Vascular disease-causing mutation R258C in ACTA2 disrupts actin dynamics and interaction with myosin

Significance

Point mutations in vascular smooth muscle α-actin are the most prevalent cause of familial thoracic aortic aneurysms leading to acute dissections, yet the molecular mechanism by which these mutations affect actin function is unknown. An underlying cause of the disease is thought to be contractile dysfunction, which initiates adaptive pathways to repair the defects in the smooth muscle cells. Here, we investigate the effects of the R258C mutation, a prevalent mutation in humans with a poor prognosis. The mutant actin shows multiple defects, including impaired interaction with myosin, formation of less stable filaments, and enhanced levels of monomer. These defects are likely to decrease cellular force production and initiate aberrant mechanosensing pathways that culminate in the disease.

Abstract

Point mutations in vascular smooth muscle α-actin (SM α-actin), encoded by the gene ACTA2, are the most prevalent cause of familial thoracic aortic aneurysms and dissections (TAAD). Here, we provide the first molecular characterization, to our knowledge, of the effect of the R258C mutation in SM α-actin, expressed with the baculovirus system. Smooth muscles are unique in that force generation requires both interaction of stable actin filaments with myosin and polymerization of actin in the subcortical region. Both aspects of R258C function therefore need investigation. Total internal reflection fluorescence (TIRF) microscopy was used to quantify the growth of single actin filaments as a function of time. R258C filaments are less stable than WT and more susceptible to severing by cofilin. Smooth muscle tropomyosin offers little protection from cofilin cleavage, unlike its effect on WT actin. Unexpectedly, profilin binds tighter to the R258C monomer, which will increase the pool of globular actin (G-actin). In an in vitro motility assay, smooth muscle myosin moves R258C filaments more slowly than WT, and the slowing is exacerbated by smooth muscle tropomyosin. Under loaded conditions, small ensembles of myosin are unable to produce force on R258C actin-tropomyosin filaments, suggesting that tropomyosin occupies an inhibitory position on actin. Many of the observed defects cannot be explained by a direct interaction with the mutated residue, and thus the mutation allosterically affects multiple regions of the monomer. Our results align with the hypothesis that defective contractile function contributes to the pathogenesis of TAAD.

Thoracic aortic aneurysms and dissections (TAAD) are the 18th most common cause of death in individuals in the United States (1). The high degree of mortality is partly due to the fact that aneurysms tend to be asymptomatic until a life-threatening acute aortic dissection occurs. Familial TAAD is an autosomal dominant disorder with variable penetrance, which is characterized by enlargement or dissection of the thoracic aorta (reviewed in 2). The most prevalent genetic cause of familial TAAD, responsible for ∼15% of all cases, are mutations in vascular smooth muscle α-actin (SM α-actin), encoded by the gene ACTA2. More than 40 mutations in ACTA2 have been identified to date (3–5). Intriguingly, ACTA2 mutations also differentially predispose individuals to occlusive vascular diseases, such as premature coronary artery disease and strokes (6). ACTA2 mutations thus can lead to either dilation of large elastic arteries like the aorta or occlusion of smaller muscular arteries.

SM α-actin is the most abundant protein in vascular smooth muscle cells, constituting ∼40% of the total protein and ∼70% of the total actin, with the rest composed of β- and γ-cytoplasmic actin. Actin is critical for contraction and force production by smooth muscle cells, as well as for their proliferation and migration. Dissected aortas show several characteristic features, namely, loss and disarray of the smooth muscle cells in the medial layer, loss of elastic fibers, and proteoglycan accumulation in the medial space (reviewed in 2). The compromised integrity of the aortic wall allows progression to dissection. In contrast, the vascular pathology in the occluded arteries of patients with ACTA2 mutations is characterized by enhanced numbers of smooth muscle cells.

Little is known about the underlying biochemical mechanisms by which mutations in SM α-actin trigger pathways that ultimately result in aortic cell loss or in the cell proliferation typical of occlusive diseases in small muscular arteries. Here, we present an in vitro characterization of the defects caused by the R258C mutation in SM α-actin. To our knowledge, we are the first to express the human SM α-actin isoform successfully using the baculovirus/insect cell system, which is a critical aspect of this study because the effect of point mutations can vary depending on the isoform in which they are present. We investigated the effect of the R258C mutation first because of its prevalence in patients (6), its relatively poor prognosis (median life expectancy of ∼35 y of age), and high penetrance (5), and because it causes TAAD as well as moyamoya-like disease, an occlusive disease of the cerebral vasculature.

The actin monomer consists of two major domains, with ATP bound in the cleft between them. In the typical view of monomeric globular actin (G-actin), R258 is located in a helix on the backside of subdomain 4 (Fig. 1A). [Note that the R258 mutation corresponds to amino acid R256 in the actin protein, due to posttranslational processing that removes both the N-terminal Met and Cys residues (7)]. In filamentous actin (F-actin), the outer domain consists of subdomains 1 and 2, as well as the inner domain of subdomains 3 and 4. The two helical strands that compose the filament are stabilized by interactions between protomers both within a strand and between strands. R258C lies at the interface between the two strands (Fig. 1 B and C). Upon polymerization, the actin protomer undergoes a conformational change: The two major domains, which are twisted in monomeric G-actin, become flatter with respect to one another in F-actin. Another feature of F-actin is that it can adopt multiple states in which the protomers adopt varied twists and tilts with respect to one another, as well as having different loop conformations (8–11).

Fig. 1.

Location of R258 in monomeric and F-actin. (A) Ribbon representation of one protomer from a model of F-actin obtained from high-resolution electron cryomicroscopy [Protein Data Bank (PDB) ID code 3J8A] (11). The four subdomains of actin are indicated. R258, whose side chain is indicated by a red stick, is located in subdomain 4. Note that R258C is the gene numbering for the mutated residue, and that the residue is amino acid R256 in the processed protein. (B) Ribbon representation of three protomers from a model of F-actin (PDB ID code 3J8A) (11). Three residues (indicated with spheres) are involved in salt bridges: R258 (red) and E197 (blue) in the same monomer and K115 (cyan) from the cross-strand monomer. (C) Close-up view of the three residues forming salt bridges shown in B.

Here, we compare the biochemical properties of expressed WT and R258C SM α-actin, with the goal of understanding how the mutation affects some of the basic functions of actin, because this initial insult ultimately culminates in vascular disease. In smooth muscle cells, force production requires both proper interaction of actin with myosin in the contractile domain and polymerization of actin in the cytoskeletal domain, where it strengthens the membrane for force transmission to the ECM (reviewed in 12). The R258C mutation adversely affects interactions with myosin and weakens filament stability. Our results predict that cells expressing the R258C mutation will have decreased force output and a larger pool of monomeric actin. These dysfunctions could trigger aberrant mechanosensing pathways that culminate in compromised aortic muscle tissue (13). Increasing the monomer/polymer ratio of actin may also have an impact on cellular phenotype, which could have implications for occlusive vascular disease.

Results

Expression of Human Vascular SM α-Actin.

Recombinant human SM α-actin was expressed using the baculovirus/Sf9 insect cell expression system. Several challenges needed to be overcome to achieve this goal: (i) The purified actin could not retain a tag, because both the N and C termini are near the myosin binding site; (ii) contamination by endogenous Sf9 cell actin must be avoided; and (iii) polymerization of actin during expression in the Sf9 cells needed to be prevented because it lowers the protein yield. In addition, expression levels of SM α-actin are considerably lower than what we obtain for β- or γ-cytoplasmic actin. Most of these problems were surmounted by using a method that was designed for expression of toxic actin mutants in Dictyostelium (14). The C terminus of actin was fused to a 43-aa actin-monomer sequestering protein, thymosin-β4, followed by a HIS tag. Binding of thymosin-β4 to the cleft between subdomains 1 and 3 renders the expressed actin monomeric in the Sf9 cell. The HIS tag allows purification of the expressed actin on a nickel-chelate column. The thymosin-β4–HIS tag is then cleaved from the C terminus of actin by proteolytic digestion with chymotrypsin, whose primary cleavage site is after the last native residue of actin, a Phe. Following ion exchange chromatography to remove the thymosin-β4 and any actin from which the tag was not cleaved, pure actin with no nonnative residues at either the N or C terminus was obtained. This strategy worked equally well for WT and the R258C mutant actin.

SDS gels of the purified WT and R258C actins show the purity of the preparations (Fig. 2A). The WT actin is shown before and after removal of the thymosin-β4–His tag (Fig. 2A, lanes 2 and 3). R258C actin is shown after final purification (Fig. 2A, lane 4). The two actins were run on a 1D isoelectric focusing gel (Fig. 2B). As expected, the R258C migrated faster than WT because the mutation replaced a positively charged residue with an uncharged residue. Their relative mobilities were consistent with the calculated isoelectric points of 5.24 for WT and 5.16 for R258C. Average yields of purified actin were ∼0.5 mg of actin for WT and ∼0.4 mg for R258C actin per 10⁹ Sf9 cells (200-mL culture).

Fig. 2.

Gel characterization of the expressed purified actin. (A) SDS gels of purified expressed human SM α-actin (lane 1, molecular mass standards; lane 2, expressed WT actin following HIS-affinity purification and before removal of thymosin-β4; lane 3, final purified WT actin following cleavage of thymosin-β4 with chymotrypsin and ion exchange chromatography; lane 4, final purified R258C actin). (B) One-dimensional isoelectric focusing gels (lane 1, WT actin; lanes 2 and 3, comigration of WT and R258C actin; lane 4, R258C actin). (C) Native gel showing similar interactions of WT and R258C actin with DNase. The faster migrating bands are free actin, and the slower migrating band (arrowhead) is the actin–DNase complex (lane 1, WT actin; lane 2, R258C actin; lanes 3–5, WT actin plus DNase; lanes 6–8, R258C actin plus DNase). Actin is constant at 10 μM, and DNase is at 10, 5, and 2.5 μM in lanes 3–5 and 6–8. In this gel system, the R258C consistently runs with a slightly faster mobility than WT actin.

The WT and mutant SM α-actins were compared with regard to their intrinsic ability to polymerize, their interaction with key actin binding proteins (e.g., smooth muscle tropomyosin, profilin, cofilin), and their ability to be moved by smooth muscle myosin under both unloaded and loaded conditions. The mutation significantly affected most of these properties of actin.

R258C Forms a Less Stable Actin Filament.

Polymerization of individual SM α-actin filaments was visualized as a function of time using total internal reflection fluorescence (TIRF) microscopy [10 mM imidazole (pH 7.5), 50 mM KCl, 4 mM MgCl₂, 1 mM EGTA, 2 mM MgATP at 37 °C]. The actin filament is polarized such that the two ends differ. The association and dissociation of subunits at the barbed end are intrinsically much faster than at the pointed end. Growth from the barbed end is 10- to 20-fold faster than from the pointed end, and thus we measure total growth from both ends. A series of images at various times illustrate growth of individual filaments (Fig. 3A and Movie S1). The filaments were not visibly different by eye, and filament breaking was rarely observed for either WT or R258C actin. Neither showed delayed polymerization, suggesting that nucleation was normal for both of them. To extract kinetic parameters, the observed increase in filament length was plotted as a function of actin concentration (Fig. 3B). The slope of each linear plot defines the polymerization rate. The y-intercept is the rate at which actin subunits disassemble from the filament ends. The x-intercept is the critical concentration (i.e., the concentration of monomeric actin in equilibrium with polymer).

Fig. 3.

Quantification of single actin filament growth observed by TIRF microscopy. (A) Growth of 1.5 μM WT G-actin as a function of time. Each panel is separated by 20 s. A yellow arrowhead follows the growth of one filament with time. (B) Rate of polymerization of WT (blue circles) and R258C (red triangles) as a function of actin concentration. Data were obtained from five experiments using three independent protein preparations each of WT and R258C actin. Values (assembly rate, disassembly rate, and critical concentration) obtained from the fits to a line are provided in Table 1. Error bars are SE.

Average data from five experiments, performed with three independent protein preparations each of WT and R258C actin, are summarized in Table 1. The assembly rate of R258C actin is ∼30% slower than WT (11 vs. 16 subunits per μM⁻¹⋅s⁻¹), whereas the disassembly rate is approximately fourfold faster (2.7 vs. 0.7 subunits per s⁻¹), suggesting that the mutant filaments are less stable than WT filaments. The critical concentration for polymerization of R258C actin is approximately fivefold higher than for WT (238 nM vs. 48 nM for WT), which will increase the G-actin concentration in the cell. The defects in polymerization are consistent with the fact that R258 is involved in salt-bridge interactions that stabilize cross-strand interactions in the filament, based on the position of this residue in the pseudoatomic model of the actin filament (Fig. 1 B and C).

Table 1.

Polymerization rates of WT and R258C actin

Actin	TM	Assembly rate, subunits per ⋅μM−1⋅s−1	Disassembly rate, subunits per s−1	Critical concentration, nM
WT	−	15.9 ± 3.4	0.7 ± 0.6	47.8 ± 44.2
R258C	−	11.2 ± 2.9	2.7 ± 1.0	238.2 ± 49.1
WT*	+	6.9 ± 0.7	0.5 ± 0.9	76
R258C*	+	6.4 ± 0.8	1.5 ± 1.4	229

Data were obtained from five experiments using three independent protein preparations each of WT and R258C actin. Conditions: 10 mM imidazole (pH 7.5), 50 mM KCl, 4 mM MgCl₂, and 1 mM EGTA (37 °C). TM, tropomyosin.

*Data were obtained from one experiment. Errors are SE of the fit.

A human heterozygous for the R258C mutation expresses both WT and mutant actin, and thus we quantified the polymerization of an equimolar ratio of WT and R258C. The rate of elongation of a 50:50 mixture of WT and R258C as a function of actin concentration was intermediate between the values obtained for the two homopolymers, but skewed more toward the rate observed for WT actin (Fig. 4A and Table 2). To fit the data, it is assumed that R258C monomers add onto the filament more slowly only when the filament end adopts a “mutant-like” conformation. This simple model is described in more detail in Materials and Methods and Discussion (Fig. 4B).

Fig. 4.

Copolymerization of equal amounts of WT and R258C actin. (A) Rate of polymerization of an equimolar mixture of WT and R258C actin (black circles). Data for the homopolymers (WT, blue circles; R258C, red triangles) were obtained at the same time. Table 2 tabulates the values obtained from the fits (solid lines). Error bars are SE. (B) Schematic to illustrate polymerization of a 50:50 mixture of WT (white circles) and R258C (red circles) actin. A WT monomer can add onto an existing filament that has either a WT or mutant protomer at the end; likewise, a R258C monomer can add onto an existing filament that has either a WT or mutant protomer at the end. The rate of assembly is slower only if the filament end adopts a mutant-like conformation, which skews the curve toward WT values. Details of the model are described in Materials and Methods. Table 2 compares the calculated and experimental values.

Table 2.

Polymerization rates of mixtures of WT and R258C

Actin	Assembly rate, subunits per μM⋅s	Disassembly rate, subunits per s	Critical concentration, nM
WT	20.4	0.70	34.4
R258C	11.5	1.91	165.9
50% WT/50% R258C	18.2	1.48	81.5
Calculated from model*	18.2	1.31	71.9

Conditions: 10 mM imidazole (pH 7.5), 50 mM KCl, 4 mM MgCl₂, and 1 mM EGTA (37 °C).

^*Details of the model are described in Discussion.

Effect of Tropomyosin on Filament Stability.

Tropomyosin, an α-helical coiled-coil protein that spans seven actin protomers, is a key component of the smooth muscle thin filament, and it also affects the binding of other actin binding proteins. A cosedimentation assay was used to measure the binding of smooth muscle tropomyosin to WT and mutant filaments. The binding constant (K_app) for tropomyosin to the R258C filaments was similar (3.3 × 10⁶ M⁻¹) to the K_app of the WT filaments (2.3 × 10⁶ M⁻¹) (Fig. 5).

Fig. 5.

Affinity of WT and R258C actin for smooth muscle tropomyosin. A sedimentation assay was used to measure the binding affinity of actin for tropomyosin at 200 mM NaCl. Maximal binding was normalized to 1. Data were fit to the Hill equation. The K_app for WT is 2.3 × 10⁶ M⁻¹, and the K_app for R258C is 3.3 × 10⁶ M⁻¹. The Hill coefficient is 1.9 for WT, and the Hill coefficient is 2.3 for R258C. Data obtained from three independent pelleting assays are shown.

When the TIRF polymerization assay was repeated in the presence of smooth muscle tropomyosin, both the assembly and disassembly rates for WT and R258C actin were reduced, but the critical concentration was similar to the critical concentration seen in the absence of tropomyosin (Table 1). This result implies that tropomyosin stabilizes the filament by slowing both the addition to and removal of actin monomers from the filament but has no effect on nucleation because tropomyosin does not bind to actin monomers or small nucleating oligomers. Despite the presence of tropomyosin, the R258C filaments remain less stable than WT.

Profilin Binding Is Strengthened by the R258C Mutation.

One of the ways in which actin polymerization is regulated is via formins, which enhance filament formation. Formins contain two conserved domains: Formin homology domain 1 (FH1) binds profilin, and formin homology domain 2 (FH2) binds actin (reviewed in 15, 16). A TIRF polymerization experiment was performed in the presence of the constitutively active FH1-FH2 fragment of mouse diaphanous 1 (mDia1). The presence of formin had no effect on the rate of polymerization of WT, R258C, R258H, or an equimolar mixture of WT and R258C, but it did help nucleate all of them, based on the observed increase in the number of filaments (Fig. 6A). The mutation thus has little or no effect on the interaction of actin with formin. We compared R258C with R258H in some assays because both mutations lead to the same disease phenotype in humans.

Fig. 6.

Profilin binds more strongly to R258C than to WT actin. (A) Rate of polymerization and total filament number (in a 54 × 54-μm area after 2 min) observed by TIRF microscopy. The polymerization was done with pure actin, actin-formin, actin-formin-profilin, or actin-profilin (WT actin, blue bars; equimolar mix of WT and R258C, gray bars; R258C actin, red bars; R258H, green bars). Error bars are SE. Conditions are 1 μM actin, 50 nM formin [mouse diaphanous 1 (mDia1) FH1-FH2], and 3 μM profilin. (B) Rate of polymerization as a function of actin concentration in the absence or presence of 3 μM profilin (WT, blue circles; R258C, red triangles). Solid lines (no profilin) are linear fits, whereas the dashed lines (plus 3 μM profilin) are the fits to a model described in Materials and Methods. Error bars are SE.

Strikingly, when profilin was added, either alone or in combination with formin, the R258C or R258H actin did not polymerize under our experimental conditions (Fig. 6A). We thus investigated the effect of profilin alone in more detail. The rate of filament growth was quantified as a function of actin concentration in the presence of 3 μM profilin, using the TIRF polymerization assay (Fig. 6B, dashed lines). Profilin decreased the polymerization rate for both WT and R258C actin (from 16 to 3.5 subunits per μM⁻¹⋅s⁻¹ for WT and from 11 to 3.8 subunits per μM⁻¹⋅s⁻¹ for R258C) (Table 3), suggesting that profilin-actin adds more slowly onto filament ends compared with the free actin monomer. The disassembly rate for both WT and R258C was faster in the presence of profilin compared with its absence (increased from 0.7 to 3 subunits per s⁻¹ for WT and from 2.7 to 8.2 subunits per s⁻¹ for R258C) (Table 3), which may be due to the unstable configuration of actin at the end of filament when profilin is still bound. Addition of profilin also increases the critical concentration from ∼50 to 330 nM for WT actin because it binds monomeric actin. Unexpectedly, the critical concentration for polymerization of R258C actin increased to a much greater extent in the presence of 3 μM profilin, from ∼240 to 1,680 nM (Table 3). This increase may have cellular implications because the level of free actin monomer is tightly controlled in vivo. The data were fitted to a simple model (Materials and Methods) in which both free actin monomer and profilin-actin can take part in the polymerization process. The parameters obtained from the fit show that profilin binds to R258C actin with ∼20-fold higher affinity than to WT (Table 3).

Table 3.

Polymerization data in the presence of 3 μM profilin

Fit parameters	WT	R258C
Assembly rate, subunits per μM−1⋅s−1	3.5	3.8
Disassembly rate, subunits per s−1	3.1	8.2
Kd, μM	3.0	0.16
Critical concentration, nM	330	1,680

Conditions: 10 mM imidazole (pH 7.5), 50 mM KCl, 4 mM MgCl₂, and 1 mM EGTA (37 °C).

R258C Filaments Are More Sensitive to the Action of Cofilin.

Cofilin is an important regulator of actin dynamics. It has a filament severing activity that increases the number of filament ends, and it depolymerizes filaments from their minus, pointed ends (reviewed in 17). We observed both types of behavior in a TIRF polymerization assay (Fig. 7 and Movie S2): Filaments were severed internally, and they also shortened from either end, with one end much faster than the other, until the filament disappeared. These two behaviors were not mutually exclusive; namely, a given filament could exhibit one or both behaviors. Both processes depended on cofilin concentration and resulted in the destruction of actin filaments.

Fig. 7.

R258C filaments are more susceptible to cofilin-induced shortening and cleavage than WT actin, both in the presence and absence of smooth muscle tropomyosin (TM). (A) Rate of actin filament shortening induced by cofilin (WT, blue bars; R258C, red bars). Lighter shaded bars are in the presence of tropomyosin. Error bars are SE. (B) Frequency of cleavage by cofilin for WT and R258C actin filaments. Frequency is reported as the number of events per 1 min per 1 μm of actin filament. Error bars are SE.

We first assessed the effect of cofilin on actin filament shortening. As little as 10 nM cofilin induced shortening of R258C filaments (Fig. 7A). At 100 nM cofilin, the process was so fast that no R258C filaments were observed by the time the slide was mounted on the microscope. In contrast, WT filaments only started to slowly shorten at 100 nM cofilin.

WT filaments were also more resistant than R258C filaments to severing by cofilin (Fig. 7B). WT filaments showed virtually no severing at 100 nM cofilin, whereas with 75 nM cofilin, severing events were frequently observed with R258C filaments.

Binding of cofilin is competitive with tropomyosin (reviewed in 17). Tropomyosin reduced the rate of shortening and strongly suppressed the severing effects of cofilin with WT actin filaments (Fig. 7). Even at 1.2 μM cofilin, there was minimal shortening and almost no severing. In contrast, the R258C filaments were much more vulnerable to attack by cofilin despite the presence of tropomyosin. Smooth muscle tropomyosin binds at least as tightly to R258C filaments as to WT filaments (Fig. 5), so low binding affinity does not explain the result.

DNase and Gelsolin Binding Are Unaffected.

To assess if the R258C mutation affects its binding to DNase and gelsolin segment-1 qualitatively, actin was incubated with varying molar ratios of actin to binding protein, and actin complexes were separated from monomers on native gels. Fig. 2C shows the results with DNase, but essentially identical gels were obtained with gelsolin segment-1. In both cases, there was no obvious difference in the amount of complex formed with WT and the R258C actin.

R258C Actin Filaments Are Propelled at Slower Speeds than WT.

An unloaded in vitro motility assay was used to measure the speed at which smooth muscle myosin moves WT vs. R258C actin. We first performed the typical motility assay using filaments that were stabilized and visualized using rhodamine-phalloidin. Mutant and WT filaments were moved at the same speed (Fig. 8A). In the presence of smooth muscle tropomyosin, WT actin moved ∼40% faster than bare actin. In contrast, the speed at which myosin moved the R258C filaments was not enhanced by tropomyosin. This increase in speed resulted in WT filaments moving ∼1.7-fold faster than R258C filaments in the presence of tropomyosin.

Fig. 8.

Smooth muscle myosin moves R258C filaments more slowly in an in vitro motility assay. (A) Actin filaments were stabilized and visualized with rhodamine-phalloidin. Speeds were determined by a semiautomated tracking program and fitted to a Gaussian distribution. (Upper, minus tropomyosin) Myosin moved WT actin at a speed of 0.53 ± 0.18 μm⋅s⁻¹ (n = 2,742) and R258C filaments at a speed of 0.50 ± 0.16 μm⋅s⁻¹ (n = 3,415). The difference in speeds was statistically significant (Student’s t test, P < 0.01) because of the large number of filaments analyzed, but it has no physiological relevance. (Lower, plus tropomyosin) In the presence of smooth muscle tropomyosin, myosin moved WT actin at a speed of 0.74 ± 0.19 μm⋅s⁻¹ (n = 1,294) and R258C filaments at the slower speed of 0.43 ± 0.15 μm⋅s⁻¹ (n = 3,743) (Student’s t test, P < 0.01). Data from five independent actin preparations were pooled. (B) Speed of movement of WT and R258C actin filaments that were not stabilized with phalloidin. Filaments were visualized by incorporation of 25% WT actin that was directly labeled with rhodamine (solid blue bar, 100% WT; striped red bar, 50% WT and 50% R258C; solid red bar, 25% WT and 75% R258C). (Left) Speed of movement of WT filaments (0.61 ± 0.13 μm⋅s⁻¹; n = 196) decreased to 0.53 ± 0.12 μm⋅s⁻¹ (n = 160) with 50% R258C and to 0.47 ± 0.13 μm⋅s⁻¹ (n = 180) with 75% R258C. The difference between all pairs was statistically significant (Student’s t test, P < 0.01). (Right) Same comparison in the presence of smooth muscle tropomyosin (TM). The speed of movement of WT filaments (0.60 ± 0.10 μm⋅s⁻¹; n = 170) decreased to 0.43 ± 0.09 μm⋅s⁻¹ (n = 80) with 50% R258C and to 0.31 ± 0.081 μm⋅s⁻¹ (n = 142) with 75% R258C.The difference between all pairs was statistically significant (Student’s t test, P < 0.01). Data from five independent actin preparations were pooled. Filament speed was tracked manually. Temperature, 30 °C.

The in vitro motility was repeated in the absence of phalloidin to test if this compound, which stabilizes the filament, moderates the defects caused by the mutation. To visualize the filaments, WT actin was directly labeled with rhodamine and incorporated into the filament as 25% of the total actin. The speed of filament movement slowed as the percentage of R258C actin in the filament increased from 50–75% (Fig. 8B). The slowing of speed by the mutant actin was accentuated in the presence of tropomyosin.

In some patients, Arg-258 is mutated to His instead of Cys. We expressed and purified R258H to test if this mutation had similar effects to R258C. In filaments containing 75% mutant actin, speeds were comparable whether the Arg residue was mutated to Cys or His (minus tropomyosin: 0.47 ± 0.13 for R258C vs. 0.50 ± 0.10 for R258H; plus tropomyosin: 0.31 ± 0.08 for R258C vs. 0.32 ± 0.09 for R258H).

To determine if the slowing of speed with the mutant actin filament was due to a change in the rate of release of ADP from myosin, a stopped-flow experiment was performed. Acto-smooth heavy meromyosin (HMM)-ADP was mixed with 2 mM MgATP. Under these conditions, the rate of acto-HMM dissociation is rate-limited by ADP dissociation. The rate of ADP release from the mutant and WT filament was the same (134 ± 1 s⁻¹ for WT actin and 127 ± 1 s⁻¹ for R258C actin).

Actomyosin Force Generation Is Severely Impaired When Tropomyosin Is Bound to the R258C Filament.

To test if the R258C mutation affects force generation by myosin, we carried out a one-bead laser trap experiment to investigate the force/velocity relationship of a small ensemble of smooth muscle heavy meromyosin with WT and R258C actin filaments (Fig. 9A). When studying actin mutants, an advantage of the one-bead setup is that the actin filaments do not require phalloidin for stabilization because they are attached to the coverslip. In the more traditional three-bead assay, the actin is suspended between two beads and stabilized with rhodamine-phalloidin.

Fig. 9.

Force dependence of smooth muscle myosin HMM interacting with WT or R258C filaments in the presence or absence of smooth muscle tropomyosin. (A) Schematic of experimental setup. Smooth muscle myosin HMM, with a biotin tag at the C terminus, was attached to 1-μm polystyrene beads coated with neutravidin. A myosin-coated bead was then captured by the laser trap and allowed to interact with actin immobilized on the surface. A force-feedback mechanism was used to keep force constant once an interaction was detected. QD, quadrant detector. (B) Sample trace showing the bead motion under different constant loads, varying from 1 to 5 pN. The solid yellow lines are linear fits to the raw data, shown in black. (C) Force-velocity curves for HMM interacting with WT (blue) or R258C (red) filaments in the presence (dashed lines, open symbols) or absence (solid lines, filled symbols) of tropomyosin. The lines are fit to the Hill equation. Error bars are SE.

We first compared the force/velocity relationship obtained with bare WT and R258C actin filaments. A force-feedback system is used to clamp the force at five levels between 1 and 5 pN. The velocity at any force level was obtained by fitting the raw data to a line (Fig. 9B). The force/velocity relationship was then fitted to the Hill equation. As expected, velocity slowed as force increased (Fig. 9C). There was virtually no difference between the curves obtained with bare WT vs. bare R258C filaments. In the presence of smooth muscle tropomyosin, the WT filament showed the same force/velocity relationship as in its absence. In contrast, velocity was greatly reduced at all force levels for R258C actin-tropomyosin filaments.

Discussion

Multiple facets of SM α-actin function are quite severely affected by the R258C mutation. Defects induced by the R258C mutation include (i) slower and less productive interactions with smooth muscle myosin, particularly in the presence of tropomyosin, that lead to reduced force output; (ii) formation of a less stable filament with greater susceptibility to cleavage by cofilin even in the presence of tropomyosin, which enhances actin filament dynamics; and (iii) higher affinity binding to profilin, which increases the monomeric pool of actin. How these in vitro defects affect smooth muscle cell function must be viewed in light of the many roles that SM α-actin plays in vascular smooth muscle cells. These roles include production of contractile force, maintenance of the integrity of the submembrane cytoskeleton, cellular mechanosensing, and regulation of cell differentiation and proliferation. Genetic mutations that predispose patients to TAAD include mutations in the contractile proteins and their regulators, ECM structural components of the aortic wall, and proteins involved in mechanosensing. It has been proposed that smooth muscle cell contractile dysfunction is one of the primary insults that activates the stress and strain pathways that initiate maladaptive remodeling of the smooth muscle cell, ultimately culminating in an aortic aneurysm (reviewed in 2, 13).

Actin is found in two distinct domains in smooth muscle cells. The “contractile” domain of smooth muscle cells consists of interdigitating myosin and stable actin filaments that cyclically interact to generate tension, controlled by phosphorylation of the regulatory light chain of smooth muscle myosin. These filaments are anchored into dense bodies, a structure characteristic of smooth muscle cells. A second “cytoplasmic” domain located in the subcortical region has a more dynamic pool of actin that is not associated with myosin. At the membranous dense plaques, this pool of actin attaches to the cytoplasmic tails of transmembrane integrins via focal adhesion proteins, which transduce mechanical force across the plasma membrane by linking the ECM with the cytoskeleton.

A unique feature of tension development in smooth muscle cells is that the independently regulated contractile domain and cytoskeletal domains need to collaborate to produce force. This view is supported by numerous studies showing that in response to stimulation, the amount of F-actin in the cell increases, and if this new actin polymerization is blocked, tension is not developed (reviewed in 12, 18–20). The mechanism by which actin polymerization facilitates force production may include an enhancement of the linkage of the actin filaments to integrins, which would strengthen force transduction between the contractile domain and the ECM. Approximately 10% of the actin undergoes polymerization, so that a resting cell has ∼70–80% F-actin, which increases to ∼80–90% after stimulation. Whether both smooth and cytoskeletal actin isoforms undergo polymerization upon stimulation remains unresolved (21, 22). Moreover, when the cell contains a mutant SM α-actin, such as R258C, which appears to be more dynamic than WT α-actin, it may more readily become part of both the contractile and cytoskeletal pools of actin. Our results suggest that the R258C mutation has the potential to disrupt the function of actin in both domains.

R258C Mutation Decreases Contractile Function by Perturbing Interactions with Both Myosin and Tropomyosin.

Under unloaded conditions, the presence of R258C in the actin filament slowed the speed of myosin-powered movement in an in vitro motility assay in a dose-dependent fashion. Phalloidin stabilization of the filament suppresses this defect. Two general mechanisms can lead to slower movement. One is a change in myosin kinetics, and the other is via a change in the properties of the actin filament per se. The kinetic step in myosin that controls actin motility speed is ADP release. Because we observed no change in the rate of ADP release, we favor a mechanism whereby the structure of the mutant filament does not allow the full displacement per working stroke of myosin to be realized, resulting in less displacement per MgATP hydrolyzed, and thus a slower speed.

Tropomyosin occupies a more inhibitory position on R258C actin filaments. In the absence of myosin and troponin, the latter of which is not present in smooth muscle cells, all tropomyosins occupy either the blocked or closed state on actin (23). In the blocked state, tropomyosin sterically prevents myosin from binding to actin. In the closed state, myosin can bind weakly, and additional binding of myosin cooperatively shifts tropomyosin to the open state, which fully exposes the myosin binding site. In the presence of tropomyosin, the difference in speed between WT and mutant filaments was accentuated. More strikingly, once load was applied, myosin was essentially unable to interact with R258C actin-tropomyosin filaments. In the loaded assay, only a small number of myosin heads interact with the actin-tropomyosin filament at any time, and this number appears to be insufficient to displace tropomyosin from an inhibitory position.

That defects in force output from the contractile domain can lead to TAAD is further supported by the fact that loss-of-function mutations in both the myosin heavy chain (MYH11) and myosin light chain kinase (MLCK), which phosphorylates the regulatory light chain of myosin and is required for actomyosin interactions, also cause autosomal dominant inheritance of TAAD (24, 25). Genes encoding proteins involved in either smooth muscle cell contraction or the TGF-β signaling pathway are the predominant causes of familial TAAD.

R258C Destabilizes the Filament.

Although R258C was not specifically looked at, immunofluorescence of cultured smooth muscle cells from individuals with ACTA2 mutations showed substantially fewer actin filaments than controls, suggesting that these mutations may interfere with actin assembly (3). Our in vitro results support this idea. R258C actin has a higher critical concentration for assembly, caused by a slower rate of assembly and a faster rate of disassembly. Based on the location of R258 in the model of the actin filament structure (Fig. 1), a polymerization defect was predictable. Recent advances in electron cryomicroscopy have led to the highest resolution structures (3.7–4.7 Å) of the actin filament to date (9, 11). In these structures, R258 (R256 in protein) forms a salt bridge with E197 (E195 in protein) in the same protomer, which, in turn, forms a salt bridge with K115 (K113 in protein) on the cross-strand actin (Fig. 1 B and C). This stabilizing salt bridge triad will be disrupted upon mutation of R258 to either Cys or His, which changes both side-chain size and charge. Mutation of R258 to either Cys or His causes similar patient phenotypes. Filament destabilization as a result of an R256H mutation (equivalent to residue Arg-258 in ACTA2) was first observed in the backbone of Saccharomyces cerevisiae actin by Malloy et al. (26). Ionic cross-strand stabilization that involves this residue thus appears to be a property common to SM α-actin and budding yeast actin.

Copolymerization of equal amounts of WT and mutant actin tempers the polymerization defect. The observation that polymerization as a function of actin concentration was skewed toward the WT values could be explained by a simple model if we make the following two assumptions (Fig. 4 and Materials and Methods). First, only when an R258C monomer adds onto a filament that adopts a mutant-like conformation (e.g., a filament with a mutant protomer at the barbed end) will it assemble at a slow rate similar to the rate observed for the R258C homopolymer. In all other scenarios, the monomer adds onto the filament with a faster rate that approximates the WT assembly rate. Second, the disassembly rate depends solely on the actin type (i.e., WT dissociates with the WT disassembly rate, R258C dissociates from filaments with the R258C disassembly rate). As shown in Table 2, the calculated results from this model agree with the experimentally observed values.

Actin Filament Dynamics Regulated by Binding Proteins.

In nonmuscle cells, the actin cytoskeleton is highly dynamic, and controlled assembly and disassembly of actin play roles both in cell migration and plasma membrane invagination during endocytosis and phagocytosis. Assembly is regulated by proteins, including formin and Arp2/3, that control nucleation and polymerization (reviewed in 15), and it is balanced by disassembly of older filaments by proteins in the ADF/cofilin family, which preferentially bind ADP-actin monomers (reviewed in 17). Cofilin accelerates remodeling of the actin network by severing actin filaments, and thus increasing the concentration of ends available for elongation and subunit exchange. Cofilin also depolymerizes filaments from the pointed ends, thereby increasing filament dynamics (27).

Cofilin is, however, also present in striated muscle cells that are considered to have considerably more stable actin filaments. In striated muscle cells, cofilin is thought to be involved in precise length control of the thin filament (28, 29). In vascular smooth muscle cells, increased cofilin activation upon dephosphorylation with slingshot phosphatase 1 has been implicated in vascular smooth muscle cell migration and neointima formation following vascular injury (30). Here, we show that R258C actin filaments are considerably more susceptible to cleavage by cofilin and show increased levels of shortening, compared with WT filaments. In essence, the increased sensitivity of R258C to cofilin should have the same effect as having more activated, unphosphorylated cofilin. Our results suggest that mutation of R258 is likely to result in enhanced actin dynamics and cell migration.

Although the molecular mechanism of severing is not known, it is affected by the structure and mechanical properties of the actin filament (31), which differ when R258 is mutated. Moreover, although binding of smooth muscle tropomyosin protects the WT filament from cofilin cleavage (32), the same is not true for the R258C filaments. This observation supports the idea that the position of tropomyosin on WT vs. R258C filaments differs, consistent with the inhibitory effect of tropomyosin on force and motion generation by myosin when it interacts with the mutant actin.

R258C Increases the Monomeric Pool of Actin by Binding Profilin Tightly.

Profilin, one of the most abundant actin binding proteins inside the cell, binds to monomeric actin with high affinity and is the predominant substrate for actin assembly in vivo. Formins accelerate polymerization by increasing the local concentration of actin-profilin at the barbed end (reviewed in 15, 16). Here, we show that the affinity of R258C actin for profilin is ∼20-fold higher than to WT. This tight binding decreases formin-mediated actin polymerization. The net result is that in cells containing R258C actin, we predict that the pool of monomeric actin is considerably larger than in WT cells. Another prediction is that more WT monomers will be incorporated into the actin filaments than the mutant in cells expressing both, potentially attenuating the detrimental effect of the mutant actin. The tighter binding to profilin is unexpected because the profilin binding site to actin is far from the location of the R258C mutation. This tight binding indicates the R258C mutation changes not only the actin filament structure but also the conformation of monomeric actin.

A previous study investigated the effect of mutating this same Arg residue to His in the backbone of budding yeast actin, which is only 86% identical to SM α-actin (26). The major defect those investigators observed in vitro was that yeast formin (Bni1) did not enhance polymerization but, instead, strongly capped the filament end. This result was consistent with their cellular results in which budding yeast cells expressing the mutant actin were delayed in their ability to rebuild their actin cytoskeleton following a challenge by the actin-depolymerizing drug latrunculin A. Although we also observed intrinsic polymerization defects with this mutation, and altered interaction of R258C with profilin and cofilin, our results did not show a defect in the mutant actin’s interaction with formin for either R258H or R258C. We conclude that the effect of vascular disease-causing mutations is best studied in the SM α-actin backbone rather than a heterologous system.

A potential downstream effect of increasing the amount of monomeric actin in the cell is altering smooth muscle cell phenotype. The subcellular localization of the actin binding transcriptional cofactor myocardin-related transcription factor (MRTF-A) depends on the relative amount of monomeric vs. F-actin. When G-actin concentration in the cytoplasm increases, it binds to MRTF-A and retains it in the cytoplasm. Conversely, upon Rho-A–dependent actin polymerization, MRTF-A accumulates in the nucleus. Once in the nucleus, it interacts with serum response factor, which binds to conserved CC[A/T]₆GG cis-elements within the smooth muscle cell-specific promoters to activate transcription of smooth muscle-specific genes (reviewed in 33, 34). Thus, a potential outcome is that cells containing R258C actin may lead to the smooth muscle cell developing a less contractile and more proliferative phenotype, because the increased monomeric pool may sequester MRTF-A in the cytoplasm.

Allosteric Effects of the R258C Mutation.

A number of the observed defects in R258C were unexpected based solely on its location in actin. These defects include altered binding of smooth muscle tropomyosin and tighter binding of the mutant actin to profilin. The mutant residue is not directly part of the myosin, tropomyosin, or profilin binding site. These latter unexpected changes suggest new pathways of allosteric communication within the actin monomer that have been disturbed by the mutation. An allosteric connection between the profilin binding site and R258 is consistent with results from a hydrogen/deuterium exchange and MS study showing that Arg-258 and residues at the profilin binding site were on a connected block of residues that display similar hydrogen/deuterium exchange kinetics (35). Reciprocally, a protein binding at the profilin site at the barbed end can propagate a conformation change that locally alters the conformation of residue 258 at an interstrand filament stabilization site, which may provide insight into how profilin at low concentrations can facilitate actin polymerization.

A prevailing argument for the extreme conservation of sequence in actin is that virtually every residue has been placed under selective pressure because of the internal networks within the actin filament that are needed to maintain such allosteric linkages (reviewed in 36). Our results provide additional evidence to support this point of view.

Perspective.

Fig. 10 shows a schematic of potential differences between a WT smooth muscle cell and a cell expressing both WT and R258C actin. The overall goal of characterizing the functional effect of actin point mutations at the molecular level is to determine if the observed defects can be correlated with vascular disease outcomes and with patient survival rates. This study is the first step toward that goal. For example, both R258C and R39H lead to TAAD and moyamoya-like cerebrovascular disease, whereas other mutations lead only to TAAD (e.g., W88R, G160D) and still others lead to TAAD and coronary artery disease (e.g., R149C, R118Q). “Loss-of-function” mutations that lead to smooth muscle cell weakness, a failure of tension sensing, and degeneration are thought to lead to thoracic aneurysms and aortic dissections. With R258C, we have evidence for loss of contractile function. In contrast, “gain-of-function” mutations that lead to cell proliferation and increased motility, or hyperplastic arterial smooth muscle cell growth, might lead to occlusive vascular diseases. With R258C, pathways that lead to cell proliferation could be mediated via the observed altered interactions of R258C filaments with cofilin and profilin, leading to increased pools of monomeric actin, and by the intrinsic lability of the mutant filament. The R258C mutation affects essentially all aspects of actin function that we measured in vitro. Cellular studies with fibroblasts and smooth muscle cells derived from patients with the R258C mutation, or mouse models, will establish how these defects are manifested in a cellular context.

Fig. 10.

Summary of likely differences in cells containing only WT actin vs. cells expressing both WT and R258C actin. The left-hand portion of the schematic cell illustrates that myosin interacts productively with WT actin-tropomyosin to produce force and motion. Most actin is filamentous, with only a small pool of monomeric actin. The filaments that are decorated with tropomyosin are resistant to severing and shortening by cofilin. In contrast, cells expressing both WT and R258C actin would be expected to show slower filament sliding and have a lower force output. Moreover, it is expected that the pool of monomeric actin will increase relative to WT cells, because R258 has a higher critical concentration, a higher affinity for profilin, and is more susceptible to severing and shortening by cofilin.

Materials and Methods

Details of cloning, expression, and purification of proteins used in this study; the tropomyosin binding assay; actin labeling and filament formation for motility; the in vitro motility assay; measurement of ADP dissociation rates; and force data acquisition and analysis are provided in SI Materials and Methods.

Actin Cloning, Expression, and Purification.

A chimeric construct composed of human SM α-actin (ACTA2 gene; accession number NP_001604) and human thymosin-β4 (accession number AAI41977.1) was cloned into pAcUW2B for expression in the baculovirus/insect cell system. The initial Cys residue of actin was not included in the construct because it is not present in the mature, processed protein. The thymosin-β4 gene was separated from the actin gene by a 14-aa linker (ASSGGSGSGGSGGA) to allow the thymosin-β4 enough flexibility to occupy the barbed end binding site on the actin monomer, thus preventing the actin from polymerizing with itself or native Sf9 cell actin. A HIS₆ tag was added at the C terminus for purification on a nickel affinity column. The final native residue of actin is Phe, which provides a site for chymotrypsin cleavage to remove the linker, thymosin-β4, and the HIS₆ tag completely. The native codons for both actin and thymosin-β4 were replaced with Drosophila preferred codons.

For expression of human SM α-actin, infected Sf9 cells (4 × 10⁹) were harvested 3 d after infection and lysed in 160 mL of 1 M Tris⋅HCl (pH 7.5 at 4 °C), 0.6 M NaCl, 0.5 mM MgCl₂, 0.5 mM Na₂ATP, 1 mM DTT, 4% (vol/vol) Triton X-100, 1 mg/mL Tween-20, and protease inhibitors [0.5 mM 4-(2-aminoethyl)benzenesulfonyl fluoride hydrochloride, 5 μg/mL leupeptin, 0.5 mM tosyllysine chloromethyl ketone hydrochloride]. The cells were stirred for 2 h and then clarified. The soluble fraction was dialyzed overnight against 10 mM Hepes (pH 7.5), 0.25 mM CaCl₂, 0.3 M NaCl, 7 mM β-mercaptoethanol, 0.25 mM Na₂ATP, and 1 μg/mL leupeptin. Following dialysis, the supernatant was applied to 5–10 mL of a nickel affinity column (HIS-Select; Sigma–Aldrich). Nonspecifically bound contaminating proteins were washed off with dialysis buffer containing 10 mM imidazole (pH 7.5). Actin was eluted with 100 mM imidazole, 10 mM Hepes (pH 7.5), 0.25 mM CaCl_2, 0.3 M NaCl, 7 mM β-mercaptoethanol, 0.25 mM Na₂ATP, and 1 μg/mL leupeptin. The peak fractions were pooled and dialyzed overnight against G-buffer [5 mM Tris (pH 8.26 at 4 °C), 0.2 mM CaCl₂, 0.1 mM NaN₃, 0.5 mM DTT, 0.2 mM Na₂ATP, 1 μg/mL leupeptin]. The protein was clarified by centrifugation, and the thymosin-HIS tag was cleaved from the actin with chymotrypsin (chymotrypsin/actin in a 1:80 weight ratio). The actin was separated from the thymosin-His tag using a Mono Q 5/50 GL column (GE Healthcare) with a gradient of 0–0.3 M NaCl in 5 mM Tris (pH 8.26 at 4 °C), 0.2 mM CaCl₂, 0.1 mM NaN₃, 0.5 mM DTT, 0.2 mM Na₂ATP, and 1 μg/mL leupeptin, followed by a step to 0.5 M NaCl. Peak fractions were pooled, concentrated, and dialyzed against 5 mM Tris (pH 8.26 at 4 °C), 0.2 mM CaCl₂, 0.1 mM NaN₃, 0.5 mM DTT, 0.2 mM Na₂ATP (pH 7), and 1 μg/mL leupeptin.

Actin Polymerization Visualized by TIRF Microscopy.

The 2× polymerization buffer consists of 20 mM imidazole (pH 7.5), 100 mM KCl, 8 mM MgCl₂, 2 mM EGTA, 0.5% methylcellulose, 4 mM MgATP, 0.26 mg/mL glucose oxidase, 0.10 mg/mL catalase, 6 mg/mL glucose, and 20 mM DTT. The rinse buffer is composed of equal volumes of G-buffer [5 mM Tris (pH 8.26 at 4 °C), 0.2 mM CaCl₂, 0.2mM Na₂ATP, 0.5 mM DTT] and 2× polymerization buffer.

Lifeact-GFP was used to visualize the actin filaments. Lifeact binds weakly to F-actin (K_d of ∼2 μM), and as much as 55 μM Lifeact was shown not to affect the rates of polymerization and depolymerization (37). This probe allows us to use unlabeled WT and mutant actin. Previous methods for visualizing actin polymerization have relied on actin that has been modified either at Cys374 or at Lys residues (38, 39). In our study, direct modification of actin is complicated both by the low yields of expressed actin and by our observation that R258C actin does not label in a manner identical to the WT actin. Actin in G-buffer was spun for 30 min at 392,148 × g. The supernatant concentration was determined by a Bio-Rad Protein Assay. The typical actin concentration is 1–2 mg/mL. To prepare the flow cell, 20 μL of 5 μg/mL N-ethyl maleimide-myosin was flowed in and incubated for 2 min. The flow cell was rinsed with 20 μL of 5 mg/mL casein, and then with 20 μL of rinse buffer. Twenty microliters of G-actin (1–5 μM, also containing 1.64 μM Lifeact) in G-buffer was mixed with 20 μL of 2× polymerization buffer to start the polymerization. The mixture was then flowed into the flow cell (two times, 20 μL each time). Excess solution was removed, and the flow cell was sealed with nail polish. The flow cell was immediately put on a Nikon ECLIPSE Ti microscope equipped with through-objective type TIRF and a temperature control unit (37 °C). The samples were excited with the TIRF field of a 488-nm laser line. The fluorescence image was observed with a 100× objective and recorded on an Andor EMCCD camera (Andor Technology USA) at a rate of one frame per second for 1–3 min with automatic focus correction. The final resolution is 0.1066 μm per pixel.

For experiments with profilin, 1 μL of 120 μM profilin (3 μM final) was added to the final 40-μL mixture. For experiments with tropomyosin, various amounts of tropomyosin were added to the mixture so that the final tropomyosin concentration was one-fourth the concentration of the actin monomer. For experiments with cofilin, actin was first polymerized at 37 °C for a few minutes in the flow cell while being monitored on the microscope. Once the filaments reached the desired length and density, 20 μL of G-buffer containing various amounts of cofilin and 1.64 μM Lifeact was mixed with 20 μL of 2× polymerization buffer and then flowed into the flow cell. The flow cell was then sealed with nail polish before observation on the microscope.

Fluorescence Data Processing.

Movies following actin polymerization or depolymerization as a function of time were imported into ImageJ (National Institutes of Health). The growth or shrinkage was followed manually. The polymerization rate was obtained by dividing the total growth by time. Rate was converted from micrometers per second to subunits per second by multiplying by the converting factor of 370 actin subunits per micrometer. For each condition, 15–50 growing filaments were measured.

Rates of polymerization were plotted against the actin monomer concentration. The data were linearly fit to Rate = a[actin] + d, in which a is the second-order assembly rate of polymerization, d is the first-order rate of disassembly, and the critical concentration is −d/a.

To determine the frequency of severing for experiments with cofilin, the total number of severing events (actin filaments that clearly broke into two pieces) was counted for each 1 min of video and then divided by the total actin filament length in the field of view at the starting frame of this 1-min video. Shortening of actin by cofilin was traced manually and then divided by time.

Modeling of Polymerization Data in the Presence of Profilin.

The following assumptions were made to model the polymerization of G-actin in the presence of profilin. Monomeric G-actin and profilin form a profilin–G-actin complex with affinity K_d. Both G-actin and profilin–G-actin can participate in the polymerization process, with rates of r_a and r_ap, respectively. Protomers dissociate from F-actin with rate r_d. The overall scheme is described in the following equations:

$$profilin\cdot actin\stackrel{Kd}{\iff }actin+profilin$$

$$f-actin\left(n\right)+actin\hspace{0.17em}\stackrel{r_a}{\to }f-actin\left(n+1\right)$$

$$f-actin\left(n\right)+profilin\cdot actin\hspace{0.17em}\stackrel{r_{ap}}{\to }f-actin\left(n+1\right)$$

$$f-actin\left(n\right)\hspace{0.17em}\stackrel{r_d}{\to }f-actin\left(n-1\right)$$

Modeling of Polymerization of Mixtures of WT and R258C.

In this scheme, as illustrated in Fig. 4B, the rate of association of G-actin to the end of an F-actin filament, and the dissociation of G-actin from F-actin, depends on the composition of the barbed end of the F-actin polymer. For polymerization, if the last actin in F-actin is WT, then both the WT and R258C G-actin monomers add to the F-actin with a rate of r_w (WT association rate). If the end unit of F-actin is R258C, then a WT G-actin will add onto it with a rate of r_w, but an R258C monomer will add to it with a rate of r_m (R258C mutant–mutant association rate). The following four equations describe the association reaction:

$$f-actin\left(n\right)\cdot WT+WT\hspace{0.17em}\stackrel{r_w}{\to }f-actin\left(n+1\right)$$

$$f-actin\left(n\right)\cdot R258C+WT\hspace{0.17em}\stackrel{r_w}{\to }f-actin\left(n+1\right)$$

$$f-actin\left(n\right)\cdot WT+R258C\hspace{0.17em}\stackrel{r_w}{\to }f-actin\left(n+1\right)$$

$$f-actin\left(n\right)\cdot R258C+R258C\hspace{0.17em}\stackrel{r_m}{\to }f-actin\left(n+1\right)$$

For depolymerization, if the end protomer in F-actin is WT, it dissociates with a rate of r_dw (WT dissociation rate), but if it is R258C, it dissociates with a rate of r_dm (R258C mutant dissociation rate). The following two equations describe the dissociation reaction:

$$f-actin\left(n\right)\cdot WT\hspace{0.17em}\stackrel{r_{dw}}{\to }f-actin\left(n-1\right)$$

$$f-actin\left(n\right)\cdot R258C\hspace{0.17em}\stackrel{r_{dm}}{\to }f-actin\left(n-1\right)$$

SI Materials and Methods

Cloning, Expression, and Purification of Other Proteins.

Lifeact-GFP.

Lifefact-GFP, the first 17 amino acids of actin binding protein 140 from S. cerevisiae (Abp140) conjugated to GFP, was used to visualize actin polymerization (37). A PCR product made using Lifeact pEGFP-N1 (a gift from Roland Wedlich-Söldner, Max Planck Institute, Martinsried, Germany) as a template was cloned into the pET3 vector (Novagen) with a C-terminal HIS₆ tag for purification. Protein was expressed in BLR(DE3) cells that were induced with 0.4 mM isopropyl-β-d-thiogalactopyranoside (IPTG) and grown overnight at 27 °C in LB broth. The pellet was sonicated in 10 mM sodium phosphate (pH 7.4), 0.3 M NaCl, 0.5% glycerol, 7% sucrose, 7 mM β-mercaptoethanol, 0.5 mM 4-(2-aminoethyl)benzenesulfonyl fluoride hydrochloride (AEBSF), and 5 μg/mL leupeptin. The cell lysate was clarified at 200,000 × g for 30 min, and the supernatant was applied to a HIS-Select nickel affinity column (Sigma–Aldrich). The resin was washed with 10 mM sodium phosphate, 10 mM imidazole (pH 7.4), 0.3 M NaCl, 0.5 mM AEBSF, 5 μg/mL leupeptin, and 5 mM benzamidine, and then with the same buffer containing 30 mM imidazole. Lifeact-GFP was eluted from the column with 10 mM sodium phosphate, 200 mM imidazole (pH 7.5), 0.3 M NaCl, and 1 μg/mL leupeptin. Purified protein was dialyzed in 10 mM imidazole (pH 7.4), 0.2 M NaCl, 1 mM DTT, and 50% glycerol, and stored at −20 °C.

Cofilin.

A PCR product of human cofilin was obtained using American Type Culture Collection clone MGC-15952 as a template and cloned into pET3a. Protein was expressed in BLR(DE3) cells that were induced with 0.4 mM IPTG and grown overnight at 27 °C. Purification was as described by Pope et al. (40).

Profilin.

A plasmid-containing human profilin pBG942 (a gift from Bruce Goode, Brandeis University, Waltham MA) was transformed into Escherichia coli strain BL21(DE3) and cultured in LB. The purification was according to the protocol of Lu and Pollard (41). Briefly, the bacteria pellet was lysed by sonication in 25 mM Tris⋅Cl (pH 7.5 at 4 °C), 50 mM sucrose, 10 mM EDTA, 5 mM DTT, 1 mM phenylmethanesulfonylfluoride, and 2 M urea. Following centrifugation, the supernatant was applied to a DE53 column equilibrated with the same buffer. Profilin was in the flow-through and was dialyzed into 20 mM Tris (pH 8.0 at 4 °C), 20 mM KCl, 1 mM EDTA, 1 mM DTT, and 1 μg/mL leupeptin. The profilin was clarified and concentrated by Centricon (Amicon) ultrafiltration. The profilin was further purified on a Superdex 200 (GE Healthcare Life Sciences) gel filtration column. Appropriate fractions were pooled, concentrated, and stored in liquid nitrogen.

Formin.

A plasmid containing a GST fusion with mouse diaphanous 1 (mDia) residues 549–1,255 (a gift from David Kovar, University of Chicago, Chicago) was transformed into Rosetta DE3 E. coli (Novagen). The protocol for bacterial expression and protein purification was described in detail by Harris et al (42) for other formin constructs. Briefly, the pelleted bacteria were resuspended in 50 mM Tris⋅HCl (pH 8), 500 mM NaCl, 5 mM EDTA, 1 mM DTT, 0.5 mM AEBSF, 0.5 mM tosyllysine chloromethyl ketone hydrochloride (TLCK), and 2 μg/mL leupeptin, and they were then sonicated and clarified. The supernatant was loaded onto a glutathione-Sepharose 4B (General Electric) column, and the formin was released by thrombin cleavage. Pooled fractions were dialyzed in 150 mM NaCl, 0.1 mM MgCl₂, 0.1 mM EGTA, 2 mM NaPO₄ (pH 7), and 1 mM DTT. Protein was concentrated by Centricon ultrafiltration, and aliquots were frozen in liquid nitrogen and stored at −80 °C.

Smooth muscle tropomyosin.

Tropomyosin was prepared as a byproduct of a chicken gizzard myosin preparation. It remained in the supernatant following myosin precipitation in 20 mM MgCl₂ (43). Isoelectric precipitation at ∼pH 4.6 was used for further purification.

Smooth muscle myosin.

Recombinant baculovirus coding for C terminally FLAG-tagged myosin heavy chain (MYH11) SM2a and another virus coding for both the aortic essential (MYL6) and regulatory (MYL9) light chains were used for expression (provided by Lee Sweeney, University of Pennsylvania, Philadelphia). Sf9 cells (2 × 10⁹) were infected with both the heavy- and light-chain viruses and were harvested after 3 d. Cells were lysed by sonication in 40 mL of 10 mM NaPO₄ (pH 7.5), 0.3 M NaCl, 1 mM EGTA, 4 mM MgCl₂, 1 mM DTT, 0.5 mM AEBSF, 5 μg/mL leupeptin, and 0.5 mM TLCK. The cell lysate was diluted with buffer containing no salt to bring the final NaCl concentration to 0.15 M. MgATP was added to 3 mM, which depolymerizes filaments to the folded monomeric 10S conformation. The solution was immediately clarified for 30 min at 184,000 × g. The clarified lysate was then loaded onto 4 mL of FLAG resin (Sigma), and nonspecifically bound proteins were washed off with 10 mM NaPO₄ (pH 7.5), 0.15 M NaCl, 1 mM EGTA, 4 mM MgCl₂, and 0.2 mM MgATP. The resin was washed with 10 mM NaPO₄ (pH 7.5), 50 mM NaCl, 1 mM EGTA, 4 mM MgCl₂, and 0.2 mM MgATP, and the myosin was then eluted with buffer containing 100 μg/mL FLAG peptide. The peak fractions were pooled and concentrated using Millipore Amicon Ultra centrifugal filters, and protein concentration was determined using the Bio-Rad Protein Assay.

For thiophosphorylation of the regulatory light chain, myosin was diluted into 10 mM NaP0₄ (pH 7.5), 25 mM NaCl, 1 mM EGTA, 4 mM MgCl₂, and 1 mM DTT. Twenty micrograms per milliliter human skeletal muscle myosin light chain kinase (MLYK2), 2 mM CaCl₂, 7.5 μg/mL calmodulin, 5 mM MgCl₂, and 3 mM ATP-γS were added, and the solution was incubated overnight at 4 °C. Complete phosphorylation was verified by gel shift on glycerol/urea gels (43). The NaCl was increased to 0.3 mM to solubilize the myosin, and the protein was stored at −20 °C in 50% glycerol, 0.3 M NaCl, 10 mM imidazole (pH 7.5), 5 mM MgCl₂, and 1 mM DTT.

Tropomyosin Binding Assay.

Tropomyosin (0.1–1.5 μM) was incubated with 3 μM actin for 10 min at room temperature in 10 mM imidazole (pH 7.5), 200 mM NaCl, 2 mM MgCl₂, and 1 mM DTT. The mixture was sedimented for 25 min at 350,000 × g (4 °C). Pellets were analyzed by SDS/PAGE gels stained with Coomassie Blue. Band intensity was quantified using Kodak 1D Image Analysis Software. The binding constant was determined by fitting the data to the Hill equation as described by Barua et al. (44).

Gel Preparation.

One-dimensional isoelectric focusing gels.

Isoelectric focusing gels were run essentially according to the method described by Otey et al. (45). A total of 2.5–5 μg of purified actin samples in 5 μL or less was diluted with 3 volumes of solubilization buffer, which contained 1% ampholyte pH 3/10, 4% ampholyte pH 5/7 (Bio-Rad), 2% Nonidet P-40, 9.5 M urea, and 5% β-mercaptoethanol. Vertical slabs (8 × 7 cm) were made according to the method of O’Farrell (46) and run at 3 V constant for 18 h at room temperature. Following electrophoresis, gels were stained for at least 2 h with 0.04% Coomassie Blue G-250 and 5% perchloric acid, and then destained with 25% isopropyl alcohol and 10% acetic acid.

Native gels.

Native gels were run according to the method of Nosworthy et al. (47). A 10% polyacrylamide gel containing 10 mM 3-(N-morpholino)propanesulfonic acid (MOPS; pH 6.8), 0.2 mM ATP, 0.2 mM CaCl₂, 0.05% ammonium persulfate, and 0.01% N,N,N′,N′-Tetramethylethylenediamine was prepared. The running buffer was 10 mM MOPS (pH 6.8), 0.2 mM ATP, and 0.2 mM CaCl₂. Actin was incubated with DNase or gelsolin segment-1 in G-buffer at room temperature for 10 min, and then mixed with an equal volume of loading buffer [62.5 mM MOPS (pH 8), 10% glycerol, 10 mM DTT]. Gels were run at 100 V for 4 h at room temperature.

Rhodamine Labeling of Actin.

Expressed actin in G-buffer [5 mM Tris (pH 8.26 at 4 °C), 0.2 mM CaCl₂, 0.2 mM Na₂ATP, 0.5 mM DTT] was polymerized by addition of 100 mM KCl and 0.5 mM MgCl₂. Fresh 0.5 mM DTT was added. The mixture was incubated at room temperature for 1 h, and then dialyzed against 100 mM NaCl, 50 mM piperazine-N,N′-bis(2-ethanesulfonic acid) (pH 6.8 at 4 °C), 0.2 mM CaCl₂, 0.2 mM Na₂ATP, and no DTT for a minimum of 4 h. The Lys-reactive dye N-hydroxysuccinimide-ester-Rhodamine (NHS-Rhodamine; Thermo Scientific, Pierce) was added in a sixfold molar excess over actin and incubated for 2 h at room temperature. A similar protocol was used when the actin was labeled with NHS-Alexa 555 (Life Technologies), except that a fivefold molar excess of dye was used and the reaction was incubated at 4 °C overnight. The reaction was stopped with 50 mM Tris (pH 7.5), and the filaments were pelleted by centrifugation for 45 min at 350,000 × g. The labeled actin was resuspended in G-buffer and dialyzed against G-buffer for 3 d to depolymerize the actin and remove excess label. The labeled actin was clarified by centrifugation (45 min at 350,000 × g), frozen in liquid nitrogen, and stored at −80 °C. The degree of labeling was determined using an extinction coefficient of 80,000 M⁻¹⋅cm⁻¹ at 555 nm for rhodamine. Actin concentration was quantified with the Bio-Rad Protein Assay. The typical degree of labeling was 0.5 mol of rhodamine per mole of actin.

Procedures for In Vitro Motility.

Formation of actin filaments for motility.

Actin in G-buffer [5 mM Tris (pH 8.26 at 4 °C), 0.2 mM CaCl₂, 0.1 mM NaN₃, 0.5 mM DTT, 0.2 mM Na₂ATP, 1 μg/mL leupeptin] was clarified by centrifugation (25 min at 350,000 × g), and its concentration was determined using the Bio-Rad Protein Assay. The actin was diluted to 12 μM, and Ca²⁺ was exchanged for Mg²⁺ by adding 1 mM EGTA and 3 mM MgCl₂ for 5 min at 4 °C. If tropomyosin was present, it was added at a 0.25 M ratio relative to actin during polymerization. Polymerization was initiated by addition of 100 mM KCl, 5 mM MgCl₂, and 10 mM imidazole (pH 7.0; 15 min at 37 °C or 1 h at room temperature). Rhodamine/phalloidin was added at a 1:1 molar ratio after polymerization. For experiments without phalloidin, NHS-Rhodamine–labeled WT actin was incorporated as 25% of the total actin.

In vitro motility.

Motility was performed at 30 °C, essentially as described previously (43). Motility buffer contained 90 mM KCl, 25 mM imidazole (pH 7.5), 4 mM MgCl₂, 1 mM EGTA, and 10 mM DTT. Full-length human smooth muscle myosin (100 μL of 0.75 mg/mL) in motility buffer with 0.3 M KCl was mixed with a twofold molar excess of F-actin and 1 mM MgATP and centrifuged for 25 min at 350,000 × g to remove ATP-insensitive myosin heads. The protein concentration of the supernatant was determined using the Bio-Rad Protein Assay, and then diluted to 150 μg/mL. The myosin was added to the flow cell and adsorbed directly onto nitrocellulose coated coverslips for 60 s. Two rinses of 0.5 mg/mL BSA in motility buffer were added to block the surface. An actin wash with fragmented, unlabeled actin was then used to block rigor heads further as described (48). F-actin (1.5–3 μg/mL) was then added to the flow cell for 30 s and rinsed with motility buffer. The assay was performed in motility buffer with 1 mM MgATP, 0.7% methylcellulose, and an oxygen scavenging system (3 mg/mL glucose, 0.13 mg/mL glucose oxidase, 0.05 mg/mL catalase). Actin movement was observed using an inverted microscope (Zeiss Axiovert 10) equipped with epifluorescence, a Rolera MGi Plus Digital camera, and a dedicated computer with the Nikon NIS Elements software package.

Data with rhodamine/phalloidin–labeled actin were analyzed using a semiautomated filament tracking program (49). The program automatically determines the trajectory of each actin filament with a speed greater than zero and movement lasting at least 10 frames. Each dataset yields a weighted distribution of filament speeds, which is fit with a Gaussian to determine a mean velocity and SD. Hundreds of filaments are thus tracked without selection bias. Filaments visualized by incorporating 25% NHS-modified WT actin, which were dimmer, were tracked by hand using ImageJ. Twenty filaments were chosen for analysis using the criteria of smooth movement for 30 or more frames.

ADP Dissociation Rate.

The stop-flow experiments were carried out on a Kintek SF-2002 stop-flow apparatus. Light scattering was measured with an exciting light of 294 ± 10 nm, and emission was monitored with a 294-nm cutoff filter. The rate of ATP-induced dissociation of the acto-HMM.ADP complex was measured by mixing a solution containing WT or R258C actin (1.2 μM), smooth muscle tropomyosin (0.25 μM), smooth muscle HMM (1 μM), and 100 μM MgADP with 2 mM MgATP. Light scattering traces were fit to single exponentials using software provided by Kintek. Buffer conditions were 10 mM imidazole (pH 7.4), 90 mM KCl, 4 mM MgCl₂, 1 mM EGTA, and 1 mM DTT at 20 °C.

Force Data Acquisition and Analysis.

Buffer A contains 10 mM imidazole (pH 7.5), 25 mM KCl, 1 mM EGTA, and 4 mM MgCl₂. Buffer B contains all of the ingredients of buffer A plus scavenger (0.13 mg/mL glucose oxidase, 0.05 mg/mL catalase, 3 mg/mL glucose), 1 mM MgATP, and the ATP regenerating system (100 U/mL pyruvate kinase, 0.5 mM phosphoenol pyruvate).

To prepare neutravidin-coated beads, 20 μL of a 5% 1-μm polystyrene bead stock (Polysciences, Inc.) was diluted into 1 mL of buffer A and spun down. The supernatant was removed, and the pelleted beads were mixed with 4 μL of 5 mg/mL neutravidin (Life Technologies). The mixture was incubated at room temperature for 2 d. Five hundred microliters of buffer A was added, and the beads were spun down. The pellet was resuspended in 500 μL of buffer A. The beads were kept on ice for further use. To bind HMM to the neutravidin-coated beads, 2 μL of the neutravidin beads was mixed with 2 μL of 0.2 mg/mL phosphorylated smooth muscle HMM with a C-bio tag (50) at room temperature for at least 10 min.

To make fluorescently labeled bundled actin for the assay, unlabeled WT (or R258C) and WT actin labeled with NHS-Alexa 555 were mixed at an 8:2 ratio, and allowed to polymerize for 15 min at 37 °C following the protocol for polymerization (final actin concentration is 1 μM). Then, fascin (4:1 actin/fascin molar ratio) was added and incubated on ice for at least 1 h before use. The use of fascin-bundled actin filaments greatly increased the success rate of obtaining myosin–actin interaction events in the laser trap experiment. Fascin was cloned and expressed as described by Hodges et al. (51).

Details of the one-bead laser trap experiment are as described earlier (52). Briefly, the flow cell was infused with 20 μL of 0.1 mg/mL N-ethyl maleimide-myosin for 2 min. After rinsing with 20 μL of buffer A, 20-μL actin bundles were flowed in and incubated for 2 min. This procedure was followed by a rinse with 20 μL of 5 mg/mL casein (Sigma–Aldrich). Beads coated with HMM were diluted 100-fold in buffer B and then infused into the flow cell. A bead was then grabbed by the laser trap and brought down to the vicinity of an actin bundle on the surface to start the reaction. A force-feedback mechanism was enabled so that once myosin engaged with actin, the force on the bead was kept constant at 1, 2, 3, 4, or 5 pN for certain periods of time.

To analyze the data, the traces of bead motion in each window of feedback level force were fitted linearly to get the velocity in Clampfit (Axon Instruments, Inc.). Velocity data for a given load for all experiments were compiled and then fit to the Hill force/velocity relation.