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Mini Review
Extracellular signaling cues for nuclear actin polymerization Matthias Plessner, Robert Grosse ∗ Institute of Pharmacology, Biochemical-Pharmacological Center (BPC), University of Marburg, Karl-von-Frisch-Str. 1, 35043 Marburg, Germany
a r t i c l e
i n f o
Keywords: Nuclear actin Myocardin-related transcription factor Formin LINC complex
a b s t r a c t Contrary to cytoplasmic actin structures, the biological functions of nuclear actin filaments remain largely enigmatic. Recent progress in the field, however, has determined nuclear actin structures in somatic cells either under steady state conditions or in response to extracellular signaling cues. These actin structures differ in size and shape as well as in their temporal appearance and dynamics. Thus, a picture emerges that suggests that mammalian cells may have different pathways and mechanisms to assemble nuclear actin filaments. Apart from serum- or LPA-triggered nuclear actin polymerization, integrin activation by extracellular matrix interaction was recently implicated in nuclear actin polymerization through the linker of nucleoskeleton and cytoskeleton (LINC) complex. Some of these extracellular cues known so far appear to converge at the level of nuclear formin activity and subsequent regulation of myocardin-related transcription factors. Nevertheless, as the precise signaling events are as yet unknown, the regulation of nuclear actin polymerization may be of significant importance for different cellular functions as well as disease conditions caused by altered nuclear dynamics and architecture. © 2015 Elsevier GmbH. All rights reserved.
1. Introduction Actin is one of the most abundant proteins in the cytoplasm and is an essential player in a plethora of cellular processes such as cellular shape, adhesion, cell motility and cytokinesis (Rottner and Stradal, 2011). Generally, actin can be found in two different states. Monomeric globular actin, termed G-actin, constitutes the G-actin pool from which in turn polymeric filamentous F-actin is assembled (Pollard and Cooper, 2009). Actin filaments display structural polarity because G-actin monomers within a filament are oriented in the same direction. Based on their appearance in electron microscopy, the terminal part is referred to as either barbed or pointed end (Bonder et al., 1983). Actin nucleation constitutes the initial step in the formation of microfilaments with formins, the Arp2/3 complex and Spire proteins being the three major classes of actin nucleators known to date. Arp2/3 and Spire both bind to pointed ends of actin filaments. While the Arp2/3 complex polymerizes actin filaments into branched arrays, Spire has the ability to assemble linear filaments (Kerkhoff, 2006). Likewise, formins polymerize linear actin filaments through processive association with the barbed end (Faix and Grosse, 2006) via their conserved formin homology (FH) 2 domain. For this, two FH2 domains form a circular head-to-tail
homo-dimer thereby stabilizing actin dimers and adding them to the barbed end in a stair-stepping process, while binding of actin by the FH1 domain increases the local G-actin concentration to accelerate actin polymerization (Campellone and Welch, 2010). Among the different classes of formins, diaphanous-related formins (DRFs) are best characterized (Bogdan et al., 2013). DRFs display a modular domain organization in which the regulatory segment is composed of a GTPase binding domain (GBD) and a diaphanous-inhibitory-domain (DID), both of which are involved in the autoinhibitory regulation of DRFs. The FH2 domain is located at the C-terminus together with the diaphanous-autoregulatorydomain (DAD). In the inactive conformation, DAD binds DID to achieve autoinhibition, while binding of a Rho GTPase was shown to release autoinhibition (Lammers et al., 2005; Kuhn and Geyer, 2014). In addition, it is now appreciated that the DID–DAD autoinhibitory module is further influenced by other cellular signaling processes such as serine/threonine kinases (Kuhn and Geyer, 2014). Many proteins such as cofilin, profilin, or capping proteins affect actin dynamics and can therefore influence the rate of actin assembly as well as the formation of secondary structures of F-actin such as actin bundles, networks of branched actin filaments or the actomyosin ring responsible for cytokinesis (Pollard and Cooper, 2009). 2. Nuclear actin and extracellular cues
∗ Corresponding author. E-mail address: [email protected] (R. Grosse).
The structure and function of actin within the mammalian cell nucleus is much less clear and hence under intense investigation
http://dx.doi.org/10.1016/j.ejcb.2015.05.009 0171-9335/© 2015 Elsevier GmbH. All rights reserved.
Please cite this article in press as: Plessner, M., Grosse, R., Extracellular signaling cues for nuclear actin polymerization. Eur. J. Cell Biol. (2015), http://dx.doi.org/10.1016/j.ejcb.2015.05.009
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Fig. 1. Visualization of native nuclear actin filaments by STED microscopy. Starved NIH 3T3 fibroblasts were stimulated with fetal calf serum (FCS) for 20 s and immediately fixed with glutaraldehyde. Endogenous F-actin was stained using phalloidin (ATTO 647N-conjugated). The sample was visualized with 633 nm pulsed excitation (confocal) and 775 nm pulsed STED (200 ps pulse width). The image was taken at the MPI for Biophysical Chemistry, Dept. of NanoBiophotonics, Göttingen.
(Belin and Mullins, 2013; Grosse and Vartiainen, 2013). One central issue is to overcome the obstacles in visualizing nuclear actin because its concentration in somatic cell nuclei is very low compared to other models utilized in nuclear actin research, i.e. Xenopus oocyte nuclei (Bohnsack et al., 2006; Miyamoto and Gurdon, 2011; Miyamoto et al., 2011). Therefore, it was previously thought that in somatic cells nuclear actin exists mainly or even only in its monomeric form (de Lanerolle and Serebryannyy, 2011; Percipalle, 2013). However, our view on nuclear actin structures is currently changing due to recently described dynamic F-actin foci (Belin and Mullins, 2013) as well as filamentous networks, which can rapidly form upon serum or lysophosphatidic acid (LPA) stimulation (Baarlink et al., 2013) evidently independent of the Arp2/3 complex. In these cases, nuclear actin was visualized by targeting actin probes such as Utrophin or LifeAct to the nucleus, thereby circumventing strong signal interference from cytoplasmic F-actin labeling (Baarlink and Grosse, 2014). Importantly, signalregulated, rapid nuclear F-actin network assembly was dependent of mDia formins and could be demonstrated without ectopic protein expression using phalloidin labeling, demonstrating the principle existence of native nuclear actin filaments and higher order structures in somatic cell nuclei (Fig. 1) (Baarlink et al., 2013). 3. MRTF-A (MAL, MKL1) is regulated by nuclear actin polymerization As part of the immediate serum response, nuclear F-actin dynamics are directly linked to myocardin-related transcription factor A (MRTF-A) (also termed megakaryocytic acute leukemia; MAL, or megakaryoblastic leukemia 1; MKL1), which acts as a critical transcriptional coactivator of the serum response factor (SRF) (Olson and Nordheim, 2010). MRTF-A is an actin-binding protein that continuously and rapidly shuttles between the cytoplasmic and nuclear compartment. This shuttling is regulated by compartmentalized actin polymerization (Miralles et al., 2003). G-actin binding to the RPEL domain of MRTF-A is necessary for nuclear export of MRTF-A to the cytoplasm while in turn
binding of cytoplasmic actin interferes with access to the nuclear localization signal (NLS) of MRTF-A. Upon release from G-actin, MRTF-A cannot be exported from the nucleus and thus is predestined for SRF-mediated transcription (Treisman, 2013). Therefore, in order to inactivate MRTF-A/SRF activity, MRTF-A requires nuclear actin binding to re-translocate to the cytoplasm (Vartiainen et al., 2007). We recently demonstrated a mechanism in which nuclear formin-dependent polymerization of actin constitutes a critical step efficiently preventing nuclear export of MRTF-A in response to extracellular signals (Baarlink et al., 2013; Baarlink and Grosse, 2014; Esnault et al., 2014). To study the functional role of endogenous nuclear formins, we generated a genetically encoded, light-switchable tool for DRF activation by fusing the LOV (light, oxygen, or voltage) J␣-domain of Avena sativa phototropin-1 (Niopek et al., 2014; Renicke et al., 2013; Wu et al., 2009) to the DAD region of mDia2. Thus, by light-induced uncaging of a DAD domain for binding to endogenous DID-containing DRFs we could spatiotemporally unleash mDia autoinhibition (Fig. 2) resulting in a reversible induction of long and unbranched nuclear actin filaments (Fig. 2). In addition, MRTF-A relocated from the cytoplasm to the nuclear compartment, demonstrating the importance and efficiency of nuclear actin polymerization for MRTF-A regulation (Fig. 3) (Baarlink et al., 2013).
4. Mechanosensing and nuclear actin dynamics MRTF transcription factors have been recognized to be involved in mechanotransduction or regulation of tension homeostasis by less well-defined mechanisms (Somogyi and Rorth, 2004; McGee et al., 2011; Janmey et al., 2013; Chan et al., 2010; Iyer et al., 2012). Mechanotransduction to the nucleus involves proteins of the nuclear envelope, which communicate with the cytoskeleton. Located in the inner and outer nuclear membrane (INM and ONM) of the nucleus, linker of nucleoskeleton and cytoskeleton (LINC) complexes are recognized for coupling of mechanical signals from the cytoplasm into the nucleus. Force or tension can be transmitted through these cytoskeletal components leading to changes in gene expression (Simon and Wilson, 2011). Fundamental components of LINC complexes are different isoforms of nesprins (nuclear envelope spectrin-repeat proteins) in the ONM and Sun (Sad1 and UNC-84) domain-containing proteins, which are located in the INM. Nesprins contain KASH (Klarsicht, ANC-1 and SYNE homology) domains, essential for binding Sun proteins (especially Sun1 and Sun2). They can also bind the aforementioned cytoskeletal components, either directly or by a variety of linker proteins (Simon and Wilson, 2011). Sun proteins and nesprins represent the core of LINC complexes, as three nesprins bind to a Sun trimer (Sosa et al., 2012). Inside the nucleus, different proteins are able to interact with LINC complexes. Two of these proteins are emerin and A-type lamins, which are thought to be responsible for mechanotransduction. Mutations in genes coding for lamin A/C or emerin can disrupt nuclear envelope stability causing diseases like laminopathies and Emery–Dreifuss muscular dystrophy. When considering nuclear actin filaments, emerin has been reported to interact with nuclear myosins (Simon and Wilson, 2011), while A-type lamins and emerin were shown to affect MRTFA function through actin presumably via modulating the state of nuclear actin polymerization (Ho et al., 2013). Furthermore, matrix elasticity and lamin A/C have been linked to SRF transcriptional activity and the control of actomyosin genes (Buxboim et al., 2014). Consistent with such a scenario, we could recently show that nuclear actin polymerization is mediated by integrin activation during cell spreading requiring the presence of lamin A/C and emerin (Plessner et al., 2015). Moreover, integrin-triggered or cell
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Fig. 2. Nuclear actin network assembly upon activation of nuclear mDia by a photoactivatable DAD (nuc.LOV-DAD). NIH3T3 cells were transfected with genetically encoded nuclear Actin-Chromobody (nAC, green, (Plessner et al., 2015)) and mCherry.nuc.LOV-DAD (red) and subjected to confocal live cell imaging. The depicted cell was repeatedly irradiated with a 488 nm laser to simultaneously activate nuc.LOV-DAD and visualize nuclear actin dynamics.
Fig. 3. Signaling pathways from the cell surface to nuclear actin dynamics.
spreading-induced nuclear F-actin formation fully depends on a functional LINC complex as assessed by silencing Sun1/2, while in turn, cell spreading promoted MRTF-A nuclear localization and activation downstream of nuclear formin activity (Plessner et al., 2015). Thus, different extracellular stimuli can induce nuclear actin assembly, which in the case of serum/LPA or integrin activation appear to converge on nuclear mDia1/2 function (Fig. 3).
However, many questions clearly still remain, such as how are these signals propagated through the cytoplasm, and how are formins finally activated in the nucleus? Alternatively, formins such as mDia2 may be also imported in a pre-activated state (Miki et al., 2009) with rapid nucleocytoplasmic shuttling being possibly a critical parameter for compartmental formin activation and inactivation.
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A study by Guilluy et al. (2014) on isolated nuclei showed that the stiffness of the nucleus adapts to force arguing for a novel mechanotransduction pathway. In this study, a pulling force is directly applied on LINC complexes of isolated nuclei using magnetic beads coated with anti-nesprin-1 antibodies, thereby mimicking force by cytoskeletal filaments. This application of force causes an activation of the small GTPase RhoA in biochemical nuclear fractions (Guilluy et al., 2014). Hence, activation of a potentially nuclear residing RhoA could trigger downstream effector systems such as formins inside the nuclear compartment (Staus et al., 2014). Another very recent study showed that nuclear shuttling of Rac1 triggers nuclear actin dynamics, which in turn would act on the nuclear envelop to deform cancer cell nuclei for efficient invasion and migratory plasticity (Navarro-Lerida et al., 2015). 5. Concluding remarks Research on nuclear actin has recently progressed due to advantages in visualizing nuclear actin structures, i.e. by using actin probes specifically targeted to the nuclear compartment. Thus, polymeric nuclear actin structures were detected in a variety of different cellular contexts and the list is likely to grow in the near future. Current strategies need to further emphasize the dynamic regulation of nuclear actin, which may be much more comparable to cytoskeletal actin as previously assumed, also since most actin-regulating factors undergo nucleocytoplasmic shuttling and can hence be detected in the cell nucleus (Grosse and Vartiainen, 2013). Thus, addressing the functional implications of nuclear actin polymerization will be of pivotal importance. The growing diversity of nuclear actin structures will likely provide exciting new insights into nuclear regulation of cell behavior. Acknowledgments We thank Haisen Ta and Stefan Hell (MPI for Biophysical Chemistry, Dept. of NanoBiophotonics, Göttingen) for the image in Fig. 1. This work was supported by the SFB 593 and GR2111/7-1 (DFG). References Baarlink, C., Grosse, R., 2014. Formin’ actin in the nucleus. Nucleus 5 (1), 15–20. Baarlink, C., Wang, H., Grosse, R., 2013. Nuclear actin network assembly by formins regulates the SRF coactivator MAL. Science 340 (6134), 864–867. Belin, B.J., Mullins, R.D., 2013. What we talk about when we talk about nuclear actin. Nucleus 4 (4), 291–297. Bogdan, S., Schultz, J., Grosshans, J., 2013. Formin’ cellular structures: physiological roles of diaphanous (Dia) in actin dynamics. Commun. Integr. Biol. 6 (6), e27634. Bohnsack, M.T., et al., 2006. A selective block of nuclear actin export stabilizes the giant nuclei of Xenopus oocytes. Nat. Cell Biol. 8 (3), 257–263. Bonder, E.M., Fishkind, D.J., Mooseker, M.S., 1983. Direct measurement of critical concentrations and assembly rate constants at the two ends of an actin filament. Cell 34 (2), 491–501. Buxboim, A., et al., 2014. Matrix elasticity regulates lamin-A C phosphorylation and turnover with feedback to actomyosin. Curr. Biol. 24 (16), 1909–1917. Campellone, K.G., Welch, M.D., 2010. A nucleator arms race: cellular control of actin assembly. Nat. Rev. Mol. Cell Biol. 11 (4), 237–251. Chan, M., Chaudary, F., Lee, W., Copeland, J., McCulloch, C., 2010. Force-induced Myofibroblast Differentiation through Collagen Receptors Is Dependent on
Mammalian Diaphanous (mDia). J. Biol. Chem. 285 (12), 9273–9281. de Lanerolle, P., Serebryannyy, L., 2011. Nuclear actin and myosins: life without filaments. Nat. Cell Biol. 13 (11), 1282–1288. Esnault, C., et al., 2014. Rho-actin signaling to the MRTF coactivators dominates the immediate transcriptional response to serum in fibroblasts. Genes Dev. 28 (9), 943–958. Faix, J., Grosse, R., 2006. Staying in shape with formins. Dev. Cell 10 (6), 693–706. Grosse, R., Vartiainen, M.K., 2013. To be or not to be assembled: progressing into nuclear actin filaments. Nat. Rev. Mol. Cell Biol. 14 (11), 693–697. Guilluy, C., et al., 2014. Isolated nuclei adapt to force and reveal a mechanotransduction pathway in the nucleus. Nat. Cell Biol. 16 (4), 376–381. Ho, C.Y., et al., 2013. Lamin A/C and emerin regulate MKL1-SRF activity by modulating actin dynamics. Nature 497 (7450), 507–511. Iyer, K., Pulford, S., Mogilner, A., Shivashankar, G., 2012. Mechanical activation of cells induces chromatin remodeling preceding MKL nuclear transport. Biophys. J. 103 (7), 1416–1428. Janmey, P.A., et al., 2013. From tissue mechanics to transcription factors. Differentiation 86 (3), 112–120. Kerkhoff, E., 2006. Cellular functions of the Spir actin-nucleation factors. Trends Cell Biol. 16 (9), 477–483. Kuhn, S., Geyer, M., 2014. Formins as effector proteins of Rho GTPases. Small GTPases 5. Lammers, M., et al., 2005. The regulation of mDia1 by autoinhibition and its release by Rho*GTP. EMBO J. 24 (23), 4176–4187. McGee, K.M., et al., 2011. Nuclear transport of the serum response factor coactivator MRTF-A is downregulated at tensional homeostasis. EMBO Rep. 12 (9), 963–970. Miki, T., et al., 2009. mDia2 shuttles between the nucleus and the cytoplasm through the importin-{alpha}/{beta}- and CRM1-mediated nuclear transport mechanism. J. Biol. Chem. 284 (9), 5753–5762. Miralles, F., et al., 2003. Actin dynamics control SRF activity by regulation of its coactivator MAL. Cell 113 (3), 329–342. Miyamoto, K., Gurdon, J.B., 2011. Nuclear actin and transcriptional activation. Commun. Integr. Biol. 4 (5), 582–583. Miyamoto, K., et al., 2011. Nuclear actin polymerization is required for transcriptional reprogramming of Oct4 by oocytes. Genes Dev. 25 (9), 946–958. Navarro-Lerida, I., et al., 2015. Rac1 nucleocytoplasmic shuttling drives nuclear shape changes and tumor invasion. Dev. Cell 32 (3), 318–334. Niopek, D., et al., 2014. Engineering light-inducible nuclear localization signals for precise spatiotemporal control of protein dynamics in living cells. Nat. Commun. 5, 4404. Olson, E.N., Nordheim, A., 2010. Linking actin dynamics and gene transcription to drive cellular motile functions. Nat. Rev. Mol. Cell Biol. 11 (5), 353–365. Percipalle, P., 2013. Co-transcriptional nuclear actin dynamics. Nucleus 4 (1), 43–52. Plessner, M., et al., 2015. Nuclear F-actin formation and reorganization upon cell spreading. J. Biol. Chem. 290 (18), 11209–11216, http://dx.doi.org/10.1074/jbc. M114.627166, Epub 2015 Mar 10. Pollard, T.D., Cooper, J.A., 2009. Actin, a central player in cell shape and movement. Science 326 (5957), 1208–1212. Renicke, C., et al., 2013. A LOV2 domain-based optogenetic tool to control protein degradation and cellular function. Chem. Biol. 20 (4), 619–626. Rottner, K., Stradal, T.E., 2011. Actin dynamics and turnover in cell motility. Curr. Opin. Cell Biol. 23 (5), 569–578. Simon, D.N., Wilson, K.L., 2011. The nucleoskeleton as a genome-associated dynamic ‘network of networks’. Nat. Rev. Mol. Cell Biol. 12 (11), 695–708. Somogyi, K., Rorth, P., 2004. Evidence for tension-based regulation of Drosophila MAL and SRF during invasive cell migration. Dev. Cell 7 (1), 85–93. Sosa, B.A., et al., 2012. LINC complexes form by binding of three KASH peptides to domain interfaces of trimeric SUN proteins. Cell 149 (5), 1035–1047. Staus, D.P., et al., 2014. Nuclear RhoA signaling regulates MRTF-dependent SMC-specific transcription. Am. J. Physiol. Heart Circ. Physiol. 307 (3), H379–H390. Treisman, R., 2013. Shedding light on nuclear actin dynamics and function. Trends Biochem. Sci. 38 (8), 376–377. Vartiainen, M.K., et al., 2007. Nuclear actin regulates dynamic subcellular localization and activity of the SRF cofactor MAL. Science 316 (5832), 1749–1752. Wu, Y.I., et al., 2009. A genetically encoded photoactivatable Rac controls the motility of living cells. Nature 461 (7260), 104–108.
Please cite this article in press as: Plessner, M., Grosse, R., Extracellular signaling cues for nuclear actin polymerization. Eur. J. Cell Biol. (2015), http://dx.doi.org/10.1016/j.ejcb.2015.05.009
ARTICLE IN PRESS European Journal of Cell Biology xxx (2015) xxx–xxx
Contents lists available at ScienceDirect
European Journal of Cell Biology journal homepage: www.elsevier.com/locate/ejcb
Mini Review
Extracellular signaling cues for nuclear actin polymerization Matthias Plessner, Robert Grosse ∗ Institute of Pharmacology, Biochemical-Pharmacological Center (BPC), University of Marburg, Karl-von-Frisch-Str. 1, 35043 Marburg, Germany
a r t i c l e
i n f o
Keywords: Nuclear actin Myocardin-related transcription factor Formin LINC complex
a b s t r a c t Contrary to cytoplasmic actin structures, the biological functions of nuclear actin filaments remain largely enigmatic. Recent progress in the field, however, has determined nuclear actin structures in somatic cells either under steady state conditions or in response to extracellular signaling cues. These actin structures differ in size and shape as well as in their temporal appearance and dynamics. Thus, a picture emerges that suggests that mammalian cells may have different pathways and mechanisms to assemble nuclear actin filaments. Apart from serum- or LPA-triggered nuclear actin polymerization, integrin activation by extracellular matrix interaction was recently implicated in nuclear actin polymerization through the linker of nucleoskeleton and cytoskeleton (LINC) complex. Some of these extracellular cues known so far appear to converge at the level of nuclear formin activity and subsequent regulation of myocardin-related transcription factors. Nevertheless, as the precise signaling events are as yet unknown, the regulation of nuclear actin polymerization may be of significant importance for different cellular functions as well as disease conditions caused by altered nuclear dynamics and architecture. © 2015 Elsevier GmbH. All rights reserved.
1. Introduction Actin is one of the most abundant proteins in the cytoplasm and is an essential player in a plethora of cellular processes such as cellular shape, adhesion, cell motility and cytokinesis (Rottner and Stradal, 2011). Generally, actin can be found in two different states. Monomeric globular actin, termed G-actin, constitutes the G-actin pool from which in turn polymeric filamentous F-actin is assembled (Pollard and Cooper, 2009). Actin filaments display structural polarity because G-actin monomers within a filament are oriented in the same direction. Based on their appearance in electron microscopy, the terminal part is referred to as either barbed or pointed end (Bonder et al., 1983). Actin nucleation constitutes the initial step in the formation of microfilaments with formins, the Arp2/3 complex and Spire proteins being the three major classes of actin nucleators known to date. Arp2/3 and Spire both bind to pointed ends of actin filaments. While the Arp2/3 complex polymerizes actin filaments into branched arrays, Spire has the ability to assemble linear filaments (Kerkhoff, 2006). Likewise, formins polymerize linear actin filaments through processive association with the barbed end (Faix and Grosse, 2006) via their conserved formin homology (FH) 2 domain. For this, two FH2 domains form a circular head-to-tail
homo-dimer thereby stabilizing actin dimers and adding them to the barbed end in a stair-stepping process, while binding of actin by the FH1 domain increases the local G-actin concentration to accelerate actin polymerization (Campellone and Welch, 2010). Among the different classes of formins, diaphanous-related formins (DRFs) are best characterized (Bogdan et al., 2013). DRFs display a modular domain organization in which the regulatory segment is composed of a GTPase binding domain (GBD) and a diaphanous-inhibitory-domain (DID), both of which are involved in the autoinhibitory regulation of DRFs. The FH2 domain is located at the C-terminus together with the diaphanous-autoregulatorydomain (DAD). In the inactive conformation, DAD binds DID to achieve autoinhibition, while binding of a Rho GTPase was shown to release autoinhibition (Lammers et al., 2005; Kuhn and Geyer, 2014). In addition, it is now appreciated that the DID–DAD autoinhibitory module is further influenced by other cellular signaling processes such as serine/threonine kinases (Kuhn and Geyer, 2014). Many proteins such as cofilin, profilin, or capping proteins affect actin dynamics and can therefore influence the rate of actin assembly as well as the formation of secondary structures of F-actin such as actin bundles, networks of branched actin filaments or the actomyosin ring responsible for cytokinesis (Pollard and Cooper, 2009). 2. Nuclear actin and extracellular cues
∗ Corresponding author. E-mail address: [email protected] (R. Grosse).
The structure and function of actin within the mammalian cell nucleus is much less clear and hence under intense investigation
http://dx.doi.org/10.1016/j.ejcb.2015.05.009 0171-9335/© 2015 Elsevier GmbH. All rights reserved.
Please cite this article in press as: Plessner, M., Grosse, R., Extracellular signaling cues for nuclear actin polymerization. Eur. J. Cell Biol. (2015), http://dx.doi.org/10.1016/j.ejcb.2015.05.009
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Fig. 1. Visualization of native nuclear actin filaments by STED microscopy. Starved NIH 3T3 fibroblasts were stimulated with fetal calf serum (FCS) for 20 s and immediately fixed with glutaraldehyde. Endogenous F-actin was stained using phalloidin (ATTO 647N-conjugated). The sample was visualized with 633 nm pulsed excitation (confocal) and 775 nm pulsed STED (200 ps pulse width). The image was taken at the MPI for Biophysical Chemistry, Dept. of NanoBiophotonics, Göttingen.
(Belin and Mullins, 2013; Grosse and Vartiainen, 2013). One central issue is to overcome the obstacles in visualizing nuclear actin because its concentration in somatic cell nuclei is very low compared to other models utilized in nuclear actin research, i.e. Xenopus oocyte nuclei (Bohnsack et al., 2006; Miyamoto and Gurdon, 2011; Miyamoto et al., 2011). Therefore, it was previously thought that in somatic cells nuclear actin exists mainly or even only in its monomeric form (de Lanerolle and Serebryannyy, 2011; Percipalle, 2013). However, our view on nuclear actin structures is currently changing due to recently described dynamic F-actin foci (Belin and Mullins, 2013) as well as filamentous networks, which can rapidly form upon serum or lysophosphatidic acid (LPA) stimulation (Baarlink et al., 2013) evidently independent of the Arp2/3 complex. In these cases, nuclear actin was visualized by targeting actin probes such as Utrophin or LifeAct to the nucleus, thereby circumventing strong signal interference from cytoplasmic F-actin labeling (Baarlink and Grosse, 2014). Importantly, signalregulated, rapid nuclear F-actin network assembly was dependent of mDia formins and could be demonstrated without ectopic protein expression using phalloidin labeling, demonstrating the principle existence of native nuclear actin filaments and higher order structures in somatic cell nuclei (Fig. 1) (Baarlink et al., 2013). 3. MRTF-A (MAL, MKL1) is regulated by nuclear actin polymerization As part of the immediate serum response, nuclear F-actin dynamics are directly linked to myocardin-related transcription factor A (MRTF-A) (also termed megakaryocytic acute leukemia; MAL, or megakaryoblastic leukemia 1; MKL1), which acts as a critical transcriptional coactivator of the serum response factor (SRF) (Olson and Nordheim, 2010). MRTF-A is an actin-binding protein that continuously and rapidly shuttles between the cytoplasmic and nuclear compartment. This shuttling is regulated by compartmentalized actin polymerization (Miralles et al., 2003). G-actin binding to the RPEL domain of MRTF-A is necessary for nuclear export of MRTF-A to the cytoplasm while in turn
binding of cytoplasmic actin interferes with access to the nuclear localization signal (NLS) of MRTF-A. Upon release from G-actin, MRTF-A cannot be exported from the nucleus and thus is predestined for SRF-mediated transcription (Treisman, 2013). Therefore, in order to inactivate MRTF-A/SRF activity, MRTF-A requires nuclear actin binding to re-translocate to the cytoplasm (Vartiainen et al., 2007). We recently demonstrated a mechanism in which nuclear formin-dependent polymerization of actin constitutes a critical step efficiently preventing nuclear export of MRTF-A in response to extracellular signals (Baarlink et al., 2013; Baarlink and Grosse, 2014; Esnault et al., 2014). To study the functional role of endogenous nuclear formins, we generated a genetically encoded, light-switchable tool for DRF activation by fusing the LOV (light, oxygen, or voltage) J␣-domain of Avena sativa phototropin-1 (Niopek et al., 2014; Renicke et al., 2013; Wu et al., 2009) to the DAD region of mDia2. Thus, by light-induced uncaging of a DAD domain for binding to endogenous DID-containing DRFs we could spatiotemporally unleash mDia autoinhibition (Fig. 2) resulting in a reversible induction of long and unbranched nuclear actin filaments (Fig. 2). In addition, MRTF-A relocated from the cytoplasm to the nuclear compartment, demonstrating the importance and efficiency of nuclear actin polymerization for MRTF-A regulation (Fig. 3) (Baarlink et al., 2013).
4. Mechanosensing and nuclear actin dynamics MRTF transcription factors have been recognized to be involved in mechanotransduction or regulation of tension homeostasis by less well-defined mechanisms (Somogyi and Rorth, 2004; McGee et al., 2011; Janmey et al., 2013; Chan et al., 2010; Iyer et al., 2012). Mechanotransduction to the nucleus involves proteins of the nuclear envelope, which communicate with the cytoskeleton. Located in the inner and outer nuclear membrane (INM and ONM) of the nucleus, linker of nucleoskeleton and cytoskeleton (LINC) complexes are recognized for coupling of mechanical signals from the cytoplasm into the nucleus. Force or tension can be transmitted through these cytoskeletal components leading to changes in gene expression (Simon and Wilson, 2011). Fundamental components of LINC complexes are different isoforms of nesprins (nuclear envelope spectrin-repeat proteins) in the ONM and Sun (Sad1 and UNC-84) domain-containing proteins, which are located in the INM. Nesprins contain KASH (Klarsicht, ANC-1 and SYNE homology) domains, essential for binding Sun proteins (especially Sun1 and Sun2). They can also bind the aforementioned cytoskeletal components, either directly or by a variety of linker proteins (Simon and Wilson, 2011). Sun proteins and nesprins represent the core of LINC complexes, as three nesprins bind to a Sun trimer (Sosa et al., 2012). Inside the nucleus, different proteins are able to interact with LINC complexes. Two of these proteins are emerin and A-type lamins, which are thought to be responsible for mechanotransduction. Mutations in genes coding for lamin A/C or emerin can disrupt nuclear envelope stability causing diseases like laminopathies and Emery–Dreifuss muscular dystrophy. When considering nuclear actin filaments, emerin has been reported to interact with nuclear myosins (Simon and Wilson, 2011), while A-type lamins and emerin were shown to affect MRTFA function through actin presumably via modulating the state of nuclear actin polymerization (Ho et al., 2013). Furthermore, matrix elasticity and lamin A/C have been linked to SRF transcriptional activity and the control of actomyosin genes (Buxboim et al., 2014). Consistent with such a scenario, we could recently show that nuclear actin polymerization is mediated by integrin activation during cell spreading requiring the presence of lamin A/C and emerin (Plessner et al., 2015). Moreover, integrin-triggered or cell
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Fig. 2. Nuclear actin network assembly upon activation of nuclear mDia by a photoactivatable DAD (nuc.LOV-DAD). NIH3T3 cells were transfected with genetically encoded nuclear Actin-Chromobody (nAC, green, (Plessner et al., 2015)) and mCherry.nuc.LOV-DAD (red) and subjected to confocal live cell imaging. The depicted cell was repeatedly irradiated with a 488 nm laser to simultaneously activate nuc.LOV-DAD and visualize nuclear actin dynamics.
Fig. 3. Signaling pathways from the cell surface to nuclear actin dynamics.
spreading-induced nuclear F-actin formation fully depends on a functional LINC complex as assessed by silencing Sun1/2, while in turn, cell spreading promoted MRTF-A nuclear localization and activation downstream of nuclear formin activity (Plessner et al., 2015). Thus, different extracellular stimuli can induce nuclear actin assembly, which in the case of serum/LPA or integrin activation appear to converge on nuclear mDia1/2 function (Fig. 3).
However, many questions clearly still remain, such as how are these signals propagated through the cytoplasm, and how are formins finally activated in the nucleus? Alternatively, formins such as mDia2 may be also imported in a pre-activated state (Miki et al., 2009) with rapid nucleocytoplasmic shuttling being possibly a critical parameter for compartmental formin activation and inactivation.
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A study by Guilluy et al. (2014) on isolated nuclei showed that the stiffness of the nucleus adapts to force arguing for a novel mechanotransduction pathway. In this study, a pulling force is directly applied on LINC complexes of isolated nuclei using magnetic beads coated with anti-nesprin-1 antibodies, thereby mimicking force by cytoskeletal filaments. This application of force causes an activation of the small GTPase RhoA in biochemical nuclear fractions (Guilluy et al., 2014). Hence, activation of a potentially nuclear residing RhoA could trigger downstream effector systems such as formins inside the nuclear compartment (Staus et al., 2014). Another very recent study showed that nuclear shuttling of Rac1 triggers nuclear actin dynamics, which in turn would act on the nuclear envelop to deform cancer cell nuclei for efficient invasion and migratory plasticity (Navarro-Lerida et al., 2015). 5. Concluding remarks Research on nuclear actin has recently progressed due to advantages in visualizing nuclear actin structures, i.e. by using actin probes specifically targeted to the nuclear compartment. Thus, polymeric nuclear actin structures were detected in a variety of different cellular contexts and the list is likely to grow in the near future. Current strategies need to further emphasize the dynamic regulation of nuclear actin, which may be much more comparable to cytoskeletal actin as previously assumed, also since most actin-regulating factors undergo nucleocytoplasmic shuttling and can hence be detected in the cell nucleus (Grosse and Vartiainen, 2013). Thus, addressing the functional implications of nuclear actin polymerization will be of pivotal importance. The growing diversity of nuclear actin structures will likely provide exciting new insights into nuclear regulation of cell behavior. Acknowledgments We thank Haisen Ta and Stefan Hell (MPI for Biophysical Chemistry, Dept. of NanoBiophotonics, Göttingen) for the image in Fig. 1. This work was supported by the SFB 593 and GR2111/7-1 (DFG). References Baarlink, C., Grosse, R., 2014. Formin’ actin in the nucleus. Nucleus 5 (1), 15–20. Baarlink, C., Wang, H., Grosse, R., 2013. Nuclear actin network assembly by formins regulates the SRF coactivator MAL. Science 340 (6134), 864–867. Belin, B.J., Mullins, R.D., 2013. What we talk about when we talk about nuclear actin. Nucleus 4 (4), 291–297. Bogdan, S., Schultz, J., Grosshans, J., 2013. Formin’ cellular structures: physiological roles of diaphanous (Dia) in actin dynamics. Commun. Integr. Biol. 6 (6), e27634. Bohnsack, M.T., et al., 2006. A selective block of nuclear actin export stabilizes the giant nuclei of Xenopus oocytes. Nat. Cell Biol. 8 (3), 257–263. Bonder, E.M., Fishkind, D.J., Mooseker, M.S., 1983. Direct measurement of critical concentrations and assembly rate constants at the two ends of an actin filament. Cell 34 (2), 491–501. Buxboim, A., et al., 2014. Matrix elasticity regulates lamin-A C phosphorylation and turnover with feedback to actomyosin. Curr. Biol. 24 (16), 1909–1917. Campellone, K.G., Welch, M.D., 2010. A nucleator arms race: cellular control of actin assembly. Nat. Rev. Mol. Cell Biol. 11 (4), 237–251. Chan, M., Chaudary, F., Lee, W., Copeland, J., McCulloch, C., 2010. Force-induced Myofibroblast Differentiation through Collagen Receptors Is Dependent on
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