Native cyclase-associated protein and actin from Xenopus laevis oocytes form a unique 4:4 complex with a tripartite structure
2021; Elsevier BV; Volume: 296; Linguagem: Inglês
10.1016/j.jbc.2021.100649
ISSN1083-351X
AutoresNoriyuki Kodera, Hiroshi Abe, Phuong Doan N. Nguyen, Shoichiro Ono,
Tópico(s)Cardiomyopathy and Myosin Studies
ResumoCyclase-associated protein (CAP) is a conserved actin-binding protein that regulates multiple aspects of actin dynamics, including polymerization, depolymerization, filament severing, and nucleotide exchange. CAP has been isolated from different cells and tissues in an equimolar complex with actin, and previous studies have shown that a CAP–actin complex contains six molecules each of CAP and actin. Intriguingly, here, we successfully isolated a complex of Xenopus cyclase-associated protein 1 (XCAP1) with actin from oocyte extracts, which contained only four molecules each of XCAP1 and actin. This XCAP1–actin complex remained stable as a single population of 340 kDa species during hydrodynamic analyses using gel filtration or analytical ultracentrifugation. Examination of the XCAP1–actin complex by high-speed atomic force microscopy revealed a tripartite structure: one middle globular domain and two globular arms. The two arms were observed in high and low states. The arms at the high state were spontaneously converted to the low state by dissociation of actin from the complex. However, when extra G-actin was added, the arms at the low state were converted to the high state. Based on the known structures of the N-terminal helical-folded domain and C-terminal CARP domain, we hypothesize that the middle globular domain corresponds to a tetramer of the N-terminal helical-folded domain of XCAP1 and that each arm in the high state corresponds to a heterotetramer containing a dimer of the C-terminal CARP domain of XCAP1 and two G-actin molecules. This novel configuration of a CAP–actin complex should help to understand how CAP promotes actin filament disassembly. Cyclase-associated protein (CAP) is a conserved actin-binding protein that regulates multiple aspects of actin dynamics, including polymerization, depolymerization, filament severing, and nucleotide exchange. CAP has been isolated from different cells and tissues in an equimolar complex with actin, and previous studies have shown that a CAP–actin complex contains six molecules each of CAP and actin. Intriguingly, here, we successfully isolated a complex of Xenopus cyclase-associated protein 1 (XCAP1) with actin from oocyte extracts, which contained only four molecules each of XCAP1 and actin. This XCAP1–actin complex remained stable as a single population of 340 kDa species during hydrodynamic analyses using gel filtration or analytical ultracentrifugation. Examination of the XCAP1–actin complex by high-speed atomic force microscopy revealed a tripartite structure: one middle globular domain and two globular arms. The two arms were observed in high and low states. The arms at the high state were spontaneously converted to the low state by dissociation of actin from the complex. However, when extra G-actin was added, the arms at the low state were converted to the high state. Based on the known structures of the N-terminal helical-folded domain and C-terminal CARP domain, we hypothesize that the middle globular domain corresponds to a tetramer of the N-terminal helical-folded domain of XCAP1 and that each arm in the high state corresponds to a heterotetramer containing a dimer of the C-terminal CARP domain of XCAP1 and two G-actin molecules. This novel configuration of a CAP–actin complex should help to understand how CAP promotes actin filament disassembly. Regulated assembly and disassembly of actin filaments are vital to the diverse function of the actin cytoskeleton (1Pollard T.D. Cooper J.A. Actin, a central player in cell shape and movement.Science. 2009; 326: 1208-1212Crossref PubMed Scopus (1265) Google Scholar). Cyclase-associated protein (CAP) is one of the actin-regulatory proteins that control multiple key aspects of actin filament dynamics (2Ono S. 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Broach J.R. SRV2, a gene required for RAS activation of adenylate cyclase in yeast.Cell. 1990; 61: 329-340Abstract Full Text PDF PubMed Scopus (189) Google Scholar). However, CAP was later recognized as an actin-binding protein in a variety of eukaryotes. CAP binds to actin monomers and inhibits polymerization (6Freeman N.L. Chen Z. Horenstein J. Weber A. Field J. An actin monomer binding activity localizes to the carboxyl-terminal half of the Saccharomyces cerevisiae cyclase-associated protein.J. Biol. Chem. 1995; 270: 5680-5685Abstract Full Text Full Text PDF PubMed Scopus (113) Google Scholar). CAP also promotes exchange of actin-bound nucleotides in competition with cofilin and increases ATP-bound actin monomers that are readily available for polymerization (7Moriyama K. Yahara I. Human CAP1 is a key factor in the recycling of cofilin and actin for rapid actin turnover.J. Cell Sci. 2002; 115: 1591-1601Crossref PubMed Google Scholar, 8Balcer H.I. Goodman A.L. Rodal A.A. Smith E. 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Aguilar R.M. Johnston A.B. Goode B.L. Species-specific functions of twinfilin in actin filament depolymerization.J. Mol. Biol. 2018; 430: 3323-3336Crossref PubMed Scopus (21) Google Scholar). CAP is involved in a number of cellular events that require actin remodeling in various cell types and tissues. For example, CAP is essential for muscle sarcomere organization in Caenorhabditis elegans (16Nomura K. Ono K. Ono S. CAS-1, a C. elegans cyclase-associated protein, is required for sarcomeric actin assembly in striated muscle.J. Cell Sci. 2012; 125: 4077-4089Crossref PubMed Scopus (20) Google Scholar) and mice (17Kepser L.J. Damar F. De Cicco T. Chaponnier C. Proszynski T.J. Pagenstecher A. Rust M.B. CAP2 deficiency delays myofibril actin cytoskeleton differentiation and disturbs skeletal muscle architecture and function.Proc. Natl. Acad. Sci. U. S. A. 2019; 116: 8397-8402Crossref PubMed Scopus (17) Google Scholar), and deficiency of CAP2, a mammalian CAP isoform, causes cardiomyopathy in mice (18Peche V. Shekar S. Leichter M. Korte H. Schroder R. Schleicher M. Holak T.A. Clemen C.S. Ramanath Y.B. Pfitzer G. Karakesisoglou I. Noegel A.A. CAP2, cyclase-associated protein 2, is a dual compartment protein.Cell Mol. Life Sci. 2007; 64: 2702-2715Crossref PubMed Scopus (57) Google Scholar, 19Field J. Ye D.Z. Shinde M. Liu F. Schillinger K.J. Lu M. Wang T. Skettini M. Xiong Y. Brice A.K. Chung D.C. Patel V.V. CAP2 in cardiac conduction, sudden cardiac death and eye development.Sci. Rep. 2015; 5: 17256Crossref PubMed Scopus (21) Google Scholar) and humans (20Aspit L. Levitas A. Etzion S. Krymko H. Slanovic L. Zarivach R. Etzion Y. Parvari R. CAP2 mutation leads to impaired actin dynamics and associates with supraventricular tachycardia and dilated cardiomyopathy.J. Med. Genet. 2019; 56: 228-235Crossref PubMed Scopus (16) Google Scholar). Intriguingly, when nonrecombinant native CAP is isolated from tissues or cells, actin is associated with CAP in a multimeric complex at an equimolar ratio and cannot be dissociated without harsh conditions. Porcine CAP (originally reported as ASP-56) was isolated from platelets as a complex with actin, and actin was finally dissociated from CAP by 3 M urea (21Gieselmann R. Mann K. ASP-56, a new actin sequestering protein from pig platelets with homology to CAP, an adenylate cyclase-associated protein from yeast.FEBS Lett. 1992; 298: 149-153Crossref PubMed Scopus (61) Google Scholar). Similar CAP–actin complex has been isolated from yeast (8Balcer H.I. Goodman A.L. Rodal A.A. Smith E. Kugler J. Heuser J.E. Goode B.L. Coordinated regulation of actin filament turnover by a high-molecular-weight Srv2/CAP complex, cofilin, profilin, and Aip1.Curr. Biol. 2003; 13: 2159-2169Abstract Full Text Full Text PDF PubMed Scopus (139) Google Scholar), bovine thymus (11Normoyle K.P. Brieher W.M. Cyclase-associated protein (CAP) acts directly on F-actin to accelerate cofilin-mediated actin severing across the range of physiological pH.J. Biol. Chem. 2012; 287: 35722-35732Abstract Full Text Full Text PDF PubMed Scopus (51) Google Scholar), and mouse brain (22Mu A. Fung T.S. Kettenbach A.N. Chakrabarti R. Higgs H.N. A complex containing lysine-acetylated actin inhibits the formin INF2.Nat. Cell Biol. 2019; 21: 592-602Crossref PubMed Scopus (29) Google Scholar). The CAP–actin complex promotes actin filament disassembly in the presence of cofilin (8Balcer H.I. Goodman A.L. Rodal A.A. Smith E. Kugler J. Heuser J.E. Goode B.L. Coordinated regulation of actin filament turnover by a high-molecular-weight Srv2/CAP complex, cofilin, profilin, and Aip1.Curr. Biol. 2003; 13: 2159-2169Abstract Full Text Full Text PDF PubMed Scopus (139) Google Scholar, 11Normoyle K.P. Brieher W.M. Cyclase-associated protein (CAP) acts directly on F-actin to accelerate cofilin-mediated actin severing across the range of physiological pH.J. Biol. Chem. 2012; 287: 35722-35732Abstract Full Text Full Text PDF PubMed Scopus (51) Google Scholar). In addition, recent studies have shown that the CAP–actin complex containing acetylated actin is an inhibitor of inverted formin 2 (INF2) (22Mu A. Fung T.S. Kettenbach A.N. Chakrabarti R. Higgs H.N. A complex containing lysine-acetylated actin inhibits the formin INF2.Nat. Cell Biol. 2019; 21: 592-602Crossref PubMed Scopus (29) Google Scholar, 23Mu A. Fung T.S. Francomacaro L.M. Huynh T. Kotila T. Svindrych Z. Higgs H.N. Regulation of INF2-mediated actin polymerization through site-specific lysine acetylation of actin itself.Proc. Natl. Acad. Sci. U. S. A. 2020; 117: 439-447Crossref PubMed Scopus (12) Google Scholar). Thus, the CAP and actin have biological functions as a complex, but how the complex is assembled and why the complex formation is important for its functions remain unknown. The native complex of yeast CAP (also known as Srv2) and actin is a 6:6 complex of ∼600 kDa (8Balcer H.I. Goodman A.L. Rodal A.A. Smith E. Kugler J. Heuser J.E. Goode B.L. Coordinated regulation of actin filament turnover by a high-molecular-weight Srv2/CAP complex, cofilin, profilin, and Aip1.Curr. Biol. 2003; 13: 2159-2169Abstract Full Text Full Text PDF PubMed Scopus (139) Google Scholar), which can be reconstituted from purified components (24Quintero-Monzon O. Jonasson E.M. Bertling E. Talarico L. Chaudhry F. Sihvo M. Lappalainen P. Goode B.L. Reconstitution and dissection of the 600-kDa Srv2/CAP complex: Roles for oligomerization and cofilin-actin binding in driving actin turnover.J. Biol. Chem. 2009; 284: 10923-10934Abstract Full Text Full Text PDF PubMed Scopus (51) Google Scholar). The CAP–actin complex from mouse brain is also in a similar size (22Mu A. Fung T.S. Kettenbach A.N. Chakrabarti R. Higgs H.N. A complex containing lysine-acetylated actin inhibits the formin INF2.Nat. Cell Biol. 2019; 21: 592-602Crossref PubMed Scopus (29) Google Scholar). The N-terminal half of yeast and mouse CAPs form a hexameric "shuriken" structure, which is mediated by oligomerization of a putative coiled-coil region at the most N-terminus (10Chaudhry F. Breitsprecher D. Little K. Sharov G. Sokolova O. Goode B.L. Srv2/cyclase-associated protein forms hexameric shurikens that directly catalyze actin filament severing by cofilin.Mol. Biol. Cell. 2013; 24: 31-41Crossref PubMed Scopus (68) Google Scholar, 25Jansen S. Collins A. Golden L. Sokolova O. Goode B.L. Structure and mechanism of mouse cyclase-associated protein (CAP1) in regulating actin dynamics.J. Biol. Chem. 2014; 289: 30732-30742Abstract Full Text Full Text PDF PubMed Scopus (37) Google Scholar) and dimerization of the helical-folded domain (HFD) (12Kotila T. Wioland H. Enkavi G. Kogan K. Vattulainen I. Jegou A. Romet-Lemonne G. Lappalainen P. Mechanism of synergistic actin filament pointed end depolymerization by cyclase-associated protein and cofilin.Nat. Comm. 2019; 10: 5320Crossref PubMed Scopus (35) Google Scholar, 26Yusof A.M. Hu N.J. Wlodawer A. Hofmann A. Structural evidence for variable oligomerization of the N-terminal domain of cyclase-associated protein (CAP).Proteins. 2005; 58: 255-262Crossref PubMed Scopus (22) Google Scholar, 27Yusof A.M. Jaenicke E. Pedersen J.S. Noegel A.A. Schleicher M. Hofmann A. Mechanism of oligomerisation of cyclase-associated protein from Dictyostelium discoideum in solution.J. Mol. Biol. 2006; 362: 1072-1081Crossref PubMed Scopus (7) Google Scholar). The C-terminal half of CAP contains a CAP and X-linked retinitis pigmentosa 2 protein (CARP) domain that dimerizes through the most C-terminal dimerization motif (28Dodatko T. Fedorov A.A. Grynberg M. Patskovsky Y. Rozwarski D.A. Jaroszewski L. Aronoff-Spencer E. Kondraskina E. Irving T. Godzik A. Almo S.C. Crystal structure of the actin binding domain of the cyclase-associated protein.Biochemistry. 2004; 43: 10628-10641Crossref PubMed Scopus (41) Google Scholar, 29Iwase S. Ono S. The C-terminal dimerization motif of cyclase-associated protein is essential for actin monomer regulation.Biochem. J. 2016; 473: 4427-4441Crossref PubMed Scopus (7) Google Scholar). The CARP domain of CAP binds to actin monomer (6Freeman N.L. Chen Z. Horenstein J. Weber A. Field J. An actin monomer binding activity localizes to the carboxyl-terminal half of the Saccharomyces cerevisiae cyclase-associated protein.J. Biol. Chem. 1995; 270: 5680-5685Abstract Full Text Full Text PDF PubMed Scopus (113) Google Scholar, 30Mattila P.K. Quintero-Monzon O. Kugler J. Moseley J.B. Almo S.C. Lappalainen P. Goode B.L. A high-affinity interaction with ADP-actin monomers underlies the mechanism and in vivo function of Srv2/cyclase-associated protein.Mol. Biol. Cell. 2004; 15: 5158-5171Crossref PubMed Scopus (80) Google Scholar, 31Iwase S. Ono S. Conserved hydrophobic residues in the CARP/β-sheet domain of cyclase-associated protein are involved in actin monomer regulation.Cytoskeleton. 2017; 74: 343-355Crossref Scopus (5) Google Scholar, 32Makkonen M. Bertling E. Chebotareva N.A. Baum J. Lappalainen P. Mammalian and malaria parasite cyclase-associated proteins catalyze nucleotide exchange on G-actin through a conserved mechanism.J. Biol. Chem. 2013; 288: 984-994Abstract Full Text Full Text PDF PubMed Scopus (48) Google Scholar), and a CARP dimer and two actin molecules form a compact globular structure (33Kotila T. Kogan K. Enkavi G. Guo S. Vattulainen I. Goode B.L. Lappalainen P. Structural basis of actin monomer re-charging by cyclase-associated protein.Nat. Comm. 2018; 9: 1892Crossref PubMed Scopus (38) Google Scholar). Although we know structures of parts of the CAP–actin complex, we still have limited knowledge on the structure of the entire complex. Furthermore, a recent study has demonstrated that the N-terminal regions of human CAP1 and CAP2 primarily form tetramers instead of hexamers (34Purde V. Busch F. Kudryashova E. Wysocki V.H. Kudryashov D.S. Oligomerization affects the ability of human cyclase-associated proteins 1 and 2 to promote actin severing by cofilins.Int. J. Mol. Sci. 2019; 20: 5647Crossref Scopus (13) Google Scholar). Therefore, whether the 6:6 configuration is conserved among CAP–actin complexes from different sources remains unknown. In this study, we purified a complex of Xenopus CAP1 and actin and demonstrated that the complex contained the two proteins in a 4:4 stoichiometric ratio, which is a novel configuration of the CAP–actin complex. We purified a native complex of CAP and actin from Xenopus oocyte extracts (Fig. 1). When Xenopus oocyte extracts were applied to a column in which glutathione S-transferase (GST)-fused Xenopus ADF/cofilin (XAC) was immobilized, several proteins specifically bound to the column as described previously (35Okada K. Obinata T. Abe H. XAIP1: A Xenopus homologue of yeast actin interacting protein 1 (AIP1), which induces disassembly of actin filaments cooperatively with ADF/cofilin family proteins.J. Cell Sci. 1999; 112: 1553-1565Crossref PubMed Google Scholar) (Fig. 1A). We reported that the 65-kDa, 42-kDa, and 19-kDa proteins were Xenopus actin-interacting protein 1 (XAIP1), actin, and XAC, respectively (35Okada K. Obinata T. Abe H. XAIP1: A Xenopus homologue of yeast actin interacting protein 1 (AIP1), which induces disassembly of actin filaments cooperatively with ADF/cofilin family proteins.J. Cell Sci. 1999; 112: 1553-1565Crossref PubMed Google Scholar). Peptide sequencing identified that the 94-kDa and 60-kDa proteins were gelsolin (36Ankenbauer T. Kleinschmidt J.A. Vandekerckhove J. Franke W.W. Proteins regulating actin assembly in oogenesis and early embryogenesis of Xenopus laevis: Gelsolin is the major cytoplasmic actin-binding protein.J. Cell Biol. 1988; 107: 1489-1498Crossref PubMed Scopus (29) Google Scholar) and cyclase-associated protein 1 (XCAP1) (37KhosrowShahian F. Hubberstey A.V. Crawford M.J. CAP1 expression is developmentally regulated in Xenopus.Mech. Dev. 2002; 113: 211-214Crossref PubMed Scopus (4) Google Scholar), respectively. We attempted to isolate XCAP1 using anion-exchange chromatography followed by hydroxyapatite chromatography, but XCAP1 and actin were not separated during these procedures and were instead purified together in an equimolar ratio (Fig. 1B). Further gel filtration chromatography using Sephadex G-200 also resulted in coelution of XCAP1 and actin in a single peak at ∼390 kDa (our unpublished observation), which is much larger than XCAP1 or actin alone, or a 1:1 complex, indicating that they form a stable multimeric complex. Native molecular mass of the XCAP1–actin complex was determined more accurately by two different methods: size-exclusion chromatography coupled with multiangle light scattering (SEC-MALS) and analytical ultracentrifugation. In SEC-MALS, the XCAP1–actin complex was resolved as a single peak with a molecular mass of 340 kDa (Fig. 1C). There were no detectable peaks that corresponded to dissociated XCAP1 or actin, indicating that the XCAP1–actin complex was stable during the SEC-MALS analysis. Likewise, in analytical ultracentrifugation, the XCAP1–actin complex was resolved as a single peak of 337 kDa (S = 10) (Fig. 1D), which agrees with the result of SEC-MALS. Considering the molecular masses of individual XCAP1 (52 kDa) and actin (42 kDa), the native molecular mass of the XCAP1–actin complex most closely matched with that of a 4:4 complex (calculated molecular mass of 376 kDa). The experimentally determined molecular mass was ∼10% smaller than the calculated molecular mass. This could be due to partial dissociation of the complex during the analyses, which is a known limitation in some cases of SEC-MALS experiments (38Folta-Stogniew E. Williams K.R. Determination of molecular masses of proteins in solution: Implementation of an HPLC size exclusion chromatography and laser light scattering service in a core laboratory.J. Biomol. Tech. 1999; 10: 51-63PubMed Google Scholar). Since CAPs are known to bind to actin monomers, the XCAP1–actin complex most likely contains G-actin. Therefore, these results indicate that the native XCAP1 and G-actin form a stable complex at a 4:4 stoichiometric ratio. Structure of the XCAP1–actin complex in its native state was examined by high-speed atomic force microscopy (HS-AFM) (Figs. 2 and 3). Typical images on mica surfaces showed that the complex consisted of three globular domains (Fig. 2A), which we designated as the middle globular domain (MGD, shown in red in Fig. 2A cartoons) and two arms (Arm 1 and Arm 2, shown in green or blue in Fig. 2A cartoons). The height of MGD was 3.6 ± 0.9 nm (n = 107) (Fig. 2, B and C) and remained relatively stable during time-lapse imaging (Fig. 2, A and G, Supplementary Movie S1). By contrast, the two arms were observed in two different states: a high state (Arm-HS, shown in blue in Fig. 2A cartoons) and a low state (Arm-LS, shown in green in Fig. 2A cartoons, also see Fig. 3). The height of Arm-HS was 7.5 ± 0.5 nm (n = 855) (Fig. 2, D and E), while that of Arm-LS was 3.3 ± 0.3 nm (n = 1078) (Figs. 2F and 3, C and D). In some cases, the arms transitioned either from Arm-LS to Arm-HS (Fig. 2, A and G, blue line at ∼0.9 s) or from Arm-HS to Arm-LS (Fig. 2, A and G, green line at 4.5 s), suggesting that association or dissociation of a component, presumably G-actin, occurred during observations. Over the periods of HS-AFM observations, Arm-HS gradually decreased, while Arm-LS predominated, suggesting that Arm-HS was converted to Arm-LS by dissociation of actin over time likely due to repeated tapping by the AFM probe and adsorption of the complex on the surface (see below). Some of the complexes had Arm-LS throughout the observations (Fig. 2A, indicated by dashed line, Fig. 3A, Supplementary Movie S2), and the height of Arm-LS was 3.3 ± 0.3 nm (n = 1078) (Fig. 3, B–D).Figure 3The XCAP1–actin complex with both arms in a low state is stable. A, time-lapse HS-AFM images of the XCAP1–actin complex containing both arms in Arm-LS on a mica surface (see Supplementary Movie S2). Scanning area was 80 × 64 nm2 with 64 × 48 pixels. Imaging rate was 66 ms/frame (∼15 fps). Bar, 20 nm. B, time course of the heights of three globular domains. C–F, cross-sectional analyses of Arm-LS (C, green) and MGD (E, red) at the straight colored lines drawn on the image in A. Height distributions of Arm-LS (D) and MGD (F) and single Gaussian fitting yielded average heights of Arm-LS and MGD as indicated in the figure. G–I, time course of the distances between the domains at their highest points (G). Distribution of the distance between two arms (H) and between MGD and one of the arms (I) and single Gaussian fitting yielded average distances as indicated in the figure.View Large Image Figure ViewerDownload Hi-res image Download (PPT) The two arms were very dynamic when they were in Arm-HS (Fig. 2A, Supplementary Movie S1), but restricted within 14.8 ± 4.9 nm (n = 214) of MGD (Fig. 2, H and I) as if the arms were connected to MGD by flexible linkers. The distance between the highest points of the two arms in Arm-HS fluctuated in a wide range with an average of 17.4 ± 7.6 nm (n = 427) (Fig. 2, H and J), further supporting the presence of flexible linkers between MGD and each arm. However, once arms were converted from Arm-HS to Arm-LS, they were stabilized in Arm-LS (Fig. 3, A–D, Supplementary Movie S2), while MGD remained unchanged (Fig. 3, B, E and F). The distance between two arms became wider [22.9 ± 5.7 nm (n = 539)], whereas that between each arm and MGD became narrower [13.0 ± 3.7 nm (n = 1078)]. These results suggest that Arm-LS was physically stabilized by adsorption to the mica surface. To test how surface adsorption affects the molecular features of the XCAP1–actin complex, we used a mica surface that was treated with APTES, which adds positive charges to the surface and causes nonspecific strong adsorption of proteins. On APTES-treated mica, the two arms were almost always detected in the low state (Arm-LS) with the height of 3.0 ± 0.4 nm (n = 2293) (Fig. 4, A–D, Supplementary Movie S3). The height and shape of MGD were indistinguishable between normal and APTES-treated mica surfaces (Fig. 4, A, B, E, and F). The distance between the two arms remained relatively constant at 25.4 ± 5.6 nm (n = 1331) (Fig. 4, G and H), which is much wider than that of two arms in Arm-HS on normal mica surfaces (Fig. 2I), suggesting that the two arms were strongly immobilized on the surface and spread apart. By contrast, the distance between MGD and an arm remained constant on the APTES-treated surface (Fig. 4, G and I) in a similar manner to MGD and Arm-LS on the normal surface (Fig. 3I). These observations suggest that strong adsorption of the XCAP1–actin complex onto a solid surface artificially converts Arm-HS to Arm-LS by causing dissociation of an arm-bound component, which we hypothesize to be G-actin. To test whether the height conversion of the arms is due to G-actin dissociation and association, we examined effects of additional free ADP-G-actin on the XCAP1–actin complex (Fig. 5). Freshly prepared samples of the XCAP1–actin complex contained mostly Arm-HS (Fig. 5A). However, after 15 min, most of them were converted to Arm-LS (Fig. 5B). After most of the arms were converted to Arm-LS (after ∼20 min), final 100 nM ADP-G-actin was added, and molecular features of the complex were observed over time (Fig. 5, C and D). In the presence of ADP-G-actin, frequent and reversible conversions of the arms between Arm-LS and Arm-HS were observed (Fig. 5C, Supplementary Movie S4). In the example shown in Figure 5C (dashed circle), initially both arms were Arm-LS (green arrowheads) at 0.6 s (Fig. 5C, top left panel), one arm was then converted to Arm-HS (blue arrowhead) at 0.8 s (Fig. 5C, top second panel), and another arm was converted to Arm-HS at 2.2 s (Fig. 5C, top fourth panel). Conversely, conversions from Arm-HS to Arm-LS were also observed (Fig. 5C, top right panel at 10.6 s and bottom second panel at 11.6 s). A plot of conversion events over time (Fig. 5D) indicates that the conversion between Arm-LS and Arm-HS was reversible and independently occurred at each arm without coordination between the two arms within a single complex. The rate of conversion events from Arm-LS to Arm-HS in the presence of ADP-G-actin was 0.14 ± 0.04 molecules−1 s−1 (Fig. 5E). Even in the absence of ADP-G-actin, conversion of Arm-LS to Arm-HS was occasionally observed at a rate of 0.03 ± 0.03 molecules−1 s−1 (Fig. 5E) most likely due to rebinding of dissociated G-actin to the complex, which is consistent with the spontaneous transition from Arm-LS to Arm-HS as demonstrated in Figure 1A. In the presence of ADP-G-actin alone or in the absence of the XCAP1–actin complex and ADP-G-actin, no objects that matched the size of Arm-LS or Arm-HS were observed (N.D.: none detected, Fig. 5E). These results strongly suggest that G-actin is a component of the two arm domains of the XCAP1–actin complex. Based on known biochemical and biophysical properties of CAP from other species, we propose a model for the structure of the XCAP1–actin complex, which is in the appearance of two "butterflies" (CARP/G-actin) and a "flower" (HFD) (Fig. 6). We hypothesize that MGD corresponds to a tetramer of the HFD of XCAP1 and that each arm domain in the high state (Arm-HS) corresponds to a heterotetramer containing a dimer of the CARP domain of XCAP1 and two G-actin molecules (Fig. 6). The HFD of CAP by itself forms a dimer (12Kotila T. Wioland H. Enkavi G. Kogan K. Vattulainen I. Jegou A. Romet-Lemonne G. Lappalainen P. Mechanism of synergistic actin filament pointed end depolymerization by cyclase-associated protein and cofilin.Nat. Comm. 2019; 10: 5320Crossref PubMed Scopus (35) Google Scholar, 26Yusof A.M. Hu N.J. Wlodawer A. Hofmann A. Structural evidence for variable oligomerization of the N-terminal domain of cyclase-associated protein (CAP).Proteins. 2005; 58: 255-262Crossref PubMed Scopus (22) Google Scholar, 27Yusof A.M. Jaenicke E. Pedersen J.S. Noegel A.A. Schleicher M. Hofmann A. Mechanism of oligomerisation of cyclase-associated protein from Dictyostel
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