- Email: [email protected]
Rapping about Mechanotransduction
Developmental Cell
Previews of nonenveloped virus fusogens that utilize several motifs (e.g., amphipathic helices, transmembrane domain, membrane destabilizing fusion peptides) to drive cell-cell fusion. Do similar motifs serve similar functions in Myomerger? Lastly, are there other examples of multicomponent fusion complexes that assign hemifusion and pore formation activities to separate proteins? Such may be the case, for example, with vaccinia virus whose fusion complex comprises 11 small proteins. REFERENCES Bi, P., Ramirez-Martinez, A., Li, H., Cannavino, J., McAnally, J.R., Shelton, J.M., Sa´nchez-Ortiz, E., Bassel-Duby, R., and Olson, E.N. (2017). Control
of muscle formation by the fusogenic micropeptide myomixer. Science 356, 323–327. Ciechonska, M., and Duncan, R. (2014). Reovirus FAST proteins: virus-encoded cellular fusogens. Trends Microbiol. 22, 715–724. Harrison, S.C. (2015). Viral membrane fusion. Virology 479-480, 498–507. Herna´ndez, J.M., and Podbilewicz, B. (2017). The hallmarks of cell-cell fusion. Development 144, 4481–4495.
Millay, D.P., O’Rourke, J.R., Sutherland, L.B., Bezprozvannaya, S., Shelton, J.M., Bassel-Duby, R., and Olson, E.N. (2013). Myomaker is a membrane activator of myoblast fusion and muscle formation. Nature 499, 301–305. Quinn, M.E., Goh, Q., Kurosaka, M., Gamage, D.G., Petrany, M.J., Prasad, V., and Millay, D.P. (2017). Myomerger induces fusion of non-fusogenic cells and is required for skeletal muscle development. Nat. Commun. 8, 15665.
Kim, J.H., Jin, P., Duan, R., and Chen, E.H. (2015). Mechanisms of myoblast fusion during muscle development. Curr. Opin. Genet. Dev. 32, 162–170.
Weber, T., Zemelman, B.V., McNew, J.A., Westermann, B., Gmachl, M., Parlati, F., So¨llner, T.H., and Rothman, J.E. (1998). SNAREpins: minimal machinery for membrane fusion. Cell 92, 759–772.
Leikina, E., Gamage, D.G., Prasad, V., Goykhberg, J., Crowe, M., Diao, J., Kozlov, M.M., Chernomordik, L.V., and Millay, D.P. (2018). Myomaker and Myomerger work independently to control distinct steps of membrane remodeling during myoblast fusion. Dev. Cell 46, this issue, 767–780.
Zhang, Q., Vashisht, A.A., O’Rourke, J., Corbel, S.Y., Moran, R., Romero, A., Miraglia, L., Zhang, J., Durrant, E., Schmedt, C., et al. (2017). The microprotein Minion controls cell fusion and muscle formation. Nat. Commun. 8, 15664.
Rapping about Mechanotransduction Consuelo Ibar1 and Kenneth D. Irvine1,* 1Waksman Institute and Department of Molecular Biology and Biochemistry, Rutgers University, Piscataway, NJ 08854, USA *Correspondence: [email protected] https://doi.org/10.1016/j.devcel.2018.09.007
Mechanical cues can regulate cell proliferation and differentiation through the Hippo-YAP signaling network. Reporting in Nature, Meng et al. (2018) show that the Ras-related GTPase RAP2 connects extracellular matrix stiffness to Hippo pathway regulation, adding to our understanding of how mechanical cues are converted into changes in YAP activity. The Hippo signaling network integrates biochemical and biomechanical cues to influence organ growth and cell fate (reviewed in Misra and Irvine, 2018). Its downstream effects are mediated by inactivation of the transcriptional co-activator proteins YAP and TAZ, through regulation of their stability and nuclear localization. YAP/TAZ are active in the nucleus, where they can induce the transcription of genes that promote cell proliferation. Mechanical cues including changes in cell shape, extracellular matrix (ECM) stiffness, cell stretching, cell density, and shear forces have all been observed to influence YAP/TAZ activity, often through effects on the levels and organization of F-actin, and tension within the actin cytoskeleton. Meng et al. (2018) have now identified the Ras-related GTPase RAP2 as a key component of the
signaling axis that connects ECM stiffness to regulation of YAP/TAZ. Their findings stem from their discovery that overexpression of RAP2A induced a cytoplasmic localization of YAP/TAZ even within cells grown on a stiff substrate, which normally have nuclear YAP/ TAZ. Conversely, loss of all three RAP2 paralogs (RAP2A, RAP2B, RAP2C) results in nuclear localization of YAP/TAZ even within cells grown on a soft substrate, which normally have cytoplasmic YAP/ TAZ. Like other small GTPases, RAP2 regulates molecular events by cycling between an inactive GDP-bound form and an active GTP-bound form. Meng and colleagues (2018) observed that low stiffness promoted binding of GTP to RAP2. This effect is mediated through previously identified activators of RAP2, PDZGEF1, and PDZGEF2, which act as guanine
678 Developmental Cell 46, September 24, 2018 ª 2018 Elsevier Inc.
nucleotide exchange factors (GEFs) for RAP2 (de Rooij et al., 1999). Moreover, PDZGEF1/2 are required for inactivation of YAP/TAZ under low stiffness. Prior studies had identified phosphatidic acid (PA) as an activator of PDZGEF and RAP2 (Gloerich et al., 2012). Consistent with this role, Meng and colleagues (2018) were able to identify influences of substrate stiffness on phospholipid metabolism and link them to regulation of PDZGEF, RAP2, and YAP/TAZ. Using a GFP reporter, they observed that low stiffness increased the cellular levels of Phosphatidylinositol-4,5bisphosphate (PI(4,5)P2), which can be converted to PA by phospholipases D1 and D2 (PLD1/2). Previous studies had linked focal adhesions to downregulation of PI(4,5)P2 through activation of PLCg1 (Zhang et al., 1999). Indeed, Meng and colleagues (2018) observed that inhibition
Developmental Cell
Previews or knockdown of PLCg1 or PLD1/2 influenced YAP/TAZ localization. ECM stiffness can influence the number and character of focal adhesions (Humphrey et al., 2014), suggesting that modulation of RAP2 activity through downregulation of PI(4,5)P2 at focal adhesions could transmit biomechanical signaling initiated by substrate stiffness. The authors also addressed the question of how RAP2 impinges on YAP/TAZ. Whether substrate stiffness acts through Hippo-dependent or Hippo-independent signaling mechanisms has remained a point of controversy since the initial discovery of biomechanical regulation of YAP/TAZ (Dupont et al., 2011). Meng et al. (2018) provide further support for Hippo pathway-dependent regulation of YAP/TAZ by ECM stiffness (Codelia et al., 2014). For example, they show that deletion of the kinases LATS1 and LATS2, which are the direct upstream regulators of YAP/TAZ within the Hippo pathway, abrogates the cytoplasmic localization of YAP normally induced by soft substrates. They also analyze changes in the transcriptome induced by substrate stiffness and show that most of the transcriptional changes induced by altered stiffness are dependent upon both YAP/TAZ and LATS1/2. MAP4K4 is one of several kinases in the Ste20 family that contribute to activation of LATS kinases, and also a previously identified downstream effector of RAP2. Meng et al. (2018) provide evidence for a RAP2-dependent activation of MAP4K4 by low stiffness. Intriguingly, a homolog of MAP4K4, Msn, has been implicated in biomechanical regulation of Hippo signaling in the Drosophila intestine (Li et al., 2018). A related RAP2 effector, TNIK/MAP4K7, has previously been identified as a regulator of F-actin (Taira et al., 2004), and F-actin levels can modulate YAP/TAZ activity. Another class of RAP2 effector is ARHGAP29, which acts as a GTPase-activating protein (GAP) for Rho. Rho has a well-established role in promoting YAP/TAZ activity, and Meng et al. (2018) now confirm that ARHGAP29 contributes to inhibition of RhoA by low stiffness. Deletion of either MAP4K4/6/7 or ARHGAP29 alone did not prevent
inhibition of YAP/TAZ by RAP2 or by low stiffness, but their combined deletion blocked inhibition of YAP/TAZ, indicating that YAP/TAZ regulation by stiffness is effected through parallel action of these downstream effectors of RAP2. Several mechanisms for focal adhesion-mediated regulation of YAP/TAZ have been described previously, including control of kinases that localize to and are activated at focal adhesions (reviewed in Misra and Irvine, 2018). For example, Integrin-linked kinase can inhibit the upstream Hippo pathway regulator Merlin, either by inhibiting the MYPT1 phosphatase or by activation of PAK and RAC. Focal adhesion kinase (FAK) can promote activation of SRC, which has been implicated in both Hippo-pathway-dependent and Hippo-pathway-independent mechanisms for promoting YAP/TAZ activity. FAK has also previously been reported to promote YAP/TAZ activity at least in part through activation of PI3K and subsequent activation of PDK1, and FAK has been linked to dephosphorylation of YAP mediated through the protein phosphatase PP1A. These alternative mechanisms for focal adhesion-mediated regulation of YAP/TAZ might explain why Meng et al. (2018) observed that deletion of RAP2 only blocked about half of the YAP/TAZdependent changes in transcription induced by altered substrate stiffness. Activation of PI3K might also contribute to the reduced PI(4,5)P2 observed under high stiffness, and consistent with this possibility, Meng et al. (2018) report that an inhibitor of FAK could increase activation of RAP2. YAP/TAZ are widely activated in diverse solid tumors, and when Meng et al. (2018) examined contributions of the RAP2 pathway to tumorigenesis, they observed effects consistent with its regulation of YAP/TAZ. Thus, loss of RAP2 contributed to increased tumor growth in xenograft assays. RAP2 deletion also suppressed differentiation of adipocytes from mesenchymal stem cells, which is the expected consequence for a treatment that induces nuclear localization of YAP/TAZ (Dupont et al., 2011). While Meng et al. (2018) focus on regulation of YAP/TAZ by substrate stiffness, it is worth noting that the RAP2
pathway they characterized might also contribute to regulation of YAP/TAZ by other treatments that influence focal adhesions, such as changes in substrate attachment and cell shape. The RAP2 pathway does not, however, appear to contribute significantly to regulation of YAP/TAZ by cell density, which instead is sensitive to biomechanical regulation through cell-cell junctions. Future investigations should reveal how widely the RAP2 pathway is deployed for biomechanical regulation of Hippo signaling. REFERENCES Codelia, V.A., Sun, G., and Irvine, K.D. (2014). Regulation of YAP by mechanical strain through Jnk and Hippo signaling. Curr. Biol. 24, 2012–2017. de Rooij, J., Boenink, N.M., van Triest, M., Cool, R.H., Wittinghofer, A., and Bos, J.L. (1999). PDZGEF1, a guanine nucleotide exchange factor specific for Rap1 and Rap2. J. Biol. Chem. 274, 38125–38130. Dupont, S., Morsut, L., Aragona, M., Enzo, E., Giulitti, S., Cordenonsi, M., Zanconato, F., Le Digabel, J., Forcato, M., Bicciato, S., et al. (2011). Role of YAP/TAZ in mechanotransduction. Nature 474, 179–183. Gloerich, M., ten Klooster, J.P., Vliem, M.J., Koorman, T., Zwartkruis, F.J., Clevers, H., and Bos, J.L. (2012). Rap2A links intestinal cell polarity to brush border formation. Nat. Cell Biol. 14, 793–801. Humphrey, J.D., Dufresne, E.R., and Schwartz, M.A. (2014). Mechanotransduction and extracellular matrix homeostasis. Nat. Rev. Mol. Cell Biol. 15, 802–812. Li, Q., Nirala, N.K., Nie, Y., Chen, H.-J., Ostroff, G., Mao, J., Wang, Q., Xu, L., and Ip, Y.T. (2018). Ingestion of food particles regulates the mechanosensing Misshapen-Yorkie pathway in Drosophila intestinal growth. Dev. Cell 45, 433–449.e6. Meng, Z., Qiu, Y., Lin, K.C., Kumar, A., Placone, J.K., Fang, C., Wang, K.C., Lu, S., Pan, M., Hong, A.W., et al. (2018). RAP2 mediates mechanoresponses of the Hippo pathway. Nature 560, 655–660. Misra, J.R., and Irvine, K.D. (2018). The Hippo signaling network and its biological functions. Annu. Rev. Genet. 52, 3.1–3.23. Taira, K., Umikawa, M., Takei, K., Myagmar, B.E., Shinzato, M., Machida, N., Uezato, H., Nonaka, S., and Kariya, K. (2004). The Traf2- and Nck-interacting kinase as a putative effector of Rap2 to regulate actin cytoskeleton. J. Biol. Chem. 279, 49488–49496. Zhang, X., Chattopadhyay, A., Ji, Q.S., Owen, J.D., Ruest, P.J., Carpenter, G., and Hanks, S.K. (1999). Focal adhesion kinase promotes phospholipase C-gamma1 activity. Proc. Natl. Acad. Sci. USA 96, 9021–9026.
Developmental Cell 46, September 24, 2018 679
Previews of nonenveloped virus fusogens that utilize several motifs (e.g., amphipathic helices, transmembrane domain, membrane destabilizing fusion peptides) to drive cell-cell fusion. Do similar motifs serve similar functions in Myomerger? Lastly, are there other examples of multicomponent fusion complexes that assign hemifusion and pore formation activities to separate proteins? Such may be the case, for example, with vaccinia virus whose fusion complex comprises 11 small proteins. REFERENCES Bi, P., Ramirez-Martinez, A., Li, H., Cannavino, J., McAnally, J.R., Shelton, J.M., Sa´nchez-Ortiz, E., Bassel-Duby, R., and Olson, E.N. (2017). Control
of muscle formation by the fusogenic micropeptide myomixer. Science 356, 323–327. Ciechonska, M., and Duncan, R. (2014). Reovirus FAST proteins: virus-encoded cellular fusogens. Trends Microbiol. 22, 715–724. Harrison, S.C. (2015). Viral membrane fusion. Virology 479-480, 498–507. Herna´ndez, J.M., and Podbilewicz, B. (2017). The hallmarks of cell-cell fusion. Development 144, 4481–4495.
Millay, D.P., O’Rourke, J.R., Sutherland, L.B., Bezprozvannaya, S., Shelton, J.M., Bassel-Duby, R., and Olson, E.N. (2013). Myomaker is a membrane activator of myoblast fusion and muscle formation. Nature 499, 301–305. Quinn, M.E., Goh, Q., Kurosaka, M., Gamage, D.G., Petrany, M.J., Prasad, V., and Millay, D.P. (2017). Myomerger induces fusion of non-fusogenic cells and is required for skeletal muscle development. Nat. Commun. 8, 15665.
Kim, J.H., Jin, P., Duan, R., and Chen, E.H. (2015). Mechanisms of myoblast fusion during muscle development. Curr. Opin. Genet. Dev. 32, 162–170.
Weber, T., Zemelman, B.V., McNew, J.A., Westermann, B., Gmachl, M., Parlati, F., So¨llner, T.H., and Rothman, J.E. (1998). SNAREpins: minimal machinery for membrane fusion. Cell 92, 759–772.
Leikina, E., Gamage, D.G., Prasad, V., Goykhberg, J., Crowe, M., Diao, J., Kozlov, M.M., Chernomordik, L.V., and Millay, D.P. (2018). Myomaker and Myomerger work independently to control distinct steps of membrane remodeling during myoblast fusion. Dev. Cell 46, this issue, 767–780.
Zhang, Q., Vashisht, A.A., O’Rourke, J., Corbel, S.Y., Moran, R., Romero, A., Miraglia, L., Zhang, J., Durrant, E., Schmedt, C., et al. (2017). The microprotein Minion controls cell fusion and muscle formation. Nat. Commun. 8, 15664.
Rapping about Mechanotransduction Consuelo Ibar1 and Kenneth D. Irvine1,* 1Waksman Institute and Department of Molecular Biology and Biochemistry, Rutgers University, Piscataway, NJ 08854, USA *Correspondence: [email protected] https://doi.org/10.1016/j.devcel.2018.09.007
Mechanical cues can regulate cell proliferation and differentiation through the Hippo-YAP signaling network. Reporting in Nature, Meng et al. (2018) show that the Ras-related GTPase RAP2 connects extracellular matrix stiffness to Hippo pathway regulation, adding to our understanding of how mechanical cues are converted into changes in YAP activity. The Hippo signaling network integrates biochemical and biomechanical cues to influence organ growth and cell fate (reviewed in Misra and Irvine, 2018). Its downstream effects are mediated by inactivation of the transcriptional co-activator proteins YAP and TAZ, through regulation of their stability and nuclear localization. YAP/TAZ are active in the nucleus, where they can induce the transcription of genes that promote cell proliferation. Mechanical cues including changes in cell shape, extracellular matrix (ECM) stiffness, cell stretching, cell density, and shear forces have all been observed to influence YAP/TAZ activity, often through effects on the levels and organization of F-actin, and tension within the actin cytoskeleton. Meng et al. (2018) have now identified the Ras-related GTPase RAP2 as a key component of the
signaling axis that connects ECM stiffness to regulation of YAP/TAZ. Their findings stem from their discovery that overexpression of RAP2A induced a cytoplasmic localization of YAP/TAZ even within cells grown on a stiff substrate, which normally have nuclear YAP/ TAZ. Conversely, loss of all three RAP2 paralogs (RAP2A, RAP2B, RAP2C) results in nuclear localization of YAP/TAZ even within cells grown on a soft substrate, which normally have cytoplasmic YAP/ TAZ. Like other small GTPases, RAP2 regulates molecular events by cycling between an inactive GDP-bound form and an active GTP-bound form. Meng and colleagues (2018) observed that low stiffness promoted binding of GTP to RAP2. This effect is mediated through previously identified activators of RAP2, PDZGEF1, and PDZGEF2, which act as guanine
678 Developmental Cell 46, September 24, 2018 ª 2018 Elsevier Inc.
nucleotide exchange factors (GEFs) for RAP2 (de Rooij et al., 1999). Moreover, PDZGEF1/2 are required for inactivation of YAP/TAZ under low stiffness. Prior studies had identified phosphatidic acid (PA) as an activator of PDZGEF and RAP2 (Gloerich et al., 2012). Consistent with this role, Meng and colleagues (2018) were able to identify influences of substrate stiffness on phospholipid metabolism and link them to regulation of PDZGEF, RAP2, and YAP/TAZ. Using a GFP reporter, they observed that low stiffness increased the cellular levels of Phosphatidylinositol-4,5bisphosphate (PI(4,5)P2), which can be converted to PA by phospholipases D1 and D2 (PLD1/2). Previous studies had linked focal adhesions to downregulation of PI(4,5)P2 through activation of PLCg1 (Zhang et al., 1999). Indeed, Meng and colleagues (2018) observed that inhibition
Developmental Cell
Previews or knockdown of PLCg1 or PLD1/2 influenced YAP/TAZ localization. ECM stiffness can influence the number and character of focal adhesions (Humphrey et al., 2014), suggesting that modulation of RAP2 activity through downregulation of PI(4,5)P2 at focal adhesions could transmit biomechanical signaling initiated by substrate stiffness. The authors also addressed the question of how RAP2 impinges on YAP/TAZ. Whether substrate stiffness acts through Hippo-dependent or Hippo-independent signaling mechanisms has remained a point of controversy since the initial discovery of biomechanical regulation of YAP/TAZ (Dupont et al., 2011). Meng et al. (2018) provide further support for Hippo pathway-dependent regulation of YAP/TAZ by ECM stiffness (Codelia et al., 2014). For example, they show that deletion of the kinases LATS1 and LATS2, which are the direct upstream regulators of YAP/TAZ within the Hippo pathway, abrogates the cytoplasmic localization of YAP normally induced by soft substrates. They also analyze changes in the transcriptome induced by substrate stiffness and show that most of the transcriptional changes induced by altered stiffness are dependent upon both YAP/TAZ and LATS1/2. MAP4K4 is one of several kinases in the Ste20 family that contribute to activation of LATS kinases, and also a previously identified downstream effector of RAP2. Meng et al. (2018) provide evidence for a RAP2-dependent activation of MAP4K4 by low stiffness. Intriguingly, a homolog of MAP4K4, Msn, has been implicated in biomechanical regulation of Hippo signaling in the Drosophila intestine (Li et al., 2018). A related RAP2 effector, TNIK/MAP4K7, has previously been identified as a regulator of F-actin (Taira et al., 2004), and F-actin levels can modulate YAP/TAZ activity. Another class of RAP2 effector is ARHGAP29, which acts as a GTPase-activating protein (GAP) for Rho. Rho has a well-established role in promoting YAP/TAZ activity, and Meng et al. (2018) now confirm that ARHGAP29 contributes to inhibition of RhoA by low stiffness. Deletion of either MAP4K4/6/7 or ARHGAP29 alone did not prevent
inhibition of YAP/TAZ by RAP2 or by low stiffness, but their combined deletion blocked inhibition of YAP/TAZ, indicating that YAP/TAZ regulation by stiffness is effected through parallel action of these downstream effectors of RAP2. Several mechanisms for focal adhesion-mediated regulation of YAP/TAZ have been described previously, including control of kinases that localize to and are activated at focal adhesions (reviewed in Misra and Irvine, 2018). For example, Integrin-linked kinase can inhibit the upstream Hippo pathway regulator Merlin, either by inhibiting the MYPT1 phosphatase or by activation of PAK and RAC. Focal adhesion kinase (FAK) can promote activation of SRC, which has been implicated in both Hippo-pathway-dependent and Hippo-pathway-independent mechanisms for promoting YAP/TAZ activity. FAK has also previously been reported to promote YAP/TAZ activity at least in part through activation of PI3K and subsequent activation of PDK1, and FAK has been linked to dephosphorylation of YAP mediated through the protein phosphatase PP1A. These alternative mechanisms for focal adhesion-mediated regulation of YAP/TAZ might explain why Meng et al. (2018) observed that deletion of RAP2 only blocked about half of the YAP/TAZdependent changes in transcription induced by altered substrate stiffness. Activation of PI3K might also contribute to the reduced PI(4,5)P2 observed under high stiffness, and consistent with this possibility, Meng et al. (2018) report that an inhibitor of FAK could increase activation of RAP2. YAP/TAZ are widely activated in diverse solid tumors, and when Meng et al. (2018) examined contributions of the RAP2 pathway to tumorigenesis, they observed effects consistent with its regulation of YAP/TAZ. Thus, loss of RAP2 contributed to increased tumor growth in xenograft assays. RAP2 deletion also suppressed differentiation of adipocytes from mesenchymal stem cells, which is the expected consequence for a treatment that induces nuclear localization of YAP/TAZ (Dupont et al., 2011). While Meng et al. (2018) focus on regulation of YAP/TAZ by substrate stiffness, it is worth noting that the RAP2
pathway they characterized might also contribute to regulation of YAP/TAZ by other treatments that influence focal adhesions, such as changes in substrate attachment and cell shape. The RAP2 pathway does not, however, appear to contribute significantly to regulation of YAP/TAZ by cell density, which instead is sensitive to biomechanical regulation through cell-cell junctions. Future investigations should reveal how widely the RAP2 pathway is deployed for biomechanical regulation of Hippo signaling. REFERENCES Codelia, V.A., Sun, G., and Irvine, K.D. (2014). Regulation of YAP by mechanical strain through Jnk and Hippo signaling. Curr. Biol. 24, 2012–2017. de Rooij, J., Boenink, N.M., van Triest, M., Cool, R.H., Wittinghofer, A., and Bos, J.L. (1999). PDZGEF1, a guanine nucleotide exchange factor specific for Rap1 and Rap2. J. Biol. Chem. 274, 38125–38130. Dupont, S., Morsut, L., Aragona, M., Enzo, E., Giulitti, S., Cordenonsi, M., Zanconato, F., Le Digabel, J., Forcato, M., Bicciato, S., et al. (2011). Role of YAP/TAZ in mechanotransduction. Nature 474, 179–183. Gloerich, M., ten Klooster, J.P., Vliem, M.J., Koorman, T., Zwartkruis, F.J., Clevers, H., and Bos, J.L. (2012). Rap2A links intestinal cell polarity to brush border formation. Nat. Cell Biol. 14, 793–801. Humphrey, J.D., Dufresne, E.R., and Schwartz, M.A. (2014). Mechanotransduction and extracellular matrix homeostasis. Nat. Rev. Mol. Cell Biol. 15, 802–812. Li, Q., Nirala, N.K., Nie, Y., Chen, H.-J., Ostroff, G., Mao, J., Wang, Q., Xu, L., and Ip, Y.T. (2018). Ingestion of food particles regulates the mechanosensing Misshapen-Yorkie pathway in Drosophila intestinal growth. Dev. Cell 45, 433–449.e6. Meng, Z., Qiu, Y., Lin, K.C., Kumar, A., Placone, J.K., Fang, C., Wang, K.C., Lu, S., Pan, M., Hong, A.W., et al. (2018). RAP2 mediates mechanoresponses of the Hippo pathway. Nature 560, 655–660. Misra, J.R., and Irvine, K.D. (2018). The Hippo signaling network and its biological functions. Annu. Rev. Genet. 52, 3.1–3.23. Taira, K., Umikawa, M., Takei, K., Myagmar, B.E., Shinzato, M., Machida, N., Uezato, H., Nonaka, S., and Kariya, K. (2004). The Traf2- and Nck-interacting kinase as a putative effector of Rap2 to regulate actin cytoskeleton. J. Biol. Chem. 279, 49488–49496. Zhang, X., Chattopadhyay, A., Ji, Q.S., Owen, J.D., Ruest, P.J., Carpenter, G., and Hanks, S.K. (1999). Focal adhesion kinase promotes phospholipase C-gamma1 activity. Proc. Natl. Acad. Sci. USA 96, 9021–9026.
Developmental Cell 46, September 24, 2018 679