
==== Front
Sci Adv
Sci Adv
sciadv
advances
Science Advances
2375-2548
American Association for the Advancement of Science

ado5788
10.1126/sciadv.ado5788
Research Article
Biomedicine and Life Sciences
SciAdv r-articles
Biophysics
Cell Biology
Biophysics
Myosin-I synergizes with Arp2/3 complex to enhance the pushing forces of branched actin networks
Myosin-I enhances the pushing forces of branched actin polymerization
https://orcid.org/0009-0004-4410-4116
Xu Mengqi Conceptualization Data curation Formal analysis Investigation Methodology Validation Visualization Writing - original draft Writing - review & editing 1 2 †
https://orcid.org/0000-0003-0430-9163
Rutkowski David M. Conceptualization Data curation Formal analysis Investigation Methodology Software Validation Visualization Writing - original draft Writing - review & editing 3 †
https://orcid.org/0000-0002-0766-6302
Rebowski Grzegorz Resources 1
https://orcid.org/0000-0003-1677-023X
Boczkowska Malgorzata Resources 1
https://orcid.org/0000-0002-3587-0714
Pollard Luther W. Conceptualization Methodology Project administration Supervision Validation Writing - review & editing 1 *
https://orcid.org/0000-0003-3186-5229
Dominguez Roberto Funding acquisition Project administration Resources Supervision Writing - review & editing 1 *
https://orcid.org/0000-0003-1802-3262
Vavylonis Dimitrios Conceptualization Data curation Funding acquisition Methodology Project administration Resources Supervision Writing - original draft Writing - review & editing 3 *
https://orcid.org/0000-0003-0544-9360
Ostap E. Michael Conceptualization Data curation Formal analysis Funding acquisition Methodology Project administration Supervision Validation Writing - original draft Writing - review & editing 1 2 *
1 Department of Physiology, Pennsylvania Muscle Institute, Perelman School of Medicine, University of Pennsylvania, Philadelphia, PA 19104, USA.
2 Center for Engineering Mechanobiology, University of Pennsylvania, Philadelphia, PA 19104, USA.
3 Department of Physics, Lehigh University, Bethlehem, PA 18015, USA.
* Corresponding author. Email: ostap@pennmedicine.upenn.edu (E.M.O.); vavylonis@lehigh.edu (D.V.); droberto@pennmedicine.upenn.edu (R.D.); luther.pollard@pennmedicine.upenn.edu (L.W.P.)
† These authors contributed equally to this work.

13 9 2024
13 9 2024
10 37 eado578809 2 2024
09 8 2024
Copyright © 2024 The Authors, some rights reserved; exclusive licensee American Association for the Advancement of Science. No claim to original U.S. Government Works. Distributed under a Creative Commons Attribution NonCommercial License 4.0 (CC BY-NC).
2024
The Authors
https://creativecommons.org/licenses/by-nc/4.0/ This is an open-access article distributed under the terms of the Creative Commons Attribution-NonCommercial license, which permits use, distribution, and reproduction in any medium, so long as the resultant use is not for commercial advantage and provided the original work is properly cited.

Class I myosins (myosin-Is) colocalize with Arp2/3 complex–nucleated actin networks at sites of membrane protrusion and invagination, but the mechanisms by which myosin-I motor activity coordinates with branched actin assembly to generate force are unknown. We mimicked the interplay of these proteins using the “comet tail” bead motility assay, where branched actin networks are nucleated by the Arp2/3 complex on the surface of beads coated with myosin-I and nucleation-promoting factor. We observed that myosin-I increased bead movement efficiency by thinning actin networks without affecting growth rates. Myosin-I triggered symmetry breaking and comet tail formation in dense networks resistant to spontaneous fracturing. Even with arrested actin assembly, myosin-I alone could break the network. Computational modeling recapitulated these observations, suggesting myosin-I acts as a repulsive force shaping the network’s architecture and boosting its force-generating capacity. We propose that myosin-I leverages its power stroke to amplify the forces generated by Arp2/3 complex–nucleated actin networks.

The myosin-I power stroke enhances the pushing forces generated by the actin cytoskeleton.

http://dx.doi.org/10.13039/100000001 National Science Foundation 2138259 http://dx.doi.org/10.13039/100000001 National Science Foundation 2138286 http://dx.doi.org/10.13039/100000001 National Science Foundation 2138307 http://dx.doi.org/10.13039/100000001 National Science Foundation 2137603 http://dx.doi.org/10.13039/100000001 National Science Foundation 2138296 http://dx.doi.org/10.13039/100000001 National Science Foundation CMMI: 15-48571 http://dx.doi.org/10.13039/100000001 National Science Foundation CMMI: 15-48571 http://dx.doi.org/10.13039/100000002 National Institutes of Health R37 GM057247 http://dx.doi.org/10.13039/100000002 National Institutes of Health R35 GM136372 http://dx.doi.org/10.13039/100000002 National Institutes of Health R01 GM073791
==== Body
pmcINTRODUCTION

Actin assembly stimulated by the Arp2/3 complex provides pushing forces for diverse cellular processes (1–8), including lamellipodial protrusion, endocytosis, phagocytosis, and cell adhesion. After being activated by membrane-associated nucleation-promoting factors (NPFs), Arp2/3 complex nucleates new actin branches from the sides of pre-existing mother filaments, generating branched, dendritic actin networks that exert pushing forces against or deform the membrane. The network geometry, assembly kinetics, and mechanical properties are dynamically adapted by a set of actin-binding proteins [e.g., capping proteins (CPs), NPFs, profilin, cofilin, and cross-linkers] that respond to mechanical loading (9–20).

Class I myosins (myosin-Is) (21, 22) frequently colocalize with Arp2/3 complex–nucleated branched actin networks near the cell membranes where they both participate in powering membrane dynamics (23–27). As an actin-activated adenosine triphosphatase (ATPase), the myosin-I motor dynamically detaches and attaches from actin filaments in an adenosine 5′-triphosphate (ATP)–dependent manner while generating force through a lever arm–mediated power stroke (28). Myosin-Is are single-headed, membrane-anchored motors that bind directly to phosphoinositide-rich membranes similar to the NPFs (29–37), thus myosin-Is colocalize with the barbed ends of actin filaments (22). Although multiple cellular studies have shown that the myosin-I motor activity cooperates with dynamic actin assembly mediated by the Arp2/3 complex at the plasma membrane (24, 25, 27, 38, 39), the functional outcome of this interaction remains unknown.

To investigate the functional interaction between Arp2/3 complex and myosin-I in actin assembly, we developed a biomimetic system using a comet tail bead motility assay (9, 12, 40). This system recapitulates the colocalization and enrichment of NPFs and myosin-I observed on cellular membranes by using micrometer-sized beads coated with both NPF and myosin-I. We investigated the impact of myosin-I on Arp2/3 complex–mediated branched actin assembly across varying network densities achieved through different CP concentrations (12, 14, 41).

Our findings revealed that myosin-I alters actin assembly kinetics by reducing Arp2/3 complex–mediated branching at NPF-coated surfaces, resulting in a sparser actin network that exhibits enhanced elongation efficiency. This effect is attributed to the pushing force exerted by the myosin-I power stroke directly on surrounding actin networks, propelling actin filaments away. Furthermore, the myosin-I power stroke generates sufficient force to disrupt the network and trigger actin shell breakdown. Notably, a computational model at the molecular level provides mechanistic insights, suggesting that myosin-I promotes force generation during Arp2/3 complex–mediated actin polymerization by exerting a repulsive force on the branched network via its power stroke.

RESULTS

Comet tail bead motility assay reveals how myosin-I affects branched actin assembly

The comet tail bead motility assay was used to investigate the effect of myosin-I activity on Arp2/3 complex–mediated branched actin assembly (9, 40). Full-length, biotinylated Drosophila Myo1d was attached to the bead surface via neutravidin (myosin-bead; see Materials and Methods; Fig. 1A) alongside a glutathione S-transferase (GST)–tagged WCA domain from human Neuronal Wiskott-Aldrich Syndrome protein (N-WASP). Drosophila Myo1d was chosen as it functions optimally at 20° to 22°C, which is the temperature range used in most in vitro reconstituted systems (42, 43). Bead-bound Myo1d activity was confirmed by processive bead movement along actin filaments (Fig. 1B and movie S1). Control-beads were made identically to the myosin-beads, except that a biotinylated far-red fluorescent dye (CF640; see Materials and Methods) was conjugated to the neutravidin, in place of myosin (Fig. 1A). Actin assembly around myosin- and control-beads was imaged simultaneously by epifluorescence microscopy (Fig. 1C).

Fig. 1. Alteration of comet tail morphology by myosin-I.

(A) Schematic of (left) control- and (right) myosin-beads coated with NPF and neutravidin, where neutravidin is further conjugated with biotinylated CF640 fluorescent dye (control-bead) or biotinylated Drosophila Myo1d (myosin-bead). (B) Time-lapse sequence of a (red) myosin-bead walking on a (green) single actin filament track. Speed ~8 nm/s. See movie S1 for a full time series. Scale bar, 2 μm. (C) Time series of the growth of actin comet tail from symmetry breaking to generation of a polarized comet tail from (top) control-bead and (bottom) myosin-bead. Scale bar, 5 μm. (D) Phase map showing representative examples of 2-μm-diameter beads coated with a range of myosin densities in the presence of 30 to 200 nM CP, 4 μM (5% rhodamine-labeled) actin, and 200 nM Arp2/3 complex. For each myosin density condition, the left column (red) shows control-beads that were acquired in the same imaging field as the myosin-beads in the right column (blue). The myosin and NPF densities were determined by sodium dodecyl sulfate–polyacrylamide gel electrophoresis (SDS-PAGE) gel, where the NPF density ~6000 μm−2 is the same for groups 0.28:1, 0.35:1, and 0.43:1, and ~4000 μm−2 for group 0.80:1. Images were acquired 20 to 35 min after mixing. Brightness and contrast were linearly set to be the same for each control- and myosin-bead pair but different among panels at different conditions for better visualization of the bead position [see fig. S1 for panels using a common lookup table (LUT)]. Scale bar, 5 μm.

Upon mixing of assay components, surface-bound NPFs stimulate Arp2/3 complex–mediated branched actin assembly. Polymerization at the bead surface displaces the branched actin arrays outward, building tension in the actin network, which ultimately breaks the symmetry, forming a polarized comet tail (Fig. 1C) (9, 14, 44–47). By varying CP concentration, we created networks of varying actin density that ranged from tightly packed and symmetry-breaking resistant (dense) to loosely woven and fracture-prone (sparse), allowing us to investigate Myo1d’s influence on symmetry breaking, comet tail formation, and morphology at different network densities.

Myosin-beads display distinct comet tail morphologies compared to the control-beads under varying CP concentrations (Fig. 1D and fig. S1), and these differences were more pronounced at higher Myo1d densities. Here, we identified three distinct regimes: high, intermediate, and low network densities, controlled by the CP concentration (200 nM, 40 to 100 nM, and 30 nM, respectively) (Fig. 1D and fig. S1) and described each regime in greater detail using the 0.43 myosin/1 NPF ratio in the following sections.

Myosin-I prevents comet tail formation from sparse actin networks

Under high CP concentrations (200 nM), actin elongation was suppressed by capping, resulting in actin arrays consisting of short branches. As a result, actin comet tails grown from control-beads were irregularly organized with sparse networks (Figs. 1D and 2A), consistent with previous reports (12, 14, 48). Myosin-beads (0.43:1 myosin/NPF) failed to generate a comet tail (Fig. 2A and movie S2) but instead formed loose actin clouds which diffused away from the bead when the flow was present in the chamber. The presence of myosin compromised the network cohesion, making it more prone to disruption and inhibiting comet tail formation.

Fig. 2. Disruption and fracturing of actin shells by myosin-I.

(A) Time series of (top) control-beads and (bottom) 0.43:1 myosin-beads acquired in the same imaging field in the presence of 200 nM CP showing the inability of myosin-beads to form a comet tail. (B) Time series of (top) control-beads and (bottom) 0.43:1 myosin-beads acquired in the same field in the presence of 25 nM CP showing fracturing of the actin shell and comet tail growth from a myosin-bead but not the control-bead. Conditions: 4 μM actin (5% rhodamine-labeled), 200 nM Arp2/3 complex, and 200 or 25 nM CP. Scale bars, 5 μm.

Myosin-I facilitates the fracturing of dense actin networks

At a low CP concentration (25 nM), long actin filaments emanating from bead surfaces become entangled, forming dense actin networks that are highly resistant to fracture by actin polymerization forces (12, 14, 46, 48). Control-beads remained encapsulated within the shell, without symmetry breaking, for >15 min after the initiation of the polymerization (Figs. 1D and 2B). Notably, myosin-beads grown under the same condition broke symmetry within 10 min of mixing (Fig. 2B and movie S3). This result suggests that myosin-I may enhance the force generation during actin assembly and/or may alter actin architecture that promotes network fracture and symmetry breaking.

Myosin-I induces efficient comet tail growth

At intermediate CP concentrations (40 to100 nM), myosin-beads generated comet tails with sparser actin networks compared to control-beads (Figs. 1D and 3, A and B, fig. S1, and movie S4). Following the time course of tail elongation, we found that the sparser network architecture of myosin-beads arises from a 0.76-fold lower actin assembly rate (P < 0.0001, n = 11; Fig. 3, E and F) as quantified by measuring the rhodamine-actin fluorescence in the comet tail over time. Despite the lower actin assembly rate, myosin comet tails in the presence of 50 nM CP elongated (0.69 ± 0.10 μm/min) at the same speed as the control-beads (0.66 ± 0.20 μm/min; Fig. 3, C and D). If we define the growth efficiency of actin comet tail as the comet tail length divided by the total amount of actin incorporated, then the myosin-beads exhibited a 1.4-fold higher growth efficiency than the control-beads (Fig. 3G).

Fig. 3. Myosin-I decreases the actin density of comet tails.

(A) Time series of (top) control and (bottom) 0.43:1 myosin-beads growing comet tails in the presence of 50 nM CP. The actin network growing from the myosin-bead is less dense but has a similar tail length as the control. Conditions: 4 μM actin (5% rhodamine-labeled), 200 nM Arp2/3 complex, and 50 nM CP. Scale bar, 5 μm. (B) Mean fluorescence intensities (Fluor. Int./area) of actin comet tails grown from control- and myosin-beads captured 20 min after mixing. The solid lines connect experimental pairs (N = 5 independent experiments, n = 11 pairs). (C) Actin comet tail length as a function of time and (D) tail growth rates derived from the slopes of the time courses in (C). (E) Comet tail fluorescence as a function of time. Control- and myosin-bead pairs are normalized to the average fluorescence level of the control-beads from 1100 to 1300 s. (F) Rate of fluorescent actin incorporation into comet tails derived from the slopes of the time courses in (E). (C and E) Large points show the averaged value at binned time intervals (every 100 s). Traces are from individual beads with each myosin-bead acquired with a control-bead in the same field of view (N = 5, n = 11). Error bars are SD. (G) Growth efficiency for control- and myosin-beads. Efficiency is defined as comet tail length per unit actin fluorescence intensity. Box plots (B, D, F, and G) show median (center line), interquartile range (box), and min-max values (whiskers). P values were calculated using a two-tailed paired t test. Each point represents a control- and myosin-bead pair acquired in the same field of view (N = 5, n = 11). See also fig. S2.

We next quantified the amount of a fluorescently labeled Arp2/3 complex (SNAP-Arp2/3 complex; see Materials and Methods) in the comet tail (Fig. 4, A to C). Myosin-beads showed substantially reduced levels of SNAP-Arp2/3 complex (P < 0.0001, n = 33, Fig. 4C) in the network. This reduced Arp2/3 complex level resulted in higher actin to Arp2/3 complex ratios on myosin-beads (Fig. 4D), indicating a sparse network organization with fewer branch points but longer filaments (15). Despite the reduced amount of Arp2/3 complex in the comet tails, we observed equivalent SNAP-Arp2/3 complex fluorescence on the control- and myosin-bead surfaces (Fig. 4E), suggesting that myosin-I does not affect the loading of Arp2/3 complex onto the bead-bound NPF. Rather, the Arp2/3 complex is recruited to NPF-coated beads and is ready for the arrival of a mother filament and G-actin to initiate branched nucleation. Together, myosin-beads have a less dense network structure as a result of reduced Arp2/3 complex–stimulated actin branching.

Fig. 4. Comets grown from myosin-beads incorporate less Arp2/3 complex.

(A) Representative actin comet tails assembled from (top) control and (bottom) myosin-beads showing (magenta) actin and (green) SNAP-Arp2/3 complex distribution. Images were acquired approximately 25 to 35 min after mixing. Brightness and contrast were set differently for actin (invert LUT) and SNAP-Arp2/3 complex (invert LUT) panels for visualization. Conditions: 4 μM actin (5% rhodamine-labeled), 200 nM Arp2/3 complex (80% SNAP-Surface 488–labeled), and 40 nM CP. Scale bars, 5 μm. (B) Total actin fluorescence intensity, (C) total SNAP-Arp2/3 fluorescence intensity, and (D) actin to SNAP-Arp2/3 fluorescence ratio over the entire comet tail region. (E) Total SNAP-Arp2/3 fluorescence intensity on bead surface. The plot shows the median (center line), interquartile range (box), and min-max values (whiskers). P values were calculated using a two-tailed paired t test. Each point represents a pair of control- and myosin-beads (N = 2 independent experiments, n = 33 pairs), with intensity values normalized to the control-beads.

We note that the myosin effect on growth efficiency depends on the CP concentration (Fig. 1D). Under low CP conditions (<50 nM), myosin-beads grew longer tails than control-beads with sparse networks, suggesting an even higher growth efficiency than the 50 nM CP case as quantified above. High CP concentrations (>50 nM) resulted in highly diffuse networks whose cohesion was easily compromised by myosin-I, causing actin dispersal, thus making their growth efficiency difficult to assess (12, 14, 17).

The myosin power stroke is required to alter network architecture

To examine whether the previously observed effects of myosin-I on actin assembly were due to its mechanochemical activity, we disabled the myosin-I motor activity by removing ATP, resulting in the population of a long-lived, actin-bound, rigor state (i.e., rigor myosin) (43). To maintain normal actin polymerization, G-actin was pretreated with ATP and gel filtered to eliminate free nucleotide (see Materials and Methods). Rigor myosin inhibited the formation of monopolar comet tails, while control-beads generated similar comet patterns as previously observed when free ATP was present (Fig. 5, A to C). At 50 nM CP, rigor myosin-beads formed dense actin shells, which subsequently fractured, forming multiple short tails of high network density (Fig. 5, A to C, fig. S3C, and movie S5). Restoring myosin motor activity by adding ATP reproduced the sparse comet architecture with high growth efficiency, as observed before (Fig. 5B and fig. S3B). At the two extreme CP conditions (15 and 200 nM), rigor myosin-beads neither enhanced symmetry breaking nor shed actin away (Fig. 5C and fig. S3D). Instead, actin shells were formed around the beads as a result of the strong actin-binding characteristics of rigor myosin. In addition to probing the effect of rigor myosin, we coupled biotinylated, Halo-tagged, actin-binding domain (Halo-ABD) of α-actinin to the beads, which binds actin filaments more dynamically than rigor myosin but does not undergo a power stroke (43, 49, 50). HaloABD-beads behaved similarly to rigor myosin (Fig. 5C, fig. S3A, and movie S6).

Fig. 5. The myosin power stroke is required for altering network architectures.

(A) Time series of actin assembly around (top) control and (bottom) rigor myosin-beads at 50 nM CP. Rigor myosin heavily delayed the growth of actin comet tails. (B) Representative images of actin comet tails grown under different ATP concentrations (top: control-beads; bottom: myosin-beads). Adding ATP back rescued comet tail growth. Images were acquired 20 to 30 min after mixing. (C) Representative actin network patterns assembled on control, myosin, rigor myosin, and HaloABD-beads under three different CP concentrations, as indicated. Images were acquired 20 to 30 min after mixing. Brightness and contrast were set to the same values for each panel. (D) Representative actin comet tails generated by (top) control and (bottom) myosin-beads in the absence and presence of 100 μM free Ca2+ at 50 nM CP. Images were acquired approximately 15 to 20 min after mixing. (E) Length and (F) network density quantified by total fluorescence intensity per area for control- and myosin-beads in the absence and presence of 100 μM free Ca2+ (N = 1 independent experiments, n = 15 pairs). Control and myosin-bead experimental pairs are normalized to the average fluorescence level of the control-beads. Box plots (E and F) show median (center line), interquartile range (box), and min-max values (whiskers). P values were calculated using a two-tailed paired t test. Conditions: 4 μM actin (5% rhodamine-labeled), 200 nM Arp2/3 complex, 15 to 200 nM CP, and 0 to 1 mM ATP, as indicated. Scale bars, 5 μm.

To further elucidate the role of myosin mechanochemistry in modulating the actin network, we uncoupled the dynamic actin binding and ATPase activity of myosin-I from its force-generating power stroke by adding calcium. Calcium weakens the affinity of lever-arm stabilizing calmodulin light chains, resulting in an inhibited power stroke while preserving ATPase activity (28, 51, 52). We confirmed calcium inhibition using the in vitro gliding assay, where the Myo1d-powered F-actin gliding (speed: 90.2 ±14.5 nm/s) was completely halted in the presence of 100 μM free calcium (fig. S3, E and F). Under these conditions, myosin-beads generated slightly shorter comet tails with network densities that were similar to control-beads (Fig. 5, D to F). We conclude that myosin power stroke is required to alter the network architecture, inducing a sparse actin organization with higher growth efficiency.

The myosin power stroke alone can fracture the actin shell

To assess the contribution of the force from the myosin power stroke independently of the force generated by actin polymerization, we arrested actin assembly around the bead using Arp2/3 complex inhibitor, CK-666, and the polymerization inhibitor, latrunculin B (LatB) (12, 53). These inhibitors were added ~100 s after the initiation of polymerization, before the fracturing of the actin shell that results in symmetry breaking, allowing us to monitor how myosin force affects the actin shell. Notably, most of the myosin-beads fractured the actin shell and ejected the bead 15 to 20 min after polymerization arrest. In contrast, control-beads remained enclosed in the actin shell without observable changes during a 40-min time window (Fig. 6A; also see fig. S4 and movies S7 and S8 acquired in the absence of phalloidin).

Fig. 6. The myosin power stroke can fracture the actin shell.

(A) Representative actin shells of control- and myosin-beads assembled under 50 nM CP, arrested by adding 20 μM (5 molar excess) of LatB and CK-666 before symmetry breaking, as well as myosin-beads assembled under the same conditions but arrested with the addition of 10 mM adenosine 5′-diphosphate (ADP) to inhibit myosin power stroke. Myosin-beads fractured and ejected from the actin shell, while control- and myosin-beads (with 10 mM ADP) remained enclosed in the shell. The control-bead assembled under 100 nM CP, showing similar network density as the myosin-bead, did not show shell fracture or bead ejection. The image was captured approximately 40 min after arrest. (B) The extents of shell breaking were classified by shell-breaking angle θ: θ = 0, no symmetry breaking; 0 < θ < 180, shell fracture; θ ≥180, bead ejected. Scale bar, 5 μm. (C) Percentage of populations with different extents of shell breaking for control (50 nM CP) (n = 182), myosin (n = 158), myosin (with 10 mM ADP) (n = 89), and control (100 nM CP) (n = 65); N = 2 independent experiments. Conditions: 4 μM actin (5% rhodamine-labeled), 200 nM Arp2/3, 50 or 100 nM CP. Twenty micromolar (5 molar excess) phalloidin and 2 mM ATP were also added to prevent actin depolymerization and preserve myosin motor activity. Actin assembly was arrested 100 s after mixing. Scale bars, 5 μm.

Shell fracture events were quantified by defining a shell-breaking angle, θ, where θ = 0 indicates no detected shell fracture; 0° < θ < 180° indicates shell fracture; and θ ≥ 180° indicates that the bead has been ejected from the shell (Fig. 6B). We found that 80% of the myosin-beads were ejected from the shell with an additional 11% showing shell fracture, while the corresponding control-beads showed only 16% shell fracture (Fig. 6C). Inhibition of myosin motor activity by adding 10 mM adenosine 5′-diphosphate (ADP) (43) together with LatB and CK-666 resulted in a substantial reduction in shell fracture events and eliminated bead ejection (Fig. 6, A and C, and fig. S4, C and D). To ensure that the shell breaking is not a result of the low actin shell density assembled around the myosin-beads, we performed control experiments with a higher CP concentration (100 nM) that created control-beads of lower actin shell densities (Fig. 6A) and found only 2% shell fracture (Fig. 6C). This further confirmed that the shell breaking was a direct result of myosin power stroke and is not due to differences in the shell network density.

Simulations show that myosin-I forces produce sparser actin networks and aid bead propulsion

We developed a filament-level computational model with an overall system size comparable to our experimental setup (see Materials and Methods; Fig. 7A and table S1). We incorporated the myosin-I power stroke as a repulsive force that pushes actin away from the bead surface and explored whether this myosin-induced pushing mechanism reproduces the experimentally observed comet tail patterns. In this model, we represented semiflexible actin filaments as beads connected by springs, polymerizing at their barbed ends and pushing against a spherical bead according to the Brownian ratchet–type force-elongation relationship. Spontaneous filament nucleation and branching at 70° angles occur close to the bead; elongation stops by capping when filaments reach a specified length. The effect of fluid and bead-filament friction was combined into a single bead friction parameter. Excluded volume interactions prevent filament crossing, resulting in tensile and compressive stresses developing within a shell of branched actin filaments that nucleate uniformly around an initially bare bead. By allowing filaments to break or debranch above a certain tensile force or branch angle threshold, we found that these networks can crack open, leading to symmetry breaking and bead propulsion (Fig. 7B and movie S9), as we observed in the experiments (Fig. 1C). To account for changes in CP concentration, we varied the average filament branch length in the simulations.

Fig. 7. The filament-level model of actin comet tail recapitulates experimental results.

(A) Schematic of the model of actin polymerization around the nucleating bead. The model includes filament-level nucleation, branching, filament fragmentation, debranching, capping, and force exerted by implicit myosin. (B) Time-lapse of the symmetry-breaking event under intermediate capping conditions (branch length = 0.5 μm) with no myosin. The color scale indicates filament tension (red: tensile; blue: compressive). (C) Tension distributions within actin comet tails formed at either control (0.0 pN) or two myosin forces (0.2 or 0.4 pN) under intermediate capping conditions (branch length = 0.5 μm). The image shows a cut through the center of the comet tail. The color scale indicates filament tension (red: tensile; blue: compressive). (D) Simulated time-lapse of epifluorescence images under different myosin forces (branch length = 0.5 μm). (E) Elongation speed, (F) actin intensity, and (G) Arp2/3 complex intensity for simulated beads as a function of myosin force (branch length = 0.5 μm). (H) Forces acting on beads along the direction of bead propulsion due to actin polymerization (green) and myosin pushing (orange) as a function of the myosin force (branch length = 0.5 μm). Error bars are SD. (I) Orientation of filaments around the beads (within 0.15 μm of the bead surface) as a function of myosin force (branch length = 0.5 μm). (J) Simulated symmetry breaking after actin polymerization and branching arrest. Actin was allowed to polymerize around the bead for 42.7 s (0.2-pN myosin force with 0.5-μm filament length) to form a shell of intermediate thickness before being halted. The time of arrest was set as t = 0 s.

The effect of the myosin-I power stroke was modeled as constant tangential pushing forces of magnitude Fmyo acting along every actin filament segment close to the bead, with equal and opposite force on the bead (Fig. 7A). With myosin-I pushing force incorporated, simulations recapitulated many of the experimental findings. For the short filament scenario (high CP concentration), simulated myosin-beads showed a notable delay or absence of symmetry breaking due to myosin pushing short filaments away from the bead surface (fig. S5, A and C, and movie S10). For the long filament scenario (low CP concentration), simulations reproduced the accumulation of a dense actin shell around the beads, which substantially inhibited the symmetry breaking of the control-beads (fig. S5, B and C, and movie S11). Myosin-beads, by contrast, promoted the growth of an asymmetric dense cloud (fig. S5, B and C), similar to the experimental observations in the early stages before bead ejection (Fig. 2B). We note that simulations do not account for the depletion of bulk actin occurring during late stages of bead ejection in the experiment (Fig. 2B). We also note a faster accumulation of actin in the simulations compared to the experiments (fig. S5B), which suggests additional factors limiting network growth not considered in the model. For the intermediate filament length scenario (i.e., intermediate CP concentration), simulations showed a similar elongation speed for the myosin-bead compared to the control-bead, albeit with a less dense actin network (Fig. 7, D to F, and movie S12), in agreement with the experimental observations (Fig. 3, A to F). Furthermore, the simulation predicted that actin filaments in the myosin comet tail experienced less stress (Fig. 7C), possibly due to the sparse actin organization, which reduces stresses arising from piling interconnected actin filaments on top of each other.

Notably, the ratio of actin to Arp2/3 complex density in simulated comet tails did not increase in the presence of myosin (Fig. 7, F to G, and fig. S6A), as observed in experiments (Fig. 4D). This fixed ratio is a consequence of our fixed-filament-length assumption to mimic a certain CP concentration. This assumption validates when the capping rate reduces equally as the actin polymerization rate in response to the opposing force, as reported by Li et al. (15, 19). However, the mismatch between our simulations and the experiments suggests an inequivalent effect on actin polymerization and CP capping due to the pushing force of myosin. We also tested whether the presence of myosin would promote actin debranching, which could also change the actin to Arp2/3 ratio. We found that myosin forces did not enhance debranching in our simulations (fig. S6). Less debranching occurred since comet tail stresses were smaller in the presence of myosin (Fig. 7C).

We next dissected the force contribution from different sources (actin polymerization or myosin power stroke) that powered the bead propulsion in our simulations. Notably, we found that while, in control-beads, the symmetry breaking and net comet propulsion were exclusively powered by actin polymerization, in myosin-beads, the myosin pushing forces contributed substantially to bead propulsion after symmetry breaking and had a predominant effect at high Fmyo (Fig. 7H and fig. S7). Simulated myosin forces also partly reorganized the network such that a larger fraction of actin filaments were polymerizing facing more perpendicular to the bead surface at larger myosin forces (Fig. 7I). When actin polymerization was arrested before shell breaking, simulations replicated the experimental findings that myosin pushing forces alone can eject the bead out of an actin shell (Fig. 7J and movie S13), provided that the shell is not too dense or too sparse (fig. S8).

Overall, our simulations with the myosin-I power stroke incorporated as an effective repulsive force replicate most of our experimental observations and support the experimental finding that myosin-I enhances force generation during branched actin assembly to promote shell breaking and enhances the efficiency of bead propulsion.

DISCUSSION

Myosin-I force generation and modulation of actin network density

The primary finding of this study is that Myo1d synergizes with the Arp2/3 complex to enhance the pushing forces of branched actin networks during assembly and that this force enhancement requires the myosin power stroke. We also found that Myo1d alters the actin network composition, producing a less dense actin network with decreased incorporation of the Arp2/3 complex.

How does myosin-I modulate Arp2/3 complex incorporation? We propose a model where myosin-I motor activity pushes the actin filaments away from the bead surface, reducing the accessibility of actin-binding sites for the NPF-activated Arp2/3 complex to bind and nucleate new branches (Fig. 8). Alternatively, the mechanochemistry of myosin-I may differently affect the incorporation of actin monomer and CP to filament ends in a way that promotes actin elongation while slowing capping (19, 54). Additional experiments that measure the actin elongation rate and capping kinetics near the NPF-coated surface are required to test this hypothesis.

Fig. 8. Schematic of how myosin-I modulates actin network structure through its power stroke.

(A) Schematic of branched actin assembly without the presence of myosin-I. (B) Schematic of branched actin assembly with the presence of myosin-I. Less dense actin networks in the presence of myosin-I result from the motor pushing actin filaments away from the NPF-coated surface through its force-generating power stroke.

We do not favor a model in which myosin-I sterically inhibits NPFs from binding and activating the Arp2/3 complex for three reasons. First, the concentration of the SNAP-Arp2/3 complex bound on beads is unchanged in the presence of myosin (Fig. 4E). Second, networks grown in the presence of calcium where the myosin power stroke was uncoupled from actin binding showed similar actin densities between myosin- and control-beads (Fig. 5, D and F), which ruled out the possibility that the sparse actin organization was due to the competition between myosin-I and the Arp2/3 complex for actin-binding sites. Last, calculations of molecular occupancy on bead surface through protein quantification verified that the steric hindrance effect of myosin-I is likely negligible (see the Supplementary Materials for detailed quantifications).

It remains possible that force generation by myosin-I debranches actin filaments, resulting in a less dense network with reduced Arp2/3 incorporation. Mechanical forces ranging from 0 to 2 pN dissociate actin branches from their mother filaments together with the Arp2/3 complex (55). A myosin-I paralog (Myo1b) that generates such forces has been shown to dissociate branches via motor activity (56). Although our simulations disfavor this hypothesis (fig. S6), a role for myosin-I–induced debranching should be explored further in the cell.

Last, it will be intriguing to explore these models further by performing experiments and computational modeling to probe network growth under mechanical load in the presence of myosin-I, especially given previous work that demonstrates the substantial loading effects on network architecture and power generation (15, 19).

Myosin-I cell biology

Myosin-Is connect the actin cytoskeleton to cellular membranes where they contribute to plasma membrane dynamics, organelle deformation, and shaping of actin network architecture. Although the molecular details of myosin-I function have been difficult to determine, recent studies suggest that some paralogs are powerful motors that have active roles in shaping membranes. For example, myosin-Is in budding yeast have mechanochemistry suitable for generating power, working with polymerizing actin to drive membrane invagination during endocytosis (24, 25, 38). Our current study confirms the ability of myosin-I to exert substantial pushing forces capable of fracturing actin networks. This power-generating capacity likely translates across species, with vertebrate Myo1c regulating actin architecture in diverse cellular regions (39, 57, 58) and Myo1e playing key roles in processes like endocytosis (59, 60), phagocytosis (27), and cancer invadosome formation (61, 62). Together, these findings point to some myosin-Is as dynamic players, actively shaping cellular structures and processes through their unique ability to both link and manipulate membranes and the actin cytoskeleton.

Not all paralogs are expected to modulate the actin networks as Myo1d. Notably, some myosin paralogs have substantially slower kinetics which are better suited for a motor that functions as a dynamic tether, providing force-dependent linkages between actin and membranes (23–25, 38, 39, 63, 64). Our experiments performed at low ATP concentrations (Fig. 5, A to C) mimic the behavior of myosins with slow motility rates and show clearly that slow kinetics can inhibit network fracturing and comet tail growth. Further studies to investigate how the intrinsic kinetic properties of different myosin-I paralogs influence actin polymerization and membrane dynamics will further reveal the diverse role of myosin-I function (24, 25, 27, 38, 39, 64).

It has been shown that myosins from classes I, III, V, VII, X, and XV function in actin protrusions (65), but their molecular roles are not clear. Recent studies demonstrate that membrane-bound myosin motors (myosin-10, -3a, and -15a) can facilitate membrane protrusion by providing an additional source of mechanical force at the distal tip to promote filipodia elongation (66, 67). We propose that myosin-I may exert a similar effect when associated with the membrane. Future research incorporating myosin-Is on lipid membranes will offer further perspectives into how myosin-I power membrane deformation within cells.

Summary

Overall, our study provides insights into how myosin-I molecules coordinate with the Arp2/3 complex, regulating the dynamics and architecture of the branched actin network and promoting the actin-based motile force generation (Fig. 8). This work sheds light on the synergy between myosin motor activity and actin polymerization, underscoring their collective role in driving morphological changes at the cellular membrane interfaces. Future work to determine the molecular mechanism by which myosin-I affects actin network architecture and polymerization forces will reveal the molecular roles of this important myosin family.

MATERIALS AND METHODS

Protein purification

Actin was purified from rabbit skeletal muscle acetone powder as previously described (68). Monomeric G-actin was purified by gel filtration on Sephacryl S-300 in G-buffer [2 mM tris-HCl (pH 8.0), 0.2 mM ATP, 0.1 mM CaCl2, 1 mM NaN3, and 0.5 mM dithiothreitol (DTT)] and used within 2 to 3 weeks. Actin was labeled with 5/6-carboxy-tetramethyl-rhodamine succinimidyl ester (NHS-rhodamine) at random surface lysine residues (69). Full-length Drosophila Myo1d, with C-terminal FLAG-Avi tags, was expressed, purified as previously described (70), and subsequently biotinylated at the C-termini Avi tag sequence via BirA biotin-protein ligase (Avidity) (43). The GST-tagged WCA domain of human WASP protein (GST-WCA) was purchased from Cytoskeleton (catalog no. VCG03-A) and used without further purification. The Arp2/3 complex was isolated from the bovine brain as previously described (71). The SNAP-tagged Arp2/3 complex was constructed and purified as described (72). The SNAP-tagged Arp2/3 complex was labeled with SNAP-Surface 488 (Biolab, catalog no. S9124S) using commercially provided protocol. Human CapZ was expressed and purified as previously described (73). Halo-ABD was constructed and purified as described in (74). (See also the Supplementary Materials for further details.)

Bead preparation

Carboxylate polystyrene beads (Polybead, 2.0-μm diameter, catalog no. 18327-10) were purchased from Polysciences. Beads were coated with NPF and neutravidin following a previous protocol (40) with modifications. Briefly, 5 to 10 μl bead slurry was washed with X buffer [10 mM Hepes (pH 7.5), 100 mM KCl, 1 mM MgCl2, 100 μM CaCl2, and 1 mM ATP], and then incubated with 50 to 100 μl of 2.3 μM GST-WCA (0.1 mg/ml) and various concentrations of neutravidin (Thermo Fisher Scientific, catalog no. 31000) [8.3 μM (0.5 mg/ml) for #myosin/NPF = 0.28:1, 16.7 μM (1 mg/ml) for #myosin/NPF = 0.35:1, 33.3 μM (2 mg/ml) for #myosin/NPF = 0.43:1, or 83.3 μM (5 mg/ml) for #myosin/NPF = 0.80:1] on a slow rotator at 4°C for 2 hours. Beads were then pelleted by spinning at 16,000g for 2 min at 4°C to remove the unreacted reagents and then resuspended in 200 to 400 μl of bovine serum albumin (BSA; 10 mg/ml), incubating on ice for 30 min to block the free space left on the bead surface. Last, beads were washed twice and stored in 50 to 100 μl of BSA (1 mg/ml) in X buffer for up to 3 days.

The NPF- and neutravidin-coated beads were next split in half, with one half coupled with biotinylated Drosophila Myo1d (myosin-bead) or biotinylated Halo-ABD (HaloABD-bead), and the other half coated with Biotin-CF640 (Biotium, catalog no. 80032) fluorescence dye (control-bead). The NPF- and neutravidin-coated beads were first washed with M buffer [20 mM Hepes (pH 7.5), 100 mM KCl, 1 mM MgCl2, 1 mM EGTA, and 2 mM ATP] to remove the free calcium in the storage X buffer and were then incubated with 1 μM biotinylated Drosophila Myo1d, 1 μM Halo-ABD, or 1 μM Biotin-CF640 fluorescence dye for 30 min on ice. Beads were then pelleted under 16,000g for 2 min at 4°C to remove the free unbound reagents, washed with BSA (1 mg/ml) in M buffer, and used immediately for motility assay on the same day after preparation.

Bead motility assay/comet tail assay

Unless specified otherwise, a typical motility mixture contained 4 μM actin (5% rhodamine-labeled), 200 nM Arp2/3 complex or SNAP-Arp2/3 complex, 6.5 to 200 nM CP, 2 μM calmodulin, and 3 μl of bead slurry (1.5 μl of myosin-beads or HaloABD-beads and 1.5 μl of control-beads), mixed in 20 mM Hepes (pH 7.5), 100 mM KCl, 1 mM MgCl2, 1 mM EGTA, 1 mM MgATP, 40 mM DTT, BSA (10 mg/ml), and 0.2% methylcellulose, contributing to a final volume of 50 μl. The activity of the SNAP-Arp2/3 complex is slightly lower than the unlabeled complex, so we changed the CP concentration (40 nM) to achieve similar tail lengths, actin densities, and growth efficiencies as observed for the native complex (Fig. 4). For experiments with calcium present, G-actin was preincubated with 200 μM EGTA and 50 μM MgCl2 for 5 min to be exchanged to Mg–G-actin before using. Calcium experiments included 1.1 mM CaCl2 in place of calmodulin to disrupt the myosin power stroke. Beads were first mixed with all other reagents, excluding actin, with a pipette, to ensure an even distribution. The reaction was then initiated by adding actin into the system, mixed thoroughly, and denoted as time 0. Slides and coverslips were wiped with 70% ethanol and ddH2O, followed by a plasma cleansing for 10 min. Upon mixing, a volume of 2.1-μl motility mixture was carefully applied between a glass slide and a coverslip (22 mm by 22 mm) forming a so-called “squeeze chamber” with a height of 4.3 μm. The squeeze chamber was then sealed with vacuum grease and imaged immediately under the microscope. Time-lapse movies were acquired of microscope fields that included both myosin- and control-beads. In most cases, the actin comet tails emerging from myosin or control-beads grew with a constant speed during the first ~10 min following symmetry breaking. As the reagents in the polymerization mixture depleted, tail elongation slowed and eventually stopped ~30 min after mixing.

Fluorescence imaging and data analysis

Fluorescence microscopy imaging was performed via Leica DMIRB epifluorescence microscope (100×, oil-immersive objective of numerical aperture 1.4) with Leica EL6000 external light source (120 w mercury metal halide short arc lamp, Osram HXP R 120 W/45 C VIS), Retiga R6 CCD camera (TELEDYNE), and Metamorph (Molecular Devices) imaging software. Movies were recorded at 25°C and acquired every 10 s for 30 to 60 min. Exposure time was 200 ms for most experiments, and 1 s to image the SNAP-Surface 488–labeled Arp2/3 complex. Images were analyzed and quantified using Fiji software. Actin comet tail length was measured manually at each frame using the segmented line draw tool (from the end of the tail to the center of the bead) and converted to micrometers. The tail growth rate and fluorescent assembly rate were calculated by fitting the first 7 to 10 data points in the time courses to get the initial slope of the growth and assembly. Growth efficiency was determined by dividing the tail growth rate by the fluorescent assembly rate. Image brightness and contrast were carefully adjusted using Fiji and Adobe Illustrator. Unless otherwise specified, the lookup table for each pair of control- and myosin-bead (or HaloABD-bead) was kept the same.

Statistical analysis

The statistical significance was calculated using paired or unpaired two-tailed Student’s t test in GraphPad Prism v9.0. Further details are described in figure legends.

Modeling methods

We simulated actin comet tail formation at the level of individual filaments, each represented as a series of segments of length l0, following earlier work (75). The pointed ends of actin filament branches are assumed to be connected to a point element of a mother filament at the location of the Arp2/3 complex. Such a filament representation allows us to model the effect of myosin as a tangential force acting along filament segments close to the nucleating bead. We thus generalize earlier filament models that did not explicitly account for filament bending mechanics (47, 76–78), earlier mechanical models that did not monitor the whole network of actin filaments (48, 79–82), or modeled the full process of symmetry breaking and propulsion (83). We did not explicitly consider diffusiophoretic contributions to bead motion (84). (Also see the Supplementary Materials for further details.)

Forces on actin filaments

The position ri of point element i of actin filaments/filament branches evolves according to dridt=Fispring+Fibend+Fiangle+Fiexcluded+Fimyo.

Here, ζ is an effective filament segment drag coefficient that allows the actin network to evolve through approximate quasi-static mechanical equilibrium while also providing numerical stability. The spring force is Fispring=kactindii+1−l0d^ii+1−kactindi−1i−l0d^i−1i where (i − 1) and (i + 1) are neighboring point elements (if they exist) before and after i, dij is the separation distance between i and j, and d^ij is the unit vector from i to j. The equilibrium length is l0, except for (i) uncapped barbed ends that elongate on average according to l0τ=l00+δr0pol τ after their initiation at τ = 0, in discrete steps of half-monomer size δ = 2.7 nm (see below) and (ii) a short branch segment joining the mother filament point element to the daughter pointed end, which has length of lbranch.

The bending force is Fibend=κ/lavg∂d^ii+1∙d^i−1i/∂ri+∂d^i−1i∙d^i−2i−1/∂ri+∂d^i+1i+2∙d^ii+1/∂ri, κ=kBTlp is the flexural rigidity, lp is the persistence length of the actin filament, and lavg is the average length of the two filament segments composing the angle. For the straight angle that exists among the first three beads in the connection between mother and daughter filaments, lavg = l0 is used for numerical stability.

An angular potential keeps Arp2/3 complex branches at 70°. The angular force is Fmangle=ϵanglecosθijk−cosθ0 ∂d^jk∙d^ij/∂rm, where ϵangle is a spring constant; i, j, and k are the indices of the point elements that make up the angle; and m is one of these indices.

The excluded volume force is due to repulsion between two actin filaments and between actin filaments and the nucleating bead. The former prevents the crossing of filaments and is exerted along the direction of vector dαβ that connects the two closest approach points on the filament segments α and β (85). It is modeled as a stiff spring force with a max range of dexcluded, with Fαexcluded,actin=−Fβexcluded,actin=kexcl dαβ−dexcludedd^αβ, where dαβ is the minimum distance between neighboring filament segments. This force is distributed to the end points of filament segments α and β (including element i) according to a lever arm rule. Although the typical diameter of actin filaments is around 7 nm, we set dexcluded to be 20 nm to mimic the effect of thermal fluctuations (not included in our simulations). The lateral distance by which thermal fluctuations bend a filament of length L held at two ends by flexible hinges is approximately 21/2L3/2/π2lp1/2 for the longest bending mode (86). For a filament segment of length 0.5 μm, this distance is 12 nm, to which we add a value of order the typical filament diameter to arrive at 20 nm. In addition, since the excluded volume is modeled as a stiff spring rather than a hard-core potential, some level of overlap between segments is possible. At a separation of order 7 nm, the excluded volume force is 22 pN. Filament segments experience a radially oriented excluded volume force with the nucleating bead if they are closer to the nucleator than R; this force can be written as Fiexcluded,bead=kexcldiN−Rd^iN , where diN is the closest approach distance between the filament segment and the nucleator bead.

Myosin forces are given by Fimyo=Fmyod^ii−1 , where i − 1 is the neighboring actin filament point element along the pointed end direction. It acts on all filament elements i closer than dmyo to the nucleator bead surface. The myosin force acts tangentially along the segment toward the pointed end. We do not model the individual binding, lever arm motion, and unbinding of myosin; instead, the magnitude of Fmyo approximates a time and ensemble average over many binding and unbinding cycles.

Nucleating bead motion

The nucleating bead evolves through time according toζnucl beaddrnucl beaddt=−∑iFiexcluded,bead+Fimyosin

where the sum is over all actin segments that contact the bead through excluded volume or myosin interactions, according to the interaction distances defined above. We assume the relative motion between the actin network and nucleating bead is dominated by frictional forces between them. We used a value of ζnucl bead that was large enough to prevent the rapid ejection of bare beads out of a shell at the onset of symmetry breaking, a phenomenon that is not seen in our experiments. We thus monitor the relative motion of the bead and actin network in the limit of quasi-static mechanical equilibrium of the actin network, including implicit transient attachment and detachment of actin filaments to the bead. This approximation does not account for the varying concentration of actin near the bead or the absolute motion of actin comet and bead in the laboratory frame, which, in reality, would be influenced by small forces between the bead or actin and the glass slide.

Actin barbed end polymerization rate

Polymerization of the barbed end of uncapped filaments occurs with the last segment of the actin filament lengthening in increments of half-monomer size δ at a rate given by the polymerization rate. If the end segment reaches a length of l0, then a new segment is added with an initial length of l00. Polymerization of the barbed end is attenuated by compressive forces on the spring bond connecting the barbed end point element to the rest of the filament. The polymerization rate is rpolT=r0polexp−T δ/kBT , where the free filament elongation rate is r0pol and T=∣Fispring∣ when Fispring is the compressive tension on the barbed end point element i [otherwise rpolT=r0pol ] (87, 88). Because each filament segment starts with an initial length of l00 (for numerical stability), the rate rpol(0) is effectively multiplied by a factor of 1.12.

Barbed end capping

Filaments polymerize until they reach a final length specified for a given simulation, Lfil (an integer multiple of l0), which we vary to simulate the effect of varying CP concentration. Here, we did not study the effects of a varying filament length distribution. The assumption that Lfil is independent of the force is based on prior experiments (19) and the fact that the filament length added by CP is close to δ. According to this evidence, the polymerization to capping rate ratio, as well as the average filament length, remains unchanged by force for a given CP concentration.

Branching

Branching occurs from actin filament point elements that are within l0 of the nucleating bead, at rate r0branch , independent of the CP concentration. The rate was chosen to approximate the timescale of symmetry breaking and comet speed at intermediate CP. The segment length connecting the mother filament point element to the daughter filament pointed end point element is lbranch. Their orientation is chosen according to a uniform angle distribution in the cone around the mother filament opening toward the barbed end. To maintain a discretization of the network at segment length l0, branches cannot branch from the barbed end point element of the filament. Branches nucleating from the same mother filament have no torsional restriction (i.e., daughter filaments can rotate about the axis of the mother filament without restriction), but implicit torsional restrictions are imposed because of the dense surrounding network limiting this motion.

De novo nucleation

Nucleation of new filaments of length l00 occurs at a rate of r0denovo . These small filaments are introduced with a uniform spherical orientation with either their pointed or barbed end touching the nucleating bead surface. This slow rate of filament introduction was tuned to allow for the startup of the network on timescales comparable to the experiment and to allow for the buildup of a thin actin cloud around the leading bead edge during actin comet propulsion, as observed in experiments. At the beginning of the simulation, 200 of these de novo filaments are added to provide enough initial filaments for the symmetric growth of the shell.

Filament fragmentation and debranching

For simplicity, filament fragmentation was assumed to occur above a certain tensile force threshold Ffrag (instead of implementing a rate of severing as a function of tension).

For filament segments, the threshold was Ffrag = 50 pN. This is lower than the experimental fragmentation force of phalloidin-labeled actin filaments, which is several hundred piconewtons (89), but we found that the shell would have difficulty breaking unless we had this lower force. Once a filament segment is fragmented, the filament with the newly created barbed end is left uncapped and can grow to the final length specified for that simulation. Debranching occurs for branches that have a branch bond with tension greater than F debranch = 30 pN or if the angle deviates by more than Δθdebranch = 25° off of the 70° equilibrium angle. The value of Δθdebranch was chosen to allow easy debranching when the branch is bent [similar to experiments where branches were bent and pulled by fluid forces of order piconewton (55)]. The value of F debranch had to be sufficiently high such that the network does not easily fall apart.

Acknowledgments

We thank D. Safer, F. A. Baez-Cruz, and R. Wike for kind help with protein purifications and technical assistance on the project. We thank everyone in Ostap Lab and Dominguez Lab for valuable inputs throughout the project duration. D.V. is a visiting scholar at the Center for Computational Biology of Flatiron Institute, Simons Foundation.

Funding: This work was supported by NIH grants R37 GM057247 (to E.M.O.) and R01 GM073791 (to R.D.). E.M.O. and M.X. were partially supported by the National Science Foundation (CMMI: 15-48571). D.V. and D.M.R. were supported by NIH grant R35 GM136372. Portions of this research were conducted on the Rockfish (Johns Hopkins) cluster through allocations MCB180021 and BIO230116 from the Advanced Cyberinfrastructure Coordination Ecosystem: Services & Support (ACCESS) program, which is supported by NSF grants #2138259, #2138286, #2138307, #2137603, and #2138296.

Author contributions: Writing—original draft: M.X., D.M.R., D.V., and E.M.O. Conceptualization: M.X., D.M.R., L.W.P., D.V., and E.M.O. Writing—review and editing: M.X., D.M.R., L.W.P., R.D., D.V., and E.M.O. Methodology: M.X., D.M.R., L.W.P., D.V., and E.M.O. Formal analysis: M.X., D.M.R., and E.M.O. Investigation: M.X. and D.M.R. Resources: G.R., M.B., D.V., and R.D. Visualization: M.X. and D.M.R. Supervision: L.W.P., R.D., D.V., and E.M.O. Data curation: M.X., D.M.R., D.V., and E.M.O. Validation: M.X., D.M.R., L.W.P., and E.M.O. Software: D.M.R. Funding acquisition: D.V., R.D., and E.M.O. Project administration: L.W.P., R.D., D.V., and E.M.O.

Competing interests: The authors declare that they have no competing interests.

Data and materials availability: All data needed to evaluate the conclusions in the paper are present in the paper and/or the Supplementary Materials. The code for the computational model can be found at https://github.com/davidmrutkowski/ActinCometTailMyo1 or https://zenodo.org/records/11289071. Other supplementary materials can be found at https://zenodo.org/records/11541677.

Supplementary Materials

The PDF file includes:

Supplementary Text

Figs. S1 to S8

Table S1

Legends for movies S1 to S13

Other Supplementary Material for this manuscript includes the following:

Movies S1 to S13
==== Refs
REFERENCES AND NOTES

1 L. Blanchoin, R. Boujemaa-Paterski, C. Sykes, J. Plastino, Actin dynamics, architecture, and mechanics in cell motility. Physiol. Rev. 94 , 235–263 (2014).24382887
2 K. Rottner, J. Faix, S. Bogdan, S. Linder, E. Kerkhoff, Actin assembly mechanisms at a glance. J. Cell Sci. 130 , 3427–3435 (2017).29032357
3 T. Svitkina, The actin cytoskeleton and actin-based motility. Cold Spring Harb. Perspect. Biol. 10 , a018267 (2018).29295889
4 V. Papalazarou, L. M. Machesky, The cell pushes back: The Arp2/3 complex is a key orchestrator of cellular responses to environmental forces. Curr. Opin. Cell Biol. 68 , 37–44 (2021).32977244
5 A. M. Gautreau, F. E. Fregoso, G. Simanov, R. Dominguez, Nucleation, stabilization, and disassembly of branched actin networks. Trends Cell Biol. 32 , 421–432 (2022).34836783
6 M. Krendel, N. C. Gauthier, Building the phagocytic cup on an actin scaffold. Curr. Opin. Cell Biol. 77 , 102112 (2022).35820329
7 M. Jin, C. Shirazinejad, B. Wang, A. Yan, J. Schoneberg, S. Upadhyayula, K. Xu, D. G. Drubin, Branched actin networks are organized for asymmetric force production during clathrin-mediated endocytosis in mammalian cells. Nat. Commun. 13 , 3578 (2022).35732852
8 D. N. Clarke, A. C. Martin, Actin-based force generation and cell adhesion in tissue morphogenesis. Curr. Biol. 31 , R667–R680 (2021).34033797
9 T. P. Loisel, R. Boujemaa, D. Pantaloni, M. F. Carlier, Reconstitution of actin-based motility of Listeria and Shigella using pure proteins. Nature 401 , 613–616 (1999).10524632
10 F. Nakamura, E. Osborn, P. A. Janmey, T. P. Stossel, Comparison of filamin A-induced cross-linking and Arp2/3 complex-mediated branching on the mechanics of actin filaments. J. Biol. Chem. 277 , 9148–9154 (2002).11786548
11 O. Chaudhuri, S. H. Parekh, D. A. Fletcher, Reversible stress softening of actin networks. Nature 445 , 295–298 (2007).17230186
12 O. Akin, R. D. Mullins, Capping protein increases the rate of actin-based motility by promoting filament nucleation by the Arp2/3 complex. Cell 133 , 841–851 (2008).18510928
13 J. Stricker, T. Falzone, M. L. Gardel, Mechanics of the F-actin cytoskeleton. J. Biomech. 43 , 9–14 (2010).19913792
14 A. Kawska, K. Carvalho, J. Manzi, R. Boujemaa-Paterski, L. Blanchoin, J. L. Martiel, C. Sykes, How actin network dynamics control the onset of actin-based motility. Proc. Natl. Acad. Sci. U.S.A. 109 , 14440–14445 (2012).22908255
15 P. Bieling, T. D. Li, J. Weichsel, R. McGorty, P. Jreij, B. Huang, D. A. Fletcher, R. D. Mullins, Force feedback controls motor activity and mechanical properties of self-assembling branched actin networks. Cell 164 , 115–127 (2016).26771487
16 J. Mueller, G. Szep, M. Nemethova, I. de Vries, A. D. Lieber, C. Winkler, K. Kruse, J. V. Small, C. Schmeiser, K. Keren, R. Hauschild, M. Sixt, Load adaptation of lamellipodial actin networks. Cell 171 , 188–200.e16 (2017).28867286
17 R. D. Mullins, P. Bieling, D. A. Fletcher, From solution to surface to filament: Actin flux into branched networks. Biophys. Rev. 10 , 1537–1551 (2018).30470968
18 J. Funk, F. Merino, M. Schaks, K. Rottner, S. Raunser, P. Bieling, A barbed end interference mechanism reveals how capping protein promotes nucleation in branched actin networks. Nat. Commun. 12 , 5329 (2021).34504078
19 T. D. Li, P. Bieling, J. Weichsel, R. D. Mullins, D. A. Fletcher, The molecular mechanism of load adaptation by branched actin networks. eLife 11 , e73145 (2022).35748355
20 A. Colin, T. Kotila, C. Guerin, M. Orhant-Prioux, B. Vianay, A. Mogilner, P. Lappalainen, M. Thery, L. Blanchoin, Recycling of the actin monomer pool limits the lifetime of network turnover. EMBO J. 42 , e112717 (2023).36912152
21 T. D. Pollard, E. D. Korn, Acanthamoeba myosin. J. Biol. Chem. 248 , 4682–4690 (1973).4268863
22 B. B. McIntosh, E. M. Ostap, Myosin-I molecular motors at a glance. J. Cell Sci. 129 , 2689–2695 (2016).27401928
23 C. G. Almeida, A. Yamada, D. Tenza, D. Louvard, G. Raposo, E. Coudrier, Myosin 1b promotes the formation of post-Golgi carriers by regulating actin assembly and membrane remodelling at the trans-Golgi network. Nat. Cell Biol. 13 , 779–789 (2011).21666684
24 R. T. A. Pedersen, D. G. Drubin, Type I myosins anchor actin assembly to the plasma membrane during clathrin-mediated endocytosis. J. Cell Biol. 218 , 1138–1147 (2019).30659101
25 H. E. Manenschijn, A. Picco, M. Mund, A. S. Rivier-Cordey, J. Ries, M. Kaksonen, Type-I myosins promote actin polymerization to drive membrane bending in endocytosis. eLife 8 , e44215 (2019).31385806
26 A. Capmany, A. Yoshimura, R. Kerdous, V. Caorsi, A. Lescure, E. Del Nery, E. Coudrier, B. Goud, K. Schauer, MYO1C stabilizes actin and facilitates the arrival of transport carriers at the Golgi complex. J. Cell Sci. 132 , jcs225029 (2019).30872458
27 S. R. Barger, N. S. Reilly, M. S. Shutova, Q. Li, P. Maiuri, J. M. Heddleston, M. S. Mooseker, R. A. Flavell, T. Svitkina, P. W. Oakes, M. Krendel, N. C. Gauthier, Membrane-cytoskeletal crosstalk mediated by myosin-I regulates adhesion turnover during phagocytosis. Nat. Commun. 10 , 1249 (2019).30890704
28 M. J. Greenberg, E. M. Ostap, Regulation and control of myosin-I by the motor and light chain-binding domains. Trends Cell Biol. 23 , 81–89 (2013).23200340
29 R. J. Adams, T. D. Pollard, Binding of myosin I to membrane lipids. Nature 340 , 565–568 (1989).2770861
30 S. K. Doberstein, T. D. Pollard, Localization and specificity of the phospholipid and actin binding sites on the tail of Acanthamoeba myosin IC. J. Cell Biol. 117 , 1241–1249 (1992).1607386
31 E. A. Feeser, C. M. G. Ignacio, M. Krendel, E. M. Ostap, Myo1e binds anionic phospholipids with high affinity. Biochemistry 49 , 9353–9360 (2010).20860408
32 S. M. Hayden, J. S. Wolenski, M. S. Mooseker, Binding of brush border myosin I to phospholipid vesicles. J. Cell Biol. 111 , 443–451 (1990).2143194
33 D. E. Hokanson, J. M. Laakso, T. Lin, D. Sept, E. M. Ostap, Myo1c binds phosphoinositides through a putative pleckstrin homology domain. Mol. Biol. Cell 17 , 4856–4865 (2006).16971510
34 H. Miyata, B. Bowers, E. D. Korn, Plasma membrane association of Acanthamoeba myosin I. J. Cell Biol. 109 , 1519–1528 (1989).2793931
35 R. Rohatgi, L. Ma, H. Miki, M. Lopez, T. Kirchhausen, T. Takenawa, M. W. Kirschner, The interaction between N-WASP and the Arp2/3 complex links Cdc42-dependent signals to actin assembly. Cell 97 , 221–231 (1999).10219243
36 T. Takenawa, S. Suetsugu, The WASP-WAVE protein network: Connecting the membrane to the cytoskeleton. Nat. Rev. Mol. Cell Biol. 8 , 37–48 (2007).17183359
37 D. A. Kramer, H. K. Piper, B. Chen, WASP family proteins: Molecular mechanisms and implications in human disease. Eur. J. Cell Biol. 101 , 151244 (2022).35667337
38 R. T. A. Pedersen, A. Snoberger, S. Pyrpassopoulos, D. Safer, D. G. Drubin, E. M. Ostap, Endocytic myosin-1 is a force-insensitive, power-generating motor. J. Cell Biol. 222 , e202303095 (2023).37549220
39 A. M. Sokac, C. Schietroma, C. B. Gundersen, W. M. Bement, Myosin-1c couples assembling actin to membranes to drive compensatory endocytosis. Dev. Cell 11 , 629–640 (2006).17084356
40 R. Boujemaa-Paterski, R. Galland, C. Suarez, C. Guerin, M. Thery, L. Blanchoin, Directed actin assembly and motility. Methods Enzymol. 540 , 283–300 (2014).24630113
41 V. Noireaux, R. M. Golsteyn, E. Friederich, J. Prost, C. Antony, D. Louvard, C. Sykes, Growing an actin gel on spherical surfaces. Biophys. J. 78 , 1643–1654 (2000).10692348
42 J. H. Lewis, T. Lin, D. E. Hokanson, E. M. Ostap, Temperature dependence of nucleotide association and kinetic characterization of myo1b. Biochemistry 45 , 11589–11597 (2006).16981718
43 F. A. Baez-Cruz, E. M. Ostap, Drosophila class-I myosins that can impact left-right asymmetry have distinct ATPase kinetics. J. Biol. Chem. 299 , 104961 (2023).37380077
44 L. A. Cameron, M. J. Footer, A. van Oudenaarden, J. A. Theriot, Motility of ActA protein-coated microspheres driven by actin polymerization. Proc. Natl. Acad. Sci. U.S.A. 96 , 4908–4913 (1999).10220392
45 A. Bernheim-Groswasser, S. Wiesner, R. M. Golsteyn, M. F. Carlier, C. Sykes, The dynamics of actin-based motility depend on surface parameters. Nature 417 , 308–311 (2002).12015607
46 J. van der Gucht, E. Paluch, J. Plastino, C. Sykes, Stress release drives symmetry breaking for actin-based movement. Proc. Natl. Acad. Sci. U.S.A. 102 , 7847–7852 (2005).15911773
47 V. Achard, J. L. Martiel, A. Michelot, C. Guerin, A. C. Reymann, L. Blanchoin, R. Boujemaa-Paterski, A “primer”-based mechanism underlies branched actin filament network formation and motility. Curr. Biol. 20 , 423–428 (2010).20188562
48 M. J. Dayel, O. Akin, M. Landeryou, V. Risca, A. Mogilner, R. D. Mullins, In silico reconstitution of actin-based symmetry breaking and motility. PLOS Biol. 7 , e1000201 (2009).19771152
49 D. H. Wachsstock, W. H. Schwarz, T. D. Pollard, Cross-linker dynamics determine the mechanical properties of actin gels. Biophys. J. 66 , 801–809 (1994).8011912
50 P. A. Kuhlman, J. Ellis, D. R. Critchley, C. R. Bagshaw, The kinetics of the interaction between the actin-binding domain of alpha-actinin and F-actin. FEBS Lett. 339 , 297–301 (1994).8112470
51 J. H. Lewis, M. J. Greenberg, J. M. Laakso, H. Shuman, E. M. Ostap, Calcium regulation of myosin-I tension sensing. Biophys. J. 102 , 2799–2807 (2012).22735530
52 S. Manceva, T. Lin, H. Pham, J. H. Lewis, Y. E. Goldman, E. M. Ostap, Calcium regulation of calmodulin binding to and dissociation from the myo1c regulatory domain. Biochemistry 46 , 11718–11726 (2007).17910470
53 B. J. Nolen, N. Tomasevic, A. Russell, D. W. Pierce, Z. Jia, C. D. McCormick, J. Hartman, R. Sakowicz, T. D. Pollard, Characterization of two classes of small molecule inhibitors of Arp2/3 complex. Nature 460 , 1031–1034 (2009).19648907
54 P. Bieling, S. D. Hansen, O. Akin, T. D. Li, C. C. Hayden, D. A. Fletcher, R. D. Mullins, WH2 and proline-rich domains of WASP-family proteins collaborate to accelerate actin filament elongation. EMBO J. 37 , 102–121 (2018).29141912
55 N. G. Pandit, W. Cao, J. Bibeau, E. M. Johnson-Chavarria, E. W. Taylor, T. D. Pollard, E. M. De La Cruz, Force and phosphate release from Arp2/3 complex promote dissociation of actin filament branches. Proc. Natl. Acad. Sci. U.S.A. 117 , 13519–13528 (2020).32461373
56 J. Pernier, A. Morchain, V. Caorsi, A. Bertin, H. Bousquet, P. Bassereau, E. Coudrier, Myosin 1b flattens and prunes branched actin filaments. J. Cell Sci. 133 , jcs247403 (2020).32895245
57 F. S. Wang, C. W. Liu, T. J. Diefenbach, D. G. Jay, Modeling the role of myosin 1c in neuronal growth cone turning. Biophys. J. 85 , 3319–3328 (2003).14581233
58 J. L. Maravillas-Montero, P. G. Gillespie, G. Patino-Lopez, S. Shaw, L. Santos-Argumedo, Myosin 1c participates in B cell cytoskeleton rearrangements, is recruited to the immunologic synapse, and contributes to antigen presentation. J. Immunol. 187 , 3053–3063 (2011).21841128
59 J. Cheng, A. Grassart, D. G. Drubin, Myosin 1E coordinates actin assembly and cargo trafficking during clathrin-mediated endocytosis. Mol. Biol. Cell 23 , 2891–2904 (2012).22675027
60 M. Krendel, E. K. Osterweil, M. S. Mooseker, Myosin 1E interacts with synaptojanin-1 and dynamin and is involved in endocytosis. FEBS Lett. 581 , 644–650 (2007).17257598
61 J. L. Ouderkirk, M. Krendel, Myosin 1e is a component of the invadosome core that contributes to regulation of invadosome dynamics. Exp. Cell Res. 322 , 265–276 (2014).24462457
62 M. E. Garone, S. E. Chase, C. Zhang, M. Krendel, Myosin 1e deficiency affects migration of 4T1 breast cancer cells. Cytoskeleton, doi.org/10.1002/cm.21819, (2023).
63 Y. Sun, A. C. Martin, D. G. Drubin, Endocytic internalization in budding yeast requires coordinated actin nucleation and myosin motor activity. Dev. Cell 11 , 33–46 (2006).16824951
64 J. M. Laakso, J. H. Lewis, H. Shuman, E. M. Ostap, Myosin I can act as a molecular force sensor. Science 321 , 133–136 (2008).18599791
65 A. Houdusse, M. A. Titus, The many roles of myosins in filopodia, microvilli and stereocilia. Curr. Biol. 31 , R586–R602 (2021).34033792
66 G. N. Fitz, M. L. Weck, C. Bodnya, O. L. Perkins, M. J. Tyska, Protrusion growth driven by myosin-generated force. Dev. Cell 58 , 18–33.e6 (2023).36626869
67 J. A. Cirilo Jr., X. Liao, B. J. Perrin, C. M. Yengo, The dynamics of actin protrusions can be controlled by tip-localized myosin motors. J. Biol. Chem. 300 , 105516 (2024).38042485
68 J. A. Spudich, S. Watt, The regulation of rabbit skeletal muscle contraction. I. Biochemical studies of the interaction of the tropomyosin-troponin complex with actin and the proteolytic fragments of myosin. J. Biol. Chem. 246 , 4866–4871 (1971).4254541
69 D. R. Kellogg, T. J. Mitchison, B. M. Alberts, Behaviour of microtubules and actin filaments in living Drosophila embryos. Development 103 , 675–686 (1988).3248521
70 G. Lebreton, C. Geminard, F. Lapraz, S. Pyrpassopoulos, D. Cerezo, P. Speder, E. M. Ostap, S. Noselli, Molecular to organismal chirality is induced by the conserved myosin 1D. Science 362 , 949–952 (2018).30467170
71 M. Boczkowska, G. Rebowski, M. V. Petoukhov, D. B. Hayes, D. I. Svergun, R. Dominguez, X-ray scattering study of activated Arp2/3 complex with bound actin-WCA. Structure 16 , 695–704 (2008).18462674
72 A. Zimmet, T. Van Eeuwen, M. Boczkowska, G. Rebowski, K. Murakami, R. Dominguez, Cryo-EM structure of NPF-bound human Arp2/3 complex and activation mechanism. Sci. Adv. 6 , eaaz7651 (2020).32917641
73 J. N. Rao, Y. Madasu, R. Dominguez, Mechanism of actin filament pointed-end capping by tropomodulin. Science 345 , 463–467 (2014).25061212
74 M. J. Greenberg, H. Shuman, E. M. Ostap, Measuring the kinetic and mechanical properties of non-processive myosins using optical tweezers. Methods Mol. Biol. 1486 , 483–509 (2017).27844441
75 D. M. Rutkowski, D. Vavylonis, Discrete mechanical model of lamellipodial actin network implements molecular clutch mechanism and generates arcs and microspikes. PLOS Comput. Biol. 17 , e1009506 (2021).34662335
76 A. E. Carlsson, Growth of branched actin networks against obstacles. Biophys. J. 81 , 1907–1923 (2001).11566765
77 J. B. Alberts, G. M. Odell, In silico reconstitution of Listeria propulsion exhibits nano-saltation. PLOS Biol. 2 , e412 (2004).15562315
78 D. Holz, D. Vavylonis, Building a dendritic actin filament network branch by branch: Models of filament orientation pattern and force generation in lamellipodia. Biophys. Rev. 10 , 1577–1585 (2018).30421277
79 K. Sekimoto, J. Prost, F. Julicher, H. Boukellal, A. Bernheim-Grosswasser, Role of tensile stress in actin gels and a symmetry-breaking instability. Eur. Phys. J. E Soft Matter 13 , 247–259 (2004).15103519
80 K. John, P. Peyla, K. Kassner, J. Prost, C. Misbah, Nonlinear study of symmetry breaking in actin gels: Implications for cellular motility. Phys. Rev. Lett. 100 , 068101 (2008).18352520
81 F. Gerbal, P. Chaikin, Y. Rabin, J. Prost, An elastic analysis of Listeria monocytogenes propulsion. Biophys. J. 79 , 2259–2275 (2000).11053107
82 J. Zhu, A. Mogilner, Mesoscopic model of actin-based propulsion. PLOS Comput. Biol. 8 , e1002764 (2012).23133366
83 N. J. Burroughs, D. Marenduzzo, Nonequilibrium-driven motion in actin networks: Comet tails and moving beads. Phys. Rev. Lett. 98 , 238302 (2007).17677942
84 K. C. Lee, A. J. Liu, New proposed mechanism of actin-polymerization-driven motility. Biophys. J. 95 , 4529–4539 (2008).18708451
85 T. Kim, W. Hwang, H. Lee, R. D. Kamm, Computational analysis of viscoelastic properties of crosslinked actin networks. PLOS Comput. Biol. 5 , e1000439 (2009).19609348
86 F. Gittes, B. Mickey, J. Nettleton, J. Howard, Flexural rigidity of microtubules and actin filaments measured from thermal fluctuations in shape. J. Cell Biol. 120 , 923–934 (1993).8432732
87 C. S. Peskin, G. M. Odell, G. F. Oster, Cellular motions and thermal fluctuations: The Brownian ratchet. Biophys. J. 65 , 316–324 (1993).8369439
88 A. Mogilner, G. Oster, Cell motility driven by actin polymerization. Biophys. J. 71 , 3030–3045 (1996).8968574
89 Y. Tsuda, H. Yasutake, A. Ishijima, T. Yanagida, Torsional rigidity of single actin filaments and actin-actin bond breaking force under torsion measured directly by in vitro micromanipulation. Proc. Natl. Acad. Sci. U.S.A. 93 , 12937–12942 (1996).8917522
