Effect of pH on the mechanism of actin polymerization

The effect of pH on the Mg2+-induced polymerization of rabbit skeletal muscle G-actin at 20 degrees C was examined. Polymerization data were obtained at various initial concentrations of Mg2+, Ca2+, and G-actin between pH 6 and 7.5. The data were found to fit a kinetic mechanism for actin polymerization previously proposed at pH 8 in which Mg2+ binding at a moderate-affinity site on actin induces an isomerization of the protein enabling more favorable nucleation [Frieden, C. (1982) J. Biol. Chem. 257, 2882-2886]. The data also suggest the formation of actin dimers induced by Mg2+ binding is over 2 orders of magnitude more favorable at pH 6 than at pH 8. Little effect on trimer formation is found over this pH range. In addition, the conformation induced by nonspecific binding of metal to low-affinity sites becomes more favorable as the pH is lowered. The critical concentration for filament formation is also decreased at lower pH. The kinetic data do not support fragmentation occurring under any of the conditions examined. Furthermore, as Mg2+ exchange for Ca2+ at a high-affinity site (Kd less than 10(-9) M) fails to alter significantly the polymerization kinetics, Ca2+ release from this site appears unnecessary for either the nucleation or the elongation of actin filaments.     

The effects of cytochalasins on actin polymerization and actin ATPase provide insights into the mechanism of polymerization

Substoichiometric concentrations of cytochalasin D inhibited the rate of polymerization of actin in 0.5 mM MgCl2, increased its critical concentration and lowered its steady state viscosity. Stoichiometric concentrations of cytochalasin D in 0.5 mM MgCl2 and even substoichiometric concentrations of cytochalasin D in 30 mM KCl, however, accelerated the rate of actin polymerization, although still lowering the final steady state viscosity. Cytochalasin B, at all concentrations in 0.5 mM MgCl2 or in 30 mM KCl, accelerated the rate of polymerization and lowered the final steady state viscosity. In 0.5 mM MgCl2, cytochalasin D uncoupled the actin ATPase activity from actin polymerization, increasing the ATPase rate by at least 20 times while inhibiting polymerization. Cytochalasin B had a very much lower stimulating effect. Neither cytochalasin D nor B affected the actin ATPase activity in 30 mM KCl. The properties of cytochalasin E were intermediate between those of cytochalasin D and B. Cytochalasin D also stimulated the ATPase activity of monomeric actin in the absence of MgCl2 and KCl and, to a much greater extent, stimulated the ATPase activity of monomeric actin below its critical concentration in 0.5 mM MgCl2. Both above and below its critical concentration and in the presence and absence of cytochalasin D, the initial rate of actin ATPase activity, when little or no polymerization had occurred, was directly proportional to the actin concentration and, therefore, apparently was independent of actin-actin interactions. To rationalize all these data, a working model has been proposed in which the first step of actin polymerization is the conversion of monomeric actin-bound ATP, A . ATP, to monomeric actin-bound ADP and Pi, A* . ADP . Pi, which, like the preferred growing end of an actin filament, can bind cytochalasins.     

Effect of phosphatidyl-L-serine and vinculin on actin polymerization

The effects of phosphatidyl-L-serine (PS) and/or vinculin on actin polymerization are examined by spectrophotometry, viscometry and electrophoresis. Actin polymerization is inhibited by PS alone and stimulated by PS and vinculin. The results suggest that actin does not directly adhere to cell membrane and that vinculin is a protein which is involved in structures connecting actin microfilaments to cell membranes.     

Effect of capping protein on the kinetics of actin polymerization

Acanthamoeba capping protein increased the rate of actin polymerization from monomers with and without calcium. In the absence of calcium, capping protein also increased the critical concentration for polymerization. Various models were evaluated for their ability to predict the effect of capping protein on kinetic curves for actin polymerization under conditions where the critical concentration was not changed. Several models, which might explain the increased rate of polymerization from monomers, were tested. Two models which predicted the experimental data poorly were (1) capping protein was similar to an actin filament, bypassing nucleation, and (2) capping protein fragmented filaments. Three models in which capping protein accelerated, but did not bypass, nucleation predicted the data well. In the best one, capping protein resembled a nondissociable actin dimer. Several lines of evidence have supported the idea that capping protein blocks the barbed end of actin filaments, preventing the addition and loss of monomers [Cooper, J. A., Blum, J. D., & Pollard, T. D. (1984) J. Cell Biol. 99, 217-225; Isenberg, G. A., Aebi, U., & Pollard, T. D. (1980) Nature (London) 288, 455-459]. This mechanism was also supported here by the effect of capping protein on the kinetics of actin polymerization which was nucleated by preformed actin filaments. Low capping protein concentrations slowed nucleated polymerization, presumably because capping protein blocked elongation at barbed ends of filaments. High capping protein concentrations accelerated nucleated polymerization because of capping protein's ability to interact with monomers and accelerate nucleation.     

Effect of muscle tropomyosin on the kinetics of polymerization of muscle actin

At saturating concentrations, tropomyosin inhibited the rate of spontaneous polymerization of ATP-actin and also inhibited by 40% the rates of association and dissociation of actin monomers to and from filaments. However, tropomyosin had no effect on the critical concentrations of ATP-actin or ADP-actin. The tropomyosin-troponin complex, with or without Ca2+, had a similar effect as tropomyosin alone on the rate of polymerization of ATP-actin. Although tropomyosin binds to F-actin and not to G-actin, the absence of an effect on the actin critical concentration is probably explicable in terms of the highly cooperative nature of the binding of tropomyosin to F-actin and its very low affinity for a single F-actin subunit relative to the affinity of one actin subunit for another in F-actin.     

Effect of cibachron blue F3GA on the polymerization of actin

Summary The effect of the dye Cibachron Blue F3GA on the G-F transformation of rabbit muscle actin has been studied with viscosimetry. The presence of the dye which is known to bind to nucleotide binding sites, decreased both the initial rate of polymerization of actin as well as the final viscosity of actin. Both these effects can be ascribed to an increase in the critical concentration of actin. The inhibitory effect of Cibachron Blue F3GA was counteracted by ATP, suggesting a competition between Cibachron Blue F3GA and ATP for the binding site/sites on actin.     

Processive capping by formin suggests a force-driven mechanism of actin polymerization

Regulation of actin polymerization is essential for cell functioning. Here, we predict a novel phenomenon-the force-driven polymerization of actin filaments mediated by proteins of the formin family. Formins localize to the barbed ends of actin filaments, but, in contrast to the standard capping proteins, allow for actin polymerization in the barbed direction. First, we show that the mechanism of such "leaky capping" can be understood in terms of the elasticity of the formin molecules. Second, we demonstrate that if a pulling force acts on the filament end via the leaky cap, the elastic stresses can drive actin polymerization. We estimate that a moderate pulling force of approximately 3.4 pN is sufficient to reduce the critical actin concentration required for barbed end polymerization by an order of magnitude. Furthermore, the pulling force increases the polymerization rate. The suggested mechanism of force-driven polymerization could be a key element in a variety of cellular mechanosensing devices.     

Effect of polymerization on the subdomain 3/4 loop of yeast actin

The Holmes F-actin model predicts a polymerization-dependent conformation change of a subdomain 3/4 loop with a hydrophobic tip (residues 266-269), allowing interaction with a hydrophobic surface on the opposing strand of the filament producing filament stabilization. We introduced cysteines in place of Val(266), Leu(267), and Leu(269) in yeast actin to allow attachment of pyrene maleimide. Pyrene at each of these positions produced differing fluorescence spectra in G-actin. Polymerization decreased the fluorescence for the 266 and 267 probes and increased that for the 269 probe. The direction of the fluorescence change was mirrored with a smaller and less hydrophobic probe, acrylodan, when attached to 266 or 269. Following polymerization, increased acrylamide quenching was observed for pyrene at 266 or 267 but not 269. The 267 probe was the least accessible of the three in G- and F-actin. F-actin quenching was biphasic for the 265, 266, and 269 but not 267 probes, suggesting that in F-actin, the pyrene samples multiple environments. Finally, in F-actin the probe at 266 interacts with one at Cys(374) on a monomer in the opposing strand, producing a pyrene excimer band. These results indicate a polymerization-dependent movement of the subdomain 3/4 loop partially consistent with Holmes' model.     

Surface-induced polymerization of actin

Living cells contain a very large amount of membrane surface area, which potentially influences the direction, the kinetics, and the localization of biochemical reactions. This paper quantitatively evaluates the possibility that a lipid monolayer can adsorb actin from a nonpolymerizing solution, induce its polymerization, and form a 2D network of individual actin filaments, in conditions that forbid bulk polymerization. G- and F-actin solutions were studied beneath saturated Langmuir monolayers containing phosphatidylcholine (PC, neutral) and stearylamine (SA, a positively charged surfactant) at PC:SA = 3:1 molar ratio. Ellipsometry, tensiometry, shear elastic measurements, electron microscopy, and dark-field light microscopy were used to characterize the adsorption kinetics and the interfacial polymerization of actin. In all cases studied, actin follows a monoexponential reaction-limited adsorption with similar time constants (approximately 10(3) s). At a longer time scale the shear elasticity of the monomeric actin adsorbate increases only in the presence of lipids, to a 2D shear elastic modulus of mu approximately 30 mN/m, indicating the formation of a structure coupled to the monolayer. Electron microscopy shows the formation of a 2D network of actin filaments at the PC:SA surface, and several arguments strongly suggest that this network is indeed causing the observed elasticity. Adsorption of F-actin to PC:SA leads more quickly to a slightly more rigid interface with a modulus of mu approximately 50 mN/m.     

Intracellular transport based on actin polymerization

In addition to the intracellular transport of particles (cargo) along microtubules, there are in the cell two actin-based transport systems. In the actomyosin system the transport is driven by myosin, which moves the cargo along actin microfilaments. This transport requires the hydrolysis of ATP in the myosin molecule motor domain that induces conformational changes in the molecule resulting in the myosin movement along the actin filament. The other actin-based transport system of the cell does not involve myosin or other motor proteins. This system is based on a unidirectional actin polymerization, which depends on ATP hydrolysis in actin polymers and is initiated by proteins bound to the surface of transported particles. Obligatory components of the actin-based transport are proteins of the WASP/Scar family and a complex of Arp2/3 proteins. Moreover, the actin-based systems often contain dynamin and cortactin. It is known that a system of actin filaments formed on the surface of particles, the so-called “comet-like tail”, is responsible for intracellular movements of pathogenic bacteria, micropinocytotic vesicles, clathrin-coated vesicles, and phagosomes. This movement is reproduced in a cell-free system containing extract of Xenopus oocytes. The formation of a comet-like structure capable of transporting vesicles from the plasma membrane into the cell depth has been studied in detail by high performance electron microscopy combined with electron tomography. A similar mechanism provides the movement of vesicles containing membrane rafts enriched with sphingolipids and cholesterol, changes in position of the nuclear spindle at meiosis, and other processes. This review will consider current ideas about actin polymerization and its regulation by actin-binding proteins and show how these mechanisms are realized in the intracellular actin-based vesicular transport system.     

Polyamine-induced actin polymerization.

Muscle actin has been found to polymerize reversibly upon addition of low concentrations of polyamines. This polymerization, studied by centrifugation, has shown a linear relationship between the actin polymerization yield and the chain length of the polyamine. Among the biological polyamines tested, spermidine and spermine are the most efficient. The polymerization of actin can also be induced by the corresponding mono or diguanidine derivatives of these polyamines but monoamines or amino acids are inactive at the same concentration. The transformation of actin from a globular to a fibrous from upon addition of spermidine is also demonstrated by the changes in the near-ultraviolet circular dichoroic spectrum of this protein. Moreover, the polyamine-induced F -actin exhibits the same properties as the salt-induced F -actin: it strongly activates the Mg2+ -ATPase of myosin, its specific viscosity is enhanced to the same extent and electron micrographs show homogeneous thin filaments.     

Influence of counterions radius on the polymerization of actin

The G–F transformation of actin was studied in terms of an interaction between different counterions and actin. macroion by means of electrometric measurements. Due to the polyelectrolyte-structure of the actin molecule, the electrostatic interactions must play a predominant role. In this sense the influence of the counterion radius has been investigated.     

The experimental basis of some recent hypotheses on the mechanism of the polymerization of actin: a reappraisal

We have studied the effect of sonication on the fluorescence of N-(1-pyrenyl)iodoacetamide-labeled F-actin as well as of native actin-pyrenyl-actin mixed oligomers in which the subunits were covalently attached to each other by phenylenebismaleimide. In both cases the fluorescence of the solution was largely decreased by sonication. We have found that this effect is due (a) to a 20-30% decrease of the specific fluorescence of the polymers. These results question the validity of the novel mechanism for the polymerization of actin recently proposed (D. Pantaloni et al. (1984) J. Biol. Chem. 259, 6274-6283). In these studies, in fact, the implicit assumption was made that the quenching of the fluorescence of the solution under sonication was due exclusively to the conversion of F-actin into G-actin.     

Role of actin DNase-I-binding loop in myosin subfragment 1-induced polymerization of G-actin: implications for the mechanism of polymerization

Proteolytic cleavage of actin between Gly(42) and Val(43) within its DNase-I-binding loop (D-loop) abolishes the ability of Ca-G-actin to spontaneously polymerize in the presence of KCl. Here we show that such modified actin is assembled into filaments, albeit at a lower rate than unmodified actin, by myosin subfragment 1 (S1) carrying the A1 essential light chain but not by S1(A2). S1 titration of pyrene-G-actin showed a diminished affinity of cleaved actin for S1, but this could be compensated for by using S1 in excess. The most significant effect of the cleavage, revealed by measuring the fluorescence of pyrene-actin and light-scattering intensities as a function of actin concentration at saturating concentrations of S1, is strong inhibition of association of G-actin-S1 complexes into oligomers. Measurements of the fluorescence of dansyl cadaverine attached to Gln(41) indicate substantial inhibition of the initial association of G-actin-S1 into longitudinal dimers. The data provide experimental evidence for the critical role of D-loop conformation in both longitudinal and lateral, cross-strand actin-actin contact formation in the nucleation reaction. Electron microscopic analysis of the changes in filament-length distribution during polymerization of actin by S1(A1) and S1(A2) suggests that the mechanism of S1-induced polymerization is not substantially different from the nucleation-elongation scheme of spontaneous actin polymerization.     

Amplification of actin polymerization forces

The actin cytoskeleton drives many essential processes in vivo, using molecular motors and actin assembly as force generators. We discuss here the propagation of forces caused by actin polymerization, highlighting simple configurations where the force developed by the network can exceed the sum of the polymerization forces from all filaments.

Introduction

Mechanical amplification is something we experience every day, in the form of gears, pulleys, and levers. While climbing a hill on a bicycle, for instance, shifting gears increases the force on the wheels while limiting the pressure required on the pedals. However, energy has to be conserved, and because mechanical work is defined as force × displacement, an increase in force can only be obtained at the expense of displacement. Thus, although shifting gears allows one to develop the additional force needed to go uphill, speed is reduced as each pedal stroke produces a smaller turn of the wheels. Cells have similarly developed microscopic force amplification strategies during evolution. Here, we discuss some amplification schemes for one of the major force generators in the cell—actin polymerization.Actin plays a ubiquitous role in cell motility and morphogenesis, spanning many scales of space and time. In fission yeast, for example, a miniature actin machinery only ∼100 nm across can induce the invagination of an endocytic vesicle in just a few seconds (Picco et al., 2015). However, to sever the entire yeast cell, a cytokinetic ring forms with an initial perimeter of ∼10 µm and requires ∼30 min to drive division (Proctor et al., 2012). These assemblies differ dramatically in both size and duration. In other species, considerably larger actin assemblies exist that reach the scale of centimeters, such as in muscle cells. Clearly, actin and its associated factors need to be specifically organized to achieve these different functions (Fig. 1). From a functional point of view, a key problem is to understand how the global architecture of an actin network allows forces that are produced at the molecular scale to be productive for the cell. In this respect, we can distinguish two sorts of components. Active components generate forces from chemical sources of energy and include molecular motors, as well as actin itself, which can push by polymerizing (Kovar and Pollard, 2004) and possibly pull while depolymerizing. Passive components, such as actin cross-linkers, are essential but can only transmit forces generated by other elements.Open in a separate windowFigure 1.Different actin networks. Networks of actin filaments are essential for many biological processes at the cellular level, and the organization of the filaments in space must be adapted to the task. Here, polymerization force (orange) of actin filaments (red) occurs near the plasma membrane (blue). Linear filopodia bundles with fascin (black) can produce high speeds, but represent a weak configuration for force generation. Lamellipodia are thin cellular extensions in which filaments are nearly parallel to the substrate on which the cell is crawling. The 2D branched network, created by Arp2/3 actin-nucleating complexes (black), can produce higher forces at the expense of displacement. During endocytosis in yeast, actin forms a 3D network at the site of the invagination that appears roughly spherical, but the organization of actin filaments in space is not known. The coat structure (yellow) enables actin to pull the membrane inward and actin polymerizes near the base of the structure, where Arp2/3 nucleators are shown in black (Picco et al., 2015). Endocytosis requires strong force amplification to pull the invagination against the turgor pressure.The forces developed by an actin meshwork are determined by the organization of its components. Ultimately, these forces must be sufficient to drive biological processes, and thus their scale depends on the physical characteristics of the cell. For example, in the case of endocytosis in yeast, the turgor pressure pushing the surface of the invagination outward reaches ∼1,000 pN, which the actin machinery must overcome (Basu et al., 2014). During cytokinesis, the actomyosin ring also works against the turgor pressure, which produces high forces on the furrow (Proctor et al., 2012). For both cases, these forces have been calculated from measured cellular parameters, particularly the turgor pressure and the dimensions over which the membrane is deformed. Hence, for these processes at least, the two ends of the problem are known: the forces produced by the molecular components make up the input and the force required for the cellular process to occur represents the output. Yet the force balance within the system must be considered to understand how the actin machinery harvests the input to produce this output.In this comment, we focus on the transmission of forces produced by the polymerization of actin, setting aside turnover and the contribution of molecular motors. We discuss specifically how the arrangement of the filaments in the system regulates the amount of productive force. In many ways, the actin machinery behaves analogously to a cyclist: though its power is limited, it can “shift gears” to favor either more displacement (high gears) or more force (low gears).

The force generated by actin polymerization

Actin polymerization can produce force. Indeed if an actin monomer in solution binds the barbed end of a filament, there is a change of free energy (ΔG_p) and polymerization will occur if ΔG_p < 0 (Fig. 2 A). This reaction depends on the concentration (C) of monomeric actin and will take place only above a critical concentration (C* of ∼0.14 µM; Pollard, 1986). It is associated with ΔG_p = −k_BT ln(C/C*), where k_B is the Boltzmann constant and T is the absolute temperature. If actin is polymerizing against a load and producing work (W), the change in free energy is ΔG_p + W. In this case, polymerization will occur spontaneously if the change is negative, i.e., ΔG_p + W < 0. Consider an actin filament pushing against a force (f) applied parallel to the filament axis (Fig. 2 B). Because the addition of one actin monomer produces a displacement (δ = 2.75 nm; Holmes et al., 1990), the mechanical work is W = f × δ. Forces that are antagonistic to elongation can impede actin assembly (Peskin et al., 1993). The critical force under which the filament would cease to elongate is called the polymerization force (f_a). Using a physiological concentration (C of ∼40 µM; Wu and Pollard, 2005), the polymerization force is thermodynamically limited to k_BT ln(C/C*)/δ = ∼9 pN (Hill, 1981). Within such limits, the force developed by polymerization will depend on the conditions of assembly. Direct measurements of the polymerization force using single-molecule techniques are scarce. A first study used optical traps on bare filaments, giving a force of ∼1 pN (Footer et al., 2007). By monitoring the buckling of filaments capped with formins, a second study found the force to be ∼1.3 pN (Kovar and Pollard, 2004). In both cases, the concentration of actin was an order of magnitude lower than in vivo, and the measured forces were in fact close to the theoretical maximum under the experimental conditions. Here, we will thus consider that f_a is within 1 and 9 pN. We further assume that an actin filament is able to elongate as long as the parallel component of the antagonistic force at its barbed end remains lower than f_a, irrespective of the perpendicular components (Fig. 2 C). We discuss various examples of force amplification in which the network develops forces that exceed f_a per filament, without breaking the thermodynamic requirement for actin polymerization (ΔG_p + W < 0).Open in a separate windowFigure 2.Polymerization mechanics. (A) During polymerization, the addition of one actin monomer (orange) corresponds to an elongation (δ) at the barbed end of an actin filament (red) and is associated with a change of free energy (ΔG_p = −k_b T ln(C/C*)). (B) The work required to push a load over a distance (h) with a force (f) is f × h, and thus assembly remains favorable as long as ΔG_p + f × h < 0. In the case where polymerization occurs straight against a load (h = δ), the maximal force (f_a) is f_a = k_b T ln(C/C*)/δ (Hill, 1981). (C) If the filament encounters the load with an angle (θ), then h = δ sinθ and the maximal force is consequently increased: f_θ = f_a/sinθ. (D) In the branched network of a lamellipod, actin grows against the leading membrane at an angle (θ = ∼54°). In the absence of friction, the force between the polymerizing tip (orange) of the actin and the membrane (blue) is perpendicular to the membrane. It can then reach a maximum magnitude of f_a/sinθ. The sum of the forces produced by the two filaments is then ∼2.5 f_a. (E) Higher forces arise by polymerizing with shallow angles. The device illustrated here is composed of a growing actin filament with a “leg” on its side. By elongating, the filament will induce rotation around the pivot point, where the leg is contacting the membrane. High forces can be exerted on a load supported at the branch point, as a result of the amplification achieved by the lever arm and contact angle. (F) The highest forces are generated if a filament polymerizes parallel to the surface. In the illustrated configuration, elongation of the filament will cause a load (green dome) to separate from the membrane. The maximal force is calculated as in E, except that anchoring has to be assumed at the pivot point to balance forces horizontally. The device can sustain high forces applied on the top of the dome because the upward movement is small compared with the elongation of the filament.

Table 1.

Physical characteristics of actin