The Filament System Its Involvement in Cell Migration and Neurotransmitter Release

Anna-Karin Johnsson

Department of , The Wenner-Gren Institute, Stockholm University, 2011

©Anna-Karin Johnsson, Stockholm 2011

ISBN 978-91-7447-292-9

US-AB, 2011

2

SUMMARY

The system consists of actin filaments as the major component and is regulated by a number of actin binding . It is juxtaposed to the plasma membrane where it forms a dense cortical weave from where it pervades into the cell interior. This filament system has multiple roles and participates in virtually all motile processes where its dynamic activities depend on receptor mediated signaling leading to constant and depolymerizations. These activities are now also known to affect gene regulation. This thesis discusses these dynamic reorganizations of the microfilament system and how components are supplied to support these motile processes. The focus is on /profilin:actin, actin and the localization of the transcripts of these proteins. The localization of profilin mRNA was examined in relation to the distribution of β-actin mRNA using fluorescent in situ hybridization. It was concluded that both these mRNAs localize to sites of massive actin polymerization called dorsal ruffles albeit the data obtained suggests that this localization must be dependent on distinct mechanisms. Additionally and cell motility was studied after depleting the two profilin isoforms 1 and 2. The activity of the transcription factor SRF is known to be coupled to microfilament system dynamics via the cofactor MAL which binds to actin monomers and is released upon receptor mediated actin polymerization. Depletion of profilin was seen to influence SRF dependent signaling, most likely because the lack of profilin enables more MAL to bind actin monomers thereby preventing SRF dependent transcription. Finally, it was also investigated how the synaptic vesicle synaptotagmin 1 which is involved in exocytosis, has a role in actin polymerization. This protein has previously been described to cause filopodia formation when ectopically expressed. A polybasic sequence motif was identified as the effector sequence for this activity and it was established that this sequence interacts with anionic lipids. It is also discussed how this sequence could have a role in neurotransmitter release and actin polymerization in the nerve synapse.

3

LIST OF PUBLICATIONS

Paper I: Microtubule-dependent Localization of Profilin mRNA to Actin Polymerization Sites in Serum- stimulated Cells Johnsson, A-K., Karlsson, R. (2010) European J of Cell Biology 89:394-401.

Paper II: Profilin I and II are Both Influencing SRF-dependent Signaling in B16 Melanoma Cells and Loss of Profilin I Interferes with Cell Migration Sadi, S., Johnsson, A-K., Karlsson, R. (2011) Manuscript.

Paper III: Synaptotagmin 1 causes Phosphatidyl-(4,5)- bisphosphate-dependent Actin Remodeling in Cultured Non-neuronal and Neuronal cells Johnsson, A-K., Karlsson, R. (2011) Submitted, under review.

4

TABLE OF CONTENTS

Introduction ...... 7

Cell motility and microfilament system organization ...... 8 Intracellular trafficking ...... 13 Actin structure ...... 14 Profilin ...... 19 PIP2 and microfilament dynamics ...... 27 Rho GTPases ...... 29 Actin regulatory proteins ...... 32 Actin depolymerizing factors ...... 46 Intracellular transport and ...... 48 Cell adhesion ...... 51 mRNA localizaiton ...... 54

Actin and profilin in the CNS ...... 59 Neuritogenesis ...... 59 Actin in the presynapse ...... 66 Actin in the post synapse ...... 70

Synaptic vesicle fusion ...... 75 SNAREs ...... 75 Synaptotagmin ...... 76

Investigations ...... 85 Aim ...... 85 Paper I: Microtubule-dependent Localization of Profilin I mRNA to Actin Polymerization Sites in Serum-stimulated Cells ...... 85 Paper II: Profilin I and II are Both Influencing SRF-dependent Signaling in B16 Melanoma Cells and Loss of Profilin I Interferes with Cell Migration ...... 87

5

Paper III: Synaptotagmin 1 causes Phosphatidyl-(4,5)-bisphosphate- dependent Actin Remodeling in Cultured Non-neuronal and Neuronal Cells ...... 89

Acknowledgements ...... 91

References ...... 92

6

INTRODUCTION

Motility, adhesion, cytokinesis and many other cellular fundamental processes are supported by three distinct protein filament systems: the microfilament, microtubule and systems, which most often are referred to as the . Not only are these filaments by themselves dynamic but the arrangements which they form are dynamically organized and the activities coupled to them tightly regulated. Therefore their collective designation as a “skeleton” is misleading but has nevertheless become general in textbooks and scientific articles. For convenience this thesis sometimes also uses the word cytoskeleton to denote either the microfilament system formed by actin or as a reference to all three filament systems. However, the reader is urged to ignore any association with a “skeleton” or “scaffolding” when reading. The microfilament system - the focus of this thesis - produces the force required for migration and for various intracellular transports, it is the “muscle” of the cell. The central building block of the microfilament system is actin. It was first known as a muscle protein but with the work by Hatano, which led to its isolation from the protist Plasmodium polycephalum, the insight grew that filaments observed in this as well as in other non-muscle cells in fact were a polymeric form of actin, i.e. actin filaments or , (Hatano and Oosawa, 1966). The field has developed massively since then and it is now more clear than ever how complex and tightly regulated the dynamic actin arrangements in fact are and how the force-generating activity produced by the microfilament system is integrated with most major cellular processes. Lately it has also been realized that there are actin-like proteins in prokaryotes (Graumann, 2009). This thesis, however, is focused on the control of actin in eukaryotic cells and in particular the role of the microfilament system arranged in direct juxtaposition to the enclosing lipid bilayer of the cell remodelled for motile activities and transport and how the polymerizing and depolymerizing processes in the cytoplasm also have a role in control of transcription.

7

CELL MOTILITY AND MICROFILAMENT SYSTEM ORGANIZATION

The microfilament system, formed by actin, and regulatory proteins, is responsible for many motile processes in eukaryotic cells. Formation of actin filaments and higher order structures by actin polymerization is one force-generating mechanism which contributes to the motility and, additionally, force is generated by acto-myosin based contractions of subcellular structures where actin and myosin filaments are organized into contractile elements. The three most commonly identified structures in cells formed by filamentous actin are the flat lamellipodia protruding from leading edges, the needle-shaped filopodia which most often project from lamellipodia and the contractile bundles called stress fibres. The abundance and organization of these actin based structures depend largely on type and metabolic state of the cell. They have been extensively studied in motile cells and it is well established that their dynamics – i.e. more or less continuous reorganization – is the result of signals which results from interactions at the cell surface, with hormones or other extracellular molecules or molecules exposed at the surface on neighbouring cells. The family of small Rho GTPases are commonly involved in signalling to the microfilament system and the formation of actin based structures is controlled by many different actin binding proteins and regulatory proteins, which in turn are under control of a diverse array of components including lipids and ions as well as posttranslational modification such phosphorylation. A balance of these actin polymerization-directed control activities, in many respects controls the behaviour of a cell. Here I will breifly describe some of the microfilament associated components and their functions in respect to actin and how the microfilaments are organized in the cell.

The lamellipodium Just below the plasma membrane there is a dense organization of filaments often referred to as cortical actin (fig. 1). Together with the lipid bilayer it forms a dynamic proteo-lipid arrangement which maintains the integrity of the cell and strongly contributes to the organization of surface molecules for instance by connecting to transmembrane receptors. The cortical actin at the front of a migrating cell forms the lamellipodium, an often seen, highly specialized veil like

8 structure extending in parallell to the substratum (Hoglund et al., 1980; Lindberg et al., 1981; Small, 1981). Since the lamellipodium is so flat, around 150-200 nm, it basically gives a ‘confocal view’ of the macromolecular architecture and is therefore an excellent structure for imaging studies of the microfilament system and consequently it is one of the most well studied “organells” in the field of actin dynamics. Lamellipodia of cells migrating in tissues have a somewhat different apperance but its protrusion still requires the force generated by the microfilament system meaning that both in the cultured and the pysiological situation the same sort of dynamic remodelling is constantly occuring. The actin filaments of the lamellipodium are all organized with their fast polymerizing ends, called (+)-ends, located just below the plasma membrane (fig. 1). Thus although they are crossing each other and have different angles in relation to the plasma membrane, the overall filament organization is highly ordered and is related to the cell advancement (Koestler et al., 2008). The cell edge progresses forward by actin monomers being incorporated at the (+)-ends and the filaments are depolymerized at the lamellipodial rear, at the filament (-)-ends exhibiting a so called treadmilling activity. The original characterization of the lamellipodial actin weave (Hoglund et al., 1980; Small, 1981) was later challenged by the presentation of the dendritic nucleation model (Mullins et al., 1998; Svitkina and Borisy, 1999). According to this conjecture, filaments are branching off from other filaments at a 70o angle in a mechansim depending on the Arp2/3 complex, which will be described further below. This model is under debate since it seems that the method used for cell-fixaton in these studies does not preserve the structure of the actin weave. Based on the original observations using a different method for preservation (Hoglund et al., 1980; Small, 1981) and especially more recent data obtained from elctron microscopy (EM) tomography where individual filaments can be followed through several focus planes at high resolution, it now seems clear that essentially no branches are present in the cortical weave (Small, 2010), the branches seen in these studies giving rise to the dendritic nucleation model were an artifact by filaments crossing each other. However, the rapid and wide acceptance of this model, which even has been included in basic text books, led to a large body of data beeing interpreted on its basis, and there will certainly take time before the re-evalutaion of this conjecture reaches full acceptance.

9

Fig 1. Representation of actin organization in the cell. (A), displaying a scanning micrograph of a spreading cell. (B), Micrograph of the lamellipodium showing the filamentous actin after removal of the plasma membrane. Note the difference in organization of the network in the lamellipodium built up by individual actin filaments versus the bundled filament organization of the filopodium. (C), A cartoon representing actin monomers organized into a filament. Illustrations are not drawn to scale. Micrographs from (Hoglund, 1985).

Filopodia Microspikes are bundles of filaments embedded in the lamellipodium and if these protrude independently of the rest of the cell periphery needle shape processes called filopodia are formed (fig. 1) (Mattila and Lappalainen, 2008; Small et al., 2002). The filaments of these structures, like in the lamellipodium, have their (+)-end facing the plasma membrane. Usually these processes consists of a bundle of 50-100 actin filaments with the lipid bilayer of the plasma mebrane smearing over the surface. They probe the cell surroundings for “information” and at their tips various recepor molceules have been observed like for instance growth factor receptors, integrins and cadherins. Hence these structures are involved in contact formation with the extracellular matrix (ECM) or with neigbouring cells. Filopodia continously connect and disconnect to the substraum, and when they are disonneced they wave around quite freely. They are also often observed to grow and shrink on minute timescales and they can do this independently of the behaviour of the lamellipodium. Microspikes are proposed to be formed by filaments of the lamellipodium converging by the forces of a so called lateral flow, a concept which means that filaments flow to the side as the lamellipodium protrudes (Nicholson-Dykstra and Higgs, 2008; Small, 1994; Steffen et al., 2006; Svitkina et al., 2003). In addition to receptor molecules, the tips of the filaments harbor components, of which filament elongators such as and VASP are important actors, these will be described further below. It is reasoned that as more filaments converge, a tip complex is formed which can start to elongate this bundle of filaments leading to the protrusion of a filopodia (Small et

10 al., 1998; Svitkina et al., 2003). However, since filopodia can form independently of lamellipodia other mechanisms must exist for their initiation (Gomez et al., 2007; Nicholson-Dykstra and Higgs, 2008; Sarmiento et al., 2008; Steffen et al., 2006). Also in such a situation a specialized assembly-machinery containing actin filament elongators is proposed to cause the actin polymerization (Faix et al., 2009; Ladwein and Rottner, 2008). The actin structures of the protruding and retracting cell edge are interconnected and they constantly form new constellations of filament arrangements. Filopodia can become retraction fibers which form if the lamellipodium withdraws but leaves the filopdium behind. Filopodia can also fold back and incorpotate into the filament arrangement of the lamella (Nemethova et al., 2008) a structure which continues just behind the lamellipodium and has a less dense organization of actin filaments but with stronger connections to the substratum. As the lamelipodial actin filaments flow laterally when the edge protrudes it causes the filament angles to change in relation to the plasma membrane thereby contributing to reorganizations of the filament assemblies. The lamellipodia and lamellae during protrusion sometimes transform into a wave-like motion called ruffling (Abercrombie et al., 1970). This phenomenon, which is most common in fibroblasts, arises when the edge become detached from the substratum because of the tensile forces which depend on actomyosin activities. Sometimes ruffles are also formed on the dorsal upper surface of the cell away from the extending edge (Buccione et al., 2004). These dorsal ruffles are most commonly seen after stimulation with growth factors and reflect extensive actin polymerizations (Legg et al., 2007); in this thesis this latter activity was utilized to study directional distribution of mRNA encoding the actin binding protein profilin (see paper I and text below)

Stress fibres Stress fibres are bundles of hundreds of antiparallel filaments overlapping with bipolar non-muscle myosin II filaments forming contractile elements that resemble a muscle fibrill and are capable of generating tension by contraction. Towards the periphery, these bundles anchor at the cytoplasmic side of the plasma membrane in multiprotein structures called focal contacts or focal adhesions where the cell, via transmembrane integrins attaches to the extracellular matrix (fig. 2) (ECM). At their interior end the stress fibres connect either to more centrally located adhesions or associate with a complex network of

11 filaments surrounding the nucleus and contributed for by all three of the filament systems of the cytoskeleton (Dupin et al. 2011). The term stress fiber refers to old cell biological litterature where these structures were anticipated to be under stress and it is used more or less for describing all contractile actin bundles within the cell. There are however different types of these structures that are assembled by different mechanisms mainly depending on where they are localizing in the cell (Hotulainen and Lappalainen, 2006). Presently, how different types of stress fibers distribute in different cell types and how they assemble, is not so well characterised. The focal adhesions originate as protein clusters containing integrin receptors at the leading edge and at tips of filopodia (Galbraith et al., 2002) where they establish contact with the substratum and grow into transient structures referred to as focal complexes (Kaverina et al., 2002). Some of these then mature into focal adhesions, which often, due to the extension of the lamellipodium, locate to the area between the lamellipodium and the lamellae. They then continue to change in character with the movement of the cell, until they are dissociated at the cell rear (more on this below (Beningo et al., 2001; Webb et al., 2004). As actin polymerizes at the leading edge, the filaments forming the cortical actin, flow in a reterograd motion towards the cell center. This process, called rearward flow or simply cortical flow, is the consequence of actin polymerization at the front and myosin II mediated contractions in the interior of the cell (Lin et al., 1997; Ponti et al., 2004). The movement of actin filaments due to rearward flow has been observed to be reduced by focal adhesions (Alexandrova et al., 2008); in one study of PtK1 epithelial cells, filamentous actin was mesured to flow rearwardly by approximately 25 nm/s in the distal lamellipodia and by 2 nm/s in the lamella (Gardel et al., 2008). The rearward actin filament flow must not be confused with the so called treadmilling phenomenon which refers to the “flow” of individual actin molecules from the fast polymerizing (+)-end to the slow growing (-)-end in a filament at steady state or during elongation. Notably, treating cells with the myosin II inhibitory drug blebbistatin, the force generated by expanding lamellipodia was actually seen to be unchanged as was filament elongation mediated protrusion, hence protrusion of the lammelipodium is not dependent on myosin II-based contraction (Gardel et al., 2008; Kaverina et al., 2002).

12

Intracellular trafficking Actin participates in most intracellular transport processes (fig. 2). The roles that have been proposed for actin in membrane trafficking include pushing of membrane invaginations, propelling vesicles, creating platforms for protein-protein interactions and regulating membrane plasticity. In this thesis vesicle trafficking is of particular interest (paper III), therefore this phenomenon is briefly dealt with here. A large body of cellular uptake occurs via clathrin mediated endocytosis, where clathrin coated vesicles are formed and pinched off at the cytosolic side of the plasma membrane. Actin polymerization is required for this and it occurs already at the initial stages of membrane invagination, promoting this process (Merrifield et al., 2002) and contributing also to constriction of the neck and scission by providing the tension required for dynamin dependent scission. Furthermore it also supports lateral mobility of the vesicle (Merrifield et al., 2002; Merrifield et al., 2005; Yarar et al., 2002). The circular dorsal ruffles mentioned above have been characterized as sites exhibiting substantial endocytosis (Buccione et al., 2004). Hence these structures with their massive actin dynamics couple actin remodeling and receptor internalization. As with clathrin mediated endocytosis, actin remodeling is also involved in other types of endocytosis. However, the clathrin mediated endocytosis is considered as the principal mechanism of recycling of the synaptic vesicles, which is interesting in relation to the discussion in paper III. The reverse process, exocytosis, mediates secretion and cell surface expansion. Regulated exocytosis occurs rapidly in response to stimulation leading to fusion of a subpopulation of vesicles already primed for release. Filamentous actin is rapidly assembled around the fusing vesicle (Sokac et al., 2003). This is triggered by compartment mixing of the two fusing membranes which recruits signaling molecules such as PKCβ and Rho GTPases (Yu and Bement, 2007). A specific targeting of the actin polymerization to the surface of the exocytosing vesicle is required since a more general increase in actin polymerization would otherwise inhibit exocytosis. The octameric exocyst complex, which tethers secretory vesicles to specific sites at the plasma membrane was seen to bind the Arp2/3 complex, a promoter for actin polymerization, indicating an early positioning of components required for polymerization (Zuo et al., 2006).

13

Fig 2. Schematic view of important sites for actin dynamics. 1, represents the organized network of filaments giving rise to the lamellipodium. 2, the long, bundled filaments of the filopodium. 3, represents the stress fibers which are built up by thick bundles of anti-parallel actin filaments engaged in myosin contraction and terminating in focal adhesions or associating with other stress fibers. 4, endocytosis, endosome trafficking and vesicle rocketing, 5, actin reorganization during exocytosis and 6, ER to Golgi transport. Red represents actin filaments and filament bundles.

Actin structure

Actin isoforms Actin is a 42 kDa ATPase present in all eukaryotic cells. It can polymerize to form the helical filaments that form the core of the microfilament system. The actin sequence is strongly conserved, yet the protein occurs in several different isoforms. These are classified as α, β or γ depending on their isoelectric point. In mammals there are six actin genes encoding six actin isoforms. α-Actin (the most acidic) is specific for muscle cells and β- and γ-actin are non-muscle isoforms. In muscle cells three variants of α-actin is expressed; α-cardiac, α-skeletal and α- smooth, and there is also a γ–smooth muscle actin. The amino acid sequences between the isoforms are virtually identical, particularly for mammalian expressed β- and γ- actin which differ only in four out of their 374 residues, and as with most actin variants the amino acid variations are clustered to the N-terminal sequence. The functional significance of the two non-muscle isoforms is not well understood, but since they are conserved between higher eukaryotic species they must reflect one or more important differences (Rubenstein, 1990). Maybe it relates to subtle variations in the binding of control proteins, in this respect it is interesting that γ- actin has a uniform distribution in cells (Otey et al., 1986) while β-actin is enriched in the lamellipodium. N-

14 terminal processing involving arginylation has been proposed as important for this localization (Karakozova et al., 2006). Also the β-actin distribution is probably at least in part dependent on localized translation of the protein (see further in a later section) which requires a so called zip-code sequence in the 3’-UTR of the mRNA and which is absent in γ-actin mRNA. Recent observations are to some extent challenging the view on how the non-muscle isoforms localize making their coexpression completely unclear (Dugina et al., 2009). The actin isoforms are interchangeable to varying degrees also adding to their enigmatic functional difference. Transgenic expression of α-cardiac actin fully rescued for the lack of α-skeletal actin in mouse (Laing et al., 2009). Additionally, mice lacking γ-actin are viable and although smaller in size and with a reduced life expectancy because of delayed development, they are otherwise appearing normal. Cells cultured from these animals migrate normally (Belyantseva et al., 2009; Bunnell and Ervasti). These examples suggest that there is certain functional redundancy between the actin isoforms. However, compensatory upregulation of the expression of other isoforms have been observed when expression of one isoform is abolished and this is likely the reason why the γ-actin knockout mouse is viable. In Drosophila, replacing a muscle specific actin with a non-muscle like isoform in the indirect flight muscle of the fly led to disturbed flight muscle structure and function (Fyrberg et al., 1998). Nor was γ-actin able to rescue for lethality in a mouse lacking α-skeletal actin (Laing et al., 2009). This points to a conserved functional variability between the different isoactins that is more pronounced between muscle and non-muscle actin than within these groups of isoactins. However, it is also clear that actin is amazingly conserved not only from sequence and structure point of view but also in its cellular function as illustrated by the fact that replacement of the endogenous actin gene in S.cerevisiae with an avian β-actin gene (encoding a β-actin identical to the mammalian isoform) supported viability albeit it caused an altered morphology and reduced growth rate of the yeast cells (Karlsson et al., 1991).

Actin protein structure The structure of the actin molecule has been determined from the crystal structure of actin together with different binding partners. Due to the property of actin to polymerize under the conditions used for crystallization it has not been possible to determine its structure without

15 the presence of a binding partner or direct modifications that interferes with its polymerization (Chik et al., 1996; Kabsch et al., 1990; Otterbein et al., 2001; Schutt et al., 1993). The actin molecule is built by two domains each of which in turn consists of two sub-domains; however none of its sub-domains can fold independently of the other and thus all parts are required to form a functional protein. The N- and C-terminus are found in the same subdomain. The molecule is stabilized by a Me2+- nucleotide complex (Mg/Ca-ATP/ADP), which binds in a cleft that nearly separates the molecule into two halves with two sub-domains contacting the nucleotide on each side, and with only a connecting bridge formed by the protein chain ‘underneath’ the nucleotide between sub-domains I and III (fig. 3).

Fig 3. Representation of the profilin:β-actin complex displayed as (A) a space fill and (B) a ribbon model. The actin polypeptide chain (red) starts in subdomain 1 and traverses the subdomains in the order of 2, 1, 3, 4, 3 and back into 1, the N- and C- termini both reside in subdomain 1. At the central cleft the nucleotide and divalent cation are visible. Profilin (blue) is built up by a seven strand β-sheet surrounded by four α -helices. The N-and C-termini form two of the α-helices and positioned at one side of the β–sheet they form the poly-L-proline binding site. The images were rendered based on the crystal structure of the closed state of the profilin:β-actin complex (PDB file 2BTF; (Schutt et al., 1993)) using MolSoft.

Consequently, the monomeric form of actin, which often is referred to as globular actin or G-actin, is an asymmetric molecule, and this asymmetry is also reflected by the molecular organization of the actin filament. Hence the filament by itself expresses directionality, an important aspect relevant for practically all actin filament functions, i.e. polymerization, interaction with other proteins and, not least, acto- myosin based force generation. All together this means that, regulation of actin polymerization is achieved both by ionic conditions and the nucleotide bound status of the molecule as well as by a number of

16 regulatory proteins. One important aspect is that to fold to its native state actin requires the eukaryotic chaperonin CCT (Sternlicht et al., 1993), which at present makes it impossible to express this protein recombinantly in bacteria.

Actin polymerization At low ionic strength actin remains unpolymerized, however at physiological salt concentrations polymerization occurs spontaneously in a process, which is favored if the actin molecule is in its ATP-bound state (Korn, 1982). The increase in ionic strength upon addition of salt most likely in combination with changes in the conformation of the actin monomers make them competent to polymerize. The polymerization reaction when initiated by salt addition to a solution of unpolymerized actin is distinguished by three phases; a lag phase referred to as the nucleation phase, followed by a phase of rapid filament elongation which finally reaches the so called steady state level where no more net filament formation occurs. The initial step of the assembly process requires that individual actin molecules come together, forming dimers, which are highly unstable, but if the protein concentration is high enough, more stable trimeric ‘nuclei’ may form and serve as starting points from which elongation rapidly proceeds. In vitro elongation of a double-stranded filament (F-actin) occurs by growth at both ends. However, the Kd for subunit addition at the two ends differs and in polymerizing solutions of actin, steady state is soon reached where growth at the (+)-end of the filament is balanced by net dissociation of subunits from the (-)-end. The rate of elongation is 10-20 times faster at the (+)-end. The concentration of G-actin at steady-state is defined as the critical concentration for actin polymerization (Acc) and is 0.1 µM under physiological salt conditions. In conjunction with polymerization, the actin ATPase is stimulated by the salt and hydrolysis occurs hand-in-hand with subunit incorporation at the (+)-end. As a result the actin molecules that dissociate from the (-)-end are ADP-bound and need to exchange the nucleotide for ATP before they can incorporate again at the (+)-end; due to this phenomenon of treadmilling (Neuhaus et al., 1983), F-actin solutions gradually consumes ATP (Le Clainche and Carlier, 2008; Schuler, 2006). So in fact, the actin filament is an asymmetric both from structural and biochemical point of views. Notably ATP hydrolysis is not a requirement for polymerization since polymerization still occurs in the presence of a non-hydrolysable ATP analogue (Cooke and Murdoch, 1973). However, for establishment of the balanced (+)-end polymerization and (-)-end depolymerization that characterizes

17 filamentous actin under treadmilling at steady-state (Carlier et al., 1986), which is believed to reflect the situation of actin in vivo in lamellipodia and filopodia (Lai et al., 2008), nucleotide hydrolysis is necessary. In addition to the polymerization kinetics, the change in nucleotide state in actin subunits along the filament is likely to influence subunit structure and therefore may be important for certain filament binding proteins (Blanchoin and Pollard, 1998), such as cofilin, Arp2/3, VASP and spire (see further below). In the cell, the concentration of unpolymerized actin is far higher than the critical concentration, reaching in some cases as high as 50 – 100 µM or even more (Carlsson et al., 1977). This is possible through binding of actin to proteins such as thymosin β4 and profilin which interferes with the polymerization process and enable the presence of monomeric actin at concentrations above Acc by preventing spontaneous nucleation and elongation as described for the test-tube situation above. Actin molecules are recruited from such monomer-bound complexes under the control of different proteins or protein complexes often referred to as nucleation and elongation promoting factors (Goley and Welch, 2006). Well studied examples are VASP, WASP-Arp2/3 and the formins which function through different mechanisms. Additionally, other proteins are readily activated to cause filament disassembly, thereby controlling the (-)-end dynamics of filament turnover. As will be dealt with below these actin regulatory proteins are under the control of several different components such as the Rho GTPases and certain phospholipids. The proteins regulating actin assembly increases actin turn-over so that it is approximately 100 times faster in cells compared to the treadmilling observed in vitro (Zigmond, 1993). Apart from being regulated by proteins and lipids, so called Ca2+-spikes (Berridge, 2006) could also have a direct influence actin dynamics, since Mg2+- and Ca2+-actin have different properties (Frieden and Patane, 1988). This type of highly localized increases in Ca2+ concentration have been shown as a crucial component in chemoattractant directed migration where the calcium is both taken up via channels in the plasma membrane and released from ER in response to IP3, the hydrolysis product of PIP2 (Wei et al., 2009).

The actin filament The actin filament is a helical polymer that can either be thought of as a left handed one start helix (short pitch) or a right handed double helix

18

(long pitch). It has still not been possible to fully determine the structure of the actin filament. A model based on the diffraction pattern of ordered gels of F-actin and the structure of monomeric actin in complex with DNase I (Kabsch et al., 1990) was presented more than 20 years ago (Holmes et al., 1990) but despite considerable efforts and several refinements (Oda et al., 2009) the orientation of the actin subunits cannot be fully established due to the low resolution of the original diffraction pattern used for the modeling. Recently improved methodology in the form of cryo EM may prove to be a fruitful approach to tackle this problem. In such a study aspects concerning interstrand connections in the original filament model by Holmes et al were challenged (Fujii et al. 2010) but a final solution to the enigmatic actin filament subunit orientation still remains to be seen. However, a structure of a polymeric form of actin has been presented based on the molecular organization in crystals of the profilin-actin complex (profilin:actin). In this structure the molecules are organized with actin- actin contacts in a ribbon arrangement, which shares many characteristics of ordinary actin and the orientation of the subunits are essentially opposite to the Holmes model (Schutt et al., 1993; Schutt et al., 1995). A ribbon-based model of the filament has not yet been published.

Profilin

Profilin isoforms are ubiquitously expressed in mammalian tissue and in fact are present in virtually all eukaryotic cells; even viruses of Vaccinia subspecies express a profilin. All profilins have a similar structure characterized by a β-sheet center formed by seven β-stands surrounded by four α-helices. The N-and C-terminal amino acid stretches form two of these α-helices which are positioned adjacent to each other on one side of the β-sheet (Cedergren-Zeppezauer et al., 1994; Schutt et al., 1993). In general, profilins interact with actin and with at least one actin related protein (Arp), the Arp2-subunit of the Arp2/3-complex, as well as with many proteins containing poly-L-proline motifs (Mahoney et al., 1997; Tanaka and Shibata, 1985) and with phosphatidylinositol lipids (Lassing and Lindberg, 1985; Lu et al., 1996; Witke, 2004). However, some profilins are at variance with this, for instance the Vaccinia virus profilin which shows poor affinity for poly-L-proline. In mammals four profilin coding genes have been characterized. These express profilin 1, 2a, 2b, 3 and 4.

19

Profilin 1 is the ubiquitous and archetype isoform, generally present though at lower levels in brain, heart, skeletal muscle and pancreas (Buss and Jockusch, 1989). Of the two profilin 2 splice variants, profilin 2a is expressed at high levels only in the brain but is found to some extent in most tissues (Honore et al., 1993; Witke et al., 2001) while the least studied of the two, 2b, is expressed in kidney and is unique among the profilins in the sense that it interacts poorly with actin but has been found to associate with tubulin (Di Nardo et al., 2000). Throughout this thesis when referring to profilin 2 it is the 2a-isoform that is discussed. Profilin 1 and 2 actually seem to have somewhat complementary expression, in tissue where profilin 2 is high, profilin 1 is lower and vice versa (Honore et al., 1993). Profilin 3 is restricted to kidney and testis and profilin 4 to testis (Hu et al., 2001; Obermann et al., 2005). Notably the majority of profilin studies have been made on profilin 1.

Profilin activities Profilin was first identified as a binding partner to actin in bovine spleen extracts and was in these early experiments recognized as an inhibitor of actin polymerization (Carlsson et al., 1977), hence its name, which is derived from is role to ’keep actin in pro-filamentous form’. It forms a 1:1 complex with G-actin and in a number of ways it regulates actin- coupled functions, both by having an effect on the actin molecule itself and by working as a recruitment factor in conjunction with other components. When binding actin, profilin stabilizes the molecule in an “open” conformation (Chik et al., 1996), facilitating nucleotide exchange and thus a rapid replacement of ADP for ATP (reviewed in (Karlsson and Lindberg, 2007). Profilin also inhibits the ATPase activity of actin (Goldschmidt-Clermont et al., 1992; Nyman et al., 2002; Tobacman and Korn, 1982). As already mentioned, during actin filament elongation ATP-containing actin monomers are added at the fast polymerizing (+)- end of the filament and ADP-containing monomers dissociate from the (-)-end. Profilin binds to the end of the actin monomer corresponding to the (+)-end (fig. 3), therefore it only blocks polymerization at the (-)- end (Korenbaum et al., 1998; Nyman et al., 2002; Pantaloni and Carlier, 1993; Pollard and Cooper, 1984; Schutt et al., 1993). Hence it is only under conditions when the (+)-end is blocked that profilin is a true inhibitor of actin polymerization. The fact that profilin also prevents self-nucleation of actin, in a situation where no filaments are present or the filament (+)-end is blocked (capped) by interaction with other components, also makes profilin an inhibitor of actin polymerization. It was proposed already in the beginning of the 1980’s that profilin delivers

20 the actin monomer to the growing filament (Pollard and Cooper, 1984) but despite a large number of studies favoring this view e.g. (Kang et al., 1999; Korenbaum et al., 1998; Pantaloni and Carlier, 1993) it was not until the generation of a covalently coupled profilin-actin complex this was demonstrated more directly (Grenklo et al., 2003; Hajkova et al., 2000). However, so far it has not been possible to capture and directly visualize profilin associated with filamentous actin by electron microscopy, probably reflecting the transient nature of the interaction. The delivery of actin to the growing filament end by profilin-actin and the subsequent incorporation of the actin into the filament must involve a complex set of processes, e.g. docking of the complex, dissociation of profilin, ATP hydrolysis and full incorporation of the actin molecule into the filament structure, whose temporal and biochemical integration currently are poorly understood. In the context of the cell, with numerous nucleation promoting factors as well as profilin binding and polymerization promoting proteins, profilin, via its influence on the actin monomer and its interactions with other proteins, clearly is important in controlling actin polymerization locally and possibly may have a stimulatory role in directed actin polymerization (see also paper II).

Profilin and phosphatidyl inositols Based on observations in vitro, interaction with PIP2 presumably provides an important control mechanism for profilin function. The PIP2 binding by profilin was first reported by (Lassing and Lindberg, 1985) who demonstrated that binding of the lipid caused dissociation of profilin:actin. Again a large number of studies from different laboratories employing a variety of techniques have contributed to map this interaction and have led to the insight that PIP2 binds profilin at two separate regions of the molecule, one present in the actin-binding surface and one being located to the N- and C-terminal helices and thus partly overlapping with the poly-L-proline binding site (Lambrechts et al., 2002; Skare and Karlsson, 2002; Witke, 2004). Studies with the covalently linked profilin-actin demonstrated that formation of a trimeric complex of PIP2-profilin-actin is possible, and led to the proposal that the lipid- induced dissociation of the complex occurs step-wise where an initial association to the N- and C-terminal helices propagates an alteration of the actin-binding surface and opens up for the second lipid interaction with release of actin as the consequence (Skare and Karlsson, 2002). Profilin 1 interacts in vitro with PI(4)P, PI(4,5)P2, PI(3,4,5)P3 and PI(3,4)P2 at increasing affinities (Lassing and Lindberg, 1985; Lu et al.,

21

1996). These lipids are localized to the inner leaflet of the plasma membrane where PI(4,5)P2 has been shown to localize to lipid rafts (Golub et al., 2004). It has been reported that PIP2 is important for profilins localization to the plasma membrane (Ostrander et al., 1995) (Bubb et al., 1998) however, although this concept has not been questioned, factual evidence supporting this conclusion are weak. Variants of PIP2 and PIP3 are formed from PI by the activities of PI3-, PI4- and PI5-kinases e.g. (Cantley, 2002; Kwiatkowska, 2010). These kinases respond rapidly to extracellular signals, in platelets for instance, changes in PI(4,5)P2 levels have been demonstrated within 10 s after thrombin stimulation (Hartwig et al., 1995; Lassing and Lindberg, 1990). This is in the same time range as the initiation of actin polymerization (Karlsson and Lindberg, 1985; Lassing and Lindberg, 1988; Lindberg et al., 2008). The ability of PI(4,5)P2 and PI(3,4,5)P3 to dissociate the profilin:actin complex in vitro (Lassing and Lindberg, 1985; Lu et al., 1996), could reflect a mechanism for release of profilin in association with growing filaments (Cedergren-Zeppezauer et al., 1994; Skare and Karlsson, 2002). Another possibility is that in vivo PI(4,5)P2 does not dissociate profilin:actin but instead clusters a somewhat destabilized profilin:actin at the plasma membrane close to sites of polymerization where it is recruited to the filament by polymer forming machineries. Yet another possibility is that PI(4,5)P2 sequesters profilin at the inner leaflet of the plasma membrane, preventing it from binding to actin. Clearly the PI(4,5)P2 interaction adds strongly to the view that profilin has role in relaying information from activated surface receptors to the actin microfilament system that is already implicated from studies of its interaction with proteins such as VASP and formins. This is further supported by the fact that profilin can influence phosphatidyl inositol turnover, since it constrains the hydrolysis of PI(4,5)P2 by phospholipase Cγ (Goldschmidt-Clermont et al., 1991; Goldschmidt-Clermont et al., 1990) and is able the increase the formation of PIP3 from PI(4,5)P2 by interacting with the regulatory subunit p85 of phosphatidylinositol 3- kinase (Singh et al., 1996a). It is unknown if the latter effect due to an increase of local PI(4,5)P2 concentration by profilin or if profilin presents the substrate to the catalytic subunit p110 in an orientation favoring the process.

Poly-L-proline binding There are numerous ligands for the poly–L-proline site of profilin, these proteins contain a proline-rich sequence canonically referred to as GP5

22 because of the glycine-pentaproline sequence observed in VASP the first profilin-binding protein identified after actin and demonstrated to interact with profilin via this motif (Reinhard et al., 1995). The poly-L- proline binding surface is formed by residues in both the α-helices building the N-and C-terminus of the molecule. This fact has seriously hampered production of functional fusion constructs of profilin with different molecules because of the interference of the fusion peptide with the folding of the helices and/or the binding of profilin to its partner at this site. Many of the proteins interacting with the poly-L-proline binding surface are also actin binding proteins and as discussed in other sections in this text are involved in the recruitment and possibly also the docking of profilin-actin to the growing end of the filament. For instance, Ena/VASP, and the formins bind to profilin and profilin:actin via the poly-L-proline binding site while at the same time they are associated to the filament (+)-end (fig. 5)(Ferron et al., 2007; Huttelmaier et al., 1999; Michelot et al., 2005) Profilin also binds to the Arp2/3 complex but this is via the actin binding site (Machesky et al., 1994).

Profilin phosphorylation Phosphorylation of profilin has been observed in in vitro studies (Hansson et al., 1988) and in the case of profilin from the plant Phaseolus vulgaris also in vivo (Guillen et al., 1999). Also activation of platelets was shown to lead to phosphorylation; PKCzeta has been demonstrated to be one kinase that recognizes profilin as a substrate, causing phosphorylation of the C-terminal residue serine 137, and the pp60c-src tyrosine kinase has been demonstrated to target tyrosine 139 (De Corte et al., 1997). The phosphorylation is stimulated by PIP2 (Hansson et al., 1988; Singh et al., 1996b) probably since PIP2-binding induces an exposure of the serine 137. Phosphorylation in activated platelets was seen to be caused by the PI3-kinase and this phosphorylation was reported to increase the affinity for actin and poly-L-proline partners while the affinity for PIP2 remained unchanged (Sathish et al., 2004). On the other hand incubation of profilin with growth factor receptor complexes immunoprecipitated from extracts of growth factor stimulated cells, led to mutually exclusive phosphorylation on serine 137 and tyrosine 139, in which case the protein did not bind poly-L-proline (Bjorkegren-Sjogren et al., 1997). The discrepancy between these two studies is unclear so far, and the possible function of phosphorylation as

23 a regulatory mechanism for profilin has not been explored to any large extent. The organization of the molecules in the profilin:actin crystal shows that actin can contact profilin at two separate sites (Schutt et al., 1993). The major contact is, as mentioned above, formed by interaction with sub- domains 1 and 3 at the (+)-end of the actin molecule, while a minor interaction is present in the crystal between actin subdomain 4 and the N-terminal helix on profilin. The latter interaction, which involves the acetylated N-terminal alanine has been proposed to function in a switch- mechanism which could release profilin from the growing filament end after docking of profilin:actin (Cedergren-Zeppezauer et al., 1994). The relevance of this hypothesis remains to be seen.

Profilin in the physiological context Studies of the conversion of monomeric actin in the form of profilin:actin into filaments after thrombin stimulation of platelets (Markey et al., 1981) led to insights that gradually developed into the concept that profilin:actin complex is the main precursor for actin filament formation in vivo (Lindberg et al., 2008). Important support for this conjecture comes from observations that microinjection of the non- dissociable, covalently coupled profilin:actin (PxA) discussed above completely incapacitated cells to respond to growth factor stimulation and surface spreading by the normal actin remodeling. Instead the injection caused a complete derangement of the microfilament system, with retracting edges and loss of stress fibers (Hajkova et al., 2000). Furthermore, PxA also efficiently interfered with the motility of the intracellular pathogen Listeria monocytogenes (Grenklo et al., 2003). Profilin is enriched at the plasma membrane, in lamellipodia and ruffles, regions with high actin dynamics (Buss et al., 1992; Karlsson and Lindberg, 2007; Li et al., 2008) and it has also been observed in association to intracellular membranes involved in intravesicular transport (Dong et al., 2000). Profilin 2 was seen as a part of the endocytic machinery in mouse hippocampal neurons (Gareus et al., 2006) where profilin 2 and dynamin form a complex, which competes for other binding partners and may therefore down-regulate synaptic vesicle endocytosis (Gareus et al., 2006) as well as neurotransmitter exocytosis (Pilo Boyl et al., 2007). The physiological relevance of profilin has been shown in a number of studies, an obvious indication of its significance is demonstrated by the knockout mouse which dies already at the two-cell stage, most likely as a

24 consequence of the cells having stopped dividing because cytokinesis is impaired (Ezezika et al., 2009) after the maternal profilin reaches too low concentrations (Witke et al., 2001). At the cellular level, the effect of profilin 1 expression seems to be dependent on the cell system used. The first organism studied for the effect of profilin depletion was Dictyostelium amobeae where a gene deletion of Profilin 1 and 2 caused impaired movement, and an increase in size (Haugwitz et al., 1994). In a human breast cancer cell line, depletion of profilin using siRNA caused a decrease in focal adhesions and an increased migration rate (Bae et al., 2009) and in human umbilical vein endothelial cells (HUVECs) profilin 1 depletion led to a reduction in filament formation, focal adhesions and proliferation, however in this cell line migration rate was impaired (Ding et al., 2006). The reason for the different outcomes on migration rate after profilin knockdown may seem surprising but in both the studies above it was concluded that the cell edge protrudes slower when profilin levels are lowered. In the breast cancer cells the protrusions were more stable, which led to faster migration, this effect was dependent on VASP, which in these cells was localized to the leading edge at four times higher levels than in the control cells(Bae et al., 2009). The HUVECs ability to form immature vascular like structures was also greatly impaired after reduction of profilin levels (Ding et al., 2006), profilin has previously been seen to be up-regulated during angiogenesis (Salazar 1999). In a later study Ding et al saw that both the poly-L-proline and the actin binding activities expressed by profilin were important for its proper functionality in HUVECs (Ding et al., 2009) and this is most likely always the case in the physiological context. When profilin 1 was overexpressed in mouse endothelial cells, an increase in adhesion formation and in stress fiber bundles in the cell periphery was seen. Also a small increase in filamentous actin was seen (Moldovan et al., 1997) an effect that was observed to an even larger extent in CHO (Chinese hamster ovary) cells (Finkel et al., 1994). Profilin 1 expression is often modified in cancer cells. Downregulation has been observed in several adenocarcinomas as for instance in aggressive mammary and pancreatic carcinoma cells (Gronborg et al., 2006; Janke et al., 2000), and increasing the profilin expression in the mammary carcinoma actually reduced tumor growth (Janke et al., 2000). One reason to this might be that profilin seems to have a stabilizing role on the CDK inhibitor p27, which causes growth arrest (Zou et al., 2010). It is still surprising that profilin is downregulated in cancer given its role in proliferation and as a component being involved in actin filament formation and thus would be expected to have stimulatory effect on migration, and also since

25 profilin seems to be involved in angiogenesis which is required for formation of solid tumors. However, this is likely to reflect that profilin plays an important fine-tuning role in the complex set of interactions pathways that control cell migration (see also paper II).

Profilin in the nucleus Nuclear profilin has been reported for different cell types (Giesemann et al., 1999; Mayboroda et al., 1997; Skare et al., 2003) and profilin 2 has been shown to accumulate in the nucleus of hippocampal neurons in an activity dependent manner (Birbach et al., 2006). Although the role of profilin in the nucleus is unclear, it is implicated in transcriptional regulation (Lederer et al., 2005) and in pre-mRNA processing, and it has been found to co-localize with markers for sub-nuclear particles such as Speckles and Cajal bodies (Giesemann et al., 1999; Sharma et al., 2005; Skare et al., 2003). Since actin itself and several other component of the microfilament system also are found in the nucleus such as cofilin, members of the family, WASP and several Arps for instance, it is likely that profilin in the nucleus participates in the regulation of actin dynamics much like in the cytoplasm although the nuclear compartment seem to hold only a minor fraction of the total cellular actin. Despite intensive work in several laboratories over the past five-six years which have generated a large body of data (Blessing et al., 2004; Grummt, 2006; Posern and Treisman, 2006; Visa and Percipalle, 2010) the role of actin and the actin binding proteins in the nucleus largely remains unclear. However, one function for nuclear actin of direct relevance for this thesis is MAL/SRF-controlled transcription where MAL is an actin binding protein that must be released from actin in order to bind SRF and together with this transcription factor activate expression of a large number of the genes that becomes upregulated after serum stimulation, for instance genes involved in actin mediated contractility and motility (Vartiainen et al., 2007). SRF regulated gene expression is also important for neuronal function where it is important during neuronal development, memory and learning (Knoll and Nordheim, 2009). The interaction of MAL with actin competes with the binding of profilin and the consequences for the balance of these two proteins for MAL/SRF- dependent gene regulation is the object of the study in paper II.

26

PIP2 and microfilament dynamics The microfilament system just below the plasma membrane is highly responsive to extracellular signals. This is due to the fact that many actin regulatory proteins are controlled by phosphoinositide (PtdIns) lipids. These lipids constitute only 5-8% of the total cellular lipid content but are essential because of their role in signaling (Cockcroft and De Matteis, 2001). PI(4,5)P2 is a major component in signaling processes, upon receptor activation it is hydrolyzed by phospholipase C (PLC) to diacylglycerol (DAG) and inositoltriphosphate (IP3). This lipid is also the main PtdIns involved in regulation of the microfilament system and it is known to accumulate at the inner leaflet of the plasma membrane at sites of cell surface motility (Higgs and Pollard, 2000; Hilpela et al., 2004; Mullins, 2000; Sechi and Wehland, 2000). PI(4,5)P2 is regenerated from DAG in the PI-cycle in a process taking place in the ER where the synthesizing are located. A constant lipid transfer is therefore cycling between the plasma membrane and ER. Ultimately the inositol ring is phosphorylated and dephosphorylated by specific enzymes residing in the plasma membrane. PI(4,5)P2 is generated by PI(4)P5’- kinase and PI(5)P4’-kinase or metabolized by PLC, 5’-phosphatases and the PI(3,4,5)P3 producing PI3-kinase (Kwiatkowska, 2010). This means that not only PIP2 but also the other intermediates, PI/PIPn, phosphatidic acid and DAG, as well as interconversion between these intermediates by kinases and phosphatases are involved in different signaling processes. PI(4,5)P2 affects microfilament dynamics both by sequestering actin binding proteins like cofilin, and by promoting their activity like in the case of N-WASP (see below) (Yin and Janmey, 2003). Also PIP3, as well a being a major signaling lipid in multiple pathways (Cantley, 2002), is known to be involved in the regulation of actin binding proteins such as WAVE and myosin-X (Oikawa et al., 2004; Plantard et al., 2010). Lipid rafts are proposed to constitute a dynamic signaling platform for PI(4,5)P2, by providing a specialized lipid environment enriched in PI(4,5)P2 and accumulating specific proteins (Simons and Gerl, 2010). Enzymes involved in the PI cycle, like PI5-kinase are recruited to these rafts by Rho GTPases (Rozelle et al., 2000). It has been shown that actin assembly at lipid rafts can be initiated efficiently and then drive membrane and vesicle movement (Taunton et al., 2000). It is not obvious how certain membrane regions become enriched in PI(4,5)P2 since this phospholipid has a large and heavily charged head group. One

27 possibility is that it accumulates in areas with more curvature (Flanagan et al., 1997), for instance at the tip of the lamellipodia, along filopodia, and especially at the tip of the latter. This also means that the membrane curvature-inducing BAR-domain proteins, that will be discussed below, could have a role in accumulating PIP2. Another possible mechanism behind the accumulation of PIP2 is that proteins containing positively charged residues and positioned at the inner plasma membrane leaflet cluster the lipid in large numbers. This correlates with what has been observed for the GAP43-like proteins GAP43, CAP23 and MARCKS, which are all regulated and also function in similar ways. They are hydrophilic and contain a basic domain which binds and clusters PIP2 to form lipid rafts and they have been seen to promote actin assembly (Caroni, 2001; Laux et al., 2000). GAP43 and CAP23 are both involved in neuronal process formation (Laux et al., 2000; Skene, 1989). MARCKS in addition to its function in neuronal process formation is also expressed in non-neuronal cells where it has a role in migration and adhesion (Myat et al., 1997). The fact that positively charged amino acids can promote actin assembly because of their effect on PIP2 is dealt with in more detail in paper III in this thesis. The PIP2 enriched rafts which are also enriched in cholesterol, cluster upon receptor signaling in a process called patching. The initial rapid patch assembly involves PIP2 together with the Rho GTPase Cdc42 which promotes this assembly through N-WASP (Golub and Caroni, 2005). These patches then cluster to larger platforms in a microtubule dependent process which in turn confine and direct actin dependent protrusions. Depleting the cholesterol with cyclodextrin disrupts these raft patches and leads to loss of lamellipodia and membrane ruffling. It was also observed that the formation of these rafts precedes lamellipodia motility. IQGAP1, a protein promoting actin polymerization (Brandt and Grosse, 2007) is targeted to the rafts and recruits microtubules. Which at their tip bring the plus-end tip tracking protein APC, which recently was proposed to be an actin nucleator (Okada et al., 2010). Thus microtubule dependent raft patch clustering is required for sustained cell surface motility and protrusion. The sustained accumulation of PIP2-rich rafts into patches also was seen to involve Cdc42, N-WASP and actin, hence actin polymerization is also involved in this process. However, the exact mechanism leading to polymerization of actin upon patching is still not known.

28

The plasma membrane has been shown to be physically connected to the cytoskeleton via PIP2. By using optical tweezers it was seen that the expression of a pleckstrin homology domain that specifically binds to and sequesters PIP2 or by decreasing PIP2 using a specific phosphatase caused a reduced adhesion between the plasma membrane and the cytoskeleton (Raucher et al., 2000) and so did depolymerization of filamentous actin. Interestingly, the downregulation of PIP2 likewise led to a decrease in filamentous actin according to phalloidin staining, however the mechanism behind was not investigated further.

Rho GTPases The Rho (Ras homologous) GTPases are as is indicated by their names a subclass of the Ras superfamily of GTPases. In mammals there are around 20 Rho GTPases, most of which are key regulators of virtually all actin dependent processes. The coordinated balance of the activities of these GTPases varies between cell types and has proven to be important for the behavior of the cell (Lozano et al., 2003; Rottner et al., 1999b; Sarmiento et al., 2008). The Rho GTPases are activated by GEF’s (Rho- guanine nucleotide exchange factors) which increases the exchange rate of the GTPase-bound nucleotide GDP in the inactivated state for GTP, and are inactivated by GAP’s (GTPase activating protein) which potentiates hydrolysis of GTP to GDP by the GTPase. Most of the Rho GTPases are active in association to membranes, either intracellular membranes or the plasma membrane. For this reason they are commonly post-translationally modified by prenylations or palmitoylations at their C-termini to enhance their membrane interaction and to define the membrane compartment for their localization (Wennerberg and Der, 2004). Rho guanine nucleotide dissociating inhibitors (GDIs) bind many Rho GTPases at the C-terminus preventing nucleotide exchange and mask the prenyl group, thereby impeding membrane interaction (DerMardirossian and Bokoch, 2005). In this also lies the ability of GDIs to target the GTPases to their right membrane location (Lin et al., 2003). Active Rho GTPases interact with many proteins involved in actin regulation, examples are protein kinases, lipid modifying enzymes and activators of actin polymerization. The response depends on which Rho GTPases that are activated. There are three well studied classes of Rho GTPases involved in microfilament system regulation. Simplified it can be said that RhoA activation leads to stress fiber formation through

29 activation of formins and myosin-II, the latter by activating ROCK (Rho associated kinase)(Ridley et al., 2003) and Rac and Cdc42 induces lamellipodia and filopodia, respectively, by activation of Arp2/3- dependent nucleation factors (Kozma et al., 1995; Ridley and Hall, 1992). However, there are overlapping functions between the different RhoGTPases. Cdc42 is not the only Rho GTPase whose activity leads to filopodia formation, also Rif, RhoD and Wrch1 are known to induce these type of processes (Aspenstrom et al., 2004; Ellis and Mellor, 2000). Furthermore, also Rho GTPases like Rho A, B, D and Cdc42 are important in different steps in vesicle trafficking like exocytosis or between intracellular compartments. Also in this latter context the abilities of these RhoGTPases to induce actin polymerization are important (Bader et al., 2004; Egea et al., 2006). Cdc42 also directs cell polarity by orienting the MTOC and Golgi apparatus in front of the nucleus (Etienne-Manneville, 2004; Raftopoulou and Hall, 2004; Ridley et al., 2003) and by stabilizing the microtubules at the cell front (Pegtel et al., 2007). The Rho GTPases also have a role in phosphatidyl inositol signaling by interacting with the enzymes involved in PI/PIPn conversion. For instance Rho and Rac activate PI(4)P5-kinase that produces PI(4,5)P2. When the cell is exposed to a gradient of a chemoattractant, local activation of the PI3-kinase leads to increased amounts of the phospholipids PI(3,4,5)P3 and PI(3,4)P2 at this location. Cdc42, which is active towards the front of a migrating cell, appears to be involved in determining where PI3-kinase and PTEN, which is a phosphatase of PI(3,4,5)P3, are active (Ridley et al., 2003). Conversely these lipids affect the activity of RhoGTPases, such as in the case of Rac which via Rac GEF’s is activated by PI3-kinase products. Rac activates WAVE, meaning that the presence of these phospholipids leads to lamellipodia formation. Since Cdc42 affects the production of PI(3,4,5)P3, Rac activation is thereby coupled to the activity of Cdc42. In a study of exocytosis in neuroendocrine cells, RhoA was seen to bind to vesicles and Cdc42 and Rac were located to the plasma membrane (Gasman et al., 1999). Interestingly both RhoA and Cdc42 induce actin polymerization but RhoA suppresses exocytosis whereas Cdc42 has an inducing role (Momboisse et al., 2011). RhoA could stabilize an actin zone surrounding the secretory vesicles by regulating the PI(4)P5-kinase producing PIP2, while Cdc42 increases release efficiency by Arp2/3-N- WASP dependent actin polymerization at the interface of the secretory

30 membrane and the plasma membrane (Gasman et al., 1998; Gasman et al., 2004; Rohatgi et al., 1999). Much of the knowledge collected on Rho GTPases comes from cell- studies where dominant negative and positive constructs were overexpressed. This type of constructs have provided valuable insights but may also been misleading in certain aspects since studies of genetically knocked out GTPases have not always been in congruence with previous results (Heasman and Ridley, 2008). Knocking out Cdc42 in mouse is embryonic lethal, therefore conditional knockouts on specific tissues have been made. The result correlates with previous data regarding an involvement of Cdc42 in filopodia formation, since several of the cell types extracted from these animals had a reduced number of filopodia including embryonic stem cells (Chen et al., 2000), neurons (Garvalov et al., 2007) and embryonic fibroblasts (Yang et al., 2006). Fibroblastoid cells derived from embryonic stem cells however did not show a reduction in filopodia, indicating a variation between cell types and that other GTPases could have a more prominent role in some cells (Czuchra et al., 2005). It was also concluded that Cdc42 is required for axon generation (Garvalov et al., 2007). Abolishing expression of the ubiquitously expressed Rac1 also requires a conditional knockout to generate a viable animal. Since there are three isoforms, Rac 1-3, this complicates the matter on evaluating Rac function. It seems however that cells from the Rac1 depleted animals migrate at normal speed with membrane ruffles, which is in contrast to the studies on dominant negative Rac where migration was impaired. However, the cells showed impaired cell spreading and chemotaxis. The latter is reflected in defects in the development of the nervous system where axonal guidance is impaired, also depletion of Rac1 using RNAi in Rac3 null hippocampal neurons inhibits formation of dendrites while axons develop normally (Gualdoni et al., 2007; Heasman and Ridley, 2008). No knockout of RhoA has been reported but it was seen that silencing of RhoA in PC12 cells reduced the levels of filamentous actin and promoted neuritogenesis (Fan et al., 2008). From these studies it can be concluded that the Rho proteins have an important role in neuronal development and synaptic plasticity by controlling neurite outgrowth and axonal guidance (Hall and Lalli, 2010).

31

Actin regulatory proteins To overcome the kinetically unfavorable step of actin nucleation several different actin nucleating machineries have evolved. The Arp2/3- dependent WASP and WAVE proteins, and the formins are maybe the most well characterized so far but the relatively recently discovered tandem monomer-binding nucleators have indicated that the mechanisms for initiating actin polymerization de novo in the cell are very diverse (Firat-Karalar and Welch, 2011).

Arp2/3 The heptameric Arp2/3 complex takes a special function in actin nucleation since its two actin related proteins Arp2 and Arp3 together exposes a surface similar to the (+)-end of the actin filament and thereby serves as a starting point for the growing filament in co-operation with its different activating binding partners, which spatiotemporally control the process (fig. 5A). Therefore the Arp2/3 complex is a (-)-end filament capper that is generally recognized as one of the major nucleators of actin polymerization in the cell. It was first purified form Acanthamoeba castellanii by profilin affinity chromatography by virtue of the Arp2- profilin interaction (Machesky and Hall, 1996), and it is highly localized to regions of lamellipodial protrusion in both A. castellanii and mammalian cells (Li et al., 2008; Machesky et al., 1994; Machesky and Hall, 1997) but it is absent from filopodia (Svitkina et al., 2003; Symons et al., 1996). In vitro, the Arp2/3 complex induces formation of branched actin filaments (Welch and Mullins, 2002), which as already mentioned in the introduction, have caused a lot of attention. After the observation of such a branched arrangement by electron microscopy of the leading edge of migrating cells (Svitkina and Borisy, 1999), this led to the dendritic model of actin polymerization for lamellipodial advancement (Pollard and Borisy, 2003). Recent and ongoing studies using imaging correlated electron microscopy (EM) and EM-tomography strongly suggests that the lamellipodium at the cell front does not contain branched filament arrangements and that these, most likely were artifacts caused by the conditions used for sample preparation (Koestler et al., 2008; Lai et al., 2008; Small, 2010). Interestingly in this context is that the ability of Arp2/3 to cause formation of branched filaments in vitro is reported to be enhanced by phalloidin (Mahaffy and Pollard, 2008), a toxin which specifically binds filamentous actin and commonly is used, labelled with

32 a fluorophore, to visualize actin filament arrangements by fluorescence microscopy (including also studies presented in this thesis). Importantly, the Arp2/3 complex is essential for lamellipodia formation, since when cells are depleted of this complex through RNA interference no lamellipodia are formed (Steffen et al., 2006). Thus the criticism put forward concerning the dendritic model should not be interpreted as Arp2/3 is not operating at the cell edge.

The WASP family of NPF’s The activity of the Arp2/3 complex requires that it binds to different nucleation promoting factors (NPFs) of which the Neuronal Wiskott- Aldrich syndrome protein (N-WASP) is one of the most well characterized. The founding member of the WASP family of NPFs is the hematopoietically expressed WASP. It was first discovered as a mutated protein causing the X-linked immune disease Wiskott-Aldrich syndrome. Most commonly this disease is caused by mutations in one of the specific sub-domains of WASP which leads to a deficiency in the microfilament system of white blood cells and platelets, causing defects in immune defenses and blood clotting (Imai et al., 2003). N-WASP which is the ubiquitously expressed homologue of WASP and WAVE 1-3 are all well known members of the WASP family (Stradal and Scita, 2006) and through more recent work the family has expanded by the new members WASH, WHAMM and JMY (Rottner et al., 2010), illustrating the diversity of the Arp2/3 complex and its function. The members of the WASP family are actin binding multi-domain proteins containing several regulatory domains and with a main function to activate the Arp2/3 complex. Although the domain structure of the WASP family proteins varies, reflecting their different functions, they all have a conserved C-terminal module called WCA (fig. 4), which constitutes the smallest sequence necessary for Arp2/3 activation. The WCA is composed of a WH2 (WASP homology) domain, which is one of the most abundant actin binding domains (Dominguez and Holmes, 2010; Machesky and Insall, 1998; Symons et al., 1996) a basic connector sequence (called C for cofilin homology domain) and an acidic stretch (A) that interacts with the Arp2/3 complex (Marchand et al., 2001). Activation of Arp2/3 occurs upon binding to the WCA modules which induces a conformational change in the complex that brings the actin monomers and the complex together. The conformational switch makes the Arp2/3 complex mimic a (+)-end free actin dimer through the Arp2

33 and Arp3 subunits. Different WCA domains have different kinetics of actin assembly even though the affinities for actin are very similar (Zalevsky et al., 2001). The rest of the protein, which constitutes a much larger part then the WCA, is involved in targeting and regulation. The original members of this family contain a proline-rich central region and an N-terminus which is the most diverse region of these proteins. The central region interacts with numerous SH3 domain containing proteins, for instance IRSp53, WISH, PLCγ, and VASP (Bear et al., 2001; Castellano et al., 2001; Padrick and Rosen, 2010; Takenawa and Miki, 2001) and binding of these proteins causes an increased activity. The central region also contains multiple profilin binding sites (Suetsugu et al., 1998). In a so called bead motility assay where the beads were coated with fragments of WASP and WAVE 1 the WCA domain was seen to be the smallest region that can support the nucleating activity of the Arp2/3 complex. However the activity was enhanced by the proline-rich and N- terminal regions, and in the case of WAVE 1 the proline-rich region was required to induce motility (Yarar et al., 2002).

N-WASP The N-terminal part of N-WASP contains an N-terminal domain called WH1, a basic region that interacts with PIP2 and a G-protein binding domain (GBD) that binds active Cdc42. It is kept in an autoinhibited state, lacking Arp2/3 stimulating activity, via intra molecular interactions between the C region of the WCA domain and the GBD (Miki et al., 1998). This autoinhibition is in vivo stabilized by the WASP interacting protein (WIP) interacting with the WH1 domain (Martinez-Quiles et al., 2001; Volkman et al., 2002). WIP is a general stabilizer of N-WASP and in the cell N-WASP is always found in complex with this protein (Dong et al., 2007). External stimuli leads to activation of N-WASP by a number of interactions, active Cdc42 interacts with the GBD domain and PIP2 with the basic region, this is enough for activation in vitro (Rohatgi et al., 2000). In the physiological context where the auto inhibition is stabilized by WIP, also interactions with the F-BAR protein Toca-1 is required for activation (Goley and Welch, 2006; Ho et al., 2004; Takenawa and Suetsugu, 2007). Toca-1 interacts with the SH3 domain of N-WASP and is activated by the same Rho GTPase, namely Cdc42, as N-WASP. Toca-1 dimerizes upon activation and the two F- BAR domains form a concave surface which interacts with the anionic lipids of the plasma membrane and induces an invagination (Shimada et al., 2007). The dimerization of Toca-1 leads to clustering of N-WASP which in turn leads to an increase in activity on top of that gained from

34 the release of the autoinhibition (Takano et al., 2008). This scenario is essential for endocytosis, and also endosome propulsion by comet tail formation is dependent on N-WASP/Arp2/3 dependent nucleation (Taunton et al., 2000). N-WASP is also regulated by casein kinase 2 and focal adhesion kinase which phosphorylates residues in the WCA region (Cory et al., 2002; Suetsugu et al., 2002). This increases the activity in the presence of Cdc42 (Torres and Rosen, 2003). N-WASP is also involved in filopodia formation (Miki et al., 1998; Zigmond, 2000). This effect is however not dependent on Arp2/3 complex activation, since a mutant lacking the WCA region still induced filopodia (Lim et al., 2008). Probably filaments already present are then used for elongation into filopodia, likely depending on the presence of the Ena/VASP family protein Mena interacting with WASP (Lim et al., 2008). The I-BAR protein IRSp53, which induces a negative curvature on membranes, is required for this N-WASP dependent filopodia formation. Actin remodeling is required for exocytosis and in a study on neuroendocrine cells N-WASP was discovered to have a role in this process. In this study treatment with actin depolymerizing drugs had a dual effect on exocytosis, low concentrations led to increased exocytosis whereas high concentration reduced this process (Gasman et al., 2004). Since cortical actin covers the inner leaflet of the plasma membrane it must be remodeled during exocytosis (Gasman et al., 2004). Actin has also been observed to separate secretory granules into a small release ready pool and a large reserve pool (Trifaro et al., 2000). This shows that both depolymerization and polymerization is important and it indicates that filamentous actin does not only work as a barrier but also has an active role in exocytosis, an important conclusion in view of paper III in this thesis. Pathogens also use N-WASP/Arp2/3 dependent actin nucleation for propulsion. This relies on the pathogens recruitment of the cellular actin machinery (reviewed in (Frischknecht and Way, 2001; Stevens et al., 2006). Both the bacterium Shigella flexneri and Vaccinia virus recruit N-WASP to their surfaces to nucleate actin comet tails.

WAVE The WAVE proteins are not kept in an auto inhibited conformational state and there was long a question on how the activity of the WAVE proteins is regulated (see (Derivery and Gautreau, 2010). WAVE forms a complex with the proteins Brk1, Nap, Abi, Sra which interact with the N-terminal WAVE homology domain (WHD) of WAVE and as well as with each other, forming a complex set of interactions that also involves

35 the WAVE WCA domain (fig. 4) (Kunda et al., 2003; Le et al., 2006). This is how WAVE is kept inactivated and its activation occurs by Rac binding to Sra (Kobayashi et al., 1998). Initially it was thought that the other components were released form WAVE after activation, but recently it has been concluded that all the components remain associated after activation, activation merely releases the WCA domain (Ismail et al., 2009). Similar to the WASP proteins a BAR-protein is involved in the function of WAVE proteins. WAVE2 has been shown to bind to the I- BAR protein IRSp53 at the plasma membrane (Miki et al., 2000). IRSp53 like Toca-1 interacts with PIP2 and dimerizes, leading to clustering of two WAVE complexes. PIP3 has also been suggested to cluster WAVE at the plasma membrane, and PIP3 and IRSp53 have been shown to synergize in activating WAVE (Suetsugu et al., 2006). Additionally, a complex pattern of phosphorylations have been shown to participate in the regulation of WAVE. For instance the casein kinase 2 mentioned to have a regulatory function on N-WASP also phosphorylates the WCA domain of WAVE and increase its affinity for the Arp2/3 complex (Pocha and Cory, 2009). WAVE localizes to ruffling membranes (Miki 1998 EMBO J), to focal adhesions (Westphal et al., 2000) and to the tips of lamellipodia, but is not present in filopodia and in retracting parts of lamellipodia (Hahne et al., 2001). The components of the WAVE complex are recruited to the plasma membrane after Rac activation (Innocenti et al., 2004; Kunda et al., 2003; Steffen et al., 2004) and then induces formation of lamellipodia (Hahne et al., 2001). Knockout of WAVE1 leads to brain deficiencies (Soderling et al., 2003). WAVE2 knockout mice suffer from embryonic lethality, brain malformations and impaired angiogenesis (Yamazaki et al., 2003; Yan et al., 2003). Profilin has been shown to increase the activity of WAVE by interacting with the proline-rich sequence N- terminal to the WCA domain and similar to N-WASP it profilin has been hypothesized to deliver actin to the WH2 domain via this interaction. The profilin binding site according to structural studies, partially overlaps with the WH2 domain which could mean that when profilin:actin has been docked via the actin-WH2 interaction, profilin is dissociated (Chereau et al., 2005).

WASH, WHAMM and JMY This group of WASP-related proteins represents recent discoveries of actin nucleation promoting components with interesting properties. WASH exists in all except yeast. This protein associates with

36 up to six other proteins as part of a pentameric subunit core complex and with the heterodimeric capping protein CapZαβ as an accessory partner (fig. 4). If CapZ is absent, the cellular level of WASH decreases (Jia et al., 2010). In vitro, CapZ promotes Arp2/3 dependent actin assembly by capping of filament (+)-ends and preventing elongation, thereby directing the actin monomers to newly forming filaments. However, CapZ’s interaction with WASH appears to inhibit this capping activity. Primarily WASH seems to be involved in endocytic recycling by promoting endosome-linked actin assembly (Jia et al., 2010), and via interaction with the actin nucleator Spire the WASH core complex appears to have capacity to link two different actin nucleation processes together (Liu et al., 2009), though the significance of this remains to be seen. It is also interesting that WASH interacts with the microtubule system and prevents tubulation of the endosome membrane in a process that is dependent on both filamentous actin and microtubules. As mentioned above, N-WASP is also involved in endocytosis but the two proteins appear to operate on different vesicle populations (Duleh and Welch, 2010). The WHAMM-protein is coupled to the secretory pathway and proposed to be involved in ER to Golgi transport (Campellone et al., 2008). It has a membrane interacting domain (WMD) and a central coiled coil region, that binds microtubules in addition to its C-terminal WCA domain (fig. 4). It is ubiquitously expressed and localizes to cis Golgi, where it appears to play a central role in organelle integrity since both its depletion and overexpression cause derangement of the Golgi apparatus into small puncta (Campellone et al., 2008). Overexpression also increases formation of tubulovesicular structures, a process that requires both its microtubule binding domain and the Arp2/3- binding/actin filament promoting WCA domain, as well as intact microtubule and actin filament systems (Campellone et al., 2008). It has been proposed that WHAMM together with microtubules deforms vesicular membranes and that these structures are promoted and stabilized by actin polymerization. Thus, with respect to membrane tubulation WHAMM appears to have the completely reverse effect to that seen for WASH. The microtubule interacting properties observed for these two recently discovered NPFs is interesting since they are likely to represent new links for communication and cellular coordination of the activities exerted by the microfilament and microtubule systems. Although at present no data are available it is possible for instance that transportation processes by the microfilament and microtubule system of

37

mRNAs encoding different actin control proteins to the cell periphery are coordinated by such mechanism (see also paper I).

Fig 4. Domain organization of the Arp2/3 dependent nucleation promoting factors. The C-termini are all composed of Arp2/3 activating WCA modules albeit with varying numbers of WH2 domains. Abbreviations: A- acidic region, B- basic region, C- connector region, Cα- and Cβ- subunits of the heterodimeric capping protein, CC- coiled coil, GBD- GTPase binding domain, PRD- proline-rich domain, P- polyproline domain, L- actin monomer binding linker, N- N-terminus, Strump- strumpellin, SWIP- Strumpellin and WASH interacting protein, TBR- tubulin binding region, W- WH2 domain, WAHD1- WASH homology domain 1, WHD- WAVE homology domain, WH1- WASP homology 1 domain, WIP- WASH interacting protein, WMD- WHAMM membrane interacting domain. This scheme was adopted from (Rottner et al., 2010).

Finally, JMY which enters the nucleus in a response to DNA damage, works as a cofactor for the histone acetyl transferase p300/CBP during transcriptional regulation in a mechanism enhancing the p53 response (Coutts et al., 2009). Interestingly JMY is also involved in activation of both Arp2/3 dependent and independent nucleation (Zuchero et al., 2009). However, if and how JMY actually couples DNA damage to actin nucleation is not known. In Arp2/3 independent nucleation JMY operates via its tandem WH2 domains which recruit actin monomers to form a nucleus. This intrinsic nucleation does however not seem to be sufficient to promote cell motility, and although JMY knockdown leads to decreased motility, the effect seems to be indirectly due to up-

38 regulation of E-cadherin (Coutts et al., 2009). The actual role for JMY in Arp2/3 dependent actin assembly is not known.

Formins The formins are involved in various motile processes like cytokinesis, adhesion and filopodia formation where they form actin filaments arranged in bundles either running in a parallel fashion like in filopodia or antiparallel like in stress fibers (Frazier and Field, 1997; Kovar et al., 2006). The formins also have an important role in stabilizing microtubules but this will not be dealt with here. In mammals the formin family of proteins is encoded by 15 different genes. On their C-terminal side these proteins contain two highly conserved domains (FH2) which nucleate actin in vitro by stabilizing actin dimers, and, more to the center of the molecule a polyproline-rich domain (FH1) is present which binds profilin (fig. 5B) (Evangelista et al., 1997; Imamura et al., 1997; Watanabe et al., 1997). Profilin is required for the function of most formins since they recruit actin for their filament elongation activity as profilin:actin which binds the formin with higher affinity than profilin alone. The N-terminus of the formins is less conserved; it is important for their cellular localization and often it contains a Rho binding domain (RBD). Furthermore, most formins are kept in an auto-inhibited state by interactions between their N-terminal so called diaphanous inhibitory domain (DID) and C-terminal diaphaneous auto regulatory domain (DAD) (Romero et al., 2007; Zigmond, 2004). For their activation, the formins require interaction with active Rho, at the RBD. There is as yet no crystal structure of an intact formin, but based on structural studies of formin fragments and biophysical data of purified proteins, a model has been suggested where the active state is proposed to be a homeodimer which binds to the fast polymerizing (+)-end of filamentous actin via the FH2 domains (fig. 5B) (Chesarone et al., 2010). By processive ‘walking’ at the growing filament tip, they elongate the filament by incorporating an actin monomer from profilin:actin into one strand of the filament at the time. This processivity has been demonstrated by formin coated quantum dots following the filament end (Paul and Pollard, 2009), and it has been postulated that the dissociation of profilin is coupled to hydrolysis of the actin-bound ATP and that the subsequent Pi-release provides the energy needed for processive actin monomer incorporation into the filament (Romero et al., 2004). However, the view that ATP hydrolysis is required for the formin activity has later been challenged by work suggesting that

39 enough energy for profilin dissociation is released by the monomer incorporation itself (Paul and Pollard, 2009). Clearly, this is a complex process and more work is needed to resolve its details. As mentioned before, spontaneous actin nucleation requires trimer formation in vitro, but in the presence of the formin FH2-domain this fragment can stabilize an actin dimer long enough to allow for proper filament nucleation (Pring et al., 2003). The binding of the FH2 is specific for the actin dimer since it lacks affinity for actin monomers, and therefore it must act by capturing and stabilizing already existing actin dimers present in solution. This means that in vivo this nucleation activity is prevented by actin monomer binding proteins like β-thymosin and profilin and additional interactions are required for the formins to work as nucleators. Recent data from a biophysical study have emerged regarding this function, where a few basic amino acids in the DAD were shown to bind to actin monomers in an interaction not competing with profilin. Hence the DAD domain, working together with the profilin:actin-recruiting FH1 domain, and the actin dimer-binding FH2 domain, could be the factor required for in vivo nucleation of actin (Gould et al., 2011). In view of the polybasic sequence in synaptotagmin characterized in paper III this is interesting although under the in vitro conditions studied the latter seemed to have no effect on actin polymerization. Formins are most well-known for supporting filopodia formation but they also have a less well characterized role in lamellipodia formation (Yang et al., 2007). Depletion of mDia2, one of the most investigated mammalian formins, was shown to inhibit the formation of filopodia (Block et al., 2008; Pellegrin and Mellor, 2005), while overexpression of constitutively active mDia2 leads to an increase in the number of filopodia. This formin has been observed to follow the tip of the filopodium as it protrudes, however it has not been formally shown if it is involved in actin nucleation or elongation or both in this context. The activity of mDia2 is promoted by the Rho GTPases Rif and Cdc42, which are both GTPases known to induce filopodia formation in their active state (Pellegrin and Mellor, 2005). In the yeast S.pombe, assembly of the contractile ring is formin dependent (Yonetani and Chang, 2010). Also in mammals, formins are involved in cell division, during centrosome separation and chromosome movements (Pollard, 2010). The activity of mDia2 is inhibited by forming a complex with WAVE and Arp2/3, so these latter components, apart from having a role in

40 lamellipodia formation, can also interfere with formation of filopodia (Beli et al., 2008), which could be an efficient mechanism to switch between different actin regulatory mechanisms in vivo. There are a few other proteins identified that inhibit formin activity in vitro, for instance the Dia interacting protein/WASP interacting SH3 protein (DIP/WISH) (Eisenmann et al., 2007) and Spire (Quinlan et al., 2007). Also tropomyosins displace formins from barbed ends by annealing filaments (Skau et al., 2009).

Ena/VASP Enabled/vasodilator stimulated phosphoproteins (Ena/VASP) is like the formins a protein family whose members recently have been realized to operate as processive filament elongators although they have been studied for more than 20 years. Ena was found in a genetic screen in Drosophila (Gertler et al., 1995) and Mena, VASP and EVL are the mammalian members of this family. VASP was originally recognized to be primarily present in platelets (Halbrugge and Walter, 1989) but actually these proteins are widely distributed in mammalian tissue. They are built up by a few conserved domains. The N-terminal Ena/VASP homology (EVH1) domain is responsible for localization through binding to polyproline-rich motifs in a variety of proteins present for instance in focal adhesions and at the plasma membrane, and a central region containing a proline-rich sequence is known to interact with profilin (fig. 5C). And finally the C-terminal part contains an EVH2 domain which binds both monomeric and filamentous actin and a coiled-coil domain at the very C-terminus responsible for multimerization of Ena/VASP proteins (Krause et al., 2003). Lately the EVH2 domain has been proposed to represent a variant of the WH2- domain in WASP-family proteins albeit with no affinity for Arp2/3 (Ferron et al., 2007). Ena/VASP proteins only work as processive elongators when attached to a surface. In vitro, in solution Ena/VASP stays attached to the side of the filament after addition of an actin monomer to the (+)-end. Initially the difference in action between soluble and attached state created a controversy regarding the anti- capping activity of Ena/VASP. It has now been concluded that in solution the protein detaches from the barbed end allowing capping protein to bind, compared to the attached situation where no capping protein can bind because Ena/VASP remains associated with the (+)- end (Breitsprecher et al., 2008; Ferron et al., 2007).

41

Fig 5. Cartoons illustrating models for cellular actin assembly described. (A), Actin filament nucleation by the Arp2/3-NPF. The pentameric Arp2/3 complex has a low nucleating activity by itself but it is activated by NPFs. The WCA domain interaction with the Arp2/3 complex induces a conformational change leading to nucleation. (B), The formins processively elongate actin filaments. They form homodimers with their FH2 domains (purple) binding to the actin filament (+)-end. By their walking they elongate the filament by recruitment of profilin:actin (depicted as blue and red respectively) via multiple polyproline-sites of their FH1 domains, adopted form (Chesarone et al., 2010). As described the formins also nucleate actin filaments. (C), Clustering of VASP at the plasma membrane leads to processive association of VASP at the filament end. Only actin monomers recruited by VASP are added to the growing filament, the filament end is blocked for spontaneous interactions with for instance profilin:actin and capping protein. However, the supply of monomers is fueled by the interaction with profilin:actin. The polyproline site of VASP is proposed to bind several profilin:actin complexes but here only one is depicted. The EVH2 domain contains both a G- and an actin filament binding site. Interactions between the EVH1 domain and plasma membrane interacting proteins (yellow) aid in positioning VASP near the membrane, adopted from (Breitsprecher et al 2011.).

42

Ena/VASP has long been known to localize to the leading edge of migrating cells, and it is now presumed to work beneath the plasma membrane by docking to membrane bound proteins such as lamellipodin via its EVH1 domain. The ability to oligomerize is essential for Ena/VASP to be able to work as a processive elongator and it is proposed to work as a tetramer (fig. 5C). At focal adhesions it contributes to actin filament assembly in maturing contacts (Reinhard et al., 1992; Worth et al., 2010) and at tips of filopodia it is likely to work as a processive filament elongator (Schirenbeck et al., 2005; Svitkina et al., 2003). Myosin-X which also localizes to tips of filopodia likely has a role in transporting VASP here (Tokuo and Ikebe, 2004), also see section about myosins. VASP was the first protein seen to interact with the profilin poly-L- proline binding site (Reinhard et al., 1995). The central proline-rich sequence motif mentioned above contains the archetype glycine- pentaproline (GP5)-motif which binds profilin (Reinhard et al., 1995). Interestingly, this region of the molecule contains several sites which has a tenfold lower Kd for profilin:actin compared to profilin alone and furthermore their binding affinity for profilin:actin increases towards the site on the molecule where it associates with filamentous actin (Chereau and Dominguez, 2006; Ferron et al., 2007). Therefore it has been suggested that VASP recruits profilin:actin for filament elongation analogous to the mechanism proposed for formins (above). Levels of VASP at the leading edge have been seen to correlate with speed of cell migration (Rottner et al., 1999a) and the mechanism suggested is that higher levels of VASP recruits more profilin:actin to these sites of high actin dynamics. However, in vitro studies on the effect of profilin on (+)- end addition of monomers in the context of VASP function have come to different conclusions. Some have observed no effect of profilin on Ena/VASP dependent actin elongation rate (Breitsprecher et al., 2008) whereas others find a significant increase in elongation upon addition of profilin (Barzik et al., 2005). It is likely that differences in preparation or storage of the recombinant VASP has led to these discrepancies (Bear and Gertler, 2009). And interestingly, as was mentioned above, in profilin knockdown breast cancer cells the rate of migration is increased in a VASP dependent mechanism (Bae et al., 2009), where VASP localization at the leading edge also increases. The interpretation of this in view of profilin function is unclear, clearly however in the absence of profilin an increased concentration of VASP can somehow compensate by recruiting actin for filament growth from other sources. VASP and

43

Mena are both present in most cells and they are implicated in neuritogenesis. Deletion of Mena leads to severe neurological defects and this means that in nerve tissue, VASP cannot functionally replace Mena. Knockout of Mena in combination with a heterozygous profilin 1 gene knockout was lethal, demonstrating the importance of the profilin- Mena interaction (Lanier et al., 1999). VASP is important for the movement of Listeria monocytogenes (Laurent et al., 1999). After infection, this intracellular pathogen moves through the cell cytoplasm using the actin polymerizing machinery of its host (Frischknecht and Way, 2001). It recruits VASP to its surface by the bacterial protein Act A. Injection of PxA to cells infected with this pathogen drastically reduced the motility of the pathogen and caused detachment of their actin tails, suggesting that profilin:actin is involved in Listeria tail elongation (Grenklo et al., 2003).

Tandem-monomer-binding nucleators Spire, cordon-bleu (Cobl), leiomodin (Lmod), JMY and APC belong to a type of quite newly discovered actin nucleators which harbor tandem WH2 motifs that brings monomers together to form a nucleus. Apart from this actin nucleating activity they constitute a heterogeneous group of actin binding proteins (reviewed in (Firat-Karalar and Welch, 2011)). Spire for instance can induce actin filament severing (Bosch et al., 2007). Since these nucleators were recently discovered, future developments in this area will be interesting to follow. No matter what, it should be realized that all these NPFs have coupled activities and their differences enables a balanced and tightly regulated activity which can respond to the diversity of signals reaching the cell.

BAR-proteins The BAR (Bin-Amphiphysin-Rvs) domain typical for the so called BAR- proteins is highly conserved. These proteins form dimers that attain a curved shape due to the biophysical properties of the BAR domain and by binding to membranes they induce membrane curvature. They are thereby involved in remodeling of membrane structure as components of the actin rich proteo-lipid zone at the cell periphery. There are three families of BAR containing proteins, two which induce membrane invaginations and contain either a so called classical BAR domain or an F-BAR (Fer/CIP4-homology-BAR) domain. The third family contains an I-BAR (Inverse-Bin-Amphiphysin-Rvs) domain and instead of inducing membrane invaginations this domain induces protrusions of the

44 membrane covered cell edge. However with the recent discovery of the srGAP2 which carries the F-BAR domain but functions like an I-BAR protein (Guerrier et al., 2009) the coupling between classification and function does not fully apply anymore and this classification therefore warrants some caution. Different BAR-domains induce different degrees of membrane curvature. For instance the F-BAR proteins induce membrane curvature in the range of the size of endocytic vesicles. One of the most well studied I-BAR domain protein, IRSp53, is typically involved in lamellipodia and filopodia formation. It has an N-terminal I- BAR domain, a partial GBD motif, an SH3 motif, a potential WH2 domain, and in some isoforms also a PDZ domain which mediates binding to certain scaffolding proteins. Through its RBD-motif, IRSp53 is regulated by Cdc42 and via its I-BAR domain it also binds Rac1 (Govind et al., 2001; Krugmann et al., 2001). Through interactions with a wide range of actin binding proteins such as WAVE 1 and 2, Mena, Dia1, Eps8 and N-WASP that all bind to its SH3 motif, and its BAR- domain, IRSp53 mediates remodeling of the actin rich proteo-lipid zone at the cell periphery (Scita et al., 2008). In the I-BAR domain there are four lysines that are involved in actin binding (lysine 142, 143, 146, 147) and this domain also has a filament bundling activity (Millard et al., 2005; Yamagishi et al., 2004). Therefore it is postulated that the modulation of membrane structure, recruitment of actin regulatory proteins and its integral bundling activity is the mechanism behind BAR-protein dependent filopodia formation. In neuronal cultures IRSp53 induces neurites that exhibit extensive lamellipodium and filopodium formation, and Cdc42 is essential for this activity (Govind et al., 2001). Filopodia formation was also observed in other cell types (Lim et al., 2008). Mutation of the lysines in the I-BAR domain abolishes filopodia formation and a mutation of the SH3 domain reduces the number of filopodia formed (Lim et al., 2008). In vitro the IRSp53 I-BAR domain together with liposomes generates membrane tubules of a size of 40-90 nm through the activity of its I-BAR domain (Mattila et al., 2007) this probably is similar to what is observed in cells where the I-BAR domain by itself induces cell surface protrusions similar in appearance to filopodia. However, these protrusions are less dynamic than filopodia normally formed, which have well organized filament bundles in contrast to the I-BAR domain induced protrusions, hence there are reasons to express concern on these I-BAR domain generated protrusions as being referred to as filopodia (Mattila et al., 2007). In the case of standard filopodia with a growing filament bundle, IRSp53 with its binding

45 partners has been associated to follow the tip of the growing filopodia. It is notable that IRSp53 was unable to induce filopodia in Mena/VASP knockout cells suggesting that it is also likely to cooperate with these proteins in its actin polymerizing function (Ahmed et al., 2010; Lim et al., 2008).

Actin depolymerizing factors The archetype protein involved in actin severing and depolymerization is gelsolin. From a structural point of view this protein has six subdomains and it caps the (+)-end of actin filaments. It is under the control of PIP2 and Ca2+ (Hartwig, 1995). Gelsolin has a well-known function in severing actin filaments during apoptosis (Geng et al., 1998; McGough et al., 1998). However, form the point of view of profilin cofilin is more interesting.

ADF/Cofilin The actin binding ADF/cofilin family of proteins is important in the control of actin dynamics in particular with respect to depolymerization of the filament. In mammals, cofilin-1 and ADF (actin depolymerizing factor) are found in non-muscle cells while cofilin-2 is muscle specific (Hotulainen et al., 2005). In vitro ADF/cofilins from different species have been characterized to bind to ADP-bound G- and F-actin with higher affinity than to the ATP-bound actin and they have been shown to increase the rate of actin dissociation from the (-)-end of the filament (Carlier et al., 1997). It has also been proposed that these proteins can cap the (+)-end and in addition to this they have been assigned a severing activity (Andrianantoandro and Pollard, 2006; Ichetovkin et al., 2002; Pope et al., 2000). In the cell ADF/cofilin localizes to protruding lamellipodia (Lai et al., 2008) and it is present at sites where new lamellipodia are about to form (DesMarais et al., 2005; DesMarais et al., 2004). Its actions here are however less characterized although several functions can be foreseen based on in vitro data. The most widely accepted role is that ADF/cofilins increases the dissociation rate from the (-)-end of the filament and thereby increase the monomer concentration so that more free actin is available to be delivered to profilin, which will compete with cofilin for binding to actin since it has a higher affinity. Profilin will upon binding, exchange the nucleotide on actin for ATP and deliver it as polymerization competent profilin:actin- ATP to polymerizing (+)-ends. Thus cofilin functions in increasing the

46 turnover of actin and despite the fact that cofilins are primarily depolymerizing factors they can cause an increased polymerization (Carlier et al., 1997). It has been suggested that cofilin and the Arp2/3 complex function together. Arp2/3 has been seen to distribute to sites for new polymerization only if cofilin is present and the Arp2/3 activity is required for the cofilin to increase actin polymerization (DesMarais et al., 2004; Sidani et al., 2007). It is unclear whether this Arp2/3 dependent activity in cooperation with cofilin requires a specific Arp2/3-dependent NPF. Cofilin is phosphorylated by LIM kinases and dephosphorylated by the phosphatases slingshot and cronophin. The phosphorylation inactivates its ability to bind actin. This regulation of cofilin versus actin is under the control of Rho GTPases. The Rho target ROCK and different Rac/Cdc42 activated PAKs activate LIM-kinases which then in turn phosphorylate cofilin (DesMarais et al., 2005; Huang et al., 2006). Also PIP2 is an important interaction partner of cofilin (van Rheenen et al., 2007). In vitro this interaction inhibits the binding to actin (Van Troys et al., 2000) and also in vivo manipulations in PIP2 levels affect the activity of cofilin. Phospholipase C dependent PIP2 hydrolysis is suggested to release membrane associated cofilin allowing for a rapid effect on actin organization, in response to receptor stimulation (Leyman et al., 2009; Mouneimne et al., 2004). The on/off rate for the coflin-PIP2 interaction is very rapid which also is important for the function for cofilin and this is reflected by the expression of a mutant having a stronger interaction with PIP2 which leads to a more persistent migration and a higher speed (van Rheenen et al., 2007). Thus it has been proposed that growth factor stimulation which causes hydrolysis of PIP2 releases cofilin which then increases actin filament depolymerization and increases the pool of actin ready for polymerization. Cofilin is inactivated upon phosphorylation and after subsequent dephosphorylation it binds to PIP2 in the plasma membrane again (van Rheenen et al., 2007). Since cofilin responds to receptor stimulation by increasing the turnover of the microfilament system it is a key protein in chemotaxis and like several other of the major proteins involved in actin control, cofilin has been associated with cancer invasion and metastasis (van Rheenen et al., 2007; Wang et al., 2006b).

47

Intracellular transport and myosins Myosins constitute a superfamily of proteins encoded by at least 40 genes in the human genome (Odronitz and Kollmar, 2007). They cooperate with actin filaments in a contractile process to generate force and are involved in organelle transport, endo- and exocytosis, cytokinesis, protein transport, transport of mRNPs and cell migration. The myosins have in common that they bind to actin filaments through their head domain which hydrolyses ATP to transduce force for the myosin power stroke. This force transduction can be used for walking along a filament and for sliding or contracting actin filaments of opposing polarity. There are both single headed and two headed myosins, the head region localizes to the N-terminus, while the C- terminal part of the protein is either cargo binding or filament forming. The latter case involves only conventional myosins that polymerize into filaments and powers actin filament contraction or sliding. These filaments are built into the actin array of muscle cells and stress fibers in non-muscle cells. This group of myosins is called myosin-II because they contain two heads (Dantzig et al., 2006). The other myosins do not form filaments and are termed unconventional myosins. Apart from a head domain these myosins have a motor regulatory center and a cargo binding tail domain. Myosin-II molecules contain two myosin heavy chains (MHCII) two regulatory light chains (RLC) and two essential light chains. In non- muscle cells the action of myosin-II is controlled by phosphorylation of the RLC which increases the ATPase activity of the MHC in the presence of actin. A number of kinases, around 20 known, including myosin light chain kinase (MLCK) and Rho associated kinase (ROCK), are able to phosphorylate myosin-II (Vicente-Manzanares et al., 2009). The phosphorylation of the RLC also affects the ability of myosin-II molecules to assemble into typical bipolar filament. There are a multitude of signaling pathways that lead to phosphorylation, for instance Ca2+- calmodulin activates the MLCK and RhoA activates ROCK. Apart from its RLC phosphorylating activity, ROCK also inhibits the major myosin phosphatase PP1, protein phosphatase 1 (Matsumura and Hartshorne, 2008). The large number of myosin kinases enables a tight control of myosin-II. As one example, the kinases already mentioned localize to different cellular compartments. MLCK is more peripherally located and ROCK is more central (Totsukawa et al., 2000). This means that there is a spatial regulation of RLC-phosphorylation and, furthermore Rac has a

48 function in inhibiting myosin-II phosphorylation (Yamada and Nelson, 2007). This concurs with the function of Rac in the fast filament turnover at protrusive edges. The activity of myosin-II is strictly organized in a moving cell where myosin-II is required for maturation of adhesions. Substrate rigidity increases the activation of myosin-II (Beningo et al., 2006) and interestingly myosin-II dependent force transduction contributes significantly to maturation of focal adhesions in non-muscle cells (Wolfenson et al., 2011). The diverse groups of motor molecules for cargo transport along the cytoskeletal filaments are constituted by myosins for actin dependent transport and kinesins and dyneins respectively for microtubule dependent transport. Both endo and exocytotic pathways require transport of vesicles (Ross et al., 2008). For instance when vesicles undergo anterograde transport they first move along the microtubule system and then are transferred to the microfilaments for final transport to the plasma membrane. Vesicles usually contain many motors, e.g several copies of the same kind and different types. By having more than one motor present the cargo can undergo continuous transport even if the motor is not processive. Continuous transport can in such cases also be accomplished by a tether which holds the cargo in place while the motor recovers form a power stroke. This tether could be formed by a region on the myosin itself and also a non processive kinesin has been shown to have this function (Hodges et al., 2009; Kincaid and King, 2006). There are many hurdles for a transporting vesicle to pass during its journey through the cell, for instance the dense order of the filament system and proteins binding to the filaments. This is probably one reason for the many different motor isoform molecules, for instance some motors prefer tropomyosin decorated filaments (Tang and Ostap, 2001) and there are also preferences for different types of filament orders among the myosins (Nagy et al., 2008b). This means that the myosins will locate to different regions of the cell based on filament structure and arrangement. The most well studied unconventional myosins for intracellular transport are probably mosin-I, V, VI, VII and X. Myosin-VI is a (-)-end directed motor, the others move towards the (+)-end of actin filaments in accordance with the classical direction of force generated by myosin-II. Myosin-I isoforms, there are eight of them in humans, are single headed and function in vesicle transport. Myosin V-genes express double headed isoforms, these work as processive

49 motors and have for instance been shown to have a role in mRNA transport. There is one isoform of myosin-VI, this myosin has a role in endocytosis and retrograde vesicle movement. The two isoforms of Myosin-VII are involved in organizing the cytoskeleton and cell adhesion. The two headed Myosin-X of which there is only one isoform, is widely expressed and has a role in nuclear positioning and filopodia formation. It works as a processive motor with a preference for bundled actin, therefore it localizes to the tips of filopodia. Myosin-X has a unique C- terminal domain which contains three pleckstrin homology domains, a myosin tail homology 4 domain which interacts with microtubules and a FERM domain which is cargo binding. The FERM domain has been seen to interact with VASP and β3-integrin and to transport these molecules (Tokuo and Ikebe, 2004; Zhang et al., 2004; Zhu et al., 2007). Overexpression of myosin-X induces formation of filopodia (Berg and Cheney, 2002; Zhang et al., 2004) and it is suggested that the transport of components used in filopodia formation contributes to this phenomenon. Filopodia are however still formed also in the absence of the cargo binding tail domain of myosin-X (Bohil et al., 2006), but these filopodia differed from those induced by full-length myosin-X, they were shorter and more unstable. It is therefore likely that several functions of myosin-X are used in the process of filopodia formation. While the initiation of filopodia formation caused by myosin-X might however not be directly coupled to its cargo binding it could instead be caused by bridging of filament tips, collecting enough filaments, around 10, which can support pushing of the plasma membrane (Watanabe et al., 2010). The myosin-X dependent transport of β-integrins would contribute to adhesion formation while VASP would add to the process of elongation, therefore the absence of the cargo binding domain leads to formation of the uncharacteristic filopodia described above. The second PH domain of myosin-X was recently shown to interact with PIP3. This interaction is important for the localization to filopodia, if it is abolished a larger fraction than normal localizes to endosomal vesicles and filopodia formation is impaired (Plantard et al., 2010). Myosin based transport has been suggested for transport of actin monomers to tips of growing filopodia (Berg and Cheney, 2002). Myosin-X is a likely candidate for this process since as was described above, filopodial growth aided by this myosin is supported experimentally (Berg and Cheney, 2002). Active transport of actin into

50 filopodia is proposed since diffusion of actin monomers is considered to be the rate limiting factor for their elongation (Lan and Papoian, 2008; Mogilner and Rubinstein, 2005). Interestingly experimentally observed filopodia are often much longer and grow faster than has been predicted by computer simulated models where diffusion of actin monomers is set as a rate limiting factor. Consequently it is easy to hypothesize that monomers are actively delivered to the filopodia tips. There are also suggestions that diffusion is not sufficient for fast lamellipodial protrusion (Zicha et al., 2003) there is however less consensus on this topic. Computer simulations have been made for myosin-X transporting VASP which in turn bind to actin monomers (Zhuravlev et al., 2010). The simulation included only two monomers per motor, but actually up to eight monomers could be transported since VASP is a tetramer and myosin-X has two cargo binding domains, and even more actin molecules could be considered if also the ability to transport profilin:actin is included. According to the simulation, a motor based supply of actin monomers is a possible mechanism to support filopodial protrusion with the criterion of a directed motor, which myosin-X is. In connection to the study on synaptotagmin 1 in paper III, a study on the role of myosin-X for the transport of profilin:actin was initiated. The observations still remain to be described, however preliminary imaging show protruberations of a cherry-tagged profilin moving along filopodia and sometimes these are connected to dynamic clusters of GFP-tagged myosin-X in agreement with an active transport of profilin and probably profilin:actin.

Cell adhesion To resist force cells make contacts to the extra cellular matrix (ECM) or to each other. These contacts constitute complex protein arrangements which in a moving cell need to be dynamic and form and dissolve on a minute time scale. Both actin and intermediate filaments connect to these structures to give support to tissues, here only the involvement of microfilament is discussed since it is most well characterized in the context of cell migration. As was mentioned above these connections are initiated by adhesion receptors called integrins, at the tip of the protruding lamellipodia or filopodia. There are also other adhesion receptors such as proteoglycans and cadherins, this section will only involve the integrins since they are best characterized and play a prominent role in actin mediated force transduction. Integrins are heterodimeric trans-membrane receptors formed by α and β-subunits.

51

There are a number of isoforms of each kind which gives a broad library of ligand specificities where different ECM-components are common ligands. Integrins are expressed in an inactive state at the cell surface. They are activated as a result of receptor activation, for instance in response to chemokines, leading to complex signaling recruiting talin to the integrin cytoplasmic tail which causes a conformational change so the integrin receptor adapts a high affinity state for its ligand (Anthis and Campbell, 2011; Shattil et al., 2010). Integrins can also be activated from the outside by binding to an ECM ligand which leads to conformational change of the integrin, causing its clustering and recruitment of talin. In the focal adhesion, talin forms one well known connection that binds the cytoplasmic part of the β-integrin and actin filaments. Other well-known proteins of the focal adhesion complex are α-actinin, vinculin, filamin, focal adhesion kinase (FAK) and paxillin. As indicated above, the integrins are receptors that signal bidirectionally across the membrane. Both directions of signaling involves the direct and indirect connections of many components which are either part of the focal adhesion or are more peripherally involved in signaling. These components are collectively called the integrin adhesome and involves more than 150 proteins (Humphries et al., 2009; Zaidel-Bar et al., 2007). At the very early stage of adhesion formation taking place in the lamellipodium, the adhesions are submicron sized, but yet contain a number of adhesion proteins, for instance vinculin and paxillin. At this stage the counteracting force comes from the rearward flowing of actin. When the nascent adhesions reach the lamella they either dissolve in the zone of depolymerizing actin or they start connecting to stress fibers and mature into small round connections. The growth and maturation of these adhesions is myosin-II dependent and a stress fiber bundle that will connect with the adhesion is built up either by the reorganization of existing filaments or by new actin polymerization (Hotulainen and Lappalainen, 2006), with the actin filament cross-linker α-actinin and myosin-II appearing at the onset of filament bundle formation (Choi et al., 2008). The focal adhesions are hence located at the termini of stress fibers and grow larger and become stretched in the direction of and in response to the force they are exposed to. The adhesions grow under tension because they are able to sense force and then recruit components to strengthen (Schwartz, 2010). The integrins themselves experience a change in their conformation in response to force leading to an increased affinity for the ECM (Campbell

52 and Humphries, 2011; Geiger and Yamada, 2011). Talin also experiences a conformational change which rapidly recruits vinculin, which is also a force sensor (del Rio et al., 2009; Galbraith et al., 2002; Grashoff et al., 2010) and the adhesion then grows in size almost at the same time scale as the recruitment of vinculin, indicating a subsequent cascade recruitment of other proteins (Riveline 2001). The force-induced conformational change of the integrin has been proposed to trigger activation of FAK, whose recruitment to adhesions is dependent on myosin-II, for activation of unbound integrins in a pathway depending on the PI3-kinase (Katsumi et al., 2005; Schwartz, 2010). Ligand binding to the integrin receptor causes tyrosine phosphorylations of its tails, this as well as the application of force on the adhesions leads to activation of many signaling pathways. For instance Rho GTPases become activated and calcium levels are elevated. The signaling events also activate certain gene expression, like cell cycle genes, genes associated with differentiation and matrix remodeling genes (Orr et al., 2006). In response to the signal transduction by the assembling adhesion, a number of GEFs and GAPs are recruited. An important focal adhesion adapter protein is P130cas which exposes its Src phosphorylation site in response to force, one of its roles is the recruitment of GEFs (Sawada et al., 2006). FAK and paxillin are examples of other proteins that undergo phosphorylation after recruitment to the forming focal adhesion and in turn recruit Rho GTPase regulatory enzymes. As the adhesion matures and tension increases, zyxin is recruited and subsequently recruits Ena/VASP proteins (Lele et al., 2006). At even higher force zyxin binds to stress fibers and, then Ena/VASP follows and increases actin polymerization at these sites (Cattaruzza et al., 2004). Zyxin also relocates to the nucleus. From this it is clear that advance remodeling occurs in the microfilament system in response to changes in force (Campbell and Humphries, 2011). The cells also influence the ECM in accordance with the applied force and collagen and fibronectin fibrils for instance align in the direction of the force applied by the connecting integrins (Gao et al., 2003). This effect is caused by the pulling on the ECM-proteins inducing a conformational change that allows the fibrils to form (Chiquet et al., 2003). As cells migrate, adhesions need to be disassembled. This usually occurs when the cell has migrated so that the contact is located at the rear. Microtubule tips are thought to be involved in disassembly (Broussard et

53 al., 2008). Also downregulation of myosin-II contractility, where vinculin seems to work as an important force sensor, is involved. As was described above, when this vinculin is under force other proteins are recruited and conversely, when it is not under force the adhesion disassemble (Grashoff et al., 2010). Also the protease calpain and endocytosis could be involved in disassembly (Bhatt et al., 2002; Burridge, 2005), but the fine details of the disassembly mechanisms remain to be resolved. mRNA localizaiton Localized translation of mRNA is a mechanism for spatio-temporal control of protein compartmentalization, it results in a sub-region of the cell being enriched in newly synthesized protein. This process has been shown to be of significant value for a number of different physiological and developmental conditions. In a screen on Drosophila embryos around 70% of the mRNAs were seen to be subcellularly localized (Lecuyer et al., 2007), and this plays a role for instance in axis formation (Roth and Lynch, 2009). In yeast, mRNA localization to the bud tip is important in mating type determination (Paquin and Chartrand, 2008) and in higher eukaryotes mRNA localization is essential for a number of processes, many of them involved in cytoskeletal remodeling. β-Actin mRNA localization to the leading edge is required for persistent migration in fibroblasts (Shestakova et al., 2001), and localization of integrin-β3 supports motility of lung carcinoma cells (Adereth et al., 2005). In neurons, localization of mRNAs to more distal regions is thought to be economical since the distances can be very long and one transcript can give rise to multiple protein copies. In agreement with this, mRNA localization has been observed to have a role in neuronal growth, axonal guidance (Willis et al., 2007) and synaptic plasticity (Gkogkas et al., 2010; Kang and Schuman, 1996; Steward and Levy, 1982). β-Actin mRNA localizing to the growth cone and dendritic spines is one transcript known to have a role in all these situations (Tiruchinapalli et al., 2003; Yoon et al., 2009). CamIIKa (Miller et al., 2002) and BDNF mRNAs (An et al., 2008; Chiaruttini et al., 2008) are other transcripts localizing to dendritic spines that are important for synaptic plasticity. Targeted messages are recognized for localization via cis-acting sequence elements, functioning as signal sequences. These are often present in the 3’-UTR of the mRNA but are also found in the 5’-UTR and the coding region. So called trans-acting factors involved in the localization process

54 will bind to the signal sequence of the mRNA, with the initial the factors binding already during transcription. Possibly these early factors in a common mechanism are recruited by the transcriptional machinery. This was shown for the RNA binding protein Shep2p in budding yeast which for instance is involved in localizing the Ash1 mRNA to the bud tip (Shen et al., 2010). During export through the nuclear pore the mRNA is unwound, at this stage additional factors residing in the cytoplasm assemble to the mRNPs forming a large complex referred to as an mRNA granule. These contain among other things motor proteins, aminoacyl tRNA synthases, elongation factors and ribosomal subunits (Sotelo-Silveira et al., 2008). Interestingly, actin, in either monomeric or a short oligomeric form is also a component of mRNPs and accompanies the mRNP into the cytoplasm (Percipalle et al., 2001). The role for actin in this context is unclear but could be in the assembly of a transport- competent granule and efficient mRNA trafficking. The binding of factors already in the nucleus could be crucial for efficient transport. In an experiment abolishing nuclear localization of Shep2p, the protein was still seen to bind the mRNA but localization was severely affected (Shen et al., 2009). However exactly what factors that assemble in the nucleus or in the cytoplasm is not clear. Motor proteins seem to bind in the cytoplasm which has been shown for Myo4p, a myosin-V homologue, responsible for transport along actin cables in budding yeast. In some cases also the splicing machinery is involved during mRNA localization (Hachet and Ephrussi, 2004). The RNA granule travels after assembly into a transport competent complex, on cytoskeletal structures to the target site for translation. This is dependent on either actin filaments or microtubules or on both. In neurons the microtubules seem to constitute the major transport track. In Cos-cells it was discovered that mRNAs without a localization signal also bind to motor complexes and are transported, but the interaction is transient and the transport less directed (Fusco et al., 2003). Perhaps the sequence containing mRNAs show a more directed transport because these mRNAs have an increased affinity for the motor complexes, or maybe the targeted mRNAs somehow change the properties of the motor complex. As was mentioned above, mRNA transport is a regulatory factor for cytoskeletal remodeling. And in this context the β-actin mRNA is probably the most well studied transcript. In the case of the growth cone, localized β-actin mRNA mediates turning towards guidance cues like was shown for netrin1 (Leung et al., 2006), and in fibroblasts it permits persistent unidirectional migration (Shestakova et al., 2001). In

55 hippocampal neurons the β-actin mRNA transport was seen to be dependent on the microtubule system (Tiruchinapalli et al., 2003) while in fibroblasts it was seen to be independent of an intact microtubule system (Latham et al., 2001) and paper I) and instead required the microfilament system with myosin-II activated by the RhoA - ROCK pathway being essential (Latham et al., 2001). In a more recent study myosin-Va was also found to be involved in β-actin mRNA transport and one suggestion is that in fibroblasts myosin-Va works as a transport motor and myosin-II captures the transcript for translation (Salerno et al., 2008). Interestingly, in motor axons β-actin mRNA has been found in transport granules together with kinesin-II and myosin-Va (Sotelo- Silveira et al., 2008) therefore the transport mechanism between neurons and fibroblasts seem to share some similarities, but many questions still remain. A factor involved in mRNA localization, zip code binding protein 1 (Zbp1) has been shown to bind the β-actin mRNA in the nucleus through a cis-acting sequence element called zip code sequence in the 3’- UTR and this factor is important in all known cases of β-actin mRNA transport (Condeelis and Singer, 2005). Zbp1 binds the mRNA during transport and acts as a translational repressor of the β-actin mRNA by preventing binding of the 80s ribosomal subunit (Huttelmaier et al., 2005). The repression is released by Src phosphorylation of Zbp1, if this process is impaired it leads to a decrease in peripheral actin and aberrant neurite outgrowth. Since activation of myosin-II is involved in the localization in fibroblasts, it is interesting to note that Src can also work as an upstream regulator of RhoA, therefore are areas with high RhoA and Src activity likely sites of β-actin mRNA translation. This concurs with the requirement for β-actin mRNA localization to myoblast cell-cell junctions where activities of RhoA and Src are known to be high (Rodriguez et al., 2006). Since loss of β-actin mRNA localization leads to impaired cell movement and defects in neuronal development one assumes that depletion of this transcript in the animal should have similar consequences. Expression of β-actin is essential for viability thus a conditional knockout mouse, where expression was depleted in motor neurons by induction of gene-excision at embryonic day 10, was constructed. Surprisingly no phenotype was seen (Cheever et al., 2011). One factor leading to the large difference between mistargeted β-actin mRNA and depletion of the mRNA could be that mistargeted localization gives a more prominent defect than no

56 protein at all. Also, the knockout was not initiated until embryonic day 10 therefore requirements for β-actin early in development could have been missed. β-actin mRNA is not the only mRNA associated with the microfilament system that actively localizes. The mRNAs for the components of the Arp2/3 complex have all been seen to localize to active cell edges, it is reasoned that synthesis of all components in the same place facilitates complex assembly (Mingle et al., 2005) and retargeting of the mRNA for Arp2 to the perinuclear region led to a loss of directional migration (Liao et al., 2011b). The transport complex for these subunits has not been characterized, but the transport is dependent on both the microtubule system and the microfilament system. In addition, many adhesion proteins seem to be translated locally (de Hoog et al., 2004; Vikesaa et al., 2006). Recently the mRNA encoding Dia1 was seen to actively locate to the ER (Liao et al., 2011a). Proteins targeted for the secretory pathway are targeted to the ER via a signal sequence in the nascent peptide chain as they are translated. The targeting sequence for mDia1 was also located in the peptide chain and targeting to ER was dependent on both this site and the ribosomal complex. RhoA also plays a role in this ER-targeting via a direct interaction in the RBD in the translating protein. What RhoA does in this context is unknown, it could be tethering the translating complex to the ER preventing it from being released into the cytoplasm. The mechanism for mDia mRNA-targeting is distinct from other known mechanisms of cytosolic proteins since translation is required. Apart from the important pathway of Zbp-dependent mRNA localization a distinct localization pathway involving adenomatous polyposis coli (APC) has been uncovered. In a screen performed on fibroblasts more than 50 transcripts were seen to accumulate in protrusions in response to migratory stimuli (Mili et al., 2008). This required an intact microtubule system and APC while it was independent of an intact microfilament system. Transcripts for β-actin or the Arp2/3 complex did not come out in this screen. The transcrips were seen anchored to the tips of microtubules and this required the APC which is known to bind to microtubule (+)-ends, if microtubules also have a role in the transport to the protrusion is not clear. Since APC is required for microtubule organization and is also emerging as an actin nucleator this protein could certainly have a bridging role between different signaling pathways. The two pathways involving either APC or Zbp1 seem to have distinct functions (Mili and Macara, 2009). This APC dependent

57 pathway could also be interesting to consider in light of the study on profilin mRNA in paper I since this mRNA localizes in a pathway distinct from β-actin mRNA. There are still many remaining questions on how and why mRNAs localize. Resolving the control and coordination of different pathways for transport will be important for understanding different aspects of cell behavior such as how polarity is maintained.

58

ACTIN AND PROFILIN IN THE CNS

Actin polymerization and actomyosin interactions are major force generating mechanisms common to all eukaryotic cells. Therefore it is not surprising that such actin dependent activities are required for process formation during development of the central nervous system as well. Additionally actin, in cells of the CNS is coupled to memory formation and learning for which actin dynamics in dendritic spines is crucial, and in the presynapse actin has a role in the tight control of neurotransmitter release. This latter subject is especially important in the context of this thesis since paper III is discussing a role for actin polymerization connected to the function of the presynaptic protein synaptotagmin.

Neuritogenesis Actin polymerization provides the necessary force for extension of neuritic processes in all steps during neuritogenesis. Initially, the neuronal cell has a more or less spherical shape and then goes through 5 identifiable morphological stages during maturation into a neuron (Dotti et al., 1988). First the cell has a unipolar distribution of lamellipodia and then in stage 2 cylindrical processes with a growth cone at their tip start to protrude and retract over a few hours. In stage 3 an axon is forming which protrudes without retraction. Stage 4 is characterized by formation of the dendritic tree and stage 5 occurs about one week after neurite induction and involves the formation of synaptic contacts. These stages have been mapped mainly in hippocampal embryonic cultures and may not look exactly the same in all cases (Kessels et al., 2010). Different factors, like signals from the extracellular environment, play a role for how this development progresses and particularly differences between primary cultures and established cell lines like the SH-SY5Y cells used in paper III are likely. However, the initiating step for neurite sprouting is common to all neurons. This involves breaking of the round cell symmetry causing the stage 1 to stage 2 transition (da Silva and Dotti, 2002). Interestingly in the light of the synaptotagmin study (Paper III), filopodia formation has been demonstrated to carry a key function for this transition, presumably by establishing contacts via integrins to the extracellular matrix (Galbraith et al., 2007). In cortical neurons lacking Ena/VASP proteins, no filopodia are formed and these cells are unable to form neurites. Ectopic expression of other factors e.g. mDia2 that

59 rescues filopodia formation also restores neurite formation in a process that depends on a dynamic microtubule system (Dent et al., 2007; Edson et al., 1993).

Actin in neuronal process formation In neurons, actin-based protrusion has been best characterized in the growth cones of advancing axons whose axonal outgrowth is dependent on guidance of cytoskeletal dynamics to form the right number and length of these processes and to guide them to correct location. The axonal growth cone is formed by a core region, which is filled with membrane organelles and growing microtubules surrounded by a fan- shaped structure formed by actin-based lamellipodia and filopodia (Dent et al., 2010; Forscher and Smith, 1988). The filopodia of the growth cone similarly to the tip-cells of sprouting arteries navigate the developing tissue as they form the highly branched networks of the nervous and the vascular systems, the growth cone and the tip cells both control guidance, are highly motile, invasive and have long filopodia. Formation of vessels and neurons also occur in a mutually correlated manner, vessels secrete factors that control their innervation and nerve derived signals control vascular patterning (Adams and Eichmann, 2010). Many of the same actin nucleation and elongation promoting factors already described in the previous section are involved in organizing the dynamic microfilament arrangement of the growth cone which in fact is often said to resemble the lamellipodia in a fibroblast (Dent et al., 2010). Furthermore treatment with the actin filament (+)-end capping drug cytochalasin B reveals the same treadmilling behavior of actin polymerization occurring at its leading edge (Forscher and Smith, 1988). The Arp2/3 complex is, as was described in a previous section, instrumental in the control of actin filament formation, especially in the lamellipodia. It has also been shown that the Arp2/3 complex together with N-WASP is essential for normal neurite formation. In immature cultures expression of N-WASP deficient in Arp2/3 activation leads to a reduced extension of neurites and conversely, a constitutively active form of N-WASP leads to an increase in the number of dendritic branches (Machesky et al., 1999; Suetsugu et al., 2002). Axonal process formation is on the contrary negatively regulated by Arp2/3. Loss of N- WASP function, or disrupted expression of Arp3 results in increased axonal length and branching (Strasser et al., 2004). N-WASP mediated neurite formation is activated by both the actin filament binding protein Abp1 and the small Rho GTPase Cdc42, which show cooperative

60 activity versus N-WASP activation (Pinyol et al., 2007). The protein syndapin is another component seen to be involved in N-WASP regulation in neurons. It is targeted to the plasma membrane via its F- BAR domain and it recruits N-WASP via its SH3 domain, thereby releasing N-WASP from its autoinhibited state, which is then further activated by Abp1 and Cdc42 (Dharmalingam et al., 2009). Thus the Arp2/3 complex with its partners is recognized to be of key importance for actin filament formation during neuritogenesis and essential for proper formation of the nervous system, but also other actin nucleators are important. Overexpression of the brain enriched actin nucleator Cobl, leads to an increase in neurite number and branching of dendritic arbors, a similar phenotype as the overexpression of N-WASP. Disruption of Cobl expression by RNAi leads to loss of branches and fewer dendrites (Ahuja et al., 2007). It has therefore been proposed that Cobl is especially important in development of the dendritic tree while the Arp2/3 complex with partners have an especially important function in axonal development (Kessels et al., 2010). Additionally the formin DAAM regulates axonal morphogenesis in Drosophila as reflected by the fact that disrupted expression of this protein reduces the number of growth cone filopodia. In mouse, loss of function of DAAM causes malformations of the CNS, like misrouted axons and malformed commissures (Matusek et al., 2008). These effects are emphasized in animals where Ena or profilin expression is also abolished, or where a loss of function mutation is introduced to Rac. It was therefore suggested that DAAM works together with Ena and profilin to regulate axonal growth in a Rac dependent process. Considering previous data Ena was suggested to interact with DAAMs FH2 domain (Schirenbeck et al., 2005) and proflin via its FH1 domain (Wallar and Alberts, 2003). Another formin, mDia1, has a positive effect on axonal elongation in cerebellar neurons, operating downstream to RhoA. This GTPase also has an inhibitory effect on neuronal process formation via ROCK activation, suggesting fine tuning of this process dependent on Rho GTPases, the details of which are not fully understood, but it seems that the Rho GTPase Rac also is involved in this (Arakawa et al., 2003). Furthermore, Dia1 has an effect on axonal guidance by interactions with the microtubule system, affecting the orientation of individual tubules (Geraldo and Gordon-Weeks, 2009; Ishizaki et al., 2001). In yeast and plant cells formins form actin cables that serve transports throughout the cell. In animal cells however, bundled actin structures of uniform polarity have only been observed in

61 the small protrusions of dendritic spines and filopodia (Pollard and Cooper, 2009). There are a number of other proteins involved in actin polymerization during the formation of neuronal networks indicating that this area still needs to be explored, see (Hall and Lalli, 2010) for review.

Profilin isoforms in neuritogenesis Profilin is an important component in neuronal process formation and as was mentioned above the profilin 1 and 2 isoforms are both expressed in the brain, where the estimated expression level for profilin 2 is 2-3 times higher than for profilin 1 (Witke et al., 2001). One of the first in vivo studies on profilin in neuronal process formation showed that mutations in chickadee, the only Drosophila profilin homologue, results in premature termination of motor axons (Wills et al., 1999). In the rat pheochromocytoma derived cell line PC12, often used for studies on neuronal differentiation, simultaneous knocking down of profilin 1 and 2 inhibits neuronal outgrowth, but this is not seen after knocking down of profilin 2 only (Sharma et al., 2005). The authors do not comment on the fact that their knockdown of profilin 1 is without effect although their data show that this is the case. The conclusion is that the two isoforms have full capacity to replace each other, which is not so surprising since these isoforms have similar biochemical properties (Lambrechts et al., 1997). However, this is very interesting in light of in vivo data from profilin knockout mice where disrupting the expression of each isoform leads to different effects. As mentioned above knocking out profilin 1 leads to 100% embryonic lethality at the pre-implantation stage (Witke et al., 2001) and as a matter of fact, knocking out just one allele of profilin 1 leads to developmental defects on brain morphology (Pilo Boyl et al., 2007). Profilin 2 knockout mice on the other hand do not show any such defects in embryonic brain development. Hence profilin 2 knockouts have a mild phenotype compared to 50% reduction in profilin 1 expression, (the synaptic effect after loss of profilin 2 expression is discussed in a later section). Why the expression of an extra profilin isoform is needed in the brain is not known. However, since profilin 2 exists in 2-3 times higher levels than profilin 1, it it seems unlikely that knocking out one allele of profilin 1 should have a larger consequence than the complete knockout of profilin 2 if cell function was dependent on the total profilin concentration. Therefore the two isoforms clearly serve different purposes which is an important topic for future research and a matter that will be discussed more in this thesis.

62

The two profilin isoforms 1 and 2 are both coupled to RhoA signaling but with different endpoints. In cultures of CA1 pyramidal neurons the profilin isoforms were both seen to work downstream of the neurotrophin receptor P75NTR which activates the RhoA-ROCK pathway. Overexpression of profilin 2 led to an increase in dendritic complexity, and knockdown of profilin 2 reduces the number of dendrites and dendritic spines, the same phenotype as was seen for P75NTR overexpression. The effect of overexpression of profilin 1 differs partly from that of profilin 2 overexpression since it increases the number of dendritic spines but has no effect on dendritic branching. Profilin 2 overexpression partly rescues the p75NTR overexpressing phenotype by increasing the number of dendrites, but the number of spines are still few. Overexpression of profilin 1 on the other hand restores the number of dendritic spines, and hence this suggest that both profilin isoforms work downstream of P75NTR (Michaelsen et al., 2010). It therefore seems reasonable that the two profilin isoforms have similar but yet complimentary function in the formation of the neuronal network. Profilin 2 was also seen to work as a ROCK effector during the onset of neuronal sprouting (Da Silva JCB 2003). It was described as a negative regulator of this process by having a stabilizing function on the microfilament system. These two functions might seem contradictory since it appears that profilin 2 has a both positive and negative role in neuronal process formation, perhaps indicating dual functions under temporal regulation for this profilin isoform. Profilin 1 has been proposed to function as an effector in RhoA signaling by Lambrechts et al and this was shown to be a consequence of its binding to PIP2 (Lambrechts et al., 2006). Treatment with the ROCK inhibitor drug Y27632 led to neurite outgrowth in PC12 cells. This effect was larger in cells overexpressing wild type profilin 1 compared to cells expressing normal levels of profilin. However, overexpression of a mutant defective in PIP2 binding led to the same increase in neurite outgrowth even without drug treatment. The inhibition of ROCK causes a decrease in PIP2 levels since the PI5-kinase, which produces PIP2, is activated downstream of ROCK. The effect seen after Y27632 treatment was therefore suggested to be caused by decreasing levels of PIP2, which frees wild type profilin:actin to contribute actin to filament formation, and analogously the mutant profilin unable to bind PIP2 was proposed to cause a constant increase in actin polymerization. (Lambrechts et al., 2006). Profilin 1 and 2 both harbor binding sites for actin, polyproline- rich motifs and PIP2, thus expressing the same biochemical properties. However, slight variations in this respect still appear to be the case since

63 the binding affinities at least for PIP2 and polyproline varies between the two with profilin 1 displaying a tighter interaction with the lipid and profilin 2 shows stronger binding to polyproline (Witke, 2004). Such differences could fine-tune the function of the different isoforms and may be the reason why analysis of brain extracts have led to the conclusion that they in fact interact with different proteins (Lambrechts et al., 1997; Michaelsen et al., 2010; Wang et al., 1999; Witke et al., 1998). It should be understood, however, that profilin isoform variation with respect to cellular function is poorly understood and clearly is a subject for future investigations.

Synapse formation In the initial stages of formation the synapse lacks characteristic features of a synapse and therefore it is difficult to visualize how the formation- process occurs. However, based on a number of observations many features of this process have been mapped. Synapses in the CNS are located along axons with the pre-synapses being formed by swellings called boutons on the axon and the post synapse formed by a meeting dendritic spine. It appears that synapse formation can be triggered either by a growing axon encountering a dendrite or vice versa (Meyer and Smith, 2006; Niell et al., 2004). Also, sites for synapse formation can be induced along the whole axon, at least in immature neurons. This is illustrated by the fact that molecules essential for synaptic vesicle release are found all along the axon during early stages of synaptogenesis (Burry, 1986). The synapse is an asymmetric structure with a pre-synapse, the synaptic cleft and post synapse. The pre-synapse, or bouton, contains a cluster of synaptic vesicles adjacent to the active zone which is a specialized region of the plasma membrane where the synaptic vesicles fuse. The active zone characterized by an electron dense protein arrangement or cytoskeletal matrix called the “cytomatrix at the active zone” (CAZ). The vesicles of the synaptic cluster are grouped into three pools. Most of the vesicles of the synaptic cluster are part of the reserve pool but some, which are docked to the plasma membrane, belong to the so called ready releasable pool. There is also a recycling pool which resides close to the plasma membrane and will become part of the ready releasable pool at moderate stimulation of the synapse. At depolarization of the membrane leading to Ca2+-influx, vesicles of the ready releasable pool fuse to the plasma membrane of the active zone. The fusion depends on a precise sequence of events involving a multipartite protein machinery that responds to Ca2+ and overcomes the energetically unfavorable step of combining the membranes of the vesicle and the

64 plasmalemma. When vesicles fuse to the plasma membrane they release their neurotransmitter in the synaptic cleft and this will be sensed by receptors in the postsynapse that are confined at the plasma membrane in an electron dense region called the post synaptic density (PSD). The CAZ and PSD both connect to trans-synaptic adhesion molecules (CAMs) and extracellular matrix proteins which stabilize the structure of the synapse. After neurotransmitter release the cell must compensate for the membrane addition and recycle the synaptic vesicle proteins, for this the fused plasma membrane and associated proteins must travel to the periactive zone where vesicles will reform through clathrin mediated endocytosis. After initial contact, according to observations made of the nerve-muscle situation, it only takes seconds to minutes for excitatory postsynaptic potentials as well as an increase in axon calcium concentration to occur (Buchanan et al., 1989; Evers et al., 1989). Then the synapse stabilizes over the following hours to days and the active zone is formed, first containing only a few docked synaptic vesicles. Dense core vesicles carrying synaptic components and coated vesicles, involved in recycling, are often observed. Subsequent maturation takes place over days-weeks and involves functional changes like types and subunit composition of voltage gated calcium channels and changes in the probability for neurotransmitter release (Mozhayeva et al., 2002). The synaptic components are transported in the axon as vesicular elements containing synaptic vesicle proteins, calcium channels and scaffolding proteins of the CAZ. When a filopodia makes contact with an appropriate target cell these transport vesicles rapidly coalesce at the site of contact to form a presynaptic site (Ahmari et al., 2000; Shapira et al., 2003; Zhai et al., 2001). Axo-dendritic contacts are in most cases transient but as the nervous system matures it becomes more and more likely that these contacts will form a synapse. This is probably a consequence of decreasing dynamics of the system as more contacts are formed (Friedman et al., 2000). It is not completely known how stabilization of a nascent contact to achieve maturation and become more permanent is regulated. Some observations imply that neuronal activity is involved in regulating the motility of the pre-and postsynapse. It was seen that motility of both axonal and dendritic filopodia is blocked by glutamate in hippocampal cultures, this effect is coupled to an influx of Ca2+ and does not affect synaptic vesicle traffic, hence synaptic transmission is unaffected (Chang and De Camilli,

65

2001; Fischer et al., 2000). In axonal filopodia this effect is dependent on AMPA and kinate type receptors and in the dendritic filopodia on AMPA and NMDA receptors. Furthermore, in hippocampal neurons it was discovered that the motility of filopodia is actually bidirectionally regulated by glutamate activation of kinate receptors; low concentrations of glutamate leads to an increased motility and high concentration to a decreased motility. As a consequence this could mean that in the young nervous system, where there is more space, glutamate released from the presynaptic terminal will be more diluted and lead to an increased, and more exploratory motility. The reverse will be the case as the system matures (Tashiro et al., 2003). In immature neurons activity of the synapse has also been shown to induce an assembly and stabilization of filamentous actin with Cdc42 likely having a regulatory role (Shen et al., 2006; Yao et al., 2006). Cellular adhesion molecules (CAM’s) like N- cadherin span the synaptic cleft and have a role in the stabilization of the early synapse (Bruses, 2000). This stabilization probably allows other proteins to interact so that maturation of the synapse can start. It is important early during the formation of the nervous system but there is a reduced actin dependence and a reduced sensitivity to defects in N- cadherin binding as it matures. Treatment of a neuronal culture for 48 hours at day 5-6 in vitro with latrunculin A leads to loss of presynaptic sites and dispersal of synaptic vesicles but at day 18-20 in vitro the culture is resistant to this treatment (Zhang and Benson, 2001). During the maturation of the CNS, expression of structural proteins like piccolo in the presynapse and PSD-95 in the post synapse correlate to the resistance to actin filament disrupting drugs (Li and Sheng, 2003) and it also coincides with the aforementioned decrease in dynamics of neurons in mature cultures.

Actin in the presynapse Two filament structures have been observed in the presynapse, one is formed by the phosphoprotein synapsin 1 and the other is identical to actin filaments, which is abundant throughout the presynaptic terminal and is also present in the active zone. Often actin filaments are associated to synapsin filaments (Landis et al., 1988; Siksou et al., 2009a) and these in turn link to the synaptic vesicle. Synapsin is also involved in the transport of vesicles from the reserve pool to the ready releasable pool (Kuromi and Kidokoro, 2005; Verstreken et al., 2005). There are both long filaments extending from the active zone appearing to subgroup the active zone vesicles into columns and there are short

66 filaments interlinking the vesicles (Cingolani and Goda, 2008). As has been shown in the lamprey synapse filamentous actin is present at a higher density around the vesicle cluster (Shupliakov et al., 2002) while it is only found at a low level within the cluster (Bloom et al., 2003). This is also in congruence with studies of cultured hippocampal neurons using GFP-actin. In this model system a dynamic shift in actin pools was seen between resting and active synapses. Using a Triton X-100 extraction procedure it was estimated that around 30% of the actin was filamentous in the resting synapse compared to around 50% in the active synapse (Sankaranarayanan et al., 2003). Monomeric actin is distributed homogenously over dendrites and axons of hippocampal neurons (Zhang and Benson, 2002). In a recent study of the Torpedo electric organ, voltage gated calcium channels were determined to make up an important connection to the submembranous cytoskeleton (Carlson et al., 2010). These ion channels are connected to many components of the CAZ, including spectrins, desmoyokin and dystrophin. Myosin-I also interacts with this complex and is suggested to constitute a dynamic link to the microfilament system in the polymerizations and depolymerizations taking place during nerve signaling. On the extracellular side, the calcium channels are linked to laminin. Voltage gated calcium channels also link to synaptic vesicle proteins such as synaptotagmin, which has a role in calcium sensing (Catterall and Few, 2008). The role of actin in the presynaptic vesicle cycle is not well understood but it has been suggested to have an organizing role of the vesicle pools both by restricting mobility and by aiding in the transfer between the pools (Cingolani and Goda, 2008). At the active zone actin could either guide vesicles to the membrane and facilitate docking, or serve as a barrier. One difficulty in mapping the functions of actin is reflected by the fact that different studies have come to different conclusions, probably because actin dependent activities actually are different in different types of synapses. In hippocampal neurons mobility of vesicles increased when filamentous actin was depolymerized (Shtrahman et al., 2005), but in the mouse neuro-muscular junction no such effect was seen (Gaffield et al., 2006). And in the Drosophila neuro-muscular junction there even seems to be a positive correlation between filamentous actin and vesicle release in a process depending on N-ethylmalemide sensitive factor (NSF) since downregulation of this factor led to a decrease in filamentous actin, and reduced vesicles to be part of the ready releasable

67 pool. Myosin-V co-immunoprecipitates with the v-SNARE synaptobrevin, and its activity is important for neurotransmitter release. Therefore it has been suggested to transport synaptic vesicles. (Bridgman, 1999; Miller and Sheetz, 2000). In chromaffin cells the t- SNARE syntaxin (see further, below) and myosin-V have been seen to interact in a Ca2+ dependent manner which could mean that at neuronal stimulation, vesicles transported on actin filaments are targeted to the plasma membrane (Watanabe et al., 2005). A short treatment with the actin destabilizing drug latrunculin A, which sequesters actin monomers, was seen to promote neurotransmitter release, the miniature excitatory postsynaptic current (mEPSC) frequency increased approximately five-fold, and the Ca2+-evoked neurotransmitter release also increased (Sankaranarayanan et al., 2003). Nonetheless the latrunculin A treatment did not lead to disruption of the vesicle cluster in a timescale of minutes, the size of the ready releasable pool did not change reflecting that there was no effect on endocytosis. Nor did the actin filament stabilizing drug jasplakinolid have an effect on either endo- or exocytosis. From this it was concluded that actin does not simply serve as a mechanical network to keep the synaptic vesicle cluster intact nor does it have a major role in vesicle recycling. Instead, by interacting with regulatory molecules like for instance bassoon and synapsin, filamentous actin serves to maintain their concentration, at the active zone. It was also suggested that actin negatively regulates vesicle release either directly or indirectly by inhibiting the late stage of priming of the vesicle and thereby preventing fusion, when the actin filaments are disrupted, more vesicles reach the final priming step and when a signal to promote fusion arrives, more release is observed (Morales et al., 2000). The effect seen was not observed with other drugs such as cytochlasin D, which targets the barbed end of the actin filament. The difference in effect is quite surprising since in a dynamic system the end result of both drug treatments should be a depolymerization of filaments. The result was interpreted as if the filaments targeted by latrunculin A were short and highly dynamic, since the effect was seen after treatment with the actin monomer binding drug (Morales et al., 2000; Sankaranarayanan et al., 2003). Clearly the matter of actin dynamics in the presynapse remains uncertain. Perhaps actin has a role in both restraining and mobilizing the reserve pool like vas seen in neuroendocrine cells (Gasman et al., 1999). If treatment with depolymerizing drugs disturbs both functions, such a dual mechanism will be masked. No matter whether actin has an active role in neurotransmission or if its function is more passive, the changes

68 in levels of filamentous actin must mean that it is under tight control. In would be very interesting for a future study to examine the actin dynamics using single molecule tracking in a way that has been performed in the postsynapse (see further below Frost 2010). Based on observations of the effect of actin drugs on longer time scales, actin depolymerization has an effect on synaptic stability (Kim and Lisman, 1999). Treatment with latrunculin B and cytochalsin D generated defects in both pre and postsynaptic transmission and inhibitory effects on long- term synaptic plasticity were seen. The exposure time for the actin inhibitors lasted for tens of minutes. Profilin 1 and 2 are as was already mentioned both expressed throughout the CNS, and these two profilins have in studies been concluded to have similar but not identical biochemical properties (Lambrechts et al., 1995). However, as was explained in a previous section they do have different functions in vivo. Profilin 2 has an important role in fine-regulation of synaptic function despite the fact that abolished expression of this protein does not affect embryonic development of the brain or LTP/LTD, learning, glycine receptor and NMDA/AMPA receptor clustering are all normal (Pilo Boyl et al., 2007), however, signaling was affected since the animals exhibited a novelty seeking behavior as a result of higher synaptic excitability with clear involvement of the striatum, and also other brain regions being affected. It further led to a morphological change at the synaptic level with larger active zones and a 30% increase in perforated synapses. At the functional level there was a slight increase in mEPSC and EPSC and a 30% increase in v-SNARE-t-SNARE interactions while the normal increase in filamentous actin seen at synaptic stimulation was absent. These results are reminiscent of the outcome in the studies conducted by (Morales et al., 2000; Sankaranarayanan et al., 2003) where Latrunculin A was used to depolymerize synaptic actin filaments. Morales et.al concluded that the disruption of the actin filaments cause more vesicle release because actin normally hinders a late priming step of the vesicles. Related to this, in the study on profilin 2 by (Pilo Boyl et al., 2007) it was concluded that profilin 2, by having an effect on actin polymerization, is also involved in priming, docking or fusion and the authors hypothesize that the reason the CNS requires an additional profilin isoform is because it is needed for a tight regulation of synaptic vesicle exocytosis. It therefore seems that profilin:actin has an important but subtle role in neurotransmitter release in higher organisms.

69

Actin is involved in synaptic vesicle recycling, as was shown in the lamprey nerve terminal and in the large calyx of Held synapses. When depolymerizing actin at the large calyx of Held there was a decrease in vesicle release as an effect of reduced vesicle recycling (Sakaba et al., 2005). In response to a strong activity of the lamprey synapse, an actin network develops from the endocytic zone, near the active zone (Merrifield et al., 2004; Shupliakov et al., 2002). These new filaments are associated with clathrin coated vesicles at the plasma membrane and uncoated vesicles more distally. If the filaments are disrupted, this cluster of vesicles is decreased in active terminals and uncoated vesicles are observed close to the plasma membrane. This suggests that actin could be involved in pinching off of the vesicles and in transporting them to the reserve pool, a transport that could either be driven by treadmilling or myosin motors. These observations are not in congruence with what has been observed in for instance hippocampal synapses were as described above no effect on endocytosis was seen after actin depolymerization (Morales et al., 2000; Sankaranarayanan et al., 2003). It might be that in these small synapses, actin polymerization is not utilized for endocytosis and this differs from what was observed in the large synapses of lamprey and the calyx of Held. Another unresolved issue is how the membrane after vesicle fusion travels to the periactive zone for endocytosis, this could require a force generating activity of actin polymerization, something that is also discussed in paper III

Actin in the post synapse The postsynaptic structure of an excitatory synapse in the CNS is formed on a dendritic spine, a type of short actin enriched protrusion connected to the main dendrite by a thin neck (Landis and Reese, 1983). The spines form and mature as synapses are established and they are divided into four main morphologies; filopodia like, mushroom shaped, thin and stubby. The common view is that mature mushroom shaped spines are formed from filopodia-like precursors. A number of actin regulatory proteins are involved in the formation of these precursors but presently not much is known about this process. It is likely that proteins that are known to lead to formation of parallel filament assemblies and that form filopodia in cell culture, like the Ena/VASP family (Dent et al., 2007) and formins (Hotulainen et al., 2009) are involved. The I-BAR protein IRSp53 which has been shown to affect spine morphogenesis in cultured neurons (Choi et al., 2005) is also likely to have a role. Actin has a number of functions in dendritic spines, all from organizing the structure

70 of the PSD (Sheng and Hoogenraad, 2007) and anchoring receptors (Renner et al., 2008) to facilitating trafficking of transport vesicles (Schlager and Hoogenraad, 2009) and localizing the translation machinery (Bramham, 2008). The morphology of the spine is correlated to synaptic function, smaller spines are more dynamic and larger ones are more sensitive because the larger surface correlates with a larger PSD and a higher prevalence of neurotransmitter receptors. Spine head volume is also correlated to the number of synaptic vesicles in the bouton, and the amount of neurotransmitter that is released (Murthy et al., 1997; Pontrello and Ethell, 2009; Schikorski and Stevens, 1997). In the adult brain the dendritic spines continue to be dynamic, they appear, withdraw, and change in size, an activity involved in memory and learning, and that is dependent on actin dynamics. Disruption of actin filaments in dendritic spines using latrunculin A results in reduction of the number of AMPA receptors (Zhou et al., 2001) and a dynamic microfilament system controls the structure and composition of the PSD by connecting to its scaffolding proteins (Allison et al., 2000; Kuriu et al., 2006). A recent study taking advantage of photoactivatable GFP-actin for single molecule tracking provides a valuable insight into the dynamics of the actin filament system in dendritic spines (Frost et al., 2010). In this study it was determined that the moving rate of the molecules observed was slower than that of diffusion and they in fact were incorporated into polymerizing filaments undergoing treadmilling similar to what has been reported for the lamellipodium in fibroblast like cells (Lai et al., 2008). The filaments of the spine are short, most of them shorter than 200 nm. Larger spines express more complex actin dynamics than small ones, indicating a functional organization occurring as the synapse strengthen. Actin polymerization in the spine head occurs mostly at the spine tip, more at the lateral tip than at the center, and the molecules of the filaments flow toward the center of the spine in a net direction. In the spine interior filament orientation is heterogenous. In the spine neck direction of polymerization is not as obvious as the spine tip, but there is a general direction of a rearward flow towards the spine center. This indicates that filaments from the spine head and neck converge in a common zone for depolymerization. Surprisingly no augmented polymerization occurred in relation to the endocytic zone, indicating that an upregulated actin activity is not required for endocytic recycling in the post synapse as is the case in at least some types of pre synapses.

71

LTP/LTD The synaptic connections building up the neuronal circuits in the brain make up the “hardware” of our thoughts and memories. Synaptic activities for instance caused by the inputs from our sensory system lead to morphological modulations in the shape and size of spines which are correlated to synaptic strength of these excitatory synapses (Kasai et al., 2010a). The mechanism for storage of information in the brain is widely accepted as being carried out by synaptic contacts being strengthened or weakened, and dendritic spines formed or removed. An increase in synaptic strength, called long-term potentiation or LTP, is induced by high frequency stimulation of the synapse. The opposite is called long- term depression (LTD). This is induced by low frequency stimulation and results in a lower efficacy of the synapses caused by a decrease in receptor density. LTP shifts the actin contents into the filamentous form and leads to an increase in spine volume, while LTD decreases actin filament content and causes spine shrinkage (Okamoto et al., 2004). These events are specific at the level of individual synapses but alterations also occur in nearby synapses, i.e. LTP induction at one synapse lowers the threshold for LTP induction at nearby spines (Harvey and Svoboda, 2007) and such a clustered plasticity is expected to be important for stimulus induced information storage. The structural plasticity of dendritic spines was summarized in 15 ‘spine learning rules’ in a recent review article (Kasai et al., 2010b). The processes required for formation, stabilization and removal of synapses are all caused by actin dependent synaptic plasticity, and of course actin regulatory proteins are required for these processes. The Arp2/3 complex is essential for spine head growth (Grove et al., 2004; Hotulainen et al., 2009). This complex is present only in low amounts in the PSD, but in higher amounts more to the center of the spine head (Racz and Weinberg, 2008). A compartmentalization of Arp2/3 might reflect the distribution of actin pools in the spine. The activity of this complex is positively correlated to spine density. For instance, Abp1 which is involved in both activation of N-WASP and phosphorylation of the Arp2/3 complex was seen to increase the density of mature spines (Haeckel et al., 2008). Also knocking down N-WASP resulted in a similar phenotype as downregulation of the Arp2/3 complex due to knockdown of the p34 subunit of the complex. In both cases spine density decreases in hippocampal neurons and the proportion of thin filopodia-like spines increases (Hotulainen et al., 2009; Wegner et al., 2008). The microfilament system of dendritic spines is also

72 managed by microtubules. Dynamic tubules of the dendrite can enter spines and thereby bring microtubule associated proteins. One example is EB3 which can promote cortactin activity, which is another protein known to activate the Arp2/3 complex (Jaworski et al., 2009). Cofilin is especially concentrated at the plasma membrane in the spine (Zhou et al., 2007) and is very important in spine plasticity. Overexpression of an inactive mutant promotes formation of mature spines (Shi et al., 2009) and this protein is also involved in the control of diffusion of excitatory AMPA receptors (Gu et al., 2010; Rust et al., 2010). The changes that occur when postsynaptic spines grow and shrink during synaptic remodeling are thought to be coupled to receptor trafficking in that more surface area gives space for more receptors. However it has recently been shown that AMPA receptor insertion is not temporally coupled to spine enlargement but instead dependent on cofilin correlated activity. An increase in cofilin activity was seen to be important for insertion of AMPA receptors which is a hallmark for LTP, however the general state for cofilin during LTP is inactivation via phosphorylation and an increase in cofilin activity is associated with spine shrinkage seen during LTD (Zhou et al., 2007). Cofilin activity must therefore be under tight temporal control during LTP (Gu et al., 2010). One process for phosphorylation of cofilin involves the small Rho GTPase Rho B. It was discovered studying RhoB knockout mice that the phosphorylation of cofilin after LTP-induction is completely abolished and that these animals have less dendritic spines in the hippocampus, the spines are also wider and longer. It is very likely that the effects seen are a result of impaired plasticity (McNair et al., 2010). Evidence suggests that another important pathway for inactivating cofilin via phosphorylation is via Eph2B receptors that activate the RhoA-ROCK-LIMK-1 pathway (Shi et al., 2009). The fact that cofilin is under such complex regulation and that it is involved in neurological disorders, e.g. Williams syndrome (Hoogenraad et al., 2004) means that further research is needed. Profilin 1 and profilin 2 are both present in dendritic spines (Neuhoff et al., 2005) but their functions are not clear. Both isoforms show an increased localization to these structures after stimulation, with the increase in profilin 2 concentration being most prominent (Ackermann and Matus, 2003; Neuhoff et al., 2005). However the studies were performed by assessment of endogenous expression of profilin 1 and a GFP construct for profilin 2, making comparisons difficult. Profilin 1

73 was found to be expressed in individual subtypes of neurons, with a high expression in hippocampal pyramidal cells where it was found in the dendritic spine center as well as the PSD and the presynapse. There is a slight increase in synaptic localization of this profilin isoform at both the pre- and postsynapses after synaptic stimulation, but the mechanism behind or the functional implications for this were not further explored (Neuhoff et al., 2005). Stimuli causing increased intracellular Ca2+ lead to targeting of profilin 2 to dendritic spines, therefore both stimuli that promotes LTP and LTD leads to spine localization and incidentally both these processes have been shown to stabilize dendritic spines (Fukazawa et al., 2003; Star et al., 2002). The stimulation causing profilin 2 targeting lead to stabilization of spines, with a block in motility within 20 minutes. This targeting was dependent on the poly-L-proline binding site since it was blocked in the presence of a polyproline peptide sequence from VASP, also this block led to a destabilization of the spine structure (Ackermann and Matus, 2003). These data indicate that profilin 2 has a role in spine stabilization during memory formation. However, in the knockout study on profilin 2 no alteration of post synaptic plasticity was seen, the LTP/LTD response was normal (Pilo Boyl et al., 2007), perhaps due to profilin 1 playing a compensatory role. Using immunogold labeling profilin 2 was seen to localize to both the pre and post synapse similar to profilin 1 (Pilo Boyl et al., 2007).

74

SYNAPTIC VESICLE FUSION

The vesicle cluster of the synapse is organized in three arrangements, the reserve pool, the readily releasable pool and the recycling pool. The readily releasable pool vesicles, which constitute only a fraction of the vesicles, around 10 per synapse, are docked at the active zone while the reserve pool vesicles are distally located. The synaptic vesicles are filled with neurotransmitter through a pH gradient generated by proton pumps which acidify the lumen of the vesicle. Loaded vesicles are targeted to the active zone, docked and primed for fusion (Schikorski and Stevens, 1997). Docked vesicles are opposed to the plasma membrane and short, thin filaments presumed to be synapsin, link the docked vesicles to the plasma membrane (Siksou et al., 2009a). The fusion to the plasma membrane is triggered by an arriving action potential leading to an influx of calcium. The sensor for the calcium signal is synaptotagmin which together with the SNARE complex mediates fusion of the synaptic vesicles (see further below). After fusion vesicles are recycled via clathrin mediated endocytosis (fig. 7A) (Schweizer and Ryan, 2006).

SNAREs The central complex mediating synaptic vesicle fusion is called the SNARE complex (soluble N-elylmalemide-sensitive factor attachment protein receptor complex) (Jahn and Scheller, 2006). It was first isolated and further characterized by Rothman and colleagues when they were searching the receptors for NSF (N-ethylmaleimide-sensitive fusion protein) and SNAPs (soluble NSF attachment proteins) by affinity purification of bovine brain extracts (Sollner et al., 1993b). The SNARE complex is involved in docking synaptic vesicles to the active zone with individual SNARE proteins being inserted in the lipid bilayer of the vesicle and the plasma membrane. Docking occurs via interactions of the SNAREs, namely via their core domains. These are α-helices that become intertwined when they interact, thus forming a so called core complex that also need other factors for functionality. In neurons the SNAREs are synaptobrevin (also known as VAMP), syntaxin and SNAP-25. Two of these, syntaxin and SNAP-25 are associated to the plasma membrane and form a complex called the target (t)-SNARE. Syntaxin is composed of a transmembrane domain and one core domain, while SNAP-25 with its two core domains is associated to the plasma

75 membrane via four palmitoylated cysteins (Nagy et al., 2008a). Synaptobrevin, via its transmembrane domain, is inserted in the synaptic vesicle and therefore called vesicle (v)-SNARE. It interacts with the t- SNARE complex via its single core domain and upon vesicle fusion the SNAREs form a stabile four helix bundle first enabling docking of the vesicle and then with further zippering of the core domains causing fusion (Weninger et al., 2008). In vitro, docking and thermal induced fusion occurs without requirement for SNAP-25 (Sakaba et al., 2005). However this protein is required for a functional Ca2+ responsive system, although it does not interact with Ca2+ itself but with synaptotagmin which is sensitive to the Ca2+-signal (see further below) (Tang et al., 2006; Weninger et al., 2008). Also vesicle docking requires the involvement of further components in vivo. Munc18 and 13 are two such proteins involved in docking and priming of the synaptic vesicle. This is related to the protein syntaxin which adopts a closed conformation, preventing it from binding to SNAP-25 prematurely. Munch 18 shifts syntaxin between its closed and open conformation and stabilizes the closed conformation unless it is dephosphorylated. Dephosphorylation exposes an additional binding site for syntaxin which enables its open conformation. By this latter type of binding munc 18 allows for the t- SNARE complex to form (Deak et al., 2009; Rickman et al., 2010). Munc 13 is proposed to stabilize the conformation of the munch18-t-SNARE complex, further promoting the formation of the core complex with synaptobrevin and thereby keeping the vesicle docked. Synapses in cultured brain slices prepared from mice lacking both munc 13 isoforms show an insignificant amount of vesicles contacting the plasma membrane (Richmond et al., 2001; Siksou et al., 2009b). When the vesicles fuse to the plasma membrane the SNARE complex form a cis complex, i.e. all the components are located on the same membrane. Once assembled the complex needs to be unfolded for recycling, this is mediated by the ATPase NSF together with the adaptor protein SNAP in an ATP dependent process (Sollner et al., 1993a).

Synaptotagmin The family of synaptotagmins is made up by at least 16 different mammalian isoforms (Craxton, 2007). The founder of the family, synaptotagmin 1 which is also the most well studied isoform was discovered by Matthew et al in 1981 when characterizing monoclonal antibodies from mice immunized against total synaptic densities (Matthew et al., 1981). The synaptotagmins are expressed in both

76 neuronal and non-neuronal cells and the different isoforms mostly localize to non-overlapping cellular compartments (Adolfsen et al., 2004). All of the 16 isoforms are expressed in brain (Mittelsteadt et al., 2009) and synaptotagmin 1 and 2 are localized to synaptic vesicles. The synaptotagmins have a short intravesicular N-terminal region, a single helix transmembrane (TM) domain, a region juxtaposed to the TM- domain rich in lysines and arginines and two homologus C2-domains termed C2A and C2B (fig. 6). Synaptotagmin 1 is the isoform studied in this thesis, this isoform works as a mediator for exocytosis in both inhibitory and excitatory synapses throughout the CNS (Maximov and Sudhof, 2005; Nishiki and Augustine, 2004b).

Fig 6. Cartoon of syt 1 inserted in a vesicle membrane, the polybasic sequences of the C2 domains are labeled in red, the red dots indicate Ca2+ (Chapman 2008 Annu Rev Biochem;77)

Expression The expression of synaptotagmin 1 (syt) coincides with the formation of the nervous system and it is expressed at high levels in the mature CNS. In cultured developing neurons syt localizes to the soma and axons and as the culture matures it is redistributed in to a punctuate pattern in neuritic processes (Littleton et al., 1993). This change in expression pattern overlaps with an increasing ability to induce evoked neurotransmitter release, which is high only in the mature culture and also coincides with synapse formation (Adolfsen et al., 2004; Basarsky et al., 1994; Littleton et al., 1993). Syt is present in synaptic vesicles, which undergo exocytosis in axons even before synapses are formed. This led to the hypothesis that syt and exocytosis are connected to axonal outgrowth and synapse formation by controlling membrane expansion. However, when axonal outgrowth was monitored in syt null Drosophila embryos, axogenesis and neuronal path finding were found to be unaffected (Littleton et al., 1995), nor was there any variation in the morphology or the number of boutons (Geppert et al., 1994). Since syt

77 expression does not seem to be crucial for neuronal development, it was put forward that syt’s early presence in these vesicles serve as a storage of components for rapid establishment of communication in the neuronal system (Kraszewski et al., 1995; Littleton et al., 1995). A number of knockout studies have been made to establish the function of syt. Mice homozygous for a syt disrupting mutation die within two days after being born because they are unable to feed from their mother. The pups are phenotypically normal with their brains having a normal number of synapses and also electron micrographs of boutons appear normal compared to the wild type control (Geppert et al., 1994). However, using cultured hippocampal neurons from these mice it was discovered that the fast synchronous Ca2+-dependent vesicle release, which occurs in less than 1 ms after calcium influx, was decreased while no change in asynchronous release was seen (which follows in the next few 100 ms). Consequently it was concluded that lack of syt affected total amount of exocytosis (Geppert et al., 1994). Yoshihara and Littelton, in a later knockout study on Drosophila, observed that evoked release was still occurring at the neuro-muscular junction but the release was completely asynchronous and it occurred at increased level (Yoshihara and Littleton, 2002). A similar result was seen in a study on cultured hippocampal neurons from syt knockout mice where the total level of exocytosis was unchanged but the rate of release was reduced and asynchronous (Nishiki and Augustine, 2004a; Nishiki and Augustine, 2004b). The loss of the fast component of neurotransmitter release of course has a major impact on nerve signaling. Consequently the Drosophila syt null mutants had severely decreased synaptic transmission and the motility of the larvae was dramatically reduced. An attempt to rescue this phenotype with syt 4 or syt 7 was unsuccessful and this lead to the conclusion that these functions are not general for synaptotagmins (Adolfsen et al., 2004). The major view now is that syt is essential for the Ca2+-induced fast synchronous vesicle release and that it has a role in preventing asynchronous release, i.e. evoked fusion still takes place in the absence of syt, but the identity of the Ca2+-sensor under these conditions is so far unclear (Xu et al., 2007).

Synaptotagmin C2-domains The C2-domains of syt have been shown to be important for most functions of syt and there is a conserved polybasic sequence in each of the C2 domains known to interact with a number of factors (Ca2+- channels, phospholipids, t-SNARES, SV2 and others). The C2 domains

78 of syt and several other synaptotagmins are calcium binding with each C2 domain of syt containing a Ca2+-binding pocket at one end and the highly polybasic sequence at the other. C2A, according to structural predictions binds three Ca2+-ions and C2B two (fig. 6)(Ubach et al., 1998) (Fernandez et al., 2001). The polybasic sequence of C2B is more pronounced than in C2A and it is known to bind both to membranes and the SNAREs, C2B also has two basic amino acids, R398, 399, on the opposite side of the Ca2+-binding domain shown to be important (Xue et al., 2008). Apart from promoting synchronous and preventing asynchronous evoked release, syt also has a function in inhibiting spontaneous release. In a reconstituted system, non-calcium bound syt inhibited vesicle fusion and upon Ca2+-binding, fusion was promoted (Tucker et al., 2004). When the polybasic sequence of C2A was mutated, spontaneous release was increased by a factor of 2.4. If the C2B-domain was removed and the C2A domain left intact there was less release than if syt was removed completely (Mace et al., 2009), hence the two C2-domains seem to have opposing roles with C2A mainly preventing spontaneous release and C2B working as the Ca2+-sensor (Mackler et al., 2002; Yoshihara and Littleton, 2002). Syt is involved in priming of vesicles and the C2B- domain is also responsible for this function of syt. A mutation in the C2B polybasic domain not affecting Ca2+-binding causes delay and slower kinetics of the synchronous vesicle release and the size of the ready releasable pool is also decreased (Neher and Penner, 1994), hence syt has also Ca2+-independent functions. Exactly how the C2B dependent priming occurs has not been determined. It is known that syt functions via interactions with both the t-SNARES syntaxin and SNAP- 25 but not the v-SNARE synaptobrevin and it can interact with the SNARE complex during all stages of assembly (Zhang et al., 2002). When binding Ca2+, syt has been observed to bind both SNAREs and membranes simultaneously (Dai et al., 2007). By co-purification experiments it has been shown that SNARE interaction takes place also in the absence of Ca2+ and that both C2-domains are required but not if tyey interact with membranes concurrently to the SNAREs (Rickman et al., 2010). Binding of Ca2+ causes the C2 domains to have a more positive electrostatic potential and a stronger attraction to membranes containing anionic phospholipids. When all of the calcium binding residues in the C2A domain were mutated no effect on neurotransmitter release was

79 seen and interestingly there was actually an increase in release for one of the mutants; D232N. The result is surprising since all the Ca2+-binding residues of the C2A domain are conserved and therefore are expected to be important (Stevens and Sullivan, 2003), probably the Ca2+-binding of C2A has a fine-tuning role of neurotransmitter release. For the C2B domain the picture is clearer, deletion of several of the Ca2+-binding residues has been shown to lead to a disrupted function of syt as a Ca2+- sensor. When trying to rescue neurotransmitter release with syt mutants incompetent in Ca2+-binding, it was seen that syt’s ability to reduce asynchronous release was intact but levels for synchronous release were decreased (Mackler et al., 2002; Nishiki and Augustine, 2004a). The inhibition of asynchronous release is hence dependent on the C2B- domain but not dependent on its Ca2+-binding ability. In a study abolishing the Ca2+-sensing of the C2B in Drosophila and rat by a mutation corresponding to Y311N in syt from rat, it was concluded that vesicle release was impaired in a stage after docking and that SNARE assembly was decreased (Littleton et al., 2001). Why the C2A- and C2B- domains have different roles in Ca2+-sensing is not known but it is reasonable to think that interactions with SNAREs and/or the phospholipids phosphatidyl serine and PIP2 play a role. In the process of fusion pore dilation where the pore can either dilate, which is what happens at full vesicle fusion, or rapidly close again in a process termed “kiss-and-run”, the Ca2+ -sensing ability of the C2A- and C2B-domains have a cooperative function. If the C2B-domain is lacking Ca2+-sensing ability only “kiss-and-run” events will take place, which is the process for which the C2A-domain is the most important component (Wang et al., 2006a). Complexin has been shown to be an interaction partner for syt and it also has a role in the regulation of SNARE dependent fusion (Weninger et al., 2008). Complexin was thought to act as a clamp to stabilize the SNARE complex in its fusion competent state while preventing fusion with syt by competing for SNARE binding in a Ca2+-dependent manner (Tang et al., 2006). In a more recent study it has been concluded that complexin is only working as a clamp in the absence of neural activity and promotes vesicle release at a depolarization signal (Martin et al., 2011). It was suggested that this happens together with syt, since knocking out either protein prevents fast synchronous vesicle release (Martin et al., 2011; Maximov et al., 2009; Sudhof and Rothman, 2009). More research is needed to unravel how this works.

80

The fusion event In a recent model for syt’s role during vesicle fusion with the presynaptic membrane (Kuo et al., 2011), the C2-domains are proposed to bind the two opposing membranes and thereby bring them closer together. This model was adapted from data obtained using site-directed spin labeling (SDSL) on PC:PS:PIP2 bilayers. The C2B-domain plays the major role, where the polybasic sequence has affinity for membranes even in the absence of Ca2+ (Bai et al., 2004). The C2B is directed towards the plasma membrane because it has been shown to interact stronger with PIP2 containing membranes (Kuo et al., 2009; Li et al., 2006), and PIP2 is present at higher concentrations in the plasma membrane than in the synaptic vesicle (James et al., 2008). An interaction with PIP2 changes the orientation of the C2B so that it penetrates deeper into the bilayer and according to the model it seems probable that this domain then works as a bridge between the plasma membrane and the vesicle using the R398, 399 opposite of the Ca2+-binding loops to interact with the vesicular membrane (Kuo et al., 2011). Since the model proposes that the C2-domains are associated with opposing membranes, and the C2B is inserted in the plasma membrane, the C2A domain must be inserted in the vesicular membrane, at least after the Ca2+-signal (Murray and Honig, 2002). Ca2+ triggers the insertion of both C2-domains into membranes, it increases the C2B affinity for membranes 20 fold and the C2A-domain might actually not interact with membranes at all unless it has bound Ca2+ (Rufener et al., 2005). Collectively this Ca2+-dependent insertion and the change in orientation of C2B upon PIP2 binding could lead to a closer positioning of the two opposing bilayers (fig. 7) (Kuo et al., 2011). This model correlates with the observation that high PIP2 levels have been seen to be required in the target membrane for optimal Ca2+- sensitivity (Lee et al., 2010). Since the C2B-domain is highly positively charged it acts to neutralize the negative charges on the membranes making it easier for the membranes to come together, a theory especially interesting in view of paper III, where the polybasic sequence juxtaposed to the transmembrane helix could act to finalize the process of fusing the membranes. In this context it is conceivable that syt also acts to trigger the assembly and zippering of the SNARE complex. An important factor to consider is that only the C2-domains are included in these studies, which could lead to misinterpretations when making models to fit the physiological system. Since buckling of the plasma membrane would be a mechanism to overcome the electrostatic repulsion and an energetically favorable way

81 to promote membrane fusion it has been proposed as a possible step in vesicle fusion. The buckling would bring a piece of the plasma membrane closer to the vesicle, creating tension and a smaller contact area which would facilitate fusion. Syt has been discovered to be able to tubulate membranes in the presence of SNAREs and maybe could have a function in altering membrane curvature (Arac et al., 2006; Martens et al., 2007). The two tandem C2-domains C2A-C2B in solution causes tubulation and fusion of membrane vesicles and they also have a preference for binding curved membranes (Hui et al., 2009). C2B by itself can tubulate membranes in a Ca2+ dependent process which further supports the model by Kuo et al where C2B bridges between membranes (Hui et al., 2009; Kuo et al., 2011). Furthermore, tubulation is inhibited by a C2A-C2B construct where C2A is unable to bind Ca2+ and a mutation in the C2B-domain abolished the vesicle fusion when the membranes were rather flat, while vesicles that were smaller i.e. more curved still fused. Targeted to presynaptic boutons of syt knockout neurons, the C2A-C2B could rescue Ca2+-dependent vesicle release. The tubulation deficient mutant was incompetent in this matter but addition of the BAR domain from endophilin, another factor that can buckle membranes, enabled the mutated C2A-C2B to facilitate release. Endophilin is normally involved in endocytosis hence from this observation it becomes obvious that endo- and exocytosis share similar intermediate mechanisms (Hui et al., 2009). Syt is involved in compensatory endocytosis by serving as a docking site for an adaptor complex for clathrin mediated endocyotosis called AP2. The role for syt in endocytosis was shown using a photo-inactivatable syt, where a decrease in endocytosis was seen after inactivation. (Fergestad and Broadie, 2001; Poskanzer et al., 2003). Synaptic vesicle protein 2 (SV2) is involved in the regulation of syt endocytosis and if SV2 is absent there will be fewer vesicles competent for Ca2+-dependent fusion. SV2 binds to syt via its tyrosine based endocytosis motif YXXφ. This tyrosine motif is also involved in regulating AP2 binding to syt and thereby its recycling. In SV2 knockout mice the amount of syt on the plasma membrane is increased. Although there is around 50% less syt in these neurons, a reduction that was seen to be independent of the SV2 endocytosis motif and could perhaps be a result of SV2 regulating the stability of the syt protein (Yao et al., 2010). The Drosophila protein stoned is another protein shown to be involved in plasma membrane retrieval of syt. It also has an YXXφ motif mediating the interaction between AP2 and syt, but stoned in addition contains other AP2 and

82 clathrin binding motifs (Fergestad and Broadie, 2001). N-terminal palmitoylations are required for endocytosis of syt, if these are absent syt localizes to the plasma membrane and is not recycled back. These palmitoylations are of stabile kind and have been shown to have a longer turnover time than six hours (Kang et al., 2004). Levels of syt is possibly also regulated via its mRNA, however not much is known so far. The C2A-domain of syt has been shown to bind the 3’-UTR of its own transcript and for in vitro translation using a reticulocyte lysate and a cDNA containing the 3’-UTR resulted in 50% reduced translation compared to that of the cDNA lacking the 3’-UTR (Sukumaran et al., 2008). Syt has multiple roles in the presynapse such as priming, clamping vesicle fusion in the absence of Ca2+ and mediating fusion when binding Ca2+ and vesicle recycling are so fare functions that have been revealed. Furthermore, syt could also have a role in inducing actin polymerization, see paper III. To conclude, only six proteins form the core of the Ca2+- triggered exocytosis apparatus; the three SNAREs, Munch 18, syt and complexin. The synaptic vesicle is docked in a process dependent on syt with the SNAREs unassembled. Aided by Munch 18 they are then assembled in the first stage of priming. From this stage the vesicle is competent only for asynchronous release. In a further priming step complexin freezes in an even more release-ready state called priming stage II. The polybasic region of the syt C2B domain interacts with the plasma membrane, but when Ca2+ then binds to syt it changes the binding of the C2B-domain to the plasma membrane causing this domain to rotate slightly. The polybasic sequence might interact with the SNAREs and R398 and 399 opposite the Ca2+-binding loops interacts 2+ with the membrane of the vesicle. The C2A is when binding Ca , inserted into the vesicular membrane and perhaps the polybasic sequence characterized in paper III, which is juxtaposed to the transmembrane domain, enhances the C2A-membrane interaction by increasing the concentration of anionic lipids through electrostatic clustering. The C2 interactions position the fusing membranes into closer proximity (fig. 7). The C2-domains also cause buckling of the plasma membrane which aids in the formation of the fusion stalk. The fusion pore is opened mostly by the C2B interactions. The polybasic sequence of syt positioned directly after the transmembrane sequence is perhaps involved in neutralizing the negative repulsion of lipids thereby bridging between the membranes at the last stage of fusion or it could be involved in the opening of the fusion pore. The C2B-domain is then

83 involved in recruiting AP2 for clathrin mediated endocytosis. The endocytic process occurs in the periactive zone, if the membrane is transported here in an active process is unknown but we speculate that it may require actin polymerization in which case syt can be involved in driving this process, see paper III.

Fig 7. (A), Model of a synapse depicting the synaptic vesicle cycle. Vesicles are docked via multi component protein complexes to specific sites at the active zone. A rise in cytosolic calcium concentration triggers fusion of the docked vesicles, the membrane from the fused vesicle moves to the periactive zone while components for clathrin mediated endocytosis are being recruited. After the formation of a clathrin coat the synaptic vesicle, containing the proteins used for fusion, is pinched off by dynamin, transported, uncoated and refilled with neurotransmitter. The uncoating and transport have been shown to depend on actin dynamics (Pechstein et al.). (B) The vesicle is docked and primed for fusion by interactions of the SNAREs syntaxin, SNAP-25 and synaptobrevin. Entry of calcium triggers a conformational change of synaptotagmin so that it bridges between the plasma membrane and the synaptic vesicle and aids on the zippering of the SNAREs allowing for fusion. The C2B domain aids in opening of the fusion pore and the vesicle integrates into the plasma membrane. The arrow labeled “polybasic” indicates the position of the polybasic sequence identified in paper III. Model was adopted from (Kuo et al., 2011).

84

INVESTIGATIONS

*For reference to the literature in this section the reader is referred to the individual manuscripts and relevant sections in the introduction

Aim In these studies of the microfilament system and the control of actin polymerization in particular, I have specifically aimed - to determine whether profilin mRNA distributes in a specific pattern in motility-stimulated cells (Paper I) - to determine the effect of profilin depletion on transcriptional serum response factor (SRF) control and cell motility of cells, considering also the profilin isoform variation (Paper II) - to determine the mechanism behind the ability of synaptotagmin 1 to induce actin polymerization when expressed in non-neuronal cells and to relate this activity to its role in neurotransmitter release (Paper III).

Paper I: Microtubule-dependent Localization of Profilin I mRNA to Actin Polymerization Sites in Serum-stimulated Cells Localization of mRNAs is a control mechanism cells use to enrich a region of the cell with newly translated protein. This could be a way of transporting fewer molecules, to avoid toxic effects of a certain protein ending up in the wrong location, to coordinate a rapid supply of protein with an increased demand and to ensure that subunits of a complex end up in close proximity to each other. This field is not so well explored but one of the most studied localized transcripts is the β-actin mRNA. This mRNA is known to localize in a myosin dependent process in fibroblasts where it localizes to active edges to support persistent migration. The hypothesis is that for persistent migration which of course is supported by actin polymerization, the cell will need freshly supplied actin monomers. As a lot of actin monomers would be present at a site of

85 localized translation, one can assume that actin regulatory proteins must also be present to inhibit spontaneous formation of filaments. In fact it has been shown that the mRNAs for all the components of the Arp2/3 complex are localized to active edges. Profilin which binds actin monomers and inhibits spontaneous nucleation but also permits elongation at the filament (+)-end should possibly also be required to buffer the newly translated actin. The aim of the studies in paper I was to examine if the mRNA for profilin localizes in a similar way as β-actin mRNA. The method used to probe for the mRNAs was florescent in situ hybridization. This is a very sensitive method since it amplifies the signal many times by a chemical reaction which deposits a fluorescent probe. This amplification facilitates the qualitative determination of how the mRNAs localize. The amplification is however also a disadvantage because the deposit only ends up in close proximity to the mRNA but maybe not exactly on the mRNA, so it is not suitable for colocalization studies. Another disadvantage is that a high hybridization temperature is used which partly destroys many cellular structures. In this case the high temperature was especially unfortunate because actin filaments do not preserve well at high temperatures. Despite this fact it was possible to use phallodin to identify the cell perimeter and to get a view of the general morphology of the cells, clearly in this situation it was not filamentous actin that this toxin bound to but if it was short pieces of actin or something else we do not know. For this study a mouse embryonic fibroblast cell line was used. After serum stimulation these cells exhibited a high degree of dorsal ruffle formation, which are sites of massive actin polymerization. These ruffles were used to study the localization of β-actin mRNA and profilin mRNA respectively and also co-hybridization with the two probes were used. From these experiments it was concluded that the mRNAs for profilin and actin co-distribute to dorsal ruffles formed after serum stimulation. However when drugs were used to disrupt the microfilament system and the microtubule system respectively it was seen that the localization of the profilin and actin mRNAs to ruffles depend on different mechanisms. The β-actin mRNA localization is dependent on the microfilament system, which concurs with previous studies while profilin mRNA localization is dependent on the microtubule system. These conclusions opens up for further investigations to learn if and how these transport processes are coordinated. Since actin requires the

86 chaperonin CCT to attain its native fold, one possibility is that the localization of the profilin mRNA correlates with the localization of the CCT. Another continuation would be to study the transport of the profilin mRNA in live cells. The mRNA of profilin has been fused to tandem repeats of bacteriophage R17/MS2 coat protein binding sites which are RNA hairpin structures. This mRNA construct can be co- expressed together with the MS2 coat protein fused to a suitable tag. In this setup GFP is fused to the NLS of SV40 so that the GFP signal will not disturb imaging in the cytoplasm. The MS2 system is also suitable for affinity purification of mRNP complexes and can be used to characterize factors binding to the profilin mRNA.

Paper II: Profilin I and II are Both Influencing SRF- dependent Signaling in B16 Melanoma Cells and Loss of Profilin I Interferes with Cell Migration Many genes encoding proteins involved in cell growth are controlled by the transcription factor serum response factor (SRF). For instance, the expression of many proteins connected to the microfilament system are controlled by SRF, including actin and profilin. The microfilament system is connected to SRF transcription via a feed-back mechanism involving megakaryocyte acute leukemia MAL, a cofactor for SRF. This cofactor works in parallel to the MAP kinase regulated cofactor TCF. MAL localizes to the cell periphery where it competes with proteins such as profilin and cofilin in binding to monomeric actin. Binding to actin inhibits MAL form working as an SRF cofactor and if cells are exposed stimuli to that induces cell motility, the levels of monomeric actin will drop and MAL will be released and enter the nucleus. Hence the nuclear MAL/SRF activity is dependent on the actin dynamics in the cytoplasm. Formally it has not been shown that profilin competes with MAL for actin in vivo. This was investigated in paper II, where RNA interference was used to deplete profilin 1 and 2 respectively from B16 mouse melanoma cells. The SRF activity was examined after depletion using an SRF dependent reporter assay. It was determined that SRF activity was drastically downregulated after depletion of either profilin isoform, with a stronger effect seen after profilin 1 depletion. Simultaneous depletion of both isoforms had a slight synergistic effect. Since at least profilin 1 is also controlled by SRF it was interesting to examine the expression levels for the profilin isoform which was not depleted. For this purpose

87 protein levels using western blot were examined. This showed that levels for the profilin isoform which was not targeted also dropped, albeit to a lesser extent than the targeted isoform. This drop was most pronounced after 24 hours after siRNA treatment. Also the migratory properties of the cells were examined. For this purpose live cell imaging was performed. From these experiments it was concluded that siRNA depletion of profilin 1 or both profilin 1 and 2 had a small but significant effect on migration. The cells moved at a slightly slower speed, with approximately the same result for either knocking down of profilin 1 or depleting both isoforms. In addition to influencing the migration speed negatively, the results also showed that profilin depletion had a positive influence on directionality. It is important to note that B16 cells are melanoma cells stimulated via an autocrine loop which is likely to keep up constant high transcription through pathways not involving SRF. Recent results, not included in the manuscript indicate that the profilin isoform 2b could be upregulated after knock down of the 2a isoform. The 2b isoform is generally referred to as being expressed only in kidney, though it may also occur in cells of other origins (DiNardo 2000). This isofom is interesting in that it has aberrant actin binding ability, which means that upregulation of this protein could work as a dominant negative component in respect to other profilin partners. This isoform also has been shown to bind tubulin. The expression levels for the different isoforms in different tissues are not at all well characterized and when it comes to cell lines, data is virtually lacking. In addition, basically nothing is known how the expression levels of the different isoforms affect each other. This study will continue in exploring these aspects, and ultimately the effects of the different isoform on the areas discussed above, i.e. SRF-regulation and actin dynamics, will be known.

88

Paper III: Synaptotagmin 1 causes Phosphatidyl- (4,5)-bisphosphate-dependent Actin Remodeling in Cultured Non-neuronal and Neuronal Cells The study in paper III involves the protein synaptotagmin 1 which works as a calcium sensor in neurotransmitter release. This protein is inserted via a transmembrane domain in synaptic vesicles and interacts with proteins and lipids of the plasma membrane of the presynapse. This study was initiated on the grounds of a previous study where synaptotagmin was seen to cause filopodia formation when ectopically expressed. Initially the intention was to use the formed filopodia to study transport of profilin:actin, and other components used in polymerization, see also page 51. However, in this study we investigated how actin polymerization used for formation of filopodia is initiated by synaptotagmin. A number of truncation constructs of synaptotagmin were designed and used to determine what region(s) of the protein is required to induce filopodia formation. This led to identification of a polybasic stretch of amino acids juxtaposed the transmembrane region solely responsible for the activity. By a number of mutations it was concluded that the electrostatic nature of this sequence causes filopodia formation. Other proteins with such a polybasic motif are known to affect the microfilament system via PIP2. Using a phosphatase that could be induced to target to the plasma membrane it was concluded that PIP2 is involved also in this case. However, exactly what actin regulatory proteins that are involved was not possible to establish. When expressing a construct including the transmembrane region and the polybasic sequence, referred to as Syt1-96, in fibroblasts that did not express N- WASP, filopodia still formed. When simultaneously depleting these cells from the formins Dia1 and 2 and VASP only 30% reduction in filopodia formation was seen, suggesting sufficient contribution from other components.

When Syt1-96 was expressed in the neuroblastoma cell line SH-SY5Y or in primary embryonic cortex cells an increase in the length and number of processes was seen. These results of course open up for the possibility that this polybasic sequence is important for the function of syt also in its natural context in nerve cells. Obviously the polybasic sequence could have a function in the already well explored context of Ca2+-induced

89 rapid and synchronous neurotransmitter release, where this sequence could facilitate the fusion of the two membranes involved. Another possibility, albeit more controversial is that synaptotagmin could have a role in causing actin polymerization at the synapse. Actually not much is known about the role for actin polymerization during neurotransmitter release, making such a role for syt an interesting possibility for further study. A continuing study would involve expression of a synaptotagmin with a mutated polybasic sequence in a model organism like Drosophila and a close examination on how this affects the synaptic cycle and neuronal development. Notably the use of syt and its polybasic sequence for studies of actin remodeling at the cell edge should not be overseen.

90

ACKNOWLEDGEMENTS

I wish to thank

Roger Karlsson, my supervisor, for a fun collaboration and inspiring science discussions. Past and present colleagues and friends at the Department of Cell Biology for being so nice to work with, especially Sara, Deike, Yu, Ebba and Julie for all the fun talks. Anna-Stina Höglund for guiding me in different microscopy techniques. Maria Eriksson for giving me cells and Ingrid Lassing for help with protein assays. Joel Schick for inviting me to his lab at the Helmholtz Institute in Munich to learn new techniques. A special thanks to my dear boyfriend Ragnar, for all help with computer stuff and for putting up with everything and to my mother and sister for always being there. My Uppsala “study”-buddies Disa, Emma, Sofia and Anna for all encouragements. Thank you to the family of Pokora Kulinska and the Sture Eriksson fund for cancer and heart research, for the scholarship is received.

The work in this thesis was financially supported by grants from the Nancy Lurie Marks Family foundation, Carl Trygger Foundation and the Magnus Bergvall Foundation to Roger Karlsson.

91

REFERENCES

Abercrombie, M., J.E. Heaysman, and S.M. Pegrum. 1970. The locomotion of fibroblasts in culture. II. "RRuffling". Exp Cell Res. 60:437-44. Ackermann, M., and A. Matus. 2003. Activity-induced targeting of profilin and stabilization of dendritic spine morphology. Nat Neurosci. 6:1194-200. Adams, R.H., and A. Eichmann. 2010. Axon guidance molecules in vascular patterning. Cold Spring Harb Perspect Biol. 2:a001875. Adereth, Y., V. Dammai, N. Kose, R. Li, and T. Hsu. 2005. RNA-dependent integrin alpha3 protein localization regulated by the Muscleblind-like protein MLP1. Nat Cell Biol. 7:1240-7. Adolfsen, B., S. Saraswati, M. Yoshihara, and J.T. Littleton. 2004. Synaptotagmins are trafficked to distinct subcellular domains including the postsynaptic compartment. J Cell Biol. 166:249-60. Ahmari, S.E., J. Buchanan, and S.J. Smith. 2000. Assembly of presynaptic active zones from cytoplasmic transport packets. Nat Neurosci. 3:445-51. Ahmed, S., W.I. Goh, and W. Bu. 2010. I-BAR domains, IRSp53 and filopodium formation. Semin Cell Dev Biol. 21:350-6. Ahuja, R., R. Pinyol, N. Reichenbach, L. Custer, J. Klingensmith, M.M. Kessels, and B. Qualmann. 2007. Cordon-bleu is an actin nucleation factor and controls neuronal morphology. Cell. 131:337-50. Alexandrova, A.Y., K. Arnold, S. Schaub, J.M. Vasiliev, J.J. Meister, A.D. Bershadsky, and A.B. Verkhovsky. 2008. Comparative dynamics of retrograde actin flow and focal adhesions: formation of nascent adhesions triggers transition from fast to slow flow. PLoS One. 3:e3234. Allison, D.W., A.S. Chervin, V.I. Gelfand, and A.M. Craig. 2000. Postsynaptic scaffolds of excitatory and inhibitory synapses in hippocampal neurons: maintenance of core components independent of actin filaments and microtubules. J Neurosci. 20:4545-54. An, J.J., K. Gharami, G.Y. Liao, N.H. Woo, A.G. Lau, F. Vanevski, E.R. Torre, K.R. Jones, Y. Feng, B. Lu, and B. Xu. 2008. Distinct role of long 3' UTR BDNF mRNA in spine morphology and synaptic plasticity in hippocampal neurons. Cell. 134:175-87. Andrianantoandro, E., and T.D. Pollard. 2006. Mechanism of actin filament turnover by severing and nucleation at different concentrations of ADF/cofilin. Mol Cell. 24:13-23. Anthis, N.J., and I.D. Campbell. 2011. The tail of integrin activation. Trends Biochem Sci. 36:191-8. Arac, D., X. Chen, H.A. Khant, J. Ubach, S.J. Ludtke, M. Kikkawa, A.E. Johnson, W. Chiu, T.C. Sudhof, and J. Rizo. 2006. Close membrane-membrane proximity induced by Ca(2+)-dependent multivalent binding of synaptotagmin-1 to phospholipids. Nat Struct Mol Biol. 13:209-17. Arakawa, Y., H. Bito, T. Furuyashiki, T. Tsuji, S. Takemoto-Kimura, K. Kimura, K. Nozaki, N. Hashimoto, and S. Narumiya. 2003. Control of axon elongation via an SDF-1alpha/Rho/mDia pathway in cultured cerebellar granule neurons. J Cell Biol. 161:381-91.

92

Aspenstrom, P., A. Fransson, and J. Saras. 2004. Rho GTPases have diverse effects on the organization of the actin filament system. Biochem J. 377:327-37. Bader, M.F., F. Doussau, S. Chasserot-Golaz, N. Vitale, and S. Gasman. 2004. Coupling actin and membrane dynamics during calcium-regulated exocytosis: a role for Rho and ARF GTPases. Biochim Biophys Acta. 1742:37-49. Bae, Y.H., Z. Ding, L. Zou, A. Wells, F. Gertler, and P. Roy. 2009. Loss of profilin-1 expression enhances breast cancer cell motility by Ena/VASP proteins. J Cell Physiol. 219:354-64. Bai, J., W.C. Tucker, and E.R. Chapman. 2004. PIP2 increases the speed of response of synaptotagmin and steers its membrane-penetration activity toward the plasma membrane. Nat Struct Mol Biol. 11:36-44. Barzik, M., T.I. Kotova, H.N. Higgs, L. Hazelwood, D. Hanein, F.B. Gertler, and D.A. Schafer. 2005. Ena/VASP proteins enhance actin polymerization in the presence of barbed end capping proteins. J Biol Chem. 280:28653-62. Basarsky, T.A., V. Parpura, and P.G. Haydon. 1994. Hippocampal synaptogenesis in cell culture: developmental time course of synapse formation, calcium influx, and synaptic protein distribution. J Neurosci. 14:6402-11. Bear, J.E., and F.B. Gertler. 2009. Ena/VASP: towards resolving a pointed controversy at the barbed end. J Cell Sci. 122:1947-53. Bear, J.E., M. Krause, and F.B. Gertler. 2001. Regulating cellular actin assembly. Curr Opin Cell Biol. 13:158-66. Beli, P., D. Mascheroni, D. Xu, and M. Innocenti. 2008. WAVE and Arp2/3 jointly inhibit filopodium formation by entering into a complex with mDia2. Nat Cell Biol. 10:849-57. Belyantseva, I.A., B.J. Perrin, K.J. Sonnemann, M. Zhu, R. Stepanyan, J. McGee, G.I. Frolenkov, E.J. Walsh, K.H. Friderici, T.B. Friedman, and J.M. Ervasti. 2009. Gamma-actin is required for cytoskeletal maintenance but not development. Proc Natl Acad Sci U S A. 106:9703-8. Beningo, K.A., M. Dembo, I. Kaverina, J.V. Small, and Y.L. Wang. 2001. Nascent focal adhesions are responsible for the generation of strong propulsive forces in migrating fibroblasts. J Cell Biol. 153:881-8. Beningo, K.A., K. Hamao, M. Dembo, Y.L. Wang, and H. Hosoya. 2006. Traction forces of fibroblasts are regulated by the Rho-dependent kinase but not by the myosin light chain kinase. Arch Biochem Biophys. 456:224-31. Berg, J.S., and R.E. Cheney. 2002. Myosin-X is an unconventional myosin that undergoes intrafilopodial motility. Nat Cell Biol. 4:246-50. Berridge, M.J. 2006. Calcium microdomains: organization and function. Cell Calcium. 40:405-12. Bhatt, A., I. Kaverina, C. Otey, and A. Huttenlocher. 2002. Regulation of focal complex composition and disassembly by the calcium-dependent protease calpain. J Cell Sci. 115:3415-25. Birbach, A., J.M. Verkuyl, and A. Matus. 2006. Reversible, activity-dependent targeting of profilin to neuronal nuclei. Exp Cell Res. 312:2279-87. Bjorkegren-Sjogren, C., E. Korenbaum, P. Nordberg, U. Lindberg, and R. Karlsson. 1997. Isolation and characterization of two mutants of human profilin I that do not bind poly(L-proline). FEBS Lett. 418:258-64. Blanchoin, L., and T.D. Pollard. 1998. Interaction of actin monomers with Acanthamoeba actophorin (ADF/cofilin) and profilin. J Biol Chem. 273:25106-11.

93

Blessing, C.A., G.T. Ugrinova, and H.V. Goodson. 2004. Actin and ARPs: action in the nucleus. Trends Cell Biol. 14:435-42. Block, J., T.E. Stradal, J. Hanisch, R. Geffers, S.A. Kostler, E. Urban, J.V. Small, K. Rottner, and J. Faix. 2008. Filopodia formation induced by active mDia2/Drf3. J Microsc. 231:506-17. Bloom, O., E. Evergren, N. Tomilin, O. Kjaerulff, P. Low, L. Brodin, V.A. Pieribone, P. Greengard, and O. Shupliakov. 2003. Colocalization of synapsin and actin during synaptic vesicle recycling. J Cell Biol. 161:737-47. Bohil, A.B., B.W. Robertson, and R.E. Cheney. 2006. Myosin-X is a molecular motor that functions in filopodia formation. Proc Natl Acad Sci U S A. 103:12411-6. Bosch, M., K.H. Le, B. Bugyi, J.J. Correia, L. Renault, and M.F. Carlier. 2007. Analysis of the function of Spire in actin assembly and its synergy with formin and profilin. Mol Cell. 28:555-68. Bramham, C.R. 2008. Local protein synthesis, actin dynamics, and LTP consolidation. Curr Opin Neurobiol. 18:524-31. Brandt, D.T., and R. Grosse. 2007. Get to grips: steering local actin dynamics with IQGAPs. EMBO Rep. 8:1019-23. Breitsprecher, D., A.K. Kiesewetter, J. Linkner, C. Urbanke, G.P. Resch, J.V. Small, and J. Faix. 2008. Clustering of VASP actively drives processive, WH2 domain- mediated actin filament elongation. EMBO J. 27:2943-54. Breitsprecher, D., A.K. Kiesewetter, J. Linkner, M. Vinzenz, T.E. Stradal, J.V. Small, U. Curth, R.B. Dickinson, and J. Faix. 2011. Molecular mechanism of Ena/VASP- mediated actin-filament elongation. EMBO J. 30:456-67. Bridgman, P.C. 1999. Myosin Va movements in normal and dilute-lethal axons provide support for a dual filament motor complex. J Cell Biol. 146:1045-60. Broussard, J.A., D.J. Webb, and I. Kaverina. 2008. Asymmetric focal adhesion disassembly in motile cells. Curr Opin Cell Biol. 20:85-90. Bruses, J.L. 2000. Cadherin-mediated adhesion at the interneuronal synapse. Curr Opin Cell Biol. 12:593-7. Bubb, M.R., I.C. Baines, and E.D. Korn. 1998. Localization of actobindin, profilin I, profilin II, and phosphatidylinositol-4,5-bisphosphate (PIP2) in Acanthamoeba castellanii. Cell Motil Cytoskeleton. 39:134-46. Buccione, R., J.D. Orth, and M.A. McNiven. 2004. Foot and mouth: podosomes, invadopodia and circular dorsal ruffles. Nat Rev Mol Cell Biol. 5:647-57. Buchanan, J., Y.A. Sun, and M.M. Poo. 1989. Studies of nerve-muscle interactions in Xenopus cell culture: fine structure of early functional contacts. J Neurosci. 9:1540- 54. Bunnell, T.M., and J.M. Ervasti. Delayed embryonic development and impaired cell growth and survival in Actg1 null mice. Cytoskeleton (Hoboken). 67:564-72. Burridge, K. 2005. Foot in mouth: do focal adhesions disassemble by endocytosis? Nat Cell Biol. 7:545-7. Burry, R.W. 1986. Presynaptic elements on artificial surfaces. A model for the study of development and regeneration of synapses. Neurochem Pathol. 5:345-60. Buss, F., and B.M. Jockusch. 1989. Tissue-specific expression of profilin. FEBS Lett. 249:31-4. Buss, F., C. Temm-Grove, S. Henning, and B.M. Jockusch. 1992. Distribution of profilin in fibroblasts correlates with the presence of highly dynamic actin filaments. Cell Motil Cytoskeleton. 22:51-61.

94

Campbell, I.D., and M.J. Humphries. 2011. Integrin structure, activation, and interactions. Cold Spring Harb Perspect Biol. 3. Campellone, K.G., N.J. Webb, E.A. Znameroski, and M.D. Welch. 2008. WHAMM is an Arp2/3 complex activator that binds microtubules and functions in ER to Golgi transport. Cell. 134:148-61. Cantley, L.C. 2002. The phosphoinositide 3-kinase pathway. Science. 296:1655-7. Carlier, M.F., V. Laurent, J. Santolini, R. Melki, D. Didry, G.X. Xia, Y. Hong, N.H. Chua, and D. Pantaloni. 1997. Actin depolymerizing factor (ADF/cofilin) enhances the rate of filament turnover: implication in actin-based motility. J Cell Biol. 136:1307-22. Carlier, M.F., D. Pantaloni, and E.D. Korn. 1986. The effects of Mg2+ at the high- affinity and low-affinity sites on the polymerization of actin and associated ATP hydrolysis. J Biol Chem. 261:10785-92. Carlson, S.S., G. Valdez, and J.R. Sanes. 2010. Presynaptic calcium channels and alpha3-integrins are complexed with synaptic cleft laminins, cytoskeletal elements and active zone components. J Neurochem. 115:654-66. Carlsson, L., L.E. Nystrom, I. Sundkvist, F. Markey, and U. Lindberg. 1977. Actin polymerizability is influenced by profilin, a low molecular weight protein in non- muscle cells. J Mol Biol. 115:465-83. Caroni, P. 2001. New EMBO members' review: actin cytoskeleton regulation through modulation of PI(4,5)P(2) rafts. EMBO J. 20:4332-6. Castellano, F., C. Le Clainche, D. Patin, M.F. Carlier, and P. Chavrier. 2001. A WASp- VASP complex regulates actin polymerization at the plasma membrane. EMBO J. 20:5603-14. Cattaruzza, M., C. Lattrich, and M. Hecker. 2004. Focal adhesion protein zyxin is a mechanosensitive modulator of gene expression in vascular smooth muscle cells. Hypertension. 43:726-30. Catterall, W.A., and A.P. Few. 2008. Calcium channel regulation and presynaptic plasticity. Neuron. 59:882-901. Cedergren-Zeppezauer, E.S., N.C. Goonesekere, M.D. Rozycki, J.C. Myslik, Z. Dauter, U. Lindberg, and C.E. Schutt. 1994. Crystallization and structure determination of bovine profilin at 2.0 A resolution. J Mol Biol. 240:459-75. Chang, S., and P. De Camilli. 2001. Glutamate regulates actin-based motility in axonal filopodia. Nat Neurosci. 4:787-93. Cheever, T.R., E.A. Olson, and J.M. Ervasti. 2011. Axonal regeneration and neuronal function are preserved in motor neurons lacking ss-actin in vivo. PLoS One. 6:e17768. Chen, F., L. Ma, M.C. Parrini, X. Mao, M. Lopez, C. Wu, P.W. Marks, L. Davidson, D.J. Kwiatkowski, T. Kirchhausen, S.H. Orkin, F.S. Rosen, B.J. Mayer, M.W. Kirschner, and F.W. Alt. 2000. Cdc42 is required for PIP(2)-induced actin polymerization and early development but not for cell viability. Curr Biol. 10:758-65. Chereau, D., and R. Dominguez. 2006. Understanding the role of the G-actin-binding domain of Ena/VASP in actin assembly. J Struct Biol. 155:195-201. Chereau, D., F. Kerff, P. Graceffa, Z. Grabarek, K. Langsetmo, and R. Dominguez. 2005. Actin-bound structures of Wiskott-Aldrich syndrome protein (WASP)- homology domain 2 and the implications for filament assembly. Proc Natl Acad Sci U S A. 102:16644-9. Chesarone, M.A., A.G. DuPage, and B.L. Goode. 2010. Unleashing formins to remodel the actin and microtubule . Nat Rev Mol Cell Biol. 11:62-74.

95

Chiaruttini, C., M. Sonego, G. Baj, M. Simonato, and E. Tongiorgi. 2008. BDNF mRNA splice variants display activity-dependent targeting to distinct hippocampal laminae. Mol Cell Neurosci. 37:11-9. Chik, J.K., U. Lindberg, and C.E. Schutt. 1996. The structure of an open state of beta- actin at 2.65 A resolution. J Mol Biol. 263:607-23. Chiquet, M., A.S. Renedo, F. Huber, and M. Fluck. 2003. How do fibroblasts translate mechanical signals into changes in extracellular matrix production? Matrix Biol. 22:73-80. Choi, C.K., M. Vicente-Manzanares, J. Zareno, L.A. Whitmore, A. Mogilner, and A.R. Horwitz. 2008. Actin and alpha-actinin orchestrate the assembly and maturation of nascent adhesions in a myosin II motor-independent manner. Nat Cell Biol. 10:1039- 50. Choi, J., J. Ko, B. Racz, A. Burette, J.R. Lee, S. Kim, M. Na, H.W. Lee, K. Kim, R.J. Weinberg, and E. Kim. 2005. Regulation of dendritic spine morphogenesis by insulin receptor substrate 53, a downstream effector of Rac1 and Cdc42 small GTPases. J Neurosci. 25:869-79. Cingolani, L.A., and Y. Goda. 2008. Actin in action: the interplay between the actin cytoskeleton and synaptic efficacy. Nat Rev Neurosci. 9:344-56. Cockcroft, S., and M.A. De Matteis. 2001. Inositol lipids as spatial regulators of membrane traffic. J Membr Biol. 180:187-94. Condeelis, J., and R.H. Singer. 2005. How and why does beta-actin mRNA target? Biol Cell. 97:97-110. Cooke, R., and L. Murdoch. 1973. Interaction of actin with analogs of . . 12:3927-32. Cory, G.O., R. Garg, R. Cramer, and A.J. Ridley. 2002. Phosphorylation of tyrosine 291 enhances the ability of WASp to stimulate actin polymerization and filopodium formation. Wiskott-Aldrich Syndrome protein. J Biol Chem. 277:45115-21. Coutts, A.S., L. Weston, and N.B. La Thangue. 2009. A transcription co-factor integrates cell adhesion and motility with the p53 response. Proc Natl Acad Sci U S A. 106:19872-7. Craxton, M. 2007. Evolutionary genomics of plant genes encoding N-terminal-TM-C2 domain proteins and the similar FAM62 genes and synaptotagmin genes of metazoans. BMC Genomics. 8:259. Czuchra, A., X. Wu, H. Meyer, J. van Hengel, T. Schroeder, R. Geffers, K. Rottner, and C. Brakebusch. 2005. Cdc42 is not essential for filopodium formation, directed migration, cell polarization, and mitosis in fibroblastoid cells. Mol Biol Cell. 16:4473- 84. da Silva, J.S., and C.G. Dotti. 2002. Breaking the neuronal sphere: regulation of the actin cytoskeleton in neuritogenesis. Nat Rev Neurosci. 3:694-704. Dai, H., N. Shen, D. Arac, and J. Rizo. 2007. A quaternary SNARE-synaptotagmin- Ca2+-phospholipid complex in neurotransmitter release. J Mol Biol. 367:848-63. Dantzig, J.A., T.Y. Liu, and Y.E. Goldman. 2006. Functional studies of individual myosin molecules. Ann N Y Acad Sci. 1080:1-18. De Corte, V., J. Gettemans, and J. Vandekerckhove. 1997. Phosphatidylinositol 4,5- bisphosphate specifically stimulates PP60(c-src) catalyzed phosphorylation of gelsolin and related actin-binding proteins. FEBS Lett. 401:191-6. de Hoog, C.L., L.J. Foster, and M. Mann. 2004. RNA and RNA binding proteins participate in early stages of cell spreading through spreading initiation centers. Cell. 117:649-62.

96

Deak, F., Y. Xu, W.P. Chang, I. Dulubova, M. Khvotchev, X. Liu, T.C. Sudhof, and J. Rizo. 2009. Munc18-1 binding to the neuronal SNARE complex controls synaptic vesicle priming. J Cell Biol. 184:751-64. del Rio, A., R. Perez-Jimenez, R. Liu, P. Roca-Cusachs, J.M. Fernandez, and M.P. Sheetz. 2009. Stretching single talin rod molecules activates vinculin binding. Science. 323:638-41. Dent, E.W., S.L. Gupton, and F.B. Gertler. 2010. The Growth Cone Cytoskeleton in Axon Outgrowth and Guidance. Cold Spring Harb Perspect Biol. Dent, E.W., A.V. Kwiatkowski, L.M. Mebane, U. Philippar, M. Barzik, D.A. Rubinson, S. Gupton, J.E. Van Veen, C. Furman, J. Zhang, A.S. Alberts, S. Mori, and F.B. Gertler. 2007. Filopodia are required for cortical neurite initiation. Nat Cell Biol. 9:1347-59. Derivery, E., and A. Gautreau. 2010. Generation of branched actin networks: assembly and regulation of the N-WASP and WAVE molecular machines. Bioessays. 32:119- 31. DerMardirossian, C., and G.M. Bokoch. 2005. GDIs: central regulatory molecules in Rho GTPase activation. Trends Cell Biol. 15:356-63. DesMarais, V., M. Ghosh, R. Eddy, and J. Condeelis. 2005. Cofilin takes the lead. J Cell Sci. 118:19-26. DesMarais, V., F. Macaluso, J. Condeelis, and M. Bailly. 2004. Synergistic interaction between the Arp2/3 complex and cofilin drives stimulated lamellipod extension. J Cell Sci. 117:3499-510. Dharmalingam, E., A. Haeckel, R. Pinyol, L. Schwintzer, D. Koch, M.M. Kessels, and B. Qualmann. 2009. F-BAR proteins of the syndapin family shape the plasma membrane and are crucial for neuromorphogenesis. J Neurosci. 29:13315-27. Di Nardo, A., R. Gareus, D. Kwiatkowski, and W. Witke. 2000. Alternative splicing of the mouse profilin II gene generates functionally different profilin isoforms. J Cell Sci. 113 Pt 21:3795-803. Ding, Z., D. Gau, B. Deasy, A. Wells, and P. Roy. 2009. Both actin and polyproline interactions of profilin-1 are required for migration, invasion and capillary morphogenesis of vascular endothelial cells. Exp Cell Res. 315:2963-73. Ding, Z., A. Lambrechts, M. Parepally, and P. Roy. 2006. Silencing profilin-1 inhibits endothelial cell proliferation, migration and cord morphogenesis. J Cell Sci. 119:4127-37. Dominguez, R., and K.C. Holmes. 2010. Actin Structure And Function. Annu Rev Biophys. Dong, J., B. Radau, A. Otto, E. Muller, C. Lindschau, and P. Westermann. 2000. Profilin I attached to the Golgi is required for the formation of constitutive transport vesicles at the trans-Golgi network. Biochim Biophys Acta. 1497:253-60. Dong, X., G. Patino-Lopez, F. Candotti, and S. Shaw. 2007. Structure-function analysis of the WIP role in T cell receptor-stimulated NFAT activation: evidence that WIP- WASP dissociation is not required and that the WIP NH2 terminus is inhibitory. J Biol Chem. 282:30303-10. Dotti, C.G., C.A. Sullivan, and G.A. Banker. 1988. The establishment of polarity by hippocampal neurons in culture. J Neurosci. 8:1454-68. Dugina, V., I. Zwaenepoel, G. Gabbiani, S. Clement, and C. Chaponnier. 2009. Beta and gamma-cytoplasmic display distinct distribution and functional diversity. J Cell Sci. 122:2980-8.

97

Duleh, S.N., and M.D. Welch. 2010. WASH and the Arp2/3 complex regulate endosome shape and trafficking. Cytoskeleton (Hoboken). 67:193-206. Dupin, I., Y. Sakamoto, and S. Etienne-Manneville. 2011. Cytoplasmic intermediate filaments mediate actin-driven positioning of the nucleus. J Cell Sci. 124:865-72. Edson, K., B. Weisshaar, and A. Matus. 1993. Actin depolymerisation induces process formation on MAP2-transfected non-neuronal cells. Development. 117:689-700. Egea, G., F. Lazaro-Dieguez, and M. Vilella. 2006. Actin dynamics at the Golgi complex in mammalian cells. Curr Opin Cell Biol. 18:168-78. Eisenmann, K.M., E.S. Harris, S.M. Kitchen, H.A. Holman, H.N. Higgs, and A.S. Alberts. 2007. Dia-interacting protein modulates formin-mediated actin assembly at the cell cortex. Curr Biol. 17:579-91. Ellis, S., and H. Mellor. 2000. The novel Rho-family GTPase rif regulates coordinated actin-based membrane rearrangements. Curr Biol. 10:1387-90. Etienne-Manneville, S. 2004. Cdc42--the centre of polarity. J Cell Sci. 117:1291-300. Evangelista, M., K. Blundell, M.S. Longtine, C.J. Chow, N. Adames, J.R. Pringle, M. Peter, and C. Boone. 1997. Bni1p, a yeast formin linking cdc42p and the actin cytoskeleton during polarized morphogenesis. Science. 276:118-22. Evers, J., M. Laser, Y.A. Sun, Z.P. Xie, and M.M. Poo. 1989. Studies of nerve-muscle interactions in Xenopus cell culture: analysis of early synaptic currents. J Neurosci. 9:1523-39. Ezezika, O.C., N.S. Younger, J. Lu, D.A. Kaiser, Z.A. Corbin, B.J. Nolen, D.R. Kovar, and T.D. Pollard. 2009. Incompatibility with formin Cdc12p prevents human profilin from substituting for fission yeast profilin: insights from crystal structures of fission yeast profilin. J Biol Chem. 284:2088-97. Faix, J., D. Breitsprecher, T.E. Stradal, and K. Rottner. 2009. Filopodia: Complex models for simple rods. Int J Biochem Cell Biol. 41:1656-64. Fan, Y.M., C.P. Pang, A.R. Harvey, and Q. Cui. 2008. Marked effect of RhoA-specific shRNA-producing plasmids on neurite growth in PC12 cells. Neurosci Lett. 440:170- 5. Fergestad, T., and K. Broadie. 2001. Interaction of stoned and synaptotagmin in synaptic vesicle endocytosis. J Neurosci. 21:1218-27. Fernandez, I., D. Arac, J. Ubach, S.H. Gerber, O. Shin, Y. Gao, R.G. Anderson, T.C. Sudhof, and J. Rizo. 2001. Three-dimensional structure of the synaptotagmin 1 C2B-domain: synaptotagmin 1 as a phospholipid binding machine. Neuron. 32:1057- 69. Ferron, F., G. Rebowski, S.H. Lee, and R. Dominguez. 2007. Structural basis for the recruitment of profilin-actin complexes during filament elongation by Ena/VASP. EMBO J. 26:4597-606. Finkel, T., J.A. Theriot, K.R. Dise, G.F. Tomaselli, and P.J. Goldschmidt-Clermont. 1994. Dynamic actin structures stabilized by profilin. Proc Natl Acad Sci U S A. 91:1510-4. Firat-Karalar, E.N., and M.D. Welch. 2011. New mechanisms and functions of actin nucleation. Curr Opin Cell Biol. 23:4-13. Fischer, M., S. Kaech, U. Wagner, H. Brinkhaus, and A. Matus. 2000. Glutamate receptors regulate actin-based plasticity in dendritic spines. Nat Neurosci. 3:887-94. Flanagan, L.A., C.C. Cunningham, J. Chen, G.D. Prestwich, K.S. Kosik, and P.A. Janmey. 1997. The structure of divalent cation-induced aggregates of PIP2 and their alteration by gelsolin and tau. Biophys J. 73:1440-7.

98

Forscher, P., and S.J. Smith. 1988. Actions of cytochalasins on the organization of actin filaments and microtubules in a neuronal growth cone. J Cell Biol. 107:1505-16. Frazier, J.A., and C.M. Field. 1997. Actin cytoskeleton: are FH proteins local organizers? Curr Biol. 7:R414-7. Frieden, C., and K. Patane. 1988. Mechanism for nucleotide exchange in monomeric actin. Biochemistry. 27:3812-20. Friedman, H.V., T. Bresler, C.C. Garner, and N.E. Ziv. 2000. Assembly of new individual excitatory synapses: time course and temporal order of synaptic molecule recruitment. Neuron. 27:57-69. Frischknecht, F., and M. Way. 2001. Surfing pathogens and the lessons learned for actin polymerization. Trends Cell Biol. 11:30-38. Frost, N.A., H. Shroff, H. Kong, E. Betzig, and T.A. Blanpied. 2010. Single-molecule discrimination of discrete perisynaptic and distributed sites of actin filament assembly within dendritic spines. Neuron. 67:86-99. Fujii, T., A.H. Iwane, T. Yanagida, and K. Namba. 2010. Direct visualization of secondary structures of F-actin by electron cryomicroscopy. Nature. 467:724-8. Fukazawa, Y., Y. Saitoh, F. Ozawa, Y. Ohta, K. Mizuno, and K. Inokuchi. 2003. Hippocampal LTP is accompanied by enhanced F-actin content within the dendritic spine that is essential for late LTP maintenance in vivo. Neuron. 38:447-60. Fusco, D., N. Accornero, B. Lavoie, S.M. Shenoy, J.M. Blanchard, R.H. Singer, and E. Bertrand. 2003. Single mRNA molecules demonstrate probabilistic movement in living mammalian cells. Curr Biol. 13:161-7. Fyrberg, E.A., C.C. Fyrberg, J.R. Biggs, D. Saville, C.J. Beall, and A. Ketchum. 1998. Functional nonequivalence of Drosophila actin isoforms. Biochem Genet. 36:271-87. Gaffield, M.A., S.O. Rizzoli, and W.J. Betz. 2006. Mobility of synaptic vesicles in different pools in resting and stimulated frog motor nerve terminals. Neuron. 51:317- 25. Galbraith, C.G., K.M. Yamada, and J.A. Galbraith. 2007. Polymerizing actin fibers position integrins primed to probe for adhesion sites. Science. 315:992-5. Galbraith, C.G., K.M. Yamada, and M.P. Sheetz. 2002. The relationship between force and focal complex development. J Cell Biol. 159:695-705. Gao, M., D. Craig, O. Lequin, I.D. Campbell, V. Vogel, and K. Schulten. 2003. Structure and functional significance of mechanically unfolded fibronectin type III1 intermediates. Proc Natl Acad Sci U S A. 100:14784-9. Gardel, M.L., B. Sabass, L. Ji, G. Danuser, U.S. Schwarz, and C.M. Waterman. 2008. Traction stress in focal adhesions correlates biphasically with actin retrograde flow speed. J Cell Biol. 183:999-1005. Gareus, R., A. Di Nardo, V. Rybin, and W. Witke. 2006. Mouse profilin 2 regulates endocytosis and competes with SH3 ligand binding to dynamin 1. J Biol Chem. 281:2803-11. Garvalov, B.K., K.C. Flynn, D. Neukirchen, L. Meyn, N. Teusch, X. Wu, C. Brakebusch, J.R. Bamburg, and F. Bradke. 2007. Cdc42 regulates cofilin during the establishment of neuronal polarity. J Neurosci. 27:13117-29. Gasman, S., S. Chasserot-Golaz, P. Hubert, D. Aunis, and M.F. Bader. 1998. Identification of a potential effector pathway for the trimeric Go protein associated with secretory granules. Go stimulates a granule-bound phosphatidylinositol 4- kinase by activating RhoA in chromaffin cells. J Biol Chem. 273:16913-20.

99

Gasman, S., S. Chasserot-Golaz, M. Malacombe, M. Way, and M.F. Bader. 2004. Regulated exocytosis in neuroendocrine cells: a role for subplasmalemmal Cdc42/N-WASP-induced actin filaments. Mol Biol Cell. 15:520-31. Gasman, S., S. Chasserot-Golaz, M.R. Popoff, D. Aunis, and M.F. Bader. 1999. Involvement of Rho GTPases in calcium-regulated exocytosis from adrenal chromaffin cells. J Cell Sci. 112 ( Pt 24):4763-71. Geiger, B., and K.M. Yamada. 2011. Molecular Architecture and Function of Matrix Adhesions. Cold Spring Harb Perspect Biol. Geng, Y.J., T. Azuma, J.X. Tang, J.H. Hartwig, M. Muszynski, Q. Wu, P. Libby, and D.J. Kwiatkowski. 1998. Caspase-3-induced gelsolin fragmentation contributes to actin cytoskeletal collapse, nucleolysis, and apoptosis of vascular smooth muscle cells exposed to proinflammatory cytokines. Eur J Cell Biol. 77:294-302. Geppert, M., Y. Goda, R.E. Hammer, C. Li, T.W. Rosahl, C.F. Stevens, and T.C. Sudhof. 1994. Synaptotagmin I: a major Ca2+ sensor for transmitter release at a central synapse. Cell. 79:717-27. Geraldo, S., and P.R. Gordon-Weeks. 2009. Cytoskeletal dynamics in growth-cone steering. J Cell Sci. 122:3595-604. Gertler, F.B., A.R. Comer, J.L. Juang, S.M. Ahern, M.J. Clark, E.C. Liebl, and F.M. Hoffmann. 1995. enabled, a dosage-sensitive suppressor of mutations in the Drosophila Abl tyrosine kinase, encodes an Abl substrate with SH3 domain-binding properties. Genes Dev. 9:521-33. Giesemann, T., S. Rathke-Hartlieb, M. Rothkegel, J.W. Bartsch, S. Buchmeier, B.M. Jockusch, and H. Jockusch. 1999. A role for polyproline motifs in the spinal muscular atrophy protein SMN. Profilins bind to and colocalize with smn in nuclear gems. J Biol Chem. 274:37908-14. Gkogkas, C., N. Sonenberg, and M. Costa-Mattioli. 2010. Translational control mechanisms in long-lasting synaptic plasticity and memory. J Biol Chem. 285:31913-7. Goldschmidt-Clermont, P.J., M.I. Furman, D. Wachsstock, D. Safer, V.T. Nachmias, and T.D. Pollard. 1992. The control of actin nucleotide exchange by thymosin beta 4 and profilin. A potential regulatory mechanism for actin polymerization in cells. Mol Biol Cell. 3:1015-24. Goldschmidt-Clermont, P.J., J.W. Kim, L.M. Machesky, S.G. Rhee, and T.D. Pollard. 1991. Regulation of phospholipase C-gamma 1 by profilin and tyrosine phosphorylation. Science. 251:1231-3. Goldschmidt-Clermont, P.J., L.M. Machesky, J.J. Baldassare, and T.D. Pollard. 1990. The actin-binding protein profilin binds to PIP2 and inhibits its hydrolysis by phospholipase C. Science. 247:1575-8. Goley, E.D., and M.D. Welch. 2006. The ARP2/3 complex: an actin nucleator comes of age. Nat Rev Mol Cell Biol. 7:713-26. Golub, T., and P. Caroni. 2005. PI(4,5)P2-dependent microdomain assemblies capture microtubules to promote and control leading edge motility. J Cell Biol. 169:151-65. Golub, T., S. Wacha, and P. Caroni. 2004. Spatial and temporal control of signaling through lipid rafts. Curr Opin Neurobiol. 14:542-50. Gomez, T.S., K. Kumar, R.B. Medeiros, Y. Shimizu, P.J. Leibson, and D.D. Billadeau. 2007. Formins regulate the actin-related protein 2/3 complex-independent polarization of the centrosome to the immunological synapse. Immunity. 26:177-90.

100

Gould, C.J., S. Maiti, A. Michelot, B.R. Graziano, L. Blanchoin, and B.L. Goode. 2011. The formin DAD domain plays dual roles in autoinhibition and actin nucleation. Curr Biol. 21:384-90. Govind, S., R. Kozma, C. Monfries, L. Lim, and S. Ahmed. 2001. Cdc42Hs facilitates cytoskeletal reorganization and neurite outgrowth by localizing the 58-kD insulin receptor substrate to filamentous actin. J Cell Biol. 152:579-94. Grashoff, C., B.D. Hoffman, M.D. Brenner, R. Zhou, M. Parsons, M.T. Yang, M.A. McLean, S.G. Sligar, C.S. Chen, T. Ha, and M.A. Schwartz. 2010. Measuring mechanical tension across vinculin reveals regulation of focal adhesion dynamics. Nature. 466:263-6. Graumann, P.L. 2009. Dynamics of bacterial cytoskeletal elements. Cell Motil Cytoskeleton. 66:909-14. Grenklo, S., M. Geese, U. Lindberg, J. Wehland, R. Karlsson, and A.S. Sechi. 2003. A crucial role for profilin-actin in the intracellular motility of Listeria monocytogenes. EMBO Rep. 4:523-9. Gronborg, M., T.Z. Kristiansen, A. Iwahori, R. Chang, R. Reddy, N. Sato, H. Molina, O.N. Jensen, R.H. Hruban, M.G. Goggins, A. Maitra, and A. Pandey. 2006. Biomarker discovery from pancreatic cancer secretome using a differential proteomic approach. Mol Cell Proteomics. 5:157-71. Grove, M., G. Demyanenko, A. Echarri, P.A. Zipfel, M.E. Quiroz, R.M. Rodriguiz, M. Playford, S.A. Martensen, M.R. Robinson, W.C. Wetsel, P.F. Maness, and A.M. Pendergast. 2004. ABI2-deficient mice exhibit defective cell migration, aberrant dendritic spine morphogenesis, and deficits in learning and memory. Mol Cell Biol. 24:10905-22. Grummt, I. 2006. Actin and myosin as transcription factors. Curr Opin Genet Dev. 16:191-6. Gu, J., C.W. Lee, Y. Fan, D. Komlos, X. Tang, C. Sun, K. Yu, H.C. Hartzell, G. Chen, J.R. Bamburg, and J.Q. Zheng. 2010. ADF/cofilin-mediated actin dynamics regulate AMPA receptor trafficking during synaptic plasticity. Nat Neurosci. 13:1208-15. Gualdoni, S., C. Albertinazzi, S. Corbetta, F. Valtorta, and I. de Curtis. 2007. Normal levels of Rac1 are important for dendritic but not axonal development in hippocampal neurons. Biol Cell. 99:455-64. Guerrier, S., J. Coutinho-Budd, T. Sassa, A. Gresset, N.V. Jordan, K. Chen, W.L. Jin, A. Frost, and F. Polleux. 2009. The F-BAR domain of srGAP2 induces membrane protrusions required for neuronal migration and morphogenesis. Cell. 138:990-1004. Guillen, G., V. Valdes-Lopez, R. Noguez, J. Olivares, L.C. Rodriguez-Zapata, H. Perez, L. Vidali, M.A. Villanueva, and F. Sanchez. 1999. Profilin in Phaseolus vulgaris is encoded by two genes (only one expressed in root nodules) but multiple isoforms are generated in vivo by phosphorylation on tyrosine residues. Plant J. 19:497-508. Hachet, O., and A. Ephrussi. 2004. Splicing of oskar RNA in the nucleus is coupled to its cytoplasmic localization. Nature. 428:959-63. Haeckel, A., R. Ahuja, E.D. Gundelfinger, B. Qualmann, and M.M. Kessels. 2008. The actin-binding protein Abp1 controls dendritic spine morphology and is important for spine head and synapse formation. J Neurosci. 28:10031-44. Hahne, P., A. Sechi, S. Benesch, and J.V. Small. 2001. Scar/WAVE is localised at the tips of protruding lamellipodia in living cells. FEBS Lett. 492:215-20. Hajkova, L., T. Nyman, U. Lindberg, and R. Karlsson. 2000. Effects of cross-linked profilin:beta/gamma-actin on the dynamics of the microfilament system in cultured cells. Exp Cell Res. 256:112-21.

101

Halbrugge, M., and U. Walter. 1989. Purification of a vasodilator-regulated phosphoprotein from human platelets. Eur J Biochem. 185:41-50. Hall, A., and G. Lalli. 2010. Rho and Ras GTPases in axon growth, guidance, and branching. Cold Spring Harb Perspect Biol. 2:a001818. Hansson, A., G. Skoglund, I. Lassing, U. Lindberg, and M. Ingelman-Sundberg. 1988. Protein kinase C-dependent phosphorylation of profilin is specifically stimulated by phosphatidylinositol bisphosphate (PIP2). Biochem Biophys Res Commun. 150:526-31. Hartwig, J.H. 1995. Actin-binding proteins. 1: Spectrin super family. Protein Profile. 2:703-800. Hartwig, J.H., G.M. Bokoch, C.L. Carpenter, P.A. Janmey, L.A. Taylor, A. Toker, and T.P. Stossel. 1995. Thrombin receptor ligation and activated Rac uncap actin filament barbed ends through phosphoinositide synthesis in permeabilized human platelets. Cell. 82:643-53. Harvey, C.D., and K. Svoboda. 2007. Locally dynamic synaptic learning rules in pyramidal neuron dendrites. Nature. 450:1195-200. Hatano, S., and F. Oosawa. 1966. Isolation and characterization of plasmodium actin. Biochim Biophys Acta. 127:488-98. Haugwitz, M., A.A. Noegel, J. Karakesisoglou, and M. Schleicher. 1994. Dictyostelium amoebae that lack G-actin-sequestering profilins show defects in F-actin content, cytokinesis, and development. Cell. 79:303-14. Heasman, S.J., and A.J. Ridley. 2008. Mammalian Rho GTPases: new insights into their functions from in vivo studies. Nat Rev Mol Cell Biol. 9:690-701. Higgs, H.N., and T.D. Pollard. 2000. Activation by Cdc42 and PIP(2) of Wiskott- Aldrich syndrome protein (WASp) stimulates actin nucleation by Arp2/3 complex. J Cell Biol. 150:1311-20. Hilpela, P., M.K. Vartiainen, and P. Lappalainen. 2004. Regulation of the actin cytoskeleton by PI(4,5)P2 and PI(3,4,5)P3. Curr Top Microbiol Immunol. 282:117-63. Ho, H.Y., R. Rohatgi, A.M. Lebensohn, M. Le, J. Li, S.P. Gygi, and M.W. Kirschner. 2004. Toca-1 mediates Cdc42-dependent actin nucleation by activating the N- WASP-WIP complex. Cell. 118:203-16. Hodges, A.R., C.S. Bookwalter, E.B. Krementsova, and K.M. Trybus. 2009. A nonprocessive class V myosin drives cargo processively when a kinesin- related protein is a passenger. Curr Biol. 19:2121-5. Hoglund, A.S. 1985. The arrangement of microfilaments and microtubules in the periphery of spreading fibroblasts and glial cells. Tissue Cell. 17:649-66. Hoglund, A.S., R. Karlsson, E. Arro, B.A. Fredriksson, and U. Lindberg. 1980. Visualization of the peripheral weave of microfilaments in glia cells. J Muscle Res Cell Motil. 1:127-46. Holmes, K.C., D. Popp, W. Gebhard, and W. Kabsch. 1990. Atomic model of the actin filament. Nature. 347:44-9. Honore, B., P. Madsen, A.H. Andersen, and H. Leffers. 1993. Cloning and expression of a novel human profilin variant, profilin II. FEBS Lett. 330:151-5. Hoogenraad, C.C., A. Akhmanova, N. Galjart, and C.I. De Zeeuw. 2004. LIMK1 and CLIP-115: linking cytoskeletal defects to Williams syndrome. Bioessays. 26:141-50. Hotulainen, P., and P. Lappalainen. 2006. Stress fibers are generated by two distinct actin assembly mechanisms in motile cells. J Cell Biol. 173:383-94. Hotulainen, P., O. Llano, S. Smirnov, K. Tanhuanpaa, J. Faix, C. Rivera, and P. Lappalainen. 2009. Defining mechanisms of actin polymerization and depolymerization during dendritic spine morphogenesis. J Cell Biol. 185:323-39.

102

Hotulainen, P., E. Paunola, M.K. Vartiainen, and P. Lappalainen. 2005. Actin- depolymerizing factor and cofilin-1 play overlapping roles in promoting rapid F- actin depolymerization in mammalian nonmuscle cells. Mol Biol Cell. 16:649-64. Hu, E., Z. Chen, T. Fredrickson, and Y. Zhu. 2001. Molecular cloning and characterization of profilin-3: a novel cytoskeleton-associated gene expressed in rat kidney and testes. Exp Nephrol. 9:265-74. Huang, T.Y., C. DerMardirossian, and G.M. Bokoch. 2006. Cofilin phosphatases and regulation of actin dynamics. Curr Opin Cell Biol. 18:26-31. Hui, E., C.P. Johnson, J. Yao, F.M. Dunning, and E.R. Chapman. 2009. Synaptotagmin-mediated bending of the target membrane is a critical step in Ca(2+)-regulated fusion. Cell. 138:709-21. Humphries, J.D., A. Byron, M.D. Bass, S.E. Craig, J.W. Pinney, D. Knight, and M.J. Humphries. 2009. Proteomic analysis of integrin-associated complexes identifies RCC2 as a dual regulator of Rac1 and Arf6. Sci Signal. 2:ra51. Huttelmaier, S., B. Harbeck, O. Steffens, T. Messerschmidt, S. Illenberger, and B.M. Jockusch. 1999. Characterization of the actin binding properties of the vasodilator- stimulated phosphoprotein VASP. FEBS Lett. 451:68-74. Huttelmaier, S., D. Zenklusen, M. Lederer, J. Dictenberg, M. Lorenz, X. Meng, G.J. Bassell, J. Condeelis, and R.H. Singer. 2005. Spatial regulation of beta-actin translation by Src-dependent phosphorylation of ZBP1. Nature. 438:512-5. Ichetovkin, I., W. Grant, and J. Condeelis. 2002. Cofilin produces newly polymerized actin filaments that are preferred for dendritic nucleation by the Arp2/3 complex. Curr Biol. 12:79-84. Imai, K., S. Nonoyama, and H.D. Ochs. 2003. WASP (Wiskott-Aldrich syndrome protein) gene mutations and phenotype. Curr Opin Allergy Clin Immunol. 3:427-36. Imamura, H., K. Tanaka, T. Hihara, M. Umikawa, T. Kamei, K. Takahashi, T. Sasaki, and Y. Takai. 1997. Bni1p and Bnr1p: downstream targets of the Rho family small G-proteins which interact with profilin and regulate actin cytoskeleton in Saccharomyces cerevisiae. EMBO J. 16:2745-55. Innocenti, M., A. Zucconi, A. Disanza, E. Frittoli, L.B. Areces, A. Steffen, T.E. Stradal, P.P. Di Fiore, M.F. Carlier, and G. Scita. 2004. Abi1 is essential for the formation and activation of a WAVE2 signalling complex. Nat Cell Biol. 6:319-27. Ishizaki, T., Y. Morishima, M. Okamoto, T. Furuyashiki, T. Kato, and S. Narumiya. 2001. Coordination of microtubules and the actin cytoskeleton by the Rho effector mDia1. Nat Cell Biol. 3:8-14. Ismail, A.M., S.B. Padrick, B. Chen, J. Umetani, and M.K. Rosen. 2009. The WAVE regulatory complex is inhibited. Nat Struct Mol Biol. 16:561-3. Jahn, R., and R.H. Scheller. 2006. SNAREs--engines for membrane fusion. Nat Rev Mol Cell Biol. 7:631-43. James, D.J., C. Khodthong, J.A. Kowalchyk, and T.F. Martin. 2008. Phosphatidylinositol 4,5-bisphosphate regulates SNARE-dependent membrane fusion. J Cell Biol. 182:355-66. Janke, J., K. Schluter, B. Jandrig, M. Theile, K. Kolble, W. Arnold, E. Grinstein, A. Schwartz, L. Estevez-Schwarz, P.M. Schlag, B.M. Jockusch, and S. Scherneck. 2000. Suppression of tumorigenicity in breast cancer cells by the microfilament protein profilin 1. J Exp Med. 191:1675-86.

103

Jaworski, J., L.C. Kapitein, S.M. Gouveia, B.R. Dortland, P.S. Wulf, I. Grigoriev, P. Camera, S.A. Spangler, P. Di Stefano, J. Demmers, H. Krugers, P. Defilippi, A. Akhmanova, and C.C. Hoogenraad. 2009. Dynamic microtubules regulate dendritic spine morphology and synaptic plasticity. Neuron. 61:85-100. Jia, D., T.S. Gomez, Z. Metlagel, J. Umetani, Z. Otwinowski, M.K. Rosen, and D.D. Billadeau. 2010. WASH and WAVE actin regulators of the Wiskott-Aldrich syndrome protein (WASP) family are controlled by analogous structurally related complexes. Proc Natl Acad Sci U S A. 107:10442-7. Kabsch, W., H.G. Mannherz, D. Suck, E.F. Pai, and K.C. Holmes. 1990. Atomic structure of the actin:DNase I complex. Nature. 347:37-44. Kang, F., D.L. Purich, and F.S. Southwick. 1999. Profilin promotes barbed-end actin filament assembly without lowering the critical concentration. J Biol Chem. 274:36963-72. Kang, H., and E.M. Schuman. 1996. A requirement for local protein synthesis in neurotrophin-induced hippocampal synaptic plasticity. Science. 273:1402-6. Kang, R., R. Swayze, M.F. Lise, K. Gerrow, A. Mullard, W.G. Honer, and A. El- Husseini. 2004. Presynaptic trafficking of synaptotagmin I is regulated by protein palmitoylation. J Biol Chem. 279:50524-36. Karakozova, M., M. Kozak, C.C. Wong, A.O. Bailey, J.R. Yates, 3rd, A. Mogilner, H. Zebroski, and A. Kashina. 2006. Arginylation of beta-actin regulates actin cytoskeleton and cell motility. Science. 313:192-6. Karlsson, R., P. Aspenstrom, and A.S. Bystrom. 1991. A chicken beta-actin gene can complement a disruption of the Saccharomyces cerevisiae ACT1 gene. Mol Cell Biol. 11:213-7. Karlsson, R., and U. Lindberg. 1985. Changes in the organization of actin and myosin in non-muscle cells induced by N-ethylmaleimide. Exp Cell Res. 157:95-115. Karlsson, R., and U. Lindberg. 2007. Profilin, an essential control element for actin polymerization. In In Actin Monomer-binding Proteins. . P. Lappalainen., editor. Landes Bioscience, Georgetown. Kasai, H., M. Fukuda, S. Watanabe, A. Hayashi-Takagi, and J. Noguchi. 2010a. Structural dynamics of dendritic spines in memory and cognition. Trends Neurosci. 33:121-9. Kasai, H., T. Hayama, M. Ishikawa, S. Watanabe, S. Yagishita, and J. Noguchi. 2010b. Learning rules and persistence of dendritic spines. Eur J Neurosci. 32:241-9. Katsumi, A., T. Naoe, T. Matsushita, K. Kaibuchi, and M.A. Schwartz. 2005. Integrin activation and matrix binding mediate cellular responses to mechanical stretch. J Biol Chem. 280:16546-9. Kaverina, I., O. Krylyshkina, and J.V. Small. 2002. Regulation of substrate adhesion dynamics during cell motility. Int J Biochem Cell Biol. 34:746-61. Kessels, M.M., L. Schwintzer, D. Schlobinski, and B. Qualmann. 2010. Controlling actin cytoskeletal organization and dynamics during neuronal morphogenesis. Eur J Cell Biol. Kim, C.H., and J.E. Lisman. 1999. A role of actin filament in synaptic transmission and long-term potentiation. J Neurosci. 19:4314-24. Kincaid, M.M., and S.J. King. 2006. Motors and their tethers: the role of secondary binding sites in processive motility. Cell Cycle. 5:2733-7. Knoll, B., and A. Nordheim. 2009. Functional versatility of transcription factors in the nervous system: the SRF paradigm. Trends Neurosci. 32:432-42.

104

Kobayashi, K., S. Kuroda, M. Fukata, T. Nakamura, T. Nagase, N. Nomura, Y. Matsuura, N. Yoshida-Kubomura, A. Iwamatsu, and K. Kaibuchi. 1998. p140Sra-1 (specifically Rac1-associated protein) is a novel specific target for Rac1 small GTPase. J Biol Chem. 273:291-5. Koestler, S.A., S. Auinger, M. Vinzenz, K. Rottner, and J.V. Small. 2008. Differentially oriented populations of actin filaments generated in lamellipodia collaborate in pushing and pausing at the cell front. Nat Cell Biol. 10:306-13. Korenbaum, E., P. Nordberg, C. Bjorkegren-Sjogren, C.E. Schutt, U. Lindberg, and R. Karlsson. 1998. The role of profilin in actin polymerization and nucleotide exchange. Biochemistry. 37:9274-83. Korn, E.D. 1982. Actin polymerization and its regulation by proteins from nonmuscle cells. Physiol Rev. 62:672-737. Kovar, D.R., E.S. Harris, R. Mahaffy, H.N. Higgs, and T.D. Pollard. 2006. Control of the assembly of ATP- and ADP-actin by formins and profilin. Cell. 124:423-35. Kozma, R., S. Ahmed, A. Best, and L. Lim. 1995. The Ras-related protein Cdc42Hs and bradykinin promote formation of peripheral actin microspikes and filopodia in Swiss 3T3 fibroblasts. Mol Cell Biol. 15:1942-52. Kraszewski, K., O. Mundigl, L. Daniell, C. Verderio, M. Matteoli, and P. De Camilli. 1995. Synaptic vesicle dynamics in living cultured hippocampal neurons visualized with CY3-conjugated antibodies directed against the lumenal domain of synaptotagmin. J Neurosci. 15:4328-42. Krause, M., E.W. Dent, J.E. Bear, J.J. Loureiro, and F.B. Gertler. 2003. Ena/VASP proteins: regulators of the actin cytoskeleton and cell migration. Annu Rev Cell Dev Biol. 19:541-64. Krugmann, S., I. Jordens, K. Gevaert, M. Driessens, J. Vandekerckhove, and A. Hall. 2001. Cdc42 induces filopodia by promoting the formation of an IRSp53:Mena complex. Curr Biol. 11:1645-55. Kunda, P., G. Craig, V. Dominguez, and B. Baum. 2003. Abi, Sra1, and Kette control the stability and localization of SCAR/WAVE to regulate the formation of actin- based protrusions. Curr Biol. 13:1867-75. Kuo, W., D.Z. Herrick, and D.S. Cafiso. 2011. Phosphatidylinositol 4,5-Bisphosphate Alters Synaptotagmin 1 Membrane Docking and Drives Opposing Bilayers Closer Together. Biochemistry. Kuo, W., D.Z. Herrick, J.F. Ellena, and D.S. Cafiso. 2009. The calcium-dependent and calcium-independent membrane binding of synaptotagmin 1: two modes of C2B binding. J Mol Biol. 387:284-94. Kuriu, T., A. Inoue, H. Bito, K. Sobue, and S. Okabe. 2006. Differential control of postsynaptic density scaffolds via actin-dependent and -independent mechanisms. J Neurosci. 26:7693-706. Kuromi, H., and Y. Kidokoro. 2005. Exocytosis and endocytosis of synaptic vesicles and functional roles of vesicle pools: lessons from the Drosophila neuromuscular junction. Neuroscientist. 11:138-47. Kwiatkowska, K. 2010. One lipid, multiple functions: how various pools of PI(4,5)P(2) are created in the plasma membrane. Cell Mol Life Sci. 67:3927-46. Ladwein, M., and K. Rottner. 2008. On the Rho'd: the regulation of membrane protrusions by Rho-GTPases. FEBS Lett. 582:2066-74. Lai, F.P., M. Szczodrak, J. Block, J. Faix, D. Breitsprecher, H.G. Mannherz, T.E. Stradal, G.A. Dunn, J.V. Small, and K. Rottner. 2008. Arp2/3 complex interactions and actin network turnover in lamellipodia. EMBO J. 27:982-92.

105

Laing, N.G., D.E. Dye, C. Wallgren-Pettersson, G. Richard, N. Monnier, S. Lillis, T.L. Winder, H. Lochmuller, C. Graziano, S. Mitrani-Rosenbaum, D. Twomey, J.C. Sparrow, A.H. Beggs, and K.J. Nowak. 2009. Mutations and polymorphisms of the skeletal muscle alpha-actin gene (ACTA1). Hum Mutat. 30:1267-77. Lambrechts, A., V. Jonckheere, D. Dewitte, J. Vandekerckhove, and C. Ampe. 2002. Mutational analysis of human profilin I reveals a second PI(4,5)-P2 binding site neighbouring the poly(L-proline) binding site. BMC Biochem. 3:12. Lambrechts, A., V. Jonckheere, C. Peleman, D. Polet, W. De Vos, J. Vandekerckhove, and C. Ampe. 2006. Profilin-I-ligand interactions influence various aspects of neuronal differentiation. J Cell Sci. 119:1570-8. Lambrechts, A., J. van Damme, M. Goethals, J. Vandekerckhove, and C. Ampe. 1995. Purification and characterization of bovine profilin II. Actin, poly(L-proline) and inositolphospholipid binding. Eur J Biochem. 230:281-6. Lambrechts, A., J.L. Verschelde, V. Jonckheere, M. Goethals, J. Vandekerckhove, and C. Ampe. 1997. The mammalian profilin isoforms display complementary affinities for PIP2 and proline-rich sequences. EMBO J. 16:484-94. Lan, Y., and G.A. Papoian. 2008. The stochastic dynamics of filopodial growth. Biophys J. 94:3839-52. Landis, D.M., A.K. Hall, L.A. Weinstein, and T.S. Reese. 1988. The organization of cytoplasm at the presynaptic active zone of a central nervous system synapse. Neuron. 1:201-9. Landis, D.M., and T.S. Reese. 1983. Cytoplasmic organization in cerebellar dendritic spines. J Cell Biol. 97:1169-78. Lanier, L.M., M.A. Gates, W. Witke, A.S. Menzies, A.M. Wehman, J.D. Macklis, D. Kwiatkowski, P. Soriano, and F.B. Gertler. 1999. Mena is required for neurulation and commissure formation. Neuron. 22:313-25. Lassing, I., and U. Lindberg. 1985. Specific interaction between phosphatidylinositol 4,5-bisphosphate and profilactin. Nature. 314:472-4. Lassing, I., and U. Lindberg. 1988. Evidence that the phosphatidylinositol cycle is linked to cell motility. Exp Cell Res. 174:1-15. Lassing, I., and U. Lindberg. 1990. Polyphosphoinositide synthesis in platelets stimulated with low concentrations of thrombin is enhanced before the activation of phospholipase C. FEBS Lett. 262:231-3. Latham, V.M., E.H. Yu, A.N. Tullio, R.S. Adelstein, and R.H. Singer. 2001. A Rho- dependent signaling pathway operating through myosin localizes beta-actin mRNA in fibroblasts. Curr Biol. 11:1010-6. Laurent, V., T.P. Loisel, B. Harbeck, A. Wehman, L. Grobe, B.M. Jockusch, J. Wehland, F.B. Gertler, and M.F. Carlier. 1999. Role of proteins of the Ena/VASP family in actin-based motility of Listeria monocytogenes. J Cell Biol. 144:1245-58. Laux, T., K. Fukami, M. Thelen, T. Golub, D. Frey, and P. Caroni. 2000. GAP43, MARCKS, and CAP23 modulate PI(4,5)P(2) at plasmalemmal rafts, and regulate cell cortex actin dynamics through a common mechanism. J Cell Biol. 149:1455-72. Le Clainche, C., and M.F. Carlier. 2008. Regulation of actin assembly associated with protrusion and adhesion in cell migration. Physiol Rev. 88:489-513. Le, J., E.L. Mallery, C. Zhang, S. Brankle, and D.B. Szymanski. 2006. Arabidopsis BRICK1/HSPC300 is an essential WAVE-complex subunit that selectively stabilizes the Arp2/3 activator SCAR2. Curr Biol. 16:895-901.

106

Lecuyer, E., H. Yoshida, N. Parthasarathy, C. Alm, T. Babak, T. Cerovina, T.R. Hughes, P. Tomancak, and H.M. Krause. 2007. Global analysis of mRNA localization reveals a prominent role in organizing cellular architecture and function. Cell. 131:174-87. Lederer, M., B.M. Jockusch, and M. Rothkegel. 2005. Profilin regulates the activity of p42POP, a novel Myb-related transcription factor. J Cell Sci. 118:331-41. Lee, H.K., Y. Yang, Z. Su, C. Hyeon, T.S. Lee, H.W. Lee, D.H. Kweon, Y.K. Shin, and T.Y. Yoon. 2010. Dynamic Ca2+-dependent stimulation of vesicle fusion by membrane-anchored synaptotagmin 1. Science. 328:760-3. Legg, J.A., G. Bompard, J. Dawson, H.L. Morris, N. Andrew, L. Cooper, S.A. Johnston, G. Tramountanis, and L.M. Machesky. 2007. N-WASP involvement in dorsal ruffle formation in mouse embryonic fibroblasts. Mol Biol Cell. 18:678-87. Lele, T.P., J. Pendse, S. Kumar, M. Salanga, J. Karavitis, and D.E. Ingber. 2006. Mechanical forces alter zyxin unbinding kinetics within focal adhesions of living cells. J Cell Physiol. 207:187-94. Leung, K.M., F.P. van Horck, A.C. Lin, R. Allison, N. Standart, and C.E. Holt. 2006. Asymmetrical beta-actin mRNA translation in growth cones mediates attractive turning to netrin-1. Nat Neurosci. 9:1247-56. Leyman, S., M. Sidani, L. Ritsma, D. Waterschoot, R. Eddy, D. Dewitte, O. Debeir, C. Decaestecker, J. Vandekerckhove, J. van Rheenen, C. Ampe, J. Condeelis, and M. Van Troys. 2009. Unbalancing the phosphatidylinositol-4,5-bisphosphate-cofilin interaction impairs cell steering. Mol Biol Cell. 20:4509-23. Li, L., O.H. Shin, J.S. Rhee, D. Arac, J.C. Rah, J. Rizo, T. Sudhof, and C. Rosenmund. 2006. Phosphatidylinositol phosphates as co-activators of Ca2+ binding to C2 domains of synaptotagmin 1. J Biol Chem. 281:15845-52. Li, Y., S. Grenklo, T. Higgins, and R. Karlsson. 2008. The profilin:actin complex localizes to sites of dynamic actin polymerization at the leading edge of migrating cells and pathogen-induced actin tails. Eur J Cell Biol. 87:893-904. Li, Z., and M. Sheng. 2003. Some assembly required: the development of neuronal synapses. Nat Rev Mol Cell Biol. 4:833-41. Liao, G., X. Ma, and G. Liu. 2011a. An RNA-zipcode-independent mechanism that localizes Dia1 mRNA to the perinuclear ER through interactions between Dia1 nascent peptide and Rho-GTP. J Cell Sci. 124:589-99. Liao, G., B. Simone, and G. Liu. 2011b. Mis-localization of Arp2 mRNA impairs persistence of directional cell migration. Exp Cell Res. 317:812-22. Lim, K.B., W. Bu, W.I. Goh, E. Koh, S.H. Ong, T. Pawson, T. Sudhaharan, and S. Ahmed. 2008. The Cdc42 effector IRSp53 generates filopodia by coupling membrane protrusion with actin dynamics. J Biol Chem. 283:20454-72. Lin, C.H., E.M. Espreafico, M.S. Mooseker, and P. Forscher. 1997. Myosin drives retrograde F-actin flow in neuronal growth cones. Biol Bull. 192:183-5. Lin, Q., R.N. Fuji, W. Yang, and R.A. Cerione. 2003. RhoGDI is required for Cdc42- mediated cellular transformation. Curr Biol. 13:1469-79. Lindberg, U., A.S. Hoglund, and R. Karlsson. 1981. On the ultrastructural organization of the microfilament system and the possible role of profilactin. Biochimie. 63:307-23. Lindberg, U., R. Karlsson, I. Lassing, C.E. Schutt, and A.S. Hoglund. 2008. The microfilament system and malignancy. Semin Cancer Biol. 18:2-11.

107

Littleton, J.T., J. Bai, B. Vyas, R. Desai, A.E. Baltus, M.B. Garment, S.D. Carlson, B. Ganetzky, and E.R. Chapman. 2001. synaptotagmin mutants reveal essential functions for the C2B domain in Ca2+-triggered fusion and recycling of synaptic vesicles in vivo. J Neurosci. 21:1421-33. Littleton, J.T., H.J. Bellen, and M.S. Perin. 1993. Expression of synaptotagmin in Drosophila reveals transport and localization of synaptic vesicles to the synapse. Development. 118:1077-88. Littleton, J.T., L. Upton, and A. Kania. 1995. Immunocytochemical analysis of axonal outgrowth in synaptotagmin mutations. J Neurochem. 65:32-40. Liu, R., M.T. Abreu-Blanco, K.C. Barry, E.V. Linardopoulou, G.E. Osborn, and S.M. Parkhurst. 2009. Wash functions downstream of Rho and links linear and branched actin nucleation factors. Development. 136:2849-60. Lozano, E., M. Betson, and V.M. Braga. 2003. Tumor progression: Small GTPases and loss of cell-cell adhesion. Bioessays. 25:452-63. Lu, P.J., W.R. Shieh, S.G. Rhee, H.L. Yin, and C.S. Chen. 1996. Lipid products of phosphoinositide 3-kinase bind human profilin with high affinity. Biochemistry. 35:14027-34. Mace, K.E., L.M. Biela, A.G. Sares, and N.E. Reist. 2009. Synaptotagmin I stabilizes synaptic vesicles via its C(2)A polylysine motif. Genesis. 47:337-45. Machesky, L.M., S.J. Atkinson, C. Ampe, J. Vandekerckhove, and T.D. Pollard. 1994. Purification of a cortical complex containing two unconventional actins from Acanthamoeba by affinity chromatography on profilin-agarose. J Cell Biol. 127:107-15. Machesky, L.M., and A. Hall. 1996. Rho: a connection between membrane receptor signalling and the cytoskeleton. Trends Cell Biol. 6:304-10. Machesky, L.M., and A. Hall. 1997. Role of actin polymerization and adhesion to extracellular matrix in Rac- and Rho-induced cytoskeletal reorganization. J Cell Biol. 138:913-26. Machesky, L.M., and R.H. Insall. 1998. Scar1 and the related Wiskott-Aldrich syndrome protein, WASP, regulate the actin cytoskeleton through the Arp2/3 complex. Curr Biol. 8:1347-56. Machesky, L.M., R.D. Mullins, H.N. Higgs, D.A. Kaiser, L. Blanchoin, R.C. May, M.E. Hall, and T.D. Pollard. 1999. Scar, a WASp-related protein, activates nucleation of actin filaments by the Arp2/3 complex. Proc Natl Acad Sci U S A. 96:3739-44. Mackler, J.M., J.A. Drummond, C.A. Loewen, I.M. Robinson, and N.E. Reist. 2002. The C(2)B Ca(2+)-binding motif of synaptotagmin is required for synaptic transmission in vivo. Nature. 418:340-4. Mahaffy, R.E., and T.D. Pollard. 2008. Influence of phalloidin on the formation of actin filament branches by Arp2/3 complex. Biochemistry. 47:6460-7. Mahoney, N.M., P.A. Janmey, and S.C. Almo. 1997. Structure of the profilin-poly-L- proline complex involved in morphogenesis and cytoskeletal regulation. Nat Struct Biol. 4:953-60. Marchand, J.B., D.A. Kaiser, T.D. Pollard, and H.N. Higgs. 2001. Interaction of WASP/Scar proteins with actin and vertebrate Arp2/3 complex. Nat Cell Biol. 3:76-82. Markey, F., T. Persson, and U. Lindberg. 1981. Characterization of platelet extracts before and after stimulation with respect to the possible role of profilactin as microfilament precursor. Cell. 23:145-53.

108

Martens, S., M.M. Kozlov, and H.T. McMahon. 2007. How synaptotagmin promotes membrane fusion. Science. 316:1205-8. Martin, J.A., Z. Hu, K.M. Fenz, J. Fernandez, and J.S. Dittman. 2011. Complexin has opposite effects on two modes of synaptic vesicle fusion. Curr Biol. 21:97-105. Martinez-Quiles, N., R. Rohatgi, I.M. Anton, M. Medina, S.P. Saville, H. Miki, H. Yamaguchi, T. Takenawa, J.H. Hartwig, R.S. Geha, and N. Ramesh. 2001. WIP regulates N-WASP-mediated actin polymerization and filopodium formation. Nat Cell Biol. 3:484-91. Matsumura, F., and D.J. Hartshorne. 2008. Myosin phosphatase target subunit: Many roles in cell function. Biochem Biophys Res Commun. 369:149-56. Matthew, W.D., L. Tsavaler, and L.F. Reichardt. 1981. Identification of a synaptic vesicle-specific membrane protein with a wide distribution in neuronal and neurosecretory tissue. J Cell Biol. 91:257-69. Mattila, P.K., and P. Lappalainen. 2008. Filopodia: molecular architecture and cellular functions. Nat Rev Mol Cell Biol. 9:446-54. Mattila, P.K., A. Pykalainen, J. Saarikangas, V.O. Paavilainen, H. Vihinen, E. Jokitalo, and P. Lappalainen. 2007. Missing-in-metastasis and IRSp53 deform PI(4,5)P2-rich membranes by an inverse BAR domain-like mechanism. J Cell Biol. 176:953-64. Matusek, T., R. Gombos, A. Szecsenyi, N. Sanchez-Soriano, A. Czibula, C. Pataki, A. Gedai, A. Prokop, I. Rasko, and J. Mihaly. 2008. Formin proteins of the DAAM subfamily play a role during axon growth. J Neurosci. 28:13310-9. Maximov, A., and T.C. Sudhof. 2005. Autonomous function of synaptotagmin 1 in triggering synchronous release independent of asynchronous release. Neuron. 48:547-54. Maximov, A., J. Tang, X. Yang, Z.P. Pang, and T.C. Sudhof. 2009. Complexin controls the force transfer from SNARE complexes to membranes in fusion. Science. 323:516-21. Mayboroda, O., K. Schluter, and B.M. Jockusch. 1997. Differential colocalization of profilin with microfilaments in PtK2 cells. Cell Motil Cytoskeleton. 37:166-77. McGough, A., W. Chiu, and M. Way. 1998. Determination of the gelsolin binding site on F-actin: implications for severing and capping. Biophys J. 74:764-72. McNair, K., R. Spike, C. Guilding, G.C. Prendergast, T.W. Stone, S.R. Cobb, and B.J. Morris. 2010. A role for RhoB in synaptic plasticity and the regulation of neuronal morphology. J Neurosci. 30:3508-17. Merrifield, C.J., M.E. Feldman, L. Wan, and W. Almers. 2002. Imaging actin and dynamin recruitment during invagination of single clathrin-coated pits. Nat Cell Biol. 4:691-8. Merrifield, C.J., D. Perrais, and D. Zenisek. 2005. Coupling between clathrin-coated-pit invagination, cortactin recruitment, and membrane scission observed in live cells. Cell. 121:593-606. Merrifield, C.J., B. Qualmann, M.M. Kessels, and W. Almers. 2004. Neural Wiskott Aldrich Syndrome Protein (N-WASP) and the Arp2/3 complex are recruited to sites of clathrin-mediated endocytosis in cultured fibroblasts. Eur J Cell Biol. 83:13-8. Meyer, M.P., and S.J. Smith. 2006. Evidence from in vivo imaging that synaptogenesis guides the growth and branching of axonal arbors by two distinct mechanisms. J Neurosci. 26:3604-14. Michaelsen, K., K. Murk, M. Zagrebelsky, A. Dreznjak, B.M. Jockusch, M. Rothkegel, and M. Korte. 2010. Fine-tuning of neuronal architecture requires two profilin isoforms. Proc Natl Acad Sci U S A. 107:15780-5.

109

Michelot, A., C. Guerin, S. Huang, M. Ingouff, S. Richard, N. Rodiuc, C.J. Staiger, and L. Blanchoin. 2005. The formin homology 1 domain modulates the actin nucleation and bundling activity of Arabidopsis FORMIN1. Plant Cell. 17:2296-313. Miki, H., T. Sasaki, Y. Takai, and T. Takenawa. 1998. Induction of filopodium formation by a WASP-related actin-depolymerizing protein N-WASP. Nature. 391:93-6. Miki, H., H. Yamaguchi, S. Suetsugu, and T. Takenawa. 2000. IRSp53 is an essential intermediate between Rac and WAVE in the regulation of membrane ruffling. Nature. 408:732-5. Mili, S., and I.G. Macara. 2009. RNA localization and polarity: from A(PC) to Z(BP). Trends Cell Biol. 19:156-64. Mili, S., K. Moissoglu, and I.G. Macara. 2008. Genome-wide screen reveals APC- associated RNAs enriched in cell protrusions. Nature. 453:115-9. Millard, T.H., G. Bompard, M.Y. Heung, T.R. Dafforn, D.J. Scott, L.M. Machesky, and K. Futterer. 2005. Structural basis of filopodia formation induced by the IRSp53/MIM homology domain of human IRSp53. EMBO J. 24:240-50. Miller, K.E., and M.P. Sheetz. 2000. Characterization of myosin V binding to brain vesicles. J Biol Chem. 275:2598-606. Miller, S., M. Yasuda, J.K. Coats, Y. Jones, M.E. Martone, and M. Mayford. 2002. Disruption of dendritic translation of CaMKIIalpha impairs stabilization of synaptic plasticity and memory consolidation. Neuron. 36:507-19. Mingle, L.A., N.N. Okuhama, J. Shi, R.H. Singer, J. Condeelis, and G. Liu. 2005. Localization of all seven messenger RNAs for the actin-polymerization nucleator Arp2/3 complex in the protrusions of fibroblasts. J Cell Sci. 118:2425-33. Mittelsteadt, T., G. Seifert, E. Alvarez-Baron, C. Steinhauser, A.J. Becker, and S. Schoch. 2009. Differential mRNA expression patterns of the synaptotagmin gene family in the rodent brain. J Comp Neurol. 512:514-28. Mogilner, A., and B. Rubinstein. 2005. The physics of filopodial protrusion. Biophys J. 89:782-95. Moldovan, N.I., E.E. Milliken, K. Irani, J. Chen, R.H. Sohn, T. Finkel, and P.J. Goldschmidt-Clermont. 1997. Regulation of endothelial cell adhesion by profilin. Curr Biol. 7:24-30. Momboisse, F., S. Houy, S. Ory, V. Calco, M.F. Bader, and S. Gasman. 2011. How important are Rho GTPases in neurosecretion? J Neurochem. Morales, M., M.A. Colicos, and Y. Goda. 2000. Actin-dependent regulation of neurotransmitter release at central synapses. Neuron. 27:539-50. Mouneimne, G., L. Soon, V. DesMarais, M. Sidani, X. Song, S.C. Yip, M. Ghosh, R. Eddy, J.M. Backer, and J. Condeelis. 2004. Phospholipase C and cofilin are required for carcinoma cell directionality in response to EGF stimulation. J Cell Biol. 166:697- 708. Mozhayeva, M.G., Y. Sara, X. Liu, and E.T. Kavalali. 2002. Development of vesicle pools during maturation of hippocampal synapses. J Neurosci. 22:654-65. Mullins, R.D. 2000. How WASP-family proteins and the Arp2/3 complex convert intracellular signals into cytoskeletal structures. Curr Opin Cell Biol. 12:91-6. Mullins, R.D., J.A. Heuser, and T.D. Pollard. 1998. The interaction of Arp2/3 complex with actin: nucleation, high affinity pointed end capping, and formation of branching networks of filaments. Proc Natl Acad Sci U S A. 95:6181-6. Murray, D., and B. Honig. 2002. Electrostatic control of the membrane targeting of C2 domains. Mol Cell. 9:145-54.

110

Murthy, V.N., T.J. Sejnowski, and C.F. Stevens. 1997. Heterogeneous release properties of visualized individual hippocampal synapses. Neuron. 18:599-612. Myat, M.M., S. Anderson, L.A. Allen, and A. Aderem. 1997. MARCKS regulates membrane ruffling and cell spreading. Curr Biol. 7:611-4. Nagy, G., I. Milosevic, R. Mohrmann, K. Wiederhold, A.M. Walter, and J.B. Sorensen. 2008a. The SNAP-25 linker as an adaptation toward fast exocytosis. Mol Biol Cell. 19:3769-81. Nagy, S., B.L. Ricca, M.F. Norstrom, D.S. Courson, C.M. Brawley, P.A. Smithback, and R.S. Rock. 2008b. A myosin motor that selects bundled actin for motility. Proc Natl Acad Sci U S A. 105:9616-20. Neher, E., and R. Penner. 1994. Mice sans synaptotagmin. Nature. 372:316-7. Nemethova, M., S. Auinger, and J.V. Small. 2008. Building the actin cytoskeleton: filopodia contribute to the construction of contractile bundles in the lamella. J Cell Biol. 180:1233-44. Neuhaus, J.M., M. Wanger, T. Keiser, and A. Wegner. 1983. Treadmilling of actin. J Muscle Res Cell Motil. 4:507-27. Neuhoff, H., M. Sassoe-Pognetto, P. Panzanelli, C. Maas, W. Witke, and M. Kneussel. 2005. The actin-binding protein profilin I is localized at synaptic sites in an activity- regulated manner. Eur J Neurosci. 21:15-25. Nicholson-Dykstra, S.M., and H.N. Higgs. 2008. Arp2 depletion inhibits sheet-like protrusions but not linear protrusions of fibroblasts and lymphocytes. Cell Motil Cytoskeleton. 65:904-22. Niell, C.M., M.P. Meyer, and S.J. Smith. 2004. In vivo imaging of synapse formation on a growing dendritic arbor. Nat Neurosci. 7:254-60. Nishiki, T., and G.J. Augustine. 2004a. Dual roles of the C2B domain of synaptotagmin I in synchronizing Ca2+-dependent neurotransmitter release. J Neurosci. 24:8542-50. Nishiki, T., and G.J. Augustine. 2004b. Synaptotagmin I synchronizes transmitter release in mouse hippocampal neurons. J Neurosci. 24:6127-32. Nyman, T., H. Schuler, E. Korenbaum, C.E. Schutt, R. Karlsson, and U. Lindberg. 2002. The role of MeH73 in actin polymerization and ATP hydrolysis. J Mol Biol. 317:577-89. Obermann, H., I. Raabe, M. Balvers, B. Brunswig, W. Schulze, and C. Kirchhoff. 2005. Novel testis-expressed profilin IV associated with acrosome biogenesis and spermatid elongation. Mol Hum Reprod. 11:53-64. Oda, T., M. Iwasa, T. Aihara, Y. Maeda, and A. Narita. 2009. The nature of the globular- to fibrous-actin transition. Nature. 457:441-5. Odronitz, F., and M. Kollmar. 2007. Drawing the tree of eukaryotic life based on the analysis of 2,269 manually annotated myosins from 328 species. Genome Biol. 8:R196. Oikawa, T., H. Yamaguchi, T. Itoh, M. Kato, T. Ijuin, D. Yamazaki, S. Suetsugu, and T. Takenawa. 2004. PtdIns(3,4,5)P3 binding is necessary for WAVE2-induced formation of lamellipodia. Nat Cell Biol. 6:420-6. Okada, K., F. Bartolini, A.M. Deaconescu, J.B. Moseley, Z. Dogic, N. Grigorieff, G.G. Gundersen, and B.L. Goode. 2010. Adenomatous polyposis coli protein nucleates actin assembly and synergizes with the formin mDia1. J Cell Biol. 189:1087-96. Okamoto, K., T. Nagai, A. Miyawaki, and Y. Hayashi. 2004. Rapid and persistent modulation of actin dynamics regulates postsynaptic reorganization underlying bidirectional plasticity. Nat Neurosci. 7:1104-12.

111

Orr, A.W., M.H. Ginsberg, S.J. Shattil, H. Deckmyn, and M.A. Schwartz. 2006. Matrix- specific suppression of integrin activation in shear stress signaling. Mol Biol Cell. 17:4686-97. Ostrander, D.B., J.A. Gorman, and G.M. Carman. 1995. Regulation of profilin localization in Saccharomyces cerevisiae by phosphoinositide metabolism. J Biol Chem. 270:27045-50. Otey, C.A., M.H. Kalnoski, J.L. Lessard, and J.C. Bulinski. 1986. Immunolocalization of the gamma isoform of nonmuscle actin in cultured cells. J Cell Biol. 102:1726-37. Otterbein, L.R., P. Graceffa, and R. Dominguez. 2001. The crystal structure of uncomplexed actin in the ADP state. Science. 293:708-11. Padrick, S.B., and M.K. Rosen. 2010. Physical mechanisms of signal integration by WASP family proteins. Annu Rev Biochem. 79:707-35. Pantaloni, D., and M.F. Carlier. 1993. How profilin promotes actin filament assembly in the presence of thymosin beta 4. Cell. 75:1007-14. Paquin, N., and P. Chartrand. 2008. Local regulation of mRNA translation: new insights from the bud. Trends Cell Biol. 18:105-11. Paul, A.S., and T.D. Pollard. 2009. Energetic requirements for processive elongation of actin filaments by FH1FH2-formins. J Biol Chem. 284:12533-40. Pechstein, A., O. Shupliakov, and V. Haucke. Intersectin 1: a versatile actor in the synaptic vesicle cycle. Biochem Soc Trans. 38:181-6. Pegtel, D.M., S.I. Ellenbroek, A.E. Mertens, R.A. van der Kammen, J. de Rooij, and J.G. Collard. 2007. The Par-Tiam1 complex controls persistent migration by stabilizing microtubule-dependent front-rear polarity. Curr Biol. 17:1623-34. Pellegrin, S., and H. Mellor. 2005. The Rho family GTPase Rif induces filopodia through mDia2. Curr Biol. 15:129-33. Percipalle, P., J. Zhao, B. Pope, A. Weeds, U. Lindberg, and B. Daneholt. 2001. Actin bound to the heterogeneous nuclear ribonucleoprotein hrp36 is associated with Balbiani ring mRNA from the gene to polysomes. J Cell Biol. 153:229-36. Pilo Boyl, P., A. Di Nardo, C. Mulle, M. Sassoe-Pognetto, P. Panzanelli, A. Mele, M. Kneussel, V. Costantini, E. Perlas, M. Massimi, H. Vara, M. Giustetto, and W. Witke. 2007. Profilin2 contributes to synaptic vesicle exocytosis, neuronal excitability, and novelty-seeking behavior. EMBO J. 26:2991-3002. Pinyol, R., A. Haeckel, A. Ritter, B. Qualmann, and M.M. Kessels. 2007. Regulation of N-WASP and the Arp2/3 complex by Abp1 controls neuronal morphology. PLoS One. 2:e400. Plantard, L., A. Arjonen, J.G. Lock, G. Nurani, J. Ivaska, and S. Stromblad. 2010. PtdIns(3,4,5)P is a regulator of myosin-X localization and filopodia formation. J Cell Sci. 123:3525-34. Pocha, S.M., and G.O. Cory. 2009. WAVE2 is regulated by multiple phosphorylation events within its VCA domain. Cell Motil Cytoskeleton. 66:36-47. Pollard, T.D. 2010. Mechanics of cytokinesis in eukaryotes. Curr Opin Cell Biol. 22:50-6. Pollard, T.D., and G.G. Borisy. 2003. Cellular motility driven by assembly and disassembly of actin filaments. Cell. 112:453-65. Pollard, T.D., and J.A. Cooper. 1984. Quantitative analysis of the effect of Acanthamoeba profilin on actin filament nucleation and elongation. Biochemistry. 23:6631-41. Pollard, T.D., and J.A. Cooper. 2009. Actin, a central player in cell shape and movement. Science. 326:1208-12.

112

Ponti, A., M. Machacek, S.L. Gupton, C.M. Waterman-Storer, and G. Danuser. 2004. Two distinct actin networks drive the protrusion of migrating cells. Science. 305:1782-6. Pontrello, C.G., and I.M. Ethell. 2009. Accelerators, Brakes, and Gears of Actin Dynamics in Dendritic Spines. Open Neurosci J. 3:67-86. Pope, B.J., S.M. Gonsior, S. Yeoh, A. McGough, and A.G. Weeds. 2000. Uncoupling actin filament fragmentation by cofilin from increased subunit turnover. J Mol Biol. 298:649-61. Posern, G., and R. Treisman. 2006. Actin' together: serum response factor, its cofactors and the link to signal transduction. Trends Cell Biol. 16:588-96. Poskanzer, K.E., K.W. Marek, S.T. Sweeney, and G.W. Davis. 2003. Synaptotagmin I is necessary for compensatory synaptic vesicle endocytosis in vivo. Nature. 426:559-63. Pring, M., M. Evangelista, C. Boone, C. Yang, and S.H. Zigmond. 2003. Mechanism of formin-induced nucleation of actin filaments. Biochemistry. 42:486-96. Quinlan, M.E., S. Hilgert, A. Bedrossian, R.D. Mullins, and E. Kerkhoff. 2007. Regulatory interactions between two actin nucleators, Spire and Cappuccino. J Cell Biol. 179:117-28. Racz, B., and R.J. Weinberg. 2008. Organization of the Arp2/3 complex in hippocampal spines. J Neurosci. 28:5654-9. Raftopoulou, M., and A. Hall. 2004. Cell migration: Rho GTPases lead the way. Dev Biol. 265:23-32. Raucher, D., T. Stauffer, W. Chen, K. Shen, S. Guo, J.D. York, M.P. Sheetz, and T. Meyer. 2000. Phosphatidylinositol 4,5-bisphosphate functions as a second messenger that regulates cytoskeleton-plasma membrane adhesion. Cell. 100:221-8. Reinhard, M., K. Giehl, K. Abel, C. Haffner, T. Jarchau, V. Hoppe, B.M. Jockusch, and U. Walter. 1995. The proline-rich focal adhesion and microfilament protein VASP is a ligand for profilins. EMBO J. 14:1583-9. Reinhard, M., M. Halbrugge, U. Scheer, C. Wiegand, B.M. Jockusch, and U. Walter. 1992. The 46/50 kDa phosphoprotein VASP purified from human platelets is a novel protein associated with actin filaments and focal contacts. EMBO J. 11:2063- 70. Renner, M., C.G. Specht, and A. Triller. 2008. Molecular dynamics of postsynaptic receptors and scaffold proteins. Curr Opin Neurobiol. 18:532-40. Richmond, J.E., R.M. Weimer, and E.M. Jorgensen. 2001. An open form of syntaxin bypasses the requirement for UNC-13 in vesicle priming. Nature. 412:338-41. Rickman, C., C.N. Medine, A.R. Dun, D.J. Moulton, O. Mandula, N.D. Halemani, S.O. Rizzoli, L.H. Chamberlain, and R.R. Duncan. 2010. t-SNARE protein conformations patterned by the lipid microenvironment. J Biol Chem. 285:13535-41. Ridley, A.J., and A. Hall. 1992. Distinct patterns of actin organization regulated by the small GTP-binding proteins Rac and Rho. Cold Spring Harb Symp Quant Biol. 57:661- 71. Ridley, A.J., M.A. Schwartz, K. Burridge, R.A. Firtel, M.H. Ginsberg, G. Borisy, J.T. Parsons, and A.R. Horwitz. 2003. Cell migration: integrating signals from front to back. Science. 302:1704-9. Rodriguez, A.J., S.M. Shenoy, R.H. Singer, and J. Condeelis. 2006. Visualization of mRNA translation in living cells. J Cell Biol. 175:67-76. Rohatgi, R., H.Y. Ho, and M.W. Kirschner. 2000. Mechanism of N-WASP activation by CDC42 and phosphatidylinositol 4, 5-bisphosphate. J Cell Biol. 150:1299-310.

113

Rohatgi, R., L. Ma, H. Miki, M. Lopez, T. Kirchhausen, T. Takenawa, and M.W. Kirschner. 1999. The interaction between N-WASP and the Arp2/3 complex links Cdc42-dependent signals to actin assembly. Cell. 97:221-31. Romero, S., D. Didry, E. Larquet, N. Boisset, D. Pantaloni, and M.F. Carlier. 2007. How ATP hydrolysis controls filament assembly from profilin-actin: implication for formin processivity. J Biol Chem. 282:8435-45. Romero, S., C. Le Clainche, D. Didry, C. Egile, D. Pantaloni, and M.F. Carlier. 2004. Formin is a processive motor that requires profilin to accelerate actin assembly and associated ATP hydrolysis. Cell. 119:419-29. Ross, J.L., M.Y. Ali, and D.M. Warshaw. 2008. Cargo transport: molecular motors navigate a complex cytoskeleton. Curr Opin Cell Biol. 20:41-7. Roth, S., and J.A. Lynch. 2009. Symmetry breaking during Drosophila oogenesis. Cold Spring Harb Perspect Biol. 1:a001891. Rottner, K., B. Behrendt, J.V. Small, and J. Wehland. 1999a. VASP dynamics during lamellipodia protrusion. Nat Cell Biol. 1:321-2. Rottner, K., A. Hall, and J.V. Small. 1999b. Interplay between Rac and Rho in the control of substrate contact dynamics. Curr Biol. 9:640-8. Rottner, K., J. Hanisch, and K.G. Campellone. 2010. WASH, WHAMM and JMY: regulation of Arp2/3 complex and beyond. Trends Cell Biol. 20:650-61. Rozelle, A.L., L.M. Machesky, M. Yamamoto, M.H. Driessens, R.H. Insall, M.G. Roth, K. Luby-Phelps, G. Marriott, A. Hall, and H.L. Yin. 2000. Phosphatidylinositol 4,5- bisphosphate induces actin-based movement of raft-enriched vesicles through WASP-Arp2/3. Curr Biol. 10:311-20. Rubenstein, P.A. 1990. The functional importance of multiple actin isoforms. Bioessays. 12:309-15. Rufener, E., A.A. Frazier, C.M. Wieser, A. Hinderliter, and D.S. Cafiso. 2005. Membrane-bound orientation and position of the synaptotagmin C2B domain determined by site-directed spin labeling. Biochemistry. 44:18-28. Rust, M.B., C.B. Gurniak, M. Renner, H. Vara, L. Morando, A. Gorlich, M. Sassoe- Pognetto, M.A. Banchaabouchi, M. Giustetto, A. Triller, D. Choquet, and W. Witke. 2010. Learning, AMPA receptor mobility and synaptic plasticity depend on n- cofilin-mediated actin dynamics. EMBO J. 29:1889-902. Sakaba, T., A. Stein, R. Jahn, and E. Neher. 2005. Distinct kinetic changes in neurotransmitter release after SNARE protein cleavage. Science. 309:491-4. Salerno, V.P., A. Calliari, D.W. Provance, Jr., J.R. Sotelo-Silveira, J.R. Sotelo, and J.A. Mercer. 2008. Myosin-Va mediates RNA distribution in primary fibroblasts from multiple organs. Cell Motil Cytoskeleton. 65:422-33. Sankaranarayanan, S., P.P. Atluri, and T.A. Ryan. 2003. Actin has a molecular scaffolding, not propulsive, role in presynaptic function. Nat Neurosci. 6:127-35. Sarmiento, C., W. Wang, A. Dovas, H. Yamaguchi, M. Sidani, M. El-Sibai, V. Desmarais, H.A. Holman, S. Kitchen, J.M. Backer, A. Alberts, and J. Condeelis. 2008. WASP family members and formin proteins coordinate regulation of cell protrusions in carcinoma cells. J Cell Biol. 180:1245-60. Sathish, K., B. Padma, V. Munugalavadla, V. Bhargavi, K.V. Radhika, R. Wasia, M. Sairam, and S.S. Singh. 2004. Phosphorylation of profilin regulates its interaction with actin and poly (L-proline). Cell Signal. 16:589-96. Sawada, Y., M. Tamada, B.J. Dubin-Thaler, O. Cherniavskaya, R. Sakai, S. Tanaka, and M.P. Sheetz. 2006. Force sensing by mechanical extension of the Src family kinase substrate p130Cas. Cell. 127:1015-26.

114

Schikorski, T., and C.F. Stevens. 1997. Quantitative ultrastructural analysis of hippocampal excitatory synapses. J Neurosci. 17:5858-67. Schirenbeck, A., T. Bretschneider, R. Arasada, M. Schleicher, and J. Faix. 2005. The Diaphanous-related formin dDia2 is required for the formation and maintenance of filopodia. Nat Cell Biol. 7:619-25. Schlager, M.A., and C.C. Hoogenraad. 2009. Basic mechanisms for recognition and transport of synaptic cargos. Mol Brain. 2:25. Schuler, H., R. Karlsson, et al. . 2006. The connection between actin ATPase and polymerization. In Advances in Molecular and Cellular Biology. Aspects of the Cytoskeleton. Vol. 37. K.a. Bittar., editor. Elsevier/Academic Press., San Diego,. Schutt, C.E., J.C. Myslik, M.D. Rozycki, N.C. Goonesekere, and U. Lindberg. 1993. The structure of crystalline profilin-beta-actin. Nature. 365:810-6. Schutt, C.E., M.D. Rozycki, J.K. Chik, and U. Lindberg. 1995. Structural studies on the ribbon-to-helix transition in profilin: actin crystals. Biophys J. 68:12S-17S; discussion 17S-18S. Schwartz, M.A. 2010. Integrins and extracellular matrix in mechanotransduction. Cold Spring Harb Perspect Biol. 2:a005066. Schweizer, F.E., and T.A. Ryan. 2006. The synaptic vesicle: cycle of exocytosis and endocytosis. Curr Opin Neurobiol. 16:298-304. Scita, G., S. Confalonieri, P. Lappalainen, and S. Suetsugu. 2008. IRSp53: crossing the road of membrane and actin dynamics in the formation of membrane protrusions. Trends Cell Biol. 18:52-60. Sechi, A.S., and J. Wehland. 2000. The actin cytoskeleton and plasma membrane connection: PtdIns(4,5)P(2) influences cytoskeletal protein activity at the plasma membrane. J Cell Sci. 113 Pt 21:3685-95. Shapira, M., R.G. Zhai, T. Dresbach, T. Bresler, V.I. Torres, E.D. Gundelfinger, N.E. Ziv, and C.C. Garner. 2003. Unitary assembly of presynaptic active zones from Piccolo-Bassoon transport vesicles. Neuron. 38:237-52. Sharma, A., A. Lambrechts, T. Hao le, T.T. Le, C.A. Sewry, C. Ampe, A.H. Burghes, and G.E. Morris. 2005. A role for complexes of survival of motor neurons (SMN) protein with gemins and profilin in neurite-like cytoplasmic extensions of cultured nerve cells. Exp Cell Res. 309:185-97. Shattil, S.J., C. Kim, and M.H. Ginsberg. 2010. The final steps of integrin activation: the end game. Nat Rev Mol Cell Biol. 11:288-300. Shen, W., B. Wu, Z. Zhang, Y. Dou, Z.R. Rao, Y.R. Chen, and S. Duan. 2006. Activity- induced rapid synaptic maturation mediated by presynaptic cdc42 signaling. Neuron. 50:401-14. Shen, Z., N. Paquin, A. Forget, and P. Chartrand. 2009. Nuclear shuttling of She2p couples ASH1 mRNA localization to its translational repression by recruiting Loc1p and Puf6p. Mol Biol Cell. 20:2265-75. Shen, Z., A. St-Denis, and P. Chartrand. 2010. Cotranscriptional recruitment of She2p by RNA pol II elongation factor Spt4-Spt5/DSIF promotes mRNA localization to the yeast bud. Genes Dev. 24:1914-26. Sheng, M., and C.C. Hoogenraad. 2007. The postsynaptic architecture of excitatory synapses: a more quantitative view. Annu Rev Biochem. 76:823-47. Shestakova, E.A., R.H. Singer, and J. Condeelis. 2001. The physiological significance of beta -actin mRNA localization in determining cell polarity and directional motility. Proc Natl Acad Sci U S A. 98:7045-50.

115

Shi, Y., C.G. Pontrello, K.A. DeFea, L.F. Reichardt, and I.M. Ethell. 2009. Focal adhesion kinase acts downstream of EphB receptors to maintain mature dendritic spines by regulating cofilin activity. J Neurosci. 29:8129-42. Shimada, A., H. Niwa, K. Tsujita, S. Suetsugu, K. Nitta, K. Hanawa-Suetsugu, R. Akasaka, Y. Nishino, M. Toyama, L. Chen, Z.J. Liu, B.C. Wang, M. Yamamoto, T. Terada, A. Miyazawa, A. Tanaka, S. Sugano, M. Shirouzu, K. Nagayama, T. Takenawa, and S. Yokoyama. 2007. Curved EFC/F-BAR-domain dimers are joined end to end into a filament for membrane invagination in endocytosis. Cell. 129:761- 72. Shtrahman, M., C. Yeung, D.W. Nauen, G.Q. Bi, and X.L. Wu. 2005. Probing vesicle dynamics in single hippocampal synapses. Biophys J. 89:3615-27. Shupliakov, O., O. Bloom, J.S. Gustafsson, O. Kjaerulff, P. Low, N. Tomilin, V.A. Pieribone, P. Greengard, and L. Brodin. 2002. Impaired recycling of synaptic vesicles after acute perturbation of the presynaptic actin cytoskeleton. Proc Natl Acad Sci U S A. 99:14476-81. Sidani, M., D. Wessels, G. Mouneimne, M. Ghosh, S. Goswami, C. Sarmiento, W. Wang, S. Kuhl, M. El-Sibai, J.M. Backer, R. Eddy, D. Soll, and J. Condeelis. 2007. Cofilin determines the migration behavior and turning frequency of metastatic cancer cells. J Cell Biol. 179:777-91. Siksou, L., A. Triller, and S. Marty. 2009a. An emerging view of presynaptic structure from electron microscopic studies. J Neurochem. 108:1336-42. Siksou, L., F. Varoqueaux, O. Pascual, A. Triller, N. Brose, and S. Marty. 2009b. A common molecular basis for membrane docking and functional priming of synaptic vesicles. Eur J Neurosci. 30:49-56. Simons, K., and M.J. Gerl. 2010. Revitalizing membrane rafts: new tools and insights. Nat Rev Mol Cell Biol. 11:688-99. Singh, S.S., A. Chauhan, N. Murakami, and V.P. Chauhan. 1996a. Profilin and gelsolin stimulate phosphatidylinositol 3-kinase activity. Biochemistry. 35:16544-9. Singh, S.S., A. Chauhan, N. Murakami, J. Styles, M. Elzinga, and V.P. Chauhan. 1996b. Phosphoinositide-dependent in vitro phosphorylation of profilin by protein kinase C. Phospholipid specificity and localization of the phosphorylation site. Recept Signal Transduct. 6:77-86. Skare, P., and R. Karlsson. 2002. Evidence for two interaction regions for phosphatidylinositol(4,5)-bisphosphate on mammalian profilin I. FEBS Lett. 522:119-24. Skare, P., J.P. Kreivi, A. Bergstrom, and R. Karlsson. 2003. Profilin I colocalizes with speckles and Cajal bodies: a possible role in pre-mRNA splicing. Exp Cell Res. 286:12-21. Skau, C.T., E.M. Neidt, and D.R. Kovar. 2009. Role of tropomyosin in formin- mediated contractile ring assembly in fission yeast. Mol Biol Cell. 20:2160-73. Skene, J.H. 1989. Axonal growth-associated proteins. Annu Rev Neurosci. 12:127-56. Small, J.V. 1981. Organization of actin in the leading edge of cultured cells: influence of osmium tetroxide and dehydration on the ultrastructure of actin meshworks. J Cell Biol. 91:695-705. Small, J.V. 1994. Lamellipodia architecture: actin filament turnover and the lateral flow of actin filaments during motility. Semin Cell Biol. 5:157-63. Small, J.V. 2010. Dicing with dogma: de-branching the lamellipodium. Trends Cell Biol. 20:628-33.

116

Small, J.V., K. Rottner, I. Kaverina, and K.I. Anderson. 1998. Assembling an actin cytoskeleton for cell attachment and movement. Biochim Biophys Acta. 1404:271-81. Small, J.V., T. Stradal, E. Vignal, and K. Rottner. 2002. The lamellipodium: where motility begins. Trends Cell Biol. 12:112-20. Soderling, S.H., L.K. Langeberg, J.A. Soderling, S.M. Davee, R. Simerly, J. Raber, and J.D. Scott. 2003. Loss of WAVE-1 causes sensorimotor retardation and reduced learning and memory in mice. Proc Natl Acad Sci U S A. 100:1723-8. Sokac, A.M., C. Co, J. Taunton, and W. Bement. 2003. Cdc42-dependent actin polymerization during compensatory endocytosis in Xenopus eggs. Nat Cell Biol. 5:727-32. Sollner, T., M.K. Bennett, S.W. Whiteheart, R.H. Scheller, and J.E. Rothman. 1993a. A protein assembly-disassembly pathway in vitro that may correspond to sequential steps of synaptic vesicle docking, activation, and fusion. Cell. 75:409-18. Sollner, T., S.W. Whiteheart, M. Brunner, H. Erdjument-Bromage, S. Geromanos, P. Tempst, and J.E. Rothman. 1993b. SNAP receptors implicated in vesicle targeting and fusion. Nature. 362:318-24. Sotelo-Silveira, J., M. Crispino, A. Puppo, J.R. Sotelo, and E. Koenig. 2008. Myelinated axons contain beta-actin mRNA and ZBP-1 in periaxoplasmic ribosomal plaques and depend on cyclic AMP and F-actin integrity for in vitro translation. J Neurochem. 104:545-57. Star, E.N., D.J. Kwiatkowski, and V.N. Murthy. 2002. Rapid turnover of actin in dendritic spines and its regulation by activity. Nat Neurosci. 5:239-46. Steffen, A., J. Faix, G.P. Resch, J. Linkner, J. Wehland, J.V. Small, K. Rottner, and T.E. Stradal. 2006. Filopodia formation in the absence of functional WAVE- and Arp2/3-complexes. Mol Biol Cell. 17:2581-91. Steffen, A., K. Rottner, J. Ehinger, M. Innocenti, G. Scita, J. Wehland, and T.E. Stradal. 2004. Sra-1 and Nap1 link Rac to actin assembly driving lamellipodia formation. EMBO J. 23:749-59. Sternlicht, H., G.W. Farr, M.L. Sternlicht, J.K. Driscoll, K. Willison, and M.B. Yaffe. 1993. The t-complex polypeptide 1 complex is a chaperonin for tubulin and actin in vivo. Proc Natl Acad Sci U S A. 90:9422-6. Steward, O., and W.B. Levy. 1982. Preferential localization of polyribosomes under the base of dendritic spines in granule cells of the dentate gyrus. J Neurosci. 2:284-91. Stevens, C.F., and J.M. Sullivan. 2003. The synaptotagmin C2A domain is part of the calcium sensor controlling fast synaptic transmission. Neuron. 39:299-308. Stevens, J.M., E.E. Galyov, and M.P. Stevens. 2006. Actin-dependent movement of bacterial pathogens. Nat Rev Microbiol. 4:91-101. Stradal, T.E., and G. Scita. 2006. Protein complexes regulating Arp2/3-mediated actin assembly. Curr Opin Cell Biol. 18:4-10. Strasser, G.A., N.A. Rahim, K.E. VanderWaal, F.B. Gertler, and L.M. Lanier. 2004. Arp2/3 is a negative regulator of growth cone translocation. Neuron. 43:81-94. Sudhof, T.C., and J.E. Rothman. 2009. Membrane fusion: grappling with SNARE and SM proteins. Science. 323:474-7. Suetsugu, S., M. Hattori, H. Miki, T. Tezuka, T. Yamamoto, K. Mikoshiba, and T. Takenawa. 2002. Sustained activation of N-WASP through phosphorylation is essential for neurite extension. Dev Cell. 3:645-58. Suetsugu, S., H. Miki, and T. Takenawa. 1998. The essential role of profilin in the assembly of actin for microspike formation. EMBO J. 17:6516-26.

117

Suetsugu, S., K. Murayama, A. Sakamoto, K. Hanawa-Suetsugu, A. Seto, T. Oikawa, C. Mishima, M. Shirouzu, T. Takenawa, and S. Yokoyama. 2006. The RAC binding domain/IRSp53-MIM homology domain of IRSp53 induces RAC-dependent membrane deformation. J Biol Chem. 281:35347-58. Sukumaran, S.S., S. Banerjee, S. Bhasker, and A. Thekkuveettil. 2008. The cytoplasmic C2A domain of synaptotagmin shows sequence specific interaction with its own mRNA. Biochem Biophys Res Commun. 373:509-14. Svitkina, T.M., and G.G. Borisy. 1999. Arp2/3 complex and actin depolymerizing factor/cofilin in dendritic organization and treadmilling of actin filament array in lamellipodia. J Cell Biol. 145:1009-26. Svitkina, T.M., E.A. Bulanova, O.Y. Chaga, D.M. Vignjevic, S. Kojima, J.M. Vasiliev, and G.G. Borisy. 2003. Mechanism of filopodia initiation by reorganization of a dendritic network. J Cell Biol. 160:409-21. Symons, M., J.M. Derry, B. Karlak, S. Jiang, V. Lemahieu, F. McCormick, U. Francke, and A. Abo. 1996. Wiskott-Aldrich syndrome protein, a novel effector for the GTPase CDC42Hs, is implicated in actin polymerization. Cell. 84:723-34. Takano, K., K. Toyooka, and S. Suetsugu. 2008. EFC/F-BAR proteins and the N- WASP-WIP complex induce membrane curvature-dependent actin polymerization. EMBO J. 27:2817-28. Takenawa, T., and H. Miki. 2001. WASP and WAVE family proteins: key molecules for rapid rearrangement of cortical actin filaments and cell movement. J Cell Sci. 114:1801-9. Takenawa, T., and S. Suetsugu. 2007. The WASP-WAVE protein network: connecting the membrane to the cytoskeleton. Nat Rev Mol Cell Biol. 8:37-48. Tanaka, M., and H. Shibata. 1985. Poly(L-proline)-binding proteins from chick embryos are a profilin and a profilactin. Eur J Biochem. 151:291-7. Tang, J., A. Maximov, O.H. Shin, H. Dai, J. Rizo, and T.C. Sudhof. 2006. A complexin/synaptotagmin 1 switch controls fast synaptic vesicle exocytosis. Cell. 126:1175-87. Tang, N., and E.M. Ostap. 2001. Motor domain-dependent localization of myo1b (myr- 1). Curr Biol. 11:1131-5. Tashiro, A., A. Dunaevsky, R. Blazeski, C.A. Mason, and R. Yuste. 2003. Bidirectional regulation of hippocampal mossy fiber filopodial motility by kainate receptors: a two-step model of synaptogenesis. Neuron. 38:773-84. Taunton, J., B.A. Rowning, M.L. Coughlin, M. Wu, R.T. Moon, T.J. Mitchison, and C.A. Larabell. 2000. Actin-dependent propulsion of endosomes and lysosomes by recruitment of N-WASP. J Cell Biol. 148:519-30. Tiruchinapalli, D.M., Y. Oleynikov, S. Kelic, S.M. Shenoy, A. Hartley, P.K. Stanton, R.H. Singer, and G.J. Bassell. 2003. Activity-dependent trafficking and dynamic localization of zipcode binding protein 1 and beta-actin mRNA in dendrites and spines of hippocampal neurons. J Neurosci. 23:3251-61. Tobacman, L.S., and E.D. Korn. 1982. The regulation of actin polymerization and the inhibition of monomeric actin ATPase activity by Acanthamoeba profilin. J Biol Chem. 257:4166-70. Tokuo, H., and M. Ikebe. 2004. Myosin X transports Mena/VASP to the tip of filopodia. Biochem Biophys Res Commun. 319:214-20. Torres, E., and M.K. Rosen. 2003. Contingent phosphorylation/dephosphorylation provides a mechanism of molecular memory in WASP. Mol Cell. 11:1215-27.

118

Totsukawa, G., Y. Yamakita, S. Yamashiro, D.J. Hartshorne, Y. Sasaki, and F. Matsumura. 2000. Distinct roles of ROCK (Rho-kinase) and MLCK in spatial regulation of MLC phosphorylation for assembly of stress fibers and focal adhesions in 3T3 fibroblasts. J Cell Biol. 150:797-806. Trifaro, J., S.D. Rose, T. Lejen, and A. Elzagallaai. 2000. Two pathways control chromaffin cell cortical F-actin dynamics during exocytosis. Biochimie. 82:339-52. Tucker, W.C., T. Weber, and E.R. Chapman. 2004. Reconstitution of Ca2+-regulated membrane fusion by synaptotagmin and SNAREs. Science. 304:435-8. Ubach, J., X. Zhang, X. Shao, T.C. Sudhof, and J. Rizo. 1998. Ca2+ binding to synaptotagmin: how many Ca2+ ions bind to the tip of a C2-domain? EMBO J. 17:3921-30. Wallar, B.J., and A.S. Alberts. 2003. The formins: active scaffolds that remodel the cytoskeleton. Trends Cell Biol. 13:435-46. van Rheenen, J., X. Song, W. van Roosmalen, M. Cammer, X. Chen, V. Desmarais, S.C. Yip, J.M. Backer, R.J. Eddy, and J.S. Condeelis. 2007. EGF-induced PIP2 hydrolysis releases and activates cofilin locally in carcinoma cells. J Cell Biol. 179:1247-59. Van Troys, M., D. Dewitte, J.L. Verschelde, M. Goethals, J. Vandekerckhove, and C. Ampe. 2000. The competitive interaction of actin and PIP2 with actophorin is based on overlapping target sites: design of a gain-of-function mutant. Biochemistry. 39:12181-9. Wang, C.T., J. Bai, P.Y. Chang, E.R. Chapman, and M.B. Jackson. 2006a. Synaptotagmin-Ca2+ triggers two sequential steps in regulated exocytosis in rat PC12 cells: fusion pore opening and fusion pore dilation. J Physiol. 570:295-307. Wang, W., G. Mouneimne, M. Sidani, J. Wyckoff, X. Chen, A. Makris, S. Goswami, A.R. Bresnick, and J.S. Condeelis. 2006b. The activity status of cofilin is directly related to invasion, intravasation, and metastasis of mammary tumors. J Cell Biol. 173:395-404. Wang, X., M. Kibschull, M.M. Laue, B. Lichte, E. Petrasch-Parwez, and M.W. Kilimann. 1999. Aczonin, a 550-kD putative scaffolding protein of presynaptic active zones, shares homology regions with Rim and Bassoon and binds profilin. J Cell Biol. 147:151-62. Vartiainen, M.K., S. Guettler, B. Larijani, and R. Treisman. 2007. Nuclear actin regulates dynamic subcellular localization and activity of the SRF cofactor MAL. Science. 316:1749-52. Watanabe, M., K. Nomura, A. Ohyama, R. Ishikawa, Y. Komiya, K. Hosaka, E. Yamauchi, H. Taniguchi, N. Sasakawa, K. Kumakura, T. Ushiki, O. Sato, M. Ikebe, and M. Igarashi. 2005. Myosin-Va regulates exocytosis through the submicromolar Ca2+-dependent binding of syntaxin-1A. Mol Biol Cell. 16:4519-30. Watanabe, N., P. Madaule, T. Reid, T. Ishizaki, G. Watanabe, A. Kakizuka, Y. Saito, K. Nakao, B.M. Jockusch, and S. Narumiya. 1997. p140mDia, a mammalian homolog of Drosophila diaphanous, is a target protein for Rho small GTPase and is a ligand for profilin. EMBO J. 16:3044-56. Watanabe, T.M., H. Tokuo, K. Gonda, H. Higuchi, and M. Ikebe. 2010. Myosin-X induces filopodia by multiple elongation mechanism. J Biol Chem. 285:19605-14. Webb, D.J., K. Donais, L.A. Whitmore, S.M. Thomas, C.E. Turner, J.T. Parsons, and A.F. Horwitz. 2004. FAK-Src signalling through paxillin, ERK and MLCK regulates adhesion disassembly. Nat Cell Biol. 6:154-61.

119

Wegner, A.M., C.A. Nebhan, L. Hu, D. Majumdar, K.M. Meier, A.M. Weaver, and D.J. Webb. 2008. N-wasp and the arp2/3 complex are critical regulators of actin in the development of dendritic spines and synapses. J Biol Chem. 283:15912-20. Wei, C., X. Wang, M. Chen, K. Ouyang, L.S. Song, and H. Cheng. 2009. Calcium flickers steer cell migration. Nature. 457:901-5. Welch, M.D., and R.D. Mullins. 2002. Cellular control of actin nucleation. Annu Rev Cell Dev Biol. 18:247-88. Weninger, K., M.E. Bowen, U.B. Choi, S. Chu, and A.T. Brunger. 2008. Accessory proteins stabilize the acceptor complex for synaptobrevin, the 1:1 syntaxin/SNAP- 25 complex. Structure. 16:308-20. Wennerberg, K., and C.J. Der. 2004. Rho-family GTPases: it's not only Rac and Rho (and I like it). J Cell Sci. 117:1301-12. Verstreken, P., C.V. Ly, K.J. Venken, T.W. Koh, Y. Zhou, and H.J. Bellen. 2005. Synaptic mitochondria are critical for mobilization of reserve pool vesicles at Drosophila neuromuscular junctions. Neuron. 47:365-78. Westphal, R.S., S.H. Soderling, N.M. Alto, L.K. Langeberg, and J.D. Scott. 2000. Scar/WAVE-1, a Wiskott-Aldrich syndrome protein, assembles an actin-associated multi-kinase scaffold. EMBO J. 19:4589-600. Vicente-Manzanares, M., X. Ma, R.S. Adelstein, and A.R. Horwitz. 2009. Non-muscle myosin II takes centre stage in cell adhesion and migration. Nat Rev Mol Cell Biol. 10:778-90. Vikesaa, J., T.V. Hansen, L. Jonson, R. Borup, U.M. Wewer, J. Christiansen, and F.C. Nielsen. 2006. RNA-binding IMPs promote cell adhesion and invadopodia formation. EMBO J. 25:1456-68. Willis, D.E., E.A. van Niekerk, Y. Sasaki, M. Mesngon, T.T. Merianda, G.G. Williams, M. Kendall, D.S. Smith, G.J. Bassell, and J.L. Twiss. 2007. Extracellular stimuli specifically regulate localized levels of individual neuronal mRNAs. J Cell Biol. 178:965-80. Wills, Z., L. Marr, K. Zinn, C.S. Goodman, and D. Van Vactor. 1999. Profilin and the Abl tyrosine kinase are required for motor axon outgrowth in the Drosophila embryo. Neuron. 22:291-9. Visa, N., and P. Percipalle. Nuclear functions of actin. 2010. Cold Spring Harb Perspect Biol. 2:a000620. Witke, W. 2004. The role of profilin complexes in cell motility and other cellular processes. Trends Cell Biol. 14:461-9. Witke, W., A.V. Podtelejnikov, A. Di Nardo, J.D. Sutherland, C.B. Gurniak, C. Dotti, and M. Mann. 1998. In mouse brain profilin I and profilin II associate with regulators of the endocytic pathway and actin assembly. EMBO J. 17:967-76. Witke, W., J.D. Sutherland, A. Sharpe, M. Arai, and D.J. Kwiatkowski. 2001. Profilin I is essential for cell survival and cell division in early mouse development. Proc Natl Acad Sci U S A. 98:3832-6. Wolfenson, H., A. Bershadsky, Y.I. Henis, and B. Geiger. 2011. Actomyosin-generated tension controls the molecular kinetics of focal adhesions. J Cell Sci. 124:1425-32. Volkman, B.F., K.E. Prehoda, J.A. Scott, F.C. Peterson, and W.A. Lim. 2002. Structure of the N-WASP EVH1 domain-WIP complex: insight into the molecular basis of Wiskott-Aldrich Syndrome. Cell. 111:565-76.

120

Worth, D.C., K. Hodivala-Dilke, S.D. Robinson, S.J. King, P.E. Morton, F.B. Gertler, M.J. Humphries, and M. Parsons. 2010. Alpha v beta3 integrin spatially regulates VASP and RIAM to control adhesion dynamics and migration. J Cell Biol. 189:369- 83. Xu, J., T. Mashimo, and T.C. Sudhof. 2007. Synaptotagmin-1, -2, and -9: Ca(2+) sensors for fast release that specify distinct presynaptic properties in subsets of neurons. Neuron. 54:567-81. Xue, M., C. Ma, T.K. Craig, C. Rosenmund, and J. Rizo. 2008. The Janus-faced nature of the C(2)B domain is fundamental for synaptotagmin-1 function. Nat Struct Mol Biol. 15:1160-8. Yamada, S., and W.J. Nelson. 2007. Localized zones of Rho and Rac activities drive initiation and expansion of epithelial cell-cell adhesion. J Cell Biol. 178:517-27. Yamagishi, A., M. Masuda, T. Ohki, H. Onishi, and N. Mochizuki. 2004. A novel actin bundling/filopodium-forming domain conserved in insulin receptor tyrosine kinase substrate p53 and missing in metastasis protein. J Biol Chem. 279:14929-36. Yamazaki, D., S. Suetsugu, H. Miki, Y. Kataoka, S. Nishikawa, T. Fujiwara, N. Yoshida, and T. Takenawa. 2003. WAVE2 is required for directed cell migration and cardiovascular development. Nature. 424:452-6. Yan, C., N. Martinez-Quiles, S. Eden, T. Shibata, F. Takeshima, R. Shinkura, Y. Fujiwara, R. Bronson, S.B. Snapper, M.W. Kirschner, R. Geha, F.S. Rosen, and F.W. Alt. 2003. WAVE2 deficiency reveals distinct roles in embryogenesis and Rac- mediated actin-based motility. EMBO J. 22:3602-12. Yang, C., L. Czech, S. Gerboth, S. Kojima, G. Scita, and T. Svitkina. 2007. Novel roles of formin mDia2 in lamellipodia and filopodia formation in motile cells. PLoS Biol. 5:e317. Yang, L., L. Wang, and Y. Zheng. 2006. Gene targeting of Cdc42 and Cdc42GAP affirms the critical involvement of Cdc42 in filopodia induction, directed migration, and proliferation in primary mouse embryonic fibroblasts. Mol Biol Cell. 17:4675-85. Yao, J., A. Nowack, P. Kensel-Hammes, R.G. Gardner, and S.M. Bajjalieh. 2010. Cotrafficking of SV2 and synaptotagmin at the synapse. J Neurosci. 30:5569-78. Yao, J., J. Qi, and G. Chen. 2006. Actin-dependent activation of presynaptic silent synapses contributes to long-term synaptic plasticity in developing hippocampal neurons. J Neurosci. 26:8137-47. Yarar, D., J.A. D'Alessio, R.L. Jeng, and M.D. Welch. 2002. Motility determinants in WASP family proteins. Mol Biol Cell. 13:4045-59. Yin, H.L., and P.A. Janmey. 2003. Phosphoinositide regulation of the actin cytoskeleton. Annu Rev Physiol. 65:761-89. Yonetani, A., and F. Chang. 2010. Regulation of cytokinesis by the formin cdc12p. Curr Biol. 20:561-6. Yoon, B.C., K.H. Zivraj, and C.E. Holt. 2009. Local translation and mRNA trafficking in axon pathfinding. Results Probl Cell Differ. 48:269-88. Yoshihara, M., and J.T. Littleton. 2002. Synaptotagmin I functions as a calcium sensor to synchronize neurotransmitter release. Neuron. 36:897-908. Yu, H.Y., and W.M. Bement. 2007. Control of local actin assembly by membrane fusion-dependent compartment mixing. Nat Cell Biol. 9:149-59. Zaidel-Bar, R., S. Itzkovitz, A. Ma'ayan, R. Iyengar, and B. Geiger. 2007. Functional atlas of the integrin adhesome. Nat Cell Biol. 9:858-67.

121

Zalevsky, J., L. Lempert, H. Kranitz, and R.D. Mullins. 2001. Different WASP family proteins stimulate different Arp2/3 complex-dependent actin-nucleating activities. Curr Biol. 11:1903-13. Zhai, R.G., H. Vardinon-Friedman, C. Cases-Langhoff, B. Becker, E.D. Gundelfinger, N.E. Ziv, and C.C. Garner. 2001. Assembling the presynaptic active zone: a characterization of an active one precursor vesicle. Neuron. 29:131-43. Zhang, H., J.S. Berg, Z. Li, Y. Wang, P. Lang, A.D. Sousa, A. Bhaskar, R.E. Cheney, and S. Stromblad. 2004. Myosin-X provides a motor-based link between integrins and the cytoskeleton. Nat Cell Biol. 6:523-31. Zhang, W., and D.L. Benson. 2001. Stages of synapse development defined by dependence on F-actin. J Neurosci. 21:5169-81. Zhang, W., and D.L. Benson. 2002. Developmentally regulated changes in cellular compartmentation and synaptic distribution of actin in hippocampal neurons. J Neurosci Res. 69:427-36. Zhang, X., M.J. Kim-Miller, M. Fukuda, J.A. Kowalchyk, and T.F. Martin. 2002. Ca2+- dependent synaptotagmin binding to SNAP-25 is essential for Ca2+-triggered exocytosis. Neuron. 34:599-611. Zhou, L., S.J. Martinez, M. Haber, E.V. Jones, D. Bouvier, G. Doucet, A.T. Corera, E.A. Fon, A.H. Zisch, and K.K. Murai. 2007. EphA4 signaling regulates phospholipase Cgamma1 activation, cofilin membrane association, and dendritic spine morphology. J Neurosci. 27:5127-38. Zhou, Q., M. Xiao, and R.A. Nicoll. 2001. Contribution of cytoskeleton to the internalization of AMPA receptors. Proc Natl Acad Sci U S A. 98:1261-6. Zhu, X.J., C.Z. Wang, P.G. Dai, Y. Xie, N.N. Song, Y. Liu, Q.S. Du, L. Mei, Y.Q. Ding, and W.C. Xiong. 2007. Myosin X regulates netrin receptors and functions in axonal path-finding. Nat Cell Biol. 9:184-92. Zhuravlev, P.I., B.S. Der, and G.A. Papoian. 2010. Design of active transport must be highly intricate: a possible role of myosin and Ena/VASP for G-actin transport in filopodia. Biophys J. 98:1439-48. Zicha, D., I.M. Dobbie, M.R. Holt, J. Monypenny, D.Y. Soong, C. Gray, and G.A. Dunn. 2003. Rapid actin transport during cell protrusion. Science. 300:142-5. Zigmond, S.H. 1993. Recent quantitative studies of actin filament turnover during cell locomotion. Cell Motil Cytoskeleton. 25:309-16. Zigmond, S.H. 2000. How WASP regulates actin polymerization. J Cell Biol. 150:F117- 20. Zigmond, S.H. 2004. Formin-induced nucleation of actin filaments. Curr Opin Cell Biol. 16:99-105. Zou, L., Z. Ding, and P. Roy. 2010. Profilin-1 overexpression inhibits proliferation of MDA-MB-231 breast cancer cells partly through p27kip1 upregulation. J Cell Physiol. 223:623-9. Zuchero, J.B., A.S. Coutts, M.E. Quinlan, N.B. Thangue, and R.D. Mullins. 2009. p53- cofactor JMY is a multifunctional actin nucleation factor. Nat Cell Biol. 11:451-9. Zuo, X., J. Zhang, Y. Zhang, S.C. Hsu, D. Zhou, and W. Guo. 2006. Exo70 interacts with the Arp2/3 complex and regulates cell migration. Nat Cell Biol. 8:1383-8.

122