Molecular mechanisms of disease-related human β-actin mutations p.R183W and p.E364K

Molecular mechanisms of disease-related human b-actinmutations p.R183W and p.E364KNikolas Hundt1, Matthias Preller1,2, Olga Swolski1, Angella M. Ang1, Hans G. Mannherz3,Dietmar J. Manstein1 and Mirco M€uller1,*

1 Institute for Biophysical Chemistry, Hannover Medical School, Hannover, Germany

2 Centre for Structural Systems Biology (CSSB), German Electron Synchrotron (DESY), Hamburg, Germany

3 Institute for Anatomy and Molecular Embryology, Ruhr-University Bochum, Bochum, Germany

Keywords

actin mutations; cytoplasmic b-actin;

molecular dynamics simulations; p.E364K;

p.R183W

Correspondence

M. M€uller, Department of Cardiology and

Angiology, OE 6887, Hannover Medical

School, Carl-Neuberg-Str. 1, D-30625

Hannover, Germany

Fax: +49 511 532 3357

Tel: +49 511 532 3270

E-mail: [email protected]

D. J. Manstein, Institute for Biophysical

Chemistry, OE4350, Hannover Medical

School, Carl-Neuberg-Str. 1, D-30625

Hannover, Germany

Fax: +49 511 532 5966

Tel: +49 511 532 3700

E-mail: [email protected]

*Present address:

Department of Cardiology and Angiology,

Hannover Medical School, Hannover,

Germany

(Received 13 May 2014, revised 6

September 2014, accepted 22 September

2014)

doi:10.1111/febs.13068

Cytoplasmic b-actin supports fundamental cellular processes in healthy and

diseased cells including cell adhesion, migration, cytokinesis and mainte-

nance of cell polarity. Mutations in ACTB, the gene encoding cytoplasmic

b-actin, lead to severe disorders with a broad range of symptoms. The two

dominant heterozygous gain-of-function b-actin mutations p.R183W and

p.E364K were identified in patients with developmental malformations,

deafness and juvenile-onset dystonia (p.R183W) and neutrophil dysfunc-

tion (p.E364K). Here, we report the recombinant production and func-

tional characterization of the two mutant proteins. Arg183 is located near

the nucleotide-binding pocket of actin. Our results from biochemical stud-

ies and molecular dynamics simulations show that replacement by a trypto-

phan residue at position 183 establishes an unusual stacking interaction

with Tyr69 that perturbs nucleotide release from actin monomers and poly-

merization behavior by inducing a closed state conformation. The replace-

ment of Glu364 by a lysine residue appears to act as an allosteric trigger

event leading to the preferred formation of the closed state. Thus, our

approach indicates that both mutations affect interdomain mobility and

nucleotide interactions as a basis for the formation of disease phenotypes

in patients.

Introduction

Actin is a highly conserved and ubiquitous protein

found in nearly all eukaryotic cells. Six actin isoforms

can be distinguished in vertebrates: three a-actin

isoforms (a-skeletal muscle, a-cardiac muscle and

a-aortic smooth muscle, also known as a-vascular),one b-isoform (b-cytoplasmic) and two c-isoforms

Abbreviations

e-ATP, 1,N6-ethenoadenosine-50-triphosphate; IC50, inhibitory concentration; MD, molecular dynamics; Ni-NTA, Ni2+-nitrilotriacetic acid; NM-

2A, nonmuscle myosin-2A isoform.

1FEBS Journal (2014) ª 2014 FEBS

(c-cytoplasmic and c-smooth muscle also known as c-enteric). Cytoplasmic b- and c-actin are essential for

cell migration, cell shape maintenance, mitosis and

intracellular transport processes and are expressed at

moderate to high levels in nearly all adult tissues [1,2].

The most abundant isoactin in many nonmuscle cells

including myeloid and neuronal cells is b-actin [2]. Cel-

lular studies show that the b-isoform is preferentially

recruited into cellular protrusions, stress fibers, circular

bundles and at cell–cell contacts [3,4]. The rapid cyto-

skeletal rearrangements observed for these structures

appear to be linked to the highly dynamic turnover of

actin filaments made from b-actin [5].

All actin isoforms share the same architecture with

four different subdomains and a common nucleotide-

binding site. In vivo, b-actin is post-translationally

modified by cleavage of the first methionine followed

by N-terminal acetylation. Actin is an ATPase and the

hydrophobic nucleotide-binding site is located in the

cleft between subdomains SD1–2 and SD3–4(Fig. 1A). In the presence of divalent cations, mono-

mers with bound ATP assemble into filaments. The

growth of actin filaments depends on the addition of

actin–ATP monomers predominantly at the fast-grow-

ing barbed ends. After monomer addition, the bound

ATP is hydrolyzed to ADP and inorganic phosphate

(Pi) followed by Pi-release. Because dissociation of

actin–ADP occurs preferentially from pointed ends,

the association and dissociation reactions result in

‘treadmilling’, a process whereby subunits migrate

through filaments [6,7]. Filament polarity is main-

tained by ATP hydrolysis and F-actin treadmilling is

accompanied by an increased rate of ATP hydrolysis

[8,9].

Mutations in actin-coding genes lead to severe disor-

ders. Compared with myopathy- and angiopathy-asso-

ciated defects of muscle isoactins, mutations of b- andc-actin isoforms lead to a wider spectrum of diseases

that include deafness, cancer and developmental disor-

ders [10,11]. Most biochemical studies in the past,

dealing with human actin mutations, have focused on

skeletal and cardiac isoactins or have aimed to charac-

terize disease-related mutations by introducing them

into the endogenous actin from yeast or other simple

eukaryotic model systems [12–14]. We describe the

recombinant production and functional characteriza-

tion of two dominant heterozygous gain-of-function

mutations in ACTB, the gene encoding human b-actin.Mutation p.E364K has been linked to neutrophil dys-

function [15] and p.R183W to a complex disease phe-

notype that includes developmental malformations,

deafness and delayed-onset dystonia [10]. The bio-

chemical properties of the mutant proteins are

described in detail. Functional differences from the

wild-type protein are explained by structural changes

observed in molecular dynamics simulations.

wt p.R183W

p.E364K

kDa2001501201008570605040

30

2520

SD4SD2

SD1

SD3

CBA R183

E364

15

10

kDa

70100130170

40

55

25

35

10

15

wt p.R183W

p.E364K

Fig. 1. Location of mutated residues and purification of human b-actin variants p.R183W and p.E364K. (A) Atomic structure of cytoplasmic

b-actin as determined in complex with profilin (PDB code: 2BTF). Actin subdomains SD1–4 are numbered. The PDB file was modified and

rendered using PYMOL 1.4 highlighting the residues R183 (close to the nucleotide ATP depicted in blue) and E364 (at the bottom of SD1).

(B) SDS/PAGE of purified, untagged b-actin wild-type (wt), p.R183W and p.E364K. (C) Immunoblot analysis of cytoplasmic b-actin

preparations.

2 FEBS Journal (2014) ª 2014 FEBS

Molecular mechanisms of human b-actin mutations N. Hundt et al.

http://www.rcsb.org/pdb/search/structidSearch.do?structureId=2BTF

Results

Recombinant production of human cytoplasmic

b-actin mutants

We used a baculovirus-driven expression of tag-free

b-actin variants in Sf9 cells, because tag-mediated

sequence changes compromise functional competence

in polymerization assays and interfere with N-terminal

processing in acetylation-competent host cells. Proper

post-translational modification of the actin N-terminus

is crucial for the native functional behavior of the pro-

tein. We previously demonstrated by MS analysis that

our recombinantly produced actin variants are N-ter-

minally acetylated [16]. A disadvantage of tag-free

actin expression in insect cells is the presence of con-

taminating endogenous actin. Our actin preparations

contain 5–15% endogenous actin, as previously shown

by 2D gel electrophoresis and MS analyses [16]. As

shown in Fig. 1B, untagged wild-type (wt) and the

mutant b-actin were purified to homogeneity. Prepara-

tions of cytoplasmic b-actin were confirmed by immu-

noblot analysis (Fig. 1C). Immunoblotting was

performed using an antibody that recognizes the N-ter-

minal acetylated epitope of human b-actin and does

not bind to insect actin. Similar yields (~ 5 mg per

3 9 109 cells) were obtained for wt-, p.R183W- and

p.E364K-b-actin from different preparations (n = 3–5).

Actin folding, stability and nucleotide release

Cytoplasmic actin filaments are highly dynamic struc-

tures undergoing constant assembly and disassembly

cycles to drive motile processes and to serve as tracks

for motor proteins. Correct folding and nucleotide

turnover of actin monomers are crucial for their ability

to assemble into filaments.

Improper folding of actin variants has been proposed

to be associated with human diseases by a number of

studies [12–14]. We tested the ability of the b-actinmutations p.R183W and p.E364K to affect protein con-

formation and stability. The DNase I inhibition assay

represents a classical approach to assess actin folding.

Only properly folded G-actin is able to inhibit DNase I

activity [17,18]. Compared with monomeric wt-b-actin(inhibitory concentration [IC50]: 63 � 4 nm) (Fig. 2A),

mutant p.R183W displayed approximately threefold

greater inhibition of DNase I-mediated nucleic acid

cleavage (IC50: 24 � 1 nm; P < 0.0001). This result

suggests an effect of the p.R183W mutation on the con-

formation of SD2, which contains the DNase I binding

loop (D-loop). Likewise, we observed an increase in

DNase I affinity for mutant p.E364K (IC50: 49 � 2 nm;

P = 0.0028) indicating a link to structural alterations

within SD2. In addition, a higher affinity to DNase I

may be linked to a more closed conformation between

SD2 and SD4 in the mutants, as outlined below.

To further assess the thermal stability of the b-G-

actin variants, we equilibrated the actin used for

DNase I inhibition assays over a range of increasing

temperatures until full denaturation was reached. The

deduced Tm values were in the range of 55.3–55.9 � 0.8 °C (Fig. 2B). Accordingly, thermal stability

and unfolding behavior of the mutant b-actins appearsto be unchanged compared with the wild-type.

Actin monomers with bound ATP preferentially

bind to the plus or barbed ends of F-actin. In fact,

actin with bound ATP has a 10-fold higher association

rate than ADP–actin [19]. The hydrolysis of ATP

A

B

Fig. 2. Folding and stability of b-actin mutants. (A) Folding of

monomeric b-actin variants was assessed by their ability to inhibit

DNase I activity. The mutant actins bind more strongly to DNase I.

(B) Thermal unfolding was monitored using the same DNase I

inhibition assay after incubating actin at the indicated temperatures.

The thermal stability of the mutant actins is unchanged.


N. Hundt et al. Molecular mechanisms of human b-actin mutations

followed by the release of inorganic phosphate occurs

after binding to barbed ends. During dynamic actin

cycling ADP–actin monomers dissociate from the

minus or pointed ends of F-actin. Exchange of ADP

for ATP is crucial for the reassembly of actin mono-

mers to the barbed ends. During nucleotide exchange

actin has to undergo transitions between open and

closed states that involve twisting of its two major

domains [20]. Any impairment of the conformational

flexibility associated with these transitions is predicted

to affect nucleotide exchange and polymerization

behavior. We measured the rate of nucleotide release

under monomeric conditions using the fluorescent

ATP analog 1,N6-ethenoadenosine-50-triphosphatee-ATP). The release kinetics of e-ATP are best

described by a double-exponential model including a

fast and a slow process. The fast process corresponds

to nucleotide exchange from free actin monomers. The

slow process most likely corresponds to the formation

of polymerization-competent actin nuclei or oligomers.

Its contribution to the signal amplitude is small (15–20%) [21]. Monomeric wt-b-actin displayed a fast pro-

cess rate of 0.062 � 0.015 s�1. The fast process was

2.4-fold slower for p.R183W (0.0254 � 0.0005 s�1;

P = 0.0113) and 2.1-fold slower for p.E364K-b-actin(0.030 � 0.003 s�1; P = 0.0177) (Fig. 3). The slow

process remained unchanged with values in the range

of 0.0016–0.0020 � 0.0004 s�1 for the actin variants.

A reduced exchange rate of ADP for ATP in actin

monomers is likely to affect the rate of polymerization.

Indeed, this behavior is reflected by our polymeriza-

tion assays, as outlined below.

Actin assembly and disassembly

The ability of actin monomers to assemble into polar

filaments is crucial for their physiological function.

G-Actin undergoes multiple conformational changes

upon polymerization. The incorporated monomer

subunits are flattened by subdomains SD1–2 undergoinga 12–13° propeller twist with respect to subdomains

SD3–4. Secondary elements such as the DNase I-bind-

ing loop are structurally reorganized. We assessed the

actin assembly behavior of both mutant b-actins by

monitoring the increase in the fluorescence of pyrene-

labeled actin that coincides with filament formation

[22]. The p.R183W mutant as well as the p.E364K var-

iant exhibited significantly slower polymerization rates

compared with wild-type actin (Fig. 4A). The apparent

half-time t1/2, pol of polymerization was increased 1.9-

fold for the p.R183W-b-actin (39.9 � 4.4 min;

P = 0.0006) and 2.6-fold in the case of p.E364K-b-actin (52.2 � 5.2 min; P < 0.0001) compared with

wild-type actin (20.6 � 2.4 min) (Fig. 4A, inset). In

addition, we monitored the ATPase activity of the fila-

mentous b-actin variants. In comparison with wt-b-actin, which showed a hydrolysis rate of

2.1 � 0.3 h�1, the p.R183W mutant exhibited a 1.7-

fold (P = 0.0141) increase in ATP turnover

(3.6 � 0.4 h�1) (Fig. 4B). Filaments made from

p.E364K-b-actin did not differ significantly from the

wild-type with an ATPase activity of 2.8 � 0.4 h�1

(P > 0.05). Depolymerization assays were performed

to analyze whether the increased ATP turnover in

p.R183W filaments and, thus, faster accumulation of

ADP–actin, affects the disassembly step. Indeed, the

depolymerization half-time of the p.R183W mutant

was significantly reduced (t1/2, depol, p.R183W =9.1 � 0.8 min; P = 0.0204), whereas wt- and p.E364K-

b-actin had similar depolymerization kinetics (t1/2, de-

pol, wt = 12.2 � 1.0 min; t1/2, depol, p.E364K = 13.0 �1.6 min; P = 0.67) (Fig. 4C). In summary, slower fila-

ment growth, higher ATP-hydrolysis and faster depo-

lymerization indicate that p.R183W actin is impaired

in forming long, stable filaments.

Interactions with profilin and myosin

Profilin is an important actin-binding protein that

regulates the dynamic reorganization of the actin

Fig. 3. Nucleotide exchange in the mutant actin monomers is

impaired. Time courses of e-ATP release from monomeric b-actin

variants in the presence of excess ATP are measured by the

decrease in e-ATP fluorescence (n = 2–5). In order to visualize the

rate differences for the fast phase, which corresponds to

nucleotide exchange in monomeric actin, the data were

normalized to the fast phase amplitudes. Only the fast phase is

shown (100 s), however, the datasets extend over a period of

25 min and were fitted using double exponential functions.



cytoskeleton and drastically increases the exchange

rate of actin-bound ADP to ATP [23]. Nunoi and

coworkers reported that the interaction of profilin with

the p.E364K variant purified from patients’ cells is

impaired [15]. By contrast, this result was not verified

by another study in which an in vitro translation

approach was applied for protein production and

interactions with different actin-binding proteins were

screened using band shift assays [13]. In order to test

the ability of our recombinant actin constructs to

interact with profilin II, we performed microscale ther-

mophoresis experiments to determine affinities [24]. As

shown in Fig. 5A, the equilibrium dissociation con-

stant Kd for wt-b-actin was 5.9 � 1.2 lM. Our experi-

ment did not show a significant change in profilin

binding for the p.E364K mutant (Kd = 8.9 � 0.8 lM;P = 0.095) in comparison with the wt-b-actin.Moreover, binding of p.R183W-b-actin to profilin II

was similar to wild-type actin with an equilibrium

dissociation constant of 6.8 � 0.5 lM.

CBA

Fig. 4. Properties of filamentous b-actin wild-type and mutants p.R183W and p.E364K. (A) Polymerization of the mutant actins is slowed.

Polymerization of 10 lM b-actin variants was initiated by a salt shift as described in the Materials and methods. Shown are the averaged

traces from ~ 10 independent measurements using two different batches of wt-, p.R183W- and p.E364K-b-actin. The t1/2 values from the

kinetic traces are depicted in the inset. (B) Intrinsic ATP hydrolysis rates of b–F-actin variants. Actin (10 lM) was freshly polymerized and

the ATP turnover was measured in a NADH-coupled assay (n = 2–3). ATP hydrolysis in p.R183W actin filaments is increased. (C) The

depolymerization rate of p.R183W actin filaments is increased. Depolymerization was induced by diluting pyrene-labeled actin filaments

below the critical concentration. The half-times of the fluorescence decrease were determined from fitting single exponential functions to

the traces. Statistically significant differences (P < 0.05) are marked with asterisks.

A CSD4 SD2

SD1SD3

SD4 SD2

SD1SD3

D-loopD-loop

E364 E364

371H371H

961Y961Y573F573F M355 M355

B

Fig. 5. Interaction with profilin II. Residue E364 does not appear to be essential for profilin binding. (A) Dissociation constants of profilin II

and monomeric b-actin variants were quantified by microscale thermophoresis (n = 2). The mutant actins bind profilin normally. The lack of

a larger effect can be explained by the peripheral location of E364 in relation to the profilin-binding site in both structures showing a closed

(B, PDB: 2BTF) and open nucleotide-binding site (C, PDB: 1HLU) for the actin–profilin complex. Actin is shown in orange and profilin in

grey. Primary actin contacts include H173, F375, (light orange spheres), Y169 and M355 (yellow orange spheres).




http://www.rcsb.org/pdb/search/structidSearch.do?structureId=1HLU

A major role of cytoplasmic b-actin is the formation

of polar tracks on which myosin motor proteins can

move in a directional fashion. The nonmuscle myosin-

2A isoform (NM-2A) colocalizes with b-actin in cells

and its ATPase activity is preferentially activated by

b-actin [16]. We tested whether the mutations p.R183W

and p.E364K in b-actin have an effect on the

activation of NM-2A ATPase activity (Fig. 6). We

found that 30 lM of filamentous wt-b-actin stimulated

the ATP turnover of NM-2A ~ 15-fold (0.189 �0.015 s�1, basal ATPase activity was 0.013 �0.0006 s�1). Activation of NM-2A ATPase activity by

p.R183W actin was reduced fourfold under the same

conditions (0.045 � 0.013 s�1), whereas activation by

mutant p.E364K resembled that observed with wt-b-actin (0.176 � 0.029 s�1).

Molecular dynamics simulations

To analyze the impact of the mutations on the struc-

ture and dynamics of b-actin monomers at the atomic

level, we carried out all-atom molecular dynamics

(MD) simulations in explicit water using NAMD 2.9

[25]. Starting with the high-resolution X-ray crystal

structure of b-actin in the ATP-bound state (PDB:

2BTF) [26], we introduced single-point mutations

using the SCHR€ODINGER SUITE [27] and replaced the

bound Sr2+∙ATP by Ca2+∙ATP, because calcium ions

were present in all our experimental setups. The analy-

sis was based on 50 ns trajectories of the energy mini-

mized and equilibrated simulation systems for each

actin monomer. Simulation of the wt-b-actin under

the same conditions as the mutants served as the con-

trol of unaffected actin. In accordance with previous

studies [28,29], the presence of Ca2+∙ATP led to an

opening of the nucleotide cleft between the SD1–SD2

and SD3–SD4 domains of wt-b-actin during our simu-

lations, which was mainly achieved by rearrangements

in SD1–SD2 and a domain rotation of the entire SD2

by ~ 37° (Fig. 7A). The final structure of the wt-b-actin monomer after 50 ns simulation time closely

resembled the crystal structure of b-actin∙Ca2+∙ATP,

solved in the open state (PDB: 1HLU; [29]), when

superimposed based on the protein Ca atoms

(Fig. 7B).

During our simulations of the p.R183W mutant, this

opening of the nucleotide cleft was significantly

impaired (Figs 7C and 8B, Movies S1 and S2). Com-

pared with the simulations with wt-b-actin, the maxi-

mal cleft opening reached only 30%. The mutated

tryptophan is located in SD4 at one side of the nucleo-

tide cleft and its bulky side chain moved into that cleft

along the trajectory, thereby blocking in part the

potential exit route for Pi. In addition, Trp183 formed

a stacking interaction with Tyr69 of SD2, which seems

to contribute to the restricted opening of the cleft in

our simulations (Fig. 7C and Movie S2). As a conse-

quence, the interaction of the critical Ser14 in the

active site with the c-phosphate of ATP remained sta-

ble throughout the simulations (Fig. 8A). This interac-

tion broke in all wild-type simulations after around

25 ns simulation time, concurrent with the progress of

the cleft opening. Furthermore, the impaired ability of

SD2 to rotate away from SD4 in order to open the

nucleotide cleft during our simulations appeared to

translate into a shift of SD3, in particular of loop

300–309 which is involved in binding the adenine base

of the nucleotide in a hydrophobic pocket. Hence,

along the trajectory, the nucleotide base lost its inter-

actions to SD1 and SD3, and slipped out of the bind-

ing pocket, whereas the triphosphate moiety remained

unaffected (Fig. 7C, transparent silhouette of ATP).

By comparison, mutation p.E364K is located in

SD1 ~ 38 �A from the nucleotide-binding pocket or the

nucleotide cleft. The p.E364K mutant also showed an

impact on cleft opening during the MD simulations,

although apparently via a different mechanism than

for p.R183W (Figs 7D and 8B,D, Movies S1 and S3).

Indeed, the SD2 tilted notably and drew even closer

towards SD4 and the nucleotide. As a consequence,

the nucleotide cleft closed to a greater extent than in

p.R183W or wt-b-actin in the closed state, narrowing

to a width of ~ 4 �A at the tightest spot (Fig. 7D). The

interaction between Ser14 and the c-phosphate of ATP

remained intact, as seen for mutant p.R183W

(Fig. 8A). In addition to the perturbed nucleotide cleft

Fig. 6. Interaction with nonmuscle myosin. The interaction of

p.R183W actin and nonmuscle myosin-2A is impaired. The

steady-state ATPase rate of the myosin was measured in the

absence (basal) and presence of 30 lM of the actin variants in a

NADH-coupled assay. *Statistical significance at P < 0.05.





opening and the stable interaction of SD1 to the nucle-

otide via Ser14 and the c-phosphate, the p.E364K

mutation led to further complex conformational

changes throughout the entire protein, which was not

observed for p.R183W (Fig. 7D and Movie S3). Fig-

ure 8D depicts the difference of the root mean square

fluctuations of wt-p.E364K, clearly showing drastic

changes in the conformational dynamics and structure

of all four subdomains. As a further control and to

exclude potential misinterpretation and bias introduced

into the trajectories by the choice of the starting rot-

amer for the mutated amino acid residues, we repeated

simulations of both mutants starting each with two

additional, distinct rotamers. Analyses of these control

simulations confirm the considerable impact of the two

mutations on cleft opening (Fig. 8C). Accordingly, the

interaction of Ser14 with the c-phosphate of ATP

remains intact in all mutant simulations.

Discussion

To study the consequences of the disease-related

human b-actin mutations p.R183W and p.E364K, we

examined the properties of the mutant proteins at the

level of the monomers, in the filamentous state, and in

the context of interactions with selected binding part-

ners. The mutant actins display normal thermal stabil-

ity indicating that the mutations do not interfere with

folding. Although the two mutations are located in dif-

ferent parts of the molecule, p.R183W and p.E364K

actin display similar changes in their biochemical

behavior compared with the wild-type protein. In the

monomeric state, both mutants show increased affinity

for DNase I and reduced exchange of nucleotides.

The results of our MD simulations suggest that the

mutated actins favor a conformation similar to the

closed state of wild-type actin (Fig. 7C,D). The con-

comitant establishment of a binding surface for

A B

C D

Fig. 7. Conformational changes of wild-type and mutant b-actins during molecular dynamics simulations. (A) Conformational changes of

wt-b-actin along the MD trajectories. Comparison of the starting structure (gray) and the conformation after 50 ns MD simulations

(colored). Structural changes are indicated by blue vectors and Mg2+∙ATP is shown as a stick representation. Note the rearrangements in

SD1 and SD2. SD2 rotates by ~ 37°. The rotation axis was determined using the program DYNDOM [50]. (B) Comparison of the wt-b-actin

conformation after 50 ns MD simulation (colored) and the high-resolution X-ray structure of wt-b-actin in the open state (PBD: 1HLU; light

gray). The final actin structure resembles closely the open state crystal structure. (C, D) Single-point mutations p.R183W and p.E364K both

interfere with the opening of the nucleotide cleft, mainly by affecting the rotation of SD2, however, through distinct mechanisms. The

p.E364K mutation appears to affect all subdomains, whereas the p.R183W mutation directly interferes with cleft opening. Color code: teal,

subdomain 1 (SD1); pink, subdomain 2 (SD2); orange, subdomain 3 (SD3); green, subdomain 4 (SD4).




DNase I and the trapping of nucleotide at the active

site correlate well with our experimental results. Our

computational analysis predicts that the tryptophan

residue in p.R183W actin forms a stacking interaction

with Tyr69 in SD2. As a result, the gap between SD2

and SD4 is closed and the hydrogen bond between

Ser14 and the c-phosphate of ATP is stabilized. The

latter interaction has been suggested to play a key role

in sensing bound ATP and to stabilize the closed con-

formation of the nucleotide-binding pocket. It is

thought to break upon ATP-hydrolysis, which facili-

tates opening of the nucleotide-binding site and prod-

uct release [30–32]. A p.S14A mutant shows an

increased nucleotide exchange rate, higher susceptibil-

ity to proteolysis, and decreased DNase I affinity

[33,34]. The mutant displays the opposite biochemical

behavior to the p.R183W mutant, most likely because

it adopts an open nucleotide-binding pocket even in

the presence of ATP.

Our MD simulations show that mutation p.E364K

induces conformational changes in SD1 that propagate

through the entire monomer. As a result, p.E364K

actin forms a closed conformation in the presence of

ATP under formation of a stable hydrogen bond

between Ser14 and the c-phosphate of ATP. However,

the mechanism is clearly different from the effect of

p.R183W. Given the distance of the mutated amino

acid residue to the nucleotide-binding cleft, the

p.E364K mutation appears to act via an allosteric

mechanism.

With regard to predicting the behavior of actin

monomers following their incorporation into the fila-

ment, the information provided by our MD simula-

tions is limited. However, our in vitro experiments still

provide insight into the mutation-derived defects in fil-

amentous actin. Reduced nucleotide exchange in

monomers of both mutant actins slows regeneration of

the ATP–actin pool from ADP–actin, which leads to

an accumulation of ADP–actin and reduces the avail-

able amount of polymerization competent actin mono-

mers. In addition, in the presence of ATP–actin, as in

our polymerization assay, both mutants display a

decreased polymerization rate. We hypothesize that

the impaired interdomain mobility affects the incorpo-

ration process of monomers into the filament, because

this process involves conformational rearrangements of

A B

C D

Fig. 8. Consequences of the b-actin mutations on the conformational dynamics. (A) Stability of the interaction between serine 14 and the

c-phosphate of ATP during the simulations of wt- (black), p.R183W- (red) and p.E364K-b-actin (blue) as indicated by the distance between

the hydroxyl group of serine 14 and the phosphorous atom of the c-phosphate. Both point mutations prevent breaking of this crucial

interaction. (B) Opening of the nucleotide cleft in the actin monomers, measured by monitoring the distance between Gly55 and Arg183

along the trajectories. Note that the shift in the cleft distance for p.E364K around 3 ns is caused by a tilting of the a helix in SD2, which

decreases the cleft width even further rather than increasing it. (C) Control simulations with two additional starting rotamers per p.R183W

and p.E364K mutant confirm the impaired opening of the cleft by the mutations. (D) Difference in the root mean square fluctuations

(RMSF) of wt- and p.E364K-b-actin averaged over the amino acid residues. The single-point mutation p.E364K leads to complex allosteric

conformational changes in all four subdomains of the protein.



the subdomains such as twisting of SD1–2 against

SD3–4 and reorganization of SD2 and its D-loop [20].

These conformational changes have been linked to the

enhanced ATPase activity of actin monomers follow-

ing their incorporation into filaments [9]. ATP-hydro-

lysis in p.R183W mutant F-actin is increased 1.7-fold

and the filament disassembly is 34% faster than in

wt-b-actin. Hence, we assume that the p.R183W

mutant forms shorter and less stable filaments. Fila-

ment instability of p.R183W actin can be linked to the

lack of lamellipodial protrusions in lymphoblastoid cell

lines derived from patients carrying this mutation [10].

Because the proper formation of lamellipodia is impor-

tant for coordinated cell migration, the observed devel-

opmental malformations in the patients with p.R183W

actin can be partially explained by our findings. Fur-

thermore, b-actin is crucial for the maintenance of ste-

reocilia in cochlear hair cells [35]. Therefore, unstable

actin filaments made from p.R183W actin are likely to

contribute to the patients’ hearing loss.

Actin dynamics are important for many intracellular

signaling pathways, particularly the chemotactic fMLP

receptor pathway in neutrophil granulocytes [36,37].

Its activation induces rapid actin polymerization and

depolymerization events. Nunoi et al. have reported

that neutrophils from patients with p.E364K actin

show a reduced fMLP receptor-mediated chemotactic

response [15]. With regard to our results, the p.E364K

mutation is likely to hinder rapid polymerization of

b-actin during fMLP receptor signaling and thereby

contributes to neutrophil dysfunction.

We examined the interactions of the mutant actins

with profilin and myosin in detail. Profilin triggers

nucleotide exchange. The complex of profilin with

ATP–actin is recruited to the barbed end of growing

filaments in the cell [38,39]. Contradictory results

regarding the interaction of p.E364K actin with profi-

lin have been published [13,15]. We tested the interac-

tion of the proteins using microscale thermophoresis.

Our results show only minor differences in the profilin

affinity of p.E364K-b-actin compared with the wild-

type. Profilin binds in a groove between SD1 and SD3

with actin residues Tyr169, His173, Met355 and

Phe375 forming the central interaction interface (see

also Refs [26,29] and Fig. 5B,C). They are less

involved in structural alterations predicted by our MD

simulations (Fig. 7C,D). As the thermophoresis experi-

ment suggests, Glu364 is not essential for profilin

binding. The structures in Fig. 5B,C show that this

residue is situated at the periphery of the actin–profilinbinding interface.

The interaction with myosin is severely impaired in

the case of p.R183W actin. Filaments formed by this

mutant show a fourfold reduced ability to activate the

ATPase of NM-2A. We assume that allosteric pertur-

bation by the p.R183W mutation affects actin’s capac-

ity to interact with the myosin.

Both nonsarcomeric myosin and cytoplasmic actin

have complementary roles in cell migration and over-

lapping functions in the maintenance of stereocilia

[35,40,41]. Therefore, a defective actomyosin interac-

tion is likely to contribute to the phenotype of the

patients carrying the p.R183W mutation [10].

Our study of disease-related b-actin variants provides

insight into the mechanisms that drive the conforma-

tional changes actin undergoes during its polymeriza-

tion and depolymerization cycle. Actin has evolved a

dense allosteric network that regulates the transition

between the monomeric and the filamentous form as

well as the interaction with many cellular components.

Our work highlights how changes in this network by sin-

gle point mutations affect the functional competence of

the molecule and thereby lead to severe disorders.

Materials and methods

Reagents

The Phusion Site-Directed-Mutagenesis Kit, goat anti-

(mouse) IgG conjugated with horseradish-peroxidase,

SuperSignal West Femto Maximum Sensitivity Substrates

and desalting columns were purchased from Thermo Fisher

Scientific (Schwerte, Germany). Ni2+-nitrilotriacetic acid

(Ni-NTA) superflow resin was purchased from Qiagen

(Hilden, Germany). e-ATP was from Life Technologies

(Carlsbad, CA, USA). Monoclonal anti-(b-actin) IgG

(mouse, clone AC-74) and standard reagents were

purchased from Sigma-Aldrich (St. Louis, MO, USA). N-

(1-pyrene)iodoacetamide and Cellfectin II were from Invi-

trogen (Carlsbad, CA, USA), recombinant bovine pancreas

DNase I and complete protease inhibitor cocktail tablets

were from Roche (Basel, Switzerland).

Plasmid construction and baculovirus generation

The pFastBac plasmid encoding tag-free human cytoplas-

mic b-actin and the human gelsolin C-terminal segments

G4–6 were generated as previously described [16]. Site-

directed mutagenesis of b-actin was performed using the

following 50-phosphorylated oligonucleotides (sequences

from 50 to 30): TACCTCATGAAGATCCTCACCGAGCG

(p.R183W sense primer), GTCAGTCAGGTCCCAGC

CAGCCAGGTC (p.R183W mutagenic antisense primer),

CAGGAGTATGACAAGTCCGGCCCCTCC (p.E364K

mutagenic sense primer), CTTGCTGATCCACATCTGCT

GGAAGGT (p.E364K antisense primer). The mutation

nomenclature follows the guidelines of the HGVS (Human



Genome Variation Society). The resulting plasmids were

double-strand sequenced to verify mutagenesis and to

exclude the possibility of amplification-associated errors.

The baculovirus transfer vectors for wild-type b-actin and

both mutants were transformed in DH10Bac

Escherichia coli cells and the recombinant bacmids were

transfected into Sf9 (Spodoptera frugiperda) insect cells. Ba-

culovirus was amplified according to the manufacturer’s

protocol and Sf9-cells were infected at MOI 25. Cells were

harvested 3 days post infection and stored at �80 °C.

Protein production

Tag-free human b-actin wild-type (wt), p.R183W and

p.E364K were purified from Sf9-cells as G-actin (globular

or monomeric actin) by affinity chromatography using the

C-terminal gelsolin half G4–6 [16,42]. Briefly, Sf9 cells were

lyzed by sonication in lysis buffer (10 mM Tris/HCl pH 8.0,

5 mM CaCl2, 4% Triton X-100, 1 mg�mL�1 Tween-20,

1 mM dithiothreitol, 1 mM ATP and protease inhibitor mix-

ture). Afterwards, 50 mM KCl, 20 mM imidazole, pH 8.0,

and a threefold excess of gelsolin G4–6 were added and

incubated at 4 °C overnight. The lysate was cleared by

centrifugation. The actin–gelsolin complex was bound to a

Ni-NTA column and washed with wash buffer (10 mM

Tris/HCl pH 8.0, 5 mM CaCl2, 50 mM KCl, 20 mM imidaz-

ole). Actin was eluted with elution buffer (10 mM Tris/HCl

pH 8.0, 50 mM KCl, 1.5 mM EGTA). The actin was dia-

lyzed against actin storage buffer (10 mM Tris/HCl pH 8.0,

0.2 mM MgCl2, 1 mM dithiothreitol), concentrated, frozen

in liquid nitrogen and stored at –80 °C. The G4–6 gelsolin

deletion mutant was expressed and purified from E. coli

according to Ohki et al. [42]. Human NM-2A and human

profilin II were prepared as described elsewhere [43,44].

The homogeneity of the protein preparations was con-

firmed by 10% SDS/PAGE and western blot analysis using

an anti-(b-actin) IgG (Fig. 1B,C).

Pyrene-actin-based assays

Actin polymerization rates were measured by the change in

fluorescence upon incorporation of pyrene-labeled actin

[22]. Using 10 lM of G-actin supplemented with 5% of

pyrene-labeled actin, polymerization was induced by the

addition of 2 mM MgCl2 and 0.1 M KCl in G-actin buffer

(10 mM Tris/HCl pH 8.0, 0.2 mM CaCl2, 7 mM b-mercapto-

ethanol, 1 mM ATP). The increase of pyrenyl-fluorescence

was monitored in a total volume of 50 lL on a microplate

at 407 nm and 21 °C using a Varian Cary Eclipse spectro-

fluorometer (Agilent Technologies, Santa Clara, CA, USA)

with an excitation wavelength of 365 nm. Actin depolymer-

ization assays were performed with filamentous actin that

was polymerized in the presence of 5% pyrene-labeled

actin. A 10 lM stock solution was diluted to 0.2 lM in

depolymerization buffer (10 mM Tris/HCl pH 8.0, 50 mM

KCl, 1 mM MgCl2, 1 mM EGTA, 2 mM dithiothreitol). The

decrease in pyrenyl-fluorescence was observed in a total

volume of 200 lL on a microplate as described above.

Apparent half-times of polymerization and depolymeriza-

tion time courses were calculated from the rate constants

obtained by approximation of a single exponential function

to the kinetic traces.

ATPase measurements

Intrinsic steady-state ATP turnover of 10 lM F-actin was

measured using an NADH-coupled assay, as described pre-

viously [45]. b-Actin variants were polymerized for at least

4 h at 21 °C and used directly in the assay. ATPase activa-

tion of NM-2A was measured in the absence and presence

of 30 lM F-actin as detailed elsewhere [16].

DNase I inhibition and thermal stability

Binding of actin to DNase I was determined with the

DNase I inhibition assay according to Morrison et al. [46].

Briefly, 0.05–0.75 lM G-actin and 150 nm recombinant

DNase I were preincubated in G-actin buffer for 30 min at

room temperature. The reactions were started in a UV-

transparent 96-well microplate by adding 80 lL of

200 lg�mL�1 salmon sperm DNA in DNA buffer (10 mM

Tris/HCl pH 8.0, 4 mM MgCl2, 1.8 mM CaCl2) into 20 lLof the actin–DNase I preincubation mix. Absorption at

260 nm was recorded for 3 min at 25 °C using a SPEC-

TROstar Omega absorbance microplate reader (BMG Lab-

tech, Ortenberg, Germany). Initial rates were compared

with a reference sample containing no actin. The actin con-

centration that causes 50% of DNase I inhibition (IC50)

was determined by fitting a Hill function to the data set.

For testing thermal stability of the actin variants the

DNase I inhibition assay was modified according to

Sch€uler et al. [18]. Briefly, G-actin was incubated for 5 min

at temperatures between 40 and 70 °C using a temperature

gradient in a PCR thermocycler (SensoQuest, G€ottingen,

Germany). Afterwards, the DNase I inhibition assay was

applied as described above using saturating actin concen-

trations. The temperatures at which inhibition of DNase I

was half-maximal were determined by nonlinear curve fit-

ting using a Boltzmann function and are regarded as mid-

points of the thermal transition into the unfolded state

(Tm).

Nucleotide exchange

Nucleotide exchange rates were determined from the

change in fluorescence after exchange of actin-bound

e-ATP by ATP [21]. First, residual nucleotides in the actin

stocks were removed by buffer exchange with 10 mM Tris/



HCl (pH 8.0) using desalting spin columns. Actin was

then incubated with 100 lM e-ATP for 2 h on ice and the

main excess of e-ATP was subsequently removed by a sec-

ond buffer exchange with 10 mM Tris/HCl (pH 8.0)

including 10 lM e-ATP. The protein concentration was

determined using a Bradford assay (Bio-Rad Protein

Assay; Bio-Rad, Munich, Germany) with BSA as stan-

dard. The e-ATP-bound actin was diluted to 2 lM and

mixed with an equal volume of G-actin buffer including

0.2 mM ATP in a High-Tech Scientific stopped-flow fluo-

rometer. The change in fluorescence at 21 °C was detected

over 25 min with an excitation wavelength of 335 nm and

a 389 nm emission cut-off filter. The signal was averaged

from at least three replicates and fitted to a double expo-

nential decay function to determine the fast and slow

phase of nucleotide exchange.

Microscale thermophoresis

Profilin II in 25 mM Hepes (pH 8.0) was labeled using the

Monolith NT Protein Labeling Kit RED NHS (NanoTem-

per Technologies, Munich, Germany). Labeling efficiency

was tested by comparing absorption at 280 and 650 nm.

The buffer of the b-actin variants was replaced by 25 mM

Hepes (pH 8.0), 20 mM KCl using desalting spin columns.

Actin was prepared as a twofold serial dilution and added

to an equal volume of 0.2 lM labeled profilin (final buffer

conditions: 25 mM Hepes pH 8.0, 10 mM KCl). After

10 min incubation time, the complex was filled into Mono-

lith NT.115 Standard Treated Capillaries (NanoTemper

Technologies) and thermophoresis was measured in a

Monolith NT.015T Microscale Thermophoresis device

(NanoTemper Technologies) at 20 °C, 75% red LED

power and 20% IR-laser power. The actin dependent

change in thermophoresis was described with a Hill func-

tion to determine the apparent dissociation constant Kd of

the actin–profilin complex.

MD simulations

MD simulations were carried out with NAMD 2.9 [25] and

the CHARMM27 force field [47]. The X-ray coordinates of G-

actin in the closed state (2BTF) were used as a starting

structure. Single-point mutations (p.R183W and p.E364K)

were inserted using the SCHR€ODINGER SUITE (2012) and the

proteins were prepared using PROTEIN PREPARATION WIZARD

[27]. N-Terminal acetylation of the protein was removed

and Sr2+∙ATP was replaced by Ca2+∙ATP. The proteins

were fully solvated with the TIP3P explicit water model

[48] and charges were neutralized by adding counter ions to

the system. Simulations were performed in a NpT ensemble

at constant temperature (310 K) and constant pressure

(1 atm) using Langevin dynamics and the Langevin piston

method. Long-range electrostatics were treated with the

particle-mesh Ewald method [49] and a 12 �A cut-off was

used for nonbonded short-range interactions. Prior to pro-

duction runs, the system was energy minimized, the solvent

was equilibrated for 200 ps with positional restraints on the

protein atoms, and subsequently the entire system was

equilibrated for 5 ns. Production runs were conducted for

50 ns per simulation system.

Computational and statistical analysis

Data analysis and graph plotting were performed using

Microsoft EXCEL 2010 (Microsoft, Redmond, WA, USA)

and ORIGIN 8.5 (OriginLab, Northampton, MA, USA).

Errors are given as standard deviations and are based on

three independent determinations (n), if not otherwise spec-

ified. Unpaired two-tailed t-tests were employed for statisti-

cal analysis of kinetic rates and affinity constants using

GraphPad Prism version 5.02 for Windows (GraphPad

Software, San Diego, CA, USA) or Microsoft EXCEL 2010

and the means were regarded as statistically significant if

P < 0.05.

Acknowledgements

This work was supported by the ‘Deutsche Fors-

chungsgemeinschaft’ Grant MA 1081/19-1 (to Dietmar

J. Manstein). The authors thank Hella Scharnhorst for

technical assistance.

Author contributions

Planned experiments: NH, MP, MM; Performed

experiments: NH, MP, OS, MM; Analyzed data: NH,

MP, OS, MM; Contributed reagents or other essential

material: NH, OS, AMA, HGM, DJM, MM; Wrote

the paper: NH, MP, DJM, MM.

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Supporting information

Additional supporting information may be found in

the online version of this article at the publisher’s web

site:Movies S1–S3. Molecular dynamics simulation of wt-

b-actin (S1), p.R183W-b-actin (S2) and p.E364K-b-actin (S3).



Documents

Molecular mechanisms of disease-related human β-actin mutations p.R183W and p.E364K