Actin is a multi-functional protein that is found in almost all eukaryotic cells. When polymerized, it forms robust filaments that participate in a variety of cellular processes. For example, actin filaments are involved in the contraction of muscles, and they are also a major component in the various structures that maintain and control the shape of cells as they move and divide. These structures include the cell cortex, a meshwork of actin filaments that is bound to the inner surface of the plasma membrane by anchor proteins. However, both the cell cortex and the plasma membrane must undergo dramatic changes when a cell divides, and the forces that drive these changes are generated by another protein, myosin II.

Myosin II contains three domains: a head domain, also known as the motor domain, that binds to actin; a neck domain; and a tail domain. Like actin, myosin II proteins also form filaments, but these myofilaments have a distinctive structure: the tail domains of two Myosin II proteins join together, with the motor domains being found at both ends of the filament. When activated, the motor domains grab actin filaments and pull against them in a ‘powerstroke’. However, the details of the interactions between the myofilament motor domains and the actin filaments in the cell cortex, which are bound to the plasma membrane, are not fully understood.

Studying these processes in living cells is extremely challenging, so Vogel et al. have built an in vitro model of the cell cortex, and then used single-molecule imaging to watch the interactions between the myofilaments and the actin filaments in this model. They show that the myofilaments move along the actin in the cortex, breaking up the filaments and compressing them in the process. They propose that tension builds up between the ends of the myofilaments, leading to compressive stress being exerted on the actin filaments. Computer simulations confirm that the forces generated are high enough to cause the actin filaments to buckle and break. The in vitro model developed by Vogel et al. should allow researchers to clarify the basic biophysical principles that underpin the structure and function of the cell cortex.

The actin cortex consists of a thin actin meshwork bound to the inner cytosolic face of the plasma membrane by various anchor proteins (Morone et al., 2006). It plays a pivotal role in providing mechanical stability to the cell membrane, and in controlling cell shape changes during cell locomotion and cell division (Wessells et al., 1971; Bray and White, 1988; Diz-Muñoz et al., 2010; Sedzinski et al., 2011). Many of these features rely on proper functioning of the actin motor myosin II (De Lozanne and Spudich, 1987; Cramer and Mitchison, 1995). Myosin II in the actin cortex functions as assemblies of motor proteins forming anti-parallely arranged bipolar filaments (myofilaments) with motor domains on both filament ends ((Verkhovsky and Borisy, 1993; Verkhovsky et al., 1995), Figure 1A). Besides being the force generator necessary for cell cortex remodeling and actomyosin ring constriction during cytokinesis, there is evidence that myofilaments also contribute to actin filament turnover during cyotkinesis (Burgess, 2005; Guha et al., 2005; Murthy and Wadsworth, 2005). In all these processes, the microscopic mechanism of the interaction between individual myofilaments and membrane-bound actin filaments is not understood. Due to the vast complexity of cellular systems, much effort has been spent to investigate the consequences of actin-myosin interactions from the in vitro perspective (Backouche et al., 2006; Smith et al., 2007; Schaller et al., 2010; Kohler et al., 2011; Soares e Silva et al., 2011; Gordon et al., 2012; Reymann et al., 2012) However, these studies focused on myosin-induced actin structure formation on a mesoscopic scale, rather than on the interaction between actin and individual myofilaments. In addition, membrane-bound minimal actin systems have only recently begun to be functionally reconstituted (Vogel and Schwille, 2012). To fill the gap in understanding individual myofilament–actin interactions, we directly visualized the action of myofilaments on membrane-bound actin filaments in a minimal in vitro system and complemented the experimental findings with a theoretical model.

Figure 1

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To mimic the cell cortex, we developed a ‘minimal actin cortex’ (MAC) consisting of actin filaments coupled to a supported lipid bilayer via biotin neutravidin bonds (Figure 1A). We used Alexa-488-phalloidin stabilized as well as non-stabilized (data not shown) actin filaments for the MAC composition. By varying the amount of biotinylated lipids present in the lipid bilayer, we have control over the density of the actin layer (Figure 1B). In order to understand the origin of contractility observed in cell cortices, we tested the response of the MAC upon addition of myosin motors. Rabbit muscle myosin II was purified and reassembled forming bipolar synthetic myofilaments with a typical length of 500–600 nm (Figure 1C). Time lapse imaging of MACs with various actin densities using total internal reflection fluorescence microscopy (TIRFM) showed a dynamic rearrangement of the actin filaments, and subsequently the formation of actomyosin foci in an ATP-dependent manner immediately after addition of myofilaments (Figure 1D, Movie 1). Actin pattern formation occurred at ATP concentrations between 0.1–1 µM in systems where ATP is enzymatically regenerated (Table 1) in 94% of the experiments (n = 45 experiments). At higher and lower ATP concentrations, actin pattern formation in the MAC is absent. Actin structure formation after ATP depletion has been also reported for experiments using actin and myofilaments in solution (Smith et al., 2007). At low ATP concentrations, myofilaments were predicted to function as active temporary crosslinkers, which may drive self-assembly of actin filaments into actin clusters.

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To understand the details of myofilament–actin interactions at low ATP concentrations during actin pattern formation, we added myofilaments to low density MACs. If crosslinking of actin by myofilaments were the only prerequisite for pattern formation, we would expect no (fast) pattern formation in low density MACs, due to the long distances between the actin filaments in relation to the length of myofilaments. Strikingly, myofilament addition to low density MACs displayed breakage events and compaction of actin filaments, resulting in their shortening over time in all experiments (n = 21, Figure 2A–C, Movie 2). After 20 min, the majority of the actin filaments have been shortened to, on average, half of their original length, and most of the fragments coalesced into single foci (Figure 2A,B, Movie 2). Note that actin filaments remained intact when imaged in the absence of myofilaments (data not shown). Upon myofilament addition, actin filaments frequently showed deformations prior to actin filament breakage (Figure 2C, yellow arrowheads, Movie 3), indicating that force is exerted by the myofilaments and stress along the actin filament may build up until the actin filament breaks. Similarly, recent evidence implied that exposure of actin/fascin bundles to myofilaments can induce their disassembling and severing by an unknown process (Backouche et al., 2006; Haviv et al., 2008; Thoresen et al., 2011).

Figure 2

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Furthermore, an increase in fluorescence intensity along the remaining actin filament is often observed after a fragmentation event (Figure 2C, white arrowheads, Figure 2D,E). The fluorescence intensity proximal to the breakage site is approximately twice as high compared to the rest of the actin filament, suggesting that the fragment is dragged along the remaining actin filament by the myofilaments leading to its compaction (Figure 2C–E, Movie 3). We propose that fragmentation and compaction contribute to the observed coalescence of actin fragments into single foci during the dynamic rearrangement of the actin filaments (Figures 1D and 2A, Movies 1 and 2).

To determine how myofilaments execute fragmentation and compaction and to test whether these processes demand (concerted) actions of a multitude of myofilaments, we reduced the myofilament concentration to the single molecule level. Alexa-647 labeled myofilaments were added to MACs with low actin density and imaged by two-color TIRFM. Upon binding of single myofilaments to individual actin filaments, we observed directed movement of myofilaments along actin filaments and actin fragmentation at low ATP concentrations (Figure 3A, Movie 4, Table 1, Figure 5). 68% of the individually observed myofilaments displayed directed movement and 50% exhibited fragmentation and compaction of an actin filament (total number of myofilaments = 152; 7 experiments). In cases where both fragmentation and compaction occurred, 95% of the observed myofilaments showed directed movement along the actin filaments, while 5% remained stationary at their original binding site (total number of fragmenting myofilaments = 75; seven experiments).

Figure 3

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In contrast, at high ATP concentrations, processive movement of myofilaments was barely visible, and fragmentation of actin filaments was absent (supplementary text in ‘Material and methods’, Figure 5, Movie 5).

Shortly after binding of myofilaments, actin filaments deformed and eventually broke, indicating that the force generated by a single myofilament is sufficient to break an actin filament (Figure 3A, yellow arrowheads, Movie 4). Subsequently, the fluorescence signal of actin filaments proximal to the breakage site increased (Figure 3A, white arrowheads, Figure 3C). The detected increase in the fluorescence intensity of actin filaments implies that actin fragments are further dragged along the remaining actin filaments by the myofilaments during their movement (Figure 3A,D, Movie 4). We analyzed our TIRFM data in greater detail by tracking single myofilaments during their directed movement along an actin filament (Figure 3B, (Rogers et al., 2007)) and determining the velocity from the trajectories (Figure 3C, red curve). Simultaneously, the fluorescence intensity in the green (actin) channel of the area occupied by the myofilament during its movement was measured, indicating breakage events as the dragged fragment leads to an increase of fluorescence proximal to the breakage site (Figure 3C, blue and black curve). The velocity fluctuates during the movement of the myofilament along the actin filament (Figure 3C, red curve), whereby acceleration events (Figure 3C, red arrowheads) are closely followed by an increase in the actin fluorescence, indicating that a breakage event occurred (Figure 3C, black arrows, blue and black curve). We hypothesize that phases of increasing tension between the ends of the myofilament lead to deformation of the actin filament and coincide with a decrease in velocity, while the phases of tension release immediately after actin filament breakage may result in acceleration of the myofilament (Figure 3C,D).

In order to explain the buckling and eventual breakage of the actin filament, we propose the following model: The myosin filament aligns parallel to the actin filament and interacts with it via the myosin heads ((Sellers and Kachar, 1990), Figure 3D). One end of the myofilament, which we will refer to as the ‘leading end’, is oriented towards the actin plus end (barbed end), as the filaments in a muscle, and can therefore walk along the actin filament while hydrolyzing ATP (Figure 3D). The other end of the myofilament, which we will refer to as the ‘trailing end’, separated from the leading end by the approximately 160 nm long bare zone (Al-Khayat et al., 2010) without myosin heads, is oriented in the opposite direction. While the trailing end still interacts with the actin filament, its opposite orientation results in a much slower directed motion towards the actin plus end (Spudich et al., 1985; Sellers and Kachar, 1990). Every myosin head independently follows the biochemical cycle consisting of ATP hydrolysis, binding to actin, powerstroke accompanied by phosphate dissociation, ADP dissociation, ATP binding, and finally detachment from actin ((Howard, 2001), Figure 8). We assume that the heads on the trailing end still interact with the actin filament and go through the same cycle, but either do not perform steps that would lead to a processive motion (probability of making a step p_st=0), or make steps with a small probability (p_st=0.1). When a myosin head makes a step (step size d = 5 nm (Howard, 2001)) it tries to move the whole myosin filament towards the actin plus end, but other attached myosin heads are holding it back, thus generating friction (Figure 3D). The myofilament trailing end functions mainly as a source of friction (an effective ‘brake’), but is also moved towards the plus end by the pulling force exerted by the leading end (Figure 3D). Since mainly the leading end is actively moving and the trailing end is either passively pulled (p_st=0) or contributes only weakly to its own motion (p_st=0.1), tension builds up within the myofilament, and is transferred onto the actin filament as a compressive force. If sufficiently high, this compressive force can cause buckling, and finally breakage, of the actin filament (Figure 3D). After the breakage, the leading end can move unhindered further towards the plus end, while dragging along the broken-off part of the actin filament attached to the trailing end (Figure 3D). The trailing end can also attach again to the actin and further breakage events on the same filament can follow.

In order to estimate the force needed to break an actin filament, we model the filaments as flexible rods with bending rigidity EI = 60 nN μm², determined from the persistence length of actin: l_p = EI/(kT) = 15 μm (Yanagida et al., 1984). The force needed to buckle and break a filament is F = π² EI/l². With the length l of the myofilament bare zone of 160 nm, this gives a force of 23 pN. Can the tension within the myofilament reach up to this force? We performed simulations of this model, describing the attached myosin head as a spring with a spring constant equal to the myosin head stiffness κ = 1 pN nm⁻¹ (Kaya and Higuchi, 2010), and equilibrate all forces after every step and every detachment of a myosin head. From the AFM image of the myofilament (Figure 1C) and the known myofilament structure (Woodhead et al., 2005), we estimated that there are n_m = 30 interacting myosin heads per filament. The result is a net movement of the whole myofilament towards the actin plus end, with fluctuating tension force, velocity and number of attached myosin heads, the mean values of which depend on the ATP concentration (Figures 4A,B and 6). We note that the tension force increases with decreasing ATP concentration, reaching forces necessary for breakage only at low (lesser than ∼3 µM) ATP concentrations, in agreement with the experiments (Figure 4A, see also Table 1). When the actin filament was allowed to bend in simulations at the point where the buckling force of 23 pN was reached (Figure 4B), thus releasing the excess tension, the filament curvature steadily increased at low ATP concentrations, reaching the critical curvature of breaking of 5.6 µm⁻¹ ((Arai et al., 1999), Figure 7A). At intermediate ATP concentrations, the actin curvature fluctuated, at times exceeding the curvature threshold (Figures 4C and 7B). At higher ATP concentrations, the threshold force was reached for too short periods for the filament to be bent to the breakage point (Figures 4D and 7C). The results do not depend significantly on whether the myosin heads on the trailing end perform steps (p_st=0.1) or not (p_st=0). The simulations thus support the idea that in the ATP concentration range used in the experiments, the differences in the interactions of the trailing and leading ends of the myofilament with actin can generate compressive forces on the actin filament, and that these forces are sufficiently high to bend and break the actin filament.

Figure 4

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In our in vitro study we provide a potential mechanism how actin turnover in cells may be mediated by myofilament driven actin fragmentation. To bend and break the actin filament, the myofilament has to be attached to the actin for sufficient time, and sufficient force has to be generated, requiring a certain mean number of heads being attached to actin at any given time. Given the small number of interacting myosin heads this translates to a requirement that a large fraction of heads is in the bound state. In our assay this was achieved with low ATP concentrations. It is tempting to speculate that actin breakage by myofilaments in vivo is governed by the ATP level in the cell. Recent evidence exists that ATP levels indeed vary significantly inside living cells (Imamura et al., 2009). However, physiological ATP concentrations, derived from methods that provide averaged ATP levels with no high spatial and temporal resolution from cell extracts, are usually found in the millimolar range (Beis and Newsholme, 1975). Here it is important to mention that skeletal muscle myosin II (used in this study) has a lower duty ratio and is therefore less processive than non-muscle myosin in cells (Harris and Warshaw, 1993; Wang et al., 2003). By lowering the ATP concentration in our assay, we increased the duty ratio of our myofilaments and thereby made them more processive (supplementary text in ‘Material and methods’, Figure 5, Movie 5), similar to non-muscle myosin, which is in line with previous studies (Humphrey et al., 2002; Smith et al., 2007; Soares e Silva et al., 2011). Moreover, processivity of myofilaments in animal cells may be also controlled by phosphorylation of myosin light chains through other proteins that modify the kinetic rates of the biochemical cycle and thereby increase the duty ratio of the myosins shifting the observed behavior to a region of high physiological ATP concentrations (Tan et al., 1992; DeBiasio et al., 1996; Matsumura et al., 2001).

Our simulations show that the model of interaction between the actin and myosin filaments described in the text is plausible given the quantitative parameter values known from literature (rate constants). Most importantly, it shows that sufficient force needed to bend and break the actin filament can be generated by a single myofilament, assuming only the difference in the interaction between the leading and trailing myofilament ends (making a step vs not making a step). This result implies that neither actin nor myosin attachment to a support or any other rigid structure is necessary for the observed process. The forces act within a single myofilament. Binding of the actin filaments to the membrane provides confinement of the filaments to a plane, without rigidly tethering the filaments to a support or a scaffold. The motion of actin is only restricted by the effectively higher viscosity of the membrane compared to the buffer solution. The simulation further yields quantitative details of the model, for example, the mean fractions of myosin heads in the six possible states (Figure 9), and describes the fluctuations of the actin filament curvature (Figure 4C,D). Other parameters that can be derived from the simulations and could be compared with new experiments include the processivity of myosin, as a function of the ATP concentration, and the dependence of the overall behavior on the length of myofilaments.

Figure 8

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Figure 9

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In conclusion, our findings show the distinct functions that myosin motors can execute. In our minimal system, myofilaments fragment and compact membrane-bound actin. We directly show that single myofilaments can interact with actin in such a way that sufficient force can be generated to break the filament, without either the myofilament or the actin being firmly attached to a solid support or scaffold. We suggest that fragmentation and compaction by myofilaments contributes to the observed large-scale pattern formation of actomyosin networks also in other in vitro systems (Backouche et al., 2006; Smith et al., 2007; Soares e Silva et al., 2011; Gordon et al., 2012). In vivo breakage of actin bundles has been shown to occur in neuronal growth cones in a myosin II dependent manner, and is thought to be important for recycling actin (Medeiros et al., 2006). We propose that the observed fragmentation and compaction of membrane-bound actin filaments by myosins in our in vitro system may serve as a possible general mechanism for actin turnover and actin cell cortex remodeling.

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Author details

1. Sven K Vogel

   Department of Cellular and Molecular Biophysics, Max Planck Institute of Biochemistry, Martinsried, Germany

   Contribution

   SKV: Conception and design, Acquisition of data, Analysis and interpretation of data, Drafting or revising the article

   For correspondence

   svogel@biochem.mpg.de

   Competing interests

   The authors have declared that no competing interests exist

2. Zdenek Petrasek

   Department of Cellular and Molecular Biophysics, Max Planck Institute of Biochemistry, Martinsried, Germany

   Contribution

   ZP: developed the theory and performed the simulations, Analysis and interpretation of data, Drafting or revising the article

   Competing interests

   The authors have declared that no competing interests exist

3. Fabian Heinemann

   Department of Cellular and Molecular Biophysics, Max Planck Institute of Biochemistry, Martinsried, Germany

   Contribution

   FH: acquired and analyzed the AFM data and helped with the analysis of the experimental data, Acquisition of data, Analysis and interpretation of data, Drafting or revising the article

   Competing interests

   The authors have declared that no competing interests exist

4. Petra Schwille

   Department of Cellular and Molecular Biophysics, Max Planck Institute of Biochemistry, Martinsried, Germany

   Contribution

   PS: Drafting or revising the article.

   For correspondence

   schwille@biochem.mpg.de

   Competing interests

   The authors have declared that no competing interests exist

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