Structural Characterization of Two Different F- Actin Cross-Linking Proteins by Electron Microscopy Cheri M

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2006 Reading Between the Filaments: Structural Characterization of Two Different F- Actin Cross-Linking Proteins by Electron Microscopy Cheri M. Hampton

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THE FLORIDA STATE UNIVERSITY

COLLEGE OF ARTS AND SCIENCES

READING BETWEEN THE FILAMENTS: STRUCTURAL CHARACTERIZATION OF TWO DIFFERENT F-ACTIN CROSS-LINKING PROTEINS BY ELECTRON MICROSCOPY

CHERI M. HAMPTON

A Dissertation submitted to the Institute of Molecular Biophysics in partial fulfillment of the requirements for the degree of Doctor of Philosophy

Degree Awarded: Fall Semester, 2006

The members of the Committee approve the Dissertation of Cheri M. Hampton defended on August 11th, 2006.

______Kenneth A. Taylor Professor Directing Dissertation

______Charles Ouimet Outside Committee Member

______P. Bryant Chase Committee Member

______Piotr Fajer Committee Member

______Hong Li Committee Member

Approved:

______

Timothy M Logan, Director, Institute of Molecular Biophysics

The Office of Graduate Studies has verified and approved the above named committee members.

ii TABLE OF CONTENTS

List of Tables ...... vi List of Figures ...... vii Abstract ...... x

1. Introduction to Actin Cross-linking Proteins and EM...... 1

1.1 The Cytoskeleton is a Cellular Scaffold...... 1 1.1.1 Actin Filaments in the Cytoskeleton ...... 1 1.1.2 Cell Structure and Tension ...... 2 1.1.3 Cell Structure and Signaling...... 3

1.2 Actin, Function and Structure ...... 6 1.2.1 Actin is the Most Highly Conserved Protein ...... 6 1.2.2 Actin Function ...... 6 1.2.3 Actin Structure ...... 7

1.3 The Spectrin Repeat Family, Function and Structure...... 10 1.3.1 Spectrin Repeat Family...... 10 1.3.2 α-Actinin Function ...... 14 1.3.3 α-Actinin Structure ...... 17

1.4 The Gelsolin Family, Function and Structure ...... 19 1.4.1 Actin Filament Severing, Capping, Nucleating,...... and Bundling Proteins...... 19 1.4.2 Villin Found Primarily in Microvilli...... 19 1.4.3 Villin is Structurally Similar to Gelsolin ...... 20 1.4.4 Gelsolin is Regulated by Calcium ...... 26

1.5 Electron Microscopy, Electron Tomography,...... And Image Analysis ...... 28 1.5.1 Electron Microscopy and Image Formation...... 28 1.5.2 The Fourier Transform ...... 29 1.5.3 The Contrast Transfer Function ...... 30 1.5.4 Tomography and the Projection Theorem...... 31 1.5.5 Averaging and Classification...... 32

2. α-Actinin is a Variable-Length Actin Cross-linker ...... 34

2.1 Background ...... 34

iii

2.2 Results ...... 37 2.2.1 Alignment and Correspondence Analysis ...... 37 2.2.2 Distribution of Cross-Link Angles and Lengths ...... 44 2.2.3 α-Actinin Binds to a Single Actin Filament ...... 49

2.3 Discussion...... 51 2.3.1 α-Actinin-Membrane Interactions ...... 52 2.3.2 Variability of Bipolar Cross-Links ...... 53 2.3.3 Significance of “Monofilament Binding”...... 54 2.3.4 Significance of Variable Length of Cross-Link...... 56

2.4 Materials and Methods...... 57 2.4.1 Protein Purification...... 57 2.4.2 EM Data Collection ...... 58 2.4.3 Image Processing ...... 58 2.4.4 Classification...... 58 2.2.5 Modeling ...... 59

3. α-Actinin Interacting Proteins ...... 61

3.1 The Alliance for Cellular Signaling...... 61

3.2 Molecule Page Navigation...... 62

3.3 α-Actinin-1 Molecule Pages ...... 68 3.3.1 α-Actinin-1 Overview...... 68 3.3.1 α-Actinin-1 Mini Molecule Page ...... 73

3.4 α-Actinin-2 Molecule Pages ...... 77 3.4.1 α-Actinin-2 Overview...... 77 3.4.2 α-Actinin-2 Summary ...... 79 3.4.3 α-Actinin-2 Network Map ...... 93 3.4.4 α-Actinin-2 States ...... 94 3.4.5 α-Actinin-2 Transitions ...... 96

3.5 α-Actinin-3 Molecule Pages ...... 98 3.5.1 α-Actinin-3 Overview...... 98 3.5.2 α-Actinin-3 Summary ...... 100 3.5.3 α-Actinin-3 Network Map ...... 104 3.5.4 α-Actinin-3 States ...... 105

iv 3.5.5 α-Actinin-3 Transitions ...... 106

3.6 α-Actinin-4 Molecule Pages ...... 107 3.6.1 α-Actinin-4 Overview...... 107 3.6.2 α-Actinin-4 Summary ...... 109 3.6.3 α-Actinin-4 Network Map ...... 118 3.6.4 α-Actinin-4 States ...... 119 3.6.5 α-Actinin-4 Transitions ...... 121

3.7 α-Actinin Interacting Proteins ...... 123

4. Villin in an Unusual Suspect in F-actin Cross-Linking ...... 135

4.1 Background ...... 135

4.2 Results ...... 138 4.2.1 Image Alignment and Volume Classification...... 138

4.2.2 Models ...... 143

4.3 Discussion...... 146 4.3.1 A Tool for Studying Actin: Actin-binding Protein Interactions 147 4.3.2 Significance of Villin-Actin Interaction Sites ...... 147 4.3.3 Indication for Multiple Modes of F-actin Binding...... 148

4.4 Materials and Methods...... 149 4.4.1 Protein Purification and Array Formation ...... 149 4.4.2 EM Data Collection ...... 149 4.4.3 Image Processing ...... 150 4.4.4 Classification...... 150 4.4.5 Modeling ...... 150

5. Summary and Future Directions...... 152

REFERENCES ...... 154

BIOGRAPHICAL SKETCH ...... 171

v LIST OF TABLES

Table 2.2.1.1: Assessment of Classification Accuracy ...... 43

Table 3.7.1: α-Actinin Interacting Proteins ...... 123

vi LIST OF FIGURES

Figure 1.1.1.1: Z Disc Architecture ...... 2

Figure 1.1.2:1: The Cytoskeletal Network ...... 3

Figure 1.1.3:1: The Focal Adhesion is a Site for Force-Regulated Cell Signaling ...... 5

Figure 1.1.3.2: Schematic of Protein Interactions Emanating from the Z Disc of the Muscle Sarcomere ...... 5

Figure 1.2.3.1: The Domain Organization of G-actin ...... 8

Figure 1.2.3.2: Domain Interaction in F-actin ...... 9

Figure 1.3.1.1: Comparison of CH Domains ...... 12

Figure 1.3.1.2: The Structure of EF-hand Domains ...... 14

Figure 1.3.3.1: Structure of the ABD from Human α-Actinin 1 ...... 18

Figure 1.3.3.2: Structure of the α-Actinin Rod ...... 18

Figure 1.4.2.1: Micrograph of a Section of Intestinal Epithelium Microvilli ...... 20

Figure 1.4.3.1: The Villin Domains ...... 21

Figure 1.4.3.2: Annotated Sequence Alignment of Villin and Gelsolin ... 23

Figure 1.4.4.1: Illustration of Two Possible Models for Gelsolin Severing and Capping F-actin ...... 27

Figure 1.5.1.1: Electron Path Through the Column of the Microscope... 28

Figure 2.2.1.1: α-Actinin-F-actin Raft ...... 38

Figure 2.2.1.2: A Global Average of 7,903 Actin Crossover Repeats and Eight Class Averages Based on the Variance ...... 40 Figure 2.2.1.3: Comparison of Whole Image Classification to

vii Left/Right Classification ...... 41

Figure 2.2.2.1: Angular Distributions of Total and Anti-parallel Cross-Links 45

Figure 2.2.2.2: Examples of 60°Cross-Links Realigned and Classified on α-Actinin ...... 46

Figure 2.2.2.3: Mapped-Back Images ...... 47

Figure 2.2.2.4: Map-Back Before and After Comparison ...... 48

Figure 2.2.2.5: Models of α-Actinin Cross-Linking a Single Actin Filament 50

Figure 3.2.1: Navigation Tool Bar ...... 63

Figure 3.2.2: Excerpt from the Protein Summary Page ...... 64

Figure 3.2.3: Excerpt from the States Page and Link to the Transitions Page ...... 65

Figure 3.2.4: Transition Page for the Association of ADAM-12 with ACTN2 66

Figure 3.2.5: Network Map for α-Actinin 3 ...... 67

Figure 3.3.1: α-Actinin 1 Overview Page...... 68

Figure 3.3.2: α-Actinin 1 Mini Molecule Page ...... 73

Figure 3.4.1: α-Actinin 2 Overview Page...... 77

Figure 3.4.2: α-Actinin 2 Summary Page ...... 79

Figure 3.4.3: α-Actinin 2 Network Map ...... 93

Figure 3.4.4: α-Actinin 2 States Page ...... 94

Figure 3.4.5: α-Actinin 2 Transitions Page ...... 96

Figure 3.5.1: α-Actinin 3 Overview ...... 98

Figure 3.5.2: α-Actinin 3 Summary Page ...... 100

Figure 3.5.3: α-Actinin 3 Network Map ...... 104

viii

Figure 3.5.4: α-Actinin 3 States Page ...... 105

Figure 3.5.5: α-Actinin 3 Transitions Page ...... 106

Figure 3.6.1: α-Actinin 4 Overview ...... 107

Figure 3.6.2: α-Actinin 4 Summary Page ...... 109

Figure 3.6.3: α-Actinin 4 Network Map ...... 118

Figure 3.6.4: α-Actinin 4 States Page ...... 119

Figure 3.6.5: α-Actinin 4 Transitions Page ...... 121

Figure 4.2.1.1: A Central Z-Section from a Raw Tomogram of Villin Cross-Linking F-actin ...... 139

Figure 4.2.1.2: Projection Images ...... 140

Figure 4.2.1.3: 3-D Classification Mask ...... 141

Figure 4.2.1.4: Class Averages Based on the Variance Within the Region Containing Villin ...... 141

Figure 4.2.1.5: Segmented Villin Density Maps ...... 142

Figure 4.2.1.6: Averaged Density Maps ...... 143

Figure 4.2.2.1: Model of Villin Cross-Linking F-actin ...... 145

Figure 4.2.2.2: Close-up of Three Villin-Actin Interactions ...... 146

Figure 4.3.2.1: Villin and Gelsolin Binding Sites do not Overlap ...... 148

ix ABSTRACT

We use a lipid monolayer system to prepare paracrystalline rafts of F-actin with two different cross-linking proteins, α-actinin and villin. α-Actinin cross-links F-actin into relatively loose networks throughout cells, while villin forms dense, tight bundles of filaments in specialized structures known as microvilli. Our monolayer system allows us to examine each of these cross-linkers as they interact with F-actin in a simplified two- component system. In order to analyze the data from these arrays, we have hybridized methods from 2-D crystal analysis with a single-particle approach to refine strategies for correspondence analysis and classification of images. For the α-actinin:F-actin rafts, we use this strategy to characterize the highly variable cross-links as 2-D projection images. The villin:F- actin rafts require further refinement of the hybrid strategy by application to 3-D volumes from electron tomography. Both protein arrays yield unique insights to the architectural arrangement of cross-linking proteins between filaments. In the α-actinin:F-actin rafts we use correspondence analysis to demonstrate that otherwise polar arrays of F-actin can have insertions of filaments of reversed polarity within them, and that these polarity differences do not influence the cross-links. We developed a method for left-right independent classification of the α-actinin cross-links to recover the high variability in the cross-link angular distribution by increasing the signal-to-noise ratio of the class averages. These averages are combined to recreate the original cross-link as it appears in a process we call “mapping-back.” Measurements reveal that the length of cross-links can vary. From the classification of cross-links we demonstrate and model the novel occurrence of α-actinin bound to successive crossover repeats on the same actin filament which we have termed “monofilament” binding. We further illustrate that the length variation of these monofilament-bound α-actinins are quantized to 55 Å, the distance between two adjacent actin monomers in the filament.

x The villin:F-actin rafts displayed homogeneous cross-linking. Docking of the homologous gelsolin atomic coordinates into the three-dimensional volumes reveals that villin does not interact with F-actin in the same manner as gelsolin. This data supports multiplicity of binding modes even in highly homologous proteins, and suggests a new mode of F-actin-binding for the villin protein.

CHAPTER 1 INTRODUCTION TO ACTIN CROSS-LINKING PROTEINS AND ELECTRON MICROSCOPY

1.1 The Cytoskeleton is a Cellular Scaffold

Beneath the cell membrane lies a complex yet organized network of filaments that function as dynamic but stable scaffolds to give the cells shape, and to serve as structural platforms onto which many other proteins bind. Three types of protein filaments are used to define a cell’s shape 1) microtubules, 2) intermediate filaments, 3) microfilaments. Microfilaments are composed of F-actin and its full complement of binding partners. They function to mediate motility (Lauffenburger and Horwitz, 1996), cell growth (Folkman and Moscona, 1978; Walker et al., 2005), apoptosis (Gourlay and Ayscough, 2005), and stem cell fate switching (McBeath et al., 2004). This body of work focuses on the structures of two different actin filament cross-linking proteins, α-actinin and villin. These are two very different members of a large family of proteins that cross- link filamentous actin (F-actin) to define cellular architecture.

1.1.1 Actin filaments in the cytoskeleton In the cytoplasm F-actin is a major component of bi-directional stress fibers that with the help of myosin II motor proteins can produce oppositely oriented contraction forces (Peterson et al., 2004) to maintain isometric tension (Schoenwaelder and Burridge, 1999). These fibers run throughout the cell, their exact location and concentration depends largely on interactions with cell-cell and cell-matrix adhesion complexes such as adherens junctions and focal adhesions, respectively. The extracellular environment, either in two-dimensional (2-D) monolayer culture or three-dimensional (3-D) matri-gel, determines the extent and location of such cellular fibers (Cukierman et al., 2001). The ends of these fibers terminate into polar arrangements of filaments at adhesion structures which are protein dense and relatively antibody inaccessible (Pavalko et al., 1995). Here, F-actin is tethered to adhesion plaques through interaction with various adaptor molecules. They in turn are directly linked through the plasma membrane via interactions with transmembrane adhesion molecules such as β-integrins in focal

adhesions (cell-matrix adhesion) and cadherins in adherens junctions (cell-cell adhesion). The precise timing and regulation of recruitment of each adaptor protein is the subject of on-going studies (Zamir and Geiger, 2001a; Zamir and Geiger, 2001b; Zimerman et al., 2004). Actin filaments are also integral to more specialized microdomain structures such as in the oppositely oriented Z-disk of the muscle sarcomere and the cytoplasmic dense bodies of smooth muscle. They also compose cellular extensions such as filipodia and villi, where different structure-specific accessory binding proteins determine the organization of the parallel filaments.

Figure 1.1.1.1. Z disc Architecture. (Z), Z disc; (M), M line; (A), A band; (I), I band; (SR), sarcoplasmic reticulum. Figure from (Epstein and Davis, 2003) and based on the 3-D reconstruction by (Luther and Squire, 2002). Inset shows EM of a section of sarcomere. Note the specific localization of α-actinin to the Z disc.

1.1.2 Cell structure and tension Although the cytoskeleton is somewhat elastic, it posses tensile properties that maintain cell shape upon deformation. Actin filaments are involved in two types of force. Protrusion forces (pushing) are produced by actin filament polymerization at the cell membrane, such as at the leading edge of crawling cells. Actin is involved in

transmitting tensile forces generated by acto-myosin and traction force produced by anchoring of the filaments to cell adhesions linked to another cell or the extracellular matrix. In the early 1990s the term tensegrity, a contraction of tensional integrity, was used by Donald Ingber to explain the push-pull forces that exist within a developing, living cell. The original concept is said to be derived from Buckminster Fuller’s geodesic sphere, modeled in Figure 1.1.2.1. Tensegrity is a state of structural pre-stress within the filament system. Perturbations of the cytoskeleton cause modifications in the structure and associated components, and this usually occurs without loss of the overall structure. Hence, cells have to be able to sense and respond to the environment without falling to pieces.

A B

Figure 1.1.2.1. A. The cytoskeletal network of actin filaments forms a geodome. Figure from (Ingber, 1998). B. Model of Fuller’s geodesic sphere.

1.1.3 Cell structure and signaling Recently, the focal adhesion structure has been redefined as a “paradigm for a signaling nexus” (Romer et al., 2006). The role of the Z disc in cell signaling is also emerging (Hoshijima, 2006). Both structures contain sites where tensile forces are generated. For the Z-disk, these forces are produced by oppositely oriented actin filaments being pulled by myosin. In the focal adhesion, stress fibers, which also have a sarcomeric arrangement, generate tension that is transmitted across the cell membrane through integrins (Riveline et al., 2001). The term mechanosensation is used to define cell signaling events in response to a mechanical stress. Classically, these events were shown to be a result of tension-

sensing ion channels in the cell membrane that open in response to stretch and allow the influx of potassium or calcium ions which act as second messenger signaling components (Lammerding et al., 2004; Kalapesi et al., 2005; Ingber, 2006; Wang and Thampatty, 2006). Other types of stretch sensors are being identified. One specific example is the cardiac stretch detector complex of α-actinin/MLP(muscle LIM protein)/ Tcap/titin in muscle Z disks. Mice with defective MLP have irregular Z disk structure and develop cardiac hypertrophy (HCM) (Geier et al., 2003; Gehmlich et al., 2004). The normal response to volume or pressure overload is stretching of the structural elements in the Z disk which triggers MAP kinase, JAK-STAT, and PI3K-AKT signaling cascades to increase gene expression and ultimately results in hypertrophic growth. The MLP deficient mice fail to localize the Tcap protein with titin and demonstrate loss of response to stretch, namely upregulation of cell survival genes and hypertrophic growth (Knoll et al., 2002). However, recent studies suggest certain cytoskeletal proteins are translocated to the nucleus in response to stretch events. MLP has been shown to dissociate from the Z disk structure in response to stretch and has been found in the nucleus as well. Another specific example is the translocation of the protein zyxin in response to stretch (Cattaruzza et al., 2004; Yoshigi et al., 2005; Lele et al., 2006). Cyclic stretch and/or shear stress causes the movement of zyxin from the focal adhesions to the actin stress fibers, which reorient and thicken in response to the stretch. Zyxin has also been demonstrated to accumulate in the nucleus of vascular smooth muscle cells in response to the same stresses where it is implicated in the regulation of gene expression. Other adhesion related proteins such as vinculin do not similarly mobilize in response to stretch.

Figure 1.1.3.1 The Focal Adhesion is a Site for Force-Regulated Cell Signaling. The focal adhesion acts as a mechanosensor. Force can be produced by pulling from the extracellular matrix, initiating intracellular signaling cascades through the transmembrane integrin receptors, or force can be generated internally by signaling from the focal adhesions to regulate myosin II-generated tension on the filaments. More than 50 proteins are known to be involved in the adhesion plaque (squares within oval). Figure from (Geiger and Bershadsky, 2002).

Figure 1.1.3.2 Schematic of Protein Interactions Emanating from the Z-disk of the Muscle Sarcomere. Figure from (Hoshijima, 2006).

1.2 Actin, Function and Structure

1.2.1 Actin is the Most Highly Conserved Protein In higher eukaryotes there are multiple isoforms of actin. They are specific to cell type, cellular location, and developmental stage. α-Actin isoforms are specific to muscle cells, while β- and -isoforms are found in non-muscle cell types. Yeast and bacteria only have a single isoform of actin. The yeast isoform, however, has 88% amino acid identity with skeletal muscle actin. Actin is also the most highly abundant protein in a cell. Non-muscle cells contain up to 5% actin in their total protein composition, while muscle cells contain 10% actin. Specialized actin structures such as filipodia and villi may have even higher local actin concentrations.

1.2.2 Actin Function As described in the previous section, actin forms filaments that compose one component of the cytoskeleton. The filaments rarely exist by themselves and are usually stabilized by interactions with multiple actin-binding proteins which cap the ends of filaments, cross-link filaments to form bundles, or act as tension generating motors. Actin is a ~375 amino acid globular protein that is referred to as G-actin in its soluble, monomeric form. The protein has four subdomains and these are separated by the ATP-magnesium ion binding cleft. Actin monomers assemble directionally to form a two- start long-pitch helix. The hydrolysis of ATP is not necessary for assembly, but does influence the polymerization kinetics. The filamentous polymer (F-actin) undergoes a phenomenon known as treadmilling where new protomers are added preferentially to one end of the filament (the +, or barbed end) while dissociating from the opposite end (the –, or pointed end). This produces a kind of molecular conveyor belt for associated actin-binding proteins. Capping proteins bound to the barbed end of the filament stabilize the addition of protomers, while capping proteins bound to the pointed end stabilize monomer dissociation.

1.2.3 Actin Structure The crystal structures of G-actin have been determined from complexes with binding proteins such as DNase I (Kabsch et al., 1985; Kabsch et al., 1990), gelsolin segment 1 (Mannherz et al., 1992; McLaughlin et al., 1993), profilin (Schutt et al., 1993; Minehardt et al., 2006) and as G-actin with an attached fluorophore (Otterbein et al., 2001). Electron microscopy provides intermediate resolution F-actin electron density maps to which the existing atomic structures can be fitted. Therefore, F-actin structure is limited to pseudo-atomic models. The first model was introduced by Holmes et al. in 1990 derived from the G-actin-DNaseI structure (Holmes et al., 1990). This model was refined by non-linear least-squares fitting of x-ray fiber diffraction data (Lorenz et al., 1993). The Holmes et al. model was further modified by fitting of atomic myosin and actin models to high resolution cryo-EM maps (Holmes et al., 2003). Crystallization of a chemically cross-linked pair of actin monomers has produced the most recent model, though it is not significantly different from the Holmes model (Kudryashov et al., 2005). While all are very similar, the main difference among the models is whether the ATP binding cleft is “open” or “shut” (Aguda et al., 2005). Additional differences occur in subdomain 2 which has a β-sheet structure when bound to DNaseI (Kabsch et al., 1990) but is α-helical in other cases (Otterbein et al., 2002). It is also a well established fact that the overall twist of the filament is variable and is influenced by binding proteins, nucleotide binding, or cation presence (Egelman and Orlova, 1995; McGough et al., 1997; Orlova et al., 2001). A conformational change within the actin monomer can change the flexibility of the entire filament, thereby suggesting large-scale cooperativity within the filament (Orlova and Egelman, 1993). The actin-binding protein ADF/cofilin also perturbs the twist of the filament, possibly increasing filament flexibility or creating strain (McGough et al., 1997).

Figure 1.2.3.1 The domain organization of G-actin. The monomer is divided into four domains as labeled. Subdomain 2, in green, tends to be disordered in most crystal structures. The cleft between subdomains 2 & 4 can be either open or closed, depending on the state of the nucleotide bound. Residues lining the interior of the hydrophobic binding pocket are shown in orange. This pocket is a “hot-spot” for the actin-binding helix of several actin capping proteins as well as subdomain 2 of the next actin monomer down the filament. Subdomain 1 is the most negatively charged subdomain (Sutoh and Yin, 1989).

Figure 1.2.3.2 Domain interaction in F-actin. Shown here are two consecutive actin monomers as they would be arranged in F-actin. The capping site at the barbed end is free; however, the same site one monomer up may be partially occluded by the variable subdomain 2 DNase-I binding loop. There is a second protein binding region that comprises subdomain 1 & 2 of one monomer and subdomain 1 of the next monomer up the filament. This is a binding site for F-actin side-binding proteins such as gelsolin G2 and α-actinin. An alternative, less well-defined protein binding region on subdomain 1 has been identified in this and other work (Renoult et al., 2001).

The crystal structures of G-actin in complex with several actin-binding proteins permit detailed analysis of modes of interaction. The structure of G-actin complexed with

DNAseI revealed the 4 subdomains of actin and that DNAseI bound specifically to actin subdomain 2 (Kabsch et al., 1985; Kabsch et al., 1990). This actin domain has been dubbed the DNase-binding loop. The next crystal structure solved was the actin-gelsolin segment 1 complex (Mannherz et al., 1992; McLaughlin et al., 1993). The binding of gelsolin segment 1 is between actin subdomains 1 and 3. This work also established the structure of the gelsolin fold. Gelsolin segment 1 is believed to play a specific role in barbed-end filament capping because its binding at this location on actin prohibits addition of new actin monomers. Furthermore, the N-terminal 3 gelsolin domains (G1- G3) (Burtnick et al., 2004), the G1 domain plus the linker region (Irobi et al., 2003) and the C-terminal 3 domains (G4-G6) (Robinson et al., 1999; Choe et al., 2002) have been independently crystallized with G-actin. The N-terminal gelsolin complex revealed two types of interactions with the G-actin monomer: the same structure of G1 in the capping position, plus a novel side-binding interaction of G2-G3. The side-binding interaction made contacts with actin subdomains 1 and 2 of G-actin and potentially subdomains 1 and 3 of the next actin protomer in a filament. G4, by homology with G1, also bound G- actin between subdomains 1 and 3 in a capping manner. The crystal structure of profilin, a G-actin binding protein, also reveals binding between subdomains 1 and 3 (Schutt et al., 1993). A more recent crystal structure of the vitamin D binding protein in complex with G-actin again shares this same binding trend (Head et al., 2002; Otterbein et al., 2002). As diagramed in Figure 1.2.3.2, there seems to be at least two distinct binding modes on F-actin. The most frequent is the subdomain 1 and 3 binding “hot- spot” or hydrophobic binding pocket and cleft.

1.3 The Spectrin Repeat Family, Function and Structure

1.3.1 Spectrin repeat family The spectrin superfamily, so named for the first protein discovered within the group, is comprised of three separate families all related by common sets of functional and structural motifs that have been conserved throughout evolution. In the post-genomic age, the shift to proteomics calls for inferring protein function from predicted domain homology determined from the gene sequence. There are far fewer known protein structures than there are genes. The spectrin-like proteins are perfect examples of how

protein diversity is achieved with only a small set of repetitive motifs. The three motifs in common to all members are the actin-binding domain, comprised of two tandem calponin-homology domains, the spectrin-like three-helix repeats, and a pair of EF-hand domains. The spectrin subfamily consists of spectrin, dystrophin, utrophin, and α- actinin, which all share the spectrin-like three-helix repeats. ABP-120 and filamin, which contain similar actin-binding domains, are separated into another family because they contain immunoglobulin-like repeats rather than three-helix repeats. Fimbrin and adducin fall into a third family with two actin-binding domains being located on the same polypeptide chain. The actin-binding domain (ABD) of all of these proteins is a tandem pair of calponin homology (CH) domains (Castresana and Saraste, 1995). Unlike the association of the smooth muscle protein calponin, which has only one such domain, spectrin family proteins require both domains to provide interaction sites for binding F-actin with affinity in the 5-50 µM range (Gimona et al., 2002). The two CH domains, while having a conserved core domain, are not functionally equivalent. The core is a compact, globular tertiary fold of four α-helices of 11-18 residues each, connected by long loops (Broderick and Winder, 2002). Pair wise comparison of helical sequences reveals greater similarity between the N-terminal CH domains (CH1) across a family than between the N-terminal and C-terminal domains (CH2) within the same protein (Bramham et al., 2002). The functional difference is that CH1 domains can bind actin independently, albeit with 10-fold less affinity than the entire ABD, while CH2 domains cannot. The difference possibly lies in sequence divergence in the exposed binding regions of the α-helices. In the dystrophin ABD three α-helices are identified as actin-binding sequences, ABS1-3 (Levine et al., 1992). Sites 1 and 2 are in CH1 and site 3 is in CH2. A helical linker separates the two CH domains. The length of this linker differs across the spectrin families. In fimbrin’s tandem CH domains, the linker contains an additional 13 residues. The crystal structure reveals a more compact density indicating a closed conformation, possibly due to the extra length allowing the domains to fold back onto themselves (Goldsmith et al., 1997). Utrophin, having a shorter linker, has a more open dumbbell shape. Yet when dimerized, the

domain-swapping phenomenon results in a compact association between CH1 of one molecule and CH2 of the other (Figure 1.3.1.1) (Keep et al., 1999).

Figure 1.3.1.1 Comparison of CH Domains. The structures (from (Broderick and Winder, 2002)) are as follows: a) CH2 domain from utrophin, b) ribbon diagram of anti-parallel utrophin dimer as an example of domain-swapping, c) utrophin monomer in the open conformation, d) ribbon diagram of fimbrin in the closed conformation. These variations in structure hint at the possibility of multiple modes of F-actin binding.

The spectrin repeats are highly conserved motifs comprised of a left-handed triple- helical coiled-coil bundle, formed from two long overlapping helices separated by a shorter helix (Speicher and Marchesi, 1984; Parry et al., 1992). The length of the individual three-helix repeat varies with spectrin having 106 residues and α-actinin having 122 (Parry et al., 1992). The number of tandem repeats also varies from 4 for α- actinin, to 24 for dystrophin. Amino acid sequence analysis suggests that the longer repeat molecules arose from an α-actinin-like common ancestor (Viel, 1999). The homology of spectrin repeats is a product of intragenic duplication followed by divergence among the species in a two-phase evolutionary model (Thomas et al., 1997). The result being that repeats across species are, again, more similar than repeats within a species. There are, fascinatingly, ~500 homologous spectrin repeat motifs appearing just in the human genome (Altmann et al., 2002). The spectrin repeats

of both α-actinin and spectrin facilitate dimerization via conserved hydrophobic residues stabilized by ionic interactions (Djinovic-Carugo et al., 2002). Utrophin and dystrophin do not dimerize due to lack of specific charged residues along the rod (Broderick and Winder, 2002). The length of the repeat domain has implication for function. In α-actinin the length of the three-helix bundle domain controls the distance separating actin filaments. Theoretically, the shorter the region, the more rigid the rod. While for the longer spectrin molecules, length contributes to the flexibility of the linkage of the actin cytoskeleton to the cell membrane, which is important for erythrocytes undergoing mechanical shear or deformation. The spectrin repeat region contains, in addition to the three-helix motifs, pleckstrin-homology domains, SH3 domains, WW or ZZ domains which serve as platforms for the docking of other structural and regulatory proteins, as well as many signaling proteins (Broderick and Winder, 2002; Djinovic-Carugo et al., 2002). The C-terminus of many spectrin family proteins consists of a pair of EF-hand domains. Each domain consists of a pair of helix-loop-helix structures. Traditionally the loop regions bind calcium or occasionally magnesium via coordinating oxygen groups. While the structural integrity of these domains may be preserved, the Ca2+ binding function is not necessarily required. For example, several of the spectrin and α-actinin isoforms have lost the ability to bind calcium in the loop domains. This implies some other conserved function for these domains. For example, EF-hands 3-4 of α-actinin, which do not bind calcium at all, act as specific binding sites for other major structural proteins (Atkinson et al., 2001). In other proteins the vestigial domain acts as a structural support (Broderick and Winder, 2002). A general role for EF-hand containing proteins in the cell is that of a calcium-sensing molecule. Many of these domains undergo a conformational change upon calcium binding. This change is characterized by an opening between the two helices accompanied by a clockwise swing of one helix to expose hydrophobic surfaces for interaction with a target protein, often helical in nature (Yap et al., 1999).

Figure 1.3.1.2 The structure of pairs of EF-hand domains. A, Calmodulin in the open conformation as with calcium bound, PDB 1CLL. B, An intermediate structure with N- terminus locked in a closed position, PDB 1Y6W. C, Apocalmodulin in the closed conformation, PDB 1QX5 chain D.

1.3.2 α-Actinin Function α-Actinin is a 94-103 kD rod-shaped antiparallel dimer which crosslinks filamentous actin at both ends in either parallel or anti-parallel orientation. α-Actinin is the smallest member of the spectrin superfamily of evolutionarily related proteins composed of similar structural domains. To date there has not been an x-ray structure produced for the entire molecule due to the flexibility associated with its modular design. However, crystal or NMR structures have been obtained for individual domains for this and other proteins within the family. The molecule is well suited for the formation of 2-D arrays for a low-resolution 3-D map using electron cryo-microscopy (Taylor and Taylor, 1993; Taylor and Taylor, 1994; Taylor et al., 2000; Tang et al., 2001; Liu et al., 2004). The existing NMR and X-ray crystal structures and homologous domains can then be docked into the low-density map to produce a 3D atomic model. This methodology is referred to as “hybrid crystallography.” Further refinement of the system would hopefully produce a model for actin binding and regulation thereof. α-Actinin was first identified by its ability to gel F-actin and super precipitate actomyosin (Ebashi and Ebashi, 1965). These somewhat rigid rods are 3-4 nm wide

and 30-40 nm long (Podlubnaya et al., 1975). Each actin-binding domain binds to two adjacent monomers of F-actin along the long-pitch, two-start helix (McGough et al., 1994). Thus, α-actinin has no affinity for G-actin. It crosslinks F-actin in either parallel or antiparallel orientation, depending on its structural role. α-Actinins can be broadly separated into two groups—sarcomeric and cytoplasmic. Sarcomeric (skeletal muscle) α-actinin is localized to the Z-discs where it functions to anchor actin filaments in an anti-parallel fashion via the giant protein titin (Lazarides, 1976; Eilertsen et al., 1997; Young et al., 1998). Cytoplasmic (non-muscle) actinins are involved in the formation of antiparallel F-actin arrays in stress fibers and formation of parallel F-actin bundles at focal adhesions. The major structural and functional difference between these two groups is that the sarcomeric isoforms have completely lost the ability to bind calcium ions in their EF-hand domains via insertions and deletions in the calcium-binding loops and thus bind actin in a calcium insensitive manner. Non- muscle isoforms are variably calcium sensitive and bind calcium at the µM level. At internal concentrations greater than 10-7M, actin cross linking is completely abolished in the most sensitive isoforms (Blanchard et al., 1989). In human and mouse there have been four isoforms of α-actinin identified. α-Actinins 2 and 3 are sarcomeric, calcium-insensitive isoforms. α-Actinin 2 was originally identified as a skeletal muscle isoform, however it has since been discovered in cardiac muscle, and in the brain at glutamatergic synapses and post-synaptic densities, which makes the muscle grouping somewhat of a misnomer (Wyszynski et al., 1998; Tiso et al., 1999). Interestingly, both of these cell types respond to calcium impulses from contraction signals and action potentials. It seems obvious that to have a calcium sensitive actin cross-linking protein in this environment could have deleterious effects on the cell structure. α-Actinin 3 has been identified only in the Z-discs of a subset of type II fast skeletal muscle and serves the same function as α-actinin 2. Because expression is not limited to a single isoform per cell type, α-actinins 2 and 3 can be found together in the type II fast skeletal muscle and instead of forming homodimers, the two can form heterodimers (Chan et al., 1998). In humans, ~18% of the population has a null mutation for α-actinin 3 due to an early stop codon (North et al., 1999). These individuals show no discernable phenotype for the mutation, suggesting that α-actinin 2

can substitute for the redundant α-actinin 3. In the mouse genome, however, the homologous α-actinin 3 is necessary for muscle development and its absence results in myopathies. Hence, α-actinin 2 does not substitute for α-actinin 3, calling into question the validity of some mouse models, especially since the mouse musculature is ~80% fast skeletal type II, versus 40-50% in humans (Mills et al., 2001). The non-muscle, calcium sensitive isoforms are α-actinin 1 and 4. α-Actinin 1 functions as an antiparallel homodimer to anchor the actin cytoskeleton to the membrane as well as link signaling proteins to the cytoskeleton. α-Actinin 1 is localized to stress fibers, microfilament bundles, and at focal adhesions and adherens type junctions. Over expression of α-actinin 1 leads to a decrease in cell motility, perhaps due to an increase in plaque formation. Reduction of its expression is linked to increased motility and tumor formation. In chicken, a single α-actinin 1 gene gives rise to two isoforms by alternative splicing. The difference lies in the second half of the first EF-hand domain. Type 1a is found in chicken smooth muscle (gizzard) while 1b is cytoskeletal and calcium sensitive like human/mouse α-actinin 1 (Parr et al., 1992; Waites et al., 1992). α-Actinin 4, also an antiparallel homodimer, was discovered as a chicken lung actinin with reduced calcium sensitivity compared to α-actinin 1 (Imamura and Masaki, 1992; Imamura et al., 1994). While absent from the focal adhesions and adherens junctions, α-actinin 4 is largely cytoplasmic and can be translocated to the nucleus in the event of induced actin depolymerization. Its expression is associated with cell motility and metastasis (Honda et al., 1998; Honda et al., 2004; Honda et al., 2005). The principal mode of regulation of the non-muscle isoforms is by calcium binding. It is supposed that the binding of calcium at the EF-hands induces a conformational change which opens the hydrophobic regions between the helices, allowing them to bind to the adjacent actin-binding domain and thus prevent actin binding. There exists only one observation in support of this hypothesis (Tang et al., 2001). In the Tang et al. model, the calmodulin-like domain binds to the two CH domains causing them to separate which would decrease their actin binding affinity. Other forms of regulation come from signaling or regulatory proteins. The enzyme focal adhesion kinase (FAK) phosphorylates α-actinin on a tyrosine residue at the actin- binding domain, thus inhibiting its cross-linking abilities (Izaguirre et al., 2001). The

second messenger PIP2 has been demonstrated to have a regulatory effect, enhancing the gelation of actin filaments (Fukami et al., 1996). It increases the ability of PKN, which also phosphorylates α-actinin, to bind to the third spectrin-like repeat and the EF- hand domain in a calcium sensitive manner (Mukai et al., 1997). The spectrin-like repeats seem to act as a platform for the binding of several proteins involved in signaling events. The PDZ/LIM domain proteins serve as adapters. They specifically recognize the spectrin repeats via the PDZ domains while recruiting and binding kinases to the LIM domain.

1.3.3 α-Actinin Structure The structure of α-actinin has yet to be solved to atomic resolution, due in part to the flexibility associated with its modular design. Yet it is this same modularity that makes this molecule well suited for hybrid crystallography and homology modeling. Because the spectrin-families are evolutionarily related to one another, the few high-resolution NMR and x-ray domains can be coupled to the lower resolution EM density maps to create pseudo-atomic models of an entire protein. The structure of the four spectrin repeats of α-actinin has been solved (Ylanne et al., 2001a; Ylanne et al., 2001b), as has the non-calcium binding EF 3-4 region (Atkinson et al., 2001). The structures for utrophin and dystrophin exhibit an extended conformation. Other more recent crystal structures for α-actinin 3 ABD (Franzot et al., 2005) and α-actinin 1 (Figure 1.3.3.1) (Borrego-Diaz et al., 2006) are available, both in the closed conformation. EM studies suggest that the ABD-actin interaction may have two modes of binding, open and closed. The compact binding theory predicts the ABD to interact with actin with few changes from the crystal structure, while the open binding theory suggests that the two domains separate from one another. Support for compact binding comes from difference mapping of ABD decorated F-actin (McGough et al., 1994; Hanein et al., 1998; Sutherland-Smith et al., 2003 ). Real Space Iterative Helical Refinement techniques have shown an extended conformation of the ABD with F-actin and further suggests that homologous ABDs may have completely different interactions with the filament and that a single ABD has multiple modes for interaction with F-actin (Galkin et al., 2002).

Figure 1.3.3.1 Structure of the ABD from human α-actinin 1. The ribbon diagram shows CH1 in blue and CH2 in red. The surface representation shows the actin-binding sequences. ABS1 is in brown, ABS2 is purple, and ABS3 is cyan. Note that ABS1 is buried in the interface between the two domains, suggesting that the domains may undergo rearrangement on binding F-actin. Figure from (Borrego-Diaz et al., 2006).

Figure 1.3.3.2. Structure of the α-actinin rod.(a) ribbon diagram of the α-actinin rod along the 2-fold symmetry axis, (b) view perpendicular to 2-fold symmetry axis, (c) diagram of how the position of repeats relate to others in the rod, (d) surface view illustrating 90º twist. Figure from (Ylanne et al., 2001a)

1.4 The Gelsolin Family, Function and Structure

1.4.1 Actin filament severing, capping, nucleating, and bundling proteins The gelsolin superfamily of proteins are characterized by their ability to bind calcium ions and actin and function to regulate actin filament length by severing and capping the barbed ends. At the cellular level this family of proteins participates in actin remodeling, cell motility, phagocytosis, apoptosis, cell development, platelet activation, and disease. The first two severing proteins described are gelsolin (Yin and Stossel, 1979) and villin (Bretscher and Weber, 1980). Other family members include adseverin, CapG, advillin, supervillin, and flightless I /severin. They all share 3 or 6 copies of a single 120 amino acid, 14 kDa, structural domain, referred to as the gelsolin-like domain. Gelsolin and villin each have 6 domains which arose evolutionarily by first a gene triplication event, followed by a duplication event resulting in two structurally homologous but functionally non-equivalent halves. CapG is the smallest, containing only 3 domains, and has no C- terminal severing ability (Yu et al., 1990). Villin is unique in this family because of a small, 76 amino acid villin head piece domain at the C-terminus of the gelsolin-like domains. This extra domain allows villin to function as an F-actin cross-linking protein.

1.4.2 Villin found primarily in microvilli Villin is an actin-binding protein localized in intestinal and kidney brush borders (Bretscher and Weber, 1980). Villin contains at least three actin-binding sites, two of which are Ca2+ dependent and located in the core domain (gelsolin repeats). The third is situated in the head piece domain and is Ca2+ independent. The in vitro activities of villin upon actin vary with the Ca2+ concentration. At high Ca2+ concentration (>10-4 M), villin severs F-actin into short filaments. This has been demonstrated to occur in vivo, as well as in vitro (Ferrary et al., 1999). At lower concentration (10-7 to 10-6 M), villin caps the fast-growing end of the actin microfilament and thereby prevents elongation. At the same Ca2+ concentration, villin nucleates microfilament growth when added to actin monomers. At very low Ca2+ concentration (<10-7 M), villin has no effect on actin polymerization but bundles actin filaments. Villin is further regulated by PIP2 binding (Janmey et al., 1992) and tyrosine phosphorylation (Khurana et al., 1997; Zhai et al., 2001; Zhai et al., 2002; Kumar et al., 2004).

Figure 1.4.2.1 Micrograph of a section of intestinal epithelium microvilli. Arrows point to actin filament bundles which are cross-linked by villin and fimbrin.

1.4.3 Villin is Structurally Similar to Gelsolin Much of what is known about the structure of villin is inferred from the structure of gelsolin. The two proteins have ~42% amino acid identity across the 6 gelsolin-like domains. There is to date no crystal structure of the intact villin protein, although villin 14T (V1) has been solved (Markus et al., 1994; Markus et al., 1997) and numerous structures of the small, fast folding head piece domain exist (McKnight et al., 1996; McKnight et al., 1997; Vardar et al., 1999; Frank et al., 2004; Vermeulen et al., 2004; Meng et al., 2005; Grey et al., 2006; Tang et al., 2006). Most of what is known about the villin core (V1-V6) structure is inferred from gelsolin G1-G6. Although structurally similar, they may have distinct functional behaviors as evidenced by villin/gelsolin chimera proteins that do not display the same behaviors (Finidori et al., 1992). Also, the villin V1 domain has a weaker affinity for actin than does the gelsolin G1 domain, despite 58% sequence identity between the two (Kwiatkowski et al., 1985; Janmey and Matsudaira, 1988). Some of the earliest structural studies of villin were investigated using hydrodynamic and spectroscopic methods in solution (Hesterberg and Weber, 1983). Their data suggest an asymmetric molecule with axial ratio of 4.5:1 and a maximum length of 84 Å

in EGTA. In the presence of 100 µM Ca2+ the molecule became more asymmetric with an axial ratio increase to 8:1 and maximum length of 123 Å. The authors also observed that the proteolytic fragment of villin comprising the core domain (V1-V6) did not undergo dramatic change in response to Ca2+, as it maintained a constant 76 Å length. This lack of change in the core region points to a major role for the actin-binding headpiece in secondary structure changes. The authors propose a hypothetical “hinge mechanism” to explain the hydrodynamic change. Here the binding of Ca2+ might exert an induced local “tightening” of the secondary structure, possibly through an increase in α-helicity, which may in turn cause another region to pivot out into solution and alter the hydrodynamic properties. Applied specifically to villin, the small headpiece domain would be compact with the core domain and addition of Ca2+ would “relax” the global compact structure by allowing the headpiece to come away from the core region. This interpretation would suggest that villin would crosslink F-actin in a compact form and the binding of Ca2+ would create a more extended, open structure that would disrupt the headpiece binding to F-actin, while still permitting Ca2+ activated filament severing by the N-terminal half of the core domain. A similar mechanism of activation has been proposed for gelsolin. Inactive, calcium-free gelsolin is globular in nature and calcium binding releases a C-terminal “tail helix unlatching” mechanism which exposes the F- actin binding sites at the N-terminus (Lin et al., 2000; Choe et al., 2002; Kiselar et al., 2003). The crystal structure of inactive gelsolin has the G6 domain folded in towards the center of the molecule where it contacts G2’s actin-binding region (Burtnick et al., 1997; Urosev et al., 2006).

Figure 1.4.3.1 The villin domains. The villin core is comprised of six conserved gelsolin- like domains (represented here by the gelsolin crystal structures) with an additional C- terminal F-actin binding domain referred to as the head piece. Figure made with Chimera.

!!AA_MULTIPLE_ALIGNMENT 1.0

MSF: 882 Type: P June 16, 2006 17:07 Check: 3329 ..

Name: VILI_CHICK Len: 882 Check: 6265 Weight: 1.0

Name: GELS_HUMAN Len: 882 Check: 7064 Weight: 1.0

VILI_CHICK 1 ...... M 1

GELS_HUMAN 1 MAPHRPAPAL LCALSLALCA LSLPVRAATA SRGASQAGAP QGRVPEARPN 50

|...Signal sequence...... |

|.....Gelsolin-like Domain 1

VILI_CHICK 2 VELSKKVTGK LDKTTPGIQI WRIENMEMVP VPTKSYGNFY EGDCYVLLST 51

GELS_HUMAN 51 SMVVEHPEFL KAGKEPGLQI WRVEKFDLVP VPTNL GDFF TGDAYVILKT 100

[Actin-severing Region | Gelsolin-like Domain 1...

Phosphotyrosine

...... |

VILI_CHICK 52 RKTGSG.FSY NIHYWLGKNS SQDEQGAAAI YTTQMDEYLG SVAVQHREVQ 100

GELS_HUMAN 101 VQLRNGNLQY DLHYWLGNEC SQDESGAAAI FTVQLDDYLN GRAVQHREVQ 150

α Helix

...... | ______

Actin-actin inter-filament contact

Figure 1.4.3.2 Annotated sequence alignment of villin and gelsolin. Alignment generated by ClustalW (Thompson et al., 1994).

{Critical for villin

actin-binding} |..

VILI_CHICK 101 GHESETFRAY FKQGLIYKQG GVASGMKHVE TNTYNVQRLL HVKGKKNVVA 150

GELS_HUMAN 151 GFESATFLGY FKSGLKYKKG GVASGFKHVV PNEVVVQRLF QVKGRRVVRA 200

______] |..

Polyphosphoinositide binding

..Gelsolin-like Domain 2...... |

VILI_CHICK 151 AEVEMSWKSF NLGDVFLLDL GQLIIQWNGP ESNRAERLRA MTLAKDIRDR 200

GELS_HUMAN 201 TEVPVSWESF NNGD FILDL GNNIHQW GS NSNRYERLKA TQVSKGIRDN 250

..Gelsolin-like Domain 2...... |

VILI_CHICK 201 ERAGRAKVGV VEGENEAASP ELMQALTHVL GEKKNIKAAT PDEQVHQALN 250

GELS_HUMAN 251 ERSGRARVHV SE...... EG TEPEAMLQVL GPKPALPAGT EDTAKEDAAN 294

|..Gelsolin-like Domain 3......

VILI_CHICK 251 .SALKLYHVS DASGNLVIQE VAIR.PLTQD MLQHEDCYIL DQAG.LKIFV 297

GELS_HUMAN 295 RKLAKLYKVS NGAGTMSVSL VADENPFAQG ALKSEDCFIL DHGKDGKIFV 344

|..Gelsolin-like Domain 3......

...... |

VILI_CHICK 298 WKGKNANKEE KQQAMSRALG FIKAKNYLAS TSVETENDGS ESAVFRQLFQ 347

GELS_HUMAN 345 WKGKQANTEE RKAALKTASD FITKMDYPKQ TQVSVLPEGG ETPLFKQFFK 394

...... | Flexible Linker Region

Figure 1.4.3.2 Continued.

VILI_CHICK 348 KWTVPNQTSG LGKTHTVGKV AKVEQVKFDA TTMHVKPEVA AQQKMVDDGS 397

GELS_HUMAN 395 NWRDPDQTDG LGLS LSSHI ANVERVPFDA ATLHTSTAMA AQHGMDDDGT 444

Phosphotyrosine [______

|...Gelsolin-like Domain 4......

VILI_CHICK 398 GEAEVWRVEN QELVPVEKRW LGHFYGGDCY LVLYTYYVGP KVNRIIYIWQ 447

GELS_HUMAN 445 GQKQIWRIEG SNKVPVDPAT GQFYGGDSY IILYNYRHGG RQGQIIYNWQ 494

Phosphotyrosine

|...Gelsolin-like Domain 4......

Calcium-Sensitive actin-binding region______

...... |

VILI_CHICK 448 GRHASTDELA ASAYQAVFLD QKYNNEPVQV RVTMGKEPAH LMAIFKG.KM 496

GELS_HUMAN 495 GAQSTQDEVA ASAILTAQLD EELGGTPVQS RVVQGKEPAH LMSLFGGKPM 544

...... |______

|...Gelsolin-like...

VILI_CHICK 497 VVYENGSSRA GGTEPASSTR LFHVHGTNEY NTKAFEVPVR AASLNSNDVF 546

GELS_HUMAN 545 IIYKGGTSRE GGQTAPASTR LFQVRANSAG ATRAVEVLPK AGALNSNDAF 594

______|...Gelsolin-like ..

Domain 5...... |

VILI_CHICK 547 VLKTPSSCYL WYGKGCSGDE REMGKMVADI ISKTEKPVVA EGQEPPEFWV 596

GELS_HUMAN 595 VLKTPSAA L WVGTGASEAE KTG.AQELLR VLRAQPVQVA EGSEPDGFWE 643

.Domain 5...... |

__Phosphotyrosine______

Figure 1.4.3.2 Continued.

|..Gelsolin-like

VILI_CHICK 597 ALGGKTSYAN SKRLQEE.NP SVPPRLFECS NKTGRFLATE IV.DFTQDDL 644

GELS_HUMAN 644 ALGGKAA RT SPRLKDKKMD AHPPRLFACS NKIGRFVIEE VPGELMQEDL 693

__Phosphotyrosine______|..Gelsolin-like

Domain 6...... |

VILI_CHICK 645 DENDVYLLDT WDQIFFWIGK GANESEKEAA AETAQEYLRS HPGSRDLDTP 694

GELS_HUMAN 694 ATDDVMLLDT WDQVFVWVGK DSQEEEKTEA LTSAKRYIET DPANRDRRTP 743

Domain 6...... | ______

VILI_CHICK 695 IIVVKQGFEP PTFTGWFMAW DPLCWSDRKS YDELKAELGD NASIGQLVSG 744

GELS_HUMAN 744 ITVVKQGFEP PSFVGWFLGW DDDYWS.VDP LDRAMAELAA ...... 782

______]

|....Villin Head Piece......

VILI_CHICK 745 LTSKNEVFTA TTTLVPTKLE TFPLDVLVNT AAEDLPRGVD PSRKENHLSD 794

GELS_HUMAN ......

{Sequence required for villin cross-linking}

...... |

VILI_CHICK 795 EDFKAVFGMT RSAFANLPLW KQQNLKKEKG LF 826

GELS_HUMAN ......

Figure 1.4.3.2 Continued.

1.4.4 Gelsolin is Regulated by Calcium In order to activate gelsolin and sever F-actin there are a series of calcium ion binding events that must occur to open 3 different latch mechanisms. The first is the aforementioned tail latch which releases the G2 F-actin side-binding domain to target gelsolin to the filament. The G1-G3 latch and the G4-G6 latch release the G1 and G4 actin severing-capping binding interfaces (Robinson et al., 1999; Choe et al., 2002). Each gelsolin domain has a type-2 Ca2+-binding site of variable affinity. Binding of Ca2+ to these sites regulates the unlatching of domains. There are also two type-1 sites in G1 and G4 that mediate the binding of these domains to actin and are sandwiched in the binding interface. The gelsolin structural data has been used to create models of how gelsolin binds to, severs, and caps F-actin. The G2 domain first binds to F-actin, bringing G1 into proximity for severing. The long, 53 residue linker between G3 and G4 is believed to wrap around the actin filament, placing G4 opposite G1 on the filament. It is the binding of G1 and G4 that initiates the cleavage of the filament by introducing torsional strain. After cleavage G1 and G4 remain bound to the barbed end as a cap. Binding of PIP2 releases the capping mechanism. This process has been modeled as shown in Figure 1.4.4.1. Villin can bind to F-actin in the absence of Ca2+ and requires nonphysiological (200 µM) levels of Ca2+ to sever F-actin (Janmey and Matsudaira, 1988). There seems to be less of a dependence Ca2+ for severing than a requirement for tyrosine phosphorylation events (Zhai et al., 2002; Kumar et al., 2004; Tomar et al., 2004). The only structural data for villin V1 (14T) predicts two possible, relatively low affinity calcium coordination areas (Markus et al., 1994).

Figure 1.4.4.1 Illustration of two possible models for gelsolin severing and capping F- actin. Figure is from (McGough et al., 2003). Step I demonstrates the release of the tail- latch mechanism. Step II illustrates the targeting of G2 to F-actin. Step III has G1 binding to actin. Steps IV a and b show alternate binding sites for G4. It is not known which actin monomer G4 binds to relative to G1. Note the linker stretches across the filament. Step V a and b are the severing and capping event. Steps VI and VII reflect the uncapping mechanism by PIP2.

1.5 Electron microscopy, electron tomography, and image analysis

1.5.1 Electron microscopy and image formation

Figure 1.5.1.1 Electron path through the column of the microscope.

The electron microscope uses a high energy electron source, ideally a 100-300 KeV field emission electron source for highest coherence, to produce short wave-length electrons that will either pass through or scatter on interaction with a very thin specimen. The column of the microscope is kept under vacuum to minimize any

additional electron scattering. The electrons emitted from the tungsten cathode must be kept coherent to ensure brightness of the beam. This is accomplished by the condenser lens and aperture. The lenses of the microscope are not made of ground glass as in light microscopy, but are electromagnetic fields generated by current passing through coils of copper wire that form a ring around the column. The specimen is inserted into the vacuum of the column just within the objective lens. Here the electrons scatter as they pass through the specimen. Electrons with no interactions pass straight through unmodified, producing a dense central spot in the diffraction pattern observed in the back focal plane, while the elastically scattered electrons in the exit wave, which are essentially shifted out of phase by 90°, form the wide angle diffraction data. As in light microscopy, a 90° phase shift applied to the scattered electrons can produce phase contrast. Unlike light microscopy, the phase shift is applied not by a “quarter wave plate” placed in the back focal plane, but instead by the contrast transfer function. Inelastically scattered electrons, which undergo a loss of energy on interaction with the specimen and hence do not come to focus in the image, contribute to amplitude contrast as do the electrons scattered at high angles that are removed by the objective aperture in the back focal plane. The unscattered and elastically scattered electrons converge onto a fluorescent viewing screen as an image, or can be recorded for analysis on micrographic film or CCD camera. Sample preparation influences the way the electrons interact with the specimen. Samples in vitreous ice produce very low contrast images, but preserve the integrity of the sample. Heavy metal negative stains such as uranyl acetate produce higher contrast images, but at the expense of smoothing and flattening the sample due to drying, pH, and ionic strength effects from the stain solution.

1.5.2 The Fourier Transform The process of image formation in the electron microscope is a function of sine and cosine waves with differences in amplitude, phase, and wavelength. The exit wave consists of both the phase-shifted scattered and unscattered electrons. The scattered electrons, consisting of elastically (characterized by no energy loss to the specimen and a wide angular distribution that contains high resolution information) and inelastically

(characterized by loss of energy to the specimen and narrow angular distribution which produces low resolution background noise) form a diffraction pattern in the objective lens back focal plane as mentioned in the previous section that can be described mathematically as a Fourier analysis of the exit wave. The propagation of this wave by the objective lens, known as a Fourier synthesis, produces an image, but the image is not perfect. The imaging process modifies the diffracted wave by the effects of defocus, energy spread, spherical aberration, etc. as the wave propagates through the imperfect lenses to the image plane in what can be described as a Fourier synthesis. These effects are collectively described in the following section as the contrast transfer function.

1.5.3 The Contrast Transfer Function (CTF) The CTF is a function that describes the combined effects of the imaging system on the exit wave such as changes in defocus, imperfect lenses, illumination divergence, and energy spread. The phase contrast transfer function in effect acts as “quarter wave plate” but its effect is more complicated than its light microscope analog. In the theory of electron image formation, the CTF multiplies the diffracted wave in the back focal plane. By the convolution theorem of the Fourier transform, the CTF produces a point spread function in real space that is convoluted across the image, producing a blurring effect that limits the overall resolution. The CTF can be calculated from knowledge of the image defocus and astigmatism, while other microscope specific factors such as spherical aberration are determined by the microscope manufacturer. A general equation follows (Frank, 1996): (α)=2π/λ[(-Cs α4) / (4 + (∆f α2)/2] where is the CTF, α is the scattering angle, λ is the wavelength, ∆ is the change in defocus, and Cs is the spherical aberration of the objective lens, which is 2.0 mm at 300 keV for the microscope used in this work. Correcting for CTF in images is not straightforward, since the function is in the form of a sine wave that oscillates between +1 and -1, crossing at nodes of zero value. Otherwise it would be a simple task to simply divide the image by the sine function. The correction factor is known as a Wiener filter which incorporates the signal-to-noise ratio

in the numerator thereby preventing over amplification near the zeroes of the CTF. The factors we measure and use in the CTF calculation are the spherical aberration of the objective lens, the wavelength in nm, the pixel size in the image, the box size of the image, the maximum and minimum defocus in nm, and the angle of astigmatism.

1.5.4 Tomography and the Projection Theorem The image created by the exit wave is a 2-D projection of the specimen. In order to obtain a 3-D reconstruction of the specimen we need either multiple views of an object, such as virus particles in random orientations on a grid, or we must tilt the same sample in space to achieve multiple views. Tomography is the semi automated process of collecting a series of tilted specimen projection images, usually spaced between ± 60°, and then aligning and merging them. From this data, 3-D volumes of density are computed by a process known as a weighted backprojection. We use a 2K x 2K CCD detector to record the data. Our lab uses a process of marker-free alignment and area matching which uses a reprojection of a backprojection computed in the previous cycle as a reference. This reference has comparable signal-to-noise ratio (SNR) or total electron counts as the sum of all the individual tilt images that comprise it. This is a superior method for image alignment of tilt series of thin specimens (Winkler and Taylor, 2006). All backprojections are weighted to decrease the low spatial frequency data that is present throughout the tilt series. The process of computing a 3-D image is possible due to the projection theorem, which defines the Fourier transform of the 2-D image projection from a 3-D object as a central section or plane of the transform of that 3-D object. Therefore, multiple central sections from multiple views of the 3-D object can be aligned and merged to produce the 3-D volume of the object. The quality of this reconstruction is dependent on the number of views of the object available and the separation between these views. The number of images required N, is a function of the diameter, D, of the object and the desired resolution, 1/d, such that N = π D/d, for an idealized cylindrical object (Crowther et al., 1970). In the case of our monolayer samples the object can be described as a slab that varies in thickness as it is tilted. Thus, we opt to tilt to angles defined by the cosine rule, or the Saxton scheme (Saxton and Baumeister, 1982), to correlate the tilt

increment with the tilt angle and specimen thickness, i.e. higher tilts are sampled at finer increments. Because the specimen holder is limited to ± 70° tilt angles, there will always be a missing wedge of data in the Fourier transformation due to a lack of central sections in this region. The effects are seen in the reconstruction as an elongated volume. Merging data sets with multiple tilt axes with respect to the specimen could potentially lessen this effect.

1.5.5 Averaging and Classification All projections have stochastic noise that reduces the signal-to-noise ratio (SNR) and hence the image resolution. Stochastic noise is generally due to low electron counts. Structural noise consists of conformational variability of the object as well as stain and other sample preparation artifacts. Averaging of like views improves the SNR by reducing the stochastic noise. Stochastic noise is random and thus decreases with averaging, while signal is reproducible and increases with averaging relative to the stochastic noise. This is the premise behind the single-particle approach of averaging defined views of molecules (Glaeser, 1971; Glaeser, 1999). In tomograms of macromolecular assemblies on the lipid monolayer, the paracrystalline arrays present a unique opportunity to merge techniques. The single- particle approach can be applied by selecting, either manually or by cross-correlation, multiple instances of a reoccurring motif across the array. Since the SNR improves as 1/N½, increasing the number of views, or repeats, of a motif stands to improve the overall quality. Difficulty can arise when not all views of a motif are identical, as is the case for proteins with heterogeneous orientations. Multivariate Statistical Analysis (MSA), or more specifically correspondence analysis, is used to identify patterns in data and when combined with hierarchical ascendant classification can cluster or bin related images. The classification process is highly dependent on the judicious use of pixel masks to define the region of interest. The best improvement in SNR is seen when a large number of repeats of limited variation are binned into relatively few but homogeneous classes and then averaged. Of course, with highly heterogeneous protein arrangements many more classes are required to capture the variation present in the sample which

results in fewer repeats being averaged together and a lower SNR. Thus, there is a balance that must be achieved between the improvements in image quality due to SNR improvement versus the capture of all variations among repeats. Suffice it to say, averaging inhomogeneous repeats blurs features rather than enhancing them. The complexity of correspondence analysis and classification increases for 3-D repeats derived from tomograms. The repeats must be aligned in Euler space, which requires 6 degrees of freedom (three angular, three rotational) in analysis, versus the usual three (two translational, one angular) required for projections. Also, masks for classification must be created in 3-D sections, and for each section some degree of variability may be lost.

33 CHAPTER 2 α-ACTININ IS A VARIABLE-LENGTH ACTIN CROSS-LINKER

We have applied correspondence analysis to electron micrographs of 2-D rafts of F- actin cross-linked with α-actinin on a lipid monolayer to investigate α-actinin: F-actin binding and cross-linking. More than 8,000 actin crossover repeats, each with 1-5 α- actinins bound, were selected, aligned, and classified to produce class averages of α- actinin cross-links with ~9-fold improvement in the signal-to-noise ratio. Measurements and comparative molecular models show variation in the length and the angle of the α- actinin cross-links. Rafts of F-actin and α-actinin formed predominantly polar 2-D arrays of actin filaments, with occasional insertion of filaments of opposite polarity. Unique to this study are the numbers of α-actinin molecules bound to successive crossovers on the same actin filament. These results suggest that α-actinin is not simply a rigid spacer between actin filaments, but rather a flexible cross-linking, scaffolding, and anchoring protein. We suggest these properties of α-actinin may contribute to tension sensing in actin bundles.

2.1 Background

α-Actinin is a modular protein belonging to the spectrin superfamliy that cross-links and bundles actin filaments in both muscle and non-muscle cells (Blanchard et al., 1989). There is no high resolution structure of the entire molecule, but atomic structures exist for most of its individual domains. α-Actinin has an N-terminal actin-binding domain (ABD) consisting of a tandem pair of non-equivalent calponin homology domains (CH1 and CH2) (Mannherz et al., 1992). Its structure has recently been solved by x-ray crystallography (Franzot et al., 2005; Borrego-Diaz et al., 2006). The placement of the ABD fragment on the actin filament has also been determined (McGough et al., 1994). In both of these structures, actin-bound and free, the ABD has a compact, closed arrangement of CH1 and CH2. In 2-D crystals, on the other hand, the ABD of intact α- actinin can adopt either an open or a closed conformation (Liu et al., 2004). The ABD is linked to the rest of the molecule by a 25-30 residue protease-sensitive flexible linker (Imamura et al., 1988) whose structure is unknown. The linker is followed by a rod-like

34 domain of four triple-helical, coiled-coil repeats (R1-R4). The R1-R4 domain lends the molecule an overall 90° left-handed twist (Ylanne et al., 2001; Liu et al., 2004) that may contribute to its role as a protein docking platform (Djinovic-Carugo et al., 2002). The C- terminus contains a calmodulin-like (CaM) domain containing a pair of structurally, but not necessarily functionally, conserved EF-hand motifs that bind Ca2+ in some isoforms (human, mouse ACTN1 & 4) while having evolutionarily lost this Ca2+-binding ability in other isoforms (human, mouse ACTN2 & 3) (Burridge and Feramisco, 1981; Blanchard et al., 1989; Witke et al., 1993). α-Actinin forms antiparallel dimers through strong ~10 pM affinity associations between R1-R4 domains (Kahana and Gratzer, 1991; Flood et al., 1997). This arrangement places the CaM domain in close proximity to the ABD and is hypothesized to influence the ABD conformation (Noegel et al., 1987; Tang et al., 2001). These existing domain structures have been combined to generate a model of the dimer to fit 3-D images obtained by cryo-EM (Liu et al., 2004). Previous studies on the bundling of actin filaments have shown that α-actinin can cross-link in any orientation. Bundles formed in solution using chicken smooth muscle α- actinin favored an antiparallel orientation (9 of 11 filaments assayed) (Meyer and Aebi, 1990), while in other studies using the same isoform, 2-D bundles (rafts) formed on a lipid monolayer overwhelmingly preferred parallel cross-links (Taylor and Taylor, 1994). Meyer and Aebi (1990) suggested that the bundle characteristics were determined solely by the α-actinin molecular length and Taylor et al. (2000) hypothesized that extrinsic factors were required to influence specificity of cross-linking orientation. α-Actinin is localized to a variety of cellular structures requiring organized actin filament polarity. In Z-disks of striated muscle (Masaki et al., 1967), cytoplasmic dense bodies of smooth muscle (Bond and Somlyo, 1982), and stress fibers of migrating cells (Lazarides and Burridge, 1975), α-actinin cross-links oppositely oriented actin filaments to form bipolar assemblies. In focal adhesion plaques at cell membranes α-actinin is thought to cross-link similarly oriented actin filaments into polar bundles and also link them specifically to integrins (Burridge et al., 1990; Otey et al., 1990; Rajfur et al., 2002). α-Actinin has been localized to these protein dense regions by fluorescent antibodies but its actin cross-linking function there is inferred.

35 α-Actinin also has numerous binding partners (Otey and Carpen, 2004; Hampton and Taylor, 2006a,b,c). Through its interaction with the β-integrin cytoplasmic domains (Otey et al., 1990; Pavalko et al., 1991; Otey et al., 1993) α-actinin is thought to play a role in the formation and stabilization of focal adhesions in migrating cells (Rajfur et al., 2002). Interactions between α-actinin and other focal adhesion and stress fiber proteins include vinculin, zyxin, CRP, paxillin, MEKKI, PIP2, and FAK (reviewed in Otey and Carpen 2004). Many of these interacting proteins are involved in cell signaling and regulation of transcription. One such protein, zyxin, has been demonstrated to mobilize from focal adhesions to stress fibers in response to cyclic stretch (Yoshigi et al., 2005). In addition to being a focal adhesion component, zyxin is also a mechanosensitive transcription factor (Cattaruzza et al., 2004). The Z-disk of striated muscle is also described as a mechanosensory signaling interface (Frank et al., 2006). Here, α-actinin cross-links opposing actin filaments to form the Z-disk lattice while also interacting with titin Z-repeats (Sorimachi et al., 1997; Young and Gautel, 2000). Recent work has suggested a role for a titin/Tcap/MLP/α- actinin complex as a stretch sensor that regulates cardiac hypertrophy. Mutations of these proteins lead to a lack of response to overstretching resulting in hypertrophic cardiomyopathy and muscular dystrophies (Hayashi et al., 2004). These observations have suggested that α-actinin is more than a simple actin cross-linker but is also a scaffolding protein on which many additional factors bind and interact to specify α- actinin’s function more precisely. The cytoskeleton is emerging as an integral component of cell tension sensing and signal transduction machinery. In order to further the understanding of α-actinin’s cross- linking function it is necessary to observe the intact molecule within an appropriate context. Here we examine the range and flexibility of α-actinin binding and cross-linking of actin filaments in a lipid monolayer environment by employing correlation averaging, correspondence analysis, and classification schemes (Frank, 1996) to increase the signal-to-noise ratio (SNR) of the averages while retaining the variability of the α-actinin cross-links. We can distinguish the individual domains within the dimer in these averages and can quantify the angular distribution and lengths of the cross-links. We observe the occasional incorporation of antiparallel filaments into otherwise parallel

36 arrays and determine that the antiparallel cross-links are no different in angular distribution or length from the parallel cross-links. We also report the intriguing and frequent occurrence of both α-actinin ABDs binding to a single actin filament. Most notable is the range of lengths measured for this molecule which suggests that α-actinin combines angular flexibility with linear elasticity in actin bundle formation.

2.2 Results

The typical micrograph had four or more actin filaments within the raft and from one to five α-actinins bound to each crossover repeat. Although the cross-links are clearly visible in the micrographs, the molecular details are not. We observed initially that the α- actinin cross-links all originate from the wide part of the actin crossover repeat with virtually no binding to the narrow portions. The cross-links are likely constrained to this periodicity by the lipid monolayer. The symmetric sites located at ±90° are blocked by the lipid on one side of the filaments while the other side faces bulk solvent, where there may be too few actin filaments free in solution to be “grabbed” by the unbound end. Both actin and α-actinin are acidic proteins and strongly partition to the basic monolayer so that the interactions are constrained to occur in the 2-D phase. This is exactly the intent of the monolayer system as by keeping the specimen planar it imitates the effect of a membrane and keeps the specimen in an ideal form for electron microscopic examination. The polarity of the filaments cannot be determined in the original micrographs and there is no obvious change in the cross-linking pattern that would suggest a change in actin filament orientation. To obtain this level of information requires computation of averages with a higher SNR.

2.2.1 Alignment and Correspondence Analysis The straightforward approach would involve alignment and classification of the crossover repeats to produce class averages. We refer to this as the simultaneous left- right approach because all cross-links in the repeat contribute to the classification. To eliminate or reduce actin filament alignment errors as one source of variability all crossover repeats were aligned on a reference actin filament and subjected to several cycles of classification and multireference alignment. During this process, it became

37 clear that some class averages, even from repeats within a single micrograph, were oppositely oriented compared to the majority. Since the linkage between repeats and their original micrograph was always maintained, it was easily determined that these oppositely oriented repeats came from a small subset of actin filaments (Fig. 2.2.1.1). Thus, bundles previously judged to have actin filaments with a single orientation were actually a mixture of parallel and antiparallel filaments. Note that there is no discernable difference in the cross-linking pattern between antiparallel actin filaments and parallel actin filaments in the original micrographs.

Figure 2.2.1.1 α-Actinin-F-actin raft. Polarity of the horizontal actin filaments is indicated by the black arrows. White arrows point to mono-filament-bound α-actinin cross-links. Note the formation of girder-like struts.

The mixed orientation of the actin filaments in these rafts contradicts earlier studies that determined the rafts contained actin filaments with a single orientation (Taylor and Taylor, 1994). That study utilized optical diffraction, changes in cross-links at inserted actin filaments within the rafts as well as the presence of actin filament spirals with α- actinin cross-links to argue bundle polarity. While cross-linking polarity in the spirals is indisputable, the other two approaches are subject to error. For example, small numbers of oppositely oriented actin filaments would not contribute significantly to the optical diffraction intensity which would report the dominant structure. The antiparallel

38 cross-links account for ~15% of the ~8,000 total in this study. This would affect the diffraction intensity by 0.152 = .02, or 2% of the intensity. In addition, as shown in this analysis, inserted actin filaments (one of which is seen at the left of Fig. 2.2.1.1) are a weak discriminator of actin polarity in α-actinin bundles since the cross-linking pattern is unchanged by changes in filament orientation (see below). Correlation averaging as used here is much more powerful for establishing actin filament polarity. Before applying correspondence analysis to the cross-links, all repeats were first rotated to the same actin orientation and aligned on the actin filament. A global average as well as eight class averages was then computed using a classification mask that included only a single crossover repeat centered on the wide part of the actin filament (Fig. 2.2.1.2). All of the averages show great detail in the actin filament, although the actin density becomes blurred at the ends indicating departures from strict linearity. Even given the detail in the actin filament, the averages do not show any α-actinin even in the wide part of the filament where the original micrographs show it to exist. The only hint of its presence is the appearance of distinct densities on either side of the actin crossover that we attribute to the ABDs. This blurring of the cross-links in the average emphasized their extreme variability. Although there are some differences between the classes, the axial position of the actin monomers is identical. This suggested that the differences were primarily in the amount of filament bending at the ends of the repeats and the azimuthal orientation of the actin filament within the plane of the raft. In the global average the resolution by the Fourier Ring Correlation was 1.6 nm indicating that the actin filament alignment is very accurate (van Heel et al., 1982).

Figure 2.2.1.2 A global average of 7,903 actin crossover repeats and eight class averages based on the variance. Some curving of the filaments is present as can be seen in the averages.

Theoretically, α-actinin can pivot a full 180° about its actin-bound ABD, thus creating a semicircle of probability centered about the crossover repeat on each side of the actin filament. This range of flexibility produces considerable variation especially when both sides of the actin filament are considered together. A battery of different classification masks that excluded the pre-aligned actin filament were applied to the aligned repeats. Of all masks tried, none worked better, as judged by detail recovered in the averages, than a simple circle with a radius long enough to contain the length of α-actinin.

Figure 2.2.1.3. Comparison of whole image classification to left/right classification. The aligned, raw repeat in the left-most column is presented next to the results of both classification methods. The center column produced using a classification mask covering both sides of the filament, but excluding the filament itself. Areas in white are included in the classification. The method in the right column classifies on the left and right sides independently and then reassembles the averages to recreate the original repeat. Overall, the results of the in-dependent left-right method compare more favorably to the raw repeat than simultaneous classification of both sides.

41 Two classification schemes were used. The initial classifications used a circular mask that included cross-links on both sides of the actin filament but excluded the pre-aligned actin filament. These classifications were not very satisfactory (Fig. 2.2.1.3) in that many α-actinins easily identified in raw repeats were not represented in the averages. Simply increasing the number of class averages only served to diminish the improvement in the SNR obtained by averaging. To improve on the number of cross-links retained in the class averages, but without sacrificing SNR improvement, we computed independent class averages of the two sides of the actin filament using semicircular left and right masks. This was based on the lack of any obvious correlation between cross-links on the left and right sides of the actin filament. Afterwards the class averages could be cut in half down the center of the aligned actin filament and “pasted” back together to regenerate the original repeat. This approach is capable of computing a “class average” for every original repeat using an optimal number of classes. The left-right reassembled classes appeared to recover cross-links in the averages with better accuracy when compared to original images than the simultaneous classification and did so without sacrificing SNR improvement. It should be noted however, that some of the α-actinins visible in the original images were not picked up by classification even with this scheme. This is likely a consequence of the extreme amount of variation present. Patterns of cross-links that are statistically too infrequent to form a cluster of their own will be grouped instead with repeats that may lack one or more of its cross-links. Averaging will then eliminate the odd cross-links. To evaluate the accuracy of the independent left-right method compared with the simultaneous left-right method, 100 original repeats containing 264 cross-links and the class averages to which they contributed were chosen at random and the number of visible cross-links in the original and two types of averages were scored (Table 2.2.1.1). Independent evaluations by the three authors scored the number of cross-links that correctly appeared in the averages, the number of false negative cross-links (present in the original repeat but not picked up by classification), and the number of false positive cross-links (present in averages but not found in the original repeat). Cross-links in class averages generated by simultaneous left-right classification were correct ~30% of

42 the time. The class averages reassembled from independent left and right classes were correct ~60% of the time. The numbers of false positive cross-links were relatively consistent between the two methods. Thus, the independent classification was twice as effective at retaining the cross-links in the reassembled averages. Table 2.2.1.1. Assessment of Classification Accuracy

Simultaneous Classification Independent Classification

Total Correct False False Correct False False

Cross- Negative Positive Negative Positive

links

1 264 72 192 17 154 110 19

2 264 88 176 33 154 110 30

3 264 68 196 15 143 121 22

Avg 76 188 22 150 114 24

% 29% 71% 57% 43%

False negatives in this analysis represent cross-links that were present in some of the members of the class but with low frequency compared with other cross-links. These cross-links are blurred out when averaged. False positives are cross-links that exist in the data as they would otherwise disappear in the average. Their presence in the class average means that they were present in most members of the class, but not in the particular original repeat that was being scored. The percentages in Table 2.2.1.1 do not represent the true error rate. Instead, they represent the percent of occurrence within a sample population. For example, a given cross-link that is absent from the class average may represent only one raw repeat out of 88-90 individual class members. The true error rate would then be on the order of ~1%.

43 2.2.2 Distribution of cross-link angles and lengths Angles were measured from left and right class averages from the ABD on the actin filament outward. Numbers of crossover repeats included in each average were tallied for each angle in 10° increments. Histograms of the numbers of cross-links falling into a 180° range of possible angles show that the majority fall into 0°, 60°, 90°, 120°, and 180° bins (Fig. 2.2.2.1 A). The 60°/120° cross-links are strut-like, while 90° cross-links appear as ladder rungs. Very often an actin filament crossover would have multiple cross-links of different angles which would suggest variations in length as well. Interestingly, the angular distribution of antiparallel cross-links is not very different from the parallel cross-links (Fig. 2.2.2.1 B). This is contrary to the prediction (Taylor et al., 2000) that the angular range should be narrower in bipolar cross-links because the helical nature of the actin filament would place alternating cross-links on opposite sides of the interfilament axis, i.e. both ends of cross-links between any pair of antiparallel actin filaments would be on the same side of the interfilament axis, either adjacent to the monolayer or in the bulk aqueous phase. For parallel cross-links, one end would be close to the monolayer and the other end closer to the bulk phase. The accuracy with which cross-links are recovered in the classes affects the accuracy of the histogram, but probably not its overall shape, nor the conclusions that can be drawn from it. The false negative rate from separate left and right classification is 43% (Table 2.2.1.1). These cross-links are lost from the distribution because they did not contribute to the class average. It is reasonable to expect that they would be spread over the distribution because concentration in a particular orientation would cause them to appear in the averages. False positives (24%), which are measured and included in the distribution, overestimate the number of cross-links at a particular angle because some members of the class lack it. Correction for this would reduce some frequencies, but even if highly biased, would not affect the conclusion which is that cross-link angle is highly variable but preferentially oriented at 0°, 60°, 90°, 120°, and 180°.

Figure 2.2.2.1 Angular distributions of total and antiparallel cross-links. A, Illustrates the angle convention used. B, Cross-link angles measured from left and right class averages of all repeats. Distribution is nearly symmetric because left and right angles are complementary, except for cases where the repeat containing the opposite end of the cross-link was not extracted. C, Cross-link angles measured from left and right class averages of antiparallel repeats. The distribution is lopsided due to twice as many filaments having right-side antiparallel cross-links. Preferred orientations overall were the ~60/120° “struts”. Total cross-links also strongly preferred 0/180° "monofilament" binding.

Figure 2.2.2.2. Examples of 60° cross-links realigned and classified on α-actinin. Note the blurring of distal actin filaments.

To obtain an accurate measurement of the length of α-actinin as a cross-linker, we selected all molecules oriented in a particular angle, 60°, and aligned each of the repeats on the α-actinin itself. However, when class averages were computed, we discovered that the distal ends of the molecules were too variable for accurate measurement (Fig.2.2.2.2). The central actin filament is usually aligned, but the second, outer actin filament is quite blurred due to translational/rotational variations. This suggests that the α-actinin length is variable and that the angle of the cross-link with respect to actin is not symmetrical with respect to the α-actinin.

Figure 2.2.2.3. Mapped-back images. A and B represent direct comparisons of the raw image cross-link on the left and the re-assembled cross-link placed into the original coordinates for both parallel and antiparallel cross-links. Each half of the cross-link is contributed by a different class average. Each panel is 157 X 78.5 nm. The thin lines in the first column are all the same fixed length and illustrate the variation in total length of the α-actinin cross-link. The heavy lines in the first row are the same length in each panel and illustrate that the reassembled averages have identical positioning to the original motif which validates our use of these averages for measurement. C, Enlarged view. One can almost discern the four spectrin repeats in the rod domain, the CaM domain density, and a tiny nub of density nestled between the actin monomers attributable to the ABD.

Figure 2.2.2.4 Map-back before and after comparison. A, The original micrograph image. B, Individual class averages (masked with a sphere) reinterpolated onto the image itself. While not extremely accurate, the filaments do line up surprisingly well. The difficulty posed by the high variability of even this highly populated class of cross- links was surmountable by “reassembling” cross-links from the left and right class averages (see Methods). In these correctly reassembled images, we were able to detect the density belonging to each ABD as well as the R1-R4 repeats because of the improved SNR. These “mapped-back” images (Figure 2.2.2.4) reveal the α-actinin

48 molecule from ABD to ABD, which is not possible in the original images. Traditionally these measurements would have been taken from the center of the actin filaments, which is the only possible measure in the original micrographs and each measurement would have measurement uncertainty because of the low SNR in the raw images. Each reassembled cross-link originates from two different class-averages (Fig. 2.2.2.3). However, because the classification and averaging is accurate only ~60% of the time, reassembled images appear to correctly line up only about 35% of the time. Cross-links measured from ABD to ABD centers averaged 33.9 ± 2.5 nm. This compares favorably to the 34.5 nm α-actinin length estimated from 2-D arrays (Taylor and Taylor, 1993; Tang et al., 2001). In many cases the ABD projection density appears to contribute only half as much intensity as might be expected from two superimposed CH domains placed on the actin filament. Our data are limited to projections at the moment so it is not possible to speculate on whether or not CH2 is actually bound to actin or to CH1 in the closed conformation.

2.2.3 α-Actinin binds to a single actin filament

An unexpected observation from these data is the frequency with which α-actinin binds with both ABDs onto single actin filaments. The α-actinins at first seemed to be aligning parallel to the actin filament as a consequence of drying in negative stain. However, a suggestion that the interaction is specific can be seen in the original micrographs (Fig. 2.2.1.1) where α-actinin ends are clearly placed on the actin filament crossovers. “Cross-links” at 0° and 180° occur with close to the same frequency as the 60° and 120° struts (Fig. 2.2.2.1). With the repeats centered on the crossover only one end of the molecule was clearly visible in the averages and although a length could have been obtained by combining successive crossovers, for better accuracy we reextracted the repeats with a larger box size to facilitate alignment centering on the narrow part of the filament between successive crossovers. Class averages could then be computed that contained both molecular ends of α-actinin. These averages show clear association of the ABD with the actin filament as a density comparable in size to that seen in a true cross-link and with both ends of the molecule placed on actin in a regular manner consistent with a specific interaction (Fig. 2.2.2.5).

Figure 2.2.2.5. Models of α-actinin cross-linking a single actin filament. A and C, class averages representing long and short α-actinin binding to actin. B and D, long and short projection images in reverse contrast generated from PDB files modeled in O. E and F, images created in Chimera.

50 The two α-actinin ABDs span successive crossovers on the same filament so that the separating distance is quantized in length increments of 5.5 nm, the distance of one actin monomer along the filament (Fig. 2.2.2.5 A, C). To help explain how α-actinin can accommodate this variation in length we constructed atomic models (Fig. 2.2.2.5 E, F). When filtered to a similar resolution and projected these models compare favorably with the class averages (Fig. 2.2.2.5 B, D). Of 4754 total repeats showing monofilament binding, 44.3%, fell into classes with a length of 35.9 nm, corresponding to 6.5 actin monomers along the filament, while 31.1% fell into classes with a length of 30.4 nm, equivalent to a length of 5.5 actin monomers. A total of 24.6% fell into classes whose lengths were ambiguous. This proportion of poorly defined images is not unusual in single particle reconstruction. In building the long monofilament model, we extended the ABD-R1 linker to its maximum length consistent with a β-strand conformation. However, this may not be the only option. Less extension would be required if the CH2 domain of the ABD adopted an open conformation, rather than the closed one modeled here. CH2 domain movement could then contribute to the separation between CH1 domains and the rod in the long monofilament model. CH2 is not necessary for actin binding and does not bind actin by itself (Way et al., 1992b; Gimona and Winder, 1998). For the short model, the ABD-R1 linker would have to be compressed and there are multiple ways to accomplish this, too many to model effectively.

2.3 Discussion

α-Actinin was originally discovered as an actin cross-linking protein (Masaki et al., 1967) but has since been shown to bind many cytoplasmic and membrane proteins. However, actin binding remains its most studied property. In vivo, α-actinin cross-links actin filaments into bipolar structures at the Z-disk, cytoplasmic dense bodies, and stress fibers. In the well ordered Z-disk cross-links of a length consistent with α-actinin are observed (Luther, 2000), while the angles of the cross-links have been estimated to be ~35° with respect to the actin filament (Goldstein et al., 1979). The cytoplasmic dense bodies of smooth muscle, which are known to be the anchor points of oppositely oriented actin filaments (Bond and Somlyo, 1982), are much less

51 well organized and clear cross-links due to α-actinin are not as easily seen as in Z- disks. Likewise, the cross-links in stress fibers are not unambiguously identifiable as α- actinin. α-Actinin’s presence in these structures is nonetheless undisputed. The role of α-actinin at membranes, when actin filaments are oriented in the same direction is much less clear. Unlike with Z-disks, the structure of F-actin-membrane contacts is very obscure. Often, the identification of α-actinin at these actin-membrane contacts is done at the light microscopic level. It has been demonstrated that protein within the focal adhesion is relatively inaccessible by antibodies (Pavalko et al., 1995) and hence localization requires the expression and correct targeting of GFP-tagged α- actinin (Dabiri et al., 1997). At structures such as focal adhesions, α-actinin could have two roles, (1) anchoring actin filaments to membrane receptors such as integrins (Otey et al., 1990) and (2) cross-linking parallel actin filaments at their ends in Z-disk-like structures of stress fibers. The present study is relevant to both polar and bipolar actin filament bundles, but is perhaps more germane to polar bundles of actin filaments at cell membranes.

2.3.1 α-Actinin-Membrane Interactions

Our monolayer system was developed to simulate interactions between cytoplasmic proteins and membrane receptors on the inner leaflet of the cell membrane. The fluidity of the lipids combined with protein-lipid interactions facilitates sorting of the proteins into energetically favorable arrangements in a 2-D phase. The system has been used to identify the β1-integrin binding site on α-actinin (Kelly and Taylor, 2005) and a ternary complex between α-actinin, the β1-integrin cytoplasmic domain and the first 258 residues of the vinculin head (Kelly et al., 2006). The present study was undertaken to test the feasibility of producing focal adhesion complexes involving F-actin and to identify a plausible image processing scheme to enhance the SNR without loosing the structural variability in low contrast, frozen hydrated specimens. The detail obtainable from the original micrographs is limited by the SNR. What has been lacking is a suitable analysis scheme to bin the highly variable cross-links for averaging. Our strategy of independent left-right classification followed by reassembly recovers enough cross-link variability that reassembled class averages can be mapped-

52 back to the original image to create large numbers of cross-links with enough detail to make measurements in a way that has not been possible before now. The accuracy of cross-link recovery could be improved by computing more class averages, but at the expense of the SNR in the averages. The 2-D rafts of F-actin and α-actinin show highly heterogeneous orientations and lengths for the cross-links, but their binding sites on actin are highly regular. The present result also shows that α-actinin can “cross-link actin crossovers,” an observation that broadens the range of possible α-actinin functions especially at the cell membrane.

2.3.2 Variability of bipolar cross-links

The present study demonstrates a promiscuous polarity preference for α-actinin: F- actin binding. Although 3-D images of the Z-disk reveal a highly regular and bi-polar arrangement of actin filaments cross-linked by α-actinin, the actual lengths of α-actinin linkers are inferred from inter-filament spacing which varies from 17-24 nm (Goldstein et al., 1979; Luther, 2000). This difference is related to differences between the small square and the basket-weave lattice form. The number of actin crossover repeats and α-actinin linkers in the Z-disk also vary across fiber types, and it was suggested that titin isoforms of varying numbers of z-repeats may be a determinant (Luther et al., 2003). This suggests that there may be a number of extrinsic factors which govern the angle and polarity of α-actinin binding in cellular structures. These factors could be binding partners or mechanical effects.

The PIP2 binding to the CH2 domain of α-actinin induces a conformation that facilitates binding to titin, the Z-disk ruler, via two distinct interactions that alleviate a “pseudoligand” interaction between the CaM domain and the ABD-R1 flexible linker domain (Young and Gautel, 2000). The Actinin-associated LIM Protein, ALP, has been shown to strengthen the binding of α-actinin to actin (Xia et al., 1997; Pomies et al., 1999; Pashmforoush et al., 2001). This protein binds α-actinin at both the CaM domain and the R1-R4 domain, crossing the flexible linker domain (Klaavuniemi et al., 2004). These authors postulate that the binding of ALP across this region may stabilize a

53 conformation of α-actinin within the Z-disk that specifically limits it to antiparallel cross- linking. Tension applied to the cross-links may also select a particular group of cross-linking orientations. The most common cross-linking orientation is the 60°-120° triangular struts which we observe in both the parallel and antiparallel cross-links. When tension is applied across the Z-disk, the 60° anti-parallel cross-links will be placed under extension, probably the more favorable case since these are retained, and the 120° cross-links will be placed under compression which may favor dissociation. Tension applied to parallel actin filaments cross-linked into a polar bundle would not experience the same effect unless different amounts of tension were applied to the individual actin filaments.

2.3.3 Significance of “monofilament binding”

Our data suggest the ability of α-actinin to bind from crossover to crossover on a single actin filament. As far as we know this is the first demonstration of this phenomenon, and raises the question of whether the length of α-actinin, which is close to the distance between actin filament crossovers, has evolved to maintain this monofilament binding capability rather than or in addition to an ability to cross-link actin filaments. Our work shows that variability in cross-linking angle is inherent in α-actinin; the separation between actin filaments in an α-actinin-actin bundle is therefore not an inherent property of α-actinin alone. Only the “struts” seem to maintain regular actin filament spacing. Other than simple coincidence, what biological role could α-actinin monofilament binding perform? One possibility may be to sterically block interaction with other proteins. That is, regions of actin with α-actinin bound alongside would be excluded from interactions with other actin-binding proteins that induce branching, severing, or depolymerization. Another possibility could be to anchor individual actin filament to membrane-associated adhesion proteins or channels. Many of these interactions, such as the α-actinin integrin interaction, have been characterized and involve the R1-R4 domain. Binding of α-actinin alongside, in fact, between the actin filament and the membrane, would present the entire length of the R1-R4 domain and hence two binding

54 sites to the membrane surface thereby facilitating binding to adhesion proteins such as integrins. Computer modeling of how α-actinin-vinculin-F-actin cross-links might appear on a lipid monolayer system suggested that simultaneous integrin binding and F-actin cross- linking was most likely to occur when the actin filaments are oriented away from the membrane whereas actin filament cross-linking alone was possible when the actin filaments run tangential to the membrane (Kelly et al., 2006). Monofilament-bound α- actinin might facilitate linkage of tangential actin filaments to transmembrane elements such as integrins without forming cross-links. Comparative modeling of the monofilament-bound α-actinins and the variation in total length of the cross-links raises the question of how this variation can be accommodated structurally. When building our comparative models for monofilament binding it was necessary to straighten this linker to accommodate the distance between the ABDs in the longer model. Although the α-actinin spectrin repeats can be unfolded with forces up to 40 pN (Ortiz et al., 2005), we do not expect that the molecules are experiencing such pulling forces at the monolayer surface in our specimens and there is no discernable change in length or morphology of the R1-R4 domain density that would suggest that this is occurring. One likely candidate then is the 25 amino acid linker region between the ABD and R1. At the moment there is no structural data on this linker. It has been demonstrated that the CaM domain can bind to this linker and its binding is relieved by PIP2 binding to the ABD, freeing the CaM domain for interaction with the titin Z-repeats (Young and Gautel, 2000). The length increase might also be accommodated if the ABD adopted an open conformation freeing CH2 to contribute to the separation. The CH2 domain is not required for actin binding as it only has one actin binding sequence and cannot bind actin on its own (Way et al., 1992a). The α-actinin ABD alone binds actin in a closed conformation; however, 2-D crystal structures suggest that the ABD can in fact assume an open conformation. Our data, which show a smaller density on actin than would be suggested by a combined CH1-CH2 ABD, suggest that CH2 may not be involved in actin binding in the intact molecule.

55 2.3.4 Significance of variable length of cross-links

α-Actinin has long been considered to be a fixed-length cross-linker based on its appearance in the Z-disk of striated muscle (Masaki et al., 1967). The present study shows that α-actinin length when bound to actin can vary by more than 5.5 nm. We think that the length variation could facilitate a role for α-actinin in sensing tension within actin filament assemblies such as Z-disks and focal adhesions. Although α-actinin is best known as an actin cross-linking protein, it has many other binding partners, some of which are known to be involved in tension sensing at the Z- disk (Hayashi et al., 2004) and at integrin-linked focal adhesions (Bershadsky et al., 2003). We think that α-actinin itself is not likely to be the tension sensor but is more likely to act as a scaffold for tension-sensing signaling proteins. The present results suggest that α-actinin has the ability to respond to a change in tension by lengthening and could do so within limits without dissociating from actin. Hence it can maintain the integrity of the structure while responding to the stress. α-Actinin binding partners can then respond by dissociating. This would be facilitated if the binding partner bound α- actinin weakly but at more than one site, for instance the ABD and R1-R4, or bound both α-actinin and F-actin weakly. Then a stretch would cause dissociation from one site which may then facilitate dissociation from the other site followed by diffusion to a distant effector. As long as tension keeps the α-actinin extended, the tension sensor would be unlikely to reassociate with the structure. An example of a tentative tension- sensing complex at the Z-disk is the titin/Tcap/MLP/α-actinin complex (Hayashi et al., 2004). Mutations in MLP LIM domains that diminish its binding with α-actinin are linked to cardiac hypertrophy (Geier et al., 2003; Gehmlich et al., 2004). An example of a tension-sensing event at focal adhesions is the relocation of zyxin, another LIM-domain protein, from focal adhesions to stress fibers in response to cyclic stress (Yoshigi et al., 2005). The specific interaction of zyxin with α-actinin has been well documented (Crawford et al., 1992; Reinhard et al., 1999; Li and Trueb, 2001). Seemingly, the PDZ and LIM domain proteins that bind α-actinin are associated with signaling cell response to cytoskeletal changes.

56 Note that tension sensing in the focal adhesions may be different from tension sensing in Z-disks. In Z-disks α-actinin is stretched by opposing tension applied to actin filaments across the structure. In the focal adhesion, α-actinin acting as a cross-linker of parallel actin filaments would most likely not respond directly to the stretch. The stretch sensing would have to occur by pulling on α-actinins that are bound to another structure, such as the integrins. Tension sensing might then involve only a subset of the α-actinins in the structure. Here we have applied a hybridization of techniques to overcome disorder in our samples. Our scheme of alignment to a central element, left/right independent classification, reassembly, and mapping-back to the original coordinates effectively captures the variability of the α-actinin cross-link. This same technique can similarly be applied to tomograms of the same samples plus other adhesion-related proteins in vitreous ice to achieve new detail in our understanding of the interactions of adhesion proteins at the membrane.

2.4 Materials and Methods

2.4.1 Protein Purification

Rabbit skeletal muscle α-actinin was purified from a myofibril preparation dissolved in

1.0 M NaCI, 25 mM Tris, 2 mM MgCl2, 0.02% β-mercaptoethanol, pH 8.0 followed by hydroxyapatite column purification (Taylor et al., 2000). Actin was prepared from rabbit muscle acetone powder (Pardee and Spudich, 1982), followed by a Superose-12 column. Fresh G-actin was prepared by dialysis overnight against Buffer A: 2 mM Tris-

Cl, 0.2 mM Na2ATP, 0.02% β-mercaptoethanol, 0.2 mM CaCl2, 0.01% NaN3, pH 8.0. The protein was clarified by high-speed centrifugation prior to sample set-up. Arrays were grown on positively charged lipid monolayers (Taylor and Taylor, 1994). A 3:7 volume ratio of DLPC: DDMA lipids (Avanti Polar Lipids) at 1 mg/mL in chloroform was layered over a solution containing 0.3 µM actin and α-actinin in phosphate-buffered polymerizing buffer. The actin was polymerized in the presence of α-actinin at 4 °C.

57 2.4.2 EM data collection The 2-D arrays were recovered using 200-300 mesh copper grids with a reticulated carbon support film (Kubalek et al., 1991). Samples were stained with 1% uranyl acetate, dried, and stabilized by a thin layer of evaporated carbon. Data were collected on Philips EM300 and EM301 electron microscopes at 80 keV. Micrographs were scanned on a Z/I Imaging PhotoScan 2000 scanner (Carl Zeiss, Oberkochen, Germany) at either 7 or 14 µm.

2.4.3 Image Processing Images were sampled and scaled to one another using the internal standard of the actin filament 5.9 nm layer line of the Fourier transform. This was achieved by cutting out single actin filaments from each micrograph and straightening using Phoelix (Whittaker et al., 1995). Filaments were then Fourier transformed and the number of pixels along the meridian to the 5.9 nm layer line was determined. All micrographs were resampled and scaled to 0.436 nm/pixel. The contrast transfer function (CTF) for each micrograph was corrected by phase flipping. Defocus and astigmatism angles were measured using the program ICE (Interactive Crystallographic Environment) (Wild et al., 1995). Images were normalized by setting the mean pixel value to 0 and the SD to 1. Repeat coordinates centered on each actin crossover from each micrograph were manually picked using the EMAN Boxer utility (Ludtke et al., 1999). Raw repeats were extracted as 180 X 80 pixel boxes and compiled into pseudo 3-D stacks. For classification of monofilament-binding α-actinins the repeats were re-extracted as 250 X 250 pixel boxes.

2.4.4 Classification Repeats were aligned to the central actin filament using in-house software for alignment and multivariate statistical analysis. Classifications were done using 8, 16, 24, 32, and 64 factors. Eigenimages were evaluated by eye and by plotting the variance and those seeming to contain only noise were left out of the classification. The resulting classes from each set were examined for completeness and resolution until it was determined that 32 factors gave the best classification result as judged by the detail

58 seen in the averages. In order to improve the amount of structure variance that can be recovered without sacrificing improvement in SNR, a left-right classification scheme was used. The CTF-corrected repeats were classified on the left side cross-links and the right-side cross-links separately. The repeats were then sorted into 88 classes left and 88 classes right; 88 being the square root of the total number of repeats, n. Because the SNR improves as the square root of the total number of images in each class average, the benefit of classifying each side separately followed by reassembly results in a SNR improvement of n¼ while capturing the variability as high as n. No images were discarded in the process as this leaves large gaps in the reassembled rafts images. The resulting class averages from left and right were combined to reassemble the original repeat with an average 9.4-fold improvement in the SNR. Measurements of angular distribution were made from these averages. These class averages, however, were somewhat blurred at the distal ends of the α- actinin molecules which prevented accurate length measurements. We first tried to align cross-links within a particular class, 60°, to the α-actinin itself. This resulted in a relatively well aligned central actin filament and α-actinin, but the distal filament was blurred due to translational differences and made the distal ABD hard to see. Instead, we reassembled these averaged repeats and mapped them back to the original micrograph. The inverse of the alignment shifts and rotation angle were applied to the repeats and each averaged repeat was placed back into the original image. First, we applied an oval mask with an apodized edge to each repeat. Then we applied the inverse of the shifts and rotations used in alignment and rescaled the repeat to its original parameters. Each repeat was then precisely inserted back onto the original micrograph from the original x and y coordinates. The reassembled image was then used to make length measurements.

2.4.5 Modeling Models of cross-links were generated in O (Jones et al., 1991). The PDB coordinates of α-actinin from Liu et al. (2004) were used to position the ABDs onto F-actin generated from the helical parameters of the averaged images which were determined by the ratio of the axial spacing of the 5.9 nm layer line to the first layer line. The MMTSB tool set

59 (Feig et al., 2004) was used for manipulating the PDB files and positioning the spectrin- repeat rod domain. The EMAN utility pdb2mrc (Ludtke et al., 1999) was used to generate projections of the models which could be filtered and compared to the averaged repeats. Final images were prepared with Chimera (Pettersen et al., 2004).

60 CHAPTER 3 α-ACTININ-INTERACTING PROTEINS

3.1 The Alliance for Cellular Signaling (AfCS)

The AfCS Project was conceived under the Glue Grant Initiative of the National Institute of General Medical Sciences. The purpose of the initiative is to provide resources to currently funded scientists to work as a group to tackle complex problems that are of central importance to biomedical science and to the mission of NIGMS, but that are beyond the means of any one research group. A high level of resources may be requested to allow participating investigators to form a consortium to address the research problem in a comprehensive and highly integrated fashion. There are five such groups currently funded: Alliance for Cell Signaling (AfCS), Cell Migration Consortium (which funds the work covered in this dissertation), Inflammation and the Host Response to Injury, Consortium for Functional Glycomics, and LIPID MAPS Consortium. The AfCS website serves the entire scientific community as a source of useful reagents (i.e. DNA constructs), signaling information (Nature Molecule Pages), lab protocols, and experimental data from hundreds of experiments that are free and accessible to the whole community. Towards the signaling information goal, the Alliance for Cellular Signaling and the Nature Publishing Group have teamed up to produce the Cell Signaling Gateway (http://www.signaling-gateway.org/). The AfCS has a list of nearly 4000 proteins involved in signaling reactions that are in a database format. This free, open access database of cell signaling molecules and their interactions (currently only ~200 peer- reviewed and published--a bottle-neck in the process) is redefining electronic publishing. Each molecule is the topic of a Molecule page with automated data taken from public databases along with invited, author-entered data from the literature. Molecule pages are peer-reviewed anonymously by Nature Publishing Group, with the assistance of the Editorial Board, and published on the Nature Signaling Gateway. They are formally citable using digital object identifiers and will be listed in PubMed. They will also require updating by the authors annually. The end-goal of such a repository is to

61 create a virtual cell, establishing connectivity in signaling networks via circuit maps and predicting effects of perturbations. In the process of reviewing the literature for this project it was determined to be of benefit to the author to combine both tasks. When the work on α-actinin was begun in 2002, there were no review-type articles involving this protein. Thus, the authoring of the Nature Molecule Pages also fills an important role for the cell biology community. Also, in consideration of potential future elements to be added to build on the the α- actinin:F-actin rafts studies here, it became necessary to list in detail all of the known α- actinin–interacting proteins. The results are listed in Table 3.1. There are no less than 92 proteins that have been demonstrated to interact with α-actinin.

3.2 Molecule Page Navigation

The molecule database is searchable by keyword or can be browsed for a protein of interest. The researcher is first linked to the overview page which reports data automatically retrieved from existing databases, such as protein sequence, molecular weight, etc. At first glance the researcher can learn the protein’s AfCS ID, name, and all other existing names, a functional categorization of the protein type, and gene symbol. The left-hand navigation tool-bar links to author-entered and peer-reviewed data, as well as most relevant automated data from multiple databases. The author-entered data are divided into six sections. The summary page is a published literature review of the protein’s basic function, how its’ activity is regulated, ligands and other interacting proteins, regulation of concentration, subcellular localization, major sites of expression, cellular phenotypes, known splice variants, and listings of known sources of antibodies. Each summary section is followed by a detailed reference section with clickable links to the PubMed accession numbers. This allows for a fast, targeted query for researchers new to a specific protein.

Figure 3.2.1 Navigation tool-bar.

63 Protein Function

α-actinin-2 (Actn2) is a member of the spectrin-family of actin crosslinking proteins. It was originally found in skeletal muscle and classified as a muscle type α-actinin. Later, it was also found in the brain at glutamatergic synapses and dendritic spines, thus making the original isoform designations “muscle” or “non-muscle” actinin obsolete. Its structure is the same as that for the other actinins. The N-terminus consists of two highly conserved tandem calponin-homology domains, which are required for actin binding, and are connected to a series of four spectrin-like coiled-coil repeats that make up the rod domain facilitating dimerization of the molecules. Once regarded as simply spacers for actin bundle formation, these spectrin repeats have a newfound role as a docking site or scaffold for many signaling proteins (Djinovic-Carugo et al. 2002). The repeats are followed by a calmodulin-like domain that is composed of two EF hand domains. The EF hands for Actn2 and Actn3 have lost the ability to bind calcium, yet the structures remain evolutionarily conserved which suggests some other major functional role. Its function in skeletal and cardiac and smooth muscle cells is to constitutively anchor the actin filaments at the Z-disks/dense bodies, while in neurons it anchors the NMDA receptor to the cytoskeleton. Actn2 exists as antiparallel homodimers, or it can form heterodimers with Actn3 in type 2B muscle cells (Beggs et al. 1992; Tiso et al. 1999; Mills et al. 2001; Dixson et al. 2003; Otey and Carpen 2004; Virel and Backman 2004).

Pub PM ID Authors Title Journal Date

Beggs AH, Byers TJ, Knoll Cloning and characterization of two human J Biol 5 May 1339456 JH, Boyce FM, Bruns GA, skeletal muscle alpha-actinin genes located on Chem, 267, 13 1992 Kunkel LM chromosomes 1 and 11.

Dixson JD, Forstner MJ, The alpha-actinin gene family: a revised J Mol Evol, Jan 12569417 Garcia DM classification. 56, 1 2003

Djinovic-Carugo K, Gautel The spectrin repeat: a structural platform for FEBS Lett, 20 Feb 11911890 M, Ylänne J, Young P cytoskeletal protein assemblies. 513, 1 2002

Figure 3.2.2 Excerpt from the protein summary page. This section highlights the protein functions and is directly linked to the PubMed citation for accelerated literature review.

The next section is the protein states page, a detailed listing of every published form of the protein and cellular location. The list begins with the protein itself, followed by dimerization (in the case of α-actinin), post-translational modifications, ligand-binding, and complexes with other interacting proteins. Although the interacting proteins are listed by their abbreviated names, each is clickable and linked to a brief descriptive file with full protein information. For each state described a link exists to a graphical depiction of the input, modification/reaction, and output creating that state.

Figure 3.2.3 Excerpt from the states page and resulting link to the transitions page. Clicking on a given state description, such as that for the interaction of the ACTN2 dimer with actin in the Z-disk, takes the user to the corresponding state summary page with literature citations.

65 The transitions producing each state are listed on the transitions page. Each entry is likewise linked to a graphic display of the modification or association. Once again, the detailed references and linked to PubMed accession numbers. As overwhelming detailed as these pages seem, they are best described visually by the network map page.

Figure 3.2.4 Transition page for the association of ADAM-12 with ACTN2.

66 Alpha-actinin-3

Transition network Version 1.0, Peer Reviewed And Published 24 Jan 2006 doi:10.1038/mp.a000196.01 How to cite this Molecule Page

Network Map

Click on ovals to see state details and on asterisks to see transition details.

See SVG version

Figure 3.2.5 Network map for α-actinin 3. Ovals link to state data, while arrows link to transitions. The network map is an interactive diagram showing each protein state as an oval with transition arrows leading to and from each state in logical procession. Clicking on an oval takes the researcher to the protein state data, while clicking on each arrow points the researcher to the transition file data.

67 3.3 α-Actinin1 Molecule Pages

3.3.1 α-Actinin1 Overview

Alpha-actinin-1

Mini Molecule Page

AfCS Protein ID A000194

Protein Name Alpha-actinin-1

Protein Aac1; AACT; ACT1; Actinin, alpha 1; actinin, alpha 1; Actn1; alpha-Actinin 1; Alpha-actinin-1; Cytoskeletal Synonyms alpha-actinin; HuActNM; Non-muscle alpha-actinin; Nonmuscle alpha-actinin 1

Author Kenneth A Taylor [email protected]

Co-Authors Cheri M. Hampton

alpha-Actinin 1 is a nonmuscle isoform. It functions as an antiparallel homodimer to bind actin filaments at either end. These crosslinks anchor the actin cytoskeleton to the membrane and link the cytoskeleton to cytoplasmic signaling proteins. The alpha-actinins are members of the spectrin family of proteins. They have an amino-terminal actin-binding domain consisting of two calponin homology (CH)domains. The central rod is Protein Function composed of four spectrin-like triple-helix repeats that facilitate dimerization. The carboxy-terminus is a calmodulin-like domain. There are four EF-hand motifs, two of which (3 and 4) have lost their ability to bind Ca(2+) but still function in protein-protein interactions. It is in EF-hands 1 and 2 that variability between the isoforms exists. Nonmuscle (ACTN1 and ACTN4) isoforms can bind calcium, while muscle (ACTN2 and ACTN3)

Figure 3.3.1 α-Actinin 1 Overview 68 isoforms cannot.

alpha-Actinin-1 is regulated by the binding of calcium ions in the micromolar range to the EF-hand domains. Calcium concentrations >10-7 completely inhibit actin binding. Because alpha-actinin functions as an Protein antiparallel homodimer, it is believed that calcium binding at the carboxy-terminus of one monomer influences Regulation the ability of actin to bind at the neighboring amino-terminus. The affinity with which alpha-actinin binds to actin can also be regulated by tyrosine phosphorylation by focal adhesion kinase (FAK). FAK phosphorylates alpha-actinin on a tyrosine residue within the actin-binding domain.

Concentration - Regulation

Subcellular alpha-Actinin-1 is localized along stress fibers, microfilament bundles, and at focal adhesions. Localization

Overexpression of alpha-actinin results in a decrease in motility, perhaps due to an increase in plaque Phenotypes formation. Reduction of alpha-actinin expression leads to increased motility and tumor formation.

In chicken, a single ACTN1 gene gives rise to two isoforms by alternative splicing. The difference lies in the Splice Variants second half of the first EF-hand domain. ACTN1a is found in chicken smooth muscle, while ACTN1b is cytoskeletal and calcium-sensitive like human/mouse ACTN1.

Mouse Gene Actn1 Symbol

Genbank 15170951 Accession

Genbank Mouse Organism

Figure 3.3.1 Continued. 69 Major Sites of brain, T lymphocytes, Platelets, Placenta Expression

Cardiac Myocyte yes (Velez C,et al (1995) Basic fibroblast and platelet-derived growth factors as modulators of actin and Expression alpha-actinin in chick myocardiocytes during development. Proc Soc Exp Biol Med 210, 57-63.)

B Lymphocyte no (-) Expression

Interactions The original role for alpha-actinin is to cross-link F-actin. Alpha-actinin binds to F-actin via its amino-terminal actin binding domain between two adjacent monomers on the long-pitch two-start actin helix. The actin binding domain at the other end of the dimer binds a neighboring filament in either parallel or antiparallel fashion. The carboxyterminus of the transmembrane metalloprotease ADAM12 interacts with the aminoterminus of ACTN1. The adhesion plaque protein vinculin has been demonstrated to bind via its head region to EF-hand 1 in actinin. Another report gives binding to the 4th 3-helix bundle. Both of these domains are in proximity to one another. The human autogen BP180 binds ACTN1 carboxyterminal domain containing EF-hands 3 and 4 (EF34) in the hemidesmosome. The intercellular adhesion proteins mediating leukocyte binding to beta-2-integrins, ICAM1 and 2, have been shown to interact with actinin. alpha-Actinin has also be demonstrated to link the cytoskeleton to the cytoplasmic tails of beta-1 and beta-2 integrin. The phospholipid PIP2 binds to the actin binding domain. Its binding strongly influences further actin crosslinking by alpha-actinin. PI3 kinase and PIP3 also bind to alpha-actinin and influence its binding to integrin, thereby restructuring the focal adhesion plaque. CLP36 is a member of the PDZ-LIM family of proteins. It is almost identical to hCLIM1 (recently renamed Elfin). It binds to actinin spectrin-like repeats 2 & 3 via its PDZ domain, leaving the LIM domain to bind cytoplasmic

Figure 3.3.1 Continued. 70 cell signaling proteins. The serine/threonine protein kinase, PKN, binds to the third spectrin-like repeat of actinin as well as to the EF- hand domain. This binding is calcium sensitive in nonmuscle actinin. PKN is activated by fatty acids and Rho, and can phosphorylate actinin. Binding of PIP2 increases the association of PKN with actinin.

Antibodies Mouse monoclonal antibody to chicken gizzard actinin, clone JLN20 (ICN, InnoGenex, BioGenex, Oncogene Research Products). Bovine mAb to mouse actinin, clone BM-75.2 (Sigma, ICN). Mouse mAb to human actinin, clone AT 6/172 (Research Diagnostics Inc, Immune Systems Ltd). Mouse mAb to quail smooth muscle actinin, 1E12 (Developmental Studies Hybridoma Bank at the University of Iowa). Mouse mAb to chicken gizzard, clone CB11 (ICN, Affiniti). Rabbit polyclonal antibody to chicken gizzard (ICN). Mouse mAb to human red blood cell ghosts, clone RBC2/1B6 (Novocastra). mAb to smooth muscle actinin, clone 1A4 (Diagnostic BioSystems). Goat polyclonal antibody to aminoterminus of human ACTN1, N-19 (Santa Cruz Biotechnology). Goat polyclonal antibody to human ACTN1 carboxyterminus, C-20 (Santa Cruz Biotechnology).

Figure 3.3.1 Continued.

71 Broderick, M.J.F. & Winder, S.J. (2002) Towards a complete atomic structure of spectrin family proteins. J. Struct. Biol. 137, 184-193. Baron MD, Davison MD, Jones P, Critchley DR. (1987). The structure and function of alpha-actinin. Biochem Soc Trans 15, 796-8. Youssoufian H, McAfee M, Kwiatkowski DJ. (1990). Cloning and chromosomal localization of the human cytoskeletal alpha-actinin gene reveals linkage to the beat-spectrin gene. Am J Hum Genet 47, 62-71. References Parr T, Waites GT, Patel B, Millake DB, Critchley DR. (1992). A chick skeletal-muscle alpha-actinin gene gives rise to two alternatively spliced isoforms which differ in the EF-hand Ca(2+)-binding domain. Eur J Biochem 210, 801-9. Waites GT, Graham IR, Jackson P, Millake DB, Patel B, Blanchard AD, Weller PA, Eperon IC, Critchley DR.(1992). Mutually exclusive splicing of calcium-binding domain exons in chick alpha-actinin. J Biol Chem 267, 6263-71.

Figure 3.3.1 Continued.

72 3.3.2 α-Actinin1 Mini Molecule Page

Alpha-actinin-1

Mini Molecule Page

AfCS Protein ID A000194

Protein Name Alpha-actinin-1

Author Kenneth A Taylor [email protected]

Co-Authors Cheri M. Hampton

Protein alpha-Actinin-1 is regulated by the binding of calcium ions in the micromolar range to the EF-hand domains. Regulation Calcium concentrations >10-7 completely inhibit actin binding. Because alpha-actinin functions as an

Figure 3.3.2 α-Actinin 1 Mini Molecule Page. 73 antiparallel homodimer, it is believed that calcium binding at the carboxy-terminus of one monomer influences the ability of actin to bind at the neighboring amino-terminus. The affinity with which alpha-actinin binds to actin can also be regulated by tyrosine phosphorylation by focal adhesion kinase (FAK). FAK phosphorylates alpha-actinin on a tyrosine residue within the actin-binding domain.

Concentration - Regulation

Subcellular alpha-Actinin-1 is localized along stress fibers, microfilament bundles, and at focal adhesions. Localization

Mouse Gene Actn1 Symbol

Genbank 15170951 Accession

Genbank Mouse Organism

Major Sites of brain, T lymphocytes, Platelets, Placenta Expression

Cardiac Myocyte yes (Velez C,et al (1995) Basic fibroblast and platelet-derived growth factors as modulators of actin and

Figure 3.3.2 Continued. 74 Expression alpha-actinin in chick myocardiocytes during development. Proc Soc Exp Biol Med 210, 57-63.)

B Lymphocyte no (-) Expression

Figure 3.3.2 Continued. 75 and can phosphorylate actinin. Binding of PIP2 increases the association of PKN with actinin.

Mouse monoclonal antibody to chicken gizzard actinin, clone JLN20 (ICN, InnoGenex, BioGenex, Oncogene Research Products). Bovine mAb to mouse actinin, clone BM-75.2 (Sigma, ICN). Mouse mAb to human actinin, clone AT 6/172 (Research Diagnostics Inc, Immune Systems Ltd). Mouse mAb to quail smooth muscle actinin, 1E12 (Developmental Studies Hybridoma Bank at the University of Iowa). Mouse mAb to chicken gizzard, clone Antibodies CB11 (ICN, Affiniti). Rabbit polyclonal antibody to chicken gizzard (ICN). Mouse mAb to human red blood cell ghosts, clone RBC2/1B6 (Novocastra). mAb to smooth muscle actinin, clone 1A4 (Diagnostic BioSystems). Goat polyclonal antibody to aminoterminus of human ACTN1, N-19 (Santa Cruz Biotechnology). Goat polyclonal antibody to human ACTN1 carboxyterminus, C-20 (Santa Cruz Biotechnology).

Broderick, M.J.F. & Winder, S.J. (2002) Towards a complete atomic structure of spectrin family proteins. J. Struct. Biol. 137, 184-193. Baron MD, Davison MD, Jones P, Critchley DR. (1987). The structure and function of alpha-actinin. Biochem Soc Trans 15, 796-8. Youssoufian H, McAfee M, Kwiatkowski DJ. (1990). Cloning and chromosomal localization of the human cytoskeletal alpha-actinin gene reveals linkage to the beat-spectrin gene. Am J Hum Genet 47, 62-71. References Parr T, Waites GT, Patel B, Millake DB, Critchley DR. (1992). A chick skeletal-muscle alpha-actinin gene gives rise to two alternatively spliced isoforms which differ in the EF-hand Ca(2+)-binding domain. Eur J Biochem 210, 801-9. Waites GT, Graham IR, Jackson P, Millake DB, Patel B, Blanchard AD, Weller PA, Eperon IC, Critchley DR.(1992). Mutually exclusive splicing of calcium-binding domain exons in chick alpha-actinin. J Biol Chem 267, 6263-71.

Figure 3.3.2 Continued.

76 3.4 α-Actinin2 Molecule Pages

3.4.1 α-Actinin2 Overview

Alpha-actinin-2

Molecule Page Overview Version 1.0, Peer Reviewed And Published 24 Jan 2006

doi:10.1038/mp.a000195.01 How to cite this Molecule Page

AfCS ID A000195

AfCS Name Alpha-actinin-2

Actinin alpha 2; Actinin, alpha 2; Actn2; Alpha-actinin 2; Alpha-actinin-2; HuActSK1; Sarcomeric actinin; Skeletal and All Names cardiac muscle alpha-actinin

Functional Category Cytoskeletal protein

Primary Symbol Actn2

Molecule Page Version 1.0, Peer Reviewed And Published 24 Jan 2006 Version

Corresponding Kenneth A Taylor [email protected] Author

Authors Cheri M Hampton, Kenneth A Taylor

Editorial Board Patrick J Casey [email protected] (editorial board member list) Member

Species Mouse

Figure 3.4.1 α-Actinin 2 Overview Page.

77 1 MNQIEPGVQY NYVYDEDEYM IQEEEWDRDL LLDPAWEKQQ RKTFTAWCNS 51 HLRKAGTQIE NIEEDFRNGL KLMLLLEVIS GERLPKPDRG KMRFHKIANV 101 NKALDYIASK GVKLVSIGAE EIVDGNVKMT LGMIWTIILR FAIQDISVEE 151 TSAKEGLLLW CQRKTAPYRN VNIQNFHTSW KDGLGLCALI HRHRPDLIDY 201 SKLNKDDPIG NINLAMEIAE KHLDIPKMLD AEDIVNTPKP DERAIMTYVS 251 CFYHAFAGAE QAETAANRIC KGLAVNQENE RLMEEYERLA SELLEWIRRT 301 IPWLENRTPE KTMQAMQKKL EDFRDYRRKH KPPKVQEKCQ LEINFNTLQT 351 KLRISNRAAF MPSEGKMVSD IAGACQRLEQ AEKGYEEWLL NEIRRLERLE 401 HLAEKFRQKA STHETWAYGK EQILLQKDYE SASLTEVRAL LRKHEAFESD Sequence 451 LAAHQDRVEQ IAAIAQELNE LDYHDAVNVN DRCQKICDQW DRLGTLTQKR 501 REALERTEKL LETIDQLHLE FAKRAAPFNN WMEGAMEDLQ DMFIVHSIEE 551 IQSLITAHEQ FKATLPEADG ERQSILAIQN EVEKVIQSYS IRISSSNPYS 601 TVTMDELRNK WDKVKQLVPV RDQSLQEELA RQHANERLRR QFAAQANAIG 651 PWIQNKMEEI ARSSIQITGA LEDQMNQLKQ YEHNIINYKN NIDKLEGDHQ 701 LIQEGLVFDN KHTNYTMEHI RVGWELLLTT IGRTINEVET QILTRDAKGI 751 TQKQMNEFRA SFNHFDRRKN GLMDHEDFRA CLISMGYDLG EAEFARIMTL 801 VDPNGQGTVT FQSFIDFMTR ETADTDTAEQ VIASFRILAS DKPYILAEEL 851 RRELPPDQAQ YCIKRMPPYS GPGSVPGALD YTAFSSALYG ESDL

Molecular Weight 103658

Isoelectric Point 5.36

Extinction Coefficient 119950 M-1cm-1

Absorption Coefficient 1.16

Aliphatic Index 84.19

Figure 3.4.1 Continued

78 3.4.2 α-Actinin2 Summary

Alpha-actinin-2

Summary Version 1.0, Peer Reviewed And Published 24 Jan 2006

doi:10.1038/mp.a000195.01 How to cite this Molecule Page

Protein Function

Pub PM ID Authors Title Journal Date

1339456 Beggs AH, Byers TJ, Knoll JH, Cloning and characterization of two human skeletal muscle J Biol Chem, 5 May Boyce FM, Bruns GA, Kunkel LM alpha-actinin genes located on chromosomes 1 and 11. 267, 13 1992

Figure 3.4.2 α-Actinin 2 Summary Page. 79 Dixson JD, Forstner MJ, Garcia Jan 12569417 The alpha-actinin gene family: a revised classification. J Mol Evol, 56, 1 DM 2003

Djinovic-Carugo K, Gautel M, The spectrin repeat: a structural platform for cytoskeletal 20 Feb 11911890 FEBS Lett, 513, 1 Ylänne J, Young P protein assemblies. 2002

Mills M, Yang N, Weinberger R, Differential expression of the actin-binding proteins, alpha- Hum Mol Genet, 15 Jun 11440986 Vander Woude DL, Beggs AH, actinin-2 and -3, in different species: implications for the 10, 13 2001 Easteal S, North K evolution of functional redundancy.

Cell Motil Jun 15083532 Otey CA, Carpen O Alpha-actinin revisited: a fresh look at an old player. Cytoskeleton, 58, 2 2004

Tiso N, Majetti M, Stanchi F, Fine mapping and genomic structure of ACTN2, the human Biochem Biophys Nov 10548523 Rampazzo A, Zimbello R, Nava A, gene coding for the sarcomeric isoform of alpha-actinin-2, Res Commun, 265, 1 1999 Danieli GA expressed in skeletal and cardiac muscle.

Jun 15014165 Virel A, Backman L Molecular evolution and structure of alpha-actinin. Mol Biol Evol, 21, 6 2004

Regulation of Activity

Unlike Actn1 and Actn4, the EF-hands of Actn2 are incapable of binding Ca2+ and are thus calcium-insensitive (Beggs et al. 1992). Its binding is largely constitutive but can be modulated by competitive binding of other proteins. It has been demonstrated that α-actinin dynamics can be regulated by the binding of PtdIns(4,5)P2 to CH2 of the actin-binding domain (Fukami et al. 1992; Fukami et al. 1996; Fraley et al. 2003). PtdIns(4,5)P2 modulates binding to actin, increases the affinity of Actn2 for titin (Young and Gautel. 2000), and inhibits Actn2 binding to CapZ (Papa et al. 1999). In neurons, Actn2 bundling activity has been shown to be enhanced by interaction with rabphillin-3A (Kato et al. 1996) and Actn2 is potentially regulated by phosphorylation at the N-terminus by PKN, which would potentially interfere with actin binding (Mukai et al. 1997).

Figure 3.4.2 Continued. 80 Pub PM ID Authors Title Journal Date

Cloning and characterization of two human skeletal Beggs AH, Byers TJ, Knoll JH, Boyce J Biol Chem, 5 May 1339456 muscle alpha-actinin genes located on chromosomes 1 and FM, Bruns GA, Kunkel LM 267, 13 1992 11.

Corgan AM, Singleton C, Santoso CB, Phosphoinositides differentially regulate alpha-actinin Biochem J, 15 Mar 14670080 Greenwood JA flexibility and function. 378, Pt 3 2004

Phosphoinositide binding regulates alpha-actinin Fraley TS, Pereira CB, Tran TC, J Biol Chem, 15 Apr 15710624 dynamics: mechanism for modulating cytoskeletal Singleton C, Greenwood JA 280, 15 2005 remodeling.

Fraley TS, Tran TC, Corgan AM, Nash Phosphoinositide binding inhibits alpha-actinin bundling J Biol Chem, 27 Jun 12716899 CA, Hao J, Critchley DR, Greenwood JA activity. 278, 26 2003

Fukami K, Furuhashi K, Inagaki M, Requirement of phosphatidylinositol 4,5-bisphosphate Nature, 10 Sep 1326084 Endo T, Hatano S, Takenawa T for alpha-actinin function. 359, 6391 1992

Identification of a phosphatidylinositol 4,5- Fukami K, Sawada N, Endo T, J Biol Chem, 2 Feb 8576235 bisphosphate-binding site in chicken skeletal muscle alpha- Takenawa T 271, 5 1996 actinin.

8943213 Kato M, Sasaki T, Ohya T, Nakanishi H, Physical and functional interaction of rabphilin-3A with J Biol Chem, 13 Dec Nishioka H, Imamura M, Takai Y alpha-actinin. 271, 50 1996

Figure 3.4.2 Continued.

81 Mukai H, Toshimori M, Shibata H, J Biol Chem, 21 Feb 9030526 Takanaga H, Kitagawa M, Miyahara M, Interaction of PKN with alpha-actinin. 272, 8 1997 Shimakawa M, Ono Y

Papa I, Astier C, Kwiatek O, Raynaud F, Alpha actinin-CapZ, an anchoring complex for thin J Muscle Res Feb 10412090 Bonnal C, Lebart MC, Roustan C, filaments in Z-line. Cell Motil, 20, 2 1999 Benyamin Y

The interaction of titin and alpha-actinin is controlled by 1 Dec 11101506 Young P, Gautel M a phospholipid-regulated intramolecular pseudoligand EMBO J, 19, 23 2000 mechanism.

Interactions with Ligands and Other Proteins

Aside from binding to F-actin via its actin-binding domain, Actn2 associates with many cellular architecture proteins. At the sarcolemma, Actn2 interacts with affixin (Yamaji et al. 2004), dystrophin (Hance et al. 1999), phospholipase D2 (Park et al. 2000), as well as L-type calcium channels (Sadeghi et al. 2002) and the Kv1.4 and 1.5 potassium channels (Maruoka et al. 2000; Cukovic et al. 2001; Mason et al. 2002). In the z-disk Actn2 interacts with a host of proteins: ADAM12 (Galliano et al. 2000), CapZ (Papa et al. 1999), ENH (Nakagawa et al. 2000; Niederlander et al. 2004), ZASP (Faulkner et al. 1999; Au et al. 2004), calsarcins (Frey et al. 2000; Frey and Olsen. 2002), myopalladin (Bang et al. 2001), elfin (Kotaka et al. 2000), ALP and MLP (Henderson et al. 2003; Klaavuniemi et al. 2004; Mohapatra et al. 2003; Geier et al. 2003; Gehmlich et al. 2004), myotillin (Salmikangas et al. 1999; Salmikangas et al. 2003), FBPase/amorphin complex (Gizak et al. 2003; Rakus et al. 2003; Mamczur et al. 2005), and titin (Sorimachi et al. 1997; Young et al. 1998; Satoh et al. 1999; Atkinson et al. 2000; Atkinson et al. 2000; Luther. 2000;Young and Gautel. 2000; Atkinson et al. 2001; Joseph et al. 2001). In neurons, Actn2 anchors and/or regulates the activity of adenosine A2A (Burgueno et al. 2003) and NMDA receptors (Wyszynski et al. 1997; Wyszynski et al. 1998; Cattabeni et al. 1999; Krupp et al. 1999; Anders et al. 2000; Dunah et al. 2000; Ratzliff and Soltesz. 2001; Leonard et al. 2002; Rycroft and Gibb. 2004) and interacts with and/or is regulated by Rabphillin3A (Kato et al. 1996) and PKN (Mukai et al. 1997). Actn2 has also been reported to interact with GRIP1, glucocorticoid receptor interacting protein 1, in the nucleus (Huang et al. 2004).

PM ID Authors Title Journal Pub Date

Figure 3.4.2 Continued. 82 Reduced ethanol inhibition of N-methyl-D- 19 Anders DL, Blevins T, Smothers CT, Woodward aspartate receptors by deletion of the NR1 C0 J Biol Chem, 10809744 May JJ domain or overexpression of alpha-actinin-2 275, 20 2000 proteins.

Atkinson RA, Joseph C, Dal Piaz F, Birolo L, Stier Binding of alpha-actinin to titin: implications for Biochemistry, 9 May 10819994 G, Pucci P, Pastore A Z-disk assembly. 39, 18 2000

Ca2+-independent binding of an EF-hand Atkinson RA, Joseph C, Kelly G, Muskett FW, Nat Struct Biol, Oct 11573089 domain to a novel motif in the alpha-actinin-titin Frenkiel TA, Nietlispach D, Pastore A 8, 10 2001 complex.

Assignment of the 1H, 13C and 15N resonances Atkinson RA, Joseph C, Kelly G, Muskett FW, J Biomol NMR, Mar 10805138 of the C-terminal EF-hands of alpha-actinin in a 14 Frenkiel TA, Pastor A 16, 3 2000 kDa complex with Z-repeat 7 of titin.

Au Y, Atkinson RA, Guerrini R, Kelly G, Joseph C, Solution structure of ZASP PDZ domain; Apr 15062084 Martin SR, Muskett FW, Pallavicini A, Faulkner G, implications for sarcomere ultrastructure and Structure, 12, 4 2004 Pastore A enigma family redundancy.

Bang ML, Mudry RE, McElhinny AS, Trombitás K, Myopalladin, a novel 145-kilodalton sarcomeric 16 Apr 11309420 Geach AJ, Yamasaki R, Sorimachi H, Granzier H, protein with multiple roles in Z-disc and I-band J Cell Biol, 153, 2 2001 Gregorio CC, Labeit S protein assemblies.

12837758 Burgueño J, Blake DJ, Benson MA, Tinsley CL, The adenosine A2A receptor interacts with the J Biol Chem, 26 Esapa CT, Canela EI, Penela P, Mallol J, Mayor F, actin-binding protein alpha-actinin. 278, 39 Sep 2003 Lluis C, Franco R, Ciruela F

Figure 3.4.2 Continued.

83 Pathophysiological implications of the Eur J Pharmacol, 30 Jun 10443587 Cattabeni F, Gardoni F, Di Luca M structural organization of the excitatory synapse. 375, 1-3 1999

A discrete amino terminal domain of Kv1.5 and 1 Jun 11389904 Cukovic D, Lu GW, Wible B, Steele DF, Fedida D Kv1.4 potassium channels interacts with the FEBS Lett, 498, 1 2001 spectrin repeats of alpha-actinin-2.

alpha-actinin-2 in rat striatum: localization and Dunah AW, Wyszynski M, Martin DM, Sheng M, Brain Res Mol 23 Jun 10925145 interaction with NMDA glutamate receptor Standaert DG Brain Res, 79, 1-2 2000 subunits.

10427098 Faulkner G, Pallavicini A, Formentin E, Comelli A, Ievolella C, Trevisan S, Bortoletto G, ZASP: a new Z-band alternatively spliced PDZ- 26 Jul J Cell Biol, 146, 2 Scannapieco P, Salamon M, Mouly V, Valle G, motif protein. 1999 Lanfranchi G

Calsarcin-3, a novel skeletal muscle-specific J Biol Chem, 19 Apr 11842093 Frey N, Olson EN member of the calsarcin family, interacts with 277, 16 2002 multiple Z-disc proteins.

Calsarcins, a novel family of sarcomeric Proc Natl Acad Sci 19 11114196 Frey N, Richardson JA, Olson EN calcineurin-binding proteins. U S A, 97, 26 Dec 2000

10788519 Binding of ADAM12, a marker of skeletal muscle Galliano MF, Huet C, Frygelius J, Polgren A, regeneration, to the muscle-specific actin-binding J Biol Chem, 5 May Wewer UM, Engvall E protein, alpha -actinin-2, is required for myoblast 275, 18 2000 fusion.

Figure 3.4.2 Continued.

84 Decreased interactions of mutant muscle LIM Gehmlich K, Geier C, Osterziel KJ, Van der Ven Cell Tissue Res, Aug 15205937 protein (MLP) with N-RAP and alpha-actinin and PF, Fürst DO 317, 2 2004 their implication for hypertrophic cardiomyopathy.

Geier C, Perrot A, Ozcelik C, Binner P, Counsell Mutations in the human muscle LIM protein D, Hoffmann K, Pilz B, Martiniak Y, Gehmlich K, van Circulation, 18 12642359 gene in families with hypertrophic der Ven PF, Fürst DO, Vornwald A, von Hodenberg 107, 10 Mar 2003 cardiomyopathy. E, Nürnberg P, Scheffold T, Dietz R, Osterziel KJ

Immunohistochemical localization of human Histol Histopathol, Jan 12507293 Gizak A, Rakus D, Dzugaj A fructose-1,6-bisphosphatase in subcellular 18, 1 2003 structures of myocytes.

15 Hance JE, Fu SY, Watkins SC, Beggs AH, alpha-actinin-2 is a new component of the Arch Biochem 10328815 May Michalak M dystrophin-glycoprotein complex. Biophys, 365, 2 1999

ALP and MLP distribution during Cell Motil Mar 12589684 Henderson JR, Pomiès P, Auffray C, Beckerle MC myofibrillogenesis in cultured cardiomyocytes. Cytoskeleton, 54, 3 2003

The enhancement of nuclear receptor Huang SM, Huang CJ, Wang WM, Kang JC, Hsu J Mol Endocrinol, Apr 15072553 transcriptional activation by a mouse actin-binding WC 32, 2 2004 protein, alpha actinin 2.

11305911 Joseph C, Stier G, O’Brien R, Politou AS, A structural characterization of the interactions Biochemistry, 24 Apr Atkinson RA, Bianco A, Ladbury JE, Martin SR, between titin Z-repeats and the alpha-actinin C- 40, 16 2001 Pastore A terminal domain.

Figure 3.4.2 Continued.

85 Kato M, Sasaki T, Ohya T, Nakanishi H, Nishioka Physical and functional interaction of rabphilin- J Biol Chem, 13 8943213 H, Imamura M, Takai Y 3A with alpha-actinin. 271, 50 Dec 1996

The ZASP-like motif in actinin-associated LIM J Biol Chem, 18 Jun 15084604 Klaavuniemi T, Kelloniemi A, Ylänne J protein is required for interaction with the alpha- 279, 25 2004 actinin rod and for targeting to the muscle Z-line.

10861853 Kotaka M, Kostin S, Ngai S, Chan K, Lau Y, Lee Interaction of hCLIM1, an enigma family J Cell Biochem, 12 Jun SM, Li H, Ng EK, Schaper J, Tsui SK, Fung K, Lee C, protein, with alpha-actinin 2. 78, 4 2000 Waye MM

Interactions of calmodulin and alpha-actinin Krupp JJ, Vissel B, Thomas CG, Heinemann SF, 15 Feb 9952395 with the NR1 subunit modulate Ca2+-dependent J Neurosci, 19, 4 Westbrook GL 1999 inactivation of NMDA receptors.

Regulation of calcium/calmodulin-dependent Leonard AS, Bayer KU, Merrill MA, Lim IA, Shea protein kinase II docking to N-methyl-D-aspartate J Biol Chem, 13 12379661 MA, Schulman H, Hell JW receptors by calcium/calmodulin and alpha- 277, 50 Dec 2002 actinin.

Three-dimensional structure of a vertebrate J Struct Biol, Feb 10675292 Luther PK muscle Z-band: implications for titin and alpha- 129, 1 2000 actinin binding.

15757649 The effect of calcium ions on subcellular 14 Mamczur P, Rakus D, Gizak A, Dus D, Dzugaj A localization of aldolase-FBPase complex in skeletal FEBS Lett, 579, 7 Mar 2005 muscle.

Figure 3.4.2 Continued. 86 alpha-actinin-2 couples to cardiac Kv1.5 12 Maruoka ND, Steele DF, Au BP, Dan P, Zhang X, 10812072 channels, regulating current density and channel FEBS Lett, 473, 2 May Moore ED, Fedida D localization in HEK cells. 2000

Modulation of Kv1.5 currents by protein kinase Mason HS, Latten MJ, Godoy LD, Horowitz B, Mol Pharmacol, Feb 11809852 A, tyrosine kinase, and protein tyrosine Kenyon JL 61, 2 2002 phosphatase requires an intact cytoskeleton.

Mohapatra B, Jimenez S, Lin JH, Bowles KR, Mutations in the muscle LIM protein and alpha- Coveler KJ, Marx JG, Chrisco MA, Murphy RT, Lurie Mol Genet Metab, 2003 14567970 actinin-2 genes in dilated cardiomyopathy and PR, Schwartz RJ, Elliott PM, Vatta M, McKenna W, 80, 1-2 Sep-Oct endocardial fibroelastosis. Towbin JA, Bowles NE

Mukai H, Toshimori M, Shibata H, Takanaga H, J Biol Chem, 21 Feb 9030526 Interaction of PKN with alpha-actinin. Kitagawa M, Miyahara M, Shimakawa M, Ono Y 272, 8 1997

ENH, containing PDZ and LIM domains, Nakagawa N, Hoshijima M, Oyasu M, Saito N, heart/skeletal muscle-specific protein, associates Biochem Biophys 7 Jun 10833443 Tanizawa K, Kuroda S with cytoskeletal proteins through the PDZ Res Commun, 272, 2 2000 domain.

Characterization of a new human isoform of the Biochem Biophys 24 15555569 Niederländer N, Fayein NA, Auffray C, Pomiès P enigma homolog family specifically expressed in Res Commun, 325, 4 Dec 2004 skeletal muscle.

10412090 Papa I, Astier C, Kwiatek O, Raynaud F, Bonnal Alpha actinin-CapZ, an anchoring complex for J Muscle Res Cell Feb C, Lebart MC, Roustan C, Benyamin Y thin filaments in Z-line. Motil, 20, 2 1999

Figure 3.4.2 Continued. 87 Cardiac phospholipase D2 localizes to Park JB, Kim JH, Kim Y, Ha SH, Yoo JS, Du G, sarcolemmal membranes and is inhibited by alpha- J Biol Chem, 14 Jul 10801846 Frohman MA, Suh PG, Ryu SH actinin in an ADP-ribosylation factor-reversible 275, 28 2000 manner.

Colocalization of muscle FBPase and muscle Biochem Biophys 14 14592412 Rakus D, Mamczur P, Gizak A, Dus D, Dzugaj A aldolase on both sides of the Z-line. Res Commun, 311, 2 Nov 2003

Differential immunoreactivity for alpha-actinin- Neuroscience, 11246149 Ratzliff AD, Soltesz I 2, an N-methyl-D-aspartate-receptor/actin binding 2001 103, 2 protein, in hippocampal interneurons.

Regulation of single NMDA receptor channel J Physiol, 557, Pt 15 Jun 15073274 Rycroft BK, Gibb AJ activity by alpha-actinin and calmodulin in rat 3 2004 hippocampal granule cells.

Regulation of the cardiac L-type Ca2+ channel Am J Physiol Cell Jun 11997265 Sadeghi A, Doyle AD, Johnson BD by the actin-binding proteins alpha-actinin and Physiol, 282, 6 2002 dystrophin.

Myotilin, a novel sarcomeric protein with two Salmikangas P, Mykkänen OM, Grönholm M, Hum Mol Genet, Jul 10369880 Ig-like domains, is encoded by a candidate gene Heiska L, Kere J, Carpén O 8, 7 1999 for limb-girdle muscular dystrophy.

12499399 Salmikangas P, van der Ven PF, Lalowski M, Myotilin, the limb-girdle muscular dystrophy 1A Hum Mol Genet, 15 Jan Taivainen A, Zhao F, Suila H, Schröder R, (LGMD1A) protein, cross-links actin filaments and 12, 2 2003 Lappalainen P, Fürst DO, Carpén O controls sarcomere assembly.

Figure 3.4.2 Continued. 88 Structural analysis of the titin gene in Satoh M, Takahashi M, Sakamoto T, Hiroe M, Biochem Biophys 27 10462489 hypertrophic cardiomyopathy: identification of a Marumo F, Kimura A Res Commun, 262, 2 Aug 1999 novel disease gene.

Sorimachi H, Freiburg A, Kolmerer B, Ishiura S, Tissue-specific expression and alpha-actinin 1 Aug 9245597 Stier G, Gregorio CC, Labeit D, Linke WA, Suzuki K, binding properties of the Z-disc titin: implications J Mol Biol, 270, 5 1997 Labeit S for the nature of vertebrate Z-discs.

Differential regional expression and Wyszynski M, Kharazia V, Shanghvi R, Rao A, ultrastructural localization of alpha-actinin-2, a 15 Feb 9454847 J Neurosci, 18, 4 Beggs AH, Craig AM, Weinberg R, Sheng M putative NMDA receptor-anchoring protein, in rat 1998 brain.

Wyszynski M, Lin J, Rao A, Nigh E, Beggs AH, Competitive binding of alpha-actinin and 30 Jan 9009191 Nature, 385, 6615 Craig AM, Sheng M calmodulin to the NMDA receptor. 1997

Yamaji S, Suzuki A, Kanamori H, Mishima W, Affixin interacts with alpha-actinin and 24 15159419 Yoshimi R, Takasaki H, Takabayashi M, Fujimaki K, mediates integrin signaling for reorganization of F- J Cell Biol, 165, 4 May Fujisawa S, Ohno S, Ishigatsubo Y actin induced by initial cell-substrate interaction. 2004

Molecular structure of the sarcomeric Z-disk: 16 9501083 Young P, Ferguson C, Bañuelos S, Gautel M two types of titin interactions lead to an EMBO J, 17, 6 Mar 1998 asymmetrical sorting of alpha-actinin.

11101506 The interaction of titin and alpha-actinin is 1 Dec Young P, Gautel M controlled by a phospholipid-regulated EMBO J, 19, 23 2000 intramolecular pseudoligand mechanism.

Figure 3.4.2 Continued.

89 Regulation of Concentration

No data available.

Subcellular Localization

α-actinin-2 is localized to the Z-disks of muscle cells (Mills et al. 2001; Sorimachi et al. 1997) and to postsynaptic densities and dendritic spines of neurons (Wyszynski et al. 1997).

Pub PM ID Authors Title Journal Date

Differential expression of the actin-binding proteins, alpha- Mills M, Yang N, Weinberger R, Vander Hum Mol 15 Jun 11440986 actinin-2 and -3, in different species: implications for the Woude DL, Beggs AH, Easteal S, North K Genet, 10, 13 2001 evolution of functional redundancy.

Sorimachi H, Freiburg A, Kolmerer B, Tissue-specific expression and alpha-actinin binding J Mol Biol, 1 Aug 9245597 Ishiura S, Stier G, Gregorio CC, Labeit D, properties of the Z-disc titin: implications for the nature of 270, 5 1997 Linke WA, Suzuki K, Labeit S vertebrate Z-discs.

Wyszynski M, Lin J, Rao A, Nigh E, Beggs Competitive binding of alpha-actinin and calmodulin to the Nature, 30 Jan 9009191 AH, Craig AM, Sheng M NMDA receptor. 385, 6615 1997

Major Sites of Expression

α-actinin-2 is expressed in all muscle fiber types (Mills et al. 2001; Sorimachi et al. 1997) and in the brain (Wyszynski et al. 1997).

Figure 3.4.2 Continued. 90 Pub PM ID Authors Title Journal Date

Wyszynski M, Lin J, Rao A, Nigh E, Beggs Competitive binding of alpha-actinin and calmodulin to the Nature, 30 Jan 9009191 AH, Craig AM, Sheng M NMDA receptor. 385, 6615 1997

Phenotypes

The genetic disease arrhythmogenic right ventricular cardiomyopathy type 2 is linked to a CA4 repeat in the last intron.

Pub PM ID Authors Title Journal Date

Beggs AH, Phillips HA, Kozman H, A (CA)n repeat polymorphism for the human skeletal muscle alpha- Genomics, Aug 1505962 Mulley JC, Wilton SD, Kunkel LM, Laing actinin gene ACTN2 and its localization on the linkage map of 13, 4 1992 NG chromosome 1.

Splice Variants

Figure 3.4.2 Continued. 91 No splice variants have been found in mammals.

Antibodies

Mouse monoclonal antibody (mAb) to rabbit skeletal α-actinin, clone EA-53 (Sigma, abcam).

Mouse mAb to chicken gizzard actinin, clone JLN20 (ICN, InnoGenex, BioGenex, Oncogene Research Products).

Bovine mAb to mouse actinin, clone BM-75.2 (Sigma, ICN).

Mouse mAb to human actinin, clone AT 6/172 (Research Diagnostics Inc, Immune Systems Ltd).

Mouse mAb to quail smooth muscle actinin, 1E12 (Developmental Studies Hybridoma Bank at the University of Iowa).

Mouse mAb to chicken gizzard, clone CB11 (ICN, Affiniti).

Rabbit polyclonal Ab to chicken gizzard (ICN).

Mouse mAb to human red blood cell ghosts, clone RBC2/1B6 (Novocastra).

Mouse mAb to smooth muscle actinin, clone 1A4 (Diagnostic BioSystems).

Goat polyclonal Ab to amino terminus of human ACTN1, N-19 (Santa Cruz Biotechnology).

Goat polyclonal Ab to human ACTN1 carboxy terminus, C-20 (Santa Cruz Biotechnology).

Figure 3.4.2 Continued.

92 3.4.3 α-Actinin2 Network Map

Alpha-actinin-2

Network Map

Figure 3.4.3 α-Actinin 2 Network Map

93 3.4.4 α-Actinin2 States

Alpha-actinin-2

State List Version 1.0, Peer Reviewed And Published 24 Jan 2006 doi:10.1038/mp.a000195.01 How to cite this Molecule Page

Functional states

State description State name Location Transition graph

Actn2 (Actn2)[2] Unknown

Actn2/Actn3 (Actn2) (Actn3) Unknown

2(Actn2) 2(Actn2) Unknown

2(Actn2)/actin(Z-disk) 2(Actn2) (Act[b,a1,a2,c1,g1,g2])[2] striated muscle thin filament

2(Actn2)/ADAM-12 2(Actn2) (Act[b,a1,a2,c1,g1,g2]) (ADAM12) striated muscle thin filament

2(Actn2)/CapZ 2(Actn2) (Act[b,a1,a2,c1,g1,g2]) (CAPZB) striated muscle thin filament

2(Actn2)/ENH 2(Actn2) (Act[b,a1,a2,c1,g1,g2]) (Enh) striated muscle thin filament

2(Actn2)/ZASP 2(Actn2) (Act[b,a1,a2,c1,g1,g2]) (LDB3) striated muscle thin filament

2(Actn2)/calsarcin 2(Actn2) (Act[b,a1,a2,c1,g1,g2]) (MYOZ[1-3]) striated muscle thin filament

2(Actn2)/myopalladin 2(Actn2) (Act[b,a1,a2,c1,g1,g2]) (MYPN) striated muscle thin filament

2(Actn2)/Elfin 2(Actn2) (Act[b,a1,a2,c1,g1,g2]) (PDLIM1) striated muscle thin filament

2(Actn2)/ALP 2(Actn2) (Act[b,a1,a2,c1,g1,g2]) (Pdlim3) striated muscle thin filament

2(Actn2)/myotilin 2(Actn2) (Act[b,a1,a2,c1,g1,g2]) (MYOT) striated muscle thin filament

2(Actn2)/MLP 2(Actn2) (Marcksl1) (Act[b,a1,a2,c1,g1,g2]) striated muscle thin filament

Figure 3.4.4 α-Actinin 2 States Page. 94 2(Actn2)/FBPase 2(Actn2) (Pfkfb2) (Act[b,a1,a2,c1,g1,g2]) striated muscle thin filament

2(Actn2)/amorphin 2(Actn2) (Pygm) (Act[b,a1,a2,c1,g1,g2]) striated muscle thin filament

2(Actn2)/Titin 2(Actn2) (Ttn) (Act[b,a1,a2,c1,g1,g2]) striated muscle thin filament

2(Actn2)/PI(4,5)P2 (Actn2) (Actn2 PIP2) (Act[b,a1,a2,c1,g1,g2]) striated muscle thin filament

2(Actn2)/actin(sarcolemma) 2(Actn2) (Act[b,a1,a2,c1,g1,g2])[1] sarcolemma

2(Actn2)/Kv1.5 2(Actn2) (Act[b,a1,a2,c1,g1,g2]) (KCNA5) sarcolemma

2(Actn2)/PLD2 2(Actn2) (Act[b,a1,a2,c1,g1,g2]) (PLD2) sarcolemma

2(Actn2)/CaChannel 2(Actn2) (Cacna1c) (Act[b,a1,a2,c1,g1,g2]) sarcolemma

2(Actn2)/dystrophin 2(Actn2) (Dmd) (Act[b,a1,a2,c1,g1,g2]) sarcolemma

2(Actn2)/actin(cytoskel) 2(Actn2) (Act[b,a1,a2,c1,g1,g2])[3] neuron projection

2(Actn2)/PKN 2(Actn2) (Act[b,a1,a2,c1,g1,g2]) (PKN1) neuron projection

2(Actn2)/Rabphilin-3A 2(Actn2) (Act[b,a1,a2,c1,g1,g2]) (Rph3a) neuron projection

2(Actn2)/A2AR 2(Actn2) (Adora2a) (Act[b,a1,a2,c1,g1,g2]) neuron projection

2(Actn2)/NMDAR 2(Actn2) (Grin1) (Grin2b) (Act[b,a1,a2,c1,g1,g2]) neuron projection

2(Actn2)/RIL 2(Actn2) (Pdlim4) (Act[b,a1,a2,c1,g1,g2]) neuron projection

2(Actn2)/actin(nuc) 2(Actn2) (Act[b,a1,a2,c1,g1,g2])[4] nucleus

2(Actn2)/GRIP1 2(Actn2) (Act[b,a1,a2,c1,g1,g2]) (Ncoa2) nucleus

Figure 3.4.4 Continued.

95 3.4.5 α-Actinin 2 Transitions

Alpha-actinin-2

State Transitions Version 1.0, Peer Reviewed And Published 24 Jan 2006 doi:10.1038/mp.a000195.01 How to cite this Molecule Page

Transitions

(Actn2)[2] --> (Actn2) (Actn3)

(Actn2)[2] --> 2(Actn2)

2(Actn2) --> 2(Actn2) (Act[b,a1,a2,c1,g1,g2])[2]

2(Actn2) --> 2(Actn2) (Act[b,a1,a2,c1,g1,g2])[1]

2(Actn2) --> 2(Actn2) (Act[b,a1,a2,c1,g1,g2])[3]

2(Actn2) --> 2(Actn2) (Act[b,a1,a2,c1,g1,g2])[4]

2(Actn2) (Act[b,a1,a2,c1,g1,g2])[2] --> 2(Actn2) (Act[b,a1,a2,c1,g1,g2]) (ADAM12)

2(Actn2) (Act[b,a1,a2,c1,g1,g2])[2] --> 2(Actn2) (Act[b,a1,a2,c1,g1,g2]) (CAPZB)

2(Actn2) (Act[b,a1,a2,c1,g1,g2])[2] --> 2(Actn2) (Act[b,a1,a2,c1,g1,g2]) (Enh)

2(Actn2) (Act[b,a1,a2,c1,g1,g2])[2] --> 2(Actn2) (Act[b,a1,a2,c1,g1,g2]) (LDB3)

2(Actn2) (Act[b,a1,a2,c1,g1,g2])[2] --> 2(Actn2) (Act[b,a1,a2,c1,g1,g2]) (MYOZ[1-3])

2(Actn2) (Act[b,a1,a2,c1,g1,g2])[2] --> 2(Actn2) (Act[b,a1,a2,c1,g1,g2]) (MYPN)

2(Actn2) (Act[b,a1,a2,c1,g1,g2])[2] --> 2(Actn2) (Act[b,a1,a2,c1,g1,g2]) (PDLIM1)

2(Actn2) (Act[b,a1,a2,c1,g1,g2])[2] --> 2(Actn2) (Act[b,a1,a2,c1,g1,g2]) (Pdlim3)

2(Actn2) (Act[b,a1,a2,c1,g1,g2])[2] --> 2(Actn2) (Act[b,a1,a2,c1,g1,g2]) (MYOT)

Figure 3.4.5 α-Actinin 2 Transitions Page 96 2(Actn2) (Act[b,a1,a2,c1,g1,g2])[2] --> 2(Actn2) (Marcksl1) (Act[b,a1,a2,c1,g1,g2])

2(Actn2) (Act[b,a1,a2,c1,g1,g2])[2] --> 2(Actn2) (Pfkfb2) (Act[b,a1,a2,c1,g1,g2])

2(Actn2) (Act[b,a1,a2,c1,g1,g2])[2] --> 2(Actn2) (Pygm) (Act[b,a1,a2,c1,g1,g2])

2(Actn2) (Act[b,a1,a2,c1,g1,g2])[2] --> 2(Actn2) (Ttn) (Act[b,a1,a2,c1,g1,g2])

2(Actn2) (Act[b,a1,a2,c1,g1,g2])[2] --> (Actn2) (Actn2 PIP2) (Act[b,a1,a2,c1,g1,g2])

2(Actn2) (Act[b,a1,a2,c1,g1,g2])[1] --> 2(Actn2) (Act[b,a1,a2,c1,g1,g2]) (KCNA5)

2(Actn2) (Act[b,a1,a2,c1,g1,g2])[1] --> 2(Actn2) (Act[b,a1,a2,c1,g1,g2]) (PLD2)

2(Actn2) (Act[b,a1,a2,c1,g1,g2])[1] --> 2(Actn2) (Cacna1c) (Act[b,a1,a2,c1,g1,g2])

2(Actn2) (Act[b,a1,a2,c1,g1,g2])[1] --> 2(Actn2) (Dmd) (Act[b,a1,a2,c1,g1,g2])

2(Actn2) (Act[b,a1,a2,c1,g1,g2])[3] --> 2(Actn2) (Act[b,a1,a2,c1,g1,g2]) (PKN1)

2(Actn2) (Act[b,a1,a2,c1,g1,g2])[3] --> 2(Actn2) (Act[b,a1,a2,c1,g1,g2]) (Rph3a)

2(Actn2) (Act[b,a1,a2,c1,g1,g2])[3] --> 2(Actn2) (Adora2a) (Act[b,a1,a2,c1,g1,g2])

2(Actn2) (Act[b,a1,a2,c1,g1,g2])[3] --> 2(Actn2) (Grin1) (Grin2b) (Act[b,a1,a2,c1,g1,g2])

2(Actn2) (Act[b,a1,a2,c1,g1,g2])[3] --> 2(Actn2) (Pdlim4) (Act[b,a1,a2,c1,g1,g2])

2(Actn2) (Act[b,a1,a2,c1,g1,g2])[4] --> 2(Actn2) (Act[b,a1,a2,c1,g1,g2]) (Ncoa2)

Figure 3.4.5 Continued.

97 3.5 α-Actinin 3 Molecule Pages

3.5.1 α-Actinin 3 Overview

Alpha-actinin-3

Molecule Page Overview Version 1.0, Peer Reviewed And Published 24 Jan 2006 doi:10.1038/mp.a000196.01 How to cite this Molecule Page

AfCS ID A000196

AfCS Name Alpha-actinin-3

All Names Acta-3; Actinin alpha 3; Actinin, alpha 3; Actn3; alpha Actinin 3; Alpha-actinin-3; ORF1

Functional Category Cytoskeletal protein

Primary Symbol Actn3

Molecule Page Version 1.0, Peer Reviewed And Published 24 Jan 2006 Version

Corresponding Author Kenneth A Taylor [email protected]

Authors Cheri M Hampton, Kenneth A Taylor

Editorial Board Patrick J Casey [email protected] (editorial board member list) Member

Species Mouse

Sequence 1 MMMVMQPEGL GAGEGPFSGG GGGEYMEQEE DWDRDLLLDP AWEKQQRKTF 51 TAWCNSHLRK AGTQIENIEE DFRNGLKLML LLEVISGERL PRPDKGKMRF 101 HKIANVNKAL DFIASKGVKL VSIGAEEIVD GNLKMTLGMI WTIILRFAIQ 151 DISVEETSAK EGLLLWCQRK TAPYRNVNVQ NFHTSWKDGL ALCALIHRHR

Figure 3.5.1 α-Actinin 3 Overview Page

98 201 PDLIDYAKLR KDDPIGNLNT AFEVAEKYLD IPKMLDAEDI VNTPKPDEKA 251 IMTYVSCFYH AFAGAEQAET AANRICKVLA VNQENEKLME EYEKLASELL 301 EWIRRTVPWL ENRVGEPSMS AMQRKLEDFR DYRRLHKPPR VQEKCQLEIN 351 FNTLQTKLRL SHRPAFMPSE GKLVSDIANA WRGLEQVEKG YEDWLLSEIR 401 RLQRLQHLAE KFQQKASLHE AWTRGKEEML NQHDYESASL QEVRALLRRH 451 EAFESDLAAH QDRVEHIAAL AQELNELDYH EAASVNSRCQ AICDQWDNLG 501 TLTQKRRDAL ERMEKLLETI DQLQLEFARR AAPFNNWLDG AIEDLQDVWL 551 VHSVEETQSL LTAHEQFKAT LPEADRERGA ILGIQGEIQK ICQTYGLRPK 601 SGNPYITLSS QDINNKWDTV RKLVPSRDQT LQEELARQQV NERLRRQFAA 651 QANAIGPWIQ GKVEEVGRLA AGLAGSLEEQ MAGLRQQEQN IINYKSNIDR 701 LEGDHQLLQE SLVFDNKHTV YSMEHIRVGW EQLLTSIART INEVENQVLT 751 RDAKGLSQEQ LNEFRASFNH FDRKRNGMME PDDFRACLIS MGYDLGEVEF 801 ARIMTMVDPN AAGVVTFQAF IDFMTRETAE TDTAEQVVAS FKILAGDKNY 851 ITPEELRREL PAEQAEYCIR RMAPYKGSGA PSGALDYVAF SSALYGESDL

Molecular Weight 103046

Isoelectric Point 5.31

Extinction Coefficient 124930 M-1cm-1

Absorption Coefficient 1.21

Aliphatic Index 84.50

Figure 3.5.1 Continued.

99 3.5.2 α-Actinin 3 Summary

Alpha-actinin-3

Summary Version 1.0, Peer Reviewed And Published 24 Jan 2006 doi:10.1038/mp.a000196.01 How to cite this Molecule Page

Protein Function

Based on high sequence identity to the other actinin isoforms and subcellular localization studies, the function of α-actinin-3 (Actn3) is to anchor actin filaments at the Z-disk. Its expression is limited to type 2B skeletal muscles fibers. It forms antiparallel homodimers or can form heterodimers with α-actinin-2 (Actn2). In human, the function of ACTN3 is overlapped by ACTN2, as evidenced by the lack of deleterious phenotype in ACTN3 null mutations. In mouse, however, these isoforms are not functionally redundant.

Pub PM ID Authors Title Journal Date

Beggs AH, Byers TJ, Knoll JH, Boyce Cloning and characterization of two human skeletal muscle 5 May 1339456 J Biol Chem, 267, 13 FM, Bruns GA, Kunkel LM alpha-actinin genes located on chromosomes 1 and 11. 1992

12569417 Dixson JD, Forstner MJ, Garcia DM The alpha-actinin gene family: a revised classification. J Mol Evol, 56, 1 Jan 2003

Cell Motil 15083532 Otey CA, Carpen O Alpha-actinin revisited: a fresh look at an old player. Jun 2004 Cytoskeleton, 58, 2

15014165 Virel A, Backman L Molecular evolution and structure of alpha-actinin. Mol Biol Evol, 21, 6 Jun 2004

Regulation of Activity

Protein activity is presumably regulated by competitive protein binding in the Z-disk.

Interactions with Ligands and Other Proteins

Based on sequence identity and subcellular localization, Actn3 binds actin via its N-terminal actin binding domain which is composed of two tandem calponin homology domains. Actn3 has been shown to associate with myozenin (FATZ, calsarcin) at Z-disks.

PM ID Authors Title Journal Pub

Figure 3.5.2 α-Actinin 3 Summary Page

100 Date

Beggs AH, Byers TJ, Knoll JH, Boyce FM, Bruns Cloning and characterization of two human skeletal muscle J Biol Chem, 5 May 1339456 GA, Kunkel LM alpha-actinin genes located on chromosomes 1 and 11. 267, 13 1992

Calsarcin-3, a novel skeletal muscle-specific member of the J Biol Chem, 19 Apr 11842093 Frey N, Olson EN calsarcin family, interacts with multiple Z-disc proteins. 277, 16 2002

Takada F, Vander Woude DL, Tong HQ, Myozenin: an alpha-actinin- and gamma-filamin-binding Proc Natl Acad Sci 13 Feb 11171996 Thompson TG, Watkins SC, Kunkel LM, Beggs AH protein of skeletal muscle Z lines. U S A, 98, 4 2001

Regulation of Concentration

No information is known about the regulation of concentration for Actn3.

Subcellular Localization

Localized to Z-disks of type 2B skeletal muscle.

Pub PM ID Authors Title Journal Date

Beggs AH, Byers TJ, Knoll JH, Boyce FM, Cloning and characterization of two human skeletal muscle alpha-actinin J Biol Chem, 5 May 1339456 Bruns GA, Kunkel LM genes located on chromosomes 1 and 11. 267, 13 1992

Differential expression of the actin-binding proteins, alpha-actinin-2 and - Mills M, Yang N, Weinberger R, Vander Hum Mol 15 Jun 11440986 3, in different species: implications for the evolution of functional Woude DL, Beggs AH, Easteal S, North K Genet, 10, 13 2001 redundancy.

Major Sites of Expression

Actn3 is only found in type 2B skeletal muscle (Beggs et al. 1992; Mills et al. 2001).

Pub PM ID Authors Title Journal Date

Mills M, Yang N, Weinberger R, Vander Differential expression of the actin-binding proteins, alpha-actinin-2 and - Hum Mol 15 Jun 11440986 Woude DL, Beggs AH, Easteal S, North K 3, in different species: implications for the evolution of functional Genet, 10, 13 2001

Figure 3.5.2 Continued.

101 redundancy.

Phenotypes

Approximately 18% of the human population carries a stop codon polymorphism (577X) resulting in a loss-of-function mutation that does not have a deleterious effect on phenotype. This is likely to be due to the functional redundancy of Actn2 which is also present in skeletal muscle sarcomeres, although this does not hold true in the mouse. Interestingly, the null mutation has been linked to a decrease in muscle power performance and increased endurance in olympic atheletes. More recent work has demonstrated that women with the 577X mutation have a reduced ability to produce high force strength, but develop dynamic strength in response to resistance training.

Pub PM ID Authors Title Journal Date

Clarkson PM, Devaney JM, Gordish-Dressman H, Thompson PD, ACTN3 genotype is associated with increases in J Appl Physiol, Jul 15718405 Hubal MJ, Urso M, Price TB, Angelopoulos TJ, Gordon PM, Moyna muscle strength in response to resistance training in 99, 1 2005 NM, Pescatello LS, Visich PS, Zoeller RF, Seip RL, Hoffman EP women.

A gene for speed? The evolution and function of Bioessays, Jul 15221860 MacArthur DG, North KN alpha-actinin-3. 26, 7 2004

Differential expression of the actin-binding proteins, Mills M, Yang N, Weinberger R, Vander Woude DL, Beggs AH, Hum Mol 15 Jun 11440986 alpha-actinin-2 and -3, in different species: implications Easteal S, North K Genet, 10, 13 2001 for the evolution of functional redundancy.

Splice Variants

No information is known about any splice variants for Actn3.

Antibodies

Mouse monoclonal antibody (mAb) to rabbit skeletal α-actinin, clone EA-53 (Sigma, abcam).

Mouse mAb to chicken gizzard actinin, clone JLN20 (ICN, InnoGenex, BioGenex, Oncogene Research Products).

Bovine mAb to mouse actinin, clone BM-75.2 (Sigma, ICN).

Mouse mAb to human actinin, clone AT 6/172 (Research Diagnostics Inc, Immune Systems Ltd).

Mouse mAb to quail smooth muscle actinin, 1E12 (Developmental Studies Hybridoma Bank at the University of Iowa).

Mouse mAb to chicken gizzard, clone CB11 (ICN, Affiniti).

Figure 3.5.2 Continued.

102 Rabbit polyclonal Ab to chicken gizzard (ICN).

Mouse mAb to human red blood cell ghosts, clone RBC2/1B6 (Novocastra).

Mouse mAb to smooth muscle actinin, clone 1A4 (Diagnostic BioSystems).

Goat polyclonal Ab to aminoterminus of human ACTN1, N-19 (Santa Cruz Biotechnology).

Goat polyclonal Ab to human ACTN1 carboxyterminus, C-20 (Santa Cruz Biotechnology).

Figure 3.5.2 Continued.

103 3.4.3 α-Actinin 3 Network Map

Alpha-actinin-3

Click on ovals to see state details and on asterisks to see transition details.

See SVG version

Figure 3.5.3 α-Actinin 3 Network Page

104 3.5.4 α-Actinin 3 States

Alpha-actinin-3

State List Version 1.0, Peer Reviewed And Published 24 Jan 2006 doi:10.1038/mp.a000196.01 How to cite this Molecule Page

Functional states

State State name Location Transition graph description

Actn3 (Actn3)[2] Unknown

Actn3/Actn2 (Actn2) (Actn3) striated muscle thin filament

2(Actn3) 2(Actn3) striated muscle thin filament

2(Actn3)/actin 2(Actn3) (Act[b,a1,a2,c1,g1,g2]) striated muscle thin filament

2(Actn3)/MYOZ1 2(Actn3) (Act[b,a1,a2,c1,g1,g2]) (MYOZ1) striated muscle thin filament

Figure 3.5.4 α-Actinin 3 States Page

105 3.5.5 α-Actinin 3 Transitions

Alpha-actinin-3

State Transitions Version 1.0, Peer Reviewed And Published 24 Jan 2006 doi:10.1038/mp.a000196.01 How to cite this Molecule Page

Transitions

(Actn3)[2] --> (Actn2) (Actn3)

(Actn3)[2] --> 2(Actn3)

2(Actn3) --> 2(Actn3) (Act[b,a1,a2,c1,g1,g2])

2(Actn3) --> 2(Actn3) (Act[b,a1,a2,c1,g1,g2]) (MYOZ1)

Figure 3.5.5 α-Actinin 3 Transitions Page

106 3.6 α-Actinin 4 Molecule Pages

3.6.1 α-Actinin 4 Overview

Alpha-actinin-4

Molecule Page Overview Version 1.0, Peer Reviewed And Published 24 Jan 2006 doi:10.1038/mp.a000197.01 How to cite this Molecule Page

AfCS ID A000197

AfCS Name Alpha-actinin-4

All Names Actinin alpha 4; Actinin, alpha 4; Actn4; Alpha-actinin-4

Functional Category Cytoskeletal protein

Primary Symbol Actn4

Molecule Page Version 1.0, Peer Reviewed And Published 24 Jan 2006 Version

Corresponding Author Kenneth A Taylor [email protected]

Authors Cheri M Hampton, Kenneth A Taylor

Editorial Board Patrick J Casey [email protected] (editorial board member list) Member

Species Mouse

Sequence 1 MVDYHAANQA YQYGPNSGGG NGAGGGGSMG DYMAQEDDWD RDLLLDPAWE 51 KQQRKTFTAW CNSHLRKAGT QIENIDEDFR DGLKLMLLLE VISGERLPKP 101 ERGKMRVHKI NNVNKALDFI ASKGVKLVSI GAEEIVDGNA KMTLGMIWTI 151 ILRFAIQDIS VEETSAKEGL LLWCQRKTAP YKNVNVQNFH ISWKDGLAFN 201 ALIHRHRPEL IEYDKLRKDD PVTNLNNAFE VAEKYLDIPK MLDAEDIVNT

Figure 3.6.1 α-Actinin 4 Overview Page 107 251 ARPDEKAIMT YVSSFYHAFS GAQKAETAAN RICKVLAVNQ ENEHLMEDYE 301 RLASDLLEWI RRTIPWLEDR VPQKTIQEMQ QKLEDFRDYR RVHKPPKVQE 351 KCQLEINFNT LQTKLRLSNR PAFMPSEGRM VSDINNGWQH LEQAEKGYEE 401 WLLNEIRRLE RLDHLAEKFR QKASIHEAWT DGKEAMLKQR DYETATLSDI 451 KALIRKHEAF ESDLAAHQDR VEQIAAIAQE LNELDYYDSH NVNTRCQKIC 501 DQWDNLGSLT HSRREALEKT EKQLETIDQL HLEYAKRAAP FNNWMESAME 551 DLQDMFIVHT IEEIEGLISA HDQFKSTLPD ADREREAILA IHKEAQRIAE 601 SNHIKLSGSN PYTTVTPQII NSKWEKVQQL VPKRDHALLE EQSKQQSNEH 651 LRRQFASQAN MVGPWIQTKM EEIGRISIEM NGTLEDQLSH LKQYERSIVD 701 YKPSLDLLEQ QHQLIQEALI FDNKHTNYTM EHIRVGWEQL LTTIARTINE 751 VENQILTRDA KGISQEQMQE FRASFNHFDK DHGGALGPEE FKACLISLGY 801 DVENDRQGDA EFNRIMSVVD PNHSGLVTFQ AFIDFMSRET TDTDTADQVI 851 ASFKVLAGDK NFITAEELRR ELPPDQAEYC IARMAPYQGP DAAPGALDYK 901 SFSTALYGES DL

Molecular Weight 104981

Isoelectric Point 5.25

Extinction Coefficient 124000 M-1cm-1

Absorption Coefficient 1.18

Aliphatic Index 80.93

Figure 3.6.1 Continued.

108 3.6.2 α-Actinin 4 Summary

Alpha-actinin-4

Protein Function

α-actinin-4 (Actn4) is a member of the spectrin-family of actin crosslinking proteins and is the most recently identified isoform (Dixson et al. 2003; Otey and Carpen 2004; Virel and Backman. 2004; Honda et al. 1998). Actn4 exists as an anti-parallel homodimer. The N-terminus consists of two highly conserved tandem calponin-homology domains (CH1, CH2) which are required for F-actin binding. The CH domains are connected to a series of four spectrin-like coiled-coil repeats which make up the rod domain that facilitates dimerization. Once regarded as simply spacers for actin bundle formation, these spectrin repeats have a newfound role as a docking site or scaffold for many signaling proteins (Djinovic-Carugo et al. 2002). The repeats are followed by a calmodulin-like domain that is composed of two EF hand domains. Aside from its role as an actin crosslinking protein, Actn4 serves to anchor integral membrane proteins or scaffolding proteins for signaling molecules to the actin cytoskeleton. Like Actn1, Actn4 is a calcium sensitive isoform (Burridge and Feramisco. 1981). It has 80% nucleotide and 86.7% amino acid similarity to Actn1. Despite this similarity, the roles of Actn4 and Actn1 are not functionally redundant as demonstrated by Kos et al. (2003).

Pub PM ID Authors Title Journal Date

Non-muscle alpha actinins are calcium- 10 Dec 7312045 Burridge K, Feramisco JR Nature, 294, 5841 sensitive actin-binding proteins. 1981

The alpha-actinin gene family: a revised Jan 12569417 Dixson JD, Forstner MJ, Garcia DM J Mol Evol, 56, 1 classification. 2003

Djinovic-Carugo K, Gautel M, Ylänne J, The spectrin repeat: a structural 20 Feb 11911890 FEBS Lett, 513, 1 Young P platform for cytoskeletal protein 2002

Figure 3.6.2 α-Actinin 4 Summary Page 109 assemblies.

Honda K, Yamada T, Endo R, Ino Y, Actinin-4, a novel actin-bundling protein 23 Mar 9508771 Gotoh M, Tsuda H, Yamada Y, Chiba H, associated with cell motility and cancer J Cell Biol, 140, 6 1998 Hirohashi S invasion.

Kos CH, Le TC, Sinha S, Henderson JM, Mice deficient in alpha-actinin-4 have J Clin Invest, Jun 12782671 Kim SH, Sugimoto H, Kalluri R, Gerszten severe glomerular disease. 111, 11 2003 RE, Pollak MR

Alpha-actinin revisited: a fresh look at Cell Motil Jun 15083532 Otey CA, Carpen O an old player. Cytoskeleton, 58, 2 2004

Molecular evolution and structure of Mol Biol Evol, Jun 15014165 Virel A, Backman L alpha-actinin. 21, 6 2004

Regulation of Activity

As Actn4 exists as an anti-parallel homodimer, the calmodulin-like domain of one molecule is in very close proximity to the actin- binding domain of the neighboring molecule. This suggests that Actn4’s ability to bind actin involves calcium binding to the EF hands (Burridge and Feramisco. 1981).

PM ID Authors Title Journal Pub Date

Burridge K, Feramisco Non-muscle alpha actinins are calcium-sensitive 10 Dec 7312045 Nature, 294, 5841 JR actin-binding proteins. 1981

Interactions with Ligands and Other Proteins

Actn4 binds to F-actin via its N-terminal actin-binding domain.

Actn4 interacts with cytoskeletal scaffolding and signaling proteins. The following interactions have been demonstrated to be specific for the Actn4 isoform: Actn4 interacts with the hemidesmosome transmembrane protein BP180 (Gonzalez et al. 2001). The

Figure 3.6.2 Continued. 110 PDZ-LIM protein Elfin/CLP-36 interacts with Actn4 at the stress fibers of colonic epithelium via binding of the PDZ domain to the spectrin repeats (Vallenius et al. 2000). PAI-1 (plasminogen activator inhibitor type 1), a protein with roles in inflammation, tumor invasion and metastasis, interacts specifically with Actn4 (Magdolen et al. 2004). Actn4 also anchors the enzyme iNOS (inducible nitric oxide synthase) to the actin cytoskeleton in macrophages (Daniliuc et al. 2003). SPA-1 is involved in the regulation of T cell activation in response to antigens through the control of Rap1 GTPase signaling, and is localized to the immunuological synapse via interaction with the Actn actin-binding domain (Harazaki et al. 2004).

Actn4 is also identified as a component of several multi-protein complexes. It is suggested that BERP, a novel RING finger protein, may anchor class V myosins to particular cell domains via its interaction with Actn4 (El-Husseini et al. 2000). Furthermore, BERP is part of the CART complex for cytoskeleton-associated recycling or transport consisting of hrs/Actn4/BERP/myosin V (Yan et al. 2005). RN-tre is a Rab5-GAP/effector that is involved in the formation of circular ruffles at the plasma membrane where it establishes a three-pronged connection with Rab5, F-actin and Actn4 (Lanzetti et al. 2004). The tight junction protein MAGI-1 binds to the C terminus of Actn4 via its fifth PDZ domain, while also binding synaptopodin in the polarized epithelial cells of the kidney (Patrie et al. 2002). The transmembrane synaptic adhesion molecule densin-180 forms a ternary complex with Ca2+/calmodulin-dependent protein kinase II (CaMKII) and Actn4 in the neural PSD and kidney podocytes. Densin-180 binds to Actn4 via its PDZ domain (Walikonis et al. 2001; Ahola et al. 2003). Another ternary complex is the NHE3/E3KARP/Actn4 in which E3KARP binds to the actin binding domain of Actn4 (maximal at 1 µM [Ca2+]i) to regulate Na/H exchanger 3 (Kim et al. 2002).

Actn4 has also been reported to interact with nuclear proteins. Actn4 interacts with NF-Y to recruit chromatin-remodeling complexes or to direct NF-Y/Actn4-targeted genes to the nuclear matrix and active transcriptional complexes (Poch et al. 2004). DNaseY is shown to interact with Actn4 and this interaction significantly enhances its endonuclease activity (Liu et al. 2004). Even more intriguing is the presence of Actn4 in the extracellular matrix as a result of cell movement. It is cleaved by monocyte-secreted urokinase to produce a 31 kDa amino-terminal fragment of Actn4 termed mactinin. Mactinin is shown to be a promoter of monocyte/macrophage maturation (Luikart et al. 1999; Masri et al. 1999; Luikart et al. 2002; Luikart et al. 2003).

Pub PM ID Authors Title Journal Date

Figure 3.6.2 Continued. 111 Ahola H, Heikkilä E, Aström E, A novel protein, densin, expressed by glomerular J Am Soc Jul 12819232 Inagaki M, Izawa I, Pavenstädt H, podocytes. Nephrol, 14, 7 2003 Kerjaschki D, Holthöfer H

Daniliuc S, Bitterman H, Rahat Hypoxia inactivates inducible nitric oxide synthase 15 J Immunol, 12960352 MA, Kinarty A, Rosenzweig D, Lahat in mouse macrophages by disrupting its interaction Sep 171, 6 N, Nitza L with alpha-actinin 4. 2003

Biochem Biophys El-Husseini AE, Kwasnicka D, BERP, a novel ring finger protein, binds to alpha- 27 Jan 10673389 Res Commun, Yamada T, Hirohashi S, Vincent SR actinin-4. 2000 267, 3

Gonzalez AM, Otey C, Edlund M, Interactions of a hemidesmosome component and J Cell Sci, Dec 11739652 Jones JC actinin family members. 114, Pt 23 2001

Harazaki M, Kawai Y, Su L, Specific recruitment of SPA-1 to the immunological Immunol Lett, 15 15081616 Hamazaki Y, Nakahata T, Minato N, synapse: involvement of actin-bundling protein 92, 3 Apr 2004 Hattori M actinin.

Ca(2+)-dependent inhibition of Na+/H+ exchanger Kim JH, Lee-Kwon W, Park JB, J Biol Chem, 28 11948184 3 (NHE3) requires an NHE3-E3KARP-alpha-actinin-4 Ryu SH, Yun CH, Donowitz M 277, 26 Jun 2002 complex for oligomerization and endocytosis.

20 Lanzetti L, Palamidessi A, Areces Rab5 is a signalling GTPase involved in actin Nature, 15152255 May L, Scita G, Di Fiore PP remodelling by receptor tyrosine kinases. 429, 6989 2004

15002038 Liu QY, Lei JX, LeBlanc J, Sodja C, Regulation of DNaseY activity by actinin-alpha4 Cell Death Jun Ly D, Charlebois C, Walker PR, during apoptosis. Differ, 11, 6 2004 Yamada T, Hirohashi S, Sikorska M

Figure 3.6.2 Continued. 112 Magdolen U, Schroeck F, Non-muscle alpha-actinin-4 interacts with Biol Chem, Sep 15493875 Creutzburg S, Schmitt M, Magdolen plasminogen activator inhibitor type-1 (PAI-1). 385, 9 2004 V

Interaction of two actin-binding proteins, 16 Patrie KM, Drescher AJ, Welihinda J Biol Chem, 12042308 synaptopodin and alpha-actinin-4, with the tight Aug A, Mundel P, Margolis B 277, 33 junction protein MAGI-1. 2002

Two distinct classes of CCAAT box elements that 15 Poch MT, Al-Kassim L, Smolinski Toxicol Appl 15364540 bind nuclear factor-Y/alpha-actinin-4: potential role in Sep SM, Hines RN Pharmacol, 199, 3 human CYP1A1 regulation. 2004

CLP-36 PDZ-LIM protein associates with nonmuscle J Biol Chem, 14 10753915 Vallenius T, Luukko K, Mäkelä TP alpha-actinin-1 and alpha-actinin-4. 275, 15 Apr 2000

Walikonis RS, Oguni A, Densin-180 forms a ternary complex with the 15 Jan 11160423 Khorosheva EM, Jeng CJ, Asuncion (alpha)-subunit of Ca2+/calmodulin-dependent J Neurosci, 21, 2 2001 FJ, Kennedy MB protein kinase II and (alpha)-actinin.

Yan Q, Sun W, Kujala P, Lotfi Y, CART: an Hrs/actinin-4/BERP/myosin V protein Mol Biol Cell, May 15772161 Vida TA, Bean AJ complex required for efficient receptor recycling. 16, 5 2005

Regulation of Concentration

No information is available yet.

Subcellular Localization

Actn4 is found at stress fibers, microfilament bundles, and sites of cell contact or adhesion. Its localization may differ somewhat from that of Actn1, the other calcium-sensitive isoform. It has also been reported that Actn4 may translocate to the nucleus in some cells.

Figure 3.6.2 Continued. 113 Pub PM ID Authors Title Journal Date

Actinin-4, a novel actin-bundling protein Honda K, Yamada T, Endo R, Ino Y, Gotoh J Cell Biol, 23 Mar 9508771 associated with cell motility and cancer M, Tsuda H, Yamada Y, Chiba H, Hirohashi S 140, 6 1998 invasion.

Bioessays, Feb 15666356 Young KG, Kothary R Spectrin repeat proteins in the nucleus. 27, 2 2005

Major Sites of Expression

Actn4 expression is widespread (Honda et al. 1998).

Pub PM ID Authors Title Journal Date

Phenotypes

Overexpression of cytoplasmic Actn4 is associated with increased cell motility and a decrease in Actn1 at sites of adhesion. Inactivation and translocation of Actn4 to the nucleus is linked to decreased metastatic potential.

A mutation in ACTN4 has been implicated in Focal Segmental Glomerulosclerosis. A possible alternative splice variant may be associated with small-cell lung carcinoma.

Pub PM ID Authors Title Journal Date

Figure 3.6.2 Continued. 114 15388893 Exp Biol Med Fan Q, Ding J, Zhang J, Guan N, Deng Effect of the knockdown of podocin mRNA on Oct (Maywood), J nephrin and alpha-actinin in mouse podocyte. 2004 229, 9

Expression of alpha-actinin-4 in acquired human Goode NP, Shires M, Khan TN, Nephrol Dial Apr 15031339 nephrotic syndrome: a quantitative Mooney AF Transplant, 19, 4 2004 immunoelectron microscopy study.

Goto H, Wakui H, Komatsuda A, Renal alpha-actinin-4: purification and Nephron Exp Jan 12411747 Ohtani H, Imai H, Sawada K, Kobayashi puromycin aminonucleoside-binding property. Nephrol, 93, 1 2003 R

J Am Soc Podocyte differentiation and hereditary Jun 12761234 Gubler MC Nephrol, 14 Suppl proteinuria/nephrotic syndromes. 2003 1

Honda K, Yamada T, Endo R, Ino Y, 23 Actinin-4, a novel actin-bundling protein J Cell Biol, 9508771 Gotoh M, Tsuda H, Yamada Y, Chiba H, Mar associated with cell motility and cancer invasion. 140, 6 Hirohashi S 1998

Honda K, Yamada T, Seike M, Alternative splice variant of actinin-4 in small cell Oncogene, 1 Jul 15122314 Hayashida Y, Idogawa M, Kondo T, Ino lung cancer. 23, 30 2004 Y, Hirohashi S

Kaplan JM, Kim SH, North KN, Rennke H, Correia LA, Tong HQ, Mathis BJ, Mutations in ACTN4, encoding alpha-actinin-4, Nat Genet, Mar 10700177 Rodríguez-Pérez JC, Allen PG, Beggs AH, cause familial focal segmental glomerulosclerosis. 24, 3 2000 Pollak MR

12617336 Komatsuda A, Wakui H, Maki N, Analysis of mutations in alpha-actinin 4 and Ren Fail, 25, 1 Jan

Figure 3.6.2 Continued. 115 Kigawa A, Goto H, Ohtani H, Hamai K, podocin genes of patients with chronic renal failure 2003 Oyama Y, Makoto H, Sawada K, Imai H due to sporadic focal segmental glomerulosclerosis.

Kos CH, Le TC, Sinha S, Henderson Mice deficient in alpha-actinin-4 have severe J Clin Invest, Jun 12782671 JM, Kim SH, Sugimoto H, Kalluri R, glomerular disease. 111, 11 2003 Gerszten RE, Pollak MR

Antitumor cytotoxic T-lymphocyte response in Mami-Chouaib F, Echchakir H, Immunol Rev, Oct 12445285 human lung carcinoma: identification of a tumor- Dorothée G, Vergnon I, Chouaib S 188 2002 associated antigen.

Menez J, Le Maux Chansac B, Mutant alpha-actinin-4 promotes tumorigenicity Oncogene, 8 Apr 15048094 Dorothée G, Vergnon I, Jalil A, Carlier and regulates cell motility of a human lung 23, 15 2004 MF, Chouaib S, Mami-Chouaib F carcinoma.

Michaud JL, Lemieux LI, Dubé M, Focal and segmental glomerulosclerosis in mice J Am Soc May 12707390 Vanderhyden BC, Robertson SJ, with podocyte-specific expression of mutant alpha- Nephrol, 14, 5 2003 Kennedy CR actinin-4.

The genetic basis of FSGS and steroid-resistant Semin Nephrol, Mar 12704574 Pollak MR nephrosis. 23, 2 2003

Alpha-actinin-4-mediated FSGS: an inherited Yao J, Le TC, Kos CH, Henderson JM, Jun 15208719 kidney disease caused by an aggregated and PLoS Biol, 2, 6 Allen PG, Denker BM, Pollak MR 2004 rapidly degraded cytoskeletal protein.

Splice Variants

A possible alternative splice variant may be associated with small cell lung carcinoma.

PM ID Authors Title Journal Pub

Figure 3.6.2 Continued. 116 Date

Honda K, Yamada T, Seike M, Hayashida Y, Alternative splice variant of actinin-4 Oncogene, 1 Jul 15122314 Idogawa M, Kondo T, Ino Y, Hirohashi S in small cell lung cancer. 23, 30 2004

Antibodies

Mouse monoclonal antibody (mAb) to chicken gizzard actinin, clone JLN20 (ICN, InnoGenex, BioGenex, Oncogene Research Products).

Mouse mAb to human actinin, clone AT 6/172 (Research Diagnostics Inc, Immune Systems Ltd).

Mouse mAb to quail smooth muscle actinin, 1E12 (Developmental Studies Hybridoma Bank at the University of Iowa).

Mouse mAb to chicken gizzard, clone CB11 (ICN, Affiniti).

Rabbit polyclonal Ab to chicken gizzard (ICN).

Mouse mAb to human red blood cell ghosts, clone RBC2/1B6 (Novocastra).

Mouse mAb to smooth muscle actinin, clone 1A4 (Diagnostic BioSystems).

Goat polyclonal Ab to aminoterminus of human ACTN1, N-19 (Santa Cruz Biotechnology).

Goat polyclonal Ab to human ACTN1 carboxyterminus, C-20 (Santa Cruz Biotechnology).

Figure 3.6.2 Continued.

117

3.6.3 α-Actinin4 Network Map

Alpha-actinin-4

Figure 3.6.3 α-Actinin 4 Network Map

118 3.6.4 α-Actinin 4 States

Alpha-actinin-4

State List Version 1.0, Peer Reviewed And Published 24 Jan 2006 doi:10.1038/mp.a000197.01 How to cite this Molecule Page

Functional states

State description State name Location Transition graph

2(Actn4) 2(Actn4)[1] actin cytoskeleton

2(Actn4)/actin (cytoskel) 2(Actn4) (Act[b,a1,a2,c1,g1,g2]) actin cytoskeleton

2(Actn4)/BP180 2(Actn4) (Act[b,a1,a2,c1,g1,g2]) (COL17A1) actin cytoskeleton

2(Actn4)/Elfin 2(Actn4) (Act[b,a1,a2,c1,g1,g2]) (PDLIM1) actin cytoskeleton

2(Actn4)/PAI-1 2(Actn4) (Act[b,a1,a2,c1,g1,g2]) (SERPINE1) actin cytoskeleton

2(Actn4)/SPA-1 2(Actn4) (Act[b,a1,a2,c1,g1,g2]) (SIPA1) actin cytoskeleton

2(Actn4)/BERP 2(Actn4) (Act[b,a1,a2,c1,g1,g2]) (Trim3) actin cytoskeleton

2(Actn4)/RN-tre 2(Actn4) (Act[b,a1,a2,c1,g1,g2]) (USP6NL) actin cytoskeleton

2(Actn4)/MAGI-1 2(Actn4) (Magi1) (Act[b,a1,a2,c1,g1,g2]) actin cytoskeleton

2(Actn4)/iNOS 2(Actn4) (Nos2) (Act[b,a1,a2,c1,g1,g2]) actin cytoskeleton

2(Actn4)/CaMK/densin180 2(Actn4) (Camk2a) (Act[b,a1,a2,c1,g1,g2]) (Lrrc7) actin cytoskeleton

2(Actn4-Ca) 2(Actn4 Ca) cytoplasm

2(Actn4-Ca)/Nherf2/E3KARP 2(Actn4 Ca) (SLC9A3) (Slc9a3r2) cytoplasm

2(Actn4) (nuc) 2(Actn4)[2] nucleus

2(Actn4)/DNaseY 2(Actn4) (Dnase1l3) nucleus

Figure 3.6.4 α-Actinin 4 States Page.

119 2(Actn4)/NF-YB 2(Actn4) (Nfyb) nucleus

2(Actn4) (ECM) 2(Actn4)[3] extracellular matrix

2(Actn4)/Urokinase 2(Actn4) (PLAU) extracellular matrix

Mactinin 2(Actn4 O) extracellular matrix

Figure 3.6.4 Continued.

120 3.6.5 α-Actinin 4 Transitions

Alpha-actinin-4

State Transitions Version 1.0, Peer Reviewed And Published 24 Jan 2006

doi:10.1038/mp.a000197.01 How to cite this Molecule Page

Transitions

2(Actn4)[1] --> 2(Actn4) (Act[b,a1,a2,c1,g1,g2])

2(Actn4)[1] --> 2(Actn4)[2]

2(Actn4)[1] --> 2(Actn4)[3]

2(Actn4) (Act[b,a1,a2,c1,g1,g2]) --> 2(Actn4) (Act[b,a1,a2,c1,g1,g2]) (COL17A1)

2(Actn4) (Act[b,a1,a2,c1,g1,g2]) --> 2(Actn4) (Act[b,a1,a2,c1,g1,g2]) (PDLIM1)

2(Actn4) (Act[b,a1,a2,c1,g1,g2]) --> 2(Actn4) (Act[b,a1,a2,c1,g1,g2]) (SERPINE1)

2(Actn4) (Act[b,a1,a2,c1,g1,g2]) --> 2(Actn4) (Act[b,a1,a2,c1,g1,g2]) (SIPA1)

2(Actn4) (Act[b,a1,a2,c1,g1,g2]) --> 2(Actn4) (Act[b,a1,a2,c1,g1,g2]) (Trim3)

2(Actn4) (Act[b,a1,a2,c1,g1,g2]) --> 2(Actn4) (Act[b,a1,a2,c1,g1,g2]) (USP6NL)

2(Actn4) (Act[b,a1,a2,c1,g1,g2]) --> 2(Actn4) (Magi1) (Act[b,a1,a2,c1,g1,g2])

2(Actn4) (Act[b,a1,a2,c1,g1,g2]) --> 2(Actn4) (Nos2) (Act[b,a1,a2,c1,g1,g2])

2(Actn4) (Act[b,a1,a2,c1,g1,g2]) --> 2(Actn4) (Camk2a) (Act[b,a1,a2,c1,g1,g2]) (Lrrc7)

2(Actn4) (Act[b,a1,a2,c1,g1,g2]) --> 2(Actn4 Ca)

Figure 3.6.5 α-Actinin 4 Transitions Page. 121 2(Actn4 Ca) --> 2(Actn4 Ca) (SLC9A3) (Slc9a3r2)

2(Actn4)[2] --> 2(Actn4) (Dnase1l3)

2(Actn4)[2] --> 2(Actn4) (Nfyb)

2(Actn4)[3] --> 2(Actn4) (PLAU)

2(Actn4) (PLAU) --> 2(Actn4 O)

Figure 3.6.5 Continued.

122 3.7 α-Actinin Interacting Proteins

Table 3.7.1 α-Actinin Interacting Proteins Interacting Which Where in the Affinity Site of interaction Function Method of PMID protein ACTN cell interaction Actin 1, 2, 3, 4 FAs, stress fibers, Z Kd=1.6 ACTN 86-117 & 350-375: bundles F-actin EM of ABD decorated 9545287; 8034735; disk uM actin interface btwn subdom1 F-actin; difference 8034744; 2124107; and 2 of two separate mapping 8449927; 3558365; monomers 9334343; 9731773; 11381081 Acetylcholine 2 plasmalemma; AChR clusters interact with co-loc.; immuno-blot; 6408100; 10771514 receptor neuromuscular the cytoskeleton co-purrification junction ADAM 12 1, 2 transmembrane ADAM12 cytoplasmic tail, ADAM12 is both a Y2H, ACTN1:11439084, ACTN-2 SR 3-C-term; 58- disintegrin and immunoprecipitation ACTN2:10788519 amino acid C-term ADAM 12: metalloprotease involved in from in vivo 27 kDa N-term ACTN-1 skeletal muscle development and regeneration; cell signalling during myoblast differentiation and fusion Adenosine 1,2,3,4 neronal cell C-term of A2AR, attachment of A2AR to the Y2H, 12837758 A2A receptor by Y2H membrane actin cytoskeleton via ACTN immunoprecipitation, for membrane clustering colocalization, pull- down expt ADIP 1 Nectin/Afadin and E- first coiled-coil domain of cell-cell Ajs Y2H, co- 12446711 cadherin based Ajs ADIP; EF hands of ACTN immunopricipitation (cell-cell Ajs) Adriamyosin 2 heart myofibrils drug; antibiotic equilibrium dialysis and 7067062 gel-filtration Affixin (B- 1,2 sarcolemma; initial Strong 249-272 affixin CH2 mediates anchoring of pull-down and blot 15159419 parvin) cell-substrate domain(phosphorylated by ACTN to F.A.s in an overlay assay, co- adhesion, FAs, tip of ILK); both N- and C-term adhesion-dependent immuno ppt, co- leading edge, fine ACTN fashion localization SFs

123 Table 3.7.1 Continued Interacting Which Where in the Affinity Site of interaction Function Method of PMID protein ACTN cell interaction ALP 2 Z-line; intercalated Kd=8.6 ALP PDZ domain; ACTN SR sarcomere; binds on either blot overlay, in vitro 12589684, 15084604, disc uM PDZ 3, also EF 3-4; ZM motif of side of actn "hinge" region binding, coimmuno ppt., 16476425, 9334352, domain ALP: alpha-actinin rod which may regulate flexibility biochemistry, Y2H 10506181 binding to (necessary and sufficient for EF3-4 localization) (Klaavunie mi et al) Amorphin 2 Sarcomeric Z-band release of glucose-1- blot overlay, co-loc. 12211109 (Phosphorylas phosphate from glycogen? e) Angeogenin 2 only used ACTN 375–894 Y2H, FRET, photo- 15737636 bleaching BERP (Trim3) 4 cytoplasmic BERP RBCC domain: ACTN suggest that BERP may Y2H, coIP, co-loc 10673389 anchor class V myosins to particular cell domains via its interaction with alpha- actinin-4 BP-180 1,4 hemidesmosome, BP180 cytoplasmic domain; adhesion Y2H, immuno-ppt., 11739652 (collagen XVII, cell-cell interactions ACTN EF hand domians immuno-blot BPAG2) Calcineurin 2 z-disks and calcineurin is tethered to a calcium-dependent 15489953; 11114196 (PP2B) neurons ACTN at the Z-line via phosphorylase calsarcins Calcium 2 mouse cardiac-in mouse cardiac myocyte ACTN modulates activity of co-loc, indirect methods 11997265 channel- the membrane over membranes channel in cariac tissue cardiac-L-type Z-line regions Calpain-1 1,2 Kd(- 80KDa subunit of calpain-1; Cleavage of C-term 5 Kda ELISA solid-phase 14622253, 12358155 largely Ca2+)= C-term of ACTN ACTN assay, co-loc chicken 0.5 uM, skeletal Kd(+Ca2+ )=0.05 uM a ten-fold increase in activity Calsarcins 2 z-line calsarcin 153-200: ACTN SR tether calcineurin to ACTN Y2H, coIP, co-loc 11842093, 11114196 (1,2,3)(FATZ, 2,3 at sarcomeric z-line of myozenin) cardiac and skeletal muscle CaMK II 4 PSD CaMKII kinase domains; Y2H, pull-down assay 11160423 (alpha and ACTN 806-871 beta)

124 Table 3.7.1 Continued Interacting Which Where in the Affinity Site of interaction Function Method of PMID protein ACTN cell interaction CapZ (beta- 2 located in nascent microM ACTN 55 kDa repetitive 2341404, 10412090 actinin) Z-lines during range; domain (SRs) skeletal muscle weakened myofibrillogenesis in the presence of phosphoin ositides alpha-catenin 1 Cadherin-mediated alpha-catenin 325-394: ACTN links cytoskeleton to Y2H, coIP, co-loc, cell-cell junctions 479-529 cadheren junctions beta-catenin 4 cytoskeleton of Regulation of interaction is co-immuno ppt., mass 16204054 colorectal cancer via E-cadherin spec. cells clathrin clathrin heavy chain: ACTN CLP-36 1, 4 stress fibers CLP-36 PDZ: ACTN SRs immuno ppt., MALDI- 10753915 TOF CRP1 actin stress Kd=1.8 CRP1 LIM1: ACTN ABD muscle differentiation; western, solution and 10926853' 16336664 (cysteine-rich fibers; +/- 0.3 uM localize CRP1 to the actin solid-phase binding protein) fibroblasts cytoskeleton; CRP-1 assays, co-loc, coIP, and smooth stabilizes ACTN:F-actin deletion mapping. FRET muscle interaction Cypher 1 and 2 Z-lines of cypher PDZ domain; ZASP is an adaptor protein biochemical, co-loc. 10391924, 11696561, 2 (ZASP) cardiac, specifically the ZM-motif : bringing PKC to Z-lines via Western, Surface 16476425 striated ACTN rod LIM domains Plasmon Resonance muscle, stress fibers dendrin GenBank dendritic spectrin repeats: dendrin suggests that it plays a role Y2H; EGFP and western 16464232 Acc. No. spines middle domain MD in linking the PSD core to blot P12814 theunderlying cytoskeleton of the spine. Densin-180 2, 4 PSD of intracellular PDZ domain of ternary complex: NMDAR- Y2H, pull-down assay, 11160423, 16120608, neurons; densin; ACTN 464-879 NR2B:densin-180:ACTN all overlay assay, 16172120 glomerular bind CaMK II podocytes DNaseY 4 nucleus DNaseY, a Ca(2+)- and Y2H, coIP, 15364540, 15002038 Mg(2+)-dependent endonuclease implicated in apoptotic DNA degradation

125 Table 3.7.1 Continued Interacting Which Where in the Affinity Site of interaction Function Method of PMID protein ACTN cell interaction desmin Z-disks, co-localization gel electrophoresis- 6889960 (skeletin) intercalated derived enzyme-linked disks; immunosorbent assay intermediate (GEDELISA) filaments of bovine heart Purkinje fibres Dystrophin 2 skeletal C-term dystrophin: binding of alpha-actinin-2 to ligand-blotting, co-loc, 10328815 muscle the carboxyl-terminus of indirect sarcolemmal dystrophin is the immunolocalization vesicles, communication link between plasma the integrins and the membranes dystrophin/dystrophin- glycoprotein complex E3KARP 4 NHE3/E3KA maximum C-term 2nd PDZ domain of calcium-dependent pull-down assay, 11948184 RP/ACTN-4 at 1 uM E3KARP; ABD ACTN with regulation of NHE3 (Na/H MALDI-TOF, immono complex [Ca2+}I calcium bound Exchanger 3) co-loc, in vivo co- intestinal immuno ppt epithelium Elfin (hCLIM1, 1,2,4 ACTN2 Z- Elfin intervening sequence; Y2H, co- ACTN2:10861853, CLP36, disks of ACTN1,4 SR 2 and 3;; Elfin immunopricipitation, blot ACTN1,4:10753915 Pdlim1) myocardium; PDZ; ACTN2 C-term EF hand overlay assay, pull- actin stress downs fibers of myoblasts; ACTN1,4 platelet ENH (Enigma 2 Z-disk of PDZ domain of ENH with scaffolding pull-down, co-loc 10833443 homologue heart and actin and alpha-actinin protein) skeletal muscle FAK 1? focal tyrosine12 ACTN phosphorylation decreases site-directed complexes affinity of ACTN for Actin mutagenesis, immuno ppt FATZ (aka 2 Z-disk of skeletal C-term FATZ; ACTN SR 3,4 Z-disk assembly Y2H, overlay and pull- Calsarcin-2) muscle (431-719) down assays, immuno ppt

126 Table 3.7.1 Continued Interacting Which Where in the Affinity Site of interaction Function Method of PMID protein ACTN cell interaction FBPase z-line, both sides strong binds as a ternary complex glyconeogenic metabolon in co-loc, co-sed, biacore 12507293, 14592412, (fructose-1,6- aldolase:FBPase:actn vertebrates' myocytes. biosensor 15757649 bisphosphatas e, Pfkfb2) GLUT1CBP 1 cell membrane PDZ domain glucose transporter Y2H 10198040 localization GpIbalpha 1- GpIb-IX complex in cytoplasmic tail GpIbalpha regulation of platelet immuno ppt 11802708 phosphor activated platelets aggregation ylated GRIP1 2 nucleus GRIP AD2 15072553 Hepatitus C cell membrane 84-95 and 466-478 HCV; replication complex deletion mutant analysis, 14623081 virus NS5B ACTN SR 4 (621-733) assembly with HCV RNA co-loc, co-immuno ppt, Y2H, siRNA HSP20 2 actin cytoskeleton vasorelaxation co-immuno ppt, co-loc 12842460 of muscle ICAM-1 ICAM-1 478-505 cell adhesion linkage to solid-phase binding, 1355095 cytoskeleton coIP, co-loc ICAM-2 1 intercellular cell ICAM-2 231-253 cytoplasmic; cell adhesion linkage to co-immuno ppt, co-loc, 8824270 adhesions ACTN cytoskeleton Integrin, beta 1, cell adhesions beta1 cytoplasmic tail : rod cell adhesion linkage to solid-phase binding, 16513669, 16430917, 1 chicken between SR 2 & 3 cytoskeleton; necessary for immuno-blot, VPM; EM 15721583' 15710624 gizzard tension development; regulation via PIP2-binding Integrin, beta leukocyte a single 11-amino acid region binding of leukocytes to 9837942, 8104223 2 (CD18) cytoskeleton (residues 736-746) is endothelial cells necessary and sufficient for alpha-actinin binding Integrin, beta 1 focal contacts 11967230 3 Kettin insect muscle z- 12560078 disks 5- 1 cell membrane for localization if inflammation Solid-phase binding 7929073 Lipoxygenase inflamation response enzyme at the cell membrane LPP (lipoma focal adhesions lesser LPP proline-rich N-term; regulation of focal blot overlay, Y2H, Y3H, 12615977 preferred affinity ACTN SR 2 and 3 adhesions co-loc partner) than zyxin

127 Table 3.7.1 Continued Interacting Which Where in the Affinity Site of interaction Function Method of PMID protein ACTN cell interaction L-Selectin plasma membrane; Kd = 550 L-selectin cytoplasmic tail The leukocyte adhesion solid phase binding 7538138 leukocyte adhesion nM molecule L-selectin assays, co ppt., mediates binding to lymph node high endothelial venules (HEV) and contributes to leukocyte rolling on endothelium at sites of inflammation MAGI-1 4 podocyte tight fifth PDZ domain of MAGI-1 in vitro binding assays, 12042308 (Baiap1) junctions binding to the C terminus of co-loc, alpha-actinin-4 MEKK 1 1, 4? strees fibers and 1-719 MEKK1; ACTN 371- linking extracellular signals Y2H, co-immuno ppt, 10401575 focal adhesions 892 to the cytoskeleton co-loc, MLP 2 z-disks MLP LIM1 domain MLP associates with ACTN 14567970, 12589684, in late stages of 15205937, 16329665, myofibrillogenesis; forms a 15582318 complex that acts as a mechanosensor myopalladin 2 Z-lines myopalladin C-term; ACTN Z-disk; tethers together the Y2H, deletion mapping 11309420 (MYPN) EF hands COOH-terminal Src homology 3 domains of nebulin and nebulette with the EF hand motifs of alpha- actinin in vertebrate Z-lines myospryn 2 costamere 16407236 myotilin (TTID) 2 Z-line of skeletal Kd=6nM 1-214 myotilin; ACTN SR 3 sarcomere archetecture Y2H, co-loc, affinity ppt, 12899871, 12499399, and cardiac muscle (high) and 4 deletion mapping 10369880 Myozenin (aka 2, 3 Z-lines and sarcomere archetecture Y2H, coIP, co-loc 10984498, 11171996, Calsarcin-2, nemaline rods 11699871, FATZ) Nebulin 2 z-disk 2387397 NCAM (neural 1 membranes C-term NCAM; cell adhesion and signaling affinity chromatography 14550299 cell adhesion molecule)140 and

128 Table 3.7.1 Continued Interacting Which Where in the Affinity Site of interaction Function Method of PMID protein ACTN cell interaction NF-Y 4 neucleus ACTN4 may assist NF-Y in solid-phase, co-IP 15364540 recruiting chromatin- remodeling complexes or may direct NF-Y/ACTN4- targeted genes to the nuclear matrix and active transcriptional complexes. NMDA 2,3 membranes, PSD, 48nM NMDA receptor NR1 C-term ACTN binding reduces 9009191, 9454847, receptor glutamatergic C0 region also NR2B channel shut time 9952395, 10809744, synapses (increases Popen) 10925145, 16510730, 16120608, 16172120, 15073274 iNOS 4 macrophage anchor to the cytoskeleton immunoprecipitation, 12960352, 16492779 submembranal mass spectroscopy, and zones confocal microscopy Palladin 1,2 focal adhesions palladin 222-280; ACTN EF ACTN targets palladin to Y2H, affinity ppt 15147863 and stress fiber 3-4 stress fibers dense regions PAI-1 1, 4 cell surface ACTN4 C-term 330-911 mononuclear phagocyte Y2H, coIP, solid phase 15493875 (plasminogen response to inflamation binding assays activator inhibitor type Pdlim2/Mystiq 1, 4 corneal epithelial pull-down assay and 15505042 ue cells, lung; stress mass spectrometry; gel fibers overlay assay with purified proteins Phosphorylase 2 Sarcomeric Z-band release of glucose-1- blot overlay, co-loc. 12211109 (Amorphin) phosphate from glycogen? PI 3-Kinase 1 cell adhesions proline-rich region of p85 cytoskeleton regulation copurification, coIP, 11802708 subunit PtdIns(4,5)-P2 1, 2? at the plasma Kd=23 uM ACTN ABD CH2(168- PIP2 modulates ACTN co-loc, inhibition of 15710624, 14670080, membrane; focal 184)[158-174]: 4th and 5th cross-linking of proteolysis 12716899, 11101506, adhesions AND position of the inositol head actin{50uM/54.1%inhibition}; 10412090 stress fibers group increases affinity of actn for titin; inhibits binding to capZ PtdIns(3,4,5)- 1 focal adhesions Kd=23 uM ACTN ABD CH2(168- PIP3 inhibits ACTN cross- fluoresence shift, 15710624, 14670080, P3 and stress fibers 184)[158-174]: 4th and 5th linking of actin[50uM/91.9% cosedimentation, 12716899 position of the inositol head inhibition].Also, disrupts mutational analysis group ACTN binding to beta- integrin

129 Table 3.7.1 Continued Interacting Which Where in the Affinity Site of interaction Function Method of PMID protein ACTN cell interaction PKD 1, 4? focal adhesions both intracellular N-&C-temini A role for polycystin-1 as a Immunocytochemistry, 15843396, 11113628 (polycystin-1, - of polycystin-2 interact with cell surface receptor sucrose density gradient 2) actn involved in cell-matrix and sedimentation, co- cell-cell interactions has immunoprecipitation been proposed. ACTN analyses and in vitro stimulates polycystin role as binding assays; GST- a calcium channel pull-down PKN (PKN1) 1 +Ca2+, neurofilaments; PKN 136-189; ACTN SR3, PKN is a fatty acid-activated Y2H, in vitro binding, ACTN1:11802708, 2 platelet EF hands{ACTN-1 1mM serine/threonine protein coIP, ACTN1,2:9030526 cytoskeleton Ca2+ Dependent}Binding is kinase, having a catalytic enhanced in the presence of domain homologous to PIP2 protein kinase C family. Potentially phosphorylates ACTN on Nterminus PLD2 2? myocardial PLD2's N-terminal 185 Purified alpha-actinin mass spectroscopy, in 10801846 sarcolemmal amino acids potently inhibits PLD2 vitro binding assay , co- membranes activity (IC(50) = 80 nm) in loc an interaction-dependent and ADP-ribosylation factorreversible manner Potassium 2 membrane/cytoskel Kv1.4 and 1.5 N-term; ACTN- Localization of channel Y2H, coIP, in vitro 11809852, 10812072, channels eton 2 SRs binding, deletion 11389904 Kv1.5, Kv1.4 mapping Prohibitin/proh 1 Kd=675n Prohibitin/prohibitone C-term; Y2H, surface plasmon 12628297 ibitone (p32/ M ACTN ? Ca2+ inhibits resonance, coIP, p37) interaction deletion mapping Rabphilin-3A 2 neurons, docking of Rabphilin-3A N-term; ACTN enhances ACTN bundling of Y2H, in vitro binding, 8943213, 16043482 synaptic vesicles, 344-859; inhibited by Rab3A F-actin. Only associates coIP, dense core with F-actin that is cross- vessicles linked by actinin Raver1 costamere, cell Raver1 is a dual in vitro binding assays 11724819, 14633994 adhesions compartment protien, binds to RNA in the nucleus; associates with the cytoskeleton in adhesions. Microfilament anchoring?

130 Table 3.7.1 Continued Interacting Which Where in the Affinity Site of interaction Function Method of PMID protein ACTN cell interaction RIL (reversion- 2? postsynaptic RIL PDZ domain RIL links AMPA receptors to 14729062, 15456832 induced LIM density; dendritic the alpha-actinin/actin protein, spines, stress cytoskeleton; RIL enhanced Pdlim4) fibers the ability of alpha-actinin to cosediment with actin filaments; a regulator of actin stress fiber turnover RN-tre 4 plasma membrane RN-tre, a Rab5-specific 15152255 GTPase-activating protein (GAP); propose that RN-tre establishes a three-pronged connection with Rab5, F- actin and actinin-4; crosslinking of actin fibres into actin networks at the plasma membrane SHP-1 1 platelet membrane establish that SHP-1 can coIP 15070900 dephosphorylate alpha- actinin in vitro and in vivo and suggest that SHP-1 may regulate the tethering of receptors to the cytoskeleton and/or the extent of cross-linking of actin filaments in cells such as platelets Skeletin intermediate gel electrophoresis- 6889960 (desmin) filaments of bovine derived enzyme-linked heart Purkinje immunosorbent assay fibres; Z-disks, (GEDELISA) intercalated disks smitin 2 muscle ACTN SR2-3 and EF 3-4 in vitro 15833278 SPA-1 4 immunological full-length SPA-1, ACTN ABD SPA-1 regulates Rap1GTP Y2H, co-loc 15081616 synapse signaling at matrix-adhesion regions SpOtx sea urchin embryos specific interaction between Otx is translocated to the Y2H 8952065 the proline- rich region of nuclei; however its SpOtx and a putative SH3 interaction with actinin in domain of the sea urchin micromeres prevents this alpha- actinin translocation and it remains in the cytoplasm

131 Table 3.7.1 Continued Interacting Which Where in the Affinity Site of interaction Function Method of PMID protein ACTN cell interaction Syndecan-4 focal adhesions, syndecan-4 V region syndecan4is a co-loc, coIP, in vitro 12493766 stress fibers transmembrane receptor; binding; Triton X-100 regulates FA and SF extraction formation; alpha-actinin immunoprecipitation and interacts with syndecan-4 in in vitro binding assays a beta-integrin-independent manner. Synemin 2 cardiac z-bands; synemin rod and tail domains immuno ppt.; 10506213, 15657940, also glioblastoma interact with ACTN 12669240 ruffles Tir plasma membrane Tir N-term: ACTN ABD Tir, an enteropathogenic coIP, blot overlay, co-loc 11093251, 12112139 E.coli protein, anchors the bacteria to the host cell's cytoskeleton Titin 1,2 Z-disk of skeletal Kd=100- titin z-repeat 7 : ACTN EF3,4 ACTN anchors titin to the CD, calorimetry, NMR 9438132, 9196024, muscle 120 nM edge of the sarcomere. Titin 9003807, 9245597, binding is inhibited by ACTN 9501083, 10819994, in the closed conformation, 10462489 enhanced with PIP2 binding. Thrombospon 1 platelet cytosol kd=6.6 nM ligand-blotting,solid- 7750553, 9351403 din phase binding Urokinase extracellular matrix urokinase is a serine in vivo uPA knock-out 12932295, 12183060, protease that digests ACTN mice; cell lysate 10029174, 10029173 to produce a 31 Kd fragment enzymology called mactinin wich acts as a macrophage maturation factor and is involved in the inflamitory response. Inhibited by Amiloride and PMSF Vinculin 1, 2 cadherin-mediated Kd=13 uM vinculin head: ACTN 713-749 cell adhesion complex low-speed cosed, blot 16430917, 8037676 cell-cell junctions overlay, cryoEM Zasp 2 heart and skeletal zasp PDZ domain/ZM-motif: ZASP is an adaptor protein Y2H, coIP, co-loc 10427098, 15062084, (LDB3)/Cyphe muscle Z-bands; ACTN Rod and C-term bringing PKC to Z-lines via 11696561, 10391924, r stress fibers LIM domains 16476425

132 Table 3.7.1 Continued Interacting Which Where in the Affinity Site of interaction Function Method of PMID protein ACTN cell interaction Zyxin 1, 2 adherens junctions, moderate zyxin: ACTN N-term a putative solid phase binding 14967842, 16247023 adhesion plaques mechanotransducer, zyxin assays, cosed, blot relocates from z-disks to the overlay, analytical gel nucleus in response to filtration cyclic stretch; at focal adhesions zyxin translocates from the adhesion plaque to F-actin in reponse to cyclic stretch Vpx (HIV-2 1 nucleus Vpx C-term: ACTN nuclear import of viral pre- Y2H, coIP, co-loc, 15483243 and integration complex western, deletion SIVmac239 mapping, in vitro binding virus) assay,

133 CHAPTER 4 VILLIN IS AN UNUSUAL SUSPECT IN F-ACTIN CROSS-LINKING

We have applied correspondence analysis to 3-D volumes generated from electron tomography of 2-D rafts of F-actin cross-linked with chicken intestinal villin on a lipid monolayer to investigate villin: F-actin binding and cross-linking. More than 6,000 paired F-actin crossover repeats with villin protein bound between them were selected, aligned, and classified to produce class averages of villin cross-links with improved signal-to-noise ratio. Villin: F-actin rafts formed predominantly polar filament arrays. The overwhelming majority of villin repeats classified together in a single class. Fifteen other classes of only 50-60 repeats each showed slight villin rotation, truncation, or differences in which actin monomers were bound. Docking of the highly homologous gelsolin domain structures plus the villin head piece structure into the global average density reveals the homogeneous villin/actin binding interfaces which are unlike most other known actin-binding protein interactions, such as cofilin, profillin, DNase I, or gelsolin segments G1 and G2. This is the first glimpse of the entire structure of villin in a cross-linking role. Up until now it has been assumed that villin interacts with F-actin in a similar fashion to its close homolog, gelsolin. This study shows this assumption to be wrong and further lends concrete evidence to the notion that there can be multiple modes of interaction with actin even among highly homologous actin-binding proteins.

4.1 Background

Villin is an ~95 kD protein found primarily in the microvilli of intestinal epithelium and kidney proximal tubules along with another cross-linking protein fimbrin, where it functions to nucleate actin polymerization, cross-link filaments, sever F-actin at high Ca2+concentration, and cap the newly formed barbed ends of filaments. Villin is one of two founding members of the gelsolin superfamily (Yin and Stossel, 1979; Bretscher and Weber, 1980). Other members include adseverin/scinderin, CapG, fragmin, flightless I /severin, and supervillin. These proteins function to regulate actin filament length by severing and/or capping the barbed ends. Each member has three or six copies of a 120 amino acid, ~14 kD structural domain. Gelsolin and villin each have six

135 such domains which arose evolutionarily by first a gene triplication, followed by a duplication event resulting in two structurally similar but functionally non-equivalent halves of the protein. Villin has 45% amino acid sequence identity with gelsolin over the six modular domains (Finidori et al., 1992) suggesting that the tertiary structure is very similar. That villin can cross-link F-actin while gelsolin cannot is due to the presence of a small 76 amino acid head piece domain at villin’s C-terminus. Much of what is inferred about villin structure comes from comparison with gelsolin, its closest homolog. Gelsolin’s function is regulated by calcium binding at up to eight different sites. At submicromolar calcium concentrations, gelsolin exists in an autoinhibited state where the C-terminal G6 domain is folded internally such that the tail-helix interacts with G2 to sterically block G2 from binding to actin. Calcium binding releases this inhibition. Further activation steps involve domain rearrangements that release “latch” mechanisms between G1 and G3, to activate the G1 actin-binding site, as well as between G4 and G6 to allow G4 to participate in actin capping (Choe et al., 2002). Gelsolin targets the F-actin through binding of G2 to the side of the filament. Further interaction of G1 and G4 with the filament leads to severing and capping. The severing mechanism requires micromolar levels of calcium (Yin and Stossel, 1979). Unlike its structural homolog, gelsolin, structural data for villin is limiting. From early hydrodynamic and spectroscopic studies it is known that villin in solution undergoes a large conformational shift on binding calcium (Hesterberg and Weber, 1983b). The molecule is reported to become more asymmetric and overall length increases from 84 to 123 Å. These data led Hesterberg and Weber to suggest a “hinge-mechanism” where calcium binding to villin creates a localized tightening of the structure through increased α-helicity and a resultant outward shift of another domain (the head piece) into solution. More recent biochemical and functional studies point to two residues, based on sequence homology to gelsolin, Asp467 and Asp715 in G4 and G6, that potentially form salt-bridges in villin domain V2 (Kumar and Khurana, 2004). The solution NMR structure of V1, a.k.a. villin 14T, has been solved (Markus et al., 1994; Markus et al., 1997), revealing two potential Ca2+-binding sites and an overall fold that is very similar to that of gelsolin G1. The villin C-terminal actin-binding head piece domain has several structures solved, as it is a small, fast-folding protein (McKnight et al., 1997; Vardar et

136 al., 1999; Frank et al., 2004; Vermeulen et al., 2004; Meng et al., 2005). Cysteine scanning mutagenesis of this domain has provided comprehensive biochemical and functional data of the surface charge distribution of basic residues in this small domain and how they are presented to actin (Doering and Matsudaira, 1996). Much of the detail about the interaction of villin with F-actin has been inferred from the structures of gelsolin. Gelsolin’s G1 domain was crystallized with G-actin (McLaughlin et al., 1993). This structure revealed that G1 bound to actin’s hydrophobic pocket between actin subdomains 1 and 3. In this position G1 would cap the barbed end of a filament by preventing actin monomer addition. The N-terminal three domains, G1-G3, have also been crystallized with calcium and G-actin to indicate how these three domains rearrange to sever actin. This structure identifies the unique G2-G3 actin side-binding position (Burtnick et al., 2004) that competes with α-actinin for F-actin binding (Way et al., 1992b; McGough and Way, 1995). EM difference mapping of G2-G6 decorated F- actin localizes G2 binding on the side of F-actin, but the rest of the molecule is disordered because it is extended away from the filament axis (McGough et al., 1998). The C-terminal half, G4-G6, has similarly been crystallized with calcium and G-actin to show that G4 binds to G-actin in a similar fashion as G1 (Choe et al., 2002). The entire protein has been crystallized in the inactive, calcium-free state which revealed a compact globular form where G6 and G2 interact to form the “tail-helix latch” (Burtnick et al., 1997). The crystal structure of ATP-bound gelsolin has also been solved (Urosev et al., 2006). Villin requires calcium concentrations as high as 200 µM to activate severing. Such concentrations are not usually found in living cells, but might occur in apoptotic cells. Villin is found in microvilli that are responsible for absorption; so, it is feasible that villin has evolved to have a higher threshold to calcium for its severing activity. Also, no clear demonstration of villin existing in a globular, autoinhibited state exists. Villin has at least three calcium-binding sites (Hesterberg and Weber, 1983a); two in the core region and one in the head piece. It has also been shown that tyrosine phosphorylation of villin can reduce of even eliminate the calcium binding requirement for severing (Kumar and Khurana, 2004), suggesting that phosphorylation may be the primary regulator of villin conformation.

137 Here we employ electron tomography of F-actin-villin 2-D rafts preserved for electron microscopy in negative stain to gain a 3-D perspective on how villin interacts with and cross-links F-actin. We use a single-particle approach by extracting sub-volumes from the tomograms and classify these volumes using correspondence analysis to characterize any variation in villin conformation of F-actin cross-linking. We show that the villin-actin contacts are relatively homogeneous and yet inconsistent with contacts for other actin-binding proteins. Without calcium to activate it, villin can cross-link F-actin and does not disrupt the F-actin structure. We discuss the implication for alternative or multiple modes of F-actin binding and the relationship to villin function.

4.2 Results

4.2.1 Image alignment and volume classification Villin forms cross-links at regularly spaced intervals along F-actin on the lipid monolayer to form rafts ≥ 1 µm across. Six different tomographic tilt series images were aligned using marker-free alignment methods (Liu et al., 1995; Winkler and Taylor, 2006) and merged into tomograms. Data sets were merged after initial alignment to the actin filament. Resolution is estimated between 26 and 30 Å as judged by the computed power spectra. Figure 4.2.1.1 shows a representative central section. At first evaluation, the villin molecules appear as long, extended “squiggles” placed at regular intervals between the actin filaments, and not crossing over or under the filaments. Villin molecules and the adjoining actin filaments were extracted by hand using the EMAN boxer utility as 64 x 144 x 40 pixel boxes. Repeats were aligned in 3-D based on the actin filaments to produce the global average seen in Figure 4.2.1.2.

138 Villin

0.1 µm

Figure 4.2.1.1 A central z-section from a raw tomogram of villin cross-linking F-actin. Actin filaments are vertical and villin cross-links appear as small tick marks between the filaments.

139

Figure 4.2.1.2 Projection images. Left, projection image of the 3-D global average of 6,336 aligned villin cross-link motifs. Right, averaged projection image of ice embedded specimens, for comparison. Image provided by Greta Ouyang. Correspondence analysis was used to characterize the structural heterogeneity in the rafts. We characterized the heterogeneity in the actin filaments separately from the heterogeneity in the villin cross-links as part of the initial alignment to the actin filament. Classification was done based on the variance calculated in the region of the villin density, but excluding the invariant density. The 3-D binary mask is shown as individual z-sections in Figure 4.2.1.3. Areas in black are excluded from the multivariate statistical analysis. The resulting class averages produced by hierarchical ascendant methods (Figure 4.2.1.4) vary in several ways. In some cases there were vacancies for the villin density, two classes showed partial villin density, and one class revealed a shifted villin density due to misalignment of the actin filaments. The partial villin density could be attributed to negative staining effects, or could be a cleavage product since villin is susceptible to cleavage of its long inter-domain linker regions. These averaged villin densities were segmented out from their accompanying actin filaments using the Watershed transform as viewed in Figures 4.2.1.5 and 4.2.1.6. The helical parameters of the actin filaments were determined and an averaged actin filament density was created to eliminate any flattening due to the missing wedge of data that is not attainable due to limitations in sample tilt angle. The actin filaments have a 13/6 helical structure and are not expected to be perturbed due to interactions with villin, as seen for some actin-binding proteins such as cofilin (McGough et al., 1997).

140

Figure 4.2.1.3 3-D Classification Mask. Montage of sections through a 3-D classification mask based on the region of variance. Left side is bottom of 3-D volume and right side is the top of the 3-D volume. Note that the regions containing the relatively invariant average villin density are excluded from the statistical analysis for classification.

Figure 4.2.1.4 Class averages based on the variance within the region containing villin. Class averages 1 and 4, labeled on the image, show incomplete “half-villin” density. Averages 3 and 6 have villin vacancies. Class average 5 has the villin density shifted downwards due to misalignment of the filament. Class 0 has 5,704 repeat volumes while classes 1-13 have only 30-50 members each.

141 A

Figure 4.2.1.5 Segmented villin density maps. Taken from class averages shown in Figure 4.2.1.4. A, as viewed from the “front.” B, a backward tilted view of the crook. C, a forward tilted view of the two actin-binding domains. D, a 180 degree flip of A to show the back side of the density maps.

142 A

Figure4.2.1.6 Averaged density maps. A, F-actin: villin average density as seen from the front. B, same density flipped 180 degrees about the filament axis. C, villin density after segmentation.

4.2.2 Models We used the symmetrized actin filament volumes to fit a generated 13/6 atomic actin filament by rigid-body fitting using Situs (Wriggers et al., 1999). We used the atomic

143 coordinates for gelsolin domains 1-6 in the calcium- and actin-bound states (PDBs 1RGI and 1H1V) to dock into the villin average density by hand, along with the crystal structure for the villin headpiece (PDB 1YU5). Although these are technically gelsolin G1-6 domains, we will refer to them in this model as V1-6, by homology. There appears to be at least three contacts of villin with the actin filament. The two contacts at the top of the density are far enough apart to accommodate two domains, but are too large for V6 + head piece. These must necessarily be V1 and V2. Because all domains are separated by flexible linker domains, it is not possible to determine with certainty the orientation of V1 and V2. The small tail region then is occupied well by the villin head piece. As the N-terminal 35 residues of the head piece tend to become disordered due to low pH and protonation of a His residue, this may explain the thin crook of density we see leading to the F-actin interacting site. Alternatively, the tail helix from V6 could be a contributor. Perhaps the most curious aspect of the villin contacts with F-actin is that the densities are not like the predicted actin binding regions. The arrangement of V1 and V2 within the density indicates an interaction with actin subdomain 1 unlike those typical of gelsolin-like domains. G1 and V1 have been crystallized with G-actin and bind in a hydrophobic pocket between actin subdomains 1 and 3. However, binding here is characteristic of end-capping proteins, and in the case of cofilin, binding in this pocket changes the helical twist of the filament (McGough et al., 1997). The ABD of α-actinin binds to two adjacent actin protomers by contacting subdomain 1 and 2 on one actin protomer and subdomain 1 of the next protomer up the filament. This corresponds to actin residues 86-117 of the lower protomer and residues 350-375 of the upper protomer (Mimura and Asano, 1987; Lebart et al., 1990; Fabbrizio et al., 1993; Lebart et al., 1993; McGough et al., 1994). The gelsolin G2 domain has been demonstrated to have a similar actin interaction (McGough et al., 1998). Although the end-loops on the ends of the actin-binding helices in V1 and V2 are in close proximity to actin residues 358-365, the rest of the protein density is oriented back and away from the rest of the α-actinin/G2 binding region. Competition-binding assays have indicated the villin head piece can displace the binding of ADF (actin depolymerizing factor), which binds in the hydrophobic binding pocket. Again, the position of the probable head piece density does not reflect this type

144 of binding interaction. Instead, the density is positioned near two α-helices, one from actin subdomain 3 and one from subdomain 4. Exhaustive mutational analysis of the head piece has defined the key actin-interacting residues, which are located on the C- terminal helix facing towards the actin filament in Figure 4.2.2.2.

Figure 4.2.2.1 Model of villin cross-linking F-actin. Domains G1-G6 were taken from PDB coordinates excluding the flexible linker domains and the actin coordinates (PDBs 1RGI and 1H1V), and the head piece domain is the crystal structure for chicken villin, (PDB 1YU5). The actin filaments were fit to the symmetrized actin density using situs.

145 Figure 4.2.2.2 Close-up of 3 villin-actin interactions. A, The end-loop following the long helix of V2, residues 222-229 points to actin subdomain 1, residues 358-365. The actin hydrophobic binding pocket is shown in orange and marked by an asterisk in each model. B, Shown from the back side of the model, an end-loop in V1, residues 90-97, similarly approaches the same actin residues. C and D show front and back views of the relative position of the villin head piece. The head piece helix residues 62-76 are positioned towards actin and are in the vicinity of helices 221-235 in subdomain 4 and helix residues 308-320 on subdomain 3.

4.3 Discussion

The crystal structures of gelsolin’s domains bound to G-actin as well as the average density of G2-6 bound to F-actin have provided good structural insight as to how gelsolin binds, severs, and caps filaments to regulate the cytoskeleton. With the exception of the EM difference map of G2-6 decorated F-actin (McGough et al., 1998),

146 there has been no direct visualization of whole gelsolin bound to F-actin. Villin, a protein with 42% amino acid sequence identity to the gelsolin domains, has been assumed to behave in a somewhat similar fashion, although direct structural evidence is very limiting.

4.3.1 A tool for studying actin: actin-binding protein interactions We have been able to use the formation of paracrystalline arrays of macromolecular assemblies on a lipid monolayer to produce a sample of F-actin cross-linked with villin. These 2-D rafts of protein can be imaged in the electron microscope and analyzed as an ordered set of single particles. Alignment on the central, relatively invariant actin filament produces a data set for multivariate statistical analysis and classification based solely on the variance in the region of the villin cross-link.

4.3.2 Significance of villin-actin interaction sites The class averages based on the villin variance were surprisingly homogeneous. Only a small fraction of the repeats were classified as varying from the global average, and even these differences were small (Figures 4.2.1.4 and 4.2.1.5). Because of this homogeneity, the observed interactions with the filament are curious since they do not resemble the types of interactions proposed for villin’s close homolog, gelsolin. Figure 4.3.2.1 clearly demonstrates the drastic differences. The glancing contacts at the outermost edge of the actin protomers would suggest a weaker binding interaction. Because there are multiple weak binding interactions, the overall cross-linking function would be stabilized. As the model is constructed, the V1 and V2 interactions are both on the same α-helix of actin subdomain 1. This subdomain is the most acidic of the four. Placement of V1 and V2 in either orientation places end-loops in close proximity to this acidic helix. These loops contain Arg and Lys residues which may form an electrostatic association with actin’s negative charge. Although the acidic actin helix is also implicated in α-actinin-like side-binding to F-actin, this type of binding also requires interactions on the lower actin protomer that are obviously lacking here. The acidic actin helix is also implicated in an alternative gelsolin binding model proposed by Renoult et al. (Renoult et al., 2001). Their model, based on peptide binding assays, places G1 and G2 on the same actin monomer. Since G1 binds in the hydrophobic binding pocket for

147 severing and capping activities, G2 residues 159-193 would then bind to the outer actin surface to residues 119-132 and 347-375. This conflicts with the models for G2 binding which require contacts on two adjacent protomers. Our data supports and confirms the plausibility of this outer surface interaction proposed by Renoult et al., at least in the second actin binding region.

Figure 4.3.2.1 Villin and gelsolin binding sites do not overlap. Villin density map shown next to gelsolin crystal structures bound to actin. A, N-terminus of gelsolin bound to actin as in the crystal structure. Compare to villin contact modeled at right. G2 binds via its long α-helix between actin subdomains 1 and 3; G3 does not make contact with actin. G1 bound in this manner effectively caps the barbed end of the filament. G1-G3 on G-actin from PDB 1RGI. B, C-terminus of gelsolin bound to actin. G4 makes contact with actin subdomain 3. G5 and G6 are not bound to actin; however, the long α-helix of G6 is in close proximity to helices of actin subdomains 3 and 4. G4-G6 on G-actin from PDB 1H1V.

4.3.3 Indication for multiple modes of F-actin binding The villin interactions in these rafts may be governed by electrostatic interactions between the basic groups of residues on villin and the particularly acidic actin subdomain 1. One could hypothesize that in the presence of high calcium, the type II Ca2+ ions, which have been determined to be complexed in the villin/gelsolin-actin interaction site, could coordinate a tighter binding interface between actin subdomains 1

148 and 3 to facilitate a cleavage event. Evidence for such variability in actin-binding proteins is currently emerging from Real Space Iterative Helical Refinement studies of decorated actin filaments in the Egelman lab (Galkin et al., 2002; Galkin et al., 2003a; Cherepanova et al., 2006). They have demonstrated multiple modes of binding for cofilin, the CH domains of utrophin, as well as Xin and nebulin-like repeats. Other studies have identified multiple conformations of the actin filament itself induced by its interaction with binding proteins (McGough et al., 1997; Galkin et al., 2001; Galkin et al., 2003b; Galkin et al., 2005). Because our filaments average to have 13/6 helical parameters, we do not expect that villin is causing any disruption of the filament. Although at first glance the villin binding interactions may seem highly uncharacteristic, our data are actually supported by the concept of multiple modes of binding interactions. That the more than 6,000 villin cross-link repeats were so homogeneous in this interaction suggests it is not an exceptional case for actin interaction in this protein, but rather a legitimate mode of interaction.

4.4 Materials and Methods

4.4.1 Protein Purification and Array Formation Villin was purified from chicken intestines (Burgess et al., 1987). Actin was prepared from rabbit muscle acetone powder (Pardee and Spudich, 1982), followed by a Superose-12 column. Fresh G-actin was prepared by dialysis overnight against Buffer

A: 2 mM Tris-Cl, 0.2 mM Na2ATP, 0.02% β-mercaptoethanol, 0.2 mM CaCl2, 0.01%

NaN3, pH 8.0. The protein was clarified by high-speed centrifugation prior to sample set- up. Arrays were grown on positively charged lipid monolayers (Taylor and Taylor, 1994). A 3:7 volume ratio of DLPC: DDMA lipids (Avanti Polar Lipids) at 1 mg/mL in chloroform was layered over a solution containing 0.35 µM G-actin and 0.14 - 0.17 µM villin in phosphate-buffered polymerizing buffer without CaCl2 and with 1 mM EGTA, pH 6.5. The actin was polymerized in the presence of villin at 4 °C.

4.4.2 EM data collection The 2-D arrays were recovered using 200-300 mesh copper grids with a reticulated carbon support film (Kubalek et al., 1991). Samples were stained with 1% uranyl

149 acetate, dried, and stabilized by a thin layer of evaporated carbon. Data were collected on a Philips CM300-FEG. Six tomograms were recorded at 24k microscope magnification followed by 1.8 x post magnification at the CCD camera. The sample was tilted using a 2° Saxton scheme and recorded on a 2k x 2k CCD camera.

4.4.3 Image Processing Tomographic images were aligned using in-house software for marker-free alignment based on cross correlation of adjacent images in the tilt series (Winkler and Taylor, 2006). Repeat coordinates centered on each actin crossover from each tomogram were manually picked using the EMAN Boxer utility (Ludtke et al., 1999). 6,336 raw repeats were extracted as 64 X 144 X 40 pixel boxes. Images were then normalized by subtracting the mean and setting the standard deviation to 1. Repeats from each tomogram were aligned in 3-D to the actin filament. After the best alignments were attained, as judged by classification of the actin filament, the repeats from each tomogram were compiled into a single data set and aligned once more against the global average.

4.4.4 Classification Repeats were aligned to the central actin filament using in-house software for alignment. We used 3-D masks for volume classification. Classifications were done using 8 - 32 factors. Resulting Eigenimages were evaluated by eye and by plotting the variance and those containing only noise were left out of the classification. The resulting classes from each set were examined for completeness and resolution until it was determined that 16 factors gave the best classification result as judged by the detail seen in the averages.

4.4.5 Modeling Several relatively straight actin filaments were boxed out of the tomogram and Fourier transformed to determine the ratio of axial spacing of the 5.9 nm layer line to the first layer line. From this we established that the filaments had 13/6 helical symmetry. Using the global average density map of villin plus F-actin, we segmented out both the villin density and the F-actin density individually using the Watershed transform (Volkmann,

150 2002). The segmented 3-D average of F-actin was resampled to 12 pixels/55 Å repeat. Successive 12 pixel segments along the filament were then aligned by rotating each by the appropriate angle and then averaged together. The resulting average was then shifted and rotated back to create the averaged, symmetrized filament. This was done to minimize flattening effects due to drying of the specimen and to equalize the anisotropic, resolution due to the missing wedge. The PDB coordinates of an actin monomer were used to generate an atomic model of F-actin with 13/6 helical symmetry. This model was fit to the symmetrized density by hand using O (Jones et al., 1991) and then rigid-body fitting was done by Situs (Wriggers et al., 1999). Models of villin cross- links were created in O (Jones et al., 1991) using the gelsolin PDB coordinates excluding the flexible linker regions and the actin coordinates (PDBs 1RGI and 1H1V) and the villin head piece coordinates (PDB 1YU5). The atomic coordinates were fit to the villin volume in O as individual domains. Domains were placed keeping in mind that the missing linker domains must be capable of bridging the distance between C-terminal residue of one domain and the N-terminal residue of the next domain.The MMTSB tool set (Feig et al., 2004) was used for manipulating the PDB files. Final images were prepared with Chimera (Pettersen et al., 2004).

151 CHAPTER 5 SUMMARY AND FUTURE DIRECTION

The two F-actin cross-linking proteins characterized in these studies demonstrate the feasibility of combining approaches to data analysis. The highly variable α-actinin:F- actin arrays yielded structural data such as variation in the length of the cross-link as well as the novel observation of “monofilament” binding of α-actinin to a single actin filament. The villin:F-actin arrays were uniquely characterized in 3-D, a process generally not pursued due to the perceived missing wedge problem in alignment. From this data we establish that villin cross-links interact with F-actin in a manner quite distinct from those characterized for its very close homolog, gelsolin. That the length of the α-actinin cross-links can vary (33.9 ± 2.5 nm) combined with the quantized length variance (30.4 nm or 35.9 nm) seen in the monofilament binding suggests that α-actinin is a flexible cross-linker. Since there were no changes in the length of the rod domains, we believe that the extra length could be accounted for by the ABD-R1 linker region and/or variation in CH domain binding to F-actin. Perhaps most novel to this work is the monofilament binding of α-actinin, i.e. α-actinin cross-links F-actin crossovers. Because the monofilament binding events occurred with such high frequency, and the ABDs were placed regularly on F-actin, suggests that the binding is specific, not an artifact of sample preparation. That the cross-linking protein has evolved to have the same length as the F-actin cross-over repeat suggests a role in maintaining the helical parameters of the filament, while presenting α-actinin’s binding sites to transmembrane proteins such as integrin. We likewise describe unusual interactions between villin and F-actin. Villin has been assumed to share similar F-actin interactions with its close homolog, gelsolin. However; comparison of the 3-D volumes of villin cross-linking F-actin with the structures of gelsolin domains on actin reveals distinct differences. We narrow the binding sites on actin to a small helix at the N-terminus of subdomain 1, which is not an established mode of interaction for the many different F-actin-binding proteins. Although unusual, these interactions support recent research that reveals multiple modes of F-actin interaction.

152 Development of these procedures for negatively stained samples sets us up for future application to electron tomography of very low contrast, ice-embedded samples. In addition to probing the structures of other known cross-linking proteins between the filaments, we will also build upon the two component system, adding additional F-actin- interacting or α-actinin-interacting proteins, such as talin, to the equation. Additionally, addition of the β integrin tail to the α-actinin:F-actin rafts may provide insight to focal adhesion attachment to the cytoskeleton. Finally, new strategies for tilt series CTF correction are under development in our lab and this raises the quality expectations for new data sets. Thus, the utility of the lipid monolayer for tomography and image processing has yet to be exhausted.

153

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BIOGRAPHICAL SKETCH

Cheri M Hampton Program in Molecular Biophysics Kasha Laboratory of Biophysics Florida State University Tallahassee, FL 32306-4380 (850) 644-5606, office (850) 644-4104, lab (850) 644-7244, fax [email protected] Education

1990-1992 Biology Major Berry College, Rome, GA

1992-1995 BS Biology University of Georgia, Athens, GA

1998-2006 PhD Molecular Biophysics, Florida State University, Tallahassee, FL Advisor, 2002-present: Kenneth A Taylor Dissertation Title: Reading Between the Filaments: Structural Characterization of Two Different F-Actin Cross-Linking Proteins by Electron Microscopy. Professional Career

1990-1991 Student Work Opportunity with Biology professor Extracting fossils from matrix 1992 Berry College Externship Award recipient CDC, Atlanta, GA 1995-1998 Laboratory Technician, UGA College of Veterinary Medicine Physiology and Pharmacology Department Molecular Biology and Genetics 2000 Teaching Assistant, Prokaryotic Diversity Lab, FSU Biology Professional Memberships American Society for Cell Biology Conferences Attended FSU-Structural Biology: 25 Years of CryoEM of Biological Macromolecules January 8-9, 1999, Turnbull Conference Center, Tallahassee, Florida

Chlamydomonas 2000. Amsterdam, The Netherlands

171 Gordon Research Conference--Motile and Contractile Systems, 2003

Gordon Research Conference–Cell Contact and Adhesion, 2003

Cell Migration Consortium, May 14-16, 2004, Washington, D.C.

Gordon Research Conference–Signaling by Adhesion Receptors, 2004

Gordon Research Conference–Three Dimensional Electron Microscopy, 2005

American Society for Cell Biology, December 10-14, 2005. San Francisco, CA Posters Presented Gordon Research Conference–Signaling by Adhesion Receptors, 2004 “The Structural Regulation of Calcium-Sensitive α-Actinin-1.”

Gordon Research Conference–Three Dimensional Electron Microscopy, 2005 “Structures of 2D Rafts of F-actin and Bundling Proteins on Lipid Monolayers.”

American Society for Cell Biology, December 10-14, 2005. San Francisco, CA Published Abstracts American Society for Cell Biology, December 10-14, 2005. San Francisco, CA Peer Reviewed Publications Cheri M Hampton and Kenneth A Taylor. Alpha-actinin-2. AfCS-Nature Molecule Pages (2006). (doi:10.1038/mp.a000195.01).

Cheri M Hampton and Kenneth A Taylor. Alpha-actinin-3. AfCS-Nature Molecule Pages (2006). (doi:10.1038/mp.a000196.01).

Cheri M Hampton and Kenneth A Taylor. Alpha-actinin-4. AfCS-Nature Molecule Pages (2006). (doi:10.1038/mp.a000197.01).

Cheri M Hampton, Dianne W Taylor, Kenneth A Taylor. -Actinin is a variable length,variable polarity actin filament cross-linker. Submitted.

Cheri M Hampton, David DeRosier, Kenneth A Taylor. Structure of villin-actin cross- links. In Preparation.

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