Actomyosin dynamics drive local membranecomponent organization in an in vitro activecomposite layerDarius Vasco Köstera, Kabir Husaina, Elda Iljazia, Abrar Bhata, Peter Bielingb,1, R. Dyche Mullinsb, Madan Raoa,c,and Satyajit Mayora,d,2

aNational Centre for Biological Sciences, Tata Institute for Fundamental Research, Bangalore 560065, India; bCellular and Molecular Pharmacology,University of California, San Francisco, CA 94143; cRaman Research Institute, Bangalore 560080, India; and dInstitute for Stem Cell Biology and RegenerativeMedicine, Bangalore 560065, India

Edited by Ronald D. Vale, Howard Hughes Medical Institute and University of California, San Francisco, CA, and approved January 26, 2016 (received forreview July 27, 2015)

The surface of a living cell provides a platform for receptor signaling,protein sorting, transport, and endocytosis, whose regulation re-quires the local control of membrane organization. Previous work hasrevealed a role for dynamic actomyosin in membrane protein andlipid organization, suggesting that the cell surface behaves as anactive composite composed of a fluid bilayer and a thin film of activeactomyosin. We reconstitute an analogous system in vitro thatconsists of a fluid lipid bilayer coupled viamembrane-associated actin-binding proteins to dynamic actin filaments andmyosinmotors. Uponcomplete consumption of ATP, this system settles into distinct phasesof actin organization, namely bundled filaments, linked apolar asters,and a lattice of polar asters. These depend on actin concentration,filament length, and actin/myosin ratio. During formation of the polaraster phase, advection of the self-organizing actomyosin networkdrives transient clustering of actin-associatedmembrane components.Regeneration of ATP supports a constitutively remodeling actomyosinstate, which in turn drives active fluctuations of coupled membranecomponents, resembling those observed at the cell surface. In amulticomponent membrane bilayer, this remodeling actomyosin layercontributes to changes in the extent and dynamics of phase-segregating domains. These results show how local membranecomposition can be driven by active processes arising from actomy-osin, highlighting the fundamental basis of the active compositemodel of the cell surface, and indicate its relevance to the study ofmembrane organization.

membrane organization | active mechanics | actin | myosin II

The cell surface mediates interactions between the cell and theoutside world by serving as the site for signal transduction. It

also facilitates the uptake and release of cargo and supportsadhesion to substrates. These diverse roles require that the cellsurface components involved in each function are spatially andtemporally organized into domains spanning a few nanometers(nanoclusters) to several micrometers (microdomains). The cellsurface itself may be considered as a fluid–lipid bilayer whereinproteins are embedded (1). In the living cell, this multicompo-nent system is supported by an actin cortex, composed of abranched network of actin and a collection of filaments (2–4).Current models of membrane organization fall into three

categories: those invoking lipid–lipid and lipid–protein interac-tions in the plasma membrane [e.g., the fluid mosaic model (1, 5)and the lipid raft hypothesis (6)], or those that appeal to themembrane-associated actin cortex (e.g., the picket fence model)(7), or a combination of these (8, 9). Although these modelsbased on thermodynamic equilibrium principles have success-fully explained the organization and dynamics of a range ofmembrane components and molecules, there is a growing classof phenomena that appears inconsistent with chemical andthermal equilibrium, which might warrant a different expla-nation. These include aspects of the organization and dynamics of

outer leaflet glycosyl-phosphatidylinositol-anchored proteins (GPI-anchored proteins) (10–13), inner leaflet Ras proteins (14),and actin-binding transmembrane proteins (13, 15, 16).Recent experimental and theoretical work has shown that

these features can be explained by taking into account that manycortical and membrane proteins are driven by ATP-consumingprocesses that drive the system out of equilibrium (13, 15, 17).The membrane models mentioned above have by-and-largeneglected this active nature of the actin cortex where actin fila-ments are being continuously polymerized and depolymerized(18–21), in addition to being persistently acted upon by a variety ofmyosin motors (22–24) that consume ATP and exert contractilestresses on cortical actin filaments, continually remodeling the ar-chitecture of the cortex (4, 21, 25). These active processes in turncan generate tangential stresses and currents on the cell surface,which could drive the dynamics and local composition of mem-brane components at different scales (22, 26–29).Actin polymerization is proposed to be driven at the membrane

by two nucleators, the Arp2/3 complex, which creates a denselybranched network, as well as formins that nucleate filaments(18, 21, 30). A number of myosin motors are also associated withthe juxtamembranous actin cortex, of which nonmuscle myosin IIis the major component in remodeling the cortex and creatingactin flows (4, 23, 25, 26, 31, 32). Based on our observations that

Significance

This manuscript addresses the role of active processes in thespatial organization and dynamics of cell surface components.Using a reconstituted minimal system, we provide experimen-tal evidence for a proposed clustering mechanism that relies onthe intrinsic, active mechanics of actin filaments and myosinmotors expected to be present at the cell cortex. The couplingbetween the actomyosin and the lipid bilayer gives rise toan emergent active composite with properties that resemblethose observed in live cells. This clustering mechanism is a keyfeature of the active composite cell surface model and furthersour understanding of the multiple ways in which the cell sur-face might regulate its composition.

Author contributions: D.V.K., M.R., and S.M. conceived the experiments; D.V.K., R.D.M.,M.R., and S.M. designed research; D.V.K., E.I., A.B., and S.M. performed research; E.I.performed some of the FRAP experiments; P.B. designed and purified capping protein;P.B. contributed new reagents/analytic tools; A.B. purified proteins; D.V.K., K.H., and M.R.analyzed data; and D.V.K., K.H., R.D.M., M.R., and S.M. wrote the paper.

The authors declare no conflict of interest.

This article is a PNAS Direct Submission.1Present address: Department of Systemic Cell Biology, Max Planck Institute of MolecularPhysiology, 44227 Dortmund, Germany.

2To whom correspondence should be addressed. Email: mayor@ncbs.res.in.

This article contains supporting information online at www.pnas.org/lookup/suppl/doi:10.1073/pnas.1514030113/-/DCSupplemental.

www.pnas.org/cgi/doi/10.1073/pnas.1514030113 PNAS | Published online February 29, 2016 | E1645–E1654

BIOPH

YSICSAND

COMPU

TATIONALBIOLO

GY

APP

LIED

PHYS

ICAL

SCIENCE

SPN

ASPL

US

Dow

nloa

ded

by g

uest

on

Oct

ober

29,

202

0

the clustering of cell surface components that couple directly orindirectly to cortical actin [e.g., GPI-anchored proteins, proteins ofthe Ezrin, Radaxin, or Moesin (ERM) family (13, 15)] depends onmyosin activity, we proposed that this clustering arises from thecoupling to contractile actomyosin platforms (called “actin as-ters”) produced at the cortex (15, 33).A coarse-grained theory describing this idea has been put

forward and corroborated by the verification of its key predic-tions in live cells (15, 33), but a systematic identification of theunderlying microscopic processes is lacking. Given the com-plexity of numerous processes acting at the membrane of a livingcell, we use an in vitro approach to study the effect of an energy-consuming actomyosin network on the dynamics of membranemolecules that directly interact with filamentous actin.A series of in vitro studies have explored the organization of

confined, dynamic filaments (both actin and microtubules) (34–39) or the role of actin architecture on membrane organization(40–46). Indeed, these studies have yielded insights into thenontrivial emergent configurations that mixtures of polar fila-ments and motors can adopt when fueled by ATP (34–37), inparticular constitutively remodeling steady states that displaycharacteristics of active mechanics (38, 39, 47). However, theeffect of linking these mechanics to the confining lipid bilayerand its organization has not been studied.The consequences of actin polymerization on membrane or-

ganization, in particular on giant unilamellar vesicles (GUVs),have been addressed in a number of studies on the propulsion ofGUVs by an actin comet tail (40, 45, 46). In those experiments,the apparent advection of membrane bound ActA or WASPtoward the site of actin polymerization is mainly due to thechange in binding affinity of WASP to actin through Arp2/3 (44)and the spherical geometry resulting in the drag of actin to onepole of the vesicle after symmetry break of the actin shell. Thatthis dynamic process changes the bulk properties of the bilayer,namely the critical temperature of a phase-separating lipid bi-layer, was shown by Liu and Fletcher (40) when the actin nu-cleator N-WASP was connected to a lipid species (PIP2) that wascapable of partitioning into one of the two phases.Besides these pioneering studies on the effects of active pro-

cesses on membrane organization, little was done to directly testthe effect of active lateral stresses as well as actomyosin remod-eling at the membrane, particularly on the dynamics and organi-zation of membrane-associated components.To this end, we build an active composite in vitro by stepwise

addition of components: a supported lipid bilayer with an actin-binding component, actin filaments, and myosin motors. By sys-tematically varying the concentrations of actin and myosin as wellas the average actin filament length, we find distinct states ofactomyosin organization at the membrane surface upon com-plete ATP consumption. More importantly, we find that theATP-fueled contractile actomyosin currents induce the transientaccumulation of actin-binding membrane components. As pre-dicted, the active mechanics of actin and myosin at physiologi-cally relevant ATP concentrations drives the system into anonequilibrium steady state with anomalous density fluctuationsand the transient clustering of actin-binding components of thelipid bilayer (15, 33). Finally, connection of this active layer ofactomyosin to a phase-segregating bilayer, influences its phasebehavior and coarsening dynamics.

ResultsDynamic Association of a Lipid Bilayer Probe with Actin. We createdsupported lipid bilayers (SLBs) composed of 1,2-dioleoyl-sn-glycero-3-phosphocholine (DOPC) doped with 2% Ni2+-chelatedlipids [DGS-NTA(Ni2+)]. The Ni2+-chelated lipids recruit histi-dine-rich proteins, including an engineered membrane–actinlinker (HYE) constructed from the actin-binding domain of Ezrin(EzrABD), yellow fluorescent protein (YFP), and 10 tandem

histidine residues allowing stable SLB binding during the time ofan experiment (Fig. S1A). We chose the EzrABD as a membrane–actin linker because we previously found that it is sufficient toconfer actin-dependent clustering to fusion proteins in vivo (15).We also created variants of HYE: one with a point mutation in theEzrABD (R579A) that abolishes actin binding (48) and ablatesthe ability to cluster transmembrane proteins (15), and another,nonfluorescent derivative, HKE, with the tripeptide KCK in placeof YFP (Fig. 1 A and B) (49). As expected, bilayers containingeither HYE or HKE recruited actin filaments (Fig. S1B), resultingin a thin film adjacent to the bilayer (Fig. 1C and Fig. S1C),whereas HYE(R579A) was unable to localize actin to the mem-brane. Actin concentrations were restricted to the range of 100–1,000 nM to limit the actin layer thickness to less than 10 actinfilaments (see Table S1 for details of the experimental conditions).We controlled the length of these filaments by adding stoichio-metric amounts of capping protein (Fig. S1D). To induce con-tractile stresses and flows, we added the motor protein myosin IIand ATP.

Membrane-Tethered Actin Filaments Alter the Mobility of HYEConstructs That Are Competent to Bind Actin as Well as Those ThatCannot Bind Actin. We first characterized the lateral mobility ofactin-binding EzrABD constructs in the absence and presenceof filamentous actin using fluorescence recovery after photo-bleaching (FRAP) and fluorescence correlation spectroscopy(FCS). The mobility of SLB-bound proteins was probed by ob-serving the diffusion of His10-tagged fluorescent proteins inFRAP experiments. FRAP curves could be well fit to a singleexponential function, from which we computed an effective dif-fusion coefficient; His10-GFP and HYE showed a similar dif-fusion coefficient in the absence of actin (Fig. S1 E–G).Adding increasing concentrations of filamentous actin de-

creased the effective diffusion coefficient of “wild-type” HYEbut had no effect on the R579A mutant (Fig. 1C and Fig. S1G),and the immobile fraction of HYE was slightly increased from12% to 16% in the presence of 1,000 nM actin (Fig. S1F). De-creasing the average length of actin filaments by using cappingprotein (CP) (Fig. S1D) increased HYE mobility as judged byFRAP (Fig. 1D) and FCS measurements (Fig. S1 H and I).Importantly, the recovery of HYE in regions covered by actinfilaments reflects the transient nature of the interaction ofEzrABD with actin. Interestingly, when we tethered actin fila-ments to bilayers with the nonfluorescent Ezrin construct, HKE,and then measured diffusion of HYE(R579A) in the same bi-layers, we observed a significant reduction in mobility. The mo-bility of HYE(R579A) decreased monotonically with increasingactin concentrations, but this effect was less pronounced than thatobserved with wild-type HYE (Fig. 1C). However, His10-GFP didnot show a significant reduction in its mobility in the presence ofactin filaments tethered via HKE to SLBs as detected by FCS (Fig.S1J). In addition, the fluidity of the lipid bilayer itself was notaffected by the membrane tethered actin network, as the recoveryof the fluorescently labeled lipid 1,2-dioleoyl-sn-glycero-3-phos-phoethanolamine-N-(lissamine rhodamine B sulfonyl) (RhoPE)was unchanged (Fig. S1K). These results are consistent with aprevious report on the effect of a dense actin network on thediffusion of other membrane-associated proteins in vitro (41) andshow that the association of actin with the membrane bilayerprovides an impedance for the mobility of membrane componentsthat can sterically and biochemically interact with actin filaments,evoking the idea of a membrane fence (7, 50). It is to be noted thatthis effect is diminished when the actin filaments are too short toform an entangled meshwork as observed when we reduced actin-filament length with titrated amounts of capping protein (Fig. 1Dand Fig. S1I).

E1646 | www.pnas.org/cgi/doi/10.1073/pnas.1514030113 Köster et al.

Dow

nloa

ded

by g

uest

on

Oct

ober

29,

202

0

Myosin Motors and Membrane-Bound Actin Filaments Self-Organizeinto Distinct Configurations. When we added bipolar myosin fila-ments—composed of ∼100–500 proteins (51) with lengths between0.5 and 1.2 μm (Fig. S2A)—to a bilayer-bound thin film of actinfilaments in the presence of a fixed amount of ATP and allowed thesystem to evolve until the ATP was consumed, we reproduciblyobserved three distinct organizational states: (i) filament bundles,(ii) linked apolar asters, and (iii) disordered lattices of polar asters(Fig. 2A). The orientation of actomyosin networks in the bundledstate was correlated over distances >5 μm (Fig. S2 B and C),whereas the polar aster state was characterized by islands of actinreflected in a local minimum of the spatial density correlation (Fig.S2D). The linked apolar aster phase was characterized by a lowcorrelation in filament orientation (Fig. S2 B and C) and by theabsence of oscillations in the spatial density correlation (Fig. S2D).These three states (after consumption of the ATP pool) wereobtained by varying (i) actin concentration, (ii) filament length, and(iii) filament-to-motor ratios (Fig. 2B and Table S2). Because thepolar asters most closely reflect the organization of the contractileplatforms that were proposed by our theoretical model, and that aresupposed to be responsible for the clustering of membrane particles(15), we focus in the following on these structures and conductedfurther experiments in conditions geared toward aster formation.

The Polar Aster Phase. We next characterized the architecture anddynamics of the polar aster phase in multiple ways. First, we notedthat the size of a single aster depends on the average actin fila-ment length (Fig. 2C), in which myosin motors have contractedmultiple filaments into a configuration that blocks further rear-rangement, indicative of a jammed state. Second, analyzing theirspatial distribution revealed that the asters formed a disorderedlattice with a characteristic spacing on the order of microns

(Fig. S2E and Materials and Methods). Third, the localization offluorescently labeled capping protein demonstrated the polar na-ture of the actin asters, in which all filaments are oriented withtheir (capped) barbed ends facing inward, toward the core of theaster while pointed ends face outward (Fig. 2D and Fig. S2G).Consistent with this interpretation, we also observed that fluo-rescently labeled myosin II—a barbed-end–directed motor—lies inthe cores of asters (Fig. 2D). Similar observations of actin andmyosin organization were obtained in samples imaged by stimu-lated emission depletion (STED) microscopy (Fig. S2F). Scanningthe sample with multipoint structural illumination microscopy(SIM) in all three spatial dimensions allowed the clear visuali-zation of the contraction of single actin filaments into asters (Fig.2D) and of the thin film of membrane bound F-actin (Fig. 2E). Inthe case of linked apolar asters, however, the distribution ofcapping protein was more spread out, as some filaments are linkedby multiple myosin filaments on both ends (Fig. S2H). Finally, inthe central region of sporadically forming larger asters, we ob-served distinct rings of capping protein (Fig. S2I), another hallmarkof myosin-driven polarity sorting and filament organization (52). Itis to be noted that the formation of actin asters (as well as the otherconfigurations obtained under different initial conditions) occurredover the major area of the sample, which was tested by the im-aging of multiple regions at random positions (Fig. S2J).

Bilayer Components Are Advected by Contractile Actomyosin Flows.We next asked whether contractile actomyosin flows advect cou-pled bilayer components. We followed the dynamics of the con-tractile system and its associated membrane components as thesystem approached the steady-state polar aster configuration de-scribed above. Particle image velocimetry (PIV) of movies of la-beled actin filaments revealed regions where the net divergence of

0.5

0

1

1.5

D(μ

m2 /

sec

)

0[actin] (nM)

100 500 750 1000

A

C

BEzrin constructs

10x His YFP EzrinABDHYE

10x His YFP EzrinABD(R579A)

HYE(R579A)

HKE10x His KCK EzrinABD

D1000 nM500 nM

actin

actin

10 nM CP 125 nM CP

0.5

0

1

1.5

D(μ

m2 /

sec

)10

0

20

30

F-actin length (μm)

0[capping protein] (nM)

10 31 62 125

lipids (PC)

functionalized lipid(Ni-NTA-DGS)membrane-actin linkerf-actin

capping protein

myosin II filaments

HYEHYE(R579A)HYE(R579A) + HKE

Fig. 1. In vitro setup. (A) Schematic of the in vitro system. (B) Schematic of the actin–membrane linker constructs. (C) Representative images of SLB-boundF-actin and plot of the diffusion coefficient HYE (n = 59–147), HYE(R579A) (n = 15–24), and a combination of HKE and HYE(R579A) (n = 10–39) at variousF-actin concentrations. (D) Representative images of SLB-bound rhodamine-labeled F-actin with capping protein (CP) and plot of the diffusion constant ofHYE (n = 50–80) and a combination of HKE and HYE(R579A) (n = 10–20) with F-actin of decreasing average length (n = 158–749). Small squares depict meanvalues; box heights, SDs; middle lines, medians; whiskers, 5–95% ranges. (Scale bars: 10 μm.)

Köster et al. PNAS | Published online February 29, 2016 | E1647

BIOPH

YSICSAND

COMPU

TATIONALBIOLO

GY

APP

LIED

PHYS

ICAL

SCIENCE

SPN

ASPL

US

Dow

nloa

ded

by g

uest

on

Oct

ober

29,

202

0

the velocity was negative at the sites of aster formation, and withmyosin-driven contractile flows of ∼0.5–1 μm/min (Fig. S3 A–C)as observed in other in vitro studies (53). A key prediction ofthe active composite model is that contractile actin currents willadvect actin-binding membrane proteins into clusters. Consistentwith this prediction, we observed that the HYE construct formsclusters that colocalize with forming, myosin-induced actin asters(Fig. 3 A and B and Movie S1). Detailed analysis of the regions ofaster formation confirmed that the clustering of HYE occurredonly when and where actin flows were contractile (i.e., at positionsand times where the divergence of actin velocity is negative; Fig.3C). Whenever the local contractile currents ceased, the associ-ated HYE clusters dissolved, indicating that cluster formation was

due to myosin-generated flows and not simply increased actindensity in the aster (Fig. 3 B and C).We further characterized the myosin dependence of density

fluctuations in membrane-associated proteins by a cross-correlationanalysis. We collected time-lapse movies of labeled actin andHYE and, for every 5 × 5 pixel region, computed (i) the tem-poral cross-correlation coefficient between the intensities ofHYE and labeled actin and (ii) the temporal variance of thelocal actin density (Materials and Methods). The latter distin-guishes between regions of static and dynamic actin. We foundthat HYE and actin dynamics were correlated only in regionswhere the actin network was dynamic (Fig. 3 D and E and Fig.S3D). These zones of strong correlation appeared only during

12

9

6

3

0.3 0.6 0.9

F-actin length (μm)

0.30.6

0.91.2

00

[Myosin II]/[F-actin] [F-actin] (nM)

filament bundleslinked, apolar asterspolar asters

1.2

2μm 3μm 4μm 5μm

aste

r siz

e, d

(μm

)

2

6

0

4

F-actin length

F-actin length, l (μm)0 1 2 3 4 5 6

B

filament bundlesA

C

polar asterslinked, apolar asters

D

slope: 0.5(+/- 0.2)

ii

ac�nCP

i

ac�nmyosin II

++

+++ +

--

-

-

-

-

myosin IIac�nCP

iviii

ac�n

E

ac�n

yzxzxy

Fig. 2. Myosin-induced formation of polar actin asters. (A) Images of F-actin after myosin II action show three distinct configurations. (Scale bars: 10 μm,2 μm.) (B) Phase diagram of actomyosin organization obtained by manual classification. (C) Diameter of polar actin asters as a function of mean filamentlength (n > 20 for each data point) and linear fit showing a linear dependence of aster size to F-actin length (slope, 0.5 ± 0.2); Insets show average projectionsof image stacks that were created by cropped single actin aster images (n = 12–20) for each indicated F-actin length. (D) Representative images depicting theorganization of polar actin asters by imaging rhodamine-actin and Atto-647-myosin II (i), or rhodamine-actin and Alexa 647 capping protein (ii) to obtain aschematic description of a typical actin aster organization (iii). (Scale bar: 2 μm.) (E) X, Y, Z scans of an actomyosin network stabilized with 0.04% glutar-aldehyde and imaged with multipoint structural illumination microscopy (SIM) showing the thin membrane-confined actin layer and the local contraction intoasters. (Scale bars: in YZ and XZ scans, 2 μm; in XY scan, 5 μm.)

E1648 | www.pnas.org/cgi/doi/10.1073/pnas.1514030113 Köster et al.

Dow

nloa

ded

by g

uest

on

Oct

ober

29,

202

0

the contractile period of aster formation and were absent beforemyosin addition as well as in the jammed steady state (Fig. 3 Dand E and Fig. S3D).Advection of membrane-associated proteins was not the result

of bulk hydrodynamic flow in the membrane, but required specificinteraction between the membrane component and filamentousactin, because HYE(R579A) did not cluster in bilayers to whichactin was linked via HKE (Fig. S3 E and F). Similarly, no effect ofactomyosin-driven HYE on bulk lipid dynamics could be detectedby the use of the fluorescently labeled lipid RhoPE and the cross-correlation analysis (Fig. S3 G and H). Both the PIV data of actinflows (Fig. 3C) and the cross-correlation analysis (Fig. 3 D and E)indicated that advection drives clustering of actin-associatedcomponents during myosin-dependent aster formation.

Emergence of an Active Actin–Membrane Composite at High Concentrationsof ATP.Upon complete ATP consumption (initial ATP, <0.01 mM),our actomyosin networks eventually reached a static steady stateconsistent with earlier reports (54). The cortex of a living cell,however, is continuously driven out of equilibrium by high con-centrations (2–5 mM) of ATP (55). To mimic this situation,we added ATP (1 mM final concentration) to preformed,membrane-associated actomyosin asters. Addition of ATP in-duced a period of reorganization during which large, static astersdissolved (Fig. 4A and Movie S2), and the system moved to anew steady state, characterized by highly dynamic actin filaments(Fig. S4A) with a statistically uniform, time-averaged density atlength scales of >5 μm (Fig. S4B). Time-lapse movies revealedtransient, localized accumulations of actin filaments that accretedand dispersed over tens of seconds (Fig. S4A,Upper, and Movie S3).

We observed the same result when we reconstituted the systemwith an ATP-regenerating system (Fig. S4A, Lower). These dynamicdensity fluctuations persisted for several minutes after additionof ATP (and much longer with the ATP regeneration system) (Fig.S4 B and C), suggesting a nonequilibrium steady state in whichcontinuous remodeling was driven by persistent myosin activity.We next asked whether the dynamics of HYE were affected by

the continuous remodeling of the actomyosin network. At highATP concentrations, the cross-correlation analysis of HYE andactin densities had a trend similar to that observed during con-tractile aster formation. That is, HYE and actin dynamics werestrongly correlated in regions of highly dynamic actin (Fig. 4 Band C and Fig. S4D). The active composite framework (33)predicts large, myosin-dependent spatial and temporal fluctu-ations in HYE density, similar to those observed for GPI-anchored proteins in living cells (15) and for other active systemsin both theory and experiment (47, 56). Compared with otherphases of the actomyosin system, the pixel intensity distributionof HYE in the driven steady state showed a broader, non-Gaussian distribution, with a significant rightward skew indica-tive of HYE clustering (Fig. 4D and Fig. S4 E and F), similar tothat observed for GPI-anchored proteins in vivo, consistent withtheoretical predictions (15).A key statistical signature of actively driven systems is the ap-

pearance of “giant number fluctuations” (56), which we previouslyobserved for GPI-anchored proteins in live cells (15). We tested forthe presence of giant number fluctuations in our reconstitutedsystem by measuring variations in HYE intensity over differentspatial scales. Briefly, we varied the size of our observation windowand measured how fluctuations in fluorescence intensity change

Aactin myoII HYE

5 m

in10

min

15 m

in 0

1

0.5

norm

aliz

ed in

tens

ity

actin

HYE

myoII

HYE ctrl

B D

HY

E-a

ctin

cros

s-co

rrel

atio

nac

tintim

e la

pse

actin + myoIIactinactin + myoII

(jammed)

1

0.5

0

time

CE

-4 -3 -2 -1

actin

-5

0

0.2

-0.2

div(

v act

in) (

1/m

in)

19 22

actin + myoII(jammed)

1.1

1

0.9

IHY

E /IHY

E (t0 )

time (min)

time (min)630 9 12 15

time (min)630 9 12 15

+ myoII

16 25

1 ImyoII /m

ax ImyoII

0

-1

div(vactin)

HYEmyoII

0.5

0

1

HY

E-a

ctin

cr

oss-

corr

elat

ion

10.5 1.5variance of actin density

actinactin + myoIIactin + myoII (jammed)

0 2

Fig. 3. Actomyosin-induced HYE accumulation during actin aster formation. (A) Snapshots of actin (Left), myosin II (Middle), and HYE (Right) during asterformation. (Scale bar: 10 μm.) (B, Top) View of the outlined region in A. (Scale bar: 2 μm.) (Bottom) Intensity profiles averaged over 40 circular regionscantered on actin asters in A, normalized to [0; 1]; control is computed from 10 circular nonaster regions; images were corrected for photobleaching by asingle exponential function. (C) Graph of contractile actin flows computed from PIV data (black; Materials and Methods) and HYE intensity (green) averagedover the same regions as in B at indicated condition. (D) Color-coded time projection of actin (Top) and corresponding HYE-actin cross-correlation (Bottom) ofSLBs before (Left) and after addition of myosin II, during contraction (Center) and after its halt (Right). (Scale bar: 10 μm.) (E) Plots of HYE-actin cross-correlationversus the variance of actin density from images in D; mean and SD computed from data within the x-axis whisker’s range.

Köster et al. PNAS | Published online February 29, 2016 | E1649

BIOPH

YSICSAND

COMPU

TATIONALBIOLO

GY

APP

LIED

PHYS

ICAL

SCIENCE

SPN

ASPL

US

Dow

nloa

ded

by g

uest

on

Oct

ober

29,

202

0

with window size (Materials and Methods). We found that, forsystems at equilibrium (i.e., no actin, actin alone, or static acto-myosin cases), HYE intensity fluctuations scaled with the windowsize b as bα with α = 0.49 (±0.02; n = 27), in good agreement withthe expected behavior of number fluctuations in an equilibriumsystem away from criticality (α = 0.5) (56). In contrast, the con-stitutively remodeling steady state exhibited α = 0.86 (±0.09; n =7), when evaluated over window sizes larger than the mean F-actinlength (≥5 μm) (Fig. 4 E and F, Fig. S4 G and H, and Table S3).We observed these giant number fluctuations in our reconstitutedsystem only at high concentrations of ATP (Fig. 4E) (30). Note thatthe average size of dynamic clusters produced in this driven steadystate was significantly smaller than our smallest interrogationwindow, and, therefore, we were analyzing fluctuations that occuron a length scale significantly larger than the cluster size (57). Thisindicates that the large number fluctuations are a consequence ofactive driving of HYE by the myosin-propelled actin filamentsupon ATP consumption, and that this actomyosin–membranesystem can be considered as an active composite system.

Consequences of an Actomyosin Layer on a Phase-Separating LipidSystem. After establishing the effect of the active actomyosinlayer on the organization of a single passive membrane compo-nent (HYE) in a homogenous bilayer, we asked whether thisactivity could have effects on the configuration of a multicom-ponent bilayer, namely the phase segregation behavior of lipidsin the SLB. For this, we formed a standard ternary lipid mixtureof DOPC, 1,2-dipalmitoyl-sn-glycero-3-phosphocholine (DPPC),and cholesterol, doped with RhoPE as a marker for liquid dis-ordered domains and DGS-NTA(Ni2+) on mica sheets (42), andimaged the system on a confocal microscope with a low magni-fication (20×, N.A. 0.75) (Fig. 5A and Materials and Methods).After formation of bilayers and subsequent addition of HYE and

actin filaments in the homogenous mixed lipid phase at 37 °C, wedecreased the temperature to 28 °C to induce phase separation,mounted the chamber on the microscope, and began imaging(with a minimal dead time of ∼60 s). The transition temperaturefor this lipid mix was about Tc = 33 °C in agreement with pre-vious reports (58), which we only could roughly estimate bywarming the sample up from T = 28 °C to T = 37 °C as our setupdid not allow controlled, repeated heating and cooling. AlthoughHYE and actin were homogeneously distributed in regions de-void of visible lipid domains (Fig. 5B, Top), HYE was excludedfrom liquid ordered (lo) domains and located in the liquid dis-ordered (ld) phase as expected for the DGS-NTA(Ni2+) lipids(Fig. 5B, Bottom). We note that actin filaments followed HYEand were excluded from lo domains as well, eventually resultingin a more bundled organization in the ld phase irrespective of theF-actin length (Fig. 5B, Fig. S5A, and Movie S4).Following the behavior of lo domains over time, a steady

growth was detected in the HYE-only bilayers (Fig. 5 C and D).This growth of the average size of lo domains was slowed downby membrane-bound actin filaments, whereat longer, entanglingactin filaments acted as barriers for lo domains (Fig. 5 C and D),but shorter actin filaments (shorter than the domain diameter)succumbed to their displacement (Fig. S5A).In contrast, the presence of active myosin motors together

with longer actin filaments stalled the increase of average do-main size even though at the outset of observation the phasesegregation was comparable to that obtained with or withoutbound actin filaments (Fig. 5 C and D and Fig. S5 B and C). Wenext compared net lo-domain area fraction in a given frame fromthe beginning of imaging (t0) to later time points. In free SLBsand those with F-actin alone, the net area of lo domains (ΣAlo)remained roughly constant over this time window but decreasedin the presence of the remodeling actomyosin network (Fig. 5E,

0-1 110-6

10-2HYE

high ATPactin + myoII

(lpixel-<Iframe>)/<Iframe> P((

l pixe

l-<I fr

ame>

)/<I fr

ame>

)

10-4

actin

-10 sec 6 sec 12 sec 28 sec 88 sec

32 41

-5

-6

-4

ln(Abox/ m2)

HYE

high ATPactin + myoII

time

1

0.5

0

slop

e (

)

actin

0.8

0.6

0.4

1

_ + + +myoII ATP

+ +highlowlow

___

slop

e,

0.25

0.5

0

0 0.5

HY

E-a

ctin

cr

oss-

corr

elat

ion

variance ofactin density

actin + myoII+ ATP

1actin time lapse HYE-actin

cross-correlation

ln(<

I2 >0.

5 /<I

>)

A

B C

D E F

Fig. 4. The remodeling actomyosin–membrane system shows features of an active composite. (A) Aster disassembly after addition of ATP to polar actinasters. (Scale bar: 10 μm.) (B) Color-coded time projection of actin (Left) and corresponding HYE-actin cross-correlation (Right) of a remodeling actomyosinnetwork. (Scale bar: 2 μm; duration, 15 min.) (C) Plot of HYE-actin cross-correlation versus the variance of actin density from images in B; mean and SDcomputed from data within the x-axis whisker’s range. (D) Individual HYE intensity probability distributions for SLBs containing HYE alone, jammed (actin plusmyosin II), or remodeling actomyosin (high ATP); dashed lines depict Gaussian fits of data from each condition. (E) Number fluctuations of HYE (ΔN∼bα)under the indicated conditions; squares depict mean values; box heights, SDs; middle lines, medians; dots, individual experiments (n = 7–12). (F) Numberfluctuations of HYE as a function of interrogation box size (Abox) obtained from one sample at conditions as indicated.

E1650 | www.pnas.org/cgi/doi/10.1073/pnas.1514030113 Köster et al.

Dow

nloa

ded

by g

uest

on

Oct

ober

29,

202

0

Fig. S5D, and Table S4 for experimental details). This is likely tobe due to a loss of small domains (compared with the actin fil-ament length) by the local remodeling of the actomyosin networkin combination with a cessation in growth of larger domains (Fig.S5B and Movie S5). The cross-correlation between actin filamentsand HYE in the presence of myosin contraction still showed thesignature of actomyosin-induced advection, indicating that it isindeed the actomyosin activity causing the loss of lo domains(Fig. 5 F and G). Interestingly, we observed this lo domain re-duction reliably only in the presence of long, entangled actin fil-ament networks, possibly a result of the size-dependent interplaybetween actin filaments and lo domains.Taken together, these experiments indicate that the en-

gagement of the actomyosin layer on a phase-segregating bi-layer influences its phase segregation characteristics in a lengthscale-dependent manner. The actin filament length scale sets alimit on the domain sizes, the actomyosin network can act on.The extent of this interplay will also depend on the strength ofactin–membrane link, and to which component of the lipidbilayer actin is linked to, underscoring the complex nature ofactive actomyosin membrane composite systems that remain to bestudied in more detail in future.

DiscussionIn the present study, we show that myosin-driven actin networksaffect the organization of membrane proteins, transforming theactin–myosin–membrane system under continuous ATP consump-tion to an active composite.Examining first the patterns formed by actin filaments and myosin

motors, we found that filament length is an important determinant inactomyosin structure formation. As observed in other in vitro works,long actin filaments form bundled structures (37, 54) or contractednetworks (35, 37, 59) under the action of myosin II, whereas at theshorter actin filament lengths used here, they gave rise to compact,isolated asters. We find that the following parameters characterizethe different static states obtained: (i) actin filament length, (ii)myosin-to-actin filament ratio, and (iii) F-actin concentration.Taking into consideration the complexity of the cell cortex com-position (25), the Arp2/3 complex and the Formin, mDia1, aremajor factors controlling actin polymerization and giving rise tobranched and straight filaments, respectively (21). The action ofactin filament severing agent cofilin could serve to further differ-entiate the population of filaments, giving rise to a hierarchy oflength scales in which smaller filaments may form structures suchas the asters observed here, whereas longer filaments would as-semble into a dense meshwork or stress fibers (2). Consistent with

A B

C D E

F G

Fig. 5. Consequences of the remodeling actomyosin on a phase-separating membrane. (A) Schematic showing the setup for SLBs with ternary lipid mixture.(B) Example fluorescent images of SLBs containing RhoPE, HYE, and binding actin with regions devoid of domains (Top) or showing lipid segregation (darkregions) (Bottom). (Scale bar: 10 μm.) T = 28 °C. (C) Examples of lo domains (dark regions) in SLBs containing HYE only (Left), bound F-actin (Center), or aremodeling actomyosin (Right). (Scale bar: 10 μm.) (D) Averaged lo domain sizes at different conditions; lines represent averages for each condition (n = 4–6,with 15–158 domains per experiment), shaded area is SD, and t0 indicates start of imaging. (E) Relative change in net lo domain area between t0 and 750 slater for indicated conditions; squares depict mean values; box heights, SDs; dots, individual experiments. (F) Color-coded time projection of actomyosinremodeling (from Fig. S5B) and the corresponding HYE-actin density cross-correlation. (Scale bar: 20 μm; duration, 2 min.) (G) Plot of HYE-actin cross-correlationversus the variance of actin density; mean and SD are from data within the x-axis whisker’s range.

Köster et al. PNAS | Published online February 29, 2016 | E1651

BIOPH

YSICSAND

COMPU

TATIONALBIOLO

GY

APP

LIED

PHYS

ICAL

SCIENCE

SPN

ASPL

US

Dow

nloa

ded

by g

uest

on

Oct

ober

29,

202

0

the system exhibiting distinct states, similar starting conditions resultfor certain parameters (e.g., [actin] = 600 nM, [myoII] = 90,Lf-actin = 3 μm), in different configurations, which most likely re-flects the lack of precise control of actin filament length in our systemand the proximity to a phase boundary. Finally and importantly, acontinuous supply of ATP and the transient binding of actin filamentsto the bilayer drives the system into a mode of continuous remodeling.Focusing on the effect of the actomyosin system on the dy-

namics of membrane components, we found that the process ofaster formation transiently accumulated a membrane componentcoupled to actin (HYE). The PIV and cross-correlation analysisindicate that the transient accumulation was driven by the con-tractile flows induced by myosin and was not merely a result ofthe increased actin density at the aster. Furthermore, the ad-vection of HYE by actomyosin did not induce hydrodynamicflows in the bilayer, as evidenced by the lack of clustering of bilayercomponents that were not capable of binding actin.At high ATP concentrations (1 mM; or in the presence of an

ATP-regenerating system), persistent motor activity continuallyexerted stresses on the filaments driving only the actin-associatedmembrane components out of equilibrium. We observed hall-marks of active dynamics on the bilayer, such as persistent ad-vection, clustering, and anomalous density fluctuations. To ourknowledge, this is the first time that such a reconstituted activecomposite system has been described. Our results suggest thatthe actin cortex may influence plasma membrane organizationnot only through the interaction of lipids and proteins with astable actin meshwork (9, 42) or polymerizing actin (40, 44), butalso through the flows of actin filaments generated by myosin-induced stresses. Importantly, the flows of myosin-driven shortactin filaments influence only those membrane components(proteins and lipids) that can bind to actin such as HYE, whereasthe mutant HYE(R579A) and the inert lipid RhoPE do not showa change in their dynamics. We note that this behavior is entirelyconsistent with our earlier observations in cells and theoret-ical predictions, where actin-associated components exhibit non-equilibrium behavior, such as giant fluctuations and temperature-independent diffusion behavior whereas unconnected componentsremain inert (13, 15). Transmembrane proteins with actin-bindingdomains in their cytoplasmic tails directly associate with actin atthe inner leaflet, and we have recently shown that long acyl chain-containing outer-leaflet GPI-anchored proteins interact with actin-associated phosphatidylserine (PS) at the cytoplasmic leaflet, inthe presence of adequate cholesterol (60), indicating that theactin-dependent clustering of membrane components requires adirect link to actin filaments and actomyosin dynamics. Thesemay be understood in terms of the active hydrodynamics of actinfilaments and myosin (56), which does not necessarily inducemeasurable hydrodynamic flows in the plasma membrane ofunconnected components. We emphasize that these observationsdo not exclude either the picket fence or phase segregationmechanisms that have been proposed earlier. However, the roleof actin and myosin-induced stresses, i.e., processes described byactive mechanics, should be included to arrive at a more com-plete description of the membrane in a living cell (33, 56).In the context of a phase-segregating membrane bilayer, we

found that the remodeling actomyosin layer contributed to achange in the size and dynamics of already formed (lo) domains(reduced/stalled growth), compared with a free membrane orone with associated static F-actin. Although at this point we donot fully understand the reason for the reduced membrane do-main segregation, it is likely that the active actomyosin churns upthe phase-segregating bilayer to create either domains smallerthan the optical limit or simply mixes up the membrane. In ad-dition, our data indicate that the membrane domains influencethe actin organization, imposing a minimal length scale for actinfilaments on the order of the membrane domain diameter, belowwhich domains will not be affected by actomyosin dynamics.

The effect of slowing of domain growth may be amplified in oursystem by frictional coupling to the substrate that is likely to in-fluence domain growth and dynamics (61). The influence of F-actinon domain dynamics is expected to be much stronger in a free-standing membrane system such as observed in GUVs (40, 43) orsupport-free planar bilayers (62). Recent theoretical work from thelaboratory of one of us suggests that the phase-segregating activecomposite membrane will exhibit specific characteristics in thedistribution of domain sizes and its scaling behavior (63), and theexperiments shown here exemplify the number of effects thatsuch an active composite system can have on their dynamics.The active composite nature of the cell surface also has impli-

cations for cellular function in the context of signaling, as severalsignaling proteins resident in the cell membrane have the capa-bility to interact with cortical actin either directly (64), via linkers(65), or indeed via transbilayer interactions of inner leaflet lipids (60).Signaling that originates at the cell surface may be influenced, en-hanced, or facilitated by interacting with actomyosin-related proteins(such as nucleators) that locally regulate the dynamics of actin suchthat cell surface proteins are organized optimally for their func-tion, resulting in enhanced reaction rates on the cell surface (66)or an efficient reading of a spatially inhomogeneous external li-gand (67); indeed, any function in which the spatiotemporal dy-namics of cell surface proteins is important is a potential target.We emphasize that the system we have described here is a

minimal realization of an active composite layer, where we haveincluded only key ingredients: small actin filaments of prescribedlength confined to a thin film on a supported membrane bilayer byusing an actin-binding binding membrane components (HYE) and,finally, ATP and myosin minifilaments. Our results on the actinconfigurations obtained after consuming the ATP in the system areconsistent with the prediction of our earlier theory [and indeedothers have predicted this for related systems (52, 68)]. The lengthscale of clustering and actomyosin structures demonstrated here arelarger than those proposed for the cell in the context of GPI-an-chored proteins, but this can be attributed to the differences inlength scales (Lactin = 2–7 μm in vitro, Lactin = 0.1–0.3 μm in vivo;LmyoII = 0.5–1.2 μm in vitro, LmyoII = ∼0.2 μm in vivo), compo-sition, and local environment between our in vitro system and thecell. The time required for aster formation at low ATP levels(30–60 s) is longer than expected in cells, likely due to thejamming of actin filaments during this process. However, theactin dynamics observed in the remodeling state (<1 s) moreclosely resembles the turnover of nanoclusters in the cell (0.1–1 s−1)(11). The ease with which this in vitro system reproduces the es-sential signatures of an active composite argues for its relevance tothe understanding of membrane organization of the living cell.

Materials and MethodsActive actin and myosin were purified from chicken breast with minor modi-fications of protocols described earlier (69, 70). His-tagged C-terminal fragmentof Ezrin and its isoforms (HYE, HKE, and HYE-RA) as well as CP were purifiedfrom bacterial lysates using affinity chromatography. Supported lipid bilayerswere formed by small unilamellar vesicles of DOPC and NTA-lipids in custom-built experimental chambers on cleaned glass slides, to localize the His-taggedproteins. F-actin length was regulated by titrated amounts of CP during F-actinpolymerization or by shear forces through pipetting F-actin. After addition ofpolymerized actin, myosin II was added to the chamber in ATP-containingbuffers and appropriate salt conditions to generate minifilaments (51). Phase-segregating lipid mixtures of multiple lipid species and cholesterol were pre-pared in chloroform, deposited on experimental chambers formed on freshlycleaved mica supports (∼100-μm thickness), and rehydrated for bilayer for-mation (42). Fluorescence imaging was carried out on a custom-designed totalinternal reflection fluorescence microscope, and electron microscopy on my-osin II filaments was performed following established protocols (71). Imageanalysis was carried out by pipelines developed specifically for the purpose ofeach experiment by using available routines in Image J and Matlab, which areavailable upon request. More detailed descriptions of the material andmethods used in this study are provided in SI Materials and Methods.

E1652 | www.pnas.org/cgi/doi/10.1073/pnas.1514030113 Köster et al.

Dow

nloa

ded

by g

uest

on

Oct

ober

29,

202

0

ACKNOWLEDGMENTS. We thank Scott Hansen (University of California,Berkeley), Tejas Gupte (InStem), C-CAMP Protein Technology Core, and theCIFF for help in protein purification and advanced light and electron microscopyimaging facilities; Enas Abu Shah, Baris Avsaroglu, Swathi Ayloo, Lena FittingKourkoutis, Alexis, Felix, Tamara Bidone, Jyongji (Physiology courses batches;2012 and 2013), and the HCIA Summer Institute 2013 for their participation inexploring the in vitro system at the Marine Biology Laboratory. We also thankRon Vale (University of California, San Francisco), Michael Rosen (University ofTexas Southwestern), Daniel Fletcher (University of California, Berkeley), and

James Spudich (Stanford University) for thoughtful discussions, andThomas van Zanten and the S.M. Laboratory [National Centre for BiologicalSciences (NCBS)] for careful reading of the manuscript. K.H. thanks PragyaSrivastava for fruitful discussions. K.H. and D.V.K. thank an old monk fordeep reflections on the matter. We thank the following sources for fundingsupport: AXA Research Fund and NCBS (D.V.K.), Simons Foundation(M.R.), J. C. Bose Fellowship from the Department for Science and Tech-nology, Government of India (S.M.), and Human Frontier ScienceProgram (S.M.).

1. Singer SJ, Nicolson GL (1972) The fluid mosaic model of the structure of cell mem-branes. Science 175(4023):720–731.

2. Morone N, et al. (2006) Three-dimensional reconstruction of the membrane skel-eton at the plasma membrane interface by electron tomography. J Cell Biol 174(6):851–862.

3. Charras GT, Hu C-K, Coughlin M, Mitchison TJ (2006) Reassembly of contractile actincortex in cell blebs. J Cell Biol 175(3):477–490.

4. Li D, et al. (2015) ADVANCED IMAGING. Extended-resolution structured illuminationimaging of endocytic and cytoskeletal dynamics. Science 349(6251):aab3500.

5. Engelman DM (2005) Membranes are more mosaic than fluid. Nature 438(7068):578–580.

6. Lingwood D, Simons K (2010) Lipid rafts as a membrane-organizing principle. Science327(5961):46–50.

7. Kusumi A, et al. (2005) Paradigm shift of the plasma membrane concept from thetwo-dimensional continuum fluid to the partitioned fluid: High-speed single-mol-ecule tracking of membrane molecules. Annu Rev Biophys Biomol Struct 34:351–378.

8. Jacobson K, Mouritsen OG, Anderson RGW (2007) Lipid rafts: At a crossroad betweencell biology and physics. Nat Cell Biol 9(1):7–14.

9. Arumugam S, Bassereau P (2015) Membrane nanodomains: Contribution of curvatureand interaction with proteins and cytoskeleton. Essays Biochem 57:109–119.

10. Sharma P, et al. (2004) Nanoscale organization of multiple GPI-anchored proteins inliving cell membranes. Cell 116(4):577–589.

11. Goswami D, et al. (2008) Nanoclusters of GPI-anchored proteins are formed by corticalactin-driven activity. Cell 135(6):1085–1097.

12. van Zanten TS, et al. (2009) Hotspots of GPI-anchored proteins and integrin nano-clusters function as nucleation sites for cell adhesion. Proc Natl Acad Sci USA 106(44):18557–18562.

13. Saha S, et al. (2015) Diffusion of GPI-anchored proteins is influenced by the activity ofdynamic cortical actin. Mol Biol Cell 26(22):4033–4045.

14. Plowman SJ, Muncke C, Parton RG, Hancock JF (2005) H-ras, K-ras, and inner plasmamembrane raft proteins operate in nanoclusters with differential dependence on theactin cytoskeleton. Proc Natl Acad Sci USA 102(43):15500–15505.

15. Gowrishankar K, et al. (2012) Active remodeling of cortical actin regulates spatio-temporal organization of cell surface molecules. Cell 149(6):1353–1367.

16. Jaqaman K, et al. (2011) Cytoskeletal control of CD36 diffusion promotes its receptorand signaling function. Cell 146(4):593–606.

17. Tian T, Plowman SJ, Parton RG, Kloog Y, Hancock JF (2010) Mathematical modeling ofK-Ras nanocluster formation on the plasma membrane. Biophys J 99(2):534–543.

18. Pollard TD, Blanchoin L, Mullins RD (2000) Molecular mechanisms controlling actinfilament dynamics in nonmuscle cells. Annu Rev Biophys Biomol Struct 29:545–576.

19. Fritzsche M, Lewalle A, Duke T, Kruse K, Charras G (2013) Analysis of turnover dy-namics of the submembranous actin cortex. Mol Biol Cell 24(6):757–767.

20. Bezanilla M, Gladfelter AS, Kovar DR, Lee W-L (2015) Cytoskeletal dynamics: A viewfrom the membrane. J Cell Biol 209(3):329–337.

21. Bovellan M, et al. (2014) Cellular control of cortical actin nucleation. Curr Biol 24(14):1628–1635.

22. Clark AG, Wartlick O, Salbreux G, Paluch EK (2014) Stresses at the cell surface duringanimal cell morphogenesis. Curr Biol 24(10):R484–R494.

23. Yumura S, et al. (2013) Cell-scale dynamic recycling and cortical flow of the actin-myosin cytoskeleton for rapid cell migration. Biol Open 2(2):200–209.

24. Luo W, et al. (2013) Analysis of the local organization and dynamics of cellular actinnetworks. J Cell Biol 202(7):1057–1073.

25. Biro M, et al. (2013) Cell cortex composition and homeostasis resolved by integratingproteomics and quantitative imaging. Cytoskeleton (Hoboken) 70(11):741–754.

26. Goehring NW, et al. (2011) Polarization of PAR proteins by advective triggering of apattern-forming system. Science 334(6059):1137–1141.

27. Yi J, Wu XS, Crites T, Hammer JA, 3rd (2012) Actin retrograde flow and actomyosin II arccontraction drive receptor cluster dynamics at the immunological synapse in JurkatT cells. Mol Biol Cell 23(5):834–852.

28. Kaizuka Y, Douglass AD, Varma R, Dustin ML, Vale RD (2007) Mechanisms for seg-regating T cell receptor and adhesion molecules during immunological synapse for-mation in Jurkat T cells. Proc Natl Acad Sci USA 104(51):20296–20301.

29. Treanor B, et al. (2010) The membrane skeleton controls diffusion dynamics andsignaling through the B cell receptor. Immunity 32(2):187–199.

30. Paul AS, Pollard TD (2009) Review of the mechanism of processive actin filamentelongation by formins. Cell Motil Cytoskeleton 66(8):606–617.

31. Beach JR, et al. (2014) Nonmuscle myosin II isoforms coassemble in living cells. CurrBiol 24(10):1160–1166.

32. Paluch E, Piel M, Prost J, Bornens M, Sykes C (2005) Cortical actomyosin breakagetriggers shape oscillations in cells and cell fragments. Biophys J 89(1):724–733.

33. Rao M, Mayor S (2014) Active organization of membrane constituents in living cells.Curr Opin Cell Biol 29:126–132.

34. Nédélec FJ, Surrey T, Maggs AC, Leibler S (1997) Self-organization of microtubules

and motors. Nature 389(6648):305–308.35. Murrell MP, Gardel ML (2012) F-actin buckling coordinates contractility and

severing in a biomimetic actomyosin cortex. Proc Natl Acad Sci USA 109(51):

20820–20825.36. Vogel SK, Heinemann F, Chwastek G, Schwille P (2013) The design of MACs (minimal

actin cortices). Cytoskeleton (Hoboken) 70(11):706–717.37. Ideses Y, Sonn-Segev A, Roichman Y, Bernheim-Groswasser A (2013) Myosin II does it

all: Assembly, remodeling, and disassembly of actin networks are governed by myosin

II activity. Soft Matter 9(29):7127–7138.38. Keber FC, et al. (2014) Topology and dynamics of active nematic vesicles. Science

345(6201):1135–1139.39. Sanchez T, Chen DTN, DeCamp SJ, HeymannM, Dogic Z (2012) Spontaneous motion in

hierarchically assembled active matter. Nature 491(7424):431–434.40. Liu AP, Fletcher DA (2006) Actin polymerization serves as a membrane domain switch

in model lipid bilayers. Biophys J 91(11):4064–4070.41. Heinemann F, Vogel SK, Schwille P (2013) Lateral membrane diffusion modulated by

a minimal actin cortex. Biophys J 104(7):1465–1475.42. Honigmann A, et al. (2014) A lipid bound actin meshwork organizes liquid phase

separation in model membranes. eLife 3:e01671.43. Arumugam S, Petrov EP, Schwille P (2015) Cytoskeletal pinning controls phase sepa-

ration in multicomponent lipid membranes. Biophys J 108(5):1104–1113.44. Delatour V, et al. (2008) Arp2/3 controls the motile behavior of N-WASP-functional-

ized GUVs and modulates N-WASP surface distribution by mediating transient links

with actin filaments. Biophys J 94(12):4890–4905.45. Giardini PA, Fletcher DA, Theriot JA (2003) Compression forces generated by actin

comet tails on lipid vesicles. Proc Natl Acad Sci USA 100(11):6493–6498.46. Upadhyaya A, Chabot JR, Andreeva A, Samadani A, van Oudenaarden A (2003)

Probing polymerization forces by using actin-propelled lipid vesicles. Proc Natl Acad

Sci USA 100(8):4521–4526.47. Stuhrmann B, Soares E Silva M, Depken M, Mackintosh FC, Koenderink GH (2012)

Nonequilibrium fluctuations of a remodeling in vitro cytoskeleton. Phys Rev E Stat

Nonlin Soft Matter Phys 86(2 Pt 1):020901.48. Saleh HS, et al. (2009) Properties of an ezrin mutant defective in F-actin binding. J Mol

Biol 385(4):1015–1031.49. Shrivastava R, Köster D, Kalme S, Mayor S, NeerathilingamM (2015) Tailor-made ezrin

actin binding domain to probe its interaction with actin in-vitro. PLoS One 10(4):

e0123428.50. Lenne P-F, et al. (2006) Dynamic molecular confinement in the plasma mem-

brane by microdomains and the cytoskeleton meshwork. EMBO J 25(14):

3245–3256.51. Craig R, Woodhead JL (2006) Structure and function of myosin filaments. Curr Opin

Struct Biol 16(2):204–212.52. Kruse K, Jülicher F (2000) Actively contracting bundles of polar filaments. Phys Rev

Lett 85(8):1778–1781.53. Murrell M, Gardel ML (2014) Actomyosin sliding is attenuated in contractile bio-

mimetic cortices. Mol Biol Cell 25(12):1845–1853.54. Smith D, et al. (2007) Molecular motor-induced instabilities and cross linkers de-

termine biopolymer organization. Biophys J 93(12):4445–4452.55. Nicotera P, Leist M, Ferrando-May E (1998) Intracellular ATP, a switch in the decision

between apoptosis and necrosis. Toxicol Lett 102-103(8):139–142.56. Marchetti MC, et al. (2013) Hydrodynamics of soft active matter. Rev Mod Phys 85(3):

1143–1189.57. Ngo S, et al. (2014) Large-scale chaos and fluctuations in active nematics. Phys Rev Lett

113(3):038302.58. Veatch SL, Polozov IV, Gawrisch K, Keller SL (2004) Liquid domains in vesicles in-

vestigated by NMR and fluorescence microscopy. Biophys J 86(5):2910–2922.59. Alvarado J, Sheinman M, Sharma A, MacKintosh FC, Koenderink GH (2013) Molecular

motors robustly drive active gels to a critically connected state. Nat Phys 9(7):1–7.60. Raghupathy R, et al. (2015) Transbilayer lipid interactions mediate nanoclustering of

lipid-anchored proteins. Cell 161(3):581–594.61. Kaizuka Y, Groves JT (2004) Structure and dynamics of supported intermembrane

junctions. Biophys J 86(2):905–912.62. Kaizuka Y, Groves JT (2010) Bending-mediated superstructural organizations in

phase-separated lipid membranes. New J Phys 12(2010):095001.63. Das A, Polley A, Rao M (2015) Phase Segregation of Passive Advective Particles in an

Active Medium. arXiv:1506.06928v2.64. Chen B, et al. (2014) The WAVE regulatory complex links diverse receptors to the actin

cytoskeleton. Cell 156(1-2):195–207.65. Neisch AL, Fehon RG (2011) Ezrin, Radixin and Moesin: Key regulators of membrane-

cortex interactions and signaling. Curr Opin Cell Biol 23(4):377–382.

Köster et al. PNAS | Published online February 29, 2016 | E1653

BIOPH

YSICSAND

COMPU

TATIONALBIOLO

GY

APP

LIED

PHYS

ICAL

SCIENCE

SPN

ASPL

US

Dow

nloa

ded

by g

uest

on

Oct

ober

29,

202

0

66. Chaudhuri A, Bhattacharya B, Gowrishankar K, Mayor S, Rao M (2011) Spatiotemporalregulation of chemical reactions by active cytoskeletal remodeling. Proc Natl Acad SciUSA 108(36):14825–14830.

67. Iyengar G, Rao M (2014) A cellular solution to an information-processing problem.Proc Natl Acad Sci USA 111(34):12402–12407.

68. Surrey T, Nedelec F, Leibler S, Karsenti E (2001) Physical properties determining self-organization of motors and microtubules. Science 292(5519):1167–1171.

69. Spudich JA, Watt S (1971) The regulation of rabbit skeletal muscle contraction. I.Biochemical studies of the interaction of the tropomyosin-troponin complex withactin and the proteolytic fragments of myosin. J Biol Chem 246(15):4866–4871.

70. Pollard TD (1982) Myosin purification and characterization. Methods Cell Biol 24:333–371.

71. Pollard TD (1982) Structure and polymerization of Acanthamoebamyosin-II filaments.J Cell Biol 95(3):816–825.

72. Pollock N, Koonce MP, de Hostos EL, Vale RD (1998) In vitro microtubule-based or-

ganelle transport in wild-type Dictyostelium and cells overexpressing a truncated

dynein heavy chain. Cell Motil Cytoskeleton 40(3):304–314.73. York AG, et al. (2013) Instant super-resolution imaging in live cells and embryos via

analog image processing. Nat Methods 10(11):1122–1126.74. Axelrod D, Koppel DE, Schlessinger J, Elson E, Webb WW (1976) Mobility measure-

ment by analysis of fluorescence photobleaching recovery kinetics. Biophys J 16(9):

1055–1069.75. Rezakhaniha R, et al. (2012) Experimental investigation of collagen waviness and

orientation in the arterial adventitia using confocal laser scanning microscopy.

Biomech Model Mechanobiol 11(3-4):461–473.76. Thielicke W, Stamhuis EJ (2014) PIVlab – Towards User-friendly, Affordable and Accurate

Digital Particle Image Velocimetry in MATLAB. J Open Res Softw 2:1–10.

E1654 | www.pnas.org/cgi/doi/10.1073/pnas.1514030113 Köster et al.

Dow

nloa

ded

by g

uest

on

Oct

ober

29,

202

0