Morphology of Actin Assemblies in Response to Polycation and Salts

Selective Formation of a Linear-Shaped Bundle of Microtubules ... Anisotropic Nucleation Growth of Actin Bundle: A Model for Determining the Well-Defi...
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Biomacromolecules 2005, 6, 3005-3009

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Morphology of Actin Assemblies in Response to Polycation and Salts Hyuck Joon Kwon,† Akira Kakugo,† Kazuhiro Shikinaka,† Yoshihito Osada,† and Jian Ping Gong*,†,‡ Graduate School of Science, Hokkaido University, Sapporo 060-0810, Japan, and SORST, JST, Sapporo, 060-0810, Japan Received May 9, 2005; Revised Manuscript Received September 26, 2005

F-actins are semi-flexible polyelectrolytes and can be assembled into a large polymer-actin complex with polymorphism through electrostatic interaction with polycations. This study investigates the structural phase behavior and the growth of polymer-actin complexes in terms of its longitudinal and lateral sizes in various polycation and KCl concentrations for a constant actin concentration. Our results show that the longitudinal growth and lateral growth of polymer-actin complexes, initiated by a common nucleation process, are dominated by different factors in subsequent growth process. This induces the structural polymorphism of polymer-actin complexes. Major factors to influence the polymorphism of polymer-actin complexes in polyelectrolyte systems have been discussed. Our results indicate that the semiflexible polyelectrolyte nature of F-actins is important for controlling the morphology and growth of actin architectures in cells. Introduction Actin, which is an abundant protein found in all eukaryotic cells, not only provides the mechanical strength of cell but also participates in cell movement such as cytokinesis, cell crawling, cytoplasmic streaming, and muscle contraction.1 In physiological conditions, globular actin (G-actin) monomers are self-assembled into linear filaments (F-actin). Moreover, actin molecules are associated into various structures such as cross-linked networks and parallel bundles. The morphology of actin assemblies is regulated by a number of actin linker proteins, which are usually classified into bundling proteins and cross-linking proteins2. However, it was reported that various actin bundles such as Drosophila bristles, nurse cell struts, stereocilia, and acrosomal process are independent of the specific type of linker proteins.3 Some linker proteins such as R-actinin and fascin show both bundling and cross-linking activities.4-6 These facts indicate that the morphology of actin assemblies is not uniquely determined by specific linker proteins but also by other factors such as the concentration of linker proteins, the environmental conditions (ionic strength or pH), and the kinetics of actin-linker proteins interaction in a nonequilibrium state in cells. It was also reported that the length and thickness of actin bundles are rigorously regulated in stereocilia, in microvilli, and in Drosophila bristles.7 Although many studies on the cellular mechanism of actin organization have been performed, it is poorly understood what the dominant factor in determining the morphology and size of the actin architecture is. F-actin is a semiflexible polymer with a persistence length of ∼10 µm.8 It is typically several micrometers in length * Corresponding author. † Hokkaido University. ‡ SORST.

and ca. 10 nm in thickness, with a linear charge density of 4 e/nm in physiological conditions. For a single F-actin, it behaves like a rodlike molecule. Longitudinal strong binding sites exist for end-to-end annealing of F-actins.1,9,10 Due to strong electrostatic repulsion, the energy barrier for the lateral growth of F-actins is much higher than that for end-to-end annealing. This results in an anisotropic growth. It has been reported that the binding of some proteins, including calponin, dystrophin, and MARKS peptide, to F-actins is due to electrostatic interactions without specific binding sites.11-13 F-actins also form large polymer-actin complexes with various polycations.11,14 It was also reported that the change in thickness of actin bundles does not affect length of actin bundles in cellular actin bundles like Drosophlia bristles.3 In our previous study, we found that the electrostatic interaction between F-actin and various polycations induces various morphologies of polymer-actin complexes.15 These results indicate that the semiflexible polyelectrolyte nature of F-actin plays a decisive role in forming the polymorphism. In this paper, we studied the effect of electrostatic interactions by regulating the polycation concentration (CP) and KCl salt concentration (CS) on growth size and morphology of polymer-actin complexes due to the polyelectrolyte nature of F-actins. The structural behavior of polymer-actin complexes is investigated systematically in a wide range of CP, which influences the electrostatic attraction between actin filaments, and CS, which influences long ranged electrostatic repulsion between actin filaments, while keeping the actin concentration (CA) as a constant. We demonstrate that the morphology of polymer-actin complexes is strongly dependent on CP and CS. We also show that the lateral growth is not accompanied by the longitudinal growth in subsequent growth processes after nucleation. The

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major factors in determining the longitudinal and lateral growth and the morphology of polymer-actin complexes are discussed. This study would provide new insight for understanding structural polymorphisms of intracellular actin architectures. Experimental Procedures Actin Preparation. G-actin was purified from scallops by the method of Spudich et al.16 Fluorescently labeled F-actin was obtained by stoichiometrically mixing G-actins and rhodamine-phalloidin (Molecular Probes No. 4171) in F-buffer (5 mM HEPES (pH 7.2), 0.2 mM ATP, 0.2 mM CaCl2, 100 mM KCl, 2 mM MgCl2) for 24 h at 4 °C. Phalloidin binds to G-actin stoichiometrically and stabilizes F-actin against depolymerization at a decreased critical concentration of actin. Polycation Preparation. Poly-N-[3-(dimethylamino)propyl] acrylamide methyl chloride quaternary (PDMAPAA -Q) was prepared by radical polymerization of a 1.0 M aqueous solution of DMAPAA-Q monomer (Tokyo Kasei Co., Ltd) in the presence of 0.2 mol % potassium persulfate (Tokyo Kasei Co., Ltd). PDMAPAA-Q is diluted by HEPES (pH7.2) buffer and then used for experiments. Sample Preparation for Observation. The rhodaminephalloidin labeled actin (later denoted as F-actin) concentration was kept constant at 0.01 mg/mL (corresponding to 2.32 × 10-7 M). The KCl concentration was varied from 0.01 to 0.4 M; the PDMAPAA-Q concentration was increased from 10-7 to 0.1 M. Following the addition of the KCl and PDMAPAA-Q to F-actin solutions, the sample was mixed and incubated at room temperature for 60 min. Fluorescence Microscopy Observation. A cover glass was placed on a slide glass equipped with two spacers 1.11.4 mm high at both sides to form a flow-cell. Solution of F-actin, PDMAPAA-Q, and KCl mixture about 10 µL was introduced into the flow-cell by a micropipet. Then the flow cell was placed on the stage of a fluorescence microscope (Olympus BX 50) and observed under a ×60 objective lens. The fluorescence images were recorded by a CCD-camera (Olympus CD-300T-RC), and measurement of the longitudinal filament assembly was performed by using a computer analyzing program (MetaMorph, Nippon ROPER). The average length L of the polymer-actin complexes, measured as long axis, was the average over 100 samples. Transmission Electron Microscopy (TEM) Observation. TEM observation was performed using a JEOL (JEM1200EX) at 120 kV acceleration voltage. After incubation of mixtures of F-actin, PDMAPAA-Q, and KCl at room temperature for 60 min, about 10 µL of the sample was dropped on carbon-coated grids (NISSHIN EM Co., Tokyo). After 3 min, 2% uranyl acetate was added to the sample, and the grid was air-dried. The average thickness D of the actin bundle was obtained over 20 samples. Results and Discussion 1. Phase Diagram of Polymer-Actin Complexes. The polymer-actin complexes exhibit a rich polymorphism in a

Figure 1. Phase diagram for the morphology of polymer-actin complexes, in which the phase behavior is summarized as a function of PDMAPAA-Q concentration CP and KCl concentration CS for a constant actin concentration CA ) 0.01 mg/mL (2.32 × 10-7M). O, F-actins; 0, coexistence (polymer-actin complex and native F-actins); 2, cross-linked structure dominant phase; (, branched structure dominant phase; and 9, parallel bundle dominant phase. The dotted line shows the possible borderline between the native F-actin and F-actin with charge inversion.

Figure 2. Typical morphologies of polymer-actin complexes in phases I, II, III, IV, and V as observed by TEM images and fluorescence images. Scale bars present 200 nm for TEM images and 25 µm for fluorescence images.

wide range of CP and CS, as elucidated by the fluorescent images and TEM images, which show micro- and nanoscales, respectively. There are five characteristic phases in the CPCS phase diagram (Figure 1). Figure 2 shows the fluorescent images and TEM images for the polymer-actin complexes in the five phases. In phase I, F-actin does not grow. In phase II, we observe the coexistence of native F-actin and polymer-actin complexes. In the coexistence phase (II), the

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fraction of native F-actins increases with the increase of CS. The borderline between phases I and II shifts to a higher CP with the increase of CS. This can be explained by screening effect of salts on the electrostatic interaction between F-actins and polycations. As CP and CS increase, the F-actins form complexes with polycations and exhibit various structures. The polymeractin complexes evolve from the cross-linked structure dominant phase (III), to the branched-structure dominant phase (IV), and then to the parallel-bundle dominant phase (V) as shown by the TEM images in Figure 2. The largeangle cross-linked structure (phase III) should be attributed to the strong electrostatic repulsion between like-charged F-actins at a relatively low CP and CS. On the other hand, when a repulsion decreases (by increasing in CS) or an electrostatic binding energy overwhelms repulsion (by increasing in CP), the branched structure, and further the parallel bundle formation is favored. Only bundle and crosslinked phase were suggested because the actin filaments were assumed to be a completely rigid rod.17 However, an actual actin filament is a semiflexible polymer with a persistent length of ∼10 µm so that it is allowed to bend. The branched bundle phase is attributed to the semiflexible nature of F-actin. In the intermediate phase, part of the actin bundle prefers to align in parallel, but part of the actin bundle favors to be apart from one another because of electrostatic repulsion. Despite the initial formation of the actin bundle, subsequent growth in length increases the flexibility of polymer-actin complexes and therefore favors electrostatic repulsion between F-actins to form branch.18 The fluorescent images in Figure 2 show that the polymeractin complexes of the cross-linked structure dominant phase (III) are in compact globule state, whereas those of the parallel bundle dominant phase are in the extended state (V). The globule size of 15∼20 µm is attributed to the persistence length of F-actin. The morphological change of polymeractin complexes from the compact globule state to the extended bundle state is due to the increase of bending rigidity that increases with the thickness D of actin bundles, varying as D4.19 Here we discuss phase I where the F-actin does not grow in detail. In the low Cp and high CS region, there is no complex formation, and this native F-actin phase should develop to the whole CP range at CS > 0.4 M by screening the interaction between F-action and the polycation, since at CS ) 0.4 M, the Debye length of the solution κ - 1 ≈ 0.5 nm, which is comparable to the distance between the two next neighboring charges on F-actins and on polycations, both of which are ca. 0.25 nm. However, the mechanism for the no-actin growth at the high Cp and low CS region in phase I should be different. In this region, for example, at a PDMAPAA-Q concentration 0.7 M in CS ) 0.01 M, it is observed that the polymeractin complex is disassembled into native actin filaments, far beyond the critical CP for formation of the polymeractin complex. We consider that this disassembly of polymeractin complexes should be attributed to charge inversion of actin filaments. The excessive addition of polycations induces positive overcharging to the F-actin, due to the competition

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Figure 3. Average length (solid line) and thickness (dashed line) of polymer-actin complexes formed in various PDMAPAA-Q concentrations CP. CS ) 0.01 M. CA ) 2.32 × 10-7 M.

between the energy gain and the entropy penalty by complexation. An increase in CS suppresses the energy gain, and this is why the charge inversion region develops to a wider range of CP with the increase in CS in Figure 1. Therefore, phase I should be divided into a sub-phase I′. Although we could not determine the exact borderline between phase I and phase I′ from fluorescence images and TEM images, it should be somewhere as shown by the dotted line in Figure 1. These kinds of phenomena are also observed in DNApolycation system.20 As the concentration of multivalent cations increases in DNA-polyelectrolyte systems, condensation happens at a critical concentration (CC) and ends at a next critical concentration (Cd) where decondensation happens and DNA dissolves into solution due to charge inversion. This result indicates that the morphology of polymer-actin complexes can be regulated by control of electrostatic interaction, based on the semi-flexible polyelectrolyte nature of F-actin without specific linker proteins. 2. Effect of CP for a Constant CS. We quantitatively investigate the length and thickness change of polymeractin complexes in a wide range of CP at a constant CS (CS ) 0.01 M). As shown in Figure 3, there exists a critical PDMAPAA-Q concentration of CP ) 10-6 M to form polymer-actin complexes. When CP > 10-5 M, the length L of polymer-actin complexes decreases with the increase of CP. The average thickness D of polymer-actin complexes increases to a value of two times as that of native F-actin from CP ) 10-6 M, which is the same as the critical concentration for longitudinal growth by end-to-end annealing of F-actins, but it remains constant from 10-5 to 10-3 M regardless of considerable change in the length of polymeractin complexes. However, the thickness of polymer-actin complexes increases significantly from CP ) 10-2 M. This result shows that the initial lateral growth of polymer-actin complexes in CP ) 10-6 M is related to the longitudinal growth since the thickness increases in the same critical CP but shows a constant value in CP ) 10-5∼10-3 M, not affected by the subsequent growth in longitudinal direction. Therefore, the decrease of longitudinal size with increasing CP in CP ) 10-5∼10-3 M is not due to the competition between the longitudinal growth and the lateral growth but due to the competition between nucleation process and

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growth process. In the nucleation-growth mechanism, a system has to overcome a free-energy barrier to nucleate a large droplet, but once this energy barrier has been overcome, there is a very low energy barrier to subsequent growth. Right above the critical CP, only a small number of nuclei exist and a relatively large amount of F-actins are available, which favors the growth of each nucleus to a large polymer-actin complex.21 Increase in CP favors the nucleation and increases the nuclei concentration to form short polymer-actin complexes. Therefore, it is thought that the longitudinal size of polymer-actin complexes is determined by the concentration of F-actins, which is available for growth, relative to the concentration of nuclei. The different factors in determining the growth in the longitudinal direction and the lateral one should be attributed to the rodlike polyelectrolyte nature of the F-actin. Because F-actin is a rodlike polymeric filament with a high negative charge density, the lateral growth of the polymer-actin complexes by side-to-side bundling will be inhibited by an electrostatic repulsion energy barrier, which is far higher than that of longitudinal end-to-end annealing. Furthermore, the gyration radius of PDMAPAA-Q measured by static light scattering is 46 nm, which is much smaller than that of the longitudinal length of F-actin that is several micrometers. Thus, at a relatively low CP ( 10-2 M indicates that the increased lateral binding of PDMAPAA-Q at this concentration is enough to neutralize the negative charges of F-actins and induces side-to-side bundling. Therefore, the critical binding constant KPD (∼102 M-1) for the lateral growth is far lower than the critical binding constant KPL (∼106 M-1) for longitudinal growth. The 4 orders of magnitude difference in the binding constants induces the polymorphism. It was reported that in Drosophlia bristles, the change in thickness of the actin bundle does not affect its length.3 This result indicates that the longitudinal growth of cellular actin bundle is independent of the lateral growth, which is consistent with our result in polyelectrolyte system. 3. Effect of CS for a Constant CP. We investigated the effect of CS on the growth of polymer-actin complexes because an increase in CS screens the electrostatic interaction between F-actins and polycations,22 and this both suppresses the longitudinal growth and screens the lateral electrostatic repulsion between F-actins. Figure 4 shows the average length L and the average thickness D of polymer-actin complexes formed in a wide range of CS for a constant CP (CP ) 0, 10-6, 10-5, 10-2, and 10-1 M). As shown in Figure 4, when CP ) 0, there is no change in the F-actin length and thickness with the increase in CS. This indicates that F-actins do not aggregate to a large complex even at high CS when there is no polycation in the system. When CP ) 10-6 M, the critical concentration for longitudinal growth, the length of polymer-actin complexes decreases with the increase in CS, indicating that salt screens the electrostatic attraction between negatively charged F-actin and polycation. When CP is well above the critical concen-

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Figure 4. Average length (a) and thickness (b) of polymer-actin complexes in various CS at a constant CP. 3, 0 M; 4, 10-6 M; ), 10-5 M; 0, 10-2 M; and O, 10-1 M. CA ) 2.32 × 10-7 M.

tration (CP ) 10-5, 10-2, and 10-1 M), the length of polymer-actin complexes increases with increasing CS at a constant CP until Cs ) 0.3 M (Figure 4a). The increase in the length of polymer-actin complexes is attributed to the decrease of the nuclei concentration by the screening effect of CS on the electrostatic interaction between F-actins and polycations. As shown in Figure 4b, the thickness of the polymer-actin complexes increases with the increase of CS in CP ) 10-5 and 10-2 M until CS ) 0.3 M, whereas D is not influenced by the increase of CS in CP ) 10-1 M. The increase of CS screens both electrostatic attraction between F-actins and polycations and electrostatic repulsion between F-actins. The result that the thickness of the polymer-actin complexes increases with the increase of CS in CP ) 10-5 and 10-2 M indicates that the increase of CS more effectively screens the electrostatic repulsion between F-actins than the lateral attraction between F-actin and polycation. With the increase of CP, the lateral binding of polycation to F-actin increases, which favors the lateral growth. The thickness of the polymer-actin complexes is less increased by the increase of CS in high CP (10-1 M). Therefore, in high CP, the electrostatic attraction due to lateral binding of polycation dominates the lateral growth. It should be pointed out that the increase in longitudinal length, as investigated by the fluorescence images, is also affected by the conformational change of polymer-actin complexes from a compact globule structure to an extended structure by the increase of CS (Figures 1 and 2). Since the total ionic strength of the solution is from both the KCl and the polycation, we summarized the data on the relationship between ionic strength and D of actin bundle in

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physical origin to determine the morphology of actin architectures in cells. Biopolymers including F-actin have a lot of potential as functional materials in biotechnology and biomaterials design.25 Recently, actin-based metallic nanowires and actuators were constructed.26-29 When using biopolymers as functional materials, it is important to design biopolymers into well-defined and stable hierarchical macroscopic structure. Our study can provide useful information for not only understanding polyelectrolyte interaction, physical origins of actin organization in cell but also designing biomaterial based on actin filaments. Figure 5. Relationship between the average thickness D of actin bundles and the total ionic strength (CP + CS)1/2(M1/2). The solid line represents the change of thickness caused by the change of CP at a constant CS ) 0.01 M, and the dashed line represents the change of thickness caused by the change of CS at a constant CP ) 10-5 M. CA ) 2.32 × 10-7 M.

(CP + CS)1/2 - D diagram. Figure 5 shows that with increasing ionic strength the thickness of the actin bundle increases. However, the increase of ionic strength by increasing CP induces more effectively the lateral growth of the actin bundle than that by increasing CS. The increase of CP not only screens electrostatic repulsion between F-actins but also increases the binding energy by increasing the electrostatic attraction, whereas the increase of CS only screens electrostatic repulsion. This indicates that the balance between the electrostatic attraction and repulsion determines the thickness of polymer-actin complexes. Conclusions Our results show that the degree of nucleation and the balance of attraction and repulsion between semiflexible F-actins are important for determining the growth size and the morphology of polymer-actin complexes. The longitudinal and lateral growth of polymer-actin complexes initially depend on nucleation process, but in subsequent growth process, they are dominated by different factors: the concentration of F-actins which is available for growth and the balance of electrostatic attraction and repulsion between F-actins. Therefore, the structural polymorphism of polymeractin complexes is attributed to the different growth factors between length and thickness. It was reported that the phosphorylation and calcium binding of linker proteins can transform the morphology of actin architectures.23,24 The chemical modification of linker proteins such as phosphorylation and calcium binding can change the electrostatic interaction between F-actin and linker proteins. Therefore, the morphological change of the actin architectures due to phosphorylation or calcium binding would be attributed to the change of balance between binding energy and electrostatic repulsion. Therefore, the morphological polymorphism of intracellular actin architectures can also be induced by the concentration or activity of linker proteins and the ionic environment that can change the balance between the binding energy and the electrostatic repulsion. This would provide various routes for regulating the polymorphism of actin architectures in cells. The balance between the binding energy and the repulsion may be the

Acknowledgment. We are grateful to Mr. Noriaki Ito for his advice and assistance on the observation of the TEM images. We also appreciate the fruitful discussion with Dr. Y. Tanaka and Dr. Y. Hayase, Hokkaido University. We gratefully acknowledge SORST and JST for the financial support of this research. References and Notes (1) Bray, D. Cell moVements: from molecules to motility, 2nd ed.; Garland Publishing: New York, 2001. (2) Lodish, H.; et al. Molecular Cell Biology, 4th ed.; Freeman: San Francisco, CA, 1999. (3) DeRosier., D. J.; Tilney, L. G. J. Cell Biol. 2000, 148, 1. (4) Meyer, R. K.; Aebi, U. J. Cell Biol. 1990, 110, 2013. (5) Pelletier, O.; Pokidysheva, E.; Hirst, L. S.; Bouxsein, N.; Safinya, C. R. Phys. ReV. Lett. 2003, 91, 148102. (6) Tseng, Y.; Fedorov, E.; McCaffery, J. M.; Almo, S. C.; Wirts, D. J. Mol. Biol. 2001, 310, 351. (7) Guild, G. M.; Connelly, P. S.; Ruggiero, L.; Vrandich, K. A.; Tilney, L. G. J. Cell Biol. 2003, 162, 1069. (8) Yanagida, T.; Nakase, M.; Nishiyama, K.; Oosawa, F. Nature 1984, 307, 58 (9) Holmes, K. C.; Popp, D.; Gebhard, W.; Kabsch, W. Nature 1990, 347, 44. (10) Lorenz, M.; Popp, D.; Holmes, K. C. J. Mol. Biol. 1993, 234, 826. (11) Tang, J. X.; Janmey, P. A. J. Biol. Chem. 1996, 271, 8556-8563. (12) Tang, J. X.; Szymanski, P.; Janmey, P. A.; Tao, T. Eur. J. Biochem. 1997, 247, 432. (13) Amann, K. J.; Renley, B. A.; Ervasti, J. M. J. Biol. Chem. 1998, 273, 28419. (14) Tang, J. X.; Wong, S.; Tran, P.; Janmey, P. A. Ber. Bunsen-Ges. Phys. Chem. 1996, 100, 796. (15) Kakugo, A.; Shikinaka, K.; Matsumoto, K.; Gong, J. P.; Osada, Y. Bioconjugate Chem. 2003, 14, 1185. (16) Spudich, J. A.; Watt, S. J. Biol. Chem. 1971, 246, 4866. (17) Borukhov, I.; Brunisma, R. F.; Gelbart, W. M.; Liu, A. J. Proc. Natl. Acad. Sci. U.S.A. 2005, 102, 3673. (18) Howard, J. Mechanics of motor proteins and the cytoskeleton; Sinauer Associates, Inc.: Massachusetts, 2001. (19) Landau, L. D.; Lifshitz, E. M. Theory of Elasticity, 3rd ed.; Pergamon: Oxford, U.K., 1986. (20) Nguyen, T. T.; Rousina, I.; Shklovskii, B. I. J. Chem. Phys, 2000, 112 (5), 2562. (21) Stokes, D. L.; DeRosier, D. J. Biophys. J. 1991, 59, 456. (22) Tang, J. X.; Ito, T.; Tao, T. T.; Janmey, P. A. Biochemisry 1997, 36, 12600. (23) Herbeck, B.; Huttelmaier, S.; Schluter, K.; Jockusch, B. M.; Illengerger, S. J. Biol. Chem. 2000, 275, 30817. (24) Furukawa, R.; Maselli, A.; Thomsom, S. A. M.; Lim, R. W. L.; Stokes, J. V.; Fechheimer, M. J. Cell Sci. 2002, 116, 187. (25) Cui, D.; Gao, H. Biotechnol. Prog. 2003, 19, 683. (26) Patolsky, F.; Weizmann, Y.; Willner, I. Nat. Mater. 2004, 3, 692. (27) Kakugo, A.; Sugimoto, S.; Gong, J. P.; Osada, Y. AdV. Mater. 2002, 14, 1124. (28) Kakugo, A.; Shikinaka, K.; Takekawa, N.; Sugimoto, S.; Osada, Y.; Gong, J. P. Biomacromolecules 2005, 6, 845. (29) Kakugo, A., Sugimoto, S., Shikinaka, K, Gong, J. P., Osada, Y. J. Biomater. Sci. Polym. Ed. 2005, 16 (2), 203.

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