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FHOD1 is a combined actin filament capping andbundling factor that selectively associates with actinarcs and stress fibers

Andre Schonichen1,*,`, Hans Georg Mannherz1,2, Elmar Behrmann1,§, Antonina J. Mazur2,", Sonja Kuhn1,3,Unai Silvan4, Cora-Ann Schoenenberger4, Oliver T. Fackler5, Stefan Raunser1, Leif Dehmelt6,7 andMatthias Geyer1,3,*1Department of Physical Biochemistry, Max Planck Institute of Molecular Physiology, 44227 Dortmund, Germany2Department of Anatomy and Molecular Embryology, Ruhr University Bochum, 44801 Bochum, Germany3Research Center Caesar, Ludwig-Erhard-Allee 2, 53175 Bonn, Germany4Maurice E. Muller Institute for Structural Biology, Biozentrum, University of Basel, 4056 Basel, Switzerland5Department of Infectious Diseases, Virology, University Hospital Heidelberg, INF 324, 69120 Heidelberg, Germany6Department of Systemic Biology, Max Planck Institute of Molecular Physiology, 44227 Dortmund, Germany7Department of Chemical Biology, Faculty of Chemistry, TU Dortmund, 44227 Dortmund, Germany

*Authors for correspondence (matthias.geyer@caesar.de; andre.schoenichen@gmail.com)`Present address: University of California San Francisco, Department of Cell and Tissue Biology, San Francisco, CA 94143, USA§Present address: Institute of Medical Physics and Biophysics, Charite, Universitatsmedizin Berlin, 10117 Berlin, Germany"Present address: Department of Cell Pathology, Faculty of Biotechnology, University of Wroclaw, PL-51-148 Wroclaw, Poland

Accepted 30 January 2013Journal of Cell Science 126, 1891–1901� 2013. Published by The Company of Biologists Ltddoi: 10.1242/jcs.126706

SummaryFormins are actin polymerization factors that are known to nucleate and elongate actin filaments at the barbed end. In the present studywe show that human FHOD1 lacks actin nucleation and elongation capacity, but acts as an actin bundling factor with capping activitytoward the filament barbed end. Constitutively active FHOD1 associates with actin filaments in filopodia and lamellipodia at the leading

edge, where it moves with the actin retrograde flow. At the base of lamellipodia, FHOD1 is enriched in nascent, bundled actin arcs aswell as in more mature stress fibers. This function requires actin-binding domains located N-terminally to the canonical FH1–FH2element. The bundling phenotype is maintained in the presence of tropomyosin, confirmed by electron microscopy showing assembly of

5 to 10 actin filaments into parallel, closely spaced filament bundles. Taken together, our data suggest a model in which FHOD1stabilizes actin filaments by protecting barbed ends from depolymerization with its dimeric FH2 domain, whereas the region N-terminalto the FH1 domain mediates F-actin bundling by simultaneously binding to the sides of adjacent F-actin filaments.

Key words: FHOD1, Actin bundling, Actin arcs, Stress fibers, Retrograde flow

IntroductionCell movements and dynamic changes in cell morphology play key

roles during tissue regeneration, immune responses, embryonic

development, and wound healing in eukaryotic organisms.

Remodeling of the actin cytoskeleton regulates many of the

associated cellular functions such as protrusion, adhesion, and

retraction. Although a wide array of actin filament assemblies

exists in multicellular organisms, three main categories can be

distinguished that play fundamental roles in cell migration and

cellular morphogenesis: (1) the lamellipodium, a veil-like

membrane protrusion at the leading edge of a migrating cell

containing a branched actin meshwork, (2) filopodia and

microvilli, finger-like protrusions of the plasma membrane,

stabilized by an F-actin bundle of varying thickness, and (3)

actin stress fibers that can form at least three different assembly

categories, such as dorsal stress fibers, actin arcs and ventral stress

fibers (Hotulainen and Lappalainen, 2006; Ridley, 2011).

A rich variety of regulators assembles, maintains and destroys

actin cytoskeletal structures. Among these, formins constitute the

largest and most diverse family of actin polymerization

promoting factors (Chesarone et al., 2010). Fifteen human

formin variants are known, many of which contain additional

splicing isoforms that cluster into eight different families

(Schonichen and Geyer, 2010). The defining element to all

formins is the presence of an FH2 domain that renders the protein

dimeric. For many formins (e.g. mDia1), it has been shown that the

FH2 domain promotes barbed end elongation of straight actin

filaments, most likely supported in combination with the proline-

rich FH1 domain, which concentrates actin–profilin units at the

site of polymerization. Diaphanous related formins (DRFs) contain

an additional array of regulating domains at their N-termini that

constitute a GTPase-binding domain (GBD) for activation, a

canonical FH3 domain composed of armadillo repeats and

additional helical elements. DRFs are negatively regulated by an

intra-molecular interaction of a C-terminal Diaphanous

autoregulation domain (DAD) with the N-terminal FH3 domain

(Alberts 2001; Watanabe et al., 1999), whose release is thought to

be mediated by competition displacement with an activated Rho-

GTPase (Chesarone et al., 2010; Schonichen and Geyer, 2010). In

contrast, activation of the human FH1/FH2 domain-containing

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protein 1 (FHOD1) was shown to require phosphorylation of serine

and threonine residues within the DAD by the Rho-associated

kinase ROCK (Takeya et al., 2008).

De novo actin filament formation requires an initial nucleation

process, i.e. the stabilization of an actin dimer or trimer that acts as

nucleating seed followed by an elongation process, during which

new actin subunits are preferentially added to the fast growing plus

or barbed ends. Actin dimer or trimer formation is kinetically

unfavorable, but strongly supported by dimeric formin FH2

domains. In addition, formins support subsequent elongation

leading to rapid polymerization of straight, unbranched actin

filaments (Chesarone et al., 2010). Based on our current

understanding, formins stay bound to the barbed end of the

growing filament as leaky cappers, where they promote filament

elongation. In cells, this behavior effectively leads to transport of the

formin through the cytoplasm or into protrusions, by transmitting

forces from actin polymerization to the plasma membrane (Romero

et al., 2004; Zigmond et al., 2003). To date, the effect of full length

formins on actin polymerization has not been analyzed in vitro,

mainly due to technical issues to purify or produce the stable,

dimeric ,125 kDa multidomain protein. Instead, only deletion

constructs containing FH1 and FH2 domains of mammalian and

ascomycota formins were investigated.

In cells, FHOD1 has been shown to induce stress fiber

formation in conjunction with either activated RhoA or Rac1

GTPases (Gasteier et al., 2003; Takeya and Sumimoto, 2003;

Westendorf, 2001). Stress fibers are contractile actomyosin

bundles, which play important roles in embryonic development,

wound healing and cell migration (Hotulainen and Lappalainen,

2006). They are composed of numerous, short actin filaments with

alternating polarity (Cramer et al., 1997). These filaments are cross-

linked by a-actinin and possibly also by other actin-bundling

proteins. a-Actinin and myosin display a periodic distribution along

stress fibers typical also for other types of contractile structures, such

as myofibrils of muscle cells. In the present study, we investigated

the function of FHOD1 by performing in vitro actin polymerization

and bundling assays and followed its distribution and movements on

actin-rich structures in living cells with high temporal and spatial

resolution. Based on the functional characterization of FHOD1

domains, a model is presented, which proposes that FHOD1

stabilizes F-actin bundles by protecting barbed ends and connecting

them to adjacent actin filaments, a function that is enforced in the

presence of tropomyosin.

ResultsFHOD1 inhibits actin polymerization in vitro

To explore FHOD1’s actin assembly properties in vitro, several

FHOD1 constructs were generated (Fig. 1A). Besides the

127 kDa full length protein (fl), we generated a truncation

mutant lacking the C-terminal DAD (DC), an N-terminal deletion

mutant composed of the canonical FH1–FH2–DAD domains

(DN), a mutant containing only the N-terminal regulation

domains (N), and the central part connecting the canonical FH3

and FH1–FH2 domains (M). The FH2 domain of mDia1 served

Fig. 1. Effect of FHOD1 activity on actin polymerization in vitro. (A) Schematic illustration of FHOD1 constructs used for in vitro and cell experiments.

(B) Coomassie-stained SDS-PAGE analysis of purified FHOD1 and mDia1 constructs. (C) Negative stains of full length FHOD1 observed in electron

micrographs. (D) Time course of actin polymerization in the presence of 300 nM FHOD1 constructs fl (red), DC (blue) and DN (dark green) as well as mDia1 FH2

(green) as a positive control. All polymerization assays contained 2 mM pyrene-labeled actin. au, arbitrary units. (E) TIRFM of polymerizing 5% Alexa488-

labeled actin (1.5 mM total) alone (top panel), and in the presence of constitutively active FHOD DC (middle panel) and FHOD1 fl (bottom panel).

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as control protein in all actin assembly assays. Although the

mDia1 FH2 domain could be expressed in E. coli, expression of

all recombinant FHOD1 constructs containing the FH2 domain

required the baculovirus expression system, indicating that

FHOD1’s FH2 domain potentially contains unknown structural

elements that require eukaryotic chaperones or posttranslational

modifications for proper maturation. However, all proteins could

be purified to homogeneity (Fig. 1B). Single particle electron

micrographs confirmed the integrity of full length FHOD1 by the

characteristic torus-shaped conformation (Fig. 1C).

We examined actin filament nucleation and elongation activities

of FHOD1 proteins by fluorescence measurements of pyrenyl-actin

(Fig. 1D). At a concentration of 2 mM, actin spontaneously

polymerized at a basal level, while the FH2 domain of mDia1

effectively stimulated actin elongation, similarly as described before

(Li and Higgs, 2003; Romero et al., 2004). In contrast, addition of

0.3 mM FHOD1 either as full length protein, or DC or DN variants,

almost completely inhibited actin polymerization, suggesting

capping activity of this formin (Fig. 1D). The inhibitory effect of

FHOD1 DC and fl on actin polymerization could be confirmed by

total internal reflection fluorescence (TIRF) microscopy, where no

indication of filament formation was seen (Fig. 1E). The N-terminal

FHOD1 domains 1–573 lacking the FH2 domain instead showed no

effect on actin polymerization in the pyrene-actin assay

(supplementary material Fig. S1).

In cells, expression of FHOD1 DC leads to a robust increase in

number and thickness of actin stress fibers, which suggests an

active role of this formin in cellular actin organization (Gasteier

et al., 2003; Schonichen et al., 2006). Thus, our finding that

FHOD1 might be a negative regulator of actin polymerization

was surprising and prompted us to investigate the molecular role

of FHOD1 on actin polymerization in more detail. In time

resolved actin polymerization assays, addition of the proposed

constitutively active FHOD1 DC variant in increasing

concentrations from 1 to 100 nM led to dose-dependent

inhibition of actin polymerization (Fig. 2A). A similar effect

Fig. 2. FHOD1 inhibits F-actin nucleation and elongation activity. (A) Time course of F-actin assembly in presence of increasing concentrations of FHOD1

DC. The F-actin elongation activity of 100 nM mDia1 FH2 is shown as a positive control. Pyrene-labeled actin was used at concentrations of 2 mM.

(B) Observation of actin polymerization over time in presence of 100 nM FHOD1 fl, phosphorylated FHOD1 or FHOD1 fl together with 1 mM DAD peptide.

300 nM mDia1 FH2 was used as a positive control for actin polymerization activity. (C) Time course of the basal actin polymerization activity at 2 mM

concentration with 100 nM FHOD1 DN, 3 mM profilin, and a mixture of 100 nM FHOD1 DN and 3 mM profilin. mDia1 FH2 activity is shown for comparison.

(D) Free barbed end elongation from 0.5 mM pyrenyl-actin and actin seeds in the presence of indicated concentrations of FHOD1 DC. (E) Free barbed end

elongation from 0.5 mM pyrenyl-actin and actin seeds in presence of 3 mM profilin and indicated concentrations of FHOD1 DC. (F) Plot of relative (rel.)

elongation rates (% s21) as a function of FHOD1 DC concentration in the absence or presence of 3 mM profilin. Slopes were taken at 20–60 s of the time courses

displayed in D and E. (G) Plot of F-actin depolymerization by 10-fold dilution in the absence and presence of FHOD1 fl and DC. (H). Concentration series of

FHOD1 DC in F-actin dilution assays. (I) Plot of depolymerization rates (au s21) as a function of FHOD1 DC concentration. Slopes were taken at 30–100 s of the

time course shown in H. au, arbitrary units.

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was observed when full length FHOD1 was applied (Fig. 2B),

indicating its ability to interact with actin despite presence of itsproposed autoinhibitory regulation. Addition of the DAD at 10-fold molar excess over FHOD1 fl to release a potential

autoinhibitory state did not stimulate polymerization, but againinhibited actin polymerization. Phosphorylation of FHOD1 fl byincubation with recombinant ROCK1, which had been shown toactivate the protein in cells by phosphorylation of the DAD

(Hannemann et al., 2008; Takeya et al., 2008), did not increasepolymerization, but suggested enhanced inhibition (Fig. 2B). Inpresence of 3 mM profilin, addition of the FHOD1 FH1–FH2–

DAD domain construct (DN) led again to significantly reducedpolymerization activity (Fig. 2C).

Actin filament elongation experiments were performed atreduced concentrations of 0.5 mM G-actin, which allowed

distinguishing between actin nucleation and elongationcapabilities, as they require the presence of actin seeds for freebarbed end elongation activity (Fig. 2D,E). Increasing

concentrations of FHOD1 DC led to dose-dependent reduction ofthe elongation activity both in the absence or presence of 3 mMprofilin. The inhibitory effect of FHOD1 DC was reduced in

presence of profilin, suggesting that FHOD1 is able to elongateactin filaments by incorporation of actin–profilin complexes(Fig. 2E). This effect was quantified, indicating comparable

exponential reduction of filament elongation by increasingamounts of FHOD1 DC in the presence or absence of profilinalbeit to different base-line activities (Fig. 2F). In conclusion, wecould not detect actin nucleation activity of FHOD1, neither as full

length protein nor in the truncated form lacking the autoregulationdomain, but found weak elongation activity in presence of profilin.

As FHOD1 was found to inhibit actin polymerization and

elongation, we wondered if FHOD1 is an actin filament cappingprotein. We thus examined FHOD1 fl and DC in F-actin dilutionassays and found that both proteins inhibited F-actindepolymerization (Fig. 2G,H). FHOD1 fl showed weaker

inhibition at 300 nM concentration than FHOD1 DC suggestingthat autoinhibition regulates association with F-actin barbed ends(Fig. 2G). A continuous decrease in the F-actin depolymerization

activity was observed upon increasing concentrations of FHOD1DC (Fig. 2H) as seen as well from the quantified depolymerizationrates (Fig. 2I).

FHOD1 bundles actin filaments

Activation of FHOD1 in cells induces stress fibers (Gasteier et al.,2003; Schonichen et al., 2006; Takeya and Sumimoto, 2003).

Because stress fibers contain F-actin bundles, we tested howFHOD1 binds to F-actin and if it is sufficient to induce F-actinbundles in vitro. High speed centrifugation assays revealed thatall tested FHOD1 constructs (fl, DC, DN, N and M), as well as

mDia1 FH2 appeared in the pellet fraction, and thus bind to F-actin (Fig. 3A). These results confirmed previous studies thatsuggested F-actin binding activities of an N-terminal FHOD1

construct lacking the FH2 domain (Takeya and Sumimoto, 2003).As FHOD1 N and DN do not overlap significantly in primarysequence, this observation suggests that two independent F-actin

binding sides are present in FHOD1. Interestingly, while FHOD1fl and DC bound F-actin very efficiently, only a small fraction ofconstructs FHOD1 M, DN and N and mDia1 FH2 was present in

the pellet fractions. In case of FHOD1 DN and the control proteinmDia1, this finding can be explained by binding exclusively tobarbed ends via the FH2 domain shared in these constructs.

Fig. 3. FHOD1 binds to and bundles F-actin in vitro. (A) Coomassie-

stained SDS-PAGE of FHOD1 constructs assayed in an F-actin binding assay.

FHOD1 constructs as indicated were incubated with F-actin and co-

sedimentations at high-speed centrifugation were analyzed. The presence of

FHOD1 in pellets (P) indicates F-actin binding, while proteins remaining in

the supernatants (S) do not bind F-actin. (B) Coomassie-stained SDS-PAGE

analysis of FHOD1 constructs tested for F-actin bundling activity after

incubation of proteins with F-actin and co-sedimentation at low speed. Actin

in pellet fractions indicates F-actin bundles. mDia1 FH2 served as a negative

control. (C) TIRFM of 5% Alexa488-labeled actin filaments (1.5 mM total)

only (top panel), and upon addition of 100 nM FHOD1 DC (middle panel), or

addition of 100 nM FHOD1 fl (bottom panel); see supplementary material

Movie 1. (D) Electron micrographs of FHOD1-induced actin filament

bundling. Actin filaments were incubated overnight on ice with the indicated

additives and negatively stained for EM display. Both the constitutively active

mutant (FHOD DC)- and full length FHOD-induced bundling of actin

filaments are independent of capping through gelsolin or tropomyosin

decoration. Scale bars: 1,000 nm for images and 200 nm for insets.

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Because the number of barbed ends is significantly smaller thanthe number of potential binding sites along actin filaments, the

amount of FH2 domain binding in the pellet fraction is expectedto be smaller. However, the independent F-actin binding site inthe FHOD1 N-terminus might be able to bind to the sides offilaments. Additional barbed-end binding or dimerization via the

FH2 domain could increase overall actin binding efficiency.Indeed, compared to FHOD1 N and M, significantly moreFHOD1 fl and DC was pulled down by F-actin, indicating that

two regions of FHOD1 can act synergistically to increase F-actinbinding efficiency.

The F-actin bundling activity of FHOD1 constructs was next

analyzed using low speed centrifugation. The data obtainedshowed that FHOD1 fl and particularly FHOD1 DC inducedsedimentation of F-actin (Fig. 3B). Low-speed sedimentation ofactin in the presence of constructs M and DN (at the limited

amounts available) was comparable to negative controls,suggesting that those N-terminal domains themselves are notable to bundle F-actin (Fig. 3B). These data suggest that two

independent actin-binding sites at the N- and C-terminal domainsof FHOD1 cooperate for actin filament bundling. Whereas the N-terminus is likely to have F-actin side-binding activity, barbed-

end binding and/or protein dimerization via the FH2 domain isrequired for the formation of stable higher-order structurescomposed of FHOD1 and actin bundles.

To examine the capability of FHOD1 for actin bundling we

applied TIRF microscopy using polymerizing actin filaments.Two FHOD1 proteins DC and fl were added at similarconcentrations to polymerizing actin filaments using actin alone

as control. Formation of thick F-actin bundles and a rigid F-actinnetwork was indeed observed upon FHOD1 addition as actinfilaments stopped fluctuating (Fig. 3C; supplementary material

Movie 1). This effect was much stronger for FHOD1 DCcompared to fl, indicating that release of the DAD from the N-terminal FH3 recognition domain is required for full bundling

activity.

In addition we tested F-actin bundling by FHOD1 usingelectron microscopy (EM) after negative staining. FHOD1 DCand fl were incubated with freshly polymerized F-actin at molar

ratios of 1:1, 1:10, 1:100 and 1:1,000 (FHOD1 versus actinmonomer concentration) for varying periods of time. F-actin waspolymerized in the presence of the barbed end capper gelsolin at

100:1 ratio to obtain filaments of similar lengths. The actinconcentration was set to 0.1 mg/ml. Furthermore, we employedF-actin decorated with skeletal muscle tropomyosin at a molar

ratio of 7:1 in some experiments. In presence of FHOD1 fl wedetected normal appearing actin filaments and occasionallybundles comprising 2 to 3 actin filaments and in between noisysprinkles and torus-shaped structures, most probably indicating

free FHOD1 (Fig. 3D, lower panels). Otherwise no indicationwas obtained for an alteration of the filament assembly andshape. In contrast, FHOD1 DC induced F-actin bundles

containing about 2 to 5 parallel actin filaments at ratios of 1:10and 1:100 after overnight incubation at room temperature(Fig. 3D, middle panels). Upon addition of gelsolin, the actin

filaments appeared shortened, potentially due to the increasingnumber of barbed end cappers, while the bundle thicknessremained similar. FHOD1 also bundled F-actin decorated with

tropomyosin as indicated by an increase in the length andthickness of F-actin bundles compared to the treatment withgelsolin only (Fig. 3D, right panels). This suggests that the

F-actin side-binding capacity of FHOD1 is maintained uponfilament stabilization with tropomyosin.

The N-terminus of FHOD1 is necessary for localization tostress fibers

Our in vitro data suggest that FHOD1 does not nucleate andassemble actin filaments de novo as other formins but ratherbundles and stabilizes existing actin filaments. We next sought to

investigate the biological significance of these in vitro activities.Previous studies applied speckle microscopy for detection ofindividual molecular complexes of related actin regulators, suchas mDia and the Arp2/3 complex (Higashida et al., 2004;

Miyoshi et al., 2006) as individual ‘speckles’ (Danuser andWaterman-Storer, 2006; Watanabe et al., 1999). We used thismethod to detect potential enrichment of several FHOD1

constructs in subcellular structures and to study its dynamiclocalization in living cells, and focused our analysis on the actin-rich cell cortex by using TIRF microscopy. We therefore

expressed FHOD1 constructs at low levels in two mammaliancell lines: (1) Neuro-2a neuroblastoma cells expressing dominantpositive Rac1, for robust formation of large, dynamic

lamellipodia and filopodia, along with less dynamic actin arcs,and (2) COS7 fibroblast-like kidney cells, which form smaller,dynamic lamellipodia along with less dynamic ventral stressfibers terminating in focal adhesions.

In both cell types, wild-type FHOD1 co-localized strongly

with actin structures known to form anti-parallel bundles: actinarcs and ventral stress fibers (Fig. 4A,B). This localizationpattern is even more pronounced for the dominant active DC

fragment. In contrast, FHOD1 DN, which contains the proposedbarbed-end binding FH1–FH2 domains, did not show anyparticular enrichment at the actin-rich cell cortex. The N-

terminal fragment 1–573, which lacks the actin barbed-endbinding FH1–FH2 domains, also strongly localized to actin arcsand ventral stress fibers. Taken together, our data suggested thatthe N-terminal region is sufficient to confer selective binding of

FHOD1 to actin-arcs and stress fibers.

Speckle microscopy reveals contrasting dynamics ofFHOD1 and mDia1 in living cells

Constitutively active mutants of the related formin mDia1 are

highly mobile in living cells due to their processive barbed-endcapping activity. Because FHOD1 elongates actin filaments onlypoorly in vitro, we examined the dynamic behavior of individual

FHOD1 DC speckles to test, whether this constitutively activemutant shows typical motility of a processive cap in living cells.It should be noted here that the FHOD1 DN mutant is notexpected to act as a constitutively active mutant, as it is lacking

the essential N-terminal F-actin binding site. mDia1 lacks suchN-terminal F-actin binding site, and a DN mutant is widelyaccepted to be constitutively active.

Kymograph analyses in lamellipodia showed that FHOD1 DC

speckles were distributed sparsely throughout the lamellipodium,where they were predominantly shifted laterally at the speedand in the direction of retrograde flow (Fig. 5A,B, left;

supplementary material Movies 2 and 3, upper panel),suggesting that FHOD1 DC interacts with preformed actinfilaments either by binding at the side of filaments, by capping

filament ends or both. FHOD1 DC speckles localized to actin arcsat the base of the lamellipodium, which was characterized bymuch slower retrograde movements (supplementary material Fig.

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S2; Movie 4). In all actin-rich regions, most FHOD1 DC speckles

were only transiently detected for one single 400 ms frame

(supplementary material Movie 3) suggesting that cortex-

associated FHOD1 DC exchanges rapidly with a cytosolic pool.

In contrast, the dynamic behavior of mDia1 DN speckles was

highly divergent from FHOD1 DC speckles. mDia1 DN did not

translocate with the retrograde flow towards the cell interior, but

rather in the opposite direction towards the leading edge

(anterograde) at much faster speeds (Fig. 5A,B, right;

supplementary material Movies 2 and 3, lower panel). As an

apparent consequence of actin filament elongation and its

processive capping function, mDia1 DN accumulated at the

very leading edge of the cell. This is in contrast to FHOD1 DC,

which accumulated at the base of the lamellipodium. Thus, our

observation, that FHOD1 does not preferentially bind to cellular

locations enriched in barbed ends, but rather interacts transiently

with preformed actin structures, suggested that FHOD1 does not

primarily function as a processive actin filament capper.

While we never observed directional, anterograde

translocation of FHOD1 DC speckles, a small subpopulation of

FHOD1 DC speckles accumulated transiently at the leading edge

of migrating cells. In a few cases, we even observed a strong

accumulation of FHOD1 DC speckles at this cellular region

(supplementary material Fig. S2; Movie 5). However, the

dynamics of the majority of FHOD1 DC speckles was in

agreement with a role for FHOD1 as an actin filament side-

binding and capping protein, which is particularly associated with

actin filament bundles in actin arcs and stress fibers.

DiscussionIn the present study we show that human FHOD1 does not de

novo polymerize actin filaments, but instead binds dynamically to

and bundles actin filaments throughout the cell. The canonical

FH2 domain exhibits a capping activity on the actin filament

barbed end, whereas the central helical region in between the

FH3 and FH1 domains interacts with sides of actin filaments.

Given that FHOD1 DC has the strongest effect on F-actin

bundling suggests that this activity is regulated by the interaction

of the DAD with the FH3 domain. In cells, activation of FHOD1

has been shown to require phosphorylation by ROCK1

(Hannemann et al., 2008; Takeya et al., 2008). In contrast, an

activation mechanism via binding to a lipidated Rho-GTPase, as

e.g. described for mDia1 (Watanabe et al., 1999), that would lead

to retention of the formin at the plasma membrane, is not

Fig. 4. Localization of wild-type and mutant FHOD1 at cellular actin structures. Two cell systems were used to study the subcellular localization of FHOD1.

(A) Neuro2a cells expressing constitutively active Rac1 form extensive lamellipodia, filopodia and actin arcs. (B) COS7 cells also form ventral stress fibers. If

expressed at very low levels using a truncated CMV promoter (DCMV), wild-type FHOD1 speckles localize to various actin-rich structures including filopodia

and actin arcs in Neuro2a cells and on a subset of stress fibers in COS7 cells (cyan arrows). Constitutively active FHOD1 DC is enriched on actin arcs in Neuro2a

cells and strongly localized to stress fibers in COS7 cells. The N-terminal region of FHOD1 1–573 is both necessary and sufficient for its localization to actin

stress fibers, while FHOD1 DN is largely cytosolic. Wild-type FHOD1, as well as its DC variant and the 1–573 regions are largely excluded from actin-rich focal

adhesions (cyan arrowheads). The corresponding FHOD1 image panels within each cell type were all acquired and scaled with identical settings to allow

comparison of expression level. Insets are magnified 36 from original panels and optimized individually for display purposes.

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observed for FHOD1 (Schulte et al., 2008). Instead, activated

FHOD1 transiently associates with various actin structures and

ultimately accumulates in actin arcs and stress fibers, where it

could protect the filament strain from depolymerization while

simultaneously binding to neighboring actin filaments. FHOD1

thus represents a new class of actin regulators by combining

capping activity at the barbed end with F-actin side-binding

activity to effectively protect actin filaments in cytoskeletal

structures.

Formins were identified as de novo actin filament nucleators

and are often referred to as actin nucleating factors in the current

literature. However, we could not confirm a ‘formin-typical’

Fig. 5. Intracellular dynamics of FHOD1 and

mDia1 formins. (A) Kymographs (right-hand panel)

show that many speckles of the constitutively active

FHOD1 DC formin undergo short but continuous

retrograde movements at speeds similar to retrograde

actin flow within the lamellipodium (supplementary

material Movie 2, upper panel). In contrast, speckles

of constitutively active mDia1 DN either remain at

the very leading edge or appear transiently for a

single image frame within the lamellipodium

(supplementary material Movie 2, lower panel).

(B) At higher frame rates, continuous anterograde

movements of mDia1 DN toward the leading edge

can be resolved, which were similar in speed to

previous reports on single molecule dynamics of this

formin mutant (Watanabe and Mitchison, 2002). Such

fast anterograde movements are not observed for

constitutively active FHOD1 DC (supplementary

material Movie 3). (C) Model for FHOD1-mediated

actin filament bundling and stress fiber stabilization.

FHOD1 is activated at the plasma membrane through

phosphorylation of its DAD by the Rho-associated

kinase ROCK1. Activated FHOD1 associates with

both barbed ends and actin filament sides and

assembles into actin arcs and stress fibers. Other

cytoskeleton stabilizing factors, such as a-actinin,

myosin II and gelsolin, might complement the

capping and bundling activity of FHOD1.

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activity for FHOD1, but found that FHOD1 inhibits de novo actinnucleation in vitro. FHOD1 does not seem to be an outlier in this

respect as some formins inhibit rather than increase actinpolymerization in vitro, including FHOD3 (Taniguchi et al.,2009), and FMNL2 (Block et al., 2012). Hence, nucleation ofactin filaments in vitro does not appear to be a common property

of formins. In fact, it is emerging that actin nucleation activity offormins may involve additional cellular co-factors: The yeastformin Bni1 interacts via the FH2 domain with the polarity factor

Bud6 (Moseley and Goode, 2005; Tu et al., 2012), the drosophilaformin Cappuccino and Spire cooperate for actin nucleation(Quinlan et al., 2007), and mDia1 synergizes with the APC

(adenomatous polyposis coli) protein for actin nucleation(Breitsprecher et al., 2012; Okada et al., 2010). Thus, wecannot rule out that FHOD1 may associate with an additionalfactor for actin nucleation in cells. FHOD1 has been shown not

only to be activated by but also to interact with ROCK1 via theN-terminal half of its FH2 domain (Hannemann et al., 2008), andthus ROCK1 may be a candidate to potentially cooperate in actin

filament nucleation.

It has been reported that the FH2 domain of certain formins,such as mDia2, is able to bundle actin filaments in vitro (Harris

et al., 2006). Although mDia2 indeed stabilizes cortical actin andfree actin filament barbed ends in focal adhesions (Gupton et al.,2007), it remains unclear whether actin filament bundling activity

by the FH2 domain is relevant in cells. However, for the FHOD1FH2 domain we neither observe actin filament bundling activityin vitro nor does it localize to F-actin in cells. In contrast, N-terminal binding regions of FHOD1 are required for in vitro

bundling activity, which is also required for F-actin localization.Hence, our data suggest that the mechanisms for actin filamentsbundling of FHOD1 and other formins differ strongly.

Live-cell imaging using EGFP–FHOD1 and mCherry–actinrevealed that FHOD1 localized near the plasma membrane toassociate with cortical F-actin. Interestingly, for a close

homologue to FHOD, ForC from Dictyostelium discoideum, theN-terminal putative GTPase binding domain was shown tointeract with phosphoinositides in vitro and was required forForC targeting to cell–cell contacts and early phagocytic cups in

cells (Dames et al., 2011). Similarly to FHOD1, this domainexhibits the ubiquitin superfold, and its appearance in domainarchitecture and structure is reminiscent to the F0 domain in

Kindlin and Talin proteins that is known to target FERM proteinsto focal adhesions (Legate et al., 2011; Perera et al., 2011).Deletion of the N-terminal region 1–115 in FHOD1 abrogates its

capability to stabilize actin fibers (Schulte et al., 2008) indicatingthat association to the plasma membrane might be required forFHOD1 function. Of note, ForC does not contain a proline-rich

FH1 domain that could recruit profilin–actin complexes for actinpolymerization, indicating a loss of actin polymerization activityin ForC, and supporting properties of FHOD1 described in thepresent paper.

The F-actin bundling mechanism identified in the presentstudy fits well to the known biological properties of FHOD1.Similar to other diaphanous formins, activated forms of FHOD1

potently induce the formation of F-actin stress fibers inmammalian cells (Gasteier et al., 2003; Schulte et al., 2008;Westendorf, 2001). As a notable difference to other DRFs, these

stress fibers are significantly thicker and FHOD1 remainsassociated with these filament bundles to decorate the stressfibers. This phenomenon can now be attributed to FHOD1’s

unique properties such as sustained barbed end association andfilament bundling. Moreover, the mobility of filament-associated

FHOD1 away from the plasma membrane observed by live cellimaging explains why FHOD1 specifically facilitates theformation of cytoplasmic F-actin structures such as stress fibersand actin arcs. Another common feature of formins consists of

their ability to induce the nuclear transcription serum responsefactor (SRF) to modulate the expression of specific target genes(Tominaga et al., 2000). This effect mirrors formin-mediated

regulation of actin dynamics since enhanced actin polymerizationliberates myocardin-related transcription factor (MRTF)cofactors from G-actin to facilitate their import into the nucleus

and subsequent activation of transcription (Jurmeister et al.,2012; Staus et al., 2011). As activated FHOD1 acts as potentinducer of SRF-mediated transcription (Gasteier et al., 2003;Westendorf, 2001) without inducing de novo actin

polymerization, our results indicate that reducing G-actin pools,e.g. by stabilization and/or bundling of F-actin structures, cansuffice for the induction of transcriptional responses.

ROCK1 activation leads to increased actin stress fiberformation via myosin light chain phosphorylation (Wojciak-Stothard et al., 2001). However, whether FHOD1 and myosin

light chain phosphorylation act cooperatively or independently toincrease actin stress fibers and whether they regulate differentarchitectures of bundled actin filaments is unknown. FHOD1

may stabilize and bundle assembled stress fibers, while myosinmotors generate force for contraction. Alternatively, FHOD1could elongate barbed ends with its FH2 domain while the N-terminal domains stay bound to adjacent actin filaments for

anchoring the formin. As FH2 domains can generate piconewtonforces, this could be an alternative mechanism for contraction(Kovar and Pollard, 2004).

The filament stabilizing function ascribed to FHOD1 in thepresent study is in line with recent analyses of FHOD1 (Mi-Miet al., 2012) and FHOD3 (Iskratsch et al., 2010; Taniguchi et al.,

2009) on the regulation of actin assembly and sarcomereorganization in muscle cells. A specific splice variant ofFHOD3 was shown to be located mainly in the Z disc of themature heart muscle and may thus assist actin turnover or

reduced barbed end depolymerization (Iskratsch et al., 2010). Inmature myofibrils, both pointed and barbed ends of the thinfilaments are capped and stabilized by tropomodulin and CapZ,

respectively. While the mechanism of thin filament assembly andmaintenance are still under debate (Sparrow and Schock, 2009),FHOD3 as a barbed end capping protein could adopt a function

similar to CapZ in muscle cells. Notably, FHOD3 is about 250amino acids longer in size than FHOD1. Whereas the GBD–FH3and FH1–FH2 elements align overall well between both

homologues, it is particularly the connecting region identifiedin the present study as F-actin side binding element (construct M,Fig. 1A) that varies substantially between both proteins. Thissuggests that the mode of bundling to neighboring filaments by

this region in FHOD1 could adopt an alternative function in theassembly mechanism mediated by its homologue FHOD3, e.g. incardiomyocytes and striated muscle cells.

On the basis of these results we propose a model, in whichFHOD1 selectively stabilizes actin filaments by associating to thegrowing actin filament at the side of filament polymerization and

travelling with the filament into cell shape maintainingcytoskeletal substructures (Fig. 5C). FHOD1 could sustain actinarc formations at nascent focal adhesions that were recently

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proposed to mediate cellular protrusions (Burnette et al., 2011),

or associate to stress fibers, which are highly decorated with

myosin II and tropomyosin (Tojkander et al., 2011). The

combination of capping activity at the filament barbed end with

F-actin side-binding activity appears complementary to other

bundling factors as a-actinin or capping factors as gelsolin,

whose expression levels are however much higher in eukaryotic

cells. These distinct cross-linkers could associate transiently to

the actin filaments to orchestrate the reorganization of stress

fibers. The diverging regulatory pathways that control these

cross-linkers are integrated into the filament reorganization

process and may allow for the cytoskeletal dynamics and

plasticity. With these features, the mammalian formin FHOD1

represents a new class of actin regulators to effectively protect

and bundle actin filaments.

Material and MethodsCloning, expression and purification of recombinant FHOD1

Detailed protocols of plasmid cloning, protein expression and purification ofrecombinant FHOD1 are provided in the supplementary material. Briefly, plasmidsencoding human FHOD1 1–573, 1–377 and 396–573 as well as mouse mDia1748–1175 were transformed into T1 phage resistant BL21(DE3) cells, expressed at20 C for 14 h and purified following standard protocols. FHOD1 constructscontaining the FH2 domain (FHOD1 fl, DC and DN) were expressed in Sf21 cellsusing a baculovirus expression system and purified as N-terminally His-taggedfusion proteins.

Plasmids encoding human FHOD1 1–573, 1–377 and 396–573 as well as mousemDia1 748–1175 were transformed into T1 phage resistant BL21(DE3) cells andexpressed at 20 C for 14 h after induction with isopropyl-b-D-1-thiogalactopyranoside. Cells were harvested by a 20 min centrifugation step at4,500 g, resuspended in buffer A (50 mM Hepes pH 7.6, 150 mM NaCl, 5 mMEDTA, 1 mM DTE, 1 mM PMSF) and lysed using a microfluidizer. The lysatewas cleared by centrifugation at 10,000 g for 45 min. GST fusion proteins werepurified using GSH Sepharose FastFlow (GE Healthcare) using Akta Prime FPLCsystems. The resin was washed with buffer A and B (50 mM NaPi pH 8, 1 MNaCl, 5 mM EDTA, 1 mM DTE, 1 mM PMSF) until A280 reached zero. GSTfusion proteins were eluted with buffer A containing 10 mM GSH. Fractionscontaining the fusion protein were pooled and treated with TEV protease (home-made) in a molar ratio of 1:200 at 4 C overnight.

FHOD1 constructs containing the FH2 domain (FHOD1 fl, DC and DN) wereexpressed in Sf21 cells (Invitrogen) using the pFastBac baculovirus expressionsystem (Invitrogen). Sf21 cells were transfected using Lipofectamine (Invitrogen)with plasmids encoding N-terminally His-tagged fusion proteins. After virusamplification, Sf21 cultures were infected with concentrated virus stock andproteins were expressed for 72 h at 27 C. Cells were harvested by centrifugation at2,500 g for 20 min at 4 C, and lysed in buffer C (50 mM Hepes pH 8, 500 mMNaCl, 20 mM imidazole, 5 mM b-mercaptoethanol, 1 mM PMSF) by sonication.The cleared lysate was applied to Ni-NTA resin (Qiagen). The resin was washedwith buffer C until the OD280 reached zero. Proteins were eluted by running agradient from 20 to 250 mM imidazole in buffer C. Peak fractions were analyzedby SDS-PAGE and fractions containing the proteins of interest were pooled,concentrated and treated with TEV protease.

After protease cleavage, the protein solution was concentrated at 3,500 g usingAmicon Ultra-15 centrifugation devices with the appropriate pore size (Millipore).2 ml protein solutions were applied to a Superdex S75 16/60 column (GEHealthcare), equilibrated in buffer D (20 mM Hepes pH 7.5, 50 KCl, 1 mMMgCl2, 1 mM EGTA, 1 mM DTT) with a GSH or Ni-NTA column attached inseries. FHOD1 proteins eluted at its expected monomer or dimer (constructsencompassing FH2 domains) weight. Homogeneity and identity of FHOD1proteins were determined by SDS-PAGE analysis and eventually electrospraymass spectrometry, respectively. Proteins containing the FH2 domain wereimmediately studied in pyrene assay measurements or stored at 4 C for less thanone week. All other proteins were stored at 280 C. Profilin was expressed as His-tagged protein, purified by standard Ni-NTA chromatography and stored at280 C. For speckle microscopy, FHOD1 constructs were subcloned into a pEGFP-based vector (Clontech), which was modified to contain a minimal CMV promoter(DCMV) (Watanabe and Mitchison, 2002).

Actin polymerization experiments

Actin nucleation and F-actin elongation assays were performed as described(Moseley et al., 2004). All proteins investigated were carefully dialyzed or gelfiltrated in buffer B to avoid artifacts due to buffer differences. Pyrene-labeledactin was purchased from Cytoskeleton Inc., and dissolved to a concentration of

20 mg/ml. 5 ml aliquots were stored at 280 C. After thawing, one aliquot wasdiluted in 225 ml G buffer and actin as depolymerized by incubation at 23 C for1 h. This aliquot was then centrifuged for 1 h at 80,000 g using a Beckman OptimaTL-100 and a TLA-100 rotor at 4 C to get rid of actin nuclei. 200 ml of thesupernatant containing 10 mM monomeric actin were taken carefully for actinassembly assays.

In polymerization experiments, monomeric pyrene-actin was used at a finalconcentration of 2 mM. After a 3 min incubation step with ME buffer (1 mMMgCl2, 1 mM EGTA) to exchange actin bound Ca2+ against Mg2+, proteins ofinterest in buffer B were added and reactions were started with 106KMEI. 100 mlof these mixtures were quickly applied to an UV cuvette and pyrene fluorescence(excited at 365 nm, emitted at 407 nm) was recorded.

For actin filament elongation assays, G-Mg buffer (G buffer with MgCl2 insteadof CaCl2), F-actin, buffer B with or without protein, and 106KMEI were mixed.The solution was sheared by passaging five times through a 27-gauge needle inorder to generate actin seeds. An invariable volume of this solution was mixedwith pre-incubated pyrenyl-actin and ME. 100 ml of this mixture were transferredinto a fluorescence cuvette. Pyrene fluorescence was monitored immediately. Finalconcentrations of actin and actin seeds used in these assays were 0.5 mM and333 nM, respectively.

In F-actin dilution assays, 10 mM actin (comprised with 5% pyrenyl-actin) waspolymerized by addition of 16 KMEI for 1 h at room temperature. F-actin was1:10 diluted into G buffer with 16KMEI in absence and presence of FHOD1 fland FHOD1 DC. The decay in fluorescence was monitored for 10 min.

For TIRFM, coverslips were gently cleaned with ethanol, Milli-Q water,Hellmanex II (Fluka, NO. 61257), and rinsed in Milli-Q water. Clean coverslipswere subsequently incubated with PEG-silane (MPEG-SIL-5000, 0.1 mg/ml in96% Ethanol) for 12 hours at room temperature. PEGylated coverslips were thenrinsed in Milli-Q water, air dried and stored at 4 C until use. Polymerizationexperiments were carried out in a Leica microscope (DMI 6000B) using 10 to 15%labeled G-actin (A488 rabbit muscle actin, Invitrogen). Images were takenevery 5 seconds using an EMCCD camera (Hamamatsu C9100-02), with theexcitation laser being shuttered between images to avoid photo-bleaching.Polymerization was started by the addition of 26 TIRF polymerization buffer(20 mM imidazole, 100 mM KCl, 2 mM MgCl2, 2 mM EGTA, 0.4 mM ATP,100 mM DTT, 2% methylcellulose, 10 mg/ml glucose oxidase, 50 mg/mlcatalase, 250 mg/ml glucose, pH 7.2–7.4). After the addition of the TIRFpolymerization buffer the sample was incubated 2 minutes on ice before it wasloaded onto the coverslip.

F-actin binding and bundling assay

Actin was isolated from acetone-dried powder prepared from rabbit skeletalmuscle as described (Mannherz et al., 2007). Actin was first depolymerized at5 mM on ice for 1 h in G buffer (5 mM Tris/HCl, pH 8.0 and 0.2 mM CaCl2,0.2 mM ATP, 0.5 mM DTT). After a centrifugation step (14,000 rpm, 30 min)actin was polymerized by addition of 106KMEI (500 mM KCl, 20 mM MgCl2,10 mM EGTA, 100 mM imidazole) for 2 h at 23 C. 5 mM phalloidin (Sigma) wasadded to the assay in order to stabilize F-actin. F-actin was added to the solutionscontaining proteins of interest using cut 200 ml tips and incubated 30 min at 23 C.After that, reaction mixtures were centrifuged at 80,000 g at 4 C for 20 min in aBeckman Optima TL-100 bench top ultracentrifuge using a TLA-100 rotor. 80 mlof the supernatants were taken and dried in a SpeedVac. The remainingsupernatant was removed and pellets were washed briefly with 100 mlpolymerization buffer. To both supernatants and pellets, 20 ml of 16 SDSsample buffer was added. 10 ml of the samples were applied to and analyzed byCoomassie-stained SDS-PAGE.

F-actin bundling assays were performed as described in the actin bindingsection. The only difference affected the centrifugation step after mixing of actinand probed proteins. Here, samples were applied to 10,000 g for 20 min at 4 Cusing a bench top centrifuge. In addition, F-actin bundling was probed by chemicalcross-linking using p-NN9-phenylene-bis-maleimide (PBM) as described (Knightand Offer, 1978) and confirmed by SDS-PAGE analysis performed as given(Hesterkamp et al., 1993). The PBM-generated ‘lower dimer’ obtained by cross-linking F-actin bundles obtained in the presence of 50 mM MgCl2 was purified bygel filtration as described (Silvan et al., 2012).

Electron microscopy

For EM analysis actin was polymerized by addition of 2 mM MgCl2 and diluted to0.1 mg/ml (2.4 mM) in polymerization buffer. Depending on the experimentalconditions gelsolin and tropomyosin were added at molar ratios of 1:100 and 1:7,respectively and incubated for 15 min at RT. Additionally, either FHOD1 DC orfull length FHOD1 at a concentration of 240 nM or 24 nM were added.

After 5 h or overnight incubation on ice, samples were prepared for electronmicroscopy similarly as described (Ohi et al., 2004). Briefly, a 3 ml drop of samplesolution was adsorbed for 45 sec to a freshly glow-discharged copper grid (AGARScientific) covered by a thin, continuous carbon film. Excess sample was blottedusing filter paper (Whatman No.4), washed twice with sample buffer and stainedfor 30 sec with 0.75% uranyl formate. Specimens were imaged at room

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temperature using a JEM1400 electron microscope (JEOL) equipped with a LaB6

electron source. The microscope was operated at an acceleration voltage of120 kV. Images were taken at a calibrated magnification of either 6,6606 or53,8006 and recorded on a 4k64k TemCam-F416 CMOS camera (TVIPS).

Life cell imaging and speckle microscopy

COS7 cells (ATCC, Teddington, UK) and Neuro-2a cells (DMSZ, Heidelberg)were cultured using standard conditions. For microscopy, 50,000 cells were platedon individual glass bottom dishes (MatTek Corp., Ashland) and transfected usingFugene-6 (Roche, Mannheim) on the following day according to themanufacturer’s protocol. The following day, cells were imaged via TIRFmicroscopy in CO2-independent growth medium containing serum and HEPES(Pan Biotech, Aidenbach) at 35 C in a home-built incubation chamber. For TIRFmicroscopy, an Olympus IX-81 microscope, equipped with a Plan APO 606 oilimmersion TIRF microscopy objective (NA51.45) and 1.6 optovar, was used. Atriple bandpass dichroic mirror (U-M3TIR405/488/561, Olympus, Hamburg) wascombined with Semrock Brightline emission filters (HC 520/35 and HC 629/53,AHF Analysentechnik, Tubingen) and CellR diode lasers 488 (20 mW), 561(25 mW). For detection, an EMCCD camera (C9100-13, Hamamatsu, Herrschingam Ammersee) was used at maximal gain. All microscope components werecontrolled by the CellR software (Olympus, Hamburg). For speckle microscopy,cells that express very low amounts of the FHOD1 constructs were selected. Inthese conditions, exposure times of at least 800 ms were required to visualizeindividual speckles. Image processing was performed using ImageJ (NIH,Bethesda) and Photoshop (Adobe, Munchen). Image manipulations were limitedto cropping, scaling, rotation, adjustment of levels, and addition of clearlyidentifiable labels or symbols. To correct for lateral drift in the stage position, animage stabilizer plugin for ImageJ was used (K. Li, ‘The image stabilizer pluginfor ImageJ’, http://www.cs.cmu.edu/,kangli/code/Image_Stabilizer.html,February 2008). The intensity levels in all image panels in time series areadjusted identically.

AcknowledgementsWe thank Karin Vogel-Bachmayr and Diana Ludwig for experttechnical assistance and Sascha Gentz for synthesis of the DADpeptide. In addition, we thank Naoki Watanabe (Tohoku University)for supplying the DCMV expression plasmid.

Author contributionsA.S. and M.G. designed the study. A.S. produced proteins and, withthe help of S.K., performed and analyzed in vitro actin assemblyexperiments. H.G.M. and A.J.M. performed cell transfectionexperiments. H.G.M., U.S. and C.A.S. did in vitro actinpolymerization TIRF microscopy. E.B., H.G.M. and S.R.performed electron microscopy measurements. L.D. performed andanalyzed live cell imaging. O.T.F. contributed with reagents anddiscussions. A.S. and M.G. wrote the manuscript with the help of allother authors. All authors discussed the results and commented onthe manuscript.

FundingThis work was supported in part by the DeutscheForschungsgemeinschaft [grant numbers GE-976/4 to M.G., FA-378/6 to O.T.F., MA-807/19 to H.G.M., and RA-1781/1 to S.R.]; andby the Fonds der Chemischen Industrie (stipend to E.B.).

Supplementary material available online at

http://jcs.biologists.org/lookup/suppl/doi:10.1242/jcs.126706/-/DC1

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