Plasma Membrane-Cortical Cytoskeleton Interactions: A Cell Biology Approach with Biophysical Considerations

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Plasma Membrane—Cortical CytoskeletonInteractions: A Cell Biology Approach withBiophysical ConsiderationsAndras Kapus*1 and Paul Janmey2

ABSTRACTFrom a biophysical standpoint, the interface between the cell membrane and the cytoskeleton isan intriguing site where a “two-dimensional fluid” interacts with an exceedingly complex three-dimensional protein meshwork. The membrane is a key regulator of the cytoskeleton, whichnot only provides docking sites for cytoskeletal elements through transmembrane proteins, lipidbinding-based, and electrostatic interactions, but also serves as the source of the signaling eventsand molecules that control cytoskeletal organization and remolding. Conversely, the cytoskeletonis a key determinant of the biophysical and biochemical properties of the membrane, including itsshape, tension, movement, composition, as well as the mobility, partitioning, and recycling of itsconstituents. From a cell biological standpoint, the membrane-cytoskeleton interplay underlies—as a central executor and/or regulator—a multitude of complex processes including chemical andmechanical signal transduction, motility/migration, endo-/exo-/phagocytosis, and other formsof membrane traffic, cell-cell, and cell-matrix adhesion. The aim of this article is to provide anoverview of the tight structural and functional coupling between the membrane and the cytoskele-ton. As biophysical approaches, both theoretical and experimental, proved to be instrumental forour understanding of the membrane/cytoskeleton interplay, this review will “oscillate” betweenthe cell biological phenomena and the corresponding biophysical principles and considerations.After describing the types of connections between the membrane and the cytoskeleton, we willfocus on a few key physical parameters and processes (force generation, curvature, tension, andsurface charge) and will discuss how these contribute to a variety of fundamental cell biologicalfunctions. C© 2013 American Physiological Society. Compr Physiol 3:1231-1281, 2013.

IntroductionThe structural and functional interconnectedness of theplasma membrane and the cytoskeleton has been long recog-nized, inasmuch that by the early 1990s this unity was concep-tualized as the “membrane-cytoskeleton trilayer” (303). Theinterface between the essentially two-dimensional (2D) andfluid-like membrane and the three-dimensional (often gel-likeyet highly dynamic) cytoskeleton is a biochemically and bio-physically intriguing locus that has emerged as a critical reg-ulator of a variety of cellular functions, including shape deter-mination, migration (edge protrusion), adhesion, cell division,endo/exocytosis, and environmental sensing, together withthe ensuing signal transduction associated with all these pro-cesses (112). Moreover, classic studies aimed at the biochem-ical characterization of membrane/cytoskeleton interactions(predominantly in the context of red blood cells) uncoveredthe existence of a “membrane skeleton,” that is, a more-or-lesstwo-dimensional meshwork, tightly connected to and paral-lel with the plasmalemma, which can be considered as themembrane’s “own” cytoskeleton (160). The discovery andcharacterization of this spectrin/ankyrin-based 2D network,first identified in erythrocytes, but present in most cell types,

and not only in the plasma membrane (Fig. 1) has provided agreat impetus for the investigation of membrane-cytoskeletoninteractions, because it revealed that inherited or acquiredalterations in the membrane skeleton can cause major defectsin cell shape, cell deformability, and transmembrane ion trans-port, which in turn are key pathogenic factors in a varietyof human diseases including hematologic disorders (hered-itary spherocytosis, ellipotocytosis, and malaria), cardiacarrhythmias, and neurological pathologies (spinocerebellarataxia) (39).

The complexity arises from the plethora of interactionsamong the lipid and protein components of the membrane,the 2D membrane skeleton, and the broader 3D cytoskeleton

*Correspondence to [email protected] Keenan Research Center in the Li Ka Shing Knowledge Institute ofthe St. Michael’s Hospital and Department of Surgery, University ofToronto, Ontario, Canada2 Institute for Medicine and Engineering (IME), University ofPennsylvania, Philadelphia, PennsylvaniaPublished online, July 2013 (comprehensivephysiology.com)DOI: 10.1002/cphy.c120015Copyright C© American Physiological Society

Volume 3, July 2013 1231


Plasma Membrane—Cortical Cytoskeleton Interactions Comprehensive Physiology

Figure 1 The two-dimensional spectrin-based membrane skeleton of red cells and its relationship to the three-dimensionalactin cytoskeleton. The triple α-helical coiled-coil structure of spectrin repeats is shown in the magnified box. Spectrin is linkedto the membrane by three mechanisms (i) the pleckstrin homology (PH) domain in β-spectrin directly binds to anionic lipidsof the inner leaflet. (ii) A variety of transmembrane proteins (including the anion exchanger (AE), and the Rhesus factor (Rh)bind to ankyrin, which interacts with α-spectrin. (iii) Transmembrane proteins also associate with another adapter complex,containing adducin, which directly or through various other connectors (4.1, 4.2, p55, and dematin) link spectrin to thecomponents of the actin skeleton, including actin, tropomyosin, and tropomodulin. Further abbreviations: GPA, GPB, andGPC glycophorin A, B, and C, respectively; GLUT 1, glucose transporter 1; RhAG, Rh-associated glycoprotein. Reproduced,with permission, from (267).

(Fig. 1). Considering the protein components of these systems,according to a conservative estimate, 15% of the 30,000 genesin the human genome encode membrane proteins (i.e., 4500genes), which translates to several-fold more polypeptides,due to alternative splicing (7). The potential interactive part-ner, that is, the cytoskeleton is composed of three distinct ele-ments: the microfilament system represented by 6 actins (with6 functional genes and > 20 pseudogenes) and 70 families ofactin-binding proteins; the microtubule (MT) system, includ-ing six α- and β-tubulins and more than a dozen families ofmicrotubule-binding proteins; and the intermediate filament(IF) system, which are encoded by 70 different genes and5 comprises families of associated proteins (121, 347). Thistally does not yet take into account the motor proteins asso-ciated with the microfilaments and microtubules (e.g., thoseencoded by ≈ 40 myosin and 40 kinesin genes and the largefamily of multisubunit dyneins) or the very large number ofregulatory/signaling proteins that control cytoskeleton andmembrane remodeling. Finally, a plethora of proteins candirectly interact with lipid components of the membrane viavarious lipid-binding domains (see Table 1 for prominent

examples). Clearly, the potential number of bi- and multilat-eral interactions between membrane and cytoskeleton com-ponents is enormous. Cognizant of this challenging fact, herewe will use an approach that concentrates on integrated func-tions or complex properties of the membrane/cytoskeletoninteractions, as viewed form a cell biological and biophysi-cal standpoint. Thus, after a summary of the various typesof the structural connections between the membrane andthe cytoskeleton and a brief overview of the 2D-membraneskeleton, this review will focus on the following functionalaspects: (i) force generation and membrane protrusion at thecell periphery; (ii) membrane curvature; (iii) membrane ten-sion; and (iv) surface charge. While the focus will be onthe aforementioned processes and functions we will mentionsome of the key methodologies that were used to obtain themain findings. Like any overview of a vast rapidly developingfield, this article is bound to be selective in its scope and lim-ited in its depth. Despite these limitations, we hope to providea bird’s eye view of the field, with its major achievementsand ongoing debates, which can orient the interested readertoward more detailed and specialized information.

1232 Volume 3, July 2013


Comprehensive Physiology Plasma Membrane—Cortical Cytoskeleton InteractionsTa

ble

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Plasma Membrane—Cortical Cytoskeleton Interactions Comprehensive PhysiologyTa

ble

1(C

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Comprehensive Physiology Plasma Membrane—Cortical Cytoskeleton InteractionsM

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Types of Connections between theMembrane and the CytoskeletonGiven the enormous crowding within the cell, where the sol-vent water occupies only around half of the volume and atypical large solute like a protein is less than a nm away fromits nearest neighbor, random collisions between elements ofthe cytoskeleton and the vast surface area of lipid bilayers,that is, random contacts between the 3D cytoskeleton and the2D membrane are likely to be very frequent. Whether thesecollisions lead to stable bonds or reactions is highly depen-dent on the type of protein, the type of lipid, and the stateof signaling. The filament-forming proteins of the cytoskele-ton, actin, tubulin, and IF subunits such as vimentin are byfar the most abundant proteins in the cell, but the filamentsthey form are generally not considered to form stable bondswith the membrane. Instead, the large array of actin-bindingproteins and other cytoskeletal linkers and regulators makespecific bonds to both protein and lipid constituents of themembrane, and often there are two or more proteins in seriesor in parallel mediating the cytoskeleton/membrane interface.

From a functional standpoint such membrane-cytoskeleton interactions are critically important as theyunderlie or contribute to a multitude of cellular functions,including receptor clustering (12, 207), signal transduction(37, 234, 437), curvature and shape changes (425), mem-brane traffic (454, 489), endo- and exocytosis (322), motility(377, 387), mechanotransduction (188) cytokinesis and celldivision (179, 313, 348), as well as cell-cell (200, 447) andcell-matrix attachment (18, 238). In principle, the linkagesbetween the cytoskeleton and membrane can be realized bythree different ways (and their combinations).

1. The cytoskeletal elements (e.g., polymers of actin, tubulin,and IF proteins) themselves interact with the lipid phaseof the membrane.

2. The cytoskeletal elements are linked to transmembraneproteins either directly or indirectly through adapter pro-teins.

3. The cytoskeletal elements are connected to the membranevia proteins that bind the inner leaflet of the plasmalemma(or the cytosolic leaflet of membranes of organelles).

Before providing a few illustrative examples of these inter-actions, it is worth mentioning that while this classificationis didactically useful, it is somewhat arbitrary. This is sobecause (i) the same protein often uses more than one type ofinteractions (e.g., β-spectrin is linked to the transmembraneanion exchanger protein through ankyrin, but it also containsa pleckstrin homology (PH) domain with which it can directlybind to membrane phospholipids (497); (ii) many membrane-interacting proteins (e.g., the Rho family regulators guaninenucleotide exchange factors) do not physically bridge the

membrane to the cytoskeleton but their topical activity hasa major impact on submembraneous cytoskeleton organiza-tion; and (iii) the concept of “cytoskeletal protein” is hardto define, and it remains a question of convention whetherthe large variety of proteins involved in membrane fusion(e.g., SNARES) and fission (e.g., the motor protein dynamin)belong to this category. Nonetheless, a brief description of themajor types of interactions according to the main categoriesis warranted. We will first review the direct interactions ofcytoskeletal polymers with the membrane, then provide someexamples of cytoskeletal interactions with transmembraneproteins, that can link the cytoskeleton to receptors, bridgeit to the extracellular matrix (ECM), or connect one cell toanother thereby forming transcellular cytoskeleton networks.Finally, we will discuss the interaction between membranelipids and various cytoskeletal elements (cytoskeletal poly-mers, other structural proteins, or cytoskeleton regulators),focusing on interactions between phosphatidylinositol (PI)4, 5 bisphosphate [PtdIns(4,5)P2; referred to as PIP2 in thisarticle], the functionally most important lipid, and the corre-sponding protein partners through well-defined lipid-binding(PH, PX, FYVE, BAR, and FERM) domains.

Binding of cytoskeletal subunits and filamentsto membranesThe prevailing models of the membrane skeleton usuallyascribe a series of coordinated events mediated by numerousproteins that link cytoskeletal filaments to the lipid bilayerbut a large body of biochemical and cell biological evidencesuggests that subunits and filaments of all three cytoskeletonpolymer types can bind membrane lipids, at least transiently.Such evidence might also warrant reconsideration. Numer-ous biochemical and biophysical studies report the binding ofactin, tubulin, and IF subunits with phospholipids and glycol-ipids of the plasma membrane.

In vitro studies in which actin is polymerized either on thesurface or within phospholipid vesicles show that the lipidbilayer can adsorb actin filaments and affect their organiza-tion (443). When lipids characteristic of the cytoplasm-facingleaflet of the plasma membrane, such as phosphatidylser-ine (PS) or phosphatidylethanolamine are contained in thebilayer, the adsorption of F-actin is stabilized by divalentcations, in particular Mg2+ at close to physiological concen-trations (175). Actin has also been found tightly bound tocell membrane preparations, suggesting that there might beconditions where actin interacts with the lipid bilayer in theabsence of intermediary protein linkers. Not only might mem-brane lipids alter actin localization and assembly, but perhapsmore intriguingly recent studies suggest that polymerizationof actin at or near the membrane alters the lateral distribu-tion of membrane lipids (115), possibly with consequencesfor the way in which proteins on the outer surface of the cellare arranged. In vitro studies show that adsorption of F-actinto liposomes containing PS and cholesterol alters the size ofcholesterol-dependent lipid domains (146). In a chemically



Comprehensive Physiology Plasma Membrane—Cortical Cytoskeleton Interactions

more complex system, assembly of branched actin networksmediated by the actin-related protein 2/3 (Arp2/3) complexand initiated by the Neuronal Wiscott-Aldrich Syndrome pro-tein N-WASP on the surface of vesicles containing PIP2 leadsto formation of membrane domains coincident with sites ofactin polymerization (259). Such studies might relate to thefinding that depolymerization of submembrane actin in cellscauses loss of membrane domains, often termed lipid rafts,and dissociation of protein clusters linked to the outer mem-brane leaflet through lipid anchors (157). Recently, it hasbeen examined whether electrostatic interactions (e.g., repul-sion) between the negatively charged PIP2 and actin mightdirectly contribute to the accumulation of PIP2 in certainactin-delimited membrane regions, such as the phagocyticcup. The diffusion rate of PIP2is much less in (or out of) thecup than in the surrounding plasmalemma, as verified by flu-orescence recovery after photobleaching (FRAP). However,computer simulations showed that despite the close proximityof actin to the membrane (< 1 nm) and its overall negativecharge, it may play only a minor role in limiting PIP2 move-ment, partly because there is charge shielding by mobile ionsand partly because actin also contains bands of positivelycharged residues (152). The connection between actin andother cytoskeletal polymers directly beneath the cell mem-brane and formation of lipid and protein clusters on bothsides of the lipid bilayers is one of the more enigmatic effectsof the membrane cytoskeleton.

Tubulin and some MT-associated proteins associate selec-tively with purified liposomes (68), and tubulin is foundtightly associated with cell membrane fractions and lipidspurified from cells (43,44, 370). Membrane binding is furthersupported by the finding that tubulin can be palmitoylated, aposttranslational modification that generally promotes inser-tion of the modified protein into the lipid bilayer (520). Thecellular function of membrane tubulin is not altogether clear,and the various hypotheses and evidence for membrane asso-ciated tubulin assembly are discussed in a recent review (507).

IFs likewise show a range of interactions with lipids inthe form of membrane bilayers as well as intracellular lipiddroplets. Traub and colleagues clearly demonstrated the bio-chemical interaction of the IF protein vimentin with puri-fied lipid bilayers that contained anionic phospholipids (340).Further studies show that vimentin interactselectrostaticallythrough its positively charged N-terminus, but also insertsinto the hydrophobic domain of the bilayer. Newly synthe-sized vimentin in a reticulocyte lysate associated tightly withadded cell membrane fractions and once bound to the mem-brane became resistant to denaturation by urea (363). In livecells such as differentiating adipocytes, vimentin forms a cagearound lipid droplets apparently associated with the dropletsurface (134). The ganglioside GM2 has also been reportedto bind vimentin at a high GM2/vimentin molar ratio and tobind tightly and selectively enough to be copurified throughimmunoprecipitation from cell extracts, although the condi-tions under which GM2 would be exposed to the intracellularcompartment under normal conditions are unclear. These data

support the hypothesis that vimentin and probably other IFsbind to the membrane where in electron micrographs they areoften observed in close association (227) suggesting that thisinteraction might occur by direct IF-lipid binding rather thanbeing mediated by another membrane bound protein. The invivo function of such binding is not yet clear but the in vitrodata suggest that this interaction deserves further study.

Membrane protein—cytoskeleton interactionsThe 3D cytoskeleton binds to multiple classes of transmem-brane proteins that act as receptors and/or adhesion sites forcell-ECM or cell-cell attachment. Before providing some con-crete examples in each category, we consider the general roleof the cytoskeleton in membrane organization. In fact, therecognition of such a role led to the reassessment and mod-ification of the classic Singer-Nicholson fluid mosaic modelof membrane architecture, and created a new paradigm inmembrane biology. For over 30 years, it has been a well-known (and for a long time neglected) fact that the diffu-sion of transmembrane proteins as well as membranelipidsis—on average—20-fold slower in real plasma membranesthan in artificial lipid bilayers such as liposomes [reviewedin (233, 236, 380)]. This finding is incompatible with theview that the membrane is 2D fluid, the components of whichperform random walk in a Brownian manner. Instead, themembrane should contain significant diffusion barriers, whilelarge protein aggregates or cholesterol-rich areas may slowdown diffusion, these by no means explain a 20-fold differ-ence (137, 233). Instead, it has been proposed that the mem-brane is constitutionally compartmentalized (235, 380). Bothmorphological and functional data provide strong support tothis concept. Namely, 3D reconstitution of the membraneskeleton/plasma membrane interface by electron tomogra-phy reveals the presence of parceled membrane areas bor-dered by actin filaments, interspersed with clathrin-coatedpits and caveolae (304) (Fig. 2A). Moreover, the develop-ment of high-speed single particle-tracking techniques pro-vided direct evidence that “free” diffusion occurs only withinconfined membrane compartments, termed as corrals, with adiameter of 40 to 100 nm. (233, 236). Beside the compatiblemorphology of the submembraneous cytoskeleton meshwork,functional measurements also suggest that membrane corralsare defined primarily by the membrane skeleton. Thus, dis-ruption of the actin skeleton enhances the overall mobilityof membrane proteins, and the diffusion coefficients foundin membrane blebs, that is, cytoskeleton-deprived membraneprotrusions, correspond to those found in liposomes (137).Based on such observations, Kusumi and colleagues proposedthe fence-and-picket model of membrane organization (233)(Fig. 2B). Submembraneous cytoskeletal filaments representthe fences that border membrane domains. Within these com-partments TM proteins move freely. However, their diffusionacross the fences is a relatively rare event, because the cytoso-lic tails of TM proteins collide with the fences. Intercompart-mental movement, termed as “hop diffusion,” occurs when




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Figure 2 The fence-and-picket model of membrane organization and the three-tiered mesoscalemodel of the plasma membrane. (A) The undercoat structure of the cytoplasmic surface of the plasmamembrane. Rapid-freeze, deep-etch electron microscopy was performed in normal rat kidney fibroblasts(a) and fetal rat skin keratocytes (b). Note the clathrin-coated pits (arrows), a caveola (∗), and the densemeshwork of fibers of the cortical actin skeleton. The latter is thought to be the structural basis ofthe corralled movement of membrane proteins; the fibers likely correspond to the fences limiting thediffusion of transmembrane proteins. Bar 100 nm, and in the inset: 50 nm. Reproduced, with permission,from (304). (B) The fence-and-picket model: the membrane skeleton (MSK) forms fences that enclosemembrane compartments. Within these sectors membrane components perform random walk (thezigzag lines indicate trajectories). Transmembrane proteins are anchored to the fence (pickets); a, b,and c represent side, bottom, and top views, respectively. (C) The three-tiered mesoscale model of themembrane. The first tier corresponds to the fences and pickets formed by the membrane skeleton. Thesecond and third tiers correspond to lipid rafts and dynamic protein complexes, respectively. B and Care reproduced, with permission, from (233).

the membrane skeleton is remodeled or temporarily separatedfrom the plasma membrane (137). According to this model,the membrane is characterized by two diffusion coefficients:one within the compartments (so called “microscopic” coef-ficient) that is valid for short distances (in the order of 10 nm)

and has a value of 5 to 10 μm2/s, in full agreement withthe Singer-Nicholson model; and another one (the so-called“macroscopic” coefficient), which is valid for the membraneas a whole, and exhibits values in the range of 0.2 to 0.5μm2/s, compatible with previous “bulk” measurements (233).




However, two observations necessitated further refinement ofthe model: first, certain TM proteins are nearly immobile, asverified by single molecule tracking or florescence recoveryafter photobleaching measurements (396). Second, lipids ofthe outer leaflet also show confined movement (137). Sincethese cannot “bump” into the submembraneous cytoskele-ton, other mechanisms had to be evoked. Both phenomenacan be explained by the existence of pickets, that is, TMproteins tethered to the fences. These fixed rods representdiffusion barriers for lipids as well. Moreover, they influencethe local distribution of lipids, because their transmembranesegments sterically inhibit the accumulation of cholesterol intheir immediate vicinity (236). Thus, according to this model,the cytoskeleton partitions the membrane both in terms of pro-tein and lipid distribution and movement. The far-reachingfunctional consequences of the fence-and-picket model arebeyond the scope of this article, and the interested readeris referred to excellent recent reviews (233, 234, 236). It isworth emphasizing, however, that such compartmentalizationcan segregate or concentrate elements of various signalingmodules (e.g., receptor subunits), and thereby it can exert amajor influence on the localization, duration and intensity oftransmembrane signaling.

Recently, Kusumi and colleagues put forward an evenmore comprehensive theory about the molecular organizationof the plasma membrane. They postulate that the membraneexhibits a hierarchical, three-tiered mesoscale (2-300 nm)domain architecture (233,234, 236) (Fig. 2C). The first tier isthe membrane compartmentalization by the actin-based mem-brane skeleton, as described above. This operates in the 40to 300 nm range. The second tier is the domains of mem-brane rafts, that is, cholesterol-rich microdomains, the size ofwhich is usually in the 2 to 20 nm range. The third tier is thedomain of dynamic protein complexes (e.g., coat proteins andscaffolding proteins) with an average diameter of 3 to 10 nm.While the cytoskeleton is primarily associated with the firsttier, evidence is accumulating that it exerts a major impact onthe other tiers as well (234, 236). Such overall organization ofthe cell surface may be one of the most significant functionsof the membrane cytoskeleton.

The following sections will provide examples of the inter-action between the cytoskeleton and various functional groupsof membrane protein, and will illustrate the above-mentionedconcepts with some experimental examples.

“Classic” transmembrane receptors

Many transmembrane receptors can—at least transiently—associate with the cytoskeleton. The epidermal growth factorreceptor (EGFR) has been shown to directly bind to actin (98),and this binding modulates both the affinity of the receptorfor its ligand and the ensuing signal transduction (379, 504).Single molecule tracking experiments revealed that EGFRdimerization, which facilitates EGF binding, occurs pref-erentially at the cell periphery. Peripheral accumulation of

EGFR dimers requires an intact actin skeleton (82). Con-versely, engagement of the EGFR by EGF initiates signalingthat leads to submembraneous actin polymerization, whichinduces the formation of protrusive structures (e.g., lamel-lipodia) at the cell periphery (214, 378). Thus, membraneprotein/cytoskeleton interactions result in regulatory circuitsthat can fine-tune signal transduction (379). Other importantexamples for transmembrane receptor/cytoskeleton linkagescan be found in the immunological synapse, the interfacebetween antigen presenting cells and lymphocytes. Here, theconnection is indirect in that the receptors do not bind to thecytoskeleton, but are linked to actin filaments through variousadaptors. These interactions regulate the dynamics of mem-brane rafts by controlling their tethering and trapping (495).Specifically, when activated, CD28, a transmembrane pro-tein that transmits costimulatory signals in T-cells, was foundto bind the actin-cross-linking protein filamin, which in turnbinds to the actin meshwork. This association is necessary toensure the recruitment of membrane rafts (containing CD28and the T cell receptor) to the immunological synapse (468).Release and subsequent coalescence of membrane rafts is alsocontrolled by regulated receptor/cytoskeleton interactions. Asan example, PAGI a raft-resident transmembrane protein inB cells is anchored to the actin filaments through the corti-cal actin-binding regulatory protein, ezrin. Upon stimulation,ezrin becomes dephosphorylated, which results in the releaseof PAGI from the cytoskeleton, allowing raft untethering andthereby greater raft mobility (167).

Even without direct binding, the cytoskeleton has a majorrole in controlling receptor diffusion, coalescence, and the for-mation of functional clusters in the membrane (12, 92, 157).For example, using single molecular tracking Jaqaman andcolleagues (207) have recently shown that CD36 (a receptorfor several ligands, including oxidized low-density lipopro-tein) exhibits cytoskeleton-restricted motion in the plane ofthe membrane. A subpopulation of receptors diffused withinlinear confinement regions, in which their freedom of move-ment was facilitated, whereas diffusion outside (perpendicularto) these linear trajectories was prohibited (“1D diffusion,”Figure 3) Motion along these preformed “molecular high-ways” was perturbed by disruption of the microfilaments orMTs. While the molecular mechanism whereby the cytoskele-ton controls receptor diffusion remains to be elucidated, theauthors propose two possibilities. In the first model, the sub-membraneous isotropic actin meshwork (which limits recep-tor diffusion) is perturbed by the presence of microtubules,leading to the formation of actin-delimited channels. Alterna-tively, the isotropic nature of the skeleton is not modified, butMTs induce local detachment of the submembraneous actinskeleton from the membrane. In any case, these examplesshow that subtle regulation of receptor movement and clus-tering do not need to depend on highly specific intermolecularinteractions; in fact this confinement, realized through randomcollisions between cytoskeletal constituents and transmem-brane proteins, may constitute a key regulatory mechanism inreceptor signaling.




Figure 3 Cytoskeleton-controlled diffusion of the scavenger receptor CD36 in humanmacrophages. The movement of antibody-labeled CD36 molecules on the cells surface was followedby particle tracking. Two measures were used to define trajectory types: the first classified trajectoryshape based on the degree of anisotropy of the scatter of particle positions along a trajectory, whilethe second extracted diffusion types using a moment scaling spectrum analysis of particle displace-ments. Trajectories, shown in (A), were classified as linear, isotropic confined, isotropic unconfined,and undetermined (B). More than 25% of the trajectories corresponded to a linear path, and morethan 75% of the particles followed linear or otherwise confined routes. A linear trajectory as moni-tored by Qdot-labeled CD36 is shown in C. The trajectory was reconstructed based on a movie with1184 frames taken in 18.9 s (the time is color coded). Scale bar 200 nm. The histogram shows thedistribution of receptor positions across the width of the linear trajectory. These findings indicate thepresence of preformed diffusion pathways in the membrane. Adapted, with permission, from (207).

Integrins

Integrins represent another important set of transmembranemolecules involved in membrane-cytoskeleton interactions.These heterodimeric proteins [composed of one of 18 α andone of 8 β-subunits, forming 24 identified heterodimers inmammals (136)] function as receptors, which connect theECM (or attachment molecules on neighboring cells) to thecytoskeleton. In contrast to “classic” membrane receptors,integrin-mediated signaling is bidirectional (410): integrinscan be activated through engagement by extracellular lig-ands (outside-in signaling) or/and by intracellular signals thatemanate from the cytosol or the cytoskeleton, and in turnalter the state and sensitivity of integrins (inside-out signal-ing). These extra- and intracellular cues impact the affinity ofintegrins for their extracellular partners as well as their cluster-ing, mobility and traffic (274). Integrins transmit and are reg-ulated by mechanical stimuli (e.g. tension) and serve as majormechanosensors. Accordingly, focal contacts and focal adhe-sions (FAs), that is, specialized, integrin-rich bridges betweenthe ECM and the cytoskeleton, function as mechanochemi-cal signaling hubs that affect many aspects of cell behavior,

including movement, survival, cell division, morphogenesis,and tissue remodeling (225). Integrins are tightly connectedto the actin skeleton, but they do not directly bind to actin.Instead, more than 50 proteins have been implicated to con-stitute the integrin/actin skeleton interface (94). (The recentlydescribed, functionally layered molecular architecture of FAsis discussed in more detail in the context of matrix attach-ment during membrane protrusion (see Section “Adhesions”).Several proteins, including parvin, paxillin, tensin, filamin,α-actinin, and talin can bind both the cytoplasmic tail of vari-ous integrin β-chains and actin (94, 505). These proteins can(in principle) anchor integrins to the actin skeleton, but theconnections are certainly not limited to such single-adaptorlinkages. The dynamics of these integrin-adaptor-actin inter-actions with regards to the various integrin-induced func-tions is an active area of current research. Strong evidencesupports the role of talin both in inside-out signaling andin mechanotransduction (312). Talin binding to a cytosolicregion of β-integrins induces a transmembrane conforma-tional change, which increases integrin affinity for externalligands (463). The membrane itself plays an important role




in this process because in addition to a membrane-proximalsalt bridge between the integrin tail and talin, the interac-tion of a positively charged patch of talin with membranephospholipids is also required for integrin activation (13, 88).Indeed, talin contains well-defined lipid-binding sequences(FERM domain, see “Membrane—cytoskeleton interactionsvia lipids”), and PIP2 was shown to alter talin’s conforma-tion (64, 275). Talin also works as a mechanotransducer:using magnetic tweezers, total internal reflection (TIRF) andatomic force microscopy (AFM), the Sheetz lab has demon-strated (93) that stretching a single talin rod with physiolog-ically relevant forces (2-12 pN) exposed cryptic-binding sitefor the FA protein vinculin (59), and enhanced talin-vinculininteraction. Vinculin in turn increases the strength and sta-bility of cell matrix attachments, primarily by immobilizingtalin, which enhances integrin clustering, and by promotingFA linkage to the actin skeleton (67, 193, 401) This resultsin decreased integrin mobility and slower cell movements.Conversely, integrin turnover is facilitated by PIP2 bindingto vinculin (73), possibly because this lipid competes for anactin-binding site and uncouples vinculin from the cytoskele-ton (339, 523). In short, both the protein and lipid componentsof the membrane/cytoskeleton interface regulate the dynamicsof cell-matrix attachments.

The state of the integrin-talin-vinculin axis has an impor-tant role in membrane dynamics as well. Specifically, focalcontacts work as a clutch: when they are “engaged” that is,linked to the cytoskeleton, actin polymerization will push themembrane forward (see Section “Adhesions”) (although toostrong attachment per se will reduce cell locomotion); in con-trast when the clutch is disengaged, polymerization results inretrograde actin flow (437). Moreover, vinculin not only bindsand bundles actin filaments (206, 209, 211), but also promotesactin polymerization. This occurs probably by two distinctmechanisms: vinculin binds monomeric actin and promotesits polymerization (502) and also interacts with and possi-bly activates the Arp2/3 complex (96, 324) a major actin-polymerizing machine (see Section “Actin polymerizationdrives membrane protrusions and intercellular movements ofpathogens”). Taken together, these examples illustrate thatintegrin activation and cytoskeleton organization are inter-dependent. While the cytoskeleton (e.g., through mechanicalforces) impacts the state of integrins, the activation of inte-grins, through a multitude of membrane-associated signalingcomplexes, exerts two major effects on the actin skeleton:it regulates the capture of actin filaments and it modulatesactin nucleation and polymerization. As the description of theunderlying very complex (and not fully understood) mecha-nisms is beyond the scope of this review, the interested readeris referred to excellent reviews (3, 17, 45, 65, 413).

Of the various cytoskeletal networks, integrins interactsprimarily with the actin skeleton, with the notable exceptionof α6β4 integrin in the hemidesmosomes, which connectscomponents of the basement membrane (laminin) throughadaptor proteins (plectin) to IFs (keratin) (258). Integrinshave not been shown to directly interact with MTs, and

MTs often appear to destabilize adhesion complexes whenthey extend to make contact. Although, unlike microfila-ments, MTs are not structurally coupled to integrins, thereis widespread functional interplay between these entities: forexample, integrins regulate Rho family GTPases, which influ-ence not only microfilaments but also microtubule dynamics(97). Conversely, MTs interact with and regulate importantsignal-transducing molecules, which impact actin dynamicsand integrin function. For example, GEF-H1, a Rho/and Racguanine nucleotide exchange factor has been shown to asso-ciate with MTs, and its dissociation from the tubulin networkresults in enhanced Rho signaling, and consequently increasedactin polymerization, contractility, and cellular tension (46,74, 230).

Cell-cell junctions

Finally, we briefly mention yet another major site for trans-membrane protein-cytoskeleton interactions, namely, the var-ious intercellular contacts, including tight and adherens junc-tions (TJs and AJs) and the desmosomes. The transmem-brane components of these structures, for example, claudins,occludin, and junctional adhesion molecules for the TJs (89,131, 297, 432, 476), cadherins for the AJs (29, 325, 422), anddesmoglein and desmocollin for desmosomes (95) are con-nected to their usually homotypic counterpart on the neigh-boring cell in the extracellular space, and to the cytoskeletonvia so-called plaque proteins in the intracellular space. Thesecytoskeleton-interacting plaque proteins include members ofthe Zonula occludens family (28, 430) for TJs and AJs, cin-gulin for TJ (91, 297), various catenin family members (α,β, and γ ) for AJs and (173), desmoplakin, desmophillin, andγ -catenin for desmosomes (451). Cytoskeletal anchorage ofthese membrane attachment loci is critically important forthe proper function of the junctions, which includes (i) themaintenance and regulation of the permeability barrier andparacellular transport (TJ); (ii) the segregation of the apicaland basolateral membrane compartments (diffusion barrierfor lipids and protein through the TJs) and thereby the main-tenance of cell polarity; and (iii) the mechanical stability andstrength of intercellular contacts, necessary for proper tissueorganization (AJs and desmosomes).

The cytoskeletal attachment and mobility of transmem-brane cell adhesion molecules can be quantitatively moni-tored by biophysical methods. To illustrate this point, it isworth mentioning the elegant experiments of Sako and col-leagues (396), who used single particle tracking and opticaltweezers to characterize the movement of wild-type (WT) E-cadherin and various E-cadherin mutants, which either lackedthe catenin/cytoskeleton-interacting regions or were perma-nently coupled to α-catenin and thereby to the actin skele-ton (Fig. 4). The extracellular part of E-cadherin moleculeswere labeled with colloidal gold- or latex-coupled antibod-ies, which were then pulled by optical tweezers. The resultsshowed that WT E-cadherin exists in two major populations,




Figure 4 The intramembrane movement of a transmembrane protein (E-cadherin).Panel I: E-cadherin can either be tethered to the cytoskeleton or corralled withincytoskeleton-delimited fences (A). Experiments using optical tweezers as shown in Acan differentiate between these modes. Using trapping forces of ≈1 pN, tetheredmolecules can only be dragged along to the extent allowed by the stretching of thecytoskeletal anchors. Corralled molecules move freely within compartments delimitedby the fences, and smaller forces (0.05-0.1 pN) might be sufficient to drag themthrough the fence. Panel II: single molecular tracking of wild-type (WT) E-cadherin andvarious E-cadherin mutants. Catenin minus lacks the α-catenin (a cytoskeletal linker)binding site, short-tailed lacks almost the entire cytosolic tail, whereas in fusion E-cadherin is covalently linked to α-catenin. Fusion represents one extreme with stronglylimited diffusion, whereas short tailed is the most diffusible species. III: interpretationof the results shown in II. Fusion is tightly linked to the cytoskeleton. WT exists in varioussubpopulations: it can be linked to the cytoskeleton, may be linked to α-catenin butnot to the actin network or may be free of α-catenin, resembling catenin minus. Theleast restricted is short tailed, which moves relatively freely not only within but alsoacross the fences. (A, B, C, and D represent the four mobility states.) Adapted, withpermission, from (396).

which were designated as either tethered or corralled. Approx-imately half of the WT molecules could be dragged onlyfor very short distances (0.14 μm) before escaping the opti-cal trap, and had very low microscopic diffusion coefficients(D = 0.2 × 10−12 cm2/s), similar to the cytoskeleton-attachedmutants. These molecules were tethered. The other half exhib-ited ≈ fivefold to tenfold higher values for both parameters.However, even these more mobile molecules were confined tomove within a limited membrane area (≈ 0.13 μm2). Thesewere corralled within membrane skeleton-delimited fences.When pulled, molecules could hop over the boundaries ofthese sectors (the fences). As mentioned, such intercompart-mental hop diffusion may also occur spontaneously, albeitwith low frequency. Short-tailed cadherin proteins were notrestricted within corrals, indicating that the cytosolic domain,

presumably due to its interaction with the submembraneouscytoskeleton, is critical for confinement.

It is important to note that these measurements of cad-herin mobility were made on E-cadherin molecules that werenot engaged in cell-cell contacts; clearly cadherin-cadherininteractions represent yet another and physiologically cruciallevel of complexity in their regulation (81, 462). Fusion ofcadherin molecules with native or photoactivatable green flu-orescence proteins (GFPs) proved to be an efficient tool tocharacterize cadherin mobility both within and outside theAJs and in different stages of AJ maturation (4, 66, 420).FRAP experiments revealed that cadherins show distinct, andbiochemically regulated mobility in nascent and mature AJs(4, 117, 473). Besides the obvious role of actin skeleton, cad-herin clustering has been shown to be regulated by MTs as




well (448). A pool of radially organized MTs extend to thecell-cell contacts, and disruption of these prevents the accu-mulation and clustering of E-cadherin in the junctions. ThusMTs also play an important role in the focal distribution ofmembrane proteins.

Membrane—cytoskeleton interactions via lipidsAs already mentioned, many of the functionally most impor-tant interactions between the membrane and the cytoskele-ton are mediated or regulated via the association of proteinswith membrane lipids. In many cases, these interactions occurthrough well-defined lipid binding domains in a dynamic andtightly regulated manner (Table 1). These interactions con-tribute both to the cytoskeleton-affecting signaling events andthe ensuing structural remodeling. Cognizant of the perplex-ing multitude of lipid-protein interactions, we will focus onthose examples, which are known to have functional signifi-cance in terms of actin polymerization and membrane dynam-ics, and therefore are known to regulate the shape of themembranes and lead to the formation of specialized protru-sions such as lamellipodial and filopodial extensions. Themechanism of these dynamic events during specialized func-tions will be discussed in Section “Membrane-CytoskeletonInteractions: Functional Aspects.” Further, we will concen-trate on PIP2, as a lipid central in membrane-cytoskeletoninteractions.

PIP2 and the membrane-cytoskeleton interface

PIP2 constitutes only on the order of 1% of the total lipidin the plasma membrane, but it is an important regulator ofcytoskeletal and membrane dynamics (139, 237, 272, 481,516). In part, the effect of PIP2 on the membrane-cytoskeletoninterface is mediated directly by its interaction with multipleactin binding proteins, but also indirectly by its effects ontransmembrane proteins, such as ion channels, transporters,and signaling receptors (220, 292, 329, 508) as well as someRho family GTPases (181) and their effectors (453).

PIP2 and actin

PIP2 regulates the organization and dynamics of the cytoskele-ton at multiple levels. There is no compelling evidence thatPIP2 directly affects any of the cytoskeletal polymers them-selves with sufficient specificity to control their assemblyin vivo, and the large anionic charge density of PIP2 wouldtend to repel the cytoskeletal polymers, all of which arestrong anionic polyelectrolytes, unless their interaction ismediated by a polyvalent cation. However, many actin- andMT-binding proteins have polycationic domains that interactstrongly with PIP2. Such interactions lead variably to localiza-tion of specific proteins to the membrane-cytoskeletal inter-face, or either positive or negative regulation of the protein’sfunction.

PIP2 binds and regulates multiple actin binding proteinsthat are involved in actin nucleation, filament severing andend capping, and reinforcement of the membrane cytoskele-tal linkages (204, 272, 392, 516). Cytoskeletal proteins wereamong the first reported physiological ligands for PIP2, start-ing with reports that profilin (270), alpha-actinin (58), vin-culin (199, 319), and components of the erythrocyte mem-brane cytoskeleton (11) bound acidic phospholipids includ-ing PIP2, and that PIP2 dissociated complexes of profilin andactin (241). Since these first reports, many more cytoskeletalproteins have been found to bind PIP2. Some are activated byPIP2, some are inhibited, and others bind PIP2 without appar-ent change in actin binding function. Studies in which PIP2

levels are altered in cells suggest that increases in cellularPIP2 promotes polymerization of cytoskeletal actin and sta-bilizes its interaction with the plasma membrane (204, 272).Decreasing cellular PIP2 levels can correspondingly decreasecellular actin assembly.

In some cases, cytoskeletal proteins bind PIP2 by domainswith homology to canonical phosphoinositide (PPI)-bindingmotifs such as PH, FYVE, PX, FERM, or ENTH/ANTHdomains. Among these well-defined PPI binding modules isthe FERM domain (for 4.1, ezrin, radixin, and moesin) (119)found in ezrin/radixin/moesin (ERM proteins) (126, 335) andtalin (119). Other actin regulatory proteins bind PIP2 usingless obviously structured motifs that contain clusters of basicand hydrophobic amino acids (204). Such proteins include theWiskott Aldrich Syndrome protein (WASP) superfamily thatpromotes actin assembly by activating the nucleating Arp2/3complex (see Section “Actin polymerization drives membraneprotrusions and intercellular movements of pathogens”); cap-ping protein/CapZ and gCap39, which cap the filament (+)ends; cofilin, which severs actin filaments and acceleratesactin treadmilling; gelsolin family proteins, which sever andcap actin filaments to promote dynamic actin reorganization;and vinculin, which regulates FA turnover.

Proteins such as gelsolin and profilin that lack traditionalPIP2-binding motifs bind both the charged inositol head groupand the hydrophobic acyl chains of PIP2 (155, 205, 241). N-WASP simultaneously binds several PIP2 via its polybasicdomain and responds to small changes in PIP2 surface density(330). Cofilin also appears to bind both the acyl chains andhead group of PIP2 in a binding pocket (155) but other studiessuggest that this binding is mainly electrostatic and involvesmultiple PIP2 head groups (364). Thus, cofilin, N-WASP, andvery likely other proteins whose activities are altered, are PIP2

sensors, which respond both to changes in PIP2 density at theplasma membrane and to other signals such as Rho familyGTPases and phosphorylation (139, 204, 237, 272, 481, 516)and so integrate multiple signals to control the link betweenthe membrane and the actin cytoskeleton.

ERM proteins link the membrane to the cell cortex atmultiple sites of membrane protrusion such as uropods andfilopodia. Their actin binding potential is activated by bothphosphorylation and PIP2. For example, binding of PIP2 toezrin at its FERM domain relieves the autoinhibited state and




Assembly initiationWASP family proteins

Membrane linkageTalinVinculinERM proteinsSpectrinBand 4.1Myosin 1Kinesin (Unc104)Monomer sequestering

Profilin

Filament cappingCapping proteinCapGGelsolin

Filament severingGelsolin family proteinsCofilin/ADF

Effects of PIP2 or PIP3 oncytoskeletal protein functionand localization

Figure 5 Summary of cytoskeletal proteins that are regulated orlocalized by polyphosphoinositides. Proteins marked in blue are acti-vated or localized at the plasma membrane by PIP2 or PIP3; thosedenoted in red are inhibited by PIP2.

allows actin binding (126, 335). Both membrane and actinbinding are required for ERM protein function. Mutation ofbasic residues in ezrin’s PIP2-binding domain prevents ezrinlocalization to actin-rich membrane structures (26). The PIP2-dependent linkage of ezrin to the transmembrane adhesionprotein ICAM 1 (178, 419) also involves the FERM domain(168).

The number of proteins affected by PIP2 and the differentways in which protein function can be affected, had made itchallenging to decide which results observed in vitro translateto analogous functions in vivo. However, there is a strikingpattern to the actin- and membrane-binding functions that areactivated or inhibited by PIP2 as summarized in Figure 5.All PIP2-sensitive actin monomer binding proteins (e.g., pro-filin), and proteins that sever actin filaments (e.g., gelsolinand cofilin) are inactivated by PIP2. In contrast, proteins thatpromote actin assembly (e.g., WASP family proteins) or thatlink F-actin to the membrane (e.g., ERM, talin, and vinculin)are activated by PIP2. In addition, some motor proteins, suchas myosin I (226) are targeted to PIP2-enriched membranes.Filament bundling proteins (e.g., alpha-actinin) are generallyinactivated by PIP2. The overall effect of this constellation ofbiochemical events is consistent with the finding that whencellular PIP2 levels decrease, actin assembly and cytoskeletallinkage are inhibited. However, there are few if any stud-ies to determine which of the many possible PIP2-mediatedreactions are preferentially stimulated under different cellularconditions.

PIP2 and microtubules

Microtubule attachment to the membrane and vesicle move-ment along MTs is also regulated by PIP2. For example, localdomains enriched in PIP2 not only stimulate N-WASP depen-dent actin assembly, but they also localize IQGAP1, which inturn stabilizes MTs to these PIP2-patches at the leading edgeof a moving cell (153). In addition, some MT motors such asthe kinesin KIF1A contain a PH domain that binds to PIP2

in a vesicle membrane (221). Such lipid-dependent anchorsfor MT motors might play a role in both vesicle transportand the extension of endoplasmic reticulum (ER) tubules thatare pulled out along MTs within the cell (see also Section“Curvature formation and sensing and its relationship to thecytoskeleton”).

Structures and properties of lipid-binding sites incytoskeletal proteins

Among the hundreds of proteins that bind selectively topolyphosphoinositides or other membrane lipids (71), manyare characterized by well-defined PPI- or PS-interactingmodules, such as pleckstrin (PH), FYVE, PX, FERM, andENTH/ANTH. However, proteins that link the cytoskeletonto the membrane (or that are functionally regulated by PIP2 orother PIPs), often lack such canonical lipid-binding motifs.Some actin-binding proteins such as spectrin and N-WASPdo possess PH domains and others such as myosin 1c andERM proteins have cationic sequences homologous to PHdomains that are involved in lipid binding. However, evensuch proteins with PH domains do not appear to bind PIP2

with the same level of specificity characteristic of truly spe-cific PIP2 ligands such as the PH domain of PLC-δ, and alsoin contrast with such specific PH domains do not bind inositolphosphates with the same affinity as the intact PIPs. Even pro-teins such as myosin 1e that possess a PH domain retain PIPbinding when their PH domain is mutated (125). More com-monly, cytoskeletal proteins bind acidic lipids, usually PIP2,at specific sites characterized by a combination of cationic andhydrophobic residues that in some causes undergo conforma-tional changes after lipid binding, but do not have strictlyhomologous sequences. The structures of PIP2-binding siteswithin cytoskeletal proteins are discussed in previous reviews(318, 516).

In addition to highly specific lipid binding by traditionalligand binding pockets, cytoskeletal-membrane linkers oftenare also stabilized by electrostatic interactions that can havehigh affinity with less specificity, except for targeting of themost highly anionic membrane sites. Several actin-bindingproteins, and notably those such as profiin, cofilin, and gel-solin, that were among the first to be shown to be function-ally regulated by PIP2, do not have canonical PIP-bindingmotifs, and require both the anionic lipid head group and thehydrophobic acyl chain domain of the lipids for binding (76,127, 155, 240).

A primarily electrostatic mode of binding to PIP2 andother anionic inner leaflet lipids has been delineated for sev-eral cytoskeletal membrane linkers. Electrostatic interactionsbetween several neighboring lipids and an unstructured poly-basic protein domain such as that in the myristoylated alanine-rich protein kinase C substrate (MARCKS) protein (242, 285)can account for the affinity and selectivity of binding andhas been the dominant model for understanding how PPIscan be localized or restricted within membrane domains andsequestered from the many other PIP-binding proteins in the




cytoplasm. Increases in the local anionic charge density onthe plasma membrane inner leaflet occurring after receptoractivation have been shown to be sufficient to localize spe-cific proteins to the plasma membrane (513, 515). Cytoskele-tal linkers that bind primarily electrostatically rather thanthrough specific head group docking sites would be morestrongly sequestered by membrane domains locally enrichedin PIP2 or other poly-PIPs compared to domains that areequally enriched in PS because of the much greater chargeof PIPs that for PIP2 reach levels of −3 to −4 comparedto a charge of −1 for PS (252). Recent demonstrations thatPIP2 forms nanoscaled clusters with very large mole fractionwithin mixed lipid membranes both in vitro (499) and in livecells (1, 480), suggests that such areas of high charge den-sity might be hot spots for recruitment of membrane linkersthat are stabilized by electrostatic attractions. The cell bio-logical aspect of electrostatic interactions and the regulationof membrane surface charge are discussed in Section “Trans-membrane potential and surface charge.”

The 2D Membrane SkeletonMost cells have a 2D meshwork composed of spectrin, actin,and several other proteins, including ankyrin, band 4.1, andadducin that link spectrin and actin to each other and anchorthem at specific sites to the lipid bilayer through transmem-brane proteins. The spectrin/actin network is most thoroughlycharacterized for red blood cells where it is the primary proteindeterminant of membrane mechanics since these cells lack a3D cytoskeleton, but most cell types have similar spectrin-based networks that line at least parts of their plasma mem-brane and the cytoplasmic face of some internal organelles,such as the Golgi (34, 446).

The 2D cytoskeleton is distinct both biochemically andphysically from the 3D cytoskeleton. The clearest physicaldifference is the great flexibility of this network. The majorelastic element in the 2D network is spectrin, rather than actin,which instead acts as a linker that bridges the large flexiblespectrin molecules. Spectrin has a persistence length (a mea-sure of the length scale over which a filament is approximatelystraight) less than 20 nm (294), whereas the persistence lengthof the filaments in the 3D cytoskeleton range from near 1 μmfor IFs to 10 μm for F-actin and mm for MTs (503).

The organization of the contacts holding spectrin to actinand the membrane is depicted in Figure 1 (90). A tetramericcomplex composed of two extended chains of alpha spectrinand two chains of beta spectrin is the fundamental networkstrand. The spectrin tetramer links to short actin polymersthrough actin-binding sites at the N-termini of the beta spec-trin chain. The short actin polymer itself is not a filamentthat links nodes within the spectrin/actin network but ratherhelps constitute the node itself and anchors the 2D spectrinnetwork to the cell membrane. The flexible structures of thespectrin tetramers and the spatial distribution of their bind-ing sites for each other and for actin filaments allow them

to assemble into a flexible 2D sheet with roughly hexagonalsymmetry. The barbed end of the actin polymer at networknodes binds adducin, which together with band 4.1 connectsthe network to the transmembrane proteins band 3 (anionexchanger 1 or AE1) and glycophorin (22). The actin polymeris kept short and stable because it is bound at the pointed endby tropomodulin and laminated on its side by tropomyosin.Spectrin forms additional bonds to transmembrane proteinsby its affinity for ankyrin which complexes with AE1 andother proteins at transmembrane sites different from thosecontacted by adducin. The complex of adducin and dematinis also reported to link the network to glucose transporter 1(216).

Spectrin-based networks are important not only for stabi-lizing the red cell membrane but also are essential at specificsites and stages of development of neurons and most otheranimal cells. Spectrin binds not only to its unique set of trans-membrane receptors but also to adhesion receptors such ascadherins at cell-cell junctions, and might contribute to bothmechanical stabilization and signaling at these sites. (39). Inaddition to its attachments to transmembrane protein com-plexes spectrin also contacts the membrane by direct contactswith phospholipids. Like many cytoskeletal proteins, spectrinand some spectrin-binding proteins bind to PIPs, and discov-ery of the complex of glycophorin, band 4.1 and PIP2 providedsome of the first evidence for a link between the cytoskeletonand poly-PIPs (11). Spectrin also binds strongly and at multi-ple sites to the much more abundant anionic phospholipid PS.Beta spectrin contains a PH domain that is sufficient to bindPIPs in lipid membranes (268, 497) but there are multiplesites in spectrin with affinity for PS, and the greater numberof these sites together with the much greater abundance of PSmakes both interactions likely important for spectrin function.Under normal conditions, most animal cells tightly restrict PSdistribution to the inner leaflet of the plasma membrane andto the cytoplasm-facing leaflet of internal organelles, and thenumerous contacts with spectrin might contribute to stabiliz-ing PS in this configuration.

The physical effect of lining the flexible spectrin-actinnetwork at multiple sites to the cell’s lipid bilayer is to pro-vide the plasma membrane with both flexibility and tensilestrength that allows large shape transformations without dam-age to the cell caused by rupture of the lipid bilayer. Sincelipid bilayers withstand only a few percent increase in areaper molecule before bursting, the cell can increase its surfacearea to volume ratio, as it does whenever it changes shapeaway from a spherical geometry, only if its plasma membranecontains excess surface area or can be rapidly increased byexocytosis. In red cells and presumably most other cell types,the excess lipid bilayer is stabilized by bonds to the spectrin-actin network, which can withstand much larger tensionalstresses and deformations without breaking. The laminationbetween lipid bilayers and 2D protein networks of differentresting surface areas also introduces interesting physical prop-erties that alter thermal fluctuations and responses to exter-nal stress (452). In red cells subjected to shear stress, the




distension of the spectrin/actin network appears to involvenot only long range changes in the end-to-end distance of thenetwork filaments but also limited unfolding of the secondarystructures within the many folded domains within the spectrinchains (210). A recent study of red cell skeletons in which thelipid bilayer was removed showed that deformations on theorder of those that occur as cells squeeze though small bloodvessels could be accomplished with minimal protein unfold-ing (5). Both network-level and protein-level changes in thestructure of spectrin maintain plasma membrane integrity ascells undergo the large strains caused by the forces to whichthey are repeatedly subjected in vivo. Finally, further studiesare warranted to explore the role of the spectrin skeleton inmembrane partitioning and the control of membrane proteinmobility (see fence-and-picket model). The spectrin mesh-work was shown to control lateral diffusion in the membraneof erythrocytes (402), and recently the mobility of CD45, atransmembrane tyrosine phosphatase with a key role in lym-phocyte activation, was found to be controlled by spectrin(61). These observations suggest that the spectrin network,in collaboration with the actin skeleton, may be an importantcontributor to membrane compartmentalization and a regula-tor of signal transduction.

Membrane-CytoskeletonInteractions: Functional AspectsThe tight functional relationship between the membrane andthe cytoskeleton roots in the facts that (i) the membraneis the site from where most of the regulatory events thatalter the cytoskeleton emanate and (ii) the ensuing cytoskele-tal changes alter the shape and other physical propertiesof the membrane, which in turn are critical for the result-ing functions. The following sections deal with some keyfunctions and the underlying biophysical and cell biologicalprocesses.

Force generation at the membrane: pushingthe envelopeActin polymerization drives membrane protrusionsand intercellular movements of pathogens

In a set of landmark papers published in the early 1980s(184–186), Hill and Kirschner considered the thermodynam-ics of polymerization reactions, and arrived at the conclusionsthat the free energy release during the polymerization processitself can be used to perform work, that is, the elongationof the polymer can move a load (for calculations in case ofactin polymerization see Section “The pushing mechanism,part A”). This idea implied that actin polymerization itselfwithout the involvement of ATP-driven motor proteins (sucha myosin) can exert sufficient force to drive membrane pro-trusions. Indeed, the estimated free energy of actin polymer-ization (�G ≈ −14 kBT/monomer, where kB is Boltzmann’s

Figure 6 Deformation of phospholipid vesicles by polymerizationof encapsulated actin or tubulin. (A) Shape change of giant unilamel-lar vesicle formed by 50:50 wt. ratio of dimyristoylphosphatidylcholinecardiolipin after polymerization of 200 μmol/L actin within the vesicleinterior. Adapted, with permission, from (298). Scale bar: 100 μm.(B) Various morphologies of vesicles composed of 40% dioleoylphos-phatidylserine and 60% dioleoylphosphatidylcholine (DOPC) afterpolymerization of 2.5 mg/mL encapsulated tubulin. Scale bar: 10 μm.(C) Protrusion of initially spherical vesicle by single microtubule poly-merization. Reproduced, with permission, from (141). Long axis of thevesicle is 15 microns.

constants and T is the absolute temperature (341) was pro-posed to be sufficient to overcome the bending energy of thelipid bilayer, thereby causing membrane deformation. Studiesin which actin was polymerized within a lipid vesicle (86, 298)have provided experimental evidence for this concept, show-ing that actin polymerization induced within a liposome dis-torted the initially spherical shape of the vesicle and in somecases resulted in the formation of long, pseudopod-like mem-brane protrusions (298) as shown in Figure 6. In other cases,the polymerized actin associated laterally along the wall ofthe liposome resembling to the cortical actin network under-lying many cell types (353). Most giant unilamellar vesiclepreparations used to encapsulate polymerizing actin are com-posed largely of phosphatidyl serine which in cells is normallyrestricted to the inner leaflet of lipid bilayers, and there is asignificant absorption to this lipid, which can be reversed byincorporation of cholesterol or lipopolymers within the lipo-some bilayers and which strongly perturbs the morphology ofactin within the liposome (256). Even more striking protru-sions of the lipid bilayer are driven by microtubule assembly,as GTP-bound tubulin polymerizes with resulting hydrolysis




of GTP to GDP (191, 213). The polymerization of tubulininitially produces long tubular extension from both sides ofan initially spherical liposome, which in the absence of MT-stabilizing MAPS can gradually transform to a liposome withone tubular extension. Tubulin assembly is able to extendthe membrane farther and straighter, because microtubulesare much stiffer than actin filaments and are, therefore, moreresistant to bucking under a mechanical load. (118, 120, 141,191). Nonetheless, in most cells actin polymerization provedto be the key mechanism underlying motility-associated mem-brane distortion, so this will be the focus of the subsequentdiscussion.

The critical role of actin polymerization in membraneprotrusion of live cells has been indicated by observationsthat treatment of cells with actin polymerization inhibitors(e.g., cytochalasin B) impairs edge protrusion and inhibitsmigration (84, 417, 475). Finally, that actin polymerizationalone, that is, without the involvement of classical ATP-drivenmolecular motors, can generate movement of physiologicallyrelevant velocities has been proven by elegant in vitro recon-stitution experiments (40, 70, 263). In these studies, actinpolymerization-driven motility was first shown in the con-text of the movement of bacteria (Shigella and Listeria) andthen of glass beads. Exploration of actin-driven movementof pathogens (both within cells and in vitro) proved to beinstrumental for our understanding of actin-driven membranedynamics, because these bacteria harness the same actin poly-merization machinery, which is normally utilized for gen-erating various membrane protrusions and processes (e.g.,lamellipodia, filopodia, microspikes, phagocytic cups, etc.).Five protein components were found to be essential for actin-propelled movement in vitro: (i) actin itself; (ii) an actinnucleation-promoting factor, which in the bacterial studieswas the cellular protein N-WASP (activated by the Shigellaprotein IscA) (465) or the Listeria protein ActA (439), whichmimics active N-WASP; (iii) the Arp2/3 complex, an actinnucleating factor composed of seven subunits, which stimu-lates filament growth by initiating a new branch at the side ofan existing filament (183, 328); (iv) a severing protein such asactin depolymerizing factor or cofilin (41); and (v) a cappingprotein, which terminates filament elongation (85). While fac-tors 1-3 are necessary for the incorporation of new monomersinto a growing filament, factors 4 and 5 are critical for main-taining a high steady-state G-actin level. ATP is also requiredfor the process. Actin binds adenine nucleotides (ATP andADP). In cells G-actin is saturated by ATP, and ATP-actinis preferentially incorporated at the barbed end. Interestingly,once captured in a filament, actin hydrolyses ATP 40,000-foldfaster than in its monomeric form (349). The phosphate thenslowly dissociates. These kinetic properties establish a gra-dient between the ATP-rich, (young, barbed) and ADP-rich(old, pointed) end of the filament, from where ADP-actin pref-erentially dissociates. When the concentration of ATP-actinfalls between the critical concentration (the ratio of the on anoff rates) of the barbed end (≈0.1 μmol/L) and that of thepointed end (≈0.6 μmol/L), then barbed end polymerization

occurs simultaneously with pointed end depolymerization,a process termed as actin treadmilling. Experimental obser-vations and theoretical considerations agree that actin-basedmotility is realized through asymmetrical addition of sub-units, which leads to unidirectional growth of actin filamentsat their barbed end that is always located adjacent to the objectto be pushed (bacterium or cell membrane), concomitant withthe continuous depolymerization at their pointed end, in theprocess of actin treadmilling.

The role of the membrane per se in the actinpolymerization-driven protrusion is at least threefold. First,the membrane is critical for generating the initial signals thatlocally turn on the actin polymerization machinery. The mem-brane contains receptors, cell/cell, and ECM/cytoskeletonconnecting elements (cadherins and integrins), the activationof which induces the generation of intracellular messengers,which topically activate nucleation promoting factors such asN-WASP and formins. In addition, a major source of suchmessengers are the lipid components of the membrane itself[e.g., PIP2 and PI (3,4,5)-trisphosphate (PIP3)]. Indeed, lipo-somes containing PIP2 are sufficient to trigger actin assemblyin a cell extract and produce an actin comet tail resemblingthat generated by Listeria (16). Second, the membrane rep-resents the load against which the polymerization machinerypushes, which is necessary for performing mechanical work.Accordingly, the biophysical properties of the membrane arekey determinants of the ensuing membrane protrusions. Forexample, thermal fluctuations of the membrane have beenproposed to be essential for submembraneous actin polymer-ization (Brownian ratchet model) [(299, 341) and see Section“The pushing mechanism, part B”), while changes in mem-brane curvature and tension may play an important role in theregulation of the polymerization process (19, 27, 411). Third,the membrane likely contains “fixed” actin skeleton-bindingsites, which may play an important role in force transductionduring protrusion (actoclampin model) (108) and see Section“The biochemical nature of ‘actoclampin”’)].

Although there is general agreement that regulated actinpolymerization provides the major driving force for themovement of the membrane, there is ongoing debate aboutthe actual mechanisms whereby this happens (510). Thereare three fundamental questions that the various modelsstrive to describe and explain: (i) What are the structuralfeatures of the actin meshwork under the membrane? (ii)How is actin polymerization coupled to membrane pro-trusion? In other words, what is the mechanism throughwhich the new monomers “squeeze” in under the membraneand get incorporated into the existing filaments? (iii) Howand where is the “pointed end” (or the middle of the fila-ment) tethered in the meshwork? The latter is a structuralrequirement necessary for net membrane expansion, sinceunless the pointed end (or the mid-region) of the filament isanchored into a mechanically rigid structure, the addition ofmonomers to the barbed end adjacent to the membrane is aslikely to push the filament back as to extend the membraneforward (246).




1. Extracellular stimuli

2. Produceactive GTPases& PIP2 3. Activate

WASp/Scar4. Activate Arp2/3complex to initiatenew filaments

ActivatedGTPases& PIP2

Arp2/3 complex

WASp/Scar

11. Pool of ATP-actin bound to profilin

8. Aging 9. ADF/cofilin severs& depolymerizesADP-filaments

5. Barbed endselongate

7. Capping proteinterminates elongation

PAK

O70o

ADP-actin

10. Profilin catalyzes exchange of ADP for ATP

6. Growing filaments push membrane forward

12. LIM-kinaseinhibits ADF/cofilin

Figure 7 The dendritic nucleation/actin treadmilling (DNAT) model for membrane protrusion. Thekey feature of the model is the generation of daughter filaments sprouting out in a 70o angle at the sideof preexistent mother filaments, as induced by the activated Arp2/3 complex. In addition to the majorstructural characteristics, the model also shows the various signaling steps leading to the activation of theArp2/3 complex (1-4), the mechanism of filament elongation, and capping associated with membraneexpansion (5-8), and the various processes that ensure and regulate the continuous recycling of actinmonomers (9-12). Several important aspects of the model are discussed in the text. Reproduced, withpermission, from (350).

The pushing structure

There are two main structural models termed as the DendriticNucleation/Array Treadmilling Model (DNAT) (49, 63, 306,351, 460) (Fig. 7) and the Linear Treadmilling Model (LIT)(479) (Fig. 8), which address F-actin organization within thelamellipodium, that is, in the flat (< 200 nm high, 5-30 μmwide) protrusion formed at the leading edge of motile fibrob-lasts and epithelial cells. The DNAT is the prevailing paradigmand it states that lammellipodial actin is primarily composedof Y-shaped branches, which are formed when a daughter fila-ment is nucleated at the side of a preexisting mother filamentby the connector and pointed end-capper Arp2/3 complex,which initiates the polymerization of the daughter filament ina 70◦ angle. In vitro and in vivo observations lend strong sup-port to this arrangement (459) (Fig. 9). Both electron micro-scopic (306) and fluorescence (10, 47) studies performed bythe Pollard lab revealed that the purified Arp2/3 complexinduces branching at the side of filaments in a 70◦ angle.Moreover, using platinum electron replica microscopy, Svitk-ina, Verkhovsky, Borisy, and their colleagues (459,460) haveshown that actin filaments formed Y-branches (with an aver-age angle of 75 ± 10◦ (306) within the densely knit meshworkat the edge of lamellipodia in fish and Xenopus keratocytesand fibroblasts, and that the Arp2/3 complex (visualized byimmunogold labeling) was located at the branch points. The

angular distribution of the short peripheral filaments exhibitedtwo peaks at ± 35◦ with respect to the normal to the mem-brane, which corresponds to the “evolutionarily optimized”angle (70◦/2) for maximal pushing by a dendritic meshworkwith 70◦ branches (271, 405).

Despite these congruent biochemical and morphologicaldata, Small and colleagues challenged the DNAT paradigm.Small, who was the first to show that the barbed end ori-ents toward the membrane (436) and that lamellipodia con-tain a dense criss-cross (diagonal) actin meshwork (434,435)recently reported that in their keratocyte preparations the fil-aments subtend angles between 15◦-90◦ to the front, whichis inconsistent with the proposed dendritic branching (224).Moreover, based on their new analysis of 3D images obtainedby using vitreously frozen cells and electron tomography, theyclaim that branching does not occur at all (479). They pro-pose that the diagonal appearance of the meshwork is due tothe crossing over of unbranched filaments, while the forma-tion of the previously reported dendritic arrays is attributableto artifacts originating from the detergent extraction, criticalpoint drying, and platinum coating of the samples, which maylead to filament breakage and rearrangement (433, 479). Theauthors argue that this theory is not only more consistent withthe wider angular distribution of the filaments with respect tothe membrane (at least behind the tip of the leading edge),but handles the lamellipodium as a 3D (and not as a 2D)



(a)(B)

(A)

(b) (c)

Key:

ActinWave

complexArp2/3

complexVASP-

profilin/actin Formin Cross-linkers

TRENDS in Cell Biology

Figure 8 The linear treadmilling (LIT) model. (A) Electron tomography of the lamel-lipodium. (a) Tomogram section of the cytoskeleton within the keratocyte lamel-lipodium. The blue rings indicate intersections of overlapping filaments, whereas thered rings mark putative filament branch points. (The arrows indicate typical examples).(b) Model constructed from the tomogram. The green lines correspond to actin fila-ments, the blue spheres label filament intersections at overlaps, while the red onesindicate putative branch points. In this particular image 320 overlaps and 4 branchpoints were identified. Based on such findings the authors suggest that the predominantarrangement of filaments in the lamellipodium is linear (nonbranching), and the previ-ously reported excessive branching might have been due to the misinterpretation of thefrequent crossovers of linear filaments. Reproduced, with permission, from (479). (B)The putative arrangement and role of the actin nucleation machinery in the context ofthe LIT model. (a) The tethered nucleation/elongation model. Actin is nucleated underthe membrane by WAVE-mediated activation of the Arp2/3 complex in a nonbranch-ing manner. The Arp2/3 complex remains at the pointed end and treadmills with thefilament. The plus end grows under the membrane while it remains tethered to WAVEand ENA-VASP (which may replace WAVE). The structure can be stabilized by shortactin cross-linkers at the membrane and by longer ones (such as filamin) deeper inthe lamellipodium. The latter cross-links filaments at the intersecting regions. At theforming filopodia filaments are extensively bundled, while actin nucleation at the tip ismediated by the collaborative action of ENA-VASP and formins. (b) and (c) show theboxes labeled in (a) with higher resolution. Reproduced, with permission, from (433).



a

(A) (C) (D)

(B)

b c

I

II

III

40 nm

IV

70°

Figure 9 Cytoskeletal structure in the lamellipodium and Arp2/3-mediated actin branch formation in vivo and in vitro. I: electron micro-scopic pictures of the actin network in the lamellipodium of Xenopuskeratocytes. In (a), the meshwork was stabilized with polyethylene glycoland phalloidin to prevent actin depolymerization from the pointed end.In (b), unprotected extraction was performed in the absence of theseF-actin-stabilizing drugs. Note the dense meshwork in the membrane-adjacent zone of the lamellipodium, which remains nearly intact evenwithout protection against pointed end depolymerization. In contrast,at the rear of the lamellipodium (>1 μm from the membrane), there issubstantial reduction in the network after unprotected extraction. Thissuggests that in the peripheral zone of the lamellipodium pointed fil-ament ends are capped (presumably by Arp2/3, see II). (c) shows theenlarged image of the boxed area in (b). Bar 1 μm. II: immunogoldlabeling of the Arp2/3 complex in the lamellipodium. Xenopus kera-tocytes were briefly treated with the barbed end capper cytochalasin D(to reduce the density of the meshwork), extracted in the presence ofphalloidin, fixed and immuno-labeled with a primary antibody againstthe p21 component of the Arp2/3 complex, and a gold-conjugatedsecondary antibody. The gold particles are highlighted with yellow.Actin filaments exhibited multiple branching, and the Arp2/3 complexlocalized at or near the branching points. Adapted, with permission,from (459). III: In vitro formation of actin branches in the presence ofthe Arp2/3 complex. (A) Electron micrograph of the purified Arp2/3complex shows globular particles of 10 × 14 nm. In the presence ofpurified Arp2/3, gelsolin-capped actin filaments form numerous end-to-side branches with a 70º angle (B and C). Linear filaments areperiodically decorated with the purified Arp2/3 complex (D). Repro-duced, with permission, from (306). IV: direct evidence of branch for-mation by the Arp2/3 complex in vitro. ADP-actin was polymerized andlabeled by Alexa-green phalloidin. Subsequently, ATP-actin monomers,Arp2/3 complex, and the Arp2/3-activating WA domain of WASP wereadded to prepolymerized green actin, and further polymerization wasmonitored with rhodamine (red)-phalloidin. As the “fresh” red labelingshows, monomers were added both at the barbed end and to the sitesof existing filaments. Reproduced, with permission, from (47).

structure and—importantly—it obviates the need for a preex-istent “mother” filament near the edge, which they regard asan ill-defined and enigmatic concept. They also point out thataccording to DNAT, as the filaments pushing the membraneelongate, they should be capped (since long filaments buckleand cannot exert sufficient force) but this means that the fila-ments oriented in the right direction are terminated and newfilaments are initiated at the sides in suboptimal directions. Itis also relevant to note that cells in which the Arp2/3 com-plex has been disabled by silencing RNA coding for essentialcomponents of the complex are viable and remain able toassemble cortical actin meshworks, albeit with somewhat dif-ferent characteristics (105, 316).

However, as Higgs mentions in his recent commentary(182), there are undisputed observations about lammellipo-dial actin polymerization, and every viable theory should takethese into account. Thus, (i) the Arp2/3 complex is enrichedat the leading edge where it acts as a major actin nucle-ation factor; (ii) Arp2/3 is at the pointed end of filamentsit nucleates; and (iii) WAVE, (a WASP-like nucleation pro-moting factor and critical Arp2/3 activator), localizes to theleading edge. In light of these facts, two major questionsarise: (i) How does the Arp2/3 complex nucleate actin at theleading edge without branching and (ii) if the Arp2/3 com-plex can induce branch formation in vitro, why does it notdo so in vivo? To address these questions, Small and col-leagues hypothesize that in vivo the Arp2/3 is associated withand is activated by WAVE, when the latter is bound to themembrane (244, 466) (as opposed to in solution in vitro,where WAVE can bind “promiscuously” and activate Arp2/3on an existing filament). WAVE is activated by phosphory-lation, prenylated Rac-GTP, and acidic phospholipids (pre-dominantly PIP3) (244), events and agents that are presentat the membrane. This way actin polymerization starts atthe membrane and the polymerizing actin remains physicallyassociated with the membrane (actoclampin model, see Sec-tions “The pushing mechanism, part C” and “The biochemi-cal nature of ‘actoclampin”’). After this initial stage, Arp2/3would treadmill with the pointed end, while WAVE wouldrecruit ENA/VASP proteins (enabled/vasodilator-stimulatedphosphoproteins), which could replace it and would takeover as a polymerization-inducing (maintaining) factor at thebarbed end (Fig. 8). ENA/VASP proteins, which emerge askey players (31, 416) in normal and pathological cell motil-ity, have a number of remarkable properties: they not onlypromote polymerization at the barbed end by recruiting pro-filin and actin, but also reduce branching, increase bundlingand behave as “leaky cappers.” This means that they inhibitfilament termination by efficient cappers such as gelsolin (ant-icapping effect), and while they remain bound to the barbedend (i.e., they cap it to certain extent), they still can promotethe incorporation of new monomers at this locus. These prop-erties are likely important for understanding the mechanism ofelongation as they allow net polymerization while the filamentremains membrane bound. The LIT model can also accountfor the properties of a protein initially called insertin (142,




390), which was later found to be a proteolytically derivedfragment of actin-binding protein tensin (471), a componentof the FAs that link the cytoskeleton to integrins at the cellmembrane (262). Kinetic analysis of insertin’s effect on actinpolymerization showed that it slowed both polymerizationand depolymerization from the actin filament barbed end, butallowed this end to grow by addition of monomers, similarto other weak cappers. Finally, the LIT model also postulatesthat stability of the filament necessary for efficient push isbrought about by cross-linking and bundling proteins (e.g.,filamin, fascin, α-actinin, etc.).

Clearly many functional assumptions of the LIT model(e.g., the mechanism of the dissociation of the Arp2/3 fromWAVE or the recruitment of VASP by WAVE and the sub-sequent replacement of WAVE) await rigorous experimentaltesting. While the debate remains on, a few aspects should beemphasized. First, albeit the models seem to be diametricallyopposite, in fact, they are not mutually exclusive. Branch-ing and nonbranching polymerization modes may coexist andtheir prevalence may be dictated by the particular stimula-tion that triggers membrane protrusions and the type of theprotrusion per se. Indeed, the DNMT also postulated thatfilopodium formation involves the merging and bundling oforiginally branched filaments into long, parallel fibers (49).Second, the very premise of the LIT model remains contro-versial: reanalyzing the published electron tomography pic-tures of the Small lab, Yang and Svitkina have concludedthat these figures do show evidence of extensive branching,which was, however, overlooked (510). If this finding provesto be unambiguous, it satisfactorily settles the debate. In fact,while this article was under revision, Vinzens et al. from theSmall group reexamined the structure of actin filaments in thelamellipodium by electron tomography, and acknowledgedthe presence of branches. However, they found that—in con-trast to previous suggestions—branch junctions are not con-centrated at the front, but are continuously distributed withinat least 1 μm from the filament plus end. This means that effi-cient membrane pushing may require actin cross-linking closeto the membrane (494). Finally, the existence of direct phys-ical linkage (proteinaceous bridge) between the membraneand the growing filament is a compelling concept that gainsincreasing support both from experimental observations andtheoretical considerations (see Section “The pushing mech-anism, part C”). This concept is compatible with dendriticand nonbranching models as well, as it concentrates on theevents that happen at the filament-membrane interface wherethe pushing takes place.

The pushing mechanism

The mechanism whereby the polymerization of actin drivesthe membrane forward has not been fully elucidated. Cur-rently two (not necessarily exclusive) paradigms exist: mod-els involving various and increasingly sophisticated versionsof the Brownian Ratchet (BR) hypothesis (299,300, 341) andthe Clamped Filament Elongation (CFE) model, also termed

as the End-Tracking Motor hypothesis (106–108). In the fol-lowing paragraphs, we will discuss the fundamental aspectsof these. However, first it is worthwhile to make some thermo-dynamic considerations, which show that not even the directsource of energy liberated during the polymerization reactionhas been fully clarified.

(A) How is the pushing force generated during poly-merization? Thermodynamic considerations In thissection, we will briefly consider some thermodynamic aspectsof actin polymerization. While these points are not specificfor actin-propelled membrane protrusion, they are importantfor the mathematical derivation of the various membrane pro-trusion models (part B). Nonetheless, the principle and quali-tative aspects of each model are summarized in the beginningof the corresponding paragraphs in part B, so those not inter-ested in the quantitative handling of the problem can skip partA and the derivations in B.

The propelling forces may originate either from the freeenergy of a monomer binding to the polymer or from ATPhydrolysis or both (106). In principle, ATP hydrolysis orsome other nonequilibrium process is needed for any use-ful mechanical work to be done, but the issue is whether theATP hydrolysis coupled to actin assembly is required only toput the system out of equilibrium and allow the preferentialassembly of monomers at the barbed end, or whether a changein actin structure directly resulting from ATP hydrolysis pro-duces the force as it does for conventional motor proteins suchas myosin or kinesin.

Considering only the binding step, the free energy change(�G) is

�G = −kBT ln[AT]/A+T,c (1)

where [AT] is the concentration of ATP-actin and A+T,c is the

critical concentration of ATP actin at the barbed end (i.e.,the equilibrium dissociation constant of the reaction, which is≈ 0. 1 μmol/L). From this, the force F at equilibrium (whichis also the stall force that can stop the filament from growing)can be calculated as

F = �G/δ = −kBT/δ ln [AT]/A+T,c (2)

where δ is the length of the filament extension by addingone monomer to the polymer. Since the actin filament is astaggered double helix (Fig. 10), the actual growth is justhalf of the diameter of the monomer, which is 5.4/2= 2.7 ×10−9 m.

How big is the force generated during elongation of afilament, and can this be achieved from monomer bindingalone without the energy of the subsequent ATP hydrolysisby actin? Various methods have been used to estimate thisparameter (both during in vitro actin polymerization and incells). In most cases, the total force generated by a popu-lation of filaments was determined and then divided by theoptically measured (2) or estimated number of the filaments.




0.4

Actin filament α β

2δv

[μm

/s]

f δω =

fD

x

0.3

0.2

0.1

00 1 2 3

KBT

4 5

Figure 10 The original Brownian Ratchet (BR) model and the cor-responding force-velocity (F-V) relationship for edge protrusion duringactin polymerization. The actin filament polymerizes against the mem-brane, which has a diffusion constant D, and which is acted upon bya load, f. The conditional probability of the association of a monomerwith the polymer (provided the gap between the membrane and thepolymer tip is sufficiently large) is α, whereas that of the dissociationis β. The gap size should be at least δ, which is half the length ofthe actin monomer, considering the “staggered” growth of the actinfilament. The y axis shows the speed of the polymerization ratchet (v),as a function of the dimensionless load force (ω). The solid line isbased on the assumption that β → 0, whereas the dashed line depictsthe F-V relationship as calculated according to equation (5), that is,when monomer association and dissociation are much slower than thediffusion. Reproduced, with permission, from (341).

The techniques used for force determination included thedeformation of lipid vesicles (478), the deflection of glassmicroneedles (273) or AFM (331, 355). The results placedthe polymerization-derived single filament force in the rangeof 4-10 pN. While highly informative, these approaches arehampered by the difficulty of accurately assessing the numberof filaments and by the finding that the ensuing network den-sity itself can be dependent on the load (331). To overcomethis problem, two groups developed methods to assess forcesgenerated by a single filament or small filament assemblies.Kovar and Pollard (228) determined the buckling forces ofsingle filaments attached to glass slides via myosin in a stablerigor bond on one end, and formin, a processive barbed endpolymerization-inducing protein on the other. Alternatively,Footer et al. (130) used optical traps to measure the forcegenerated by a few parallel actin filaments apposed against arigid barrier. Both studies proposed that the forces are in the1-2 pN range.

So can monomer binding alone account for the genera-tion of 1-10 pN forces? The answer is a probable yes at thelower, and a possible no at the higher limit of this inter-val. The force, as determined by Eq. (2) depends on theconcentration of ATP-actin ([AT]). Cytosolic actin concen-tration can be as high as 100 μmol/L (350), however the“free,” polymerization-competent concentration is certainlymuch less (albeit not exactly known). Nonetheless, if all this100 μmol/L were available and were bound to ATP, the poly-merization could provide ≈10 pN force. However, this upper

limit is very likely not achieved, for two reasons: (i) a large partof actin is bound to sequestering proteins (e.g., thymosin-β)and (ii) treadmilling, which does occur in cells, necessitatesthat the actin concentration (at least at the pointed end) beless than the A+

T,c, which (at least in pure actin solutions) is0.6 μmol/L. This puts the upper limit of [AT] at this level(although a gradient between the two ends is possible), whichcorresponds to ≈ 2.8 pN, if the efficiency of the reaction were100%. Thus, monomer binding may not be the only sourceof force generation under all conditions, and ATP hydrolysismay also contribute. While this point awaits verification, theinvolvement of a hydrolysis-dependent step would be highlycompatible with the end-tracking motor model of protrusion(see Section “The pushing mechanism, part C”). In any case,these calculations unambiguously show that even with themost conservative estimates, the ATP-G-actin/F-actin poly-mer system is thermodynamically adequate for generatingsufficient protrusive force to extend the cell membrane.

(B) The Brownian Ratchet models

The classic BR model The BR model, postulated in a pio-neering paper by Peskin, Odell, and Oster (341), provideda consistent and mathematically formulated theory aboutthe actual mechanism whereby polymerization is convertedto movement. Beside the fundamental qualitative insight,the great value of this approach was that it predicted aload force/velocity relationship in actin-propelled movement,which served as the basis to test this and other hypothesesagainst experimental data. The principle of the idea is that anactin monomer, which randomly diffuses to the tip of a poly-mer is captured by the barbed end, and since this event pre-vents the backdiffusion of the monomer, the process becomes“ratcheted.” However, polymerization will be able to performwork, that is, to move a load only if the monomer can be inter-calated between the load and the tip of the polymer and theload will be moved only if the polymer is fixed in space. Howis the gap generated into which the monomer can “squeeze”in? According to the original BR model, the gap formationis due to the thermal fluctuations (Brownian movement) ofthe load (e.g., the plasma membrane or bacterial surface).Thus, the random fluctuations of the load are rectified by thepolymerization process, which provides the free energy toimplement the ratchet. If the polymerizing filament is per-pendicular to the load, the minimal gap size needed to allowprotrusion is δ = 2.7 nm. Intuitively, the rate of the expan-sion will depend on the probability of the formation of a largeenough gap, where the gap width x must be greater than δ (x ≥δ). Ultimately, the rate of elongation will depend on the ratesof monomer addition and dissociation, but the former will beweighted by the probability of the formation of a sufficientgap at any given load, and this probability will depend on thegeometry and diffusion constant of the load particle. Thus,without load, the velocity of the elongation is

v = kon[AT] − β (3)




where β is the constant dissociation rate (a zero order process).However, if there is a load, then the rate will be:

v = kon(x)[AT] − β (4)

where kon(x) [AT] is the conditional probability of adding amonomer, when a gap with a width of x is formed. The authorsdenote kon(x) [AT] = α, when there is a sufficient gap to accepta monomer (x ≥ δ), and they set it at 0, when the gap is toosmall (x <δ).

From this, they derive the velocity of elongation as

v = δ(αe−ω − β), (5)

where ω is the dimensionless work performed against theload during the insertion of one monomer. [See the detailedderivation in the appendix of (341).] This is an importantresult, because it provides a relationship between the loadforce (f), and the velocity,

since ω = f δ/kBT (6)

and from (5) and (6) the stall force f0 (the load force at whichthe movement is stopped, so v = 0) can be calculated as:

f0 = −kBT/δ ln(β/α) (7)

It is worth noting that (5) can be expressed in a moregeneralized form (106):

V = kon [AT]∫ ∞

δ

p(x)dx − koff (8)

which shows even more plastically that the on-rate (kon [AT]) isweighted by a probability factor, p(x) denoting the probabilitydensity for a gap opening at a distance x between the filamenttip and the load. Here, p(x) is the steady-state solution of theFokker-Planck equation (which describes the time evolutionof the probability density function of particles, and can beseparately solved for each type of the various BR hypotheses.)

The BR model gives a convex force-velocity (F-V) rela-tionship, where the velocity sharply drops at small loads andmore slowly at the larger ones (Fig. 10). This correspondswell with the experimentally determined relationship for actinpolymerization on N-WASP-coated polystyrene beads (273)and for Listeria moving in tissue extracts of varying viscoelas-ticity (280). (Comparison of F-V relationships as predictedby the various models with the experimentally observed datain the lamellipodia of moving cells is addressed in Section“Force-velocity relationship in lamellipodia.”)

The elastic BR model (EBR) According to the original BRmodel, the gap is generated exclusively by the fluctuations ofthe load, while the filaments are treated as completely stiffrods that are perpendicular to the load. However, two obser-vations prompted Mogilner and Oster to refine the model,

thereby developing the EBR hypothesis. BR predicted thatthe velocity depends on the load through its diffusion coef-ficient. However, two actin-propelled bacteria, Listeria andShigella moved with the same speed despite their differentsize (458, 521), which should correspond to different loadfluctuations. Further, the lamellipodium contains a meshworkof actin filaments (not parallel arrays), and these are not abut-ting the membrane perpendicularly. Accordingly, the EBRcontains two major modifications: it predicts that the ther-mal fluctuation of the filaments and not that of the load (e.g.,membrane) represents the gap-generating mechanism; and ittakes into consideration that the pushing may not occur ata right angle. The model assumes that actin filaments at theedge behave as elastic rods, with a bending modulus that isproportional to their persistence length (λ, taken as 1 μm),that they are anchored to a cross-linked rigid actin gel, theyall have the same “free” length (l) and they reach the load atthe same angle (θ ) (Fig. 11 and Figure 12). A single filamentcan then be seen as a tiny spring (with an elastic constantthat depends on λ and l), which performs thermal motion.Since it is much harder to compress the filament than to bendit, the fluctuation will manifest as bending, and this bendingaway from the load will generate the gap necessary for actinintercalation. If the filament impinges on the load at an angleθ , the minimum required gap size is δcosθ (2.7 nm × cosθ ),that is, a smaller gap is sufficient than predicted by BR. Theanalysis then follows the same path as in BR, that is, the like-lihood of the formation of a gap equal or larger than δcosθis calculated, except this time the gap size will be a functionof the elastic properties of the “actin spring.” In analogy toEq. (4), the velocity of elongation is given by the followingformula:

v = δ cos θ (kon[AT]p − koff), (9)

where p is the probability of the gap of sufficient size, whichdepends on θ , the elastic constant of the spring and the equi-librium distance of the load from the tip [see (299) for thefull expression and the solution of the corresponding Fokker-Planck equations].

The model predicts the following relationship for the stallforce:

f0 ≈ kBT/δ√

l2 + 4λδ (10)

The authors also computed the force/velocity relation for amodel lamellipodium (Fig. 12) operating according to EBR,assuming that it is composed of 5000 actin filaments in amembrane area of 5× 0.2 μm at a free actin concentration of45 μmol/L. The resulting curve predicts a shallow dependenceat low loads followed by a steep part at bigger ones.

The existence of a gap between load and filament is a fun-damental feature of both BR and EBR. However, experimen-tal observations of actin-propelled pathogens, beads, and lipidvesicles cast doubt on the existence of such a gap. Specifically,(i) the actin tail was found to be tightly attached to Listeria




fθ

δ

Actinfilament tip

Loadforce

Load

Thermal fluctuations

V

x

k

l

y

Monomer concentration, M

Rigidactinnetwork

(A)

(B)

Koff

Kon

Figure 11 The Elastic Brownian Ratchet (EBR) model. (A) Accordingto the EBR model, the thermal fluctuations of the “free” filament end(rather than that of the membrane) generate the gap (δ) between thetip of the filaments and the membrane. The moving portion of thefilament (with a length of l) is anchored to a rigid base, that is, a cross-linked portion of the actin meshwork. In addition, the filament end isnot perpendicular to the membrane but impinges on it in an angleof θ . (B) The mechanical equivalent of the EBR model. The fluctuatingfilaments is modeled by a spring with a spring constant κ, which has anequilibrium position at a distance y from the membrane. The deviationfrom the equilibrium is denoted by x. Reproduced, with permission,from (299).

(148), or ActA-covered beads (62); (ii) the separation of theactin tail (by optical traps) required substantial force (>10 pN)(323); (iii) ActA- or WASP-coated vesicles are deformed anddevelop tension during actin polymerization (478); (iv) thethermal fluctuation of propelled bacteria are strongly sup-pressed (232); and (v) the movement of Listeria occurs as astepwise motion, with 5.4 nm “jumps.” These findings werenot readily compatible with the presence of a gap betweenthe load and the actin meshwork. But if there is no gap,how can the new monomer be incorporated under the load?To address these problems, Mogilner and Oster formulated

Figure 12 Lamellipodial protrusion according to the EBR model. (A)Lamellipodial actin is modeled as a biorthogonal array of filaments,oriented at angle θ to the membrane normal. The system pushes themembrane against a load force f. The force-velocity curve was calcu-lated for a hypothetical lamellipodium, containing 5000 filaments act-ing on a membrane area of 5 μm × 0.2 μm, and at a local monomerconcentration of 45 μmol/L. Reproduced, with permission, from (299).

the Elastic Ratchet and Tethered Filaments—or for short—Tethered Ratchet (TR) model (300). This posits that there aretwo populations of actin filaments: one that is attached to theload, and does not perform work, and one that is not attached,and pushes the load forward, performing work. The attachedfilaments are proposed to be tethered to the load (e.g., mem-brane) by nucleation promoting factors (e.g., WASP), whichbind to Arp2/3 at the side of the filament. In contrast, theworking filaments are not linked to the membrane, and theypush it forward according to an EBR mechanism. The work-ing filaments put the attached filaments into tension (the latterbehave like stiff Hookean springs), and eventually break theirattachment to the load. In the TR model, attached filamentsexert force against protrusion. Accordingly, the force balanceequation is

FL + faa = fww (11)

where FL is the total external load force (which may originate,e.g., from viscous drag), a and w are the number of attachedand working filaments, and fa, and fw is the force exerted by a




single attached or working filament, respectively. The criticalissue is the balance between the two populations. The changein the number of attached filaments depends on the rate ofnucleation (which couples the filament to the load), and therate of filament dissociation from these sites:

da/dt = n − kdissa (12)

where kdiss is the filament dissociation rate constant, a force-dependent parameter.

The change in the number of working filaments dependson rate of filament association (since these released filamentscan start working) and on the rate of capping, which terminatesmonomer incorporation and pushing:

dw/dt = kdissa − kcapw (13)

where, kcap is the rate constant for capping, which meansthat at steady state, the rate of nucleation is equal to the rateof dissociation, which is equal to the rate of capping (n =kdissa = kcapw).

Based on these considerations, a F-V relation can bederived (300). The TR model offers an improved hypothe-sis, which not only deals with the apparent contradiction ofconstant attachments versus monomer incorporation, but alsotakes capping into consideration, which is an experimentallyverified determinant of lamellipodium protrusion (33). It alsoprovides explanation for various phenomena observed dur-ing actin-propelled movement, including the findings that thevelocity of ActA-coated beads does not depend on the den-sity of coating and that smaller beads move slower (62, 63).However, the model cannot explain the stepwise motion ofthese particles.

(C) Clamped filament elongation/end-trackingmotor models The models aimed to describe the micro-scopic mechanisms of actin-driven movement span a wholespectrum, surmising that the load is untethered (BR and EBR),partially tethered (TR) or fully and permanently tethered(CFE) to the filament. Evidence supporting the strong cou-pling between the filament and the pushed surface promptedDickinson and Purich to formulate an alternative to the Brow-nian model (108) (Fig. 13). According to this, a tetheringapparatus, referred to as the clamp, links the load to the fil-ament tip. The clamp is locked to the terminal actin doubletand allows sufficient space for a new monomer to be loadedonto the tip. The addition of the new monomer triggers ATPhydrolysis on the clamped subunit, which in turn decreasesthe affinity of the clamp, allowing the translocation of theclamp to the newly incorporated, terminal, ATP-containingsubunit. The clamp is now locked there and the cycle repeats.This elegant hypothesis represents a new mechanochemicalmodel, which the authors term as the Lock, Load, & Firemechanism. The locking step is the tight binding of the clampto the last ATP-actin subunit; during the loading step a newATP-monomer(s) is added to the tip, and this initiates the

firing step, which involves the conformation change neces-sary for the drop in the affinity for the clamp and its subse-quent translocation. Thus, the load is driven by a three-strokeengine. Since various filaments attached to the load exist indifferent phases of this three-step process, some filaments arecompressed (favoring pushing), while others are stretched anddevelop tensile forces (pulling back). The surface advanceswhen a force imbalance occurs, which may be associated withthe “firing” of the most lagging filament, resulting in a discretestep forward.

The model treats the transition in the clamp position as if itwere the diffusion of the filament end along a 5.4 nm path (thediameter of the monomer), where it falls into a deep energywell. The rate of this “diffusion” depends on tensile andcompressive forces, the former facilitating whereas the latterhindering clamp translocation. Based on these approaches, astochastic simulation has been carried out, which predictedstepwise elongation and a F-V relationship wherein at smallopposing forces the velocity is force independent (i.e., lim-ited only by the time for monomer addition and ATP hydrol-ysis but not by the translocation step). In addition to theseaccurate predictions (see Section “Force-velocity relation-ship in lamellipodia”), this model offers several fundamentalinsights as well. For example, it explains why actin polymer-ization occurs (in a regulated manner) locally, at the surfaceto be pushed (e.g., the membrane); the clamp must interactnot only with actin (hence the name actoclampin) but alsowith the specific surface. Thus, the actoclampin should repre-sent a well-defined and regulatable link between the filamentand the membrane. The CFE also proposes a role for actin-mediated ATP hydrolysis. While treadmilling is known torequire ATP, it consumes only 10% of the available energy; therole (if any) of the other 90% remained an enigma. Accord-ing to the Stop Lock and Fire mechanism, ATP hydrolysismay fuel the clamp translocation step. Finally, the model isvery useful for the identification of the actual actocampin(s),because it can predict its biochemical and biophysicalproperties.

Since the clamp binds specifically to the end of the fila-ment, it must be a barbed end-tracking protein, and accord-ingly the mechanism whereby it translocates from the penul-timate to the terminal actin subunit is referred to as anend-tracking stepping motor. How can such a molecularmachine work? In the simplest case, the monomer directlybinds to the polymer tip, while the tracking protein (clampor part of the clamp) binds to the filament through an F-actin-binding domain (FAB). Alternatively, the tracking pro-tein may be involved in monomer binding as well, whichoccurs through a G-actin-binding domain (GAB), from wherethe monomer is transferred to the filament (referred to asthe “direct-transfer motor” (106) (Fig. 14). Thus, the puta-tive actoclampin should be a membrane-associated, presum-ably G- and F-actin binding, barbed end-tracking, processiveactin polymerase. Intriguingly, proteins from the ENA/VASPfamily meet all these criteria. ENA/VASP family members(Mena, VASP, EVL, and Drosophila ENA) (149) are modular




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Figure 13 Clamped filament elongation (CFE)/end-tracking motor (ETM) Model. (A) According tothe actoclampin hypothesis, the membrane is directly linked to the polymerization machinery through a“clamp,” whose affinity for actin is dynamically modulated, and which contains or is associated with amembrane-tethering domain. (B) In the CFT/EMT model, the actoclampin system pushes the membraneas a “three-stroke” engine through a “Load-Lock-Fire” mechanism. The central features of the model arethat (i) there is enough space between the membrane and the actin-associated region of the clamp formonomers to be added to the clamped filament; (ii) the affinity of the clamp changes according to thehydrolysis of the actin-associated adenine nucleotide (high affinity for actin-ATP, low affinity for actin-ADP+Pi or actin-ADP); and (iii) the clamp translocates from the low-affinity penultimate actin subunits tothe high affinity, ultimate actin subunits. For further details, see the figure and the text. Reproduced, withpermission, from (108).

proteins consisting of an N-terminal ENA/VASP Homology1 (EVH1) domain, which is involved in subcellular targetingof the molecule, a C-terminal EVH2 domain, which medi-ates tetramerization and also contains G- and F-actin bind-ing sites (GAB and FAB), and a central proline-rich region,which interacts with the monomer-binding protein, profilin(31, 113). ENA/VASP is localized to the lamellipodia andthe tip of filopodia, promoting actin polymerization at theseloci, and it is thought to be a central regulator of filopodialprotrusions, which contain parallel, nonbranching actin fibers(31, 385, 406). VASP has been shown to work as a barbed endpolymerase, which—in a cytosol-like environment—operatesin a profilin-actin-dependent manner (172). Moreover, in thepresence of VASP, monomer addition to the filament occursin a VASP-controlled manner (and not spontaneously), whichis consistent with a direct-transfer motor mode of action.

The biochemical nature of “actoclampin”

While these observations suggest that VASP is anactoclampin-type molecule (or part of such a molecularassembly), the identity of the actoclampin(s) is far from beingresolved. For example, although VASP increases the velocity

of WASP- or ActA-covered beads several-fold (70, 399), it isnot needed for the in vitro reconstruction of movement. Thisimplies that other actoclampins are in effect (e.g., WASP itselfmay fulfill such a function) and/or BR-type mechanisms mayalso contribute. Further support for the existence of alternativemechanism comes from observations that downregulation ofVASP does not stop lamellipodial expansion (in fact, it favorslamellipodium formation over filopodia), albeit it interfereswith normal lamellipodial dynamics (32, 33).

Another key and outstanding issue is the nature of thelink between the membrane and the actin clamp. The EVH1domain of VASP binds directly to a proline-rich region (the“FPPPP motif”) in ActA. Similar proline-rich motifs arefound in eukaryotic binding partners of VASP as well, includ-ing zyxin, vinculin, and palladin (53, 372, 373). These pro-teins are localized to FAs and cell-cell contacts sites (36,314), where they can interact (e.g., through α-actinin) withtransmembrane proteins, such as integrins (35). However,although overexpressed zyxin may localize to the lamellipo-dia (320), the endogenous protein does not seem to do so(386). A more likely candidate is lamellipodin (Lpd) whichalso binds to VASP through a polyproline region, but itcontains a PH domain as well, which specifically associates




Figure 14 Hypothetical actin polymerization motors involved inclamped filament elongation. (A) According to the end-tracking step-ping motor model, the clamp has a regulated affinity for F-actinthrough an F-actin-binding region (FAB). This domain translocates tothe ultimate actin subunits as they get incorporated in the filament. (B)According to the direct transfer end-trapping motor model, the trackingprotein has a G-actin binding region (GAB) as well, which captures themonomer (or the G-actin-profilin complex, as shown here). This stepreduces the affinity of the GAB on the adjacent actin polymer. The FABregion helps localize the GAB to the filament end. Reproduced, withpermission, from Dickinson (106).

with PI(3,4)P2, and is sufficient to target Lpd to the plasmamembrane (229, 291). Since Lpd recruits VASP and is crit-ical for lamellipodium formation and organization (229), itmay well be part of the actoclampin system. Finally, a recentintriguing report (245) shows that filopodia-like structures(FLS) can form in vitro from lipid bilayers containing 10%PIP2 in the presence of frog egg extracts. The elongatingactin filaments self-organize under the bilayer into paral-lel bundles. The proposed sequence of events is that theF-BAR protein toca (198) recruits N-WASP followed byArp2/3 and actin. Subsequently, the end-tracking proteinsformin and VASP, together with the filament bundling proteinfascin are recruited, and filament elongation occurs. Whilethe acidic membrane lipid is indispensable for FLS formation,no preformed membrane microdomains seem to be necessary.Clearly, future research should address not only the identity of

the additional proteins that link the membrane to the polymer-ization machinery, but also their biochemical and biophysicalregulation. The “absence of a gap” means the presence ofregulated linkers. It will be of particular interest to clarifyhow the external load or the compressive and tensile forces ofthe cytoskeleton affect the interaction of the membrane withthese linker proteins and what functional consequences suchinteractions have.

Force-velocity relationship in lamellipodia

After some indirect approaches (48, 239), the first direct deter-mination of the F-V relationship in the lamellipodia of migrat-ing cells was carried out in fish keratocytes using atomic forcemicrocopy (355) (Fig. 15). A cantilever was placed in the pathof a migrating cell. Upon contact with the lamellipodium, thecantilever is deflected and exerts a Hookean force on thelamellipodium. Knowing the deflection and the stiffness ofthe cantilever, the F-V relationship can be determined. Fur-ther, knowing the lamellipodial area, which is in contact withthe cantilever (estimated to be 3 μm × 0. 2 μm), the numberof filaments abutting the membrane in this area (120-240 perμm) and the stall force, the maximal force exerted by individ-ual filaments can be calculated (≈ 4 pN). The F-V measure-ments gave surprising results. Interestingly, the lamellipodiumslowed down approximately eightfold before it appeared toreach the cantilever. Thus, when the cell arrived within 200 nmof the cantilever, its rear continued to advance at ≈160 nm/s,while the speed of the front dropped to ≈20 nm/s. It is pos-sible that there is a narrow invisible rim at the very tip,which gets into contact with the cantilever, and the ensu-ing very small force (< 100 pN, i.e., below the resolutionof the measurement) is enough to induce this large drop invelocity. This might be due to various possibilities includinga cantilever-associated dampening of the thermal fluctuationsof the membrane, or by local osmotic or hydrostatic pressuresthat might contribute to membrane expansion (see Section“Not only actin polymerization: other mechanisms support-ing membrane protrusion.”). Alternatively, the cells may sensethe vicinity of the cantilever and could accordingly calibrateback force generation. In any case, another report also foundthat very weak forces (few pN/μm) were able to counteractlamellipodial protrusion, (48) without actually stopping actinpolymerization. These results were interpreted to mean thatthese small forces (generated by fluid flow from a pipette)disrupted the formation of weak nascent adhesions at thevery tip, which may be indispensable for protrusion. Theseobservations suggest that lamellipodium expansion cannot beexplained by submembraneous actin polymerization alone.

Notwithstanding the unexplained behavior of the lamel-lipodium at these undetectably small forces, Prass et al. (355)were able to obtain the F-V relationship between the reducedspeed (20 nm/s) and stalling (0 nm/s), where the cantileverdeflections were measurable. This showed that the veloc-ity is force independent at small loads and drops along aconcave line at greater ones. This relationship is different




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Figure 15 Measurement of the lamellipodial stalling force in migrating cells. (A) Experimental setup to measurethe stalling force. The cantilever of the atomic force microscope was placed in front of a migrating keratocyte. Thelamellipodium deflects the cantilever. Measuring the deflection and knowing the spring constant of the cantilever, theforce exerted on the lamellipodium can be calculated. Monitoring the movement of the lamellipodium, the stallingforce (at which the lamellipodial protrusion is prevented) can be determined. (B) Visualization of the interactionbetween the lamellipodium and the cantilever. The keratocyte was fixed shortly after making contact with thecantilever and stained with labeled phalloidin. At the site of contact, the cantilever caused an indentation in thelamellipodium (boxed area), while the adjacent regions of the lamellipodium continued to move forward. (C)Deflection (force)-time curve for the moving keratocytes. (D) The gray box shown in C is magnified. Note that uponinitial contact, there is a sharp deflection, which peaks after a few seconds when the lamellipodium stalls (shadedarea). Subsequently, the lamellipodium escapes (sneaks around) the cantilever and the deflection falls to zero. Thisphase is followed by a second rise, when the cell body (that continues to move) hits the probe. Adapted, withpermission, from (355).

from the predictions of the original BR models (299, 341)and from observations made on Listeria (280) and ActA-covered beads (273), but is similar to simulations based onthe CFE/ETM models (at least in a range of selected param-eters) (106–108) (Fig. 16) and with experiments in whichdeflections of ActA-covered cantilevers was measured duringthe in vitro formation of a branched actin meshwork. None ofthese models have explained the initial drop in velocity at verysmall forces.

It has been increasingly recognized that while the basicmechanisms of pushing may be interpreted according to theBrownian and Actoclampin models, a wide variety of othercytoskeleton-related events have great impact on the over-all membrane protrusion process. These include polymeriza-tion kinetics in complex networks (branching, capping, nucle-ation, and shrinkage), the formation, and breaking of adhe-sive contacts (focal contacts and adhesions), and the so-calledexcluded volume effect (a packing constrains that originates

from the fact that the filaments cannot intersect each other). Arecent stochastic simulation model, which takes into consid-eration the above-mentioned processes, has yielded an F-Vrelationship that rather accurately matches the experimentalobservations (411), although the great decrease in velocity atvery small forces has not been addressed by this model.

Adhesions

In most (but not all) cells, a prerequisite for lamellipodialextension (as well as for tail retraction) during locomotionis the presence and dynamic turnover of cell-ECM adhe-sions, which act as a “molecular clutch” (243). Only when thisclutch is engaged can the force generated by actin assemblybe converted into protrusion, while disengagement is asso-ciated with slippage and retrograde actin flow. Focal con-tacts at the base of the lamellipodium are constantly formedand then either disassembled (to reform again in a forward




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Figure 16 Force-velocity relationships. (A) Typical atomic force microscopy recording showing can-tilever deflection as a function of time in the fish keratocyte lamellipodium, determined as describedin Figure 14. (B) Force-velocity relationship of the lamellipodium obtained from the experiments shownin A. Reproduced, with permission, from Prass M. et al. J Cell Biol. 2006, 174(6):767-72 (355). (C)Force-velocity relationship derived from the theoretical considerations of the filament end-tracking motormodel (Fig. 13). The monomer transfer/edge advancement steps is assumed to be a function of a force(F)-dependent rate constant (kt = kt0 e−Fd/kBT, where d = 2.7 nm). The numbers beside the curves indicatevarious ratios between the baseline transfer rate constant (kt0) and the monomer binding rate. At smallcompressive forces the force-independent monomer binding step is slower than the force-dependenttransfer/advancement step and, therefore, filament elongation is force independent. At larger forces, theadvancement step becomes rate limiting, and ultimately no elongation occurs. The experimental datashown in B are compatible with the end-tracking motor model, whereas they do not correspond wellto the predictions of the original BR model (shown on the graph as well). Reproduced, with permission,from (106).

location) or reinforced to develop into FAs. These loci areimportant force transducers and mechanosensors whose for-mation and turnover is regulated by tensile forces (387), asdirectly demonstrated by single molecule fluorescence forcemicroscopy (159), or high temporal resolution FRAP (506).Other membrane-associated cytoskeleton-matrix interactionsites include podosomes and invadopodia, which are criti-cal for tumor cell migration and invasion (307). While moredetailed discussion of various adhesions is found in reviewson cytoskeleton, cell migration, and mechanotransduction, itis worth mentioning here that new techniques allow insightinto the fine structure of FAs. Using cryo-electron tomog-raphy, four layers have been distinguished. Outermost arethe membrane-embedded integrins, the extracellular domainsof which bind to the ECM. Their cytosolic surface and theadjacent inner leaflet of the membrane interact with

doughnut-shaped FA associated particles (25 nm in diame-ter), which in turn contact short tangential actin filaments thatassociate with stress fibers (acto-myosin bundles) (334). Toresolve the molecular architecture of FAs, Kanchanawaonget al. used three-dimensional super-resolution fluorescencemicroscopy (212). They found that the plaque connectingintegrins to the actin skeleton contains various protein spe-cific strata: closest to the membrane are the integrin cyto-plasmic tails, FA kinase, and paxillin; this is followed by aforce-transduction layer containing talin and vinculin; furtherinside there is an actin-regulatory layer, composed of zyxin,VASP, and α-actinin. Future investigations of focal contactswill be instrumental for our understanding of the critical inter-face between the very dynamic and actively pushing region ofthe cortical actin filaments and the more “static” meshworkfrom which the former originate.




Not only actin polymerization: Other mechanismssupporting membrane protrusion

As mentioned previously, hydrostatic and osmotic forces mayalso contribute to membrane expansion. Such idea is sup-ported by two sets of observations. First, contrary to theprevious paradigm, spatio-temporally localized variations inhydrostatic pressures (e.g., induced by contraction) do existeven in mammalian cells. They occur in the scale of 10 μm and10 s and underlie the formation of membrane blebs (75); Sec-ond, cell migration seems to involve osmotic swelling at thefront (219) and shrinkage at the rear (409, 412). What can bethe mechanism for osmotically driven swelling at the lamel-lipodium? There are two possibilities: an osmotic imbalancegenerated by a biochemical reaction within the lamellipodiumor (and) transmembrane transport of osmotic equivalents fol-lowed the influx of water. The solation-gel expansion modelproposed by Oster and Perelson (327) corresponds to the firstpossibility and states that the osmotic pressure of the actingel drives the lamellipodium (326). At rest, the osmotic pres-sure of the gel is balanced by elastic forces. The osmoticpressure is due to the accumulation of mobile counterions,which neutralize the immobile negative charges of the actingel. If the elasticity is suddenly decreased, for example, dueto severing, unbundling, and capping of filaments, an imbal-ance develops between the osmotic pressure and the (reduced)elastic forces and the gel expands. This expansion can pushthe membrane forward and the cycle can be repeated. Interest-ingly, the process—at least temporarily—can be faster thanthe rate of monomer diffusion to the gel.

The other alternative, that is, the contribution of mem-brane transport-coupled water influx is supported by sev-eral observations. The leading edge of migrating cells showsenriched distribution for the Na+/H+ exchanger (NHE) andthe Cl−/HCO3

− antiporter, the combined operation of whichleads to NaCl uptake followed by osmotically obliged water(99, 162, 219). Importantly, topical inhibition of these trans-porters halts lamellipodium expansion and migration (219).Moreover, NHE activation (383) and the ensuing substan-tial water influx through aquaporin 9 (264) were found tobe indispensable for neutrophil migration, and the effect ofchemotactic agents can be mimicked by hypotonic swelling(383).

How could local hydration support membrane protrusion?Topical swelling may be a non-Brownian mechanism thatgenerates gaps between the filament tips and the membrane(219, 264). This way the osmotic and the polymerization-powered mechanisms might cooperate, explaining the needfor both processes for lamellipodium expansion.

Nonetheless, this issue remains controversial as Hayashiet al. found that NHE affects neutrophil migration via itsimpact on intracelluar pH (pHi) and not on cell volume (176).The mechanism underlying local pHi-dependent effects onactin dynamics is emerging. Submembraneous alkalizationdirectly affects cofilin activity and thereby local filament sev-ering (79), and regulates the interaction of cofilin with the

nucleation regulator cortactin (269). In addition, NHE anchorsthe plasma membrane to actin filaments through ERM pro-teins (99,100). On the other hand, osmotic forces or the hydra-tion state of the cell have a strong impact on the cytoskeletonper se. Physiologically relevant changes in osmotic concen-tration have been described to alter signaling via Rho familysmall GTPases (102–104, 255, 336) and ERM proteins (365)which in turn provoke profound remodeling of the cytoskele-ton. Usually, hyperosmolarity (shrinkage) induces net actinpolymerization whereas hypo-osmolarity (swelling) results indepolymerization (albeit cell-specific differences exist) (103,337, 338). In addition, large osmotic pressures may directlychange the hydration of actin and thereby its polymerization(138).

Conclusions

The last two decades brought enormous progress in our under-stating of the biochemistry and biophysics of membrane pro-trusion. Nonetheless, many fundamental questions are stillopen. Future research should elucidate the F-actin structurewithin the lamellipodium (branching vs. nonbranching), themodes of protrusion (Brownian models vs. actoclampin mech-anisms) and provide insight into the submolecular nature ofthe bidirectional relationship between mechanical forces andbiochemical reactions at the leading edge and in cell adhesionsites.

Membrane curvatureCell membranes exist in a variety of shapes and forms, includ-ing flat or curved sheets, tubules, and vesicles (15, 425, 524).Moreover, local membrane areas constantly undergo dynamicshape changes such as invagination, tubulation, vesicula-tion, and budding, which transform these shapes into oneanother, and underlie key cellular processes including endo-/exocytosis and vesicular transport (123, 425, 426, 491). Thelocal curvature of a membrane area can be characterized bytwo circular arcs, which are perpendicular to each other, anddefined by their radii R1 and R2. The so-called principle cur-vatures are then defined as the reciprocals of these radii, thatis, C1 = 1/R1 and C2 = 1/R2 (524) (Fig. 17) If the value ofR is much larger than the thickness of the bilayer (h ≈ 4 nm)then the curvature is small, and conversely, if h and R are notvery far apart, then the curvature is large. As an example, theh/R ratio for the relatively flat plasma membrane is around10−4, whereas for highly curved vesicular membranes, it isapproximately 10−1 (158). For typical lipid compositions, thelowest radius that a bilayer can maintain without rupture isapproximately 30 to 50 nm. In spherical structures, R1 andR2 are similar, whereas in cylinders such as tubules of the ERor filopodia, one of the R − s is 0 (no curve along the axis),while the other has a finite value.

There are four basic shapes that can be characterized bythe combination of the principal curvatures, and each of theseis present in various natural membranes (Fig. 17). These




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Figure 17 Membrane curvature. (A) The membrane is a two-dimensional surface in a three-dimensionalspace, and to characterize its curvature in a vicinity of a given point on it, two principal planes can be definedthat are perpendicular to the surface and each other and have some other special properties (Euler’s theorem).The intersections of these planes with the surface define two perpendicular arcs with radii R1 and R2, and thereciprocals of these are the principal curvatures. Reproduced, with permission, from (524). (B) Basic curvedshapes. These shapes are characterized and distinguished by their principal curvatures (c1 and c2), their totalcurvature J (defined as the sum of the principal curvatures) and their Gaussian curvature K, (defined as theproduct of the principal curvatures). Adapted, with permission, from (425).




shapes are (i) the flat (e.g., the nearly flat, noninvaginatedstretches of the plasma membrane); (ii) the cylindrical (e.g.,the neck of budding vesicles or invaginated pits, usually sur-rounded by the membrane-pinching GTPase, dynamin (128,187), and tubular structures formed in the secretory path-way, such as transport intermediates between the ER, theGolgi apparatus, and the plasma membrane (345, 346, 356);(iii) the saddle-like (e.g., transitional areas between the necksof the budding vesicles and the membrane they arise from),and (iv) the spherical (e.g., clathrin-coated endocytic vesicles,lysosomes, and coatomer (COPI and COPII)-coated vesiclesinvolved in ER-Golgi anterograde and retrograde transport(218, 247, 384, 408, 441, 461).

A symmetric lipid bilayer tends to be flat, thus the bend-ing of the membrane requires work and the correspondingmolecular machinery that performs it (30). Regarding thethermodynamics of membrane bending, please see an excel-lent recent review (15). From a mechanistic standpoint, twoquestions arise (i) what are the ways to generate membranecurvature and (ii) what is the role of the cytoskeleton in theseprocesses? In the following sections, we describe the majorcurvature-generating mechanisms and briefly allude to theknown or hypothesized contribution of the cytoskeleton tothese. Subsequently, we focus on the regulation of curvatureduring the formation or maintenance of plasma membraneprotrusions, in which the involvement of the cytoskeleton isprominent. With regard to curvature regulation during vesic-ular transport the reader is referred to excellent reviews (201,248, 260, 344, 418, 425).

Finally, two general thoughts warrant mentioning. First,generation, maintenance, and sensing of membrane curva-ture are interconnected processes (15). In fact, the exact roleof several curvature-regulating proteins—as generators ver-sus sensors—remains to be clarified. Second, while a varietyof protein complexes, including cytoskeletal ones, regulatemembrane curvature, there is strong evidence that the curva-ture per se is an important regulatory factor that affects lipidsorting (485), vesicular coat protein assembly (344), mem-brane fusion (288), the recruitment, and function of dynamin(133, 388, 518), and other processes, which in turn can impacton many functions, including cytoskeleton organization (seeSection “Actin-driven membrane processes and BAR domainproteins”). This interdependence, termed the mechanochemi-cal cross-talk during membrane dynamics (260) is a rich areaof current research.

Curvature formation and sensing and itsrelationship to the cytoskeletonThere are several mechanisms whereby membrane curvaturecan be induced (158, 287) (Fig. 18).

1. Altering the lipid composition of the membrane can pro-mote curvature formation, because it causes asymmetrybetween the inner and the other leaflets (due to the uneven

size of the phospholipid head groups or the different con-formation of the acyl chains), which in turn facilitatesmembrane bending. Such asymmetry is generated andmaintained by lipid-modifying enzymes (e.g., phospho-lipases, acyltransferases, and flippases) (484). The recruit-ment of these might be influenced by the cytoskeleton, butthis is an underexplored area that warrants further study.

2. Transmembrane proteins with conical shape promotemembrane curving. For example, the nicotinic acetyl-choline receptor (AChR), a large wedge-shaped molec-ular complex (477) is thought to contribute to the highlyfolded membrane morphology of the neuromuscular junc-tion (287). Importantly, the cytoskeleton has been shownto play a key role in the clustering and retention of AChRin cholinergic synapses (21, 343, 429). The receptor isanchored to the cytoskeleton by the adaptor protein rap-syn, which binds to the spectrin-family member ACF-7,which in turn directly interacts with actin (14). More-over, the MT network also participates in the membranedelivery and stabilization of certain AChRs. This involvesa large molecular complex containing the adenomatosispolyposis coli protein, the microtubule plus end-bindingprotein EB1, and several other adaptors, which interactwith the α-subunit of AChR (382). The integrity of bothactin and the MT skeleton is required for normal AChRdistribution.

3. The cytoskeleton can be a direct inducer of membranecurvature by two mechanisms (Fig. 18):

a. the polymerization of cytoskeletal subunits (actin, tubu-lin) into filaments can push or pull the membrane, asdetailed in Section “Force-generation at the membrane:pushing the envelope” and further addressed in Section“Actin-driven membrane processes and BAR domainproteins”;

b. the membrane can be pulled by molecular motorsthat move along cytoskeletal tracks. Actin-activatedmyosin I motors have been shown to directly con-tribute to the invagination of the forming endo-cytic vesicle (196, 361). Interestingly, force gener-ation through WASP-promoted actin polymerizationand acto-myosin-mediated contraction collaborate inthe induction of the shape change. The MT motorproteins, kinesins, and dyneins play a central rolein vesicular traffic and endocytic sorting (194, 309,423, 489), processes that are associated with dramaticchanges in membrane curvature. It seems likely thatthese motors not only drive the movement of the vesi-cles but also affect their genesis and shape. Indeed,endosomes, lysosomes, and the plasma membrane wereshown to undergo microtubule-dependent tubulation(116, 482, 496). Nonetheless, the available informa-tion in this respect is relatively scarce, which calls formore research in this area.




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Membrane proteinshape

Membrane proteinoligomerization

Scaffolding

Amphipathic helixinsertion

Acyl chaincomposition

Headgroupcomposition

Indirect scaffolding

Direct scaffolding(negative) Direct scaffolding

(positive)

Figure 18 Various mechanisms underlying membrane curvature formation. Curvature can be generated by (A) asymmetric lipid distribution;(B) intrinsic shape of transmembrane proteins or their complexes; (C) pushing or pulling the membrane by polymerization of cytoskeletal filamentsor by cytoskeletal motors; (D) scaffolding, and (E) insertion of amphipathic helices. Further explanation can be found in the text. Reproduced, withpermission, from (287).

4. Membrane bending by scaffold proteins is also a potentmechanism to induce curvature. The classical coat pro-teins such as clathrin, COPI and COPII complexes canself-assemble into cages or lattices with intrinsic curva-ture (403), which can bend the membrane (or keep itin a curved configuration), working as an “exoskeleton”(287). However, several factors necessitate the participa-tion of other components, including cytoskeletal ones, inthe induction of efficient shape change. First, many of thescaffolds do not directly bind to the membrane so theyrequire lipid-binding adaptors (296). Second, although thescaffolds adopt a curved shape, some of them (e.g., theCOP complexes) may not have sufficient rigidity (whichhas to be greater than that of the membrane) to maintaina bent conformation (524). Third, the formed pits even-tually close, pinch off, and move away from the plasmamembrane, and these processes, which are also regulatedby the curvature itself, require many additional molecules.Thus, it is not surprising that there is a multitude of inter-actions between the coat complexes and the cytoskele-ton, both at the plasma membrane (143, 360, 404) and atendomembranes (ER/Golgi) (177, 309, 445). A dynamicactin skeleton is essential for endocytosis in yeast andexerts significant modulatory effects on the process inmammalian cells (143). Actin polymerization and remod-eling have been proposed to affect all steps of endocytosisincluding initiation and propagation of coat assembly, bud-ding, scission, detachment, and the cytosolic movementof the endocytic vesicle (83, 359). While the details arebeyond the scope of this review, it is important to note that

endocytosis regulates actin dynamics, which in turn con-trols and coordinates the steps of the endocytic process.One important link between the endocytic machinery andthe actin skeleton is dynamin itself, which is recruited tothe bud, partially in a curvature-dependent manner (388).Dynamin in turn can locally modulate actin dynamics bybinding to F-actin (164), to actin-binding proteins such ascortactin (289, 302) (which interacts with and regulatesboth WASP and Arp2/3), and to several GEFs that affectRho and Rac activity [see (128)]. Furthermore, recentlythe WASP and Scar-homolog protein WASH has beenproposed to link actin polymerization and microtubuledynamics with retromer-mediated sorting and retrograde(endosome-to-Golgi) transport (154, 470).

Caveolins (1-3) constitute another important set ofcurvature-related scaffold proteins, which reside in lipidrafts and are necessary for the formation of caveolae,that is, flask-shaped, cholesterol-rich invaginations of theplasma membrane (171, 407). Caveolins are somewhat dif-ferent from the other coat proteins (and represent a transi-tion to the fifth group, see below), because they not onlyoligomerize (like the others), but are also directly embed-ded in the membrane. They exhibit an unusual topologyin that their N- and C-termini are cytosolic while theirelongated middle region is immersed in the inner leaflet.Caveolins are acetylated and bind substantial amounts ofcholesterol. Approximately 144 molecules of caveolin-1and 20,000 molecules of cholesterol are estimated to residein a single caveola. How caveolins induce flask shapemembrane invaginations (a very complex geometry) is




not clear, but recent research indicates that they act incollaboration with cavins, a family of proteins that containhydrophobic heptad repeats followed by a basic region thatlikely serve as membrane-targeting sequences (171). Elec-tron microscopy tomography and 3D reconstitution revealsa close relationship between caveolae and microfilamentsas well as microtubules (376). Actin filaments (along withcross-bridges of 3 nm diameter) were proposed to regu-late the clustering of caveolae by linking them togetherand thereby forming a larger caveolar unit. While thestructural basis of the caveolae-cytoskeleton associationremains to be further elucidated, a recent study shows thatthe ATPase Eps-15 homology domain-containing protein 2links caveolae to actin filaments (450). Further, caveolin-1has been shown to bind to the actin cross-linker filamin,and Rho stimulation-induced alignment of caveolae withstress fibers, suggesting a tight mechanical coupling (444).Moreover, ERM proteins and talin were found to localizeto the neck region of caveolae (132). Thus, caveolae in par-ticular and lipid rafts in general are intimately connected tothe cytoskeleton (78, 254). These structural features maybe related to the central role of caveolae in mechanosens-ing and mechanotransduction (55, 332, 519). Indeed cave-olin null animals show defects in flow-, shear stress-, andstretch-induced signaling and adaptive responses (8, 519)[and see (332) for further references]. Regarding the under-lying mechanisms, one intriguing hypothesis is that caveo-lae are involved in the regulation of membrane elasticity, aparameter that is altered by mechanical stress (332). Elas-ticity is determined by the lipid (e.g., cholesterol) compo-sition of the membrane, and it regulates the hydrophobiccoupling between membrane-spanning proteins and thebilayer. This in turn may have profound effects on signal-ing processes. This view is compatible with the other keyfunction of caveolae as well, namely, their role as regula-tors of lipid traffic and membrane lipid composition (197,440). While caveolin concentrates cholesterol in the mem-brane, retrieval of lipids from the membrane can occurthrough caveolar endocytosis leading to the formation of“caveosomes” (315). The cytoskeleton likely participatesin this process as well (171). Clearly, shape regulationand mechanotransduction by caveolae, and the complexrelationship between these processes and the cytoskeletonremain an area of intensive research.

Arguably, the most important scaffolds in sculpting orsensing membrane curvature are proteins containing BARdomains (135, 277, 361, 456). This domain was iden-tified in proteins that function in endocytosis and bindperipherally to the inner leaflet of the plasma membraneat sites where the membrane exhibits positive curvature,that is, it bends locally inward to the cytoplasm. There aretwo mechanisms whereby BAR domain-containing pro-teins can induce membrane deformation: they can act asintrinsically curved, rigid scaffolds that bind to the mem-brane through electrostatic interactions, and they may alsocontain amphipathic helices, which embed into the bilayer.

Due to their importance and widespread relationship withother scaffolds, signaling molecules, and the cytoskeletonwe will discuss various BAR-domain proteins in a separateSection “BAR-domain proteins: some general considera-tions.”

5. Insertion of amphipathic helices into the bilayer (similarto wedge-shaped transmembrane proteins) induces curva-ture. The prototypic representative of membrane sculptorsthat works through this mechanism is the endocytic pro-tein Epsin (190). Epsin family members contain an EpsinN-terminal Homology (ENTH) domain, which has an α-helical superstructure (7+1 alpha helices, named α1-α7and α0). The ENTH also contains a PIP2-binding region,which, when engaged, induces a conformation change thatexposes the α0 helix and allows it to insert into the innerleaflet. While the ENTH domain binds to and bends themembrane, the other parts of the molecule harbor protein-protein interaction domains, through which epsin can bindto clathrin, the membrane-associated adaptor protein AP2,and also to the transmembrane endocytic cargo molecules.In short, epsin not only induces curvature, but also con-nects the key components of the endocytic vesicle. More-over, epsin coordinates vesicular maturation with actindynamics through the regulation of actin-binding adaptorproteins (56). Other important molecules with N-terminalamphipathic helices include the Arf GTPases (which regu-late COP complexes) (357). Interestingly, they may insertpreferentially into the membranes with a given curvature,thus they may function more as sensors than inducers.Finally, several BAR domain proteins (BDPs) (called N-BARs) comprise amphipathic helices in conjunction withthe BAR domains (see below).

BAR-domain proteins: Some general considerations

The common structural feature of BDPs is a positively chargedbanana- or boomerang-shaped region, which interacts withand molds the membrane (135, 277, 361, 456). BAR domainsform dimers (BAR modules), the core of which is a 6-helixbundle, composed of 3 helices contributed by each monomerin a symmetrical manner (278, 342) (Fig. 19). This structurenot only possesses an intrinsic curvature, but also exhibitssufficient rigidity (278) to overcome the bending resistanceof the membrane (123). There are then several variations onthis basic theme, which distinguish groups as well as indi-vidual BDPs within the superfamily (Fig. 19). The threemajor groups, based on sequence similarities, are the BAR/N-BAR, the F-BAR, and the I-BAR families. Crystal struc-ture analyses show that the curvature of the BAR domainsgreatly varies from diameters as low as 15 to 22 nm (e.g.,for arfaptin, endophilin, pacsisns and amphiphysin) in theclassical BAR/N-BAR proteins, through a medium range of50 nm (pacsisns) to very shallow values of 200 nm (e.g.,FCHo and syndapins) in the F-BAR family (180, 342, 361,428, 501). I-BAR (where I stands for inverse) domains have




Cla

ssic

al B

AR

N-B

AR

BA

R-P

HP

X-B

AR

F-B

AR

I-B

AR

Lateral view Top view

Arfaptin(1149)

Endophilin(1X03)

Amphiphysin(2FIC)

APPL(2Q13)

SNX9(3DYT)

FCHo(2V0O)

FBP17(2EFL)

Syndapin(3HAH)

MIM(2D1L)

61°

37°

90°

PX

PH

Figure 19 BAR domain proteins. BAR domains are composed of a symmetric arrangement of twotriple helices. Various BAR domains can be classified into different families and possess characteristiccurvatures. They bend the membrane as scaffolds but may also contain amphipathic helices. Reproduced,with permission, from (361).

the opposite (convex) curvature, which appears to stabilize orinduce negative membrane curvature, that is, protrusion. Thedifferences in the magnitude and direction of their in-builtcurvature predispose various BDPs to preferentially interactwith different membrane surfaces. Further molecular versa-tility stems from the fact that many BDPs are multimodu-lar and contain other lipid- and protein-interaction domains.As already mentioned, N-BAR proteins have an N-terminalamphipathic region, which can bind PIP2 and also insert into

the bilayer (144). Thus, these proteins are targeted to spe-cific lipid environments where they induce enhanced bending.Other lipid-binding modules can be found in the PX-BAR(e.g., sorting nexins) and BAR-PH (e.g., APPLs) proteins,as the names indicate. Through protein-protein interactiondomains BAR proteins link membrane surfaces of definedcurvatures to adaptors or regulators involved in endocytosisand cytoskeleton organization. For example, amphysin bindsto clathrin (293), while TUBA (a BDP, which also has Cdc42




GEF activity) contains several SH3 domains through which itinteracts with dynamin, WASP, and ENA-VASP (397), coor-dinating membrane scission with actin polymerization.

In vitro many BDPs have been shown to induce tubula-tion or vesiculation of liposomes (124, 278, 342). The aver-age diameter varies with the concavity of the particular BARdomain, and the density will determine whether tubulationor vesiculation occurs. Furthermore, experiments and com-putational studies (20) imply that BAR domain dimers canassemble into larger molecular complexes. These results sug-gest that BAR proteins induce not only local bending butalso anisotropic membrane remodeling over larger distances(123), for example, around entire organelles. Observationsabout the diameter-dependent interactions of BAR proteinswith liposomes, together with the involvement of multipleBAR proteins during membrane remodeling processes formthe basis of the so called BAR domain hypothesis (361). Thispostulates that there is a spatial and temporal hierarchy for therecruitment of various BDPs, which is dictated by the actualcurvature of the surface that undergoes remodeling. UsingTIRF microscopy, Taylor and colleagues (469) examined the“recruitment signature” of 34 proteins including ten BARdomain-containing ones during endocytosis. The results sug-gest that the earliest events indeed involve the recruitment ofthe molecules with shallowest curvature, however in the mid-stage of the process BAR proteins with different curvatures getrecruited simultaneously. Thus, the recruitment/release pro-cess is very complex, and although curvature is an importantinput affecting the spatiotemporal dynamics of endocytosis,other factors (e.g., multimerization of BAR proteins and otherprotein-protein interactions) also play key roles.

Actin-driven membrane processes and BARdomain proteins

The shapes of liposomes in which actin or tubulin has poly-merized, as seen in Figure 6, do not closely resemble the typesof protrusions formed by live cells. For example, actin poly-merization by itself leads to relatively modest shape changes,and not the broad lamella or the sharp filopodia, characteristicof polymerization-driven protrusion in cells. This differenceis due to at least three factors. Random polymerization ofactin does not reproduce the polarized growth outward at themembrane surface that is triggered by signals derived fromN-WASP, VASP, or formins. The flexibility of actin filamentsmeans that even a few pN force will buckle a filament that ismore than a μm long, and it is not yet possible to produce thevery high actin concentration and cross-linking density thatlimits the length of a free actin filaments at the cell membraneto a few hundred nms between cross-linking points. A thirdfactor is the finite bending rigidity of the cell membrane thatis difficult to overcome by the force of a single actin filament,and that in the cell appears to be promoted by the function ofvarious BAR superfamily proteins (342, 522).

In fact, a new view is emerging that interprets the com-plex dynamics of actin-driven protrusions in terms of a close

synergy between BDPs and actin nucleators or nucleationregulators (414, 455, 522). What constitutes this synergy?Several key points can be mentioned. First, BDPs can link thepolymerization machinery to curved surfaces. Second, theinteraction of BDPs with the cytoskeleton may determine thedirectionality of the protrusion, attributing a sign (+ or −) ora vectorial character to the polymerization process. Remark-ably, the composition of the molecular machinery associatedwith the formation of deep invaginations such as podosomes,or membrane protrusions such as filopodia or lamellipodia isvery similar. However, podosomes contain a classical (con-cave) BDP [ASAP1 (208)], whereas filopodia are rich in the(convex) I-BAR protein IRSp53, which is also present inlamellipodia (6, 414). Moreover, these BDPs play an essen-tial role in the formation of these actin-based processes,since downregulation of ASAP1 prevents podosome assembly(208), while silencing IRSp53 leads to the loss of filopodia,and—remarkably—overexpression of IRSp53 is sufficient toinduce filopodium formation (109, 231, 414). However, thescenario is more complex, since filopodia are also induced bycertain F-BAR family proteins (166, 358) although F-BARdomains usually sense/induce concave curvatures. This mayreflect the fact that the filopodial membrane contains regionsof positive curvature and/or that certain F-BAR domainsappear to interact with the membrane through their convex-ity [so-called “inverse F-BAR domain” (166)]. Third, BDPsare active regulators of the actin polymerization (and orga-nization) process itself. This may happen through a varietyof mechanisms, because different BDPs can (i) bind to actinthrough their WASP Homology 2 (WH2) domain (279); (ii)have actin bundling capacity (295, 509); (iii) interact, throughtheir SH3 domains, with nucleation promoting factors or theirregulators such as N-WASP, WAVE, Ena-VASP, and formins[see (414, 455)]; (iv) harbor direct binding sites (e.g., CRIBdomain) for small GTPases (e.g., Cdc42 or Rac); (v) possessGAP or GEF activities [see (456)]; (vi) have tyrosine kinaseactivities (87). Obviously, not all these capacities are realizedin any single BDP; rather each BDP represents a particular setof selected capacities. The overall versatility is large and hasa major impact on the formation of local actin networks. Theconcept of curvature-dependent actin polymerization was ele-gantly demonstrated by Takano et al. (464), who showed thatF-BAR proteins promoted in vitro, N-WASP-dependent actinpolymerization on PS-containing vesicles, and this effect wasdependent on the diameter of the liposomes.

The challenge is to understand the various BAR/actinmachinery interactions in the context of building or remodel-ing of particular structures. For example, filopodia formationrequires strong convex curvature at the filopodial tip and con-cave curvature at the filopodial base. It also requires the assem-bly of long nonbranching and bundled filaments. In short, thegenesis of filopodia necessitates the coordinated functions ofactin nucleators, linear elongation machines (formins), fila-ment cross-linkers, membrane linkers, and curvature induc-ing proteins [as illustrated 20 and discussed in more detailin recent reviews (290)]. As mentioned, the presence of the




Formin

VASP

Actin

I-BAR

N-BAR

Figure 20 The structure of the filopodium. The formation of thefilopodium requires the coordinated function of various cytoskeletaland membrane-shaping proteins. The core of the filopodium is com-posed of cross-linked (bundled), nonbranching actin filaments, thegrowth of which is brought about by leaky capper nucleators such asENA-VASP and linear actin polymerization machines such as formins,localized at the tip. I-BAR and N-BAR proteins participate in the forma-tion of negative and positive curvatures at the filopodium tip and base,respectively.

I-BAR protein IRSp53 is a prerequisite for filopodium gener-ation, presumably due to its interaction with the membrane aswell with Cdc42 and other actin nucleators/modulators (mDia,Ena-VASP, and Eps8). During the formation of lamellipodia(i.e., structures with much shallower curvatures), the Arp2/3complex initially localizes to the membrane through its activa-tion by N-WASP, which binds both PIP2 and Cdc42 at the lipidbilayers, while formins appear to interact autonomously withthe membrane either by their binding to PIP2 and other acidicphospholipids (156, 364), or by transmembrane domainspresent in the base of some formin isoforms (77). Nonetheless,while not indispensable, IRSp53 participates also in shapinglamellipodia, presumably by modulating the local Rac activity(140). Clearly, the vast area of structure generation throughBAR/cytoskeleton interactions represents a rich and impor-tant field for future research.

Membrane tensionThe lipid bilayer, which is the continuous phase of the cellmembrane, into which such objects as transmembrane pro-teins are imbedded, generally has the properties of a 2D fluid.Namely, individual lipids or perhaps patches of lipids diffusefreely within the bilayer, with some restrictions due to themore rigid obstacles imbedded in or bound to its surface. Suchfluid membranes have no elastic resistance to shear deforma-tions, but they do have important solid-like elastic resistanceto deformations that alter membrane curvature or the aver-age area per molecule within the plane of the membrane.Resistance to curvature is characterized by an elastic-bendingconstant, often denoted B or κB that quantifies the force

Figure 21 Measurement of membrane tension. A probe such as abead in an optical trap or a magnetic particle is attached to the outersurface of the cell membrane. A force ft is applied to the probe sufficientto pull out a membrane tether. The tether, with a radius Rt is separatedfrom the cytoskeleton to which the rest of the plasma membrane isattached. The force required to pull such a tether depends on both thebending constant and the tension of the membrane.

required to change the radius of curvature of a bilayer awayfrom its equilibrium configuration. This equilibrium config-uration need not be flat, and depends on the chemical com-position of the bilayer, which generally contains hundreds orthousands of different lipids, some of which are not stable in atotally flat membrane and therefore induce spontaneous cur-vature. The other elastic constant, generally denoted Tm is anisotropic stretching constant that quantifies resistance to areachange or, equivalently, measures membrane tension. Bothtypes of elastic response are governed by the free energycost associated with deforming lipid bilayers in any waythat increases exposure of the hydrophobic acyl chains orsteroid rings of the lipids to water, or that increases steric orelectrostatic repulsions among lipid head groups (251, 257,394). The elastic parameters describing bending and stretch-ing are both affected by interactions with the underlying 3Dor 2D cytoskeletons or other peripheral membrane bindingproteins (261, 424). Coordinated changes in actin assemblyand changes in membrane curvature or tension are intimatelyrelated to cell shape changes, creation of cell polarity, and themechanism of motility (27, 147, 215).

Membrane tension in particular has attracted much recentattention as a physical factor that limits or controls shapechanges that are initiated by forces produced by cytoskeletalpolymerization or motor activity. In a simple lipid vesicle,membrane tension arises when the area of the bilayer beginsto exceed the minimal amount needed to enclose the volumewithin it. A series of elegant studies have shown that mem-brane tension in a lipid bilayer vesicle can be calculated fromthe force required to pull a tether out from the surface of thevesicle. The critical force needed to keep a tether extendedfrom the rest of the vesicle depends on the tension withinthe bilayer, the bending constant of the bilayers, and on theadhesion of the vesicle to a rigid surface (438). In the sim-plest case, the force is related to the membrane tension Tm,the bending constant B, and the radius of the tether Rt (seeFigure 21) by the relation

ft = 2π RtTm + π B/Rt




For most cells, the surface area of the plasma membrane isgreater than the minimum required to enclose the cytoplasm,and therefore the membrane tension would be expected tobe near zero unless the cell is osmotically swollen or under-goes a massive volume-conserving shape change sufficient topull out all of the excess area. However, contacts between theplasma membrane and the underlying cytoskeleton changethis picture, and numerous studies suggest that the plasmamembrane is more closely modeled as a film that adheres atmultiple sites or perhaps even continuously to the dense mesh-work of proteins beneath it. Therefore, the effective mem-brane tension of a cell is a combination of purely bilayereffects as seen in liposomes and the dynamic interactionsof the membrane with the cytoskeleton. All aspects of thecytoskeleton-membrane interaction potentially contribute tomembrane tension, from the most nonspecific electrostaticor hydrophobic contacts formed during thermally driven col-lisions to specific anchor sites such as the transmembraneglycophorin-based complex that pins the red cell membranebilayers to the spectrin-actin cytoskeleton at distinct pointsalong a hexagonal meshwork. As in many aspects of themembrane/cytoskeletal interface, poly-PIPs, and in particu-lar PIP2 emerge as major factors linking the membrane to thecytoskeleton (367, 368). The consequences of linking a 2Dfluid bilayer to solid or viscoelastic network are probably bestcharacterized for the erythrocyte and discussed in excellentreviews on this topic (110, 395).

The impact of the 3D cytoskeleton on membrane tensionis equally important and has many effects on the degree towhich cytoskeletal changes can alter cell shape. In broad out-line, membrane tension limits the extent of cell spreading andcounteracts the force generated by formins, N-WASP-Arp2/3and other proteins that promote actin polymerization that cangenerate a propulsive force at the leading edge of the cell (27,192, 366). Therefore, the rapid spreading response of a cellas for example, it readheres to a surface after suspension intothe medium is characterized by low membrane tension thatincreases as the area/volume ratio increases during spread-ing and eventually slows the cytoskeleton-mediated extensionof the cell periphery. Thereafter, local and global cell shapechanges (215) acquisition of cell polarity (192, 301), and pro-cesses such as endocytosis and exocytosis (54, 147, 427) thatchange the area of the plasma membrane all appear to betightly controlled by the membrane tension. What sets thistension is not clear, but osmotic regulation and the activity ofmotor proteins such as myosin I or myosin X that directly linkthe cytoskeleton to membrane lipids are increasingly impli-cated in membrane tension generation.

Transmembrane potential and surface chargeThe membrane potential, that is, the potential differenceacross the plasma membrane is primarily an electrodiffusional(Nernst-type) potential, which results from the unequal dis-tribution of and selective membrane permeability for mobilecharge carriers (ions) (333). This potential difference exerts

two major effects: it is a component of the driving force(positive or negative) for all electrogenic transport processesacross the membrane, and it regulates a large variety of pro-teins (e.g., voltage-sensitive channels) the conformation ofwhich is altered by changes in the transmembrane electricfield (42, 50, 72). Because alterations in the membrane poten-tial initiate a multitude of signaling events (e.g., Ca2+influxthrough voltage-gated Ca2+ channels), which in turn can pro-voke cytoskeletal effects (e.g., actin-myosin interaction andcontraction), the transmembrane potential has a major butindirect role in the control of the cytoskeleton. Conversely,the cytoskeleton (e.g., by transmitting forces) participates inthe regulation of mechanosensitive channels and other trans-porters, which in turn affect the membrane potential (222,223,393). While very important, this mutual interdependence rep-resents an indirect form of regulation. The rest of this reviewconcentrates on a different kind of regulation, wherein theelectrical properties (charge distribution) of the membraneitself constitute the signal that impacts the cytoskeleton. Mem-brane surface charge (see Section “Electrical double layer atthe inner leaflet of the plasma membrane”) emerges as a keyfactor in cytoskeleton-modifying signal transduction duringcomplex membrane-remodeling processes, including endo-cytosis, autophagy, and phagocytosis [for recent reviews, see(151, 161, 512, 514)].

Electrical double layer at the inner leaflet of theplasma membrane

The plasmalemma contains a substantial amount of anionicphospholipids, the headgroups of which carry a net nega-tive charge at physiological pH. The most abundant of theseare PS (constituting approx 2%-10% of total cellular lipids,and as much as 15% of the inner leaflet of the membrane)and various PIPs, including PI (≈8% total and 10% of theinner leaflet), and its phosphorylated derivatives such as PI4-phosphate (PI4P) and PIP2, each of which represent 1% to5% of the lipid content of the inner leaflet (286, 362, 490,512). Importantly, with increasing phosphorylation of theinositol ring, the molecules become increasingly negative,for example, PIP2 carries −3.5 charges, due to an equimo-lar presence of tri- and tetravalent ionic species. The mem-brane is highly asymmetric, meaning that the vast majorityof the PIPs resides in the inner leaflet (362, 490). The conse-quence of this lipid distribution is the formation of a doublelayer at the cytosolic surface of the membrane, comprisedof a (quasi) fixed monolayer of negative charges (the surfacecharge), which in turn attracts mobile cations and positivelycharged peptides/proteins, leading to their submembraneousaccumulation. The potential profile of such a charged mono-layer is described by the Gouy-Chapman-Stern theory (281).The entire potential difference between the charged surfaceand the bulk (the cytosol) is referred to as the surface poten-tial. Various components of this can be distinguished basedon the finer structure of the transition between the chargedsurface and the bulk phase (Fig. 22). Thus, the presence of




Figure 22 Surface charge and the components of the surfacepotential. The distribution of differentially mobile layers of ions inthe vicinity of a charged particle (or the negatively charged innerleaflet of the membrane) and the corresponding components ofthe surface potential are shown. Modified from Life Science Leaderhttp://www.bioresearchonline.com/article.mvc/Automated-Protein-Characterization-With-The-M-0002. Reprinted, with permission, fromMalvern Instruments Ltd., www.malvern.com.

fixed charges at the membrane immobilizes a layer of coun-tercharges (the Stern layer) in its immediate vicinity. This isfollowed by a so-called slipping plane, where charge distribu-tion is still skewed towards counterions, which, however, arenot “immobilized.” The potential difference between the bulkand the Stern layer is called the Stern potential, and betweenthe bulk and the outer boundary of the slipping plane is thezeta potential. Charge separation at the membrane creates asubstantial electric field (105 V/m), but the potential dissipatesin a very short distance (281). The surface potential decays(according to the original Gouy-Chapman model) exponen-tially, and it can be characterized by the so-called Debyelength, at which it drops to 1/e of its original value. Thislength, which is directly proportional to the charge densityat the surface and inversely correlates with the ionic strengthof the medium (cytosol), is a good indicator of how effec-tively the surface charge is screened. It is estimated to be1 nm, implying that the membrane can have a strong impacton the distribution of charged molecules within this narrowstrip.

Biological impact of surface charge: Membranelocalization of key regulators

The major biological importance of the electrostatic proper-ties of the membrane has been recognized and emphasized byMcLaughlin, Murray, and their colleagues (281, 284). Theseauthors showed that polybasic regions, present in a varietyof proteins including MARCKS (217) [an actin filament andplasma membrane cross-linking protein involved in mem-brane ruffling and spreading (174, 310, 311)], c-Src and v-Src(products of the Src (proto)oncogene) (60) and the tail of the

EGFR (284) can associate, by virtue of electrostatic interac-tions, with lipid vesicles that contain anionic phospholipids.Similarly, Silvius and colleagues demonstrated that the poly-basic carboxy terminal region of the small GTPase proto-oncogene K-Ras (which can rapidly and reversibly bind tothe plasma membrane (517)) exhibits strong preference forlipid vesicles with more negative surface charge (253, 389),inasmuch that a 10 mol% difference in anionic lipid contentcaused a 45-fold change in binding. While the net charge wascritical for this change, the type of the particular anionic lipidwas not important. Direct evidence that membrane localiza-tion of K-Ras requires the polybasic motif came from elegantmutational studies, in which the net positive charge (+8) ofthe tail was gradually diminished by lysine-to-glutamine sub-stitutions (170, 203, 389). This resulted in the dissociation ofK-Ras from the membrane and the loss of its cell-transformingcapacity. Investigating the intracellular distribution of 125 flu-orescently tagged small GTPases, the Meyer lab found that48 of these localized to the plasma membrane and 37 proteinsin the latter set contained positively charged amino acid clus-ters (181). The majority of these proteins dissociated from theplasma membrane only when both PIP2 and PIP3 levels weredrastically reduced by overexpression of a PIP2 phosphataseand inhibition of PIP3 synthesis. While these observationsimply that the polybasic clusters are critical for membranetargeting, substantial evidence indicates that they are often notsufficient, and a second signal is also necessary. This is usu-ally a covalent lipid modification of the proteins via acylation(e.g., palmitoylation), myristoylation, or prenylation (i.e., theaddition of farnesyl or geranylgeranyl moieties). For exam-ple, many Rho and Ras family members (Rho, Rac, Cdc42,K-Ras, etc.) are prenylated at a conserved cysteine in theirC-terminal CAAX motif, which is adjacent to a polycationicregion, while Src and MARCKS are myristoylated in their N-terminus and harbor polybasic clusters nearby or further away,respectively (169, 202, 251, 308, 374, 375). These observa-tions revealed that the plasma membrane targeting of manyproteins is supported both by hydrophilic and electrostaticinteractions (dual signal), constituting a “coincidence detec-tor” mechanism, which may serve to ensure that proteins areaccurately targeted to the inner leaflet and do not end up asso-ciating with other positively charged structures, for example,the DNA (512). It is worth noting that firm membrane attach-ment may be brought about by dual lipid modification as well,for example, H-Ras, in contrast to K-Ras, has no positivelycharged stretch but is dually palmitoylated, which endows thisGTPase with a high and charge-independent affinity for theplasma membrane (80, 512).

Many key regulators of the cytoskeleton, for example,(Rho GTPases, K-Ras, and Src) contain polybasic clustersand are activated at the plasma membrane, which raises thepossibility that electrostatic interactions may play an impor-tant role in the control of these signal transducers and therebycytoskeletal organization. In addition to the central role ofRho proteins in the regulation of the actin and tubulin orga-nization in a plethora of physiological functions (317, 398,




442), a key feature of malignant transformation by oncogenes(e.g., Ras and Src) is a dramatic remodeling of the cytoskele-ton, which manifests in increased motility, invasiveness, anddecreased adhesion (69, 114, 165, 352). Thus, a key questionis how the electrostatic interactions themselves are regulated.This is addressed in Section “Regulation and monitoring ofmembrane surface charge-dependent interactions.”

Regulation and monitoring of membrane surfacecharge-dependent interactions

Changes in surface charge have been postulated to operateas “binary electrostatic switches” which can turn on andoff the membrane localization of various polypeptides (282,421). In principle, the electrostatic interactions of proteinswith the plasma membrane can be regulated either by chang-ing the charge of the protein or that of the membrane. Thefirst mode of regulation can be brought about either by post-translational modification or by protein-protein interactions.The most common charge-altering modification is phospho-rylation, which reduces the positive (or increases the nega-tive) electrostatic character of proteins. This mechanism isexemplified by protein kinase C-mediated phosphorylation ofMARCKS, which reduces the net charge of the polycationicmotif and results in the dissociation of MARCKS from themembrane (174, 381, 472). Little is known about the roleof other potentially charge-affecting modifications, such asacetylation and ubiquitination of lysine residues, a topic wor-thy of future investigations. MARCKS are also regulated bythe alternative mechanism, that is, protein-protein interaction:upon a rise in intracellular [Ca2+], calcium-calmodulin com-plex is formed, which in turn binds to the polybasic regionof MARCKS. This event effectively shields the charge of thecationic motif and releases the protein from the membrane(16, 174, 217). Detaching of MARCKS (and other surfacecharge-regulated proteins) unmasks those anionic headgroupsto which it had been bound, making them accessible for otherelectrostatic interactions (145, 150, 498). Similar calcium-calmodulin-dependent mechanisms have been proposed withregards to the cytosolic tail of the EGFR and other proteinsas well (283, 284, 457).

Whether changes of the surface charge of the membraneper se can play a regulatory role in membrane-protein interac-tion and protein targeting, was a difficult question to address,because the lack of appropriate methodology to monitor sur-face charge. A major breakthrough in this area was made pos-sible by three important advancements: (i) a major step wasthe development of genetically encoded fluorescent probes(fusions between a particular lipid-binding domain and GFP),which are highly specific for various lipids (e.g., differentPIPs) and can visualize rapid, localized changes in live cells[for excellent reviews, see (23–25, 195, 487, 488)]. Manyof these probes were developed by the laboratories of TamasBalla and Tobias Meyer (ii). The use of these probes led to thediscovery and characterization of profound topical changesin inositol lipids during complex membrane-remodeling

processes, particularly phagocytosis and autophagy [forreviews see (101, 111, 161, 321, 431, 492, 512, 514). (iii)Finally, in addition to these lipid-specific indicators, sensitiveprobes have been developed to monitor in situ changes inmembrane surface charge (515).

We will illustrate the principles and significance of sur-face charge measurement in the context of phagocytosis, asit was developed in the laboratory of Sergio Grinstein. Dur-ing phagocytosis, a particle (e.g., bacterium, IgG-coated bead,and opsonized red cell) binds to the surface of a phagocyte andactivates phagocytic receptors, which in turn induce the for-mation of actin-rich pseudopods around the particle (phago-cytic cup), eventually engulfing it in a membrane-enclosedphagosome. As revealed by lipid-specific probes, the forma-tion of the phagocytic cup, the sealing of the phagosomeas well as its subsequent internalization and maturation areaccompanied by dramatic changes in the lipid compositionof the phagocytic membrane areas. Briefly, PIP2, which isabundant in the resting membrane, exhibits a transient initialincrease during phagocytosis, which peaks at the middle ofcup formation, followed by a major decrease during the seal-ing process, dropping much below the level observed in theresting (nonphagocytic) membrane areas. In contrast, PIP3 isvery low in the resting membrane, starts rising during cupformation, reaches its maximum during sealing and gradu-ally declines thereafter. Diacylglycerol and phosphatidic acidshow further delayed (right-shifted) transients. The interestedreader is referred to excellent reviews and original papersregarding the numerous changes in the metabolic and syn-thetic processes underlying these fluctuations of the phos-pholipid levels (51, 52, 122, 492, 514). These robust changesin the concentration of various phospholipids raise the pos-sibility that the net surface charge of the membrane mayindeed change during phagocytosis. Many proteins show tran-sient association with the nascent or mature phagosome: forexample, from a cytoskeletal point of view, Rac is transientlyrecruited to the forming phagosome (189, 276, 415) and actinitself shows a robust accumulation during cup formation andthe sealing phase, followed by rapid loss at the bottom of thephagosome and complete disappearance later (189, 276, 415).The generation of the actin meshwork is the driving force ofpseudopod formation, and the submembraneous actin mayalso help propel the nascent vesicle inward. However, at laterstages the actin shell should disappear to allow the matura-tion and fusion of the phagosome. Similarly, the various lipidkinases and phospholipases responsible for the ensuing lipidchanges are recruited and released in a spatially and tem-porally tightly controlled fashion. This scenario raises twoquestions: (i) does the surface potential change during phago-cytosis? And if so, (ii) is the surface charge of the membranea genuine regulated and regulatory factor in protein traffic? Inother words, are certain proteins recruited/released primarilydue to changes in membrane surface charge, as opposed to(or in addition to) changes in the concentration of specificlipids? Are there both lipid-specific and potential-dependentrecruitment mechanisms?




To address these questions, Tony Yeung in the Grinsteinlab generated novel surface charge probes (515). The pro-totypic probe was based on the C-terminus of K-Ras [con-taining 15 amino acids, including the polybasic region andthe prenylation site (pre)] fused to Red Fluorescence Protein(RFP). To prevent phosphorylation and ubiquitination, serineand threonine (T) residues were replaced by alanines (A),while lysines (K) were substituted with arginines (R). Thefusion protein (called R-pre-RFP) had a charge of +8 andshowed nearly exclusive accumulation in the plasma mem-brane. In vitro liposome association assays also confirmedthat the probe accumulates in PS or PIP2 containing vesicles,and shows no preference for either lipid. Moreover, dissi-pation of the surface potential, either by a Ca+ ionophore(which induces charge shielding at the membrane, activatesthe aminophospholipid scramblase and induces PIP2 hydrol-ysis) or ATP depletion [which abolishes PIP2 accumulation inthe inner leaflet by inhibiting the lipid flippase(s) (265, 354)]induced complete dissociation of the probe from the plas-malemma. All these experiments confirmed that the probewas a true indicator of surface charge. Importantly, duringphagocytosis of IgG opsonized beads by macrophages, R-pre-RFP was initially present at the base of the cup and in thegrowing pseudopods, but was then depleted from the base, andfully disappeared from the vesicle upon completion of engulf-ment. These observations verified that the phagosome mem-brane undergoes major changes in surface charge. Moreover,three labeled cellular proteins, K-Ras, Rac, and Src, each ofwhich harbors a polycationic motif, showed essentially iden-tical spatial and temporal patterns of phagosomal associationand dissociation as R-pre-RFP. Remarkably, the constitutivelyactive (GTPase-deficient) Rac mutant (Q61L) behaved simi-larly, indicating that the effect of Rac is more likely to be ter-minated by its surface potential-regulated dissociation fromthe phagosome than by the cessation of its activity. Finally,the use of probes with less net negative charge [−4 and −2]verified a hierarchy in the electronegativity of cellular mem-branes, the plasmalemma being the most negative, followedby a tubulovesicular endocytic compartment, the maturingphagosome, and finally the mitochondria and vesicles of thesecretory pathway. This hierarchy may determine the relativepartitioning (membrane association) of proteins with differentnet charge, signifying an “address code” for the delivery andretention of various cationic ligands (161, 512).

What is the reason for the drop in surface potential inthe forming phagosome? To address this question, Yeung andcolleagues sought to develop a PS-specific probe, since PSis the most abundant anionic lipid in the membrane. Thisgoal was achieved by fusing a specific, PS-binding region,the C2 domain of lactadherin, to GFP (511). Using this noveltool, they found that although PS is a significant componentof the negative surface charge in essentially all membranesfrom the plasmalemma to the ER (511), its level does notshow major variations in the phagosome during phagocytosis(513). It is therefore likely that the change in PIP2 is the mostimportant component in drop of the surface charge during

phagocytosis. This means that PIP2 has an important dualrole in the regulation of protein traffic: it specifically bindsto proteins that bear PIP2-interacting domains (e.g., certainPH domains (249, 250, 400, 493), and it is a major partner instrong yet nonlipid-specific electrostatic interactions.

In summary, membrane surface charge emerges as adynamically regulated factor that (together with the posttrans-lational modification of the charge profile of polypeptides) isa key determinant of membrane-protein interactions. The roleof electrostatic switches in the membrane recruitment andrelease of a large numbers of signaling molecules (e.g., lipidkinases, phosphatases, and phospholipases, which are not onlyregulated by surface charge but are also key regulators of thisparameter) is an important area of future study.

ConclusionIn the era of whole-genome sequencing and genome-widemicroarray analysis, the demand of “making sense” of theperplexing wealth of the newly generated information—of this “inventory of life”—keeps rapidly increasing. Thechallenge is to integrate the multitude of molecular play-ers and their interactions into complex structures and func-tions, which can serve as the basis to interpret many funda-mental phenomena of physiology and pathology. Biophys-ical approaches have played and keep playing an essentialand increasing role in this integrative process. In addition tothe classic repertoire of physical and physicochemical meth-ods to study protein and lipid structure and function, a vari-ety of novel functional imaging techniques has indeed rev-olutionized research in the field of membrane/cytoskeletoninteractions. These methods include (but are certainly notrestricted to): atomic force microscopy (AFM) (9, 305) andthe related single molecule-force spectroscopy (57, 449), opti-cal tweezers (483), TIRF microscopy (129, 163, 371) fluores-cence correlation microscopy (474), single-molecule trackingtechniques (391) and quantum dot technology (486), specklemicroscopy (500), fluorescence resonance energy transfer(266, 467), FRAP (369) electron tomography (38), and others.The development of many of these techniques went hand inand hand and was facilitated by the ever-increasing sophisti-cation in confocal microscopy. Together, these methods haveprovided insight into membrane and cytoskeletal dynamicswith a resolution of tens-of-nanometer distances, microsec-ond times, and pN forces. Literally, a new vision is beinggenerated by these biophysical approaches, and the currenttrend suggests that the best is yet to come.

AcknowledgementsWe are indebted to Dr. Monika Lodyga for her contributionto assembling Table 1 and to Dr. Katalin Szaszi for criti-cal reading of the article. The original studies by A. Kapuscited in the review were supported by the National Engi-neering and Research Council (NSERC) of Canada and the




Canadian Institutes of Health Research (CIHR). P. Janmeywas supported by NIH grants GM096971 and GM083272.

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Plasma Membrane-Cortical Cytoskeleton Interactions: A Cell Biology Approach with Biophysical Considerations

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