C.M., Rosen J.M.. Ror2-mediated alternative Wnt signaling regulates cell fate and adhesion during mammary tumor progression. Oncogene. 2017 doi:10.1038/onc.2017.206 PubMed
116. Beurel E., Grieco S.F., Jope R.S.. Glycogen synthase kinase-3 (GSK3): Regulation, actions, and diseases. Pharmacol. Ther.. 2015 doi:10.1016/j.pharmthera.2014.11.016 PubMed
117. Deng S., Leong H.C., Datta A., Gopal V., Kumar A.P., Yap C.T.. PI3K/AKT Signaling Tips the Balance of Cytoskeletal Forces for Cancer Progression. Cancers. 2022 doi:10.3390/cancers14071652 PubMed
118. May-Simera H.L., Kelley M.W.. Cilia, Wnt signaling, and the cytoskeleton. Cilia. 2012 doi:10.1186/2046-2530-1-7 PubMed
119. Shapiro L.. The multi-talented beta-catenin makes its first appearance. Structure. 1997 doi:10.1016/S0969-2126(97)00278-5 PubMed
120. Shi J., Wei L.. Rho Kinases in Embryonic Development and Stem Cell Research. Arch. Immunol. Ther. Exp.. 2022 doi:10.1007/s00005-022-00642-z PubMed
121. Tang L., Dai F., Liu Y., Yu X., Huang C., Wang Y. et al. RhoA/ROCK signaling regulates smooth muscle phenotypic modulation and vascular remodeling via the JNK pathway and vimentin cytoskeleton. Pharmacol. Res.. 2018 doi:10.1016/j.phrs.2018.05.011 PubMed
122. Schofield A.V., Bernard O.. Rho-associated coiled-coil kinase (ROCK) signaling and disease. Crit. Rev. Biochem. Mol. Biol.. 2013 doi:10.3109/10409238.2013.786671 PubMed
123. Sit S.-T., Manser E.. Rho GTPases and their role in organizing the actin cytoskeleton. J. Cell Sci.. 2011 doi:10.1242/jcs.064964 PubMed
124. Sarasa-Renedo A., Tunç-Civelek V., Chiquet M.. Role of RhoA/ROCK-dependent actin contractility in the induction of tenascin-C by cyclic tensile strain. Exp. Cell Res.. 2006 doi:10.1016/j.yexcr.2005.12.025 PubMed
125. Woods A., Wang G., Beier F.. RhoA/ROCK signaling regulates Sox9 expression and actin organization during chondrogenesis. J. Biol. Chem.. 2005 doi:10.1074/jbc.M409158200 PubMed
126. Da Silva J.S., Medina M., Zuliani C., Di Nardo A., Witke W., Dotti C.G.. RhoA/ROCK regulation of neuritogenesis via profilin IIa-mediated control of actin stability. J. Cell Biol.. 2003 doi:10.1083/jcb.200304021 PubMed
127. Amano M., Fukata Y., Kaibuchi K.. Regulation and functions of Rho-associated kinase. Exp. Cell Res.. 2000 doi:10.1006/excr.2000.5046 PubMed
128. Yang L., Tang L., Dai F., Meng G., Yin R., Xu X. et al. Raf-1/CK2 and RhoA/ROCK signaling promote TNF-α-mediated endothelial apoptosis via regulating vimentin cytoskeleton. Toxicology. 2017 doi:10.1016/j.tox.2017.07.010 PubMed
129. Lei S., Tian Y.P., Xiao W.D., Li S., Rao X.C., Zhang J.L. et al. ROCK is Involved in Vimentin Phosphorylation and Rearrangement Induced by Dengue Virus. Cell Biochem. Biophys.. 2013 doi:10.1007/s12013-013-9665-x PubMed
130. Hirose M., Ishizaki T., Watanabe N., Uehata M., Kranenburg O., Moolenaar W.H. et al. Molecular dissection of the Rho-associated protein kinase (p160ROCK)- regulated neurite remodeling in neuroblastoma N1E-115 cells. J. Cell Biol.. 1998 doi:10.1083/jcb.141.7.1625 PubMed
131. Becker K.N., Pettee K.M., Sugrue A., Reinard K.A., Schroeder J.L., Eisenmann K.M.. The Cytoskeleton Effectors Rho-Kinase (ROCK) and Mammalian Diaphanous-Related (mDia) Formin Have Dynamic Roles in Tumor Microtube Formation in Invasive Glioblastoma Cells. Cells. 2022 doi:10.3390/cells11091559 PubMed
132. Heng Y.W., Lim H.H., Mina T., Utomo P., Zhong S., Lim C.T. et al. TPPP acts downstream of RhoA-ROCK-LIMK2 to regulate astral microtubule organization and spindle orientation. J. Cell Sci.. 2012 doi:10.1242/jcs.096818 PubMed
133. Fonseca A.V., Freund D., Bornhäuser M., Corbeil D.. Polarization and migration of hematopoietic stem and progenitor cells rely on the RhoA/ROCK I pathway and an active reorganization of the microtubule network. J. Biol. Chem.. 2010 doi:10.1074/jbc.M110.145037 PubMed
134. Takesono A., Heasman S.J., Wojciak-Stothard B., Garg R., Ridley A.J.. Microtubules regulate migratory polarity through Rho/ ROCK signaling in T cells. PLoS ONE. 2010 doi:10.1371/journal.pone.0008774 PubMed
135. Han D., Sun J., Fan D., Zhang C., Du S., Zhang W.. Simvastatin ameliorates oxygen glucose deprivation/reoxygenation-induced pulmonary endothelial barrier dysfunction by restoring cell-cell junctions and actin cytoskeleton dynamics via the PI3K/Akt signaling pathway. Am. J. Transl. Res.. 2020 PubMed
136. Lien E.C., Dibble C.C., Toker A.. PI3K signaling in cancer: Beyond AKT. Curr. Opin. Cell Biol.. 2017 doi:10.1016/j.ceb.2017.02.007 PubMed
137. Kakinuma N., Roy B.C., Zhu Y., Wang Y., Kiyama R.. Kank regulates RhoA-dependent formation of actin stress fibers and cell migration via 14-3-3 in PI3K-Akt signaling. J. Cell Biol.. 2008 doi:10.1083/jcb.200707022 PubMed
138. Qian Y., Zhong X., Flynn D.C., Zheng J.Z., Qiao M., Wu C. et al. ILK mediates actin filament rearrangements and cell migration and invasion through PI3K/Akt/Rac1 signaling. Oncogene. 2005 doi:10.1038/sj.onc.1208525 PubMed
139. Qian Y., Corum L., Meng Q., Blenis J., Zheng J.Z., Shi X. et al. PI3K induced actin filament remodeling through Akt and p70S6K1: Implication of essential role in cell migration. Am. J. Physiol. Cell Physiol.. 2004 doi:10.1152/ajpcell.00142.2003 PubMed
140. Krasilnikov M.A.. Phosphatidylinositol-3 kinase dependent pathways: The role in control of cell growth, survival, and malignant transformation. Biochemistry. 2000 PubMed
141. Roux A., Loranger A., Lavoie J.N., Marceau N.. Keratin 8/18 regulation of insulin receptor signaling and trafficking in hepatocytes through a concerted phosphoinositide-dependent Akt and Rab5 modulation. FASEB J.. 2017 doi:10.1096/fj.201700036R PubMed
142. Wang R.C., Wei Y., An Z., Zou Z., Xiao G., Bhagat G. et al. Akt-mediated regulation of autophagy and tumorigenesis through Beclin 1 phosphorylation. Science. 2012 doi:10.1126/science.1225967 PubMed
143. Kong L., Schäfer G., Bu H., Zhang Y., Zhang Y., Klocker H.. Lamin A/C protein is overexpressed in tissue-invading prostate cancer and promotes prostate cancer cell growth, migration and invasion through the PI3K/AKT/PTEN pathway. Carcinogenesis. 2012 doi:10.1093/carcin/bgs022 PubMed
144. Tseng Y.H., Yang C.C., Lin S.C., Cheng C.C., Lin S.H., Liu C.J. et al. Areca nut extract upregulates vimentin by activating PI3K/AKT signaling in oral carcinoma. J. Oral Pathol. Med.. 2011 doi:10.1111/j.1600-0714.2010.00978.x PubMed
145. Chakrabarty S., Nag D., Ganguli A., Das A., Ghosh Dastidar D., Chakrabarti G.. Theaflavin and epigallocatechin-3-gallate synergistically induce apoptosis through inhibition of PI3K/Akt signaling upon depolymerizing microtubules in HeLa cells. J. Cell. Biochem.. 2019 doi:10.1002/jcb.27886 PubMed
146. Fu Y.F., Liu X., Gao M., Zhang Y.N., Liu J.. Endoplasmic reticulum stress induces autophagy and apoptosis while inhibiting proliferation and drug resistance in multiple myeloma through the PI3K/Akt/mTOR signaling pathway. Oncotarget. 2017 doi:10.18632/oncotarget.17862 PubMed
147. Kitagishi Y., Nakanishi A., Ogura Y., Matsuda S.. Dietary regulation of PI3K/AKT/GSK-3β pathway in Alzheimer’s disease. Alzheimers Res. Ther.. 2014 doi:10.1186/alzrt265 PubMed
148. Onishi K., Higuchi M., Asakura T., Masuyama N., Gotoh Y.. The PI3K-Akt pathway promotes microtubule stabilization in migrating fibroblasts. Genes Cells. 2007 doi:10.1111/j.1365-2443.2007.01071.x PubMed
149. Fujiwara Y., Hosokawa Y., Watanabe K., Tanimura S., Ozaki K.I., Kohno M.. Blockade of the phosphatidylinositol-3-kinase-Akt signaling pathway enhances the induction of apoptosis by microtubule-destabilizing agents in tumor cells in which the pathway is constitutively activated. Mol. Cancer Ther.. 2007 doi:10.1158/1535-7163.MCT-06-0639 PubMed
150. Zhang J., Hu Q., Jiang X., Wang S., Zhou X., Lu Y. et al. Actin Alpha 2 Downregulation Inhibits Neural Stem Cell Proliferation and Differentiation into Neurons through Canonical Wnt/β-Catenin Signaling Pathway. Oxid. Med. Cell. Longev.. 2022 doi:10.1155/2022/7486726 PubMed
151. Galli C., Piemontese M., Lumetti S., Ravanetti F., MacAluso G.M., Passeri G.. Actin cytoskeleton controls activation of Wnt/β-catenin signaling in mesenchymal cells on implant surfaces with different topographies. Acta Biomater.. 2012 doi:10.1016/j.actbio.2012.04.043 PubMed
152. Lehmann M., Hu Q., Hu Y., Hafner K., Costa R., van den Berg A. et al. Chronic WNT/β-catenin signaling induces cellular senescence in lung epithelial cells. Cell. Signal.. 2020 doi:10.1016/j.cellsig.2020.109588 PubMed
153. Tian Z., Zhang X., Zhao Z., Zhang F., Deng T.. The Wnt/β-catenin signaling pathway affects the distribution of cytoskeletal proteins in Aβ treated PC12 cells. J. Integr. Neurosci.. 2019 doi:10.31083/j.jin.2019.03.168 PubMed
154. Bermeo S., Vidal C., Zhou H., Duque G.. Lamin A/C Acts as an Essential Factor in Mesenchymal Stem Cell Differentiation Through the Regulation of the Dynamics of the Wnt/β-Catenin Pathway. J. Cell. Biochem.. 2015 doi:10.1002/jcb.25185 PubMed
155. Prasad C.P., Mirza S., Sharma G., Prashad R., DattaGupta S., Rath G. et al. Epigenetic alterations of CDH1 and APC genes: Relationship with activation of Wnt/β-catenin Pathway in invasive ductal carcinoma of breast. Life Sci.. 2008 doi:10.1016/j.lfs.2008.06.019 PubMed
156. Bierie B., Nozawa M., Renou J.P., Shillingford J.M., Morgan F., Oka T. et al. Activation of β-catenin in prostate epithelium induces hyperplasias and squamous transdifferentiation. Oncogene. 2003 doi:10.1038/sj.onc.1206426 PubMed
157. Puri D., Ponniah K., Biswas K., Basu A., Dey S., Lundquist E.A. et al. Wnt signaling establishes the microtubule polarity in neurons through regulation of kinesin-13. J. Cell Biol.. 2021 doi:10.1083/jcb.202005080 PubMed
158. Ou D., Chen L., He J., Rong Z., Gao J., Li Z. et al. CDK11 negatively regulates Wnt/β-catenin signaling in the endosomal compartment by affecting microtubule stability. Cancer Biol. Med.. 2020 doi:10.20892/j.issn.2095-3941.2019.0229 PubMed
159. Huang P., Senga T., Hamaguchi M.. A novel role of phospho-β-catenin in microtubule regrowth at centrosome. Oncogene. 2007 doi:10.1038/sj.onc.1210217 PubMed
160. Ciani L., Krylova O., Smalley M.J., Dale T.C., Salinas P.C.. A divergent canonical WNT-signaling pathway regulates microtubule dynamics: Dishevelled signals locally to stabilize microtubules. J. Cell Biol.. 2004 doi:10.1083/jcb.200309096 PubMed
161. Peifer M., Polakis P.. Wnt signaling in oncogenesis and embryogenesis—A look outside the nucleus. Science. 2000 doi:10.1126/science.287.5458.1606 PubMed
162. Hajka D., Budziak B., Pietras Ł., Duda P., McCubrey J.A., Gizak A.. GSK3 as a Regulator of Cytoskeleton Architecture: Consequences for Health and Disease. Cells. 2021 doi:10.3390/cells10082092 PubMed
163. Yoshino Y., Suzuki M., Takahashi H., Ishioka C.. Inhibition of invasion by glycogen synthase kinase-3 beta inhibitors through dysregulation of actin re-organisation via down-regulation of WAVE2. Biochem. Biophys. Res. Commun.. 2015 doi:10.1016/j.bbrc.2015.06.142 PubMed
164. Watanabe T., Noritake J., Kakeno M., Matsui T., Harada T., Wang S. et al. Phosphorylation of CLASP2 by GSK-3β regulates its interaction with IQGAP1, EB1 and microtubules. J. Cell Sci.. 2009 doi:10.1242/jcs.046649 PubMed
165. Sun T., Rodriguez M., Kim L.. Glycogen synthase kinase 3 in the world of cell migration. Dev. Growth Differ.. 2009 doi:10.1111/j.1440-169X.2009.01141.x PubMed
166. Vaidya R.J., Ray R.M., Johnson L.R.. Akt-mediated GSK-3β inhibition prevents migration of polyamine-depleted intestinal epithelial cells via Rac1. Cell. Mol. Life Sci.. 2006 doi:10.1007/s00018-006-6379-x PubMed
167. Sen S., Lagas S., Roy A., Kumar H.. Cytoskeleton saga: Its regulation in normal physiology and modulation in neurodegenerative disorders. Eur. J. Pharmacol.. 2022 doi:10.1016/j.ejphar.2022.175001 PubMed
168. Lee W.C., Kan D., Chen Y.Y., Han S.K., Lu K.S., Chien C.L.. Suppression of extensive neurofilament phosphorylation rescues α-internexin/peripherin-overexpressing PC12 cells from neuronal cell death. PLoS ONE. 2012 doi:10.1371/journal.pone.0043883 PubMed
169. Kim J.Y., Kim Y.M., Yang C.H., Cho S.K., Lee J.W., Cho M.. Functional regulation of Slug/Snail2 is dependent on GSK-3β-mediated phosphorylation. FEBS J.. 2012 doi:10.1111/j.1742-4658.2012.08674.x PubMed
170. Sasaki T., Taoka M., Ishiguro K., Uchida A., Saito T., Toshiaki Isobe D. et al. In vivo and in vitro phosphorylation at Ser-493 in the glutamate (E)-segment of neurofilament-H subunit by glycogen synthase kinase 3β. J. Biol. Chem.. 2002 doi:10.1074/jbc.M206674200 PubMed
171. Guidato S., Tsai L.H., Woodgett J., Miller C.C.J.. Differential cellular phosphorylation of neurofilament heavy side-arms by glycogen synthase kinase-3 and cydin-dependent kinase-5. J. Neurochem.. 1996 doi:10.1046/j.1471-4159.1996.66041698.x PubMed
172. Fumoto K., Hoogenraad C.C., Kikuchi A.. GSK-3β-regulated interaction of BICD with dynein is involved in microtubule anchorage at centrosome. EMBO J.. 2006 doi:10.1038/sj.emboj.7601459 PubMed
173. Grimes C.A., Jope R.S.. The multifaceted roles of glycogen synthase kinase 3β in cellular signaling. Prog. Neurobiol.. 2001 doi:10.1016/S0301-0082(01)00011-9 PubMed
174. Joe S.Y., Yang S.G., Lee J.H., Park H.J., Koo D.B.. Stabilization of F-Actin Cytoskeleton by Paclitaxel Improves the Blastocyst Developmental Competence through P38 MAPK Activity in Porcine Embryos. Biomedicines. 2022 doi:10.3390/biomedicines10081867 PubMed
175. Yang C., Patel K., Harding P., Sorokin A., Glass Ii W.F.. Regulation of TGF-β1/MAPK-mediated PAI-1 gene expression by the actin cytoskeleton in human mesangial cells. Exp. Cell Res.. 2007 doi:10.1016/j.yexcr.2007.01.011 PubMed
176. Fujiwara M., Jin E., Ghazizadeh M., Kawanami O.. Activation of PAR4 induces a distinct actin fiber formation via p38 MAPK in human lung endothelial cells. J. Histochem. Cytochem.. 2005 doi:10.1369/jhc.4A6592.2005 PubMed
177. Paliga A.J.M., Natale D.R., Watson A.J.. p38 mitogen-activated protein kinase (MAPK) first regulates filamentous actin at the 8-16-cell stage during preimplantation development. Biol. Cell. 2005 doi:10.1042/BC20040146 PubMed
178. Tania M., Khan M.A., Fu J.. Epithelial to mesenchymal transition inducing transcription factors and metastatic cancer. Tumour Biol.. 2014 doi:10.1007/s13277-014-2163-y PubMed
179. Wang P.B., Chen Y., Ding G.R., Du H.W., Fan H.Y.. Keratin 18 induces proliferation, migration, and invasion in gastric cancer via the MAPK signalling pathway. Clin. Exp. Pharmacol. Physiol.. 2021 doi:10.1111/1440-1681.13401 PubMed
180. Wöll S., Windoffer R., Leube R.E.. p38 MAPK-dependent shaping of the keratin cytoskeleton in cultured cells. J. Cell Biol.. 2007 doi:10.1083/jcb.200703174 PubMed
181. Schechter R., Yanovitch T., Abboud M., Johnson III G., Gaskins J.. Effects of brain endogenous insulin on neurofilament and MAPK in fetal rat neuron cell cultures. Brain Res.. 1998 doi:10.1016/S0006-8993(98)00842-7 PubMed
182. Cheng T.J., Lai Y.K.. Identification of mitogen-activated protein kinase-activated protein kinase-2 as a vimentin kinase activated by okadaic acid in 9L rat brain tumor cells. J. Cell. Biochem.. 1998 doi:10.1002/(SICI)1097-4644(19981101)71:2<169::AID-JCB3>3.0.CO;2-T PubMed
183. Li L., Hu J., He T., Zhang Q., Yang X., Lan X. et al. P38/MAPK contributes to endothelial barrier dysfunction via MAP4 phosphorylation-dependent microtubule disassembly in inflammation-induced acute lung injury. Sci. Rep.. 2015 doi:10.1038/srep08895 PubMed
184. Hu J.Y., Chu Z.G., Han J., Dang Y.M., Yan H., Zhang Q. et al. The p38/MAPK pathway regulates microtubule polymerization through phosphorylation of MAP4 and Op18 in hypoxic cells. Cell. Mol. Life Sci.. 2010 doi:10.1007/s00018-009-0187-z PubMed
185. Lee S.E., Kim J.H., Kim N.H.. Inactivation of MAPK affects centrosome assembly, but not actin filament assembly, in mouse oocytes maturing in vitro. Mol. Reprod. Dev.. 2007 doi:10.1002/mrd.20695 PubMed
186. Fan M., Chambers T.C.. Role of mitogen-activated protein kinases in the response of tumor cells to chemotherapy. Drug Resist. Updates. 2001 doi:10.1054/drup.2001.0214 PubMed
187. Reszka A.A., Seger R., Diltz C.D., Krebs E.G., Fischer E.H.. Association of mitogen-activated protein kinase with the microtubule cytoskeleton. Proc. Natl. Acad. Sci. USA. 1995 doi:10.1073/pnas.92.19.8881 PubMed
188. Butt E., Gambaryan S., Göttfert N., Galler A., Marcus K., Meyer H.E.. Actin binding of human LIM and SH3 protein is regulated by cGMP- and cAMP-dependent protein kinase phosphorylation on serine 146. J. Biol. Chem.. 2003 doi:10.1074/jbc.M209009200 PubMed
189. Butt E., Immler D., Meyer H.E., Kotlyarov A., Laaß K., Gaestel M.. Heat shock protein 27 is a substrate of cGMP-dependent protein kinase in intact human platelets. Phosphorylation-induced actin polymerization caused by HSP27 mutants. J. Biol. Chem.. 2001 doi:10.1074/jbc.M009234200 PubMed
190. Sandau K.B., Gantner F., Brüne B.. Nitric oxide-induced F-actin disassembly is mediated via cGMP, cAMP, and protein kinase A activation in rat mesangial cells. Exp. Cell Res.. 2001 doi:10.1006/excr.2001.5378 PubMed
191. Pryzwansky K.B., Wyatt T.A., Lincoln T.M.. Cyclic guanosine monophosphate-dependent protein kinase is targeted to intermediate filaments and phosphorylates vimentin in A23187-stimulated human neutrophils. Blood. 1995 doi:10.1182/blood.V85.1.222.bloodjournal851222 PubMed
192. MacMillan-Crow L.A., Lincoln T.M.. High-affinity binding and localization of the cyclic GMP-dependent protein kinase with the intermediate filament protein vimentin. Biochemistry. 1994 doi:10.1021/bi00192a007 PubMed
193. Wyatt T.A., Lincoln T.M., Pryzwansky K.B.. Regulation of human neutrophil degranulation by LY-83583 and L-arginine: Role of cGMP-dependent protein kinase. Am. J. Physiol. Cell Physiol.. 1993 doi:10.1152/ajpcell.1993.265.1.C201 PubMed
194. Wyatt T.A., Lincoln T.M., Pryzwansky K.B.. Vimentin is transiently co-localized with and phosphorylated by cyclic GMP-dependent protein kinase in formyl-peptide-stimulated neutrophils. J. Biol. Chem.. 1991 doi:10.1016/S0021-9258(18)54851-1 PubMed
195. Xia C., Nguyen M., Garrison A.K., Zhao Z., Wang Z., Sutherland C. et al. CNP/cGMP signaling regulates axon branching and growth by modulating microtubule polymerization. Dev. Neurobiol.. 2013 doi:10.1002/dneu.22078 PubMed
196. Gong K., Xing D., Li P., Hilgers R.H., Hage F.G., Oparil S. et al. cGMP inhibits TGF-beta signaling by sequestering Smad3 with cytosolic beta2-tubulin in pulmonary artery smooth muscle cells. Mol. Endocrinol.. 2011 doi:10.1210/me.2011-1009 PubMed
197. Guo J., Wenk M.R., Pellegrini L., Onofri F., Benfenati F., De Camilli P.. Phosphatidylinositol 4-kinase type IIα is responsible for the phosphatidylinositol 4-kinase activity associated with synaptic vesicles. Proc. Natl. Acad. Sci. USA. 2003 doi:10.1073/pnas.0230488100 PubMed
198. Michalczyk I., Sikorski A.F., Kotula L., Junghans R.P., Dubielecka P.M.. The emerging role of protein kinase Cθ in cytoskeletal signaling. J. Leukoc. Biol.. 2013 doi:10.1189/jlb.0812371 PubMed
199. Liao J.K., Seto M., Noma K.. Rho kinase (ROCK) inhibitors. J. Cardiovasc. Pharmacol.. 2007 doi:10.1097/FJC.0b013e318070d1bd PubMed
200. Apodaca G.. Endocytic traffic in polarized epithelial cells: Role of the actin and microtubule cytoskeleton. Traffic. 2001 doi:10.1034/j.1600-0854.2001.020301.x PubMed
201. Ridley A.J.. Rho proteins: Linking signaling with membrane trafficking. Traffic. 2001 doi:10.1034/j.1600-0854.2001.002005303.x PubMed
202. Etienne-Manneville S., Hall A.. Rho GTPases in cell biology. Nature. 2002 doi:10.1038/nature01148 PubMed
203. Jaffe A.B., Hall A.. Rho GTPases: Biochemistry and biology. Annu. Rev. Cell Dev. Biol.. 2005 doi:10.1146/annurev.cellbio.21.020604.150721 PubMed
204. Burridge K., Wennerberg K.. Rho and Rac take center stage. Cell. 2004 doi:10.1016/S0092-8674(04)00003-0 PubMed
205. Bishop A.L., Hall A.. Rho GTPases and their effector proteins. Biochem. J.. 2000 doi:10.1042/bj3480241 PubMed
206. Narumiya S., Tanji M., Ishizaki T.. Rho signaling, ROCK and mDia1, in transformation, metastasis and invasion. Cancer Metastasis Rev.. 2009 doi:10.1007/s10555-008-9170-7 PubMed
207. Wittinghofer A., Vetter I.R.. Structure-function relationships of the G domain, a canonical switch motif. Annu. Rev. Biochem.. 2011 doi:10.1146/annurev-biochem-062708-134043 PubMed
208. Zheng Y.. Dbl family guanine nucleotide exchange factors. Trends Biochem. Sci.. 2001 doi:10.1016/S0968-0004(01)01973-9 PubMed
209. Hodge R.G., Ridley A.J.. Regulating Rho GTPases and their regulators. Nat. Rev. Mol. Cell Biol.. 2016 doi:10.1038/nrm.2016.67 PubMed
210. Jiu Y., Peränen J., Schaible N., Cheng F., Eriksson J.E., Krishnan R. et al. Vimentin intermediate filaments control actin stress fiber assembly through GEF-H1 and RhoA. J. Cell Sci.. 2017 doi:10.1242/jcs.196881 PubMed
211. Inaba H., Yamakawa D., Tomono Y., Enomoto A., Mii S., Kasahara K. et al. Regulation of keratin 5/14 intermediate filaments by CDK1, Aurora-B, and Rho-kinase. Biochem. Biophys. Res. Commun.. 2018 doi:10.1016/j.bbrc.2018.03.016 PubMed
212. Dharmawardhane S.. Rho family GTPases in cancer. Cancers. 2021 doi:10.3390/cancers13061271 PubMed
213. Ellenbroek S.I.J., Collard J.G.. Rho GTPases: Functions and association with cancer. Clin. Exp. Metastasis. 2007 doi:10.1007/s10585-007-9119-1 PubMed
214. El-Sibai M., Pertz O., Pang H., Yip S.C., Lorenz M., Symons M. et al. RhoA/ROCK-mediated switching between Cdc42- and Rac1-dependent protrusion in MTLn3 carcinoma cells. Exp. Cell Res.. 2008 doi:10.1016/j.yexcr.2008.01.016 PubMed
215. Bustelo X.R., Sauzeau V., Berenjeno I.M.. GTP-binding proteins of the Rho/Rac family: Regulation, effectors and functions in vivo. Bioessays. 2007 doi:10.1002/bies.20558 PubMed
216. Pinyol R., Haeckel A., Ritter A., Qualmann B., Kessels M.M.. Regulation of N-WASP and the Arp2/3 complex by Abp1 controls neuronal morphology. PLoS ONE. 2007 doi:10.1371/journal.pone.0000400 PubMed
217. Koka S., Neudauer C.L., Li X., Lewis R.E., McCarthy J.B., Westendorf J.J.. The formin-homology-domain-containing protein FHOD1 enhances cell migration. J. Cell Sci.. 2003 doi:10.1242/jcs.00386 PubMed
218. Royal I., Lamarche-Vane N., Lamorte L., Kaibuchi K., Park M.. Activation of cdc42, rac, PAK, and rho-kinase in response to hepatocyte growth factor differentially regulates epithelial cell colony spreading and dissociation. Mol. Biol. Cell. 2000 doi:10.1091/mbc.11.5.1709 PubMed
219. Watanabe N., Madaule P., Reid T., Ishizaki T., Watanabe G., Kakizuka A. et al. p140mDia, a mammalian homolog of Drosophila diaphanous, is a target protein for Rho small GTPase and is a ligand for profilin. EMBO J.. 1997 doi:10.1093/emboj/16.11.3044 PubMed
220. ROCK1, Rho Associated Coiled-Coil Containing Protein Kinase 1 [Homo Sapiens (Human)]. 2022
221. ROCK2, Rho Associated Coiled-Coil Containing Protein Kinase 1 [Homo Sapiens (Human)]. 2022
222. Totsukawa G., Yamakita Y., Yamashiro S., Hartshorne D.J., Sasaki Y., Matsumura F.. Distinct roles of ROCK (Rho-kinase) and MLCK in spatial regulation of MLC phosphorylation for assembly of stress fibers and focal adhesions in 3T3 fibroblasts. J. Cell Biol.. 2000 doi:10.1083/jcb.150.4.797 PubMed
223. Ohashi K., Nagata K., Maekawa M., Ishizaki T., Narumiya S., Mizuno K.. Rho-associated kinase ROCK activates LIM-kinase 1 by phosphorylation at threonine 508 within the activation loop. J. Biol. Chem.. 2000 doi:10.1074/jbc.275.5.3577 PubMed
224. Qiao Y.N., He W.Q., Chen C.P., Zhang C.H., Zhao W., Wang P. et al. Myosin phosphatase target subunit 1 (MYPT1) regulates the contraction and relaxation of vascular smooth muscle and maintains blood pressure. J. Biol. Chem.. 2014 doi:10.1074/jbc.M113.525444 PubMed
225. Lee J.H., Katakai T., Hara T., Gonda H., Sugai M., Shimizu A.. Roles of p-ERM and Rho-ROCK signaling in lymphocyte polarity and uropod formation. J. Cell Biol.. 2004 doi:10.1083/jcb.200403091 PubMed
226. Fukata Y., Oshiro N., Kinoshita N., Kawano Y., Matsuoka Y., Bennett V. et al. Phosphorylation of adducin by Rho-kinase plays a crucial role in cell motility. J. Cell Biol.. 1999 doi:10.1083/jcb.145.2.347 PubMed
227. Shibukawa Y., Yamazaki N., Daimon E., Wada Y.. Rock-dependent calponin 3 phosphorylation regulates myoblast fusion. Exp. Cell Res.. 2013 doi:10.1016/j.yexcr.2012.12.022 PubMed
228. Ikenoya M., Hidaka H., Hosoya T., Suzuki M., Yamamoto N., Sasaki Y.. Inhibition of rho-kinase-induced myristoylated alanine-rich C kinase substrate (MARCKS) phosphorylation in human neuronal cells by H-1152, a novel and specific Rho-kinase inhibitor. J. Neurochem.. 2002 doi:10.1046/j.1471-4159.2002.00801.x PubMed
229. Izawa T., Fukata Y., Kimura T., Iwamatsu A., Dohi K., Kaibuchi K.. Elongation factor-1 alpha is a novel substrate of rho-associated kinase. Biochem. Biophys. Res. Commun.. 2000 doi:10.1006/bbrc.2000.3772 PubMed
230. Vahebi S., Kobayashi T., Warren C.M., de Tombe P.P., Solaro R.J.. Functional effects of rho-kinase-dependent phosphorylation of specific sites on cardiac troponin. Circ. Res.. 2005 doi:10.1161/01.RES.0000162457.56568.7d PubMed
231. Shao J., Welch W.J., Diprospero N.A., Diamond M.I.. Phosphorylation of profilin by ROCK1 regulates polyglutamine aggregation. Mol. Cell. Biol.. 2008 doi:10.1128/MCB.00079-08 PubMed
232. Amano M., Ito M., Kimura K., Fukata Y., Chihara K., Nakano T. et al. Phosphorylation and activation of myosin by Rho-associated kinase (Rho-kinase). J. Biol. Chem.. 1996 doi:10.1074/jbc.271.34.20246 PubMed
233. Shi J., Wu X., Surma M., Vemula S., Zhang L., Yang Y. et al. Distinct roles for ROCK1 and ROCK2 in the regulation of cell detachment. Cell Death Dis.. 2013 doi:10.1038/cddis.2013.10 PubMed
234. Wang J., Liu X.-H., Yang Z.-J., Xie B., Zhong Y.-S.. The effect of ROCK-1 activity change on the adhesive and invasive ability of Y79 retinoblastoma cells. BMC Cancer. 2014 doi:10.1186/1471-2407-14-89 PubMed
235. Rochelle T., Daubon T., Van Troys M., Harnois T., Waterschoot D., Ampe C. et al. p210bcr-abl induces amoeboid motility by recruiting ADF/destrin through RhoA/ROCK1. FASEB J.. 2013 doi:10.1096/fj.12-205112 PubMed
236. Yasui Y., Amano M., Nagata K., Inagaki N., Nakamura H., Saya H. et al. Roles of Rho-associated kinase in cytokinesis; mutations in Rho-associated kinase phosphorylation sites impair cytokinetic segregation of glial filaments. J. Cell Biol.. 1998 doi:10.1083/jcb.143.5.1249 PubMed
237. Hashimoto R., Nakamura Y., Goto H., Wada Y., Sakoda S., Kaibuchi K. et al. Domain- and site-specific phosphorylation of bovine NF-L by Rho-associated kinase. Biochem. Biophys. Res. Commun.. 1998 doi:10.1006/bbrc.1998.8446 PubMed
238. Goto H., Kosako H., Inagaki M.. Regulation of intermediate filament organization during cytokinesis: Possible roles of Rho-associated kinase. Microsc. Res. Tech.. 2000 doi:10.1002/(SICI)1097-0029(20000415)49:2<173::AID-JEMT10>3.0.CO;2-A PubMed
239. Bordeleau F., Myrand Lapierre M.E., Sheng Y., Marceau N.. Keratin 8/18 regulation of cell stiffness-extracellular matrix interplay through modulation of Rho-mediated actin cytoskeleton dynamics. PLoS ONE. 2012 doi:10.1371/journal.pone.0038780 PubMed
240. Sin W.C., Chen X.Q., Leung T., Lim L.. RhoA-binding kinase alpha translocation is facilitated by the collapse of the vimentin intermediate filament network. Mol. Cell. Biol.. 1998 doi:10.1128/MCB.18.11.6325 PubMed
241. Amano M., Kaneko T., Maeda A., Nakayama M., Ito M., Yamauchi T. et al. Identification of Tau and MAP2 as novel substrates of Rho-kinase and myosin phosphatase. J. Neurochem.. 2003 doi:10.1046/j.1471-4159.2003.02054.x PubMed
242. Arimura N., Ménager C., Kawano Y., Yoshimura T., Kawabata S., Hattori A. et al. Phosphorylation by Rho kinase regulates CRMP-2 activity in growth cones. Mol. Cell. Biol.. 2005 doi:10.1128/MCB.25.22.9973-9984.2005 PubMed
243. Amano M., Tsumura Y., Taki K., Harada H., Mori K., Nishioka T. et al. A proteomic approach for comprehensively screening substrates of protein kinases such as Rho-kinase. PLoS ONE. 2010 doi:10.1371/journal.pone.0008704 PubMed
244. Zhang J., Dong X.P.. Dysfunction of microtubule-associated proteins of MAP2/tau family in Prion disease. Prion. 2012 doi:10.4161/pri.20677 PubMed
245. Lin P.C., Chan P.M., Hall C., Manser E.. Collapsin response mediator proteins (CRMPs) are a new class of microtubule-associated protein (MAP) that selectively interacts with assembled microtubules via a taxol-sensitive binding interaction. J. Biol. Chem.. 2011 doi:10.1074/jbc.M111.283580 PubMed
246. Schofield A.V., Steel R., Bernard O.. Rho-associated coiled-coil kinase (ROCK) protein controls microtubule dynamics in a novel signaling pathway that regulates cell migration. J. Biol. Chem.. 2012 doi:10.1074/jbc.M112.394965 PubMed
247. Huang C.H., Cheng J.C., Chen J.C., Tseng C.P.. Evaluation of the role of Disabled-2 in nerve growth factor-mediated neurite outgrowth and cellular signalling. Cell. Signal.. 2007 doi:10.1016/j.cellsig.2007.01.019 PubMed
248. OuYang C., Xie Y., Fu Q., Xu G.. SYNPO2 suppresses hypoxia-induced proliferation and migration of colorectal cancer cells by regulating YAP-KLF5 axis. Tissue Cell. 2021 doi:10.1016/j.tice.2021.101598 PubMed
249. Kai F., Tanner K., King C., Duncan R.. Myopodin isoforms alter the chemokinetic response of PC3 cells in response to different migration stimuli via differential effects on Rho-ROCK signaling pathways. Carcinogenesis. 2012 doi:10.1093/carcin/bgs268 PubMed
250. Maldonado H., Calderon C., Burgos-Bravo F., Kobler O., Zuschratter W., Ramirez O. et al. Astrocyte-to-neuron communication through integrin-engaged Thy-1/CBP/Csk/Src complex triggers neurite retraction via the RhoA/ROCK pathway. Biochim. Biophys. Acta Mol. Cell Res.. 2017 doi:10.1016/j.bbamcr.2016.11.006 PubMed
251. Fusella F., Seclì L., Busso E., Krepelova A., Moiso E., Rocca S. et al. The IKK/NF-κB signaling pathway requires Morgana to drive breast cancer metastasis. Nat. Commun.. 2017 doi:10.1038/s41467-017-01829-1 PubMed
252. Barry D.M., Koo Y., Norden P.R., Wylie L.A., Xu K., Wichaidit C. et al. Rasip1-Mediated Rho GTPase Signaling Regulates Blood Vessel Tubulogenesis via Nonmuscle Myosin II. Circ. Res.. 2016 doi:10.1161/CIRCRESAHA.116.309094 PubMed
253. de Kreuk B.J., Gingras A.R., Knight J.D., Liu J.J., Gingras A.C., Ginsberg M.H.. Heart of glass anchors Rasip1 at endothelial cell-cell junctions to support vascular integrity. Elife. 2016 doi:10.7554/eLife.11394 PubMed
254. Yamaguchi H., Kasa M., Amano M., Kaibuchi K., Hakoshima T.. Molecular Mechanism for the Regulation of Rho-Kinase by Dimerization and Its Inhibition by Fasudil. Structure. 2006 doi:10.1016/j.str.2005.11.024 PubMed
255. Wang J., Liu X., Zhong Y.. Rho/Rho-associated kinase pathway in glaucoma (Review). Int. J. Oncol.. 2013 doi:10.3892/ijo.2013.2100 PubMed
256. Feng Y., LoGrasso P.V., Defert O., Li R.. Rho Kinase (ROCK) Inhibitors and Their Therapeutic Potential. J. Med. Chem.. 2016 doi:10.1021/acs.jmedchem.5b00683 PubMed
257. Liu Y., Gray N.S.. Rational design of inhibitors that bind to inactive kinase conformations. Nat. Chem. Biol.. 2006 doi:10.1038/nchembio799 PubMed
258. Zhang T., Inesta-Vaquera F., Niepel M., Zhang J., Ficarro S.B., Machleidt T. et al. Discovery of potent and selective covalent inhibitors of JNK. Chem. Biol.. 2012 doi:10.1016/j.chembiol.2011.11.010 PubMed
259. Moshirfar M., Parker L., Birdsong O.C., Ronquillo Y.C., Hofstedt D., Shah T.J. et al. Use of Rho kinase Inhibitors in Ophthalmology: A Review of the Literature. Med. Hypothesis Discov. Innov. Ophthalmol.. 2018 PubMed
260. Zhao J., Zhou D., Guo J., Ren Z., Zhou L., Wang S. et al. Effect of fasudil hydrochloride, a protein kinase inhibitor, on cerebral vasospasm and delayed cerebral ischemic symptoms after aneurysmal subarachnoid hemorrhage. Neurol. Med. Chir.. 2006 doi:10.2176/nmc.46.421 PubMed
261. Satoh S., Ueda Y., Koyanagi M., Kadokami T., Sugano M., Yoshikawa Y. et al. Chronic inhibition of Rho kinase blunts the process of left ventricular hypertrophy leading to cardiac contractile dysfunction in hypertension-induced heart failure. J. Mol. Cell. Cardiol.. 2003 doi:10.1016/S0022-2828(02)00278-X PubMed
262. Rikitake Y., Oyama N., Wang C.Y., Noma K., Satoh M., Kim H.H. et al. Decreased perivascular fibrosis but not cardiac hypertrophy in ROCK1+/- haploinsufficient mice. Circulation. 2005 doi:10.1161/CIRCULATIONAHA.105.584623 PubMed
263. Hannan J.L., Albersen M., Kutlu O., Gratzke C., Stief C.G., Burnett A.L. et al. Inhibition of Rho-kinase improves erectile function, increases nitric oxide signaling and decreases penile apoptosis in a rat model of cavernous nerve injury. J. Urol.. 2013 doi:10.1016/j.juro.2012.09.104 PubMed
264. Wingard C.J., Johnson J.A., Holmes A., Prikosh A.. Improved erectile function after Rho-kinase inhibition in a rat castrate model of erectile dysfunction. Am. J. Physiol. Regul. Integr. Comp. Physiol.. 2003 doi:10.1152/ajpregu.00041.2003 PubMed
265. Shin J.Y., Kim Y.I., Cho S.J., Lee M.K., Kook M.C., Lee J.H. et al. MicroRNA 135a suppresses lymph node metastasis through down-regulation of ROCK1 in early gastric cancer. PLoS ONE. 2014 doi:10.1371/journal.pone.0085205 PubMed
266. Li E., Zhang J., Yuan T., Ma B.. MiR-145 inhibits osteosarcoma cells proliferation and invasion by targeting ROCK1. Tumour Biol.. 2014 doi:10.1007/s13277-014-2031-9 PubMed
267. Li J., Song Y., Wang Y., Luo J., Yu W.. MicroRNA-148a suppresses epithelial-to-mesenchymal transition by targeting ROCK1 in non-small cell lung cancer cells. Mol. Cell. Biochem.. 2013 doi:10.1007/s11010-013-1682-y PubMed
268. Cascione M., De Matteis V., Toma C.C., Pellegrino P., Leporatti S., Rinaldi R.. Morphomechanical and structural changes induced by ROCK inhibitor in breast cancer cells. Exp. Cell Res.. 2017 doi:10.1016/j.yexcr.2017.09.020 PubMed
269. Ueno K., Hirata H., Shahryari V., Chen Y., Zaman M.S., Singh K. et al. Tumour suppressor microRNA-584 directly targets oncogene Rock-1 and decreases invasion ability in human clear cell renal cell carcinoma. Br. J. Cancer. 2011 doi:10.1038/sj.bjc.6606028 PubMed
270. Ogawa T., Tashiro H., Miyata Y., Ushitora Y., Fudaba Y., Kobayashi T. et al. Rho-associated kinase inhibitor reduces tumor recurrence after liver transplantation in a rat hepatoma model. Am. J. Transplant.. 2007 doi:10.1111/j.1600-6143.2006.01647.x PubMed
271. Kroiss A., Vincent S., Decaussin-Petrucci M., Meugnier E., Viallet J., Ruffion A. et al. Androgen-regulated microRNA-135a decreases prostate cancer cell migration and invasion through downregulating ROCK1 and ROCK2. Oncogene. 2015 doi:10.1038/onc.2014.222 PubMed
272. Patel R.A., Forinash K.D., Pireddu R., Sun Y., Sun N., Martin M.P. et al. RKI-1447 is a potent inhibitor of the Rho-associated ROCK kinases with anti-invasive and antitumor activities in breast cancer. Cancer Res.. 2012 doi:10.1158/0008-5472.CAN-12-0954 PubMed
273. McLeod R., Kumar R., Papadatos-Pastos D., Mateo J., Brown J.S., Garces A.H.I. et al. First-in-Human Study of AT13148, a Dual ROCK-AKT Inhibitor in Patients with Solid Tumors. Clin. Cancer Res.. 2020 doi:10.1158/1078-0432.CCR-20-0700 PubMed
274. Garg R., Riento K., Keep N., Morris J.D., Ridley A.J.. N-terminus-mediated dimerization of ROCK-I is required for RhoE binding and actin reorganization. Biochem. J.. 2008 doi:10.1042/BJ20071342 PubMed
275. Jacobs M., Hayakawa K., Swenson L., Bellon S., Fleming M., Taslimi P. et al. The structure of dimeric ROCK I reveals the mechanism for ligand selectivity. J. Biol. Chem.. 2006 doi:10.1074/jbc.M508847200 PubMed
276. Gu Z., Yan T., Yan F.. Rational design and improvement of the dimerization-disrupting peptide selectivity between ROCK-I and ROCK-II kinase isoforms in cerebrovascular diseases. J. Mol. Recognit.. 2020 doi:10.1002/jmr.2835 PubMed
277. Hartmann S., Ridley A.J., Lutz S.. The Function of Rho-Associated Kinases ROCK1 and ROCK2 in the Pathogenesis of Cardiovascular Disease. Front. Pharmacol.. 2015 doi:10.3389/fphar.2015.00276 PubMed
278. Couzens A.L., Saridakis V., Scheid M.P.. The hydrophobic motif of ROCK2 requires association with the N-terminal extension for kinase activity. Biochem. J.. 2009 doi:10.1042/BJ20081376 PubMed