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Actin-binding channels
I.D. Neumann and R. Landgraf (Eds.) Progress in Brain Research, Vol. 170 ISSN 0079-6123 Copyright r 2008 Elsevier B.V. All rights reserved
CHAPTER 42
Actin-binding channels Yumi Noda1,2, and Sei Sasaki1 1
Department of Nephrology, Graduate School of Medicine, Tokyo Medical and Dental University, 1-5-45 Yushima, Bunkyo-ku, Tokyo, Japan 2 COE Program for Brain Integration and its Disorders, Graduate School of Medicine, Tokyo Medical and Dental University, 1-5-45 Yushima, Bunkyo-ku, Tokyo, Japan
Abstract: Channel proteins play essential roles in physiology including ion and volume homeostasis and signal transduction and in pathophysiology of many human diseases. Recently, the importance of the actin cytoskeleton in the channel protein regulation has been highlighted. The actin cytoskeleton and its reorganization have been reported to be required for the regulation of both channel activity and its intracellular trafficking. Furthermore, there are increasing evidences showing the direct interactions of channel to actin. This review focuses on actin-binding channel proteins and the role of the actin interaction in the channel protein regulation. Keywords: actin remodelling; membrane trafficking; aquaporin
been reported to bind to actin directly: ClC2 and short isoform of ClC3 (sClC3) chloride channel, vacuolar type H+-ATPase (V-ATPase), voltagedependent anion channel (VDAC) and a subunit of epithelial sodium channel (ENaC). In this review, we describe current findings indicating actin-binding channels and the role of the actin interaction in the channel regulation.
Introduction Channel proteins are essential for a variety of physiological and pathophysiological processes including maintenance of intra- and extracellular ionic gradients, volume homeostasis and signal transduction. There are two targets for channel regulation: channel activity and its intracellular localization. For both ways of regulation, the importance of the actin cytoskeleton was shown using actin-modulating drugs. Furthermore, recent development of molecular and proteomic approaches has revealed the direct interaction of several channels to actin itself. We previously showed that aquaporin-2 (AQP2) directly binds to actin (Noda et al., 2004b, 2005; Noda and Sasaki, 2006). To date, in addition to AQP2, other five channels have
ClC2 and sClC3 chloride channel ClC2 and sClC3 (short ClC3 isoform) are hypotonic cell swelling-sensitive channels belonging to a voltage-regulated chloride channel family (Duan et al., 1997; Furukawa et al., 1998; Jentsch et al., 1999). The activation of these channels contributes to the maintenance of physiological cell volume. These channels have different biophysical properties: ClC2 exhibits inward rectification, whereas sClC3 exhibits outward rectification. However,
Corresponding author. Tel.: +81-3-5803-5214; Fax: +81-3-5803-5215; E-mail: [email protected]
DOI: 10.1016/S0079-6123(08)00442-1
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actin remodelling is reported to be involved in the activation of both channels (Ahmed et al., 2000; Wang et al., 2005). Furthermore, it has been demonstrated that ClC2 N terminus and sClC3 C terminus directly bind to actin, respectively (Ahmed et al., 2000; McCloskey et al., 2007). Jentsch and co-workers show that mutations in the N-terminus and in the the cytoplasmic loop between transmembrane spans 7 and 8 of ClC2 lead to a constitutive activation with a loss of swelling- and voltage-sensitivity and suggest that the interaction between these domains inhibits the channel opening (Gru¨nder et al., 1992; Jordt and Jentsch, 1997). Bear and co-workers show that the N terminal domain fused to glutathione S-transferase (GST) is capable of binding actin in overlay and cosedimentation assay (Ahmed et al., 2000). Furthermore, the chloride channel activity of ClC2 in Xenopus oocyte is enhanced by actin-disrupting agents cytochalasin and latrunculin. Although the binding of the whole molecule of ClC2 to actin is not confirmed, these findings suggest that the intramolecular association between the N-terminus and the loop between transmembrane spans 7 and 8 of ClC2 may be stabilized through associations with the actin cytoskeleton, which results in the channel inactivation. Yamboliev and co-workers show a strong binding between cytosolic C terminus of sClC3 fused with GST (GST-sClC3-CT) and F-actin by cosedimentation assays (McCloskey et al., 2007). Inhibition of the interaction by synthetic peptides corresponding to the binding domain leads to a reduced hypotonic activation of sClC3. The C terminus of sClC3 also contains a tandem repeat of cystathionineb-synthase (CBS)-like domains, which have been proposed to affect multimerization and sorting of proteins, channel gating and ligand binding (Ignoul and Eggermont, 2005). It is speculated that F-actin binding to the C terminus of sClC3 may induce ordered folding of the CBS tandem and lead to the hypotonic activation of sClC3.
V-ATPase Vacuolar type H+-ATPase (V-ATPase) is the most versatile proton pump, highly conserved among all
eukaryotic organisms, and expressed in the intracellular membrane systems and in the plasma membrane (Nishi and Forgac, 2002). V-ATPase is essential for cellular pH homeostasis and creates an electrochemical proton gradient. Acidification of organelles by V-ATPase is critical for many cellular processes including neurotransmitter uptake into synaptic vesicles (Beyenbach and Wieczorek, 2006). V-ATPase also is able to acidify the extracelllular compartment that serves a number of roles such as bone reabsorption by osteoclasts and urinary acidification (Beyenbach and Wieczorek, 2006). V-ATPase is composed of a catalytic ATP-hydrolysing V1 complex residing on the cytoplasmic side of the membrane, and a membrane-bound protontranslocating V0 complex. The V1 complex contains eight different subunits, A–H, whereas the V0 complex consists of four different subunits a and c–e (Merzendorfer et al., 2000). V-ATPase activity is down-regulated by reversible dissociation of the V1 complex from the membrane and during this process subunit C gets lost from the V1 complex (Kane, 1995, 2000; Sumner et al., 1995; Merzendorfer et al., 2000). However, subunit C is necessary for reassembly of the two complexes into a functional holoenzyme. There are several reports showing the interactions of V-ATPase with actin. During osteoclast activation, V-ATPase directly binds to F-actin with high affinity and this association is correlated with V-ATPase transport from the cytoplasm to the plasma membrane (Lee et al., 1999). Because the complex with F-actin and V-ATPase also contains myosin II, Holliday and co-workers suggest that myosin II-powered contraction of microfilaments transports V-ATPase to the site of nascent ruffled membranes. The same group identifies an F-actin binding site in the amino-terminal domain of the subunit B (Holliday et al., 2000). In addition to subunit B, subunit C, which gets released during reversible dissociation of the holoenzyme, is also shown to bind to F-actin (Vitavska et al., 2003). Furthermore, Vitavska et al. (2005) demonstrate that subunit C binds not only to F-actin but also to monomeric G-actin. Subunit C has at least two actin-binding sites in its N- and C-terminal halves and is shown to cross-link actin filaments. This interaction may be involved in stabilising V-ATPase
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in its assembled state because subunit C appears to bridge the V1 with the V0 complex (Inoue and Forgac, 2005). Recently, recombinant subunit C is shown to be phosphorylated by protein kinase A (PKA; Voss et al., 2007). Furthermore, subunit C is phosphorylated by cAMP-analogue 8-CPTcAMP and the neurohormone serotonin in salivary glands of the blowfly Calliphora vicina in which V-ATPase reassembly and activity is regulated by serotonin via PKA (Voss et al., 2007). Voss et al. suggest that subunit C phosphorylation may facilitate the reassembly to an active V1V0 holoenzyme. In addition, subunit C interaction with actin may be regulated by its phosphorylation and be involved in the regulation of V-ATPase.
VDAC VDAC is a channel-forming protein in the mitochondrial outer membrane responsible for metabolic flux through that membrane (Colombini, 2004). VDAC has been implicated in the initiation of the mitochondrially mediated pathway of apoptosis. Although the mechanism of action is still controversial, closure of the VDAC and the subsequent decrease in metabolic flux may cause the permeabilization of the outer membrane to relatively small proteins, leading to apoptosis (Rostovtseva et al., 2005). The gating of VDAC is influenced by a variety of proteins including Bcl-xL (Vander Heiden et al., 2001), heat shock protein mtHSP70 (Schwarzer et al., 2002), dynein light chain (Schwarzer et al., 2002) and G-actin (Xu et al., 2001). G-actin induces the channel closure, and DNase-I, a protein that binds tightly to actin, blocks this effect (Xu et al., 2001). Surface plasmon resonance experiments show the direct binding of VDAC to G-actin (Roman et al., 2006). How this interaction regulates VDAC function remains unclear.
ENaC Amiloride-sensitive ENaC functions in the movement of sodium across epithelial cells in a variety
of tissues and is importantly involved in the control of extracellular fluid volume and blood pressure (Snyder, 2002). Several lines of evidence have indicated the involvement of actin in the regulation of ENaC. Patch-clamp experiments showed that ENaC activity in cell-attached patches from A6 cells was increased by cytochalasin D and short actin filaments, respectively (Cantiello et al., 1991). Short actin filaments increased open probability of abgENaC reconstituted in planar lipid bilayers (Berdiev et al., 1996). Using a series of truncation and deletion mutants of aENaC reconstituted in planar lipid bilayers, Copeland et al. (2001) showed that the region between residues 631 and 644 in the carboxy terminus was important for the regulation by actin. The involvement of actin in the intracellular trafficking of ENaC has also been reported. In mouse cortical collecting duct epithelial cells, an actin-depolymerizing agent latrunculin A inhibits ENaC exocytosis (Butterworth et al., 2005). Mazzochi et al. (2006a, b) show the direct interaction of aENaC with actin. In MDCK cells stably expressing abgENaC, aENaC is colocalized with cortical F-actin and F-actin is coimmunoprecipitated with aENaC. Gel overlay assays and cosedimentation assays show that F-actin binds directly and specifically to the C terminus of aENaC. The colocalization of aENaC and F-actin in the subapical cytoplasm suggests that in addition to regulating ENaC activity, the direct interaction between F-actin and ENaC may function in the intracellular trafficking of ENaC from a subapical pool to the plasma membrane.
AQP2 AQP2 is the predominant vasopressin-regulated water channel (Fushimi et al., 1993). Trafficking of AQP2 to the apical membrane and its vasopressin and PKA-dependent regulation in renal collecting ducts is critical for body water homeostasis (Noda and Sasaki, 2006). There are many reports showing the involvement of actin in the intracellular trafficking of AQP2. Forskolin and okadaic
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acid stimulate AQP2 translocation by inducing a reorganization of the apical actin network, respectively (Valenti et al., 2000). Actin depolymerization is a prerequisite for cAMP-dependent translocation of AQP2 (Klussmann et al., 2001; Tamma et al., 2001). These findings indicate a role of the actin depolymerization in the vasopressininduced translocation of AQP2 from intracellular vesicles to the cell surface. Furthermore, RhoA inhibition is required for AQP2 translocation (Tamma et al., 2003a). Stimulation of prostaglandin E3 receptors inhibits vasopressin-induced Rho inactivation, vasopressin-induced F-actin depolymerization, and vasopressin-, cAMP- and forskolin-induced AQP2 translocation (Tamma et al., 2003b). Rho activation by Bradykinin stabilizes cortical F-actin and inhibits AQP2 trafficking (Tamma et al., 2005). Not only inhibitory effects but also facilitatory effects of F-actin assembly on AQP2 trafficking have been suggested. Actin-depolymerizing agent cytochalasin D did not enhance water permeability of the epithelium of toad urinary bladder (Franki et al., 1992). The authors speculate that F-actin has at least two hypothetical pools: one is involved in the barrier function and the other pool is involved in the transport of AQP2. Tajika et al. (2005) show that actin depolymerization caused by cytochalasin D or latrunculin B inhibits AQP2 translocation from EEA1-positive early endosomes to Rab11-positive subapical storage vesicles. Furthermore, vasopressin signalling induces myosin light chain phosphorylation which is known to enhance myosin–actin filament interaction and the formation of actin fibres (Chou et al., 2004). Recently, myosin is shown to be critical for AQP2 recycling (Nedvetsky et al., 2007). Since both stimulatory and inhibitory effects of actin assembly are observed in AQP2 trafficking, regulation of actin remodelling may be different by time and localization during the translocation. In the process that the physiological trafficking of AQP2 occurs in vivo, the actin dynamics may change in a restricted narrow area around AQP2 molecule and may vary among different areas within the cell.
Recently, we have clarified that AQP2 directly binds to actin and SPA-1 (Noda et al., 2004a, b, 2005). SPA-1 is a GTPase-activating protein (GAP) for Rap1 and the GAP activity of SPA-1 is required for AQP2 trafficking to the apical membrane. Since Rap1 affects the assembly of F-actin (Tsukamoto et al., 1999; Pak et al., 2001; Harazaki et al., 2004; Kometani et al., 2004; Noda and Sasaki, 2006), SPA-1 binding to AQP2 may reduce the levels of Rap1GTP that trigger F-actin disassembly in a restricted area, resulting in the promotion of the AQP2 trafficking. Indeed, AQP2 trafficking to the apical membrane is impaired in the collecting duct principal cells of SPA-1 deficient mice (Noda et al., 2004a). SPA-1 deficient mice show marked bilateral hydronephrosis due to polyuria (Noda et al., 2004a; Kometani et al., 2006). In addition to actin and SPA-1, AQP2 is shown to form a complex with 11 proteins: ionized calcium binding adapter molecule 2, myosin regulatory light chain smooth muscle isoforms 2-A and 2-B, a-tropomyosin 5b, annexin A2 and A6, scinderin, gelsolin, a-actinin 4, a-II spectrin and myosin heavy chain nonmuscle type A (Noda et al., 2005). Since these proteins have actin binding abilities, each interaction of these proteins in the complex may be dynamic and this dynamic assembly acts as a key point for the regulation of the AQP2 trafficking. To clarify the biophysical mechanisms which provide force driving AQP2 movement, we are now examining the interaction dynamics of AQP2 with these binding proteins at the resolution of single molecule during AQP2 translocation.
Concluding remarks In recent years evidence has accumulated to demonstrate the direct binding of channel proteins to actin. However, the roles of the direct interaction in the regulation of many channels have not been confirmed. There are several reasons for the difficulty in examining the roles. For example, high concentrations of actin molecules in the cell make the microscopic colocalization assessment difficult.
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Actin-modulating agents inevitably affect the overall cell architecture which makes the experiments examining the roles of actin interaction in channel regulation difficult. The roles and the regulation mechanisms will be uncovered by application of time lapse imaging with high spatiotemporal resolution and other methods that enable to assess the real-time molecular dynamics in a small area in live cells.
Acknowledgement We thank Dr. Saburo Horikawa for useful discussions and comments.
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Actin-binding channels Yumi Noda1,2, and Sei Sasaki1 1
Department of Nephrology, Graduate School of Medicine, Tokyo Medical and Dental University, 1-5-45 Yushima, Bunkyo-ku, Tokyo, Japan 2 COE Program for Brain Integration and its Disorders, Graduate School of Medicine, Tokyo Medical and Dental University, 1-5-45 Yushima, Bunkyo-ku, Tokyo, Japan
Abstract: Channel proteins play essential roles in physiology including ion and volume homeostasis and signal transduction and in pathophysiology of many human diseases. Recently, the importance of the actin cytoskeleton in the channel protein regulation has been highlighted. The actin cytoskeleton and its reorganization have been reported to be required for the regulation of both channel activity and its intracellular trafficking. Furthermore, there are increasing evidences showing the direct interactions of channel to actin. This review focuses on actin-binding channel proteins and the role of the actin interaction in the channel protein regulation. Keywords: actin remodelling; membrane trafficking; aquaporin
been reported to bind to actin directly: ClC2 and short isoform of ClC3 (sClC3) chloride channel, vacuolar type H+-ATPase (V-ATPase), voltagedependent anion channel (VDAC) and a subunit of epithelial sodium channel (ENaC). In this review, we describe current findings indicating actin-binding channels and the role of the actin interaction in the channel regulation.
Introduction Channel proteins are essential for a variety of physiological and pathophysiological processes including maintenance of intra- and extracellular ionic gradients, volume homeostasis and signal transduction. There are two targets for channel regulation: channel activity and its intracellular localization. For both ways of regulation, the importance of the actin cytoskeleton was shown using actin-modulating drugs. Furthermore, recent development of molecular and proteomic approaches has revealed the direct interaction of several channels to actin itself. We previously showed that aquaporin-2 (AQP2) directly binds to actin (Noda et al., 2004b, 2005; Noda and Sasaki, 2006). To date, in addition to AQP2, other five channels have
ClC2 and sClC3 chloride channel ClC2 and sClC3 (short ClC3 isoform) are hypotonic cell swelling-sensitive channels belonging to a voltage-regulated chloride channel family (Duan et al., 1997; Furukawa et al., 1998; Jentsch et al., 1999). The activation of these channels contributes to the maintenance of physiological cell volume. These channels have different biophysical properties: ClC2 exhibits inward rectification, whereas sClC3 exhibits outward rectification. However,
Corresponding author. Tel.: +81-3-5803-5214; Fax: +81-3-5803-5215; E-mail: [email protected]
DOI: 10.1016/S0079-6123(08)00442-1
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actin remodelling is reported to be involved in the activation of both channels (Ahmed et al., 2000; Wang et al., 2005). Furthermore, it has been demonstrated that ClC2 N terminus and sClC3 C terminus directly bind to actin, respectively (Ahmed et al., 2000; McCloskey et al., 2007). Jentsch and co-workers show that mutations in the N-terminus and in the the cytoplasmic loop between transmembrane spans 7 and 8 of ClC2 lead to a constitutive activation with a loss of swelling- and voltage-sensitivity and suggest that the interaction between these domains inhibits the channel opening (Gru¨nder et al., 1992; Jordt and Jentsch, 1997). Bear and co-workers show that the N terminal domain fused to glutathione S-transferase (GST) is capable of binding actin in overlay and cosedimentation assay (Ahmed et al., 2000). Furthermore, the chloride channel activity of ClC2 in Xenopus oocyte is enhanced by actin-disrupting agents cytochalasin and latrunculin. Although the binding of the whole molecule of ClC2 to actin is not confirmed, these findings suggest that the intramolecular association between the N-terminus and the loop between transmembrane spans 7 and 8 of ClC2 may be stabilized through associations with the actin cytoskeleton, which results in the channel inactivation. Yamboliev and co-workers show a strong binding between cytosolic C terminus of sClC3 fused with GST (GST-sClC3-CT) and F-actin by cosedimentation assays (McCloskey et al., 2007). Inhibition of the interaction by synthetic peptides corresponding to the binding domain leads to a reduced hypotonic activation of sClC3. The C terminus of sClC3 also contains a tandem repeat of cystathionineb-synthase (CBS)-like domains, which have been proposed to affect multimerization and sorting of proteins, channel gating and ligand binding (Ignoul and Eggermont, 2005). It is speculated that F-actin binding to the C terminus of sClC3 may induce ordered folding of the CBS tandem and lead to the hypotonic activation of sClC3.
V-ATPase Vacuolar type H+-ATPase (V-ATPase) is the most versatile proton pump, highly conserved among all
eukaryotic organisms, and expressed in the intracellular membrane systems and in the plasma membrane (Nishi and Forgac, 2002). V-ATPase is essential for cellular pH homeostasis and creates an electrochemical proton gradient. Acidification of organelles by V-ATPase is critical for many cellular processes including neurotransmitter uptake into synaptic vesicles (Beyenbach and Wieczorek, 2006). V-ATPase also is able to acidify the extracelllular compartment that serves a number of roles such as bone reabsorption by osteoclasts and urinary acidification (Beyenbach and Wieczorek, 2006). V-ATPase is composed of a catalytic ATP-hydrolysing V1 complex residing on the cytoplasmic side of the membrane, and a membrane-bound protontranslocating V0 complex. The V1 complex contains eight different subunits, A–H, whereas the V0 complex consists of four different subunits a and c–e (Merzendorfer et al., 2000). V-ATPase activity is down-regulated by reversible dissociation of the V1 complex from the membrane and during this process subunit C gets lost from the V1 complex (Kane, 1995, 2000; Sumner et al., 1995; Merzendorfer et al., 2000). However, subunit C is necessary for reassembly of the two complexes into a functional holoenzyme. There are several reports showing the interactions of V-ATPase with actin. During osteoclast activation, V-ATPase directly binds to F-actin with high affinity and this association is correlated with V-ATPase transport from the cytoplasm to the plasma membrane (Lee et al., 1999). Because the complex with F-actin and V-ATPase also contains myosin II, Holliday and co-workers suggest that myosin II-powered contraction of microfilaments transports V-ATPase to the site of nascent ruffled membranes. The same group identifies an F-actin binding site in the amino-terminal domain of the subunit B (Holliday et al., 2000). In addition to subunit B, subunit C, which gets released during reversible dissociation of the holoenzyme, is also shown to bind to F-actin (Vitavska et al., 2003). Furthermore, Vitavska et al. (2005) demonstrate that subunit C binds not only to F-actin but also to monomeric G-actin. Subunit C has at least two actin-binding sites in its N- and C-terminal halves and is shown to cross-link actin filaments. This interaction may be involved in stabilising V-ATPase
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in its assembled state because subunit C appears to bridge the V1 with the V0 complex (Inoue and Forgac, 2005). Recently, recombinant subunit C is shown to be phosphorylated by protein kinase A (PKA; Voss et al., 2007). Furthermore, subunit C is phosphorylated by cAMP-analogue 8-CPTcAMP and the neurohormone serotonin in salivary glands of the blowfly Calliphora vicina in which V-ATPase reassembly and activity is regulated by serotonin via PKA (Voss et al., 2007). Voss et al. suggest that subunit C phosphorylation may facilitate the reassembly to an active V1V0 holoenzyme. In addition, subunit C interaction with actin may be regulated by its phosphorylation and be involved in the regulation of V-ATPase.
VDAC VDAC is a channel-forming protein in the mitochondrial outer membrane responsible for metabolic flux through that membrane (Colombini, 2004). VDAC has been implicated in the initiation of the mitochondrially mediated pathway of apoptosis. Although the mechanism of action is still controversial, closure of the VDAC and the subsequent decrease in metabolic flux may cause the permeabilization of the outer membrane to relatively small proteins, leading to apoptosis (Rostovtseva et al., 2005). The gating of VDAC is influenced by a variety of proteins including Bcl-xL (Vander Heiden et al., 2001), heat shock protein mtHSP70 (Schwarzer et al., 2002), dynein light chain (Schwarzer et al., 2002) and G-actin (Xu et al., 2001). G-actin induces the channel closure, and DNase-I, a protein that binds tightly to actin, blocks this effect (Xu et al., 2001). Surface plasmon resonance experiments show the direct binding of VDAC to G-actin (Roman et al., 2006). How this interaction regulates VDAC function remains unclear.
ENaC Amiloride-sensitive ENaC functions in the movement of sodium across epithelial cells in a variety
of tissues and is importantly involved in the control of extracellular fluid volume and blood pressure (Snyder, 2002). Several lines of evidence have indicated the involvement of actin in the regulation of ENaC. Patch-clamp experiments showed that ENaC activity in cell-attached patches from A6 cells was increased by cytochalasin D and short actin filaments, respectively (Cantiello et al., 1991). Short actin filaments increased open probability of abgENaC reconstituted in planar lipid bilayers (Berdiev et al., 1996). Using a series of truncation and deletion mutants of aENaC reconstituted in planar lipid bilayers, Copeland et al. (2001) showed that the region between residues 631 and 644 in the carboxy terminus was important for the regulation by actin. The involvement of actin in the intracellular trafficking of ENaC has also been reported. In mouse cortical collecting duct epithelial cells, an actin-depolymerizing agent latrunculin A inhibits ENaC exocytosis (Butterworth et al., 2005). Mazzochi et al. (2006a, b) show the direct interaction of aENaC with actin. In MDCK cells stably expressing abgENaC, aENaC is colocalized with cortical F-actin and F-actin is coimmunoprecipitated with aENaC. Gel overlay assays and cosedimentation assays show that F-actin binds directly and specifically to the C terminus of aENaC. The colocalization of aENaC and F-actin in the subapical cytoplasm suggests that in addition to regulating ENaC activity, the direct interaction between F-actin and ENaC may function in the intracellular trafficking of ENaC from a subapical pool to the plasma membrane.
AQP2 AQP2 is the predominant vasopressin-regulated water channel (Fushimi et al., 1993). Trafficking of AQP2 to the apical membrane and its vasopressin and PKA-dependent regulation in renal collecting ducts is critical for body water homeostasis (Noda and Sasaki, 2006). There are many reports showing the involvement of actin in the intracellular trafficking of AQP2. Forskolin and okadaic
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acid stimulate AQP2 translocation by inducing a reorganization of the apical actin network, respectively (Valenti et al., 2000). Actin depolymerization is a prerequisite for cAMP-dependent translocation of AQP2 (Klussmann et al., 2001; Tamma et al., 2001). These findings indicate a role of the actin depolymerization in the vasopressininduced translocation of AQP2 from intracellular vesicles to the cell surface. Furthermore, RhoA inhibition is required for AQP2 translocation (Tamma et al., 2003a). Stimulation of prostaglandin E3 receptors inhibits vasopressin-induced Rho inactivation, vasopressin-induced F-actin depolymerization, and vasopressin-, cAMP- and forskolin-induced AQP2 translocation (Tamma et al., 2003b). Rho activation by Bradykinin stabilizes cortical F-actin and inhibits AQP2 trafficking (Tamma et al., 2005). Not only inhibitory effects but also facilitatory effects of F-actin assembly on AQP2 trafficking have been suggested. Actin-depolymerizing agent cytochalasin D did not enhance water permeability of the epithelium of toad urinary bladder (Franki et al., 1992). The authors speculate that F-actin has at least two hypothetical pools: one is involved in the barrier function and the other pool is involved in the transport of AQP2. Tajika et al. (2005) show that actin depolymerization caused by cytochalasin D or latrunculin B inhibits AQP2 translocation from EEA1-positive early endosomes to Rab11-positive subapical storage vesicles. Furthermore, vasopressin signalling induces myosin light chain phosphorylation which is known to enhance myosin–actin filament interaction and the formation of actin fibres (Chou et al., 2004). Recently, myosin is shown to be critical for AQP2 recycling (Nedvetsky et al., 2007). Since both stimulatory and inhibitory effects of actin assembly are observed in AQP2 trafficking, regulation of actin remodelling may be different by time and localization during the translocation. In the process that the physiological trafficking of AQP2 occurs in vivo, the actin dynamics may change in a restricted narrow area around AQP2 molecule and may vary among different areas within the cell.
Recently, we have clarified that AQP2 directly binds to actin and SPA-1 (Noda et al., 2004a, b, 2005). SPA-1 is a GTPase-activating protein (GAP) for Rap1 and the GAP activity of SPA-1 is required for AQP2 trafficking to the apical membrane. Since Rap1 affects the assembly of F-actin (Tsukamoto et al., 1999; Pak et al., 2001; Harazaki et al., 2004; Kometani et al., 2004; Noda and Sasaki, 2006), SPA-1 binding to AQP2 may reduce the levels of Rap1GTP that trigger F-actin disassembly in a restricted area, resulting in the promotion of the AQP2 trafficking. Indeed, AQP2 trafficking to the apical membrane is impaired in the collecting duct principal cells of SPA-1 deficient mice (Noda et al., 2004a). SPA-1 deficient mice show marked bilateral hydronephrosis due to polyuria (Noda et al., 2004a; Kometani et al., 2006). In addition to actin and SPA-1, AQP2 is shown to form a complex with 11 proteins: ionized calcium binding adapter molecule 2, myosin regulatory light chain smooth muscle isoforms 2-A and 2-B, a-tropomyosin 5b, annexin A2 and A6, scinderin, gelsolin, a-actinin 4, a-II spectrin and myosin heavy chain nonmuscle type A (Noda et al., 2005). Since these proteins have actin binding abilities, each interaction of these proteins in the complex may be dynamic and this dynamic assembly acts as a key point for the regulation of the AQP2 trafficking. To clarify the biophysical mechanisms which provide force driving AQP2 movement, we are now examining the interaction dynamics of AQP2 with these binding proteins at the resolution of single molecule during AQP2 translocation.
Concluding remarks In recent years evidence has accumulated to demonstrate the direct binding of channel proteins to actin. However, the roles of the direct interaction in the regulation of many channels have not been confirmed. There are several reasons for the difficulty in examining the roles. For example, high concentrations of actin molecules in the cell make the microscopic colocalization assessment difficult.
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Actin-modulating agents inevitably affect the overall cell architecture which makes the experiments examining the roles of actin interaction in channel regulation difficult. The roles and the regulation mechanisms will be uncovered by application of time lapse imaging with high spatiotemporal resolution and other methods that enable to assess the real-time molecular dynamics in a small area in live cells.
Acknowledgement We thank Dr. Saburo Horikawa for useful discussions and comments.
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