Regulation of growth factor signaling and cell cycle progression by cell adhesion and adhesion-dependent changes in cellular tension

Cytokine & Growth Factor Reviews 16 (2005) 395–405 www.elsevier.com/locate/cytogfr

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Regulation of growth factor signaling and cell cycle progression by cell adhesion and adhesion-dependent changes in cellular tension Janice L. Walker, Alaina K. Fournier, Richard K. Assoian * Department of Pharmacology, University of Pennsylvania School of Medicine, 3620 Hamilton Walk, 167 Johnson Pavillion, Philadelphia, PA 19104-6084, USA Available online 10 May 2005

Abstract The proliferation of most non-transformed cell types requires cell adhesion and cellular tension as well as exposure to mitogenic growth factors. Integrins and cadherins provide the adhesion signals, which ultimately allow for the cytoskeletal changes that control cellular tension. This review discusses the roles of integrins, cadherins, and the actin cytoskeleton as mediators of the mechanical tension critical for growth factor-dependent signaling and cell cycle progression. # 2005 Elsevier Ltd. All rights reserved. Keywords: Integrin; Cadherin; Receptor tyrosine kinase; Mechanical force; G1 phase; Cyclin

Contents 1. 2. 3. 4. 5. 6. 7. 8. 9. 10.

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Integrins, cell adhesion, cell spreading, and regulation of cellular tension. . . . . . . . . . . . . . . . . . . . . . . . Mechanisms of mechanical signaling . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Links between integrin- and growth factor-dependent signaling in G1 phase . . . . . . . . . . . . . . . . . . . . . . Role of the actin cytoskeleton, cell shape, and ECM rigidity in regulating G1 phase cell cycle progression Distinct pathways to cyclin D1 differ in their requirements for cellular tension . . . . . . . . . . . . . . . . . . . . Fibroblasts in 3D culture . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Regulation of adhesion, tension, and cell cycle progression by cell–cell adhesion . . . . . . . . . . . . . . . . . . Epithelial cells in 3D . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Concluding comments . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Acknowledgements . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

1. Introduction Most non-transformed cell types require growth factor stimulation, adhesion to the extracellular matrix (ECM) and organization of the actin cytoskeleton for cell cycle progression. In addition, epithelial and endothelial cells can control G1 phase progression through cadherin-mediated cell–cell adhesion. Finally, cell shape can influence growth factor-dependent G1 phase progression, since inhibition of * Corresponding author. Tel.: +1 215 898 7157; fax: +1 215 573 5656. E-mail address: [email protected] (R.K. Assoian). 1359-6101/$ – see front matter # 2005 Elsevier Ltd. All rights reserved. doi:10.1016/j.cytogfr.2005.03.003

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cell spreading also leads to G1 arrest in growth factor-treated cells [1]. Many studies now indicate that the adhesion and shape requirements for proliferation have a similar molecular basis and reflect alterations in signaling and cytoskeletal organization mediated by integrins and cadherins. Integrins are the class of transmembrane receptors that mediate the binding of cells to ECM proteins such as collagen, fibronectin, laminin, and vitronectin. In addition to anchoring the cell to the ECM, the binding of ECM proteins to integrins triggers outside–in signal transduction cascades conceptually analogous to those stimulated by ligand-dependent activation of growth factor receptor tyrosine kinases (RTKs). However,

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integrins are bidirectional signaling receptors which also transmit inside–out signals. These inside–out signals reflect the fact that the cytosolic domains of integrins are coupled to the actin cytoskeleton, so changes in the actin cytoskeleton can modulate integrin clustering and/or affinity [2,3]. Classical cadherins are attached to the actin cytoskeleton via b- and a-catenins [4], suggesting that these receptors also mediate both outside–in and inside–out signaling. Thus, in addition to their ability to initiate chemical signaling, integrins and cadherins can act as mechanical sensors, responding to changes in cell shape and changes in cytoskeleton-derived tension. Importantly, these alterations in tension influence growth factor-mediated signaling. This review focuses on the effects of integrins, cadherins, the actin cytoskeleton, and cytoskeletal-dependent tension in regulating growth factor-dependent signal transduction and G1 phase cell cycle progression via the G1 phase cyclin-dependent kinases (cdks). G1 phase cell cycle progression has been extensively reviewed [5–8], and the reader is directed to one of these reviews for details on the G1 phase cyclin-dependent kinases.

2. Integrins, cell adhesion, cell spreading, and regulation of cellular tension Integrins are a large family of heterodimeric transmembrane proteins comprised of a- and b-chains [9], and the heterodimerization of different a- and b-chains results in ligand specificity. For example, a5b1 is a receptor for fibronectin, a1b1 and a2b1 are receptors for collagen, and avb3 receptor is a multi-ligand receptor binding fibronectin, vitronectin, osteopontin, and other matrix proteins. With the exception of a6b4 (which participates in hemidesmosome formation), the cytoplasmic domains of the a- and b-chains are small (13–70 amino acids) [3,9]. These cytoplasmic tails lack intrinsic kinase activity characteristic of RTKs but initiate signaling by recruiting kinases such as focal adhesion kinase (FAK; see below). Integrins cluster at discrete sites of cell contact with the substratum. Different integrin–matrix adhesion structures vary in size, location, composition, and tension-bearing properties [10,11]. It is important to understand the different physical properties of these distinct integrin-containing structures since they can contain RTKs and likely transmit mechanical signals (through the actin cytoskeleton) necessary for RTK-mediated S phase entry. Upon initial contact between the integrin and the ECM, small nascent adhesion structures containing talin and actin are formed at the leading edge of the cell [12–14]. These small adhesion structures exert a large amount of force, which decreases as they develop into larger, more mature structures [15,16]. Focal complexes develop from these initial integrin–ECM contacts in a Rac-dependent manner [17] and are identified by the recruitment of vinculin, paxillin, and phosphoproteins such as FAK [18]. Galbraith

et al. [19] demonstrated that recruitment of vinculin to focal complexes is required to transmit force. Focal adhesions (also known as focal contacts) develop from focal complexes, and these structures accumulate additional proteins, are more elongated, centrally located, and attach to bundles of actin stress fibers that traverse the cell [17,20]. Unlike nascent adhesions, focal adhesions increase in size proportionally to the applied force [21,22]. Focal adhesion and stress fiber formation occurs in a Rho-dependent manner [23] and require myosin light chain (MLC)dependent cell contractility [24–26]. Two effectors of Rho, Rho kinase and the formin-homology protein, mDia, can substitute for Rho to regulate focal adhesion and stress fiber formation [27]. Rho kinase stimulates myosin II-driven contractility by direct phosphorylation of MLC [28,29] and by phosphorylating and inactivating MLC phosphatase [30,31]. mDia interacts with profilin to stimulate actin polymerization [27] and, additionally, stabilizes microtubules [32]. The requirement for Rho kinase-mediated actin–myosin contractility can be bypassed if an external force is applied to the cell [33]. Under these circumstances mDia is necessary and sufficient to regulate focal adhesion formation [33]. Fibrillar adhesions (also known as ECM contacts) emerge from focal adhesions and are marked by accumulation of a5b1 integrin, tensin, and low levels of other focal adhesion proteins such as vinculin and paxillin [20,34,35]. Fibrillar adhesions mediate fibronectin fibrillogenesis in a tensindependent manner; a dominant-negative inhibitor of tensin blocks fibrillogenesis, translocation of a5b1 integrin and fibrillar adhesion formation without affecting focal adhesion structures [36]. Rho-generated contractility is also important for the formation (but not the maintenance) of fibrillar adhesions [20,34] and fibronectin matrix assembly [37]. Integrin-containing structures can influence growth factor-mediated regulation of G1 phase cell cycle progression. For instance, integrin linked kinase (ILK), a serine/ threonine cytoplasmic kinase, and at least two of its binding partners, PINCH and ILKBP, localize to focal and fibrillar adhesions [38]. If their interactions are perturbed, these molecules no longer localize to adhesion structures and growth factor-mediated S phase entry is inhibited [38]. Thus, the molecular composition of a matrix adhesion can influence G1 phase cell cycle progression by RTKs. Additionally, integrin-mediated adhesion structures can affect S phase progression by regulating cell shape. Cell shape has been tightly linked to regulation of growth factormediated S phase progression (see below), and the force regulated by each integrin–ECM adhesion structure provides the cell with a mechanism to control cell spreading.

3. Mechanisms of mechanical signaling Molecules that detect or respond to mechanical signals have been called mechanosensors, and integrins fall into this category [11,39] (Fig. 1). Tugging on an integrin receptor

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cytoskeletal connection can translate mechanical signals into chemical signals. Changes in actin polymerization provide another way for mechanical tension to regulate cell proliferation (Fig. 1). The transcriptional co-activator, MAL, is retained in the cytoplasm as a consequence of its association with actin. Decreases in the amount of G actin associated with actin polymerization results in the nuclear accumulation of MAL, where it co-activates transcription of genes containing serum response factor (SRF) sites in their promoters [49]. SRFregulated genes include immediate–early genes associated with G1 phase progression, and MAL co-activates a subset of these genes including SRF itself, vinculin, and Cyr61.

4. Links between integrin- and growth factordependent signaling in G1 phase

Fig. 1. Interplay between mechanosensors and the actin cytoskeleton. Potential mechanosensors transmit signals between the actin cytoskeleton and the external environment. Integrins clustered in focal adhesion complexes sense tensional forces from the extracellular matrix (ECM) to the actin cytoskeleton. Of the proteins in focal adhesions (such as talin, vinculin, paxillin and FAK), FAK has received the most attention as a mechanosensor since its phosphorylation changes in response to mechanical force in 2D and 3D environments. Other molecules, such as Rho, may act as intracellular mechanosensors by altering actin cytoskeletal organization, thereby affecting the formation of focal adhesions and adherens junctions. Within the adherens junctions, cadherins link to the actin cytoskeleton via bcatenin and a-catenin, transmitting mechanical force signals between neighboring cells and the actin cytoskeleton. Thus, mechanical forces can act at multiple levels to influence formation and composition of cell adhesion structures, which ultimately can lead to changes in signaling through associated RTKs.

with a magnetic twisting device can alter cytoskeletal organization and nuclear form, suggesting that integrins, the actin cytoskeleton, and the nucleus are interconnected [40]. Moreover, Feldherr and Akin [41] showed that mechanically induced changes in nuclear shape can affect nuclear import of transcription factors, and nuclear size has been correlated with initiation of cell growth [42]. b1 integrin binding to fibronectin-bound beads can recruit mRNAs and ribosomes to focal adhesions, and this recruitment likely depends on tensional forces since drugs that inhibit actin–myosin contraction blocks recruitment of mRNA to the integrin [43]. We and others have demonstrated that the translation of cyclin D1 mRNA occurs in an adhesion-dependent manner [44–46], and it would be interesting to know if cyclin D1 mRNA is recruited to focal adhesions for translation. Sheetz and co-workers have reported that stretch promotes the binding of several phosphoproteins (including FAK, paxillin and p130Cas) to Triton X-100 cytoskeletons [47]. Stretch was also sufficient to activate the Crk/C3G/Rap1 pathway at sites of cell–ECM contact [48]. Thus, the ECM–integrin–

Sites of integrin–ECM adhesion serve as points of convergence for integrins and RTKs. In fact, RTKs themselves have been identified within these adhesion structures, and interactions between RTKs and integrins have been documented biochemically and functionally. avb3 can interact with receptors for IGF-1, PDGF, and VEGF [50–53], and PDGF-induced S phase entry measured by thymidine incorporation is enhanced on vitronectin (an avb3 ligand) compared to collagen [51]. Interactions between integrins and EGF receptors have been extensively studied in epithelial cells. Moro et al. [54] demonstrated that the EGF receptor can associate with b1 integrin and that EGF receptor intrinsic kinase activity is activated by integrin-mediated adhesion even in the absence of EGF. This ligand-independent activation of the EGF receptor results in ERK1 activation and tyrosine phosphorylation of Shc, but fails to drive cell cycle progression into G2/M without the addition of EGF or serum. In a subsequent study, Moro et al. [55] identified a macromolecular complex containing EGF receptor, avb3 integrin, p130CAS, and Src. Inhibition of Src activity or EGF receptor tyrosine kinase activity inhibited the association of the EGF receptor with avb3, as did loss of p130CAS. Most recently, Bill et al. [56] demonstrated that integrindependent activation of the EGF receptor in epithelial cells activates ERK- and AKT-dependent G1 phase progression as measured by the induction of cyclin D1, phosphorylation of Rb and activation of cdk4. However, integrin-dependent activation of EGF receptor is insufficient to downregulate p27, and as a consequence these cells arrest in G1 phase with inactive cyclin E-cdk2 and hypophosphorylated Rb. Addition of EGF, or overexpression of myc or the EGF receptor, rescues S phase entry. Functional interactions between RTKs and integrins have also been documented downstream of the receptors themselves. Perhaps the best-studied target of RTK/integrin synergism is the ERK subfamily of MAP kinases. Several labs have shown that integrins, like RTKs, transiently

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activate ERK1 and ERK2 [57–59] In addition, integrins can enhance the level of ERK activity seen in response to a mitogenic stimulus [54,60–66]. The induction of cyclin D1 mRNA requires a sustained activation of ERK1/2 [67–71] and a synergistic signaling by RTKs and integrins [44,72]. In this setting, the activation of ERK by RTKs alone or integrins alone is insufficient to induce cyclin D1 mRNA because the ERK signal is not sufficiently sustained [70]. In contrast, when both RTKs and integrins are ligated concomitantly, the ERK signal is sustained for >6 h and cyclin D1 mRNA is induced [70]. Thus, there appear to be three distinct effects of integrin signaling on ERK1/2 activation: integrins alone activate ERK; integrins increase the amplitude of the ERK signal induced by a mitogenic stimulus; and integrins sustain the ERK signal in cells treated with an optimal growth factor stimulus. The points of RTK–integrin synergism are not clear, with reports of synergistic effects both upstream and downstream of Ras– GTP loading [63,64,73,74]. Renshaw et al. [75] initially reported that FAK activity is required for the synergistic effect of RTKs and integrins on S phase entry. Guan and co-workers [76–78] subsequently showed that cyclin D1 mRNA is the major target for FAK activity in G1 phase. FAK stimulates both Ets and KLF8dependent cyclin D1 promoter activity. Consistent with these reports, we find that FAK activity is required for sustained ERK activation in MEFs (unpublished results). However, Barberis et al. [79] showed that PDGF will sustain ERK in fibroblasts expressing a b1 integrin that fails to recruit FAK. Similarly, sustained ERK activity and induction of cyclin D1 is independent of FAK in NIH3T3 cells [73]. The likely conclusion of these experiments is that cells probably have FAK-dependent and -independent mechanisms to regulate ERK activity and cyclin D1 induction. FAK-dependent mechanisms may involve Srcfamily kinases and B-Raf [79]. A FAK-independent mechanism involving caveolin and Fyn has been described by Wary et al. [80,81]. Other studies indicate that FAK also regulates levels of the cdk2 inhibitor, p27, by controlling expression of Skp2, the E3 ubiquitin ligase responsible for p27 degradation [82].

5. Role of the actin cytoskeleton, cell shape, and ECM rigidity in regulating G1 phase cell cycle progression Several studies have indicated that organization of the actin cytoskeleton and the consequent cell spreading is required for RTK-mediated G1 phase progression. In fibroblasts, this requirement for cytoskeletal integrity lasts slightly beyond the phosphorylation of Rb [72]. Inhibition of actin polymerization, Rho kinase, or MLC-dependent contractility prevents growth factor-dependent induction of cyclin D1 mRNA and protein in fibroblasts [72,73]. Similarly, growth factors fail to stimulate sustained activation of the Ras–Raf–MEK–ERK

pathway in the absence of an organized actin cytoskeleton or when actin–myosin contractility is inhibited [73]. Antibodymediated clustering of a5b1 can rescue sustained ERK activity and cyclin D1 expression in cells treated with MLC kinase inhibitors [73]. FAK autophosphorylation is inhibited when contractility is blocked, and this effect is also rescued by antibody-induced clustering of a5b1 [73]. These data support a model by Burridge and co-workers [25,83], which proposes that actin–myosin contractility is required for the clustering (and hence signaling) of integrins. Integrin-dependent ERK activation in the absence of growth factors is also blocked by depolymerization of the cytoskeleton [57–59]. The requirement for an organized actin cytoskeleton extends beyond ERK and cyclin D1 in endothelial cells; ERK activation is required to mid-G1 phase while an intact cytoskeleton is required longer, up to 3 h prior to the restriction point [84]. Moreover, the expression of cyclin D1 mRNA is not inhibited when mitogen-treated endothelial cells are plated on low density fibronectin (a condition that allows for cell adhesion but not cell spreading) [45], but the expression of cyclin D1 protein is reduced. Mitogen-treated endothelial cells plated on low density fibronectin also fail to downregulate p27 or enter S phase. All these events occur readily when growth factor-treated endothelial cells are spread on high-density fibronectin [45]. Similarly, Mettouchi et al. [46] showed that cyclin D1 mRNA is efficiently translated when endothelial cells are attached to fibronectin (a substratum supporting cell spreading) but not to laminin (a substratum that does not support efficient cell spreading). Recently, Mammoto et al. [85] concluded that the effect of endothelial cell shape on p27 is mediated by Rho, and that a balance between Rho-kinase and mDia activities (two Rho effectors) regulates the expression of Skp2, the p27 E3 ligase. Several other studies have also indicated that cell spreading is required for growth factor-dependent mitogenesis. Koyoma et al. [86] showed that growth factor-treated smooth muscle cells arrest in G1 phase when plated on polymerized type I collagen fibrils (a condition in which the cells are poorly spread) whereas plating on monomeric collagen films (where cells are well spread) is permissive for growth factor-stimulated proliferation. The G1 phase arrest seen with polymerized collagen was associated with inhibition of cyclin E-cdk2 activation and increased levels of the cdk2 inhibitors, p27 and p21. These effects may be related to the fact that FAK inhibition prevents Skp2 expression in smooth muscle cells [82]. In hepatocytes, growth factors efficiently induce cyclin D1 mRNA and protein when the cells are spread on collagen films, but not when their spreading is blocked by plating the cells on collagen gels [87]. S phase entry in these hepatocytes correlates with the degree of cell spreading and cyclin D1 expression. Cyclin D1 expression in hepatocytes is dependent on Rho, organization of the actin cytoskeleton and actin–myosin contractility [87,88], as it is in fibroblasts [71]. Moreover, hepatocytes plated on collagen films show

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two peaks of ERK activity during G1 phase progression, and only the second (mid-G1 phase) peak of ERK activity is critical for cyclin D1 expression and S phase entry [89]. These results are in good agreement with those in fibroblasts showing that integrin signaling is required for cyclin D1 induction because it is needed to sustain ERK activity (see above). Taken together, the spreading/tension requirements for ERK-dependent cyclin D1 expression and p27 downregulation probably explain why Rb hyperphosphorylation, cyclin A gene expression and S phase entry are all associated with cell spreading. S phase entry induced by mechanical strain can depend on specific ECM–integrin interactions. When endothelial cells are exposed to sustained hydrostatic pressure they increase av expression and exhibit altered distribution of av (av-containing focal adhesions were smaller and more elongated under pressure than controls). Entry into S phase is reduced by an antagonist of av [90]. Similarly, strain induces DNA synthesis in smooth muscle cells plated on fibronectin, vitronectin, and collagen, but not on laminin or elastin [91]. Adding soluble fibronectin, RGD peptides, or function blocking antibodies to avb5 and b3 integrin inhibits strain-induced proliferation. In this system, RGD peptides inhibit strain-induced autocrine production of PDGF by suppressing the PDGF A promoter. Thus, tension can stimulate G1 phase progression by regulating growth factor gene expression. Kumar et al. [92] demonstrated that mechanical cyclic strain increases cell proliferation of myoblasts (C2C12 cells) by inducing the expression of cyclin D1 and cyclin A, and by increasing the activity of cdk2. Cyclic strain activated FAK, Rac, and NFkB, and inhibition of these signaling molecules blocks cyclic straininduced myoblast proliferation. Sechler and Schwarzbauer [93] used CHO cells to compare the effects of native fibronectin and fibronectin delta III1-7 on actin assembly and cell cycle progression. Incubating cells with native fibronectin led to the formation of actin stress fibers and colocalization of a5b1 integrin, FAK, vinculin, and paxillin to focal adhesions. When cells were incubated with fibronectin delta III1-7, a5b1 integrins, and FAK were again clustered but actin reorganization and focal adhesion formation were delayed. The altered fibronectin matrix inhibited S phase entry (relative to native fibronectin) and also had a dominantnegative effect on growth stimulation by native fibronectin. Alterations in FAK tyrosine phosphorylation were also noted, with more FAK tyrosine phosphorylation occurring in response to native fibronectin. Thus, alterations in integrin signaling that result from changes in ECM architecture can affect S phase progression. These data indicate that FAK phosphorylation is a mechanosensor in growth factor-mediated G1 phase cell cycle progression (Fig. 1). FAK-null cells fail to reorient their focal adhesions in response to an externally applied force [94], consistent with work showing that FAK is required for the disassembly of focal adhesions [95,96].

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6. Distinct pathways to cyclin D1 differ in their requirements for cellular tension As outlined above, growth factors stimulate sustained ERK activity when added to quiescent, spread fibroblastic cells, and this sustained ERK signal leads to the induction of cyclin D1 in mid-G1phase [70,73]. The ERK pathway to cyclin D1 is dependent on the presence of actin stress fibers (the typical situation when quiescent fibroblasts are growth factor-stimulated in culture). However, Rac can also induce cyclin D1 in fibroblasts [71]. This pathway is normally cryptic, but can be revealed by inhibition of Rho or Rho kinase. Since Rho/Rho kinase inhibition prevents stress fiber formation, the Rac-dependent pathway to cyclin D1 must be independent of the cellular tension associated with stress fibers [97]. Thus, while both pathways require growth factor signaling, the ERK and Rac pathways to cyclin D1 expression can be distinguished by their tensional requirements. These results raise the possibility that these superficially redundant pathways to cyclin D1 may co-exist to allow for G1 phase progression in varying tensional environments.

7. Fibroblasts in 3D culture Culturing cells in three dimensions (3D) provides another way to study the consequences of changing tension on growth factor-dependent G1 phase progression. Human fibroblasts plated on cell or tissue-derived 3D matrices form distinct integrin structures called ‘‘3D-matrix adhesions’’ and have higher proliferation rates than cells grown in 2D (monolayer) cultures [98]. Mechanical compression of 3D cultures results in the loss of 3D matrix adhesions and a decreased rate of proliferation in growth factor-treated cells. Thus, growth factor-dependent responses in 3D cultures are affected by the rigidity or pliability of the substrate. It is worth noting that although FAK localizes to these 3D matrix adhesions, it is not efficiently autophosphorylated at Y397 as compared to cells plated on fibronectin in 2D. FAK phosphorylation may therefore be a mechanosensor in both 2D and 3D environments (Fig. 1). Fibroblasts grown in 3D collagen matrices exhibit distinct growth properties depending on whether the cells are cultured under mechanically loaded (attached) or unloaded (floating) conditions [99]. Under mechanically loaded conditions, growth factor-treated fibroblasts develop stress fibers, and replicate DNA whereas growth factortreated fibroblasts cultured under unloaded conditions have low levels of DNA synthesis [100]. Fibroblasts under unloaded conditions have reduced autophosphorylation of PDGF receptor (decreased receptor kinase activity) in response to PDGF even though normal levels of PDGF receptors remain on the cell surface [100,101]. Similarly, growth factors poorly induce cyclin D1 and downregulate p27 in mechanically unloaded fibroblasts [102]. These

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effects can be traced to an inefficient activation of Raf and MEK [103]. Many of the signaling and cell cycle defects seen in 3D fibroblast cultures are reminiscent of those seen in suspended (nonadherent) cells [44]. However, in contrast to results obtained in 2D [62,72], depolymerization of actin does not significantly inhibit ERK activation, cyclin D1 expression or p27 downregulation in mechanically loaded 3D fibroblast cultures [102]. In related studies, fibroblasts were cultured for several days in mechanically loaded collagen gels prior to release of tension. Under these conditions, growth factor-stimulated fibroblasts remain spread although they lack stress fibers and do not undergo DNA synthesis [104]. The absence of DNA synthesis in these spread cells suggests that spreading is necessary but not sufficient for growth factor-dependent mitogenesis. Davey et al. [105] reached a similar conclusion by showing that NRK fibroblasts lacking a5b1 integrin spread fully when plated on collagen and stimulated with serum, yet progress through G1 phase poorly. Thus, it seems that integrins have distinct effects on G1 phase progression, with some related to integrin-specific signaling while others are related to a general integrin-dependent polymerization of actin, cell spreading, and generation of isometric tension.

8. Regulation of adhesion, tension, and cell cycle progression by cell–cell adhesion Like fibroblasts, epithelial and endothelial cells maintain adhesion to the ECM via integrins. However, epithelial and endothelial cells also rely on cell–cell adhesion to maintain tissue integrity and function [4,106,107]. Cadherins are plasma membrane-spanning proteins that mediate cell–cell adhesion by binding to identical cadherins on the surface of neighboring cells. The cytoplasmic domains of cadherins are linked to the actin cytoskeleton and can influence both integrin–matrix adhesions and growth factor signaling. Cadherin-mediated cell–cell adhesion is fortified by catenin molecules that bind to the cytoplasmic tail of cadherins and link it to the actin cytoskeleton. Cadherins cluster at sites of cell–cell contact called adherens junctions and attach to the actin cytoskeleton, allowing for the development of mechanical stress within the epithelium [39,108]. This source of mechanical stress is probably dynamic and responsive in nature to the external environment, e.g. half of the E-cadherin protein within the cytoplasm of L cells expressing wild type E-cadherin is connected to the cytoskeleton while the other half remains free [109]. Ecadherin associates with the EGF receptor via b-catenin in several epithelial cell lines [110–112], and this interaction can lead to a ligand-independent activation of the EGF receptor [112] that is conceptually analogous to the ligandindependent of EGF receptors by integrins (see above). Conversely, the binding of EGF to the EGF receptor can uncouple E-cadherin from the actin cytoskeleton by phosphorylating b-catenin and g-catenin [111].

In addition to the EGF receptor, focal adhesions and adherens junctions share several signaling and structural proteins, including a-actinin, Arp2/3, zyxin, moesin, and vinculin (reviewed in [113]). Thus, these two types of junctions are probably in dynamic competition with one another, leading to a regulated balance between cell–cell adhesion and ECM–cell adhesion. In fact, Ryan et al. [114] found that increasing cell–cell adhesion leads to a decrease in cell–substrate adhesion and cell spreading. These results fit well with studies showing that overexpression of Ecadherin leads to growth arrest while inhibition of cadherin function or loss of adherens junction assembly induces tumorigenicity (reviewed in [115]). Thus, changes in cadherin-mediated cell–cell adhesion have the potential to alter cellular tension through direct changes in the actin cytoskeleton and through indirect changes in ECM– substrate adhesion (Fig. 1). Competition for shared focal adhesion and adherens junction components likely contribute to these growth effects. Since the decreased cell spreading that accompanies cell– cell adhesion is associated with G1 phase growth arrest (see above), it has been difficult to isolate the specific effect of cadherin-mediated signaling on cell proliferation. Nelson and Chen [116] were able to uncouple cell–cell adhesion from cell-spreading using micropatterned substrates in which cell spreading is constant and only cell–cell adhesion is varied. These authors showed that, under these conditions, cell–cell adhesion mediated by VE-cadherin stimulates S phase entry. This stimulatory effect of VE-cadherin requires an intact actin cytoskeleton and actin–myosin contractility [117]. Thus, cadherin-mediated cell–cell adhesion has the potential to both stimulate and inhibit cell proliferation, and the direction of its effect may well depend on the degree to which ECM/integrin-mediated cell spreading is affected. Cadherins sequester b-catenin, a molecule that can translocate to the nucleus and activate TCF/LEF-dependent genes such as cyclin D1 [118,119]. The effects of cadherin expression and actin-dependent stress on b-catenin-dependent transcription are poorly understood.

9. Epithelial cells in 3D In 2D culture, fibroblasts and epithelial cells behave very similarly. They proliferate in response to a coordination of adhesion and growth factor signals, while remaining undifferentiated and unorganized. Mammary epithelial cells in mechanically loaded 3D collagen gels spread with a morphology similar to that seen in 2D culture, have high Rho activity, FAK autophosphorylation, and proliferate in response to exogenous mitogens [120]. When mammary epithelial cells are cultured in floating 3D matrices, they go through an initial burst of cell proliferation and then become growth arrested even if growth factors and an ECM are present [121,122]. However, in contrast to fibroblasts, growth-arrested epithelial cells in floating collagen gels

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undergo morphogenesis, forming polarized, tissue-like acini or tubes with hollow lumens toward the apical surface and a basement membrane at the basal surface [123–125]. Differentiated mammary epithelial cells in unloaded 3D cultures have lower levels of Rho activity and FAK autophosphorylation than their counterparts in mechanically loaded collagen gels [120]. Under these released conditions, vinculin and p16 of the Arp2/3 complex are localized to cell–cell junctions and the cell periphery rather than 3D matrix adhesions. When extracellular tension is increased (by increasing collagen density or by leaving the gel attached to the dish), FAK, vinculin, and p16 become localized to 3D matrix adhesions, and FAK is autophosphorylated [120]. Thus, rigidity of the matrix appears to regulate the formation of 3D matrix adhesions, which, in turn, mediates downstream signaling events that control growth factor-mediated cell cycle progression. Bissell and co-workers have championed the use of 3D mammary cell cultures to explore the relationships between integrins, cadherins and tumorigenicity using a tumor progression series in which continual passage of a nonmalignant mammary epithelial line (S-1) gave rise to a tumorigenic derivative (T4-2) that overexpresses b1 integrin. Weaver et al. [126] showed that the T4-2 cells could be reverted to a normal, non-tumorigenic phenotype by blocking b1 integrin signaling with a function-blocking antibody. Like the S-1 cell line, the reverted cells became growth arrested and differentiated into acinar structures, indicating that increased b1 integrin signaling had contributed to increased proliferation. In a follow-up study, Wang et al. [127] showed that the EGF receptor is also upregulated in the tumorigenic cells and that b1 integrin and the EGF receptor are jointly regulated in this system. This receptor co-regulation is not seen in 2D culture, suggesting that the cell shape provided by the 3D environment may be regulating localization of these receptors, and that specific localization and context may be required for receptor expression. Additionally, E-cadherin is not properly localized at cell–cell junctions in cells overexpressing b1 integrin but is relocalized upon reversion. Liu et al. [128] used the reversion model to demonstrate that inhibition of phosphatidylinositol 3-kinase (PI3K) prevents the loss of tissue polarity and increased proliferation characteristic of the breast tumor phenotype. PI3K inhibition also downregulates levels of b1 integrin and the EGF receptor. Downstream of PI3K, AKT signaling regulated proliferation, whereas Rac signaling controlled polarity [128]. The conversion from proliferation to morphogenesis and acini formation is also detected when MCF10A mammary epithelial cells are cultured in 3D matrices. In this setting, ectopic expression of cyclin D1 or E7 (which inactivates the retinoblastoma protein) overrides the cell cycle arrest seen in late-stage cultures undergoing morphogenesis into acini [122]. Activated AKT, although minimally proliferative in itself, enhanced the late stage proliferative responses seen in cyclin D1- or E7-expressing cells [129]. Ectopic expression

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of activated CSF-1 receptor or Erb2 also increased cell proliferation in late-stage 3D cultures [130,131]. But in contrast to the effects seen with cyclin D1 and E7, AKT, CSF-1, and Erb2 activation disrupted morphogenesis [122,129–131]. The morphogenic effects of CSF-1 receptor signaling was associated with the redistribution of Ecadherin away from the plasma membrane and a disordered distribution of laminin-5 [130]. CSF-1 receptor signaling in these 3D mammary epithelial cell cultures provides a potentially powerful system for studying relationships between adhesion-dependent morphogenesis and proliferation.

10. Concluding comments Two of the major adhesion receptors, integrins and cadherins, are linked to the actin cytoskeleton and can respond to the mechanical environment (Fig. 1). In addition, the adhesion structures containing integrins and cadherins are regulated by intracellular mechanosensors such as FAK and Rho, which are, in turn, affected by cellular architecture (Fig. 1). There may well be a continual interplay between integrins, cadherins, the actin cytoskeleton, and intracellular mechanosensors, with each component existing in a dynamic equilibrium with the others (Fig. 1). Since integrins and cadherins are physically coupled to RTKs (Fig. 1), it becomes easy to visualize how remodeling of the ECM, altered expression of integrins and cadherins, or changes in the actin cytoskeleton could affect RTK-dependent signaling. As discussed above, cell spreading and mechanical loading are necessary for most cells to proliferate, leading to the conclusion that cell architecture and accompanying changes in tension are critical determinants of adhesion- and growth factor-mediated cell cycle progression. Using cyclin D1 expression in fibroblasts as a paradigm, we propose a threshold model by which mechanical force regulates growth factor-dependent progression through G1 phase (Fig. 2). In conditions that are below the tension threshold (such as seen in suspended cells or cells treated with actin depolymerizing agents), cells are unable to induce cyclin D1 or progress through G1 phase. Above this threshold, different amounts of tension may lead to activation of different signaling pathways that control G1 phase progression. For example, actin polymerization in the absence of stress fibers is sufficient to allow for Racdependent induction of cyclin D1, while the formation of actin stress fibers at the highest levels of tension allow for the sustained activation of ERK and an ERK-dependent induction of cyclin D1. This model provides an explanation for the apparent redundancy in the signaling pathways to cyclin D1 by emphasizing that the use of Rac- versus ERKdependent signaling gives the cell plasticity in regulating proliferation in response to its mechanical environment. It will be interesting to see if a threshold model can predict the

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Fig. 2. Threshold model of mechanical force regulating growth factormediated G1 phase cell cycle progression. In this model, cells cultured below a minimum amount of tension (conditions A–D) will not progress through G1 phase despite the presence of a suitable mitogenic stimulus. These conditions include incubation of cells in suspension, incubating cells with inhibitors of actin polymerization, precluding spreading of an attached cell in 2D, or incubating cells in floating or tension-released 3D matrices. Growth factor-dependent G1 phase progression will occur when cells are placed in tensioned (attached) 3D cultures (condition E) or when they are in 2D with polymerized actin (conditions F and G). In this model, proliferating cells may be under different amounts of tension that result in activation of distinct growth factor-dependent signaling pathways. For instance, the RhoERK-cyclin D1 signaling pathway requires high tension associated with actin stress fibers (condition G) while the Rac-cyclin D1 pathway can induce cyclin D1 without stress fiber-derived tension (condition F).

cell cycle behavior of epithelial cells (which typically proliferate in the absence of actin stress fibers) as well as the behavior of epithelial cells in 3D environments where cell cycle progression, arrest, and differentiation can be manipulated by altering cellular tension.

Acknowledgements Work in our laboratory is supported by grants from the National Institutes of Health. JW was supported by NIH post-doctoral fellowship F32 GM065031-03 and AKF is supported by predoctoral training Grant R25-CA-101871.

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