Smooth Muscle Titin Interactions with Alpha-Actinin Richard Jue-Hsien Chi

Home , Smooth muscle

Florida State University Libraries

Electronic Theses, Treatises and Dissertations The Graduate School

2007 Smooth Muscle Titin Interactions with Alpha-Actinin Richard Jue-Hsien Chi

Follow this and additional works at the FSU Digital Library. For more information, please contact [email protected]

THE FLORIDA STATE UNIVERSITY

COLLEGE OF ARTS AND SCIENCES

SMOOTH MUSCLE TITIN INTERACTIONS WITH ALPHA-ACTININ

Richard Jue-Hsien Chi

A Dissertation submitted to the Department of Biological Science In partial fulfillment of the requirements for the degree of Doctor of Philosophy

Degree Awarded: Fall Semester, 2007

The members of the Committee approve the dissertation of Richard Jue-Hsien Chi defended on October 8, 2007

Thomas C.S. Keller, III Professor Directing Dissertation

Michael Blaber Outside Committee Member

Kenneth A. Taylor Committee Member

Piotr G. Fajer Committee Member

P. Bryant Chase Committee Member

Approved:

Timothy S. Moerland, Chair, Department of Biological Science

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

The research presented in this dissertation is dedicated to the memory of my mother, Lily Chi.

iii

ACKNOWLEDGEMENTS

I would like to thank the other members of the T. Keller lab for advice and encouragement, Margaret Seavy for help with protein purification, Scott Olenych and Rani Dhanarajan for help with molecular cloning, Kim Riddle for help with immunofluorescence microcopy, XiXi Jia for help with ultrathin cryosections, and Ewa Bienkiewicz for help with surface plasmon resonance and experimental design. This work was funded by grants from the FSU Cornerstone Program, National Institute of Health and American Heart Association. I would also like to thank the members of my committee for their time, expertise, insight and unwavering support. I would especially like to thank Tom Keller. Dr. Keller has not only been a great mentor to me throughout the program but also throughout my life development. He has happily held the capacity of mentor/teacher/role-model/accountant/father-figure/friend, and for that I will always be grateful. Finally, I would like to thank my family and friends; it was long and difficult journey but with you guys by my side, I had a ton of fun doing it!

TABLE OF CONTENTS

LIST OF FIGURES ...... vi ABSTRACT ...... viii

CHAPTERS

1. GENERAL INTRODUCTION...... 1

2. SMOOTH MUSCLE α-ACTININ INTERACTION WITH SM-TITN...... 10 Introduction ...... 10 Materials and Methods...... 12 Results ...... 16 Discussion ...... 19

3. SMOOTH MUSCLE TITIN ZQ DOMAIN INTERACTION WITH THE SMOOTH MUSCLE α-ACTININ CENTRAL ROD ...... 30 Introduction ...... 30 Materials and Methods...... 32 Results ...... 36 Discussion ...... 40

4. SUMMARY AND CONCLUSION ...... 56

REFERENCES ...... 61 BIOGRAPHICAL SKETCH ...... 66

LIST OF FIGURES

1. Three-filament model of the scarcomere and domain architecture of human soleus titin according to Labeit and Kolmerer (1995) ...... 2

2. Striated muscle titin interaction with α-actinin ...... 5

3. Schematic rendition of the organization of the cytoskeleton and contractile apparatus of the smooth muscle cell proposed by JV Small (1995)...... 6

4. Purposed model of parallel contractile units in smooth muscle tissue syncytium by Seow (2005)...... 7

5. Affinity of smooth muscle α-actinin for sm-titin-myosin coassemblies ...... 22

6. α-Actinin binds only to sm-titin in far western overlay analysis ...... 23

7. Assay of α-actinin actin-binding and triple-helical rod domain interactions with sm-titin ...... 24

8. Sites of α-actinin interaction with sm-titin mapped using expressed GST-α-actinin fragments ...... 25

9. Sites of α-actinin interaction with sm-titin mapped using a solid-phase protein-binding assay...... 27

10. Both the α-actinin R2-R3 triple-helical repeat and C-terminus domains are necessary to out compete native α-actinin for sm-titin binding...... 28

11. Interaction of smooth muscle and striated muscle α-actinins with sm-titin and titin -PIP2 dependence ...... 29

12. Smooth muscle titin RT-PCR products...... 43

13. Anti-TKZ reactivity of a high molecular weight band in smooth muscle extracts ...... 44

14. Localization of smooth muscle titin in cryosections of smooth muscle and cultured smooth muscle cells ...... 45

15. Model representation of the smooth muscle α-actinin central rod spectrin repeat-like R2 and R3 domains and key loop residues...... 47

16. Sm-titin Zq domain binding to wild type and mutated smooth muscle α-actinin R2-R3 domains in vitro...... 48

17. Solid Phase Binding Assay confirms a decrease in binding in mutated R2-R3 Loops...... 49

18. Predicted sm-titin Zq domain contains highly conserved residues...... 50

19. Sm-titin Zq domain binds to α-actinin R2-R3 domain with nanomolar affinity...... 51

20. Mutations in the sm-titin Zq domain significantly decrease affinity for the α-actinin R2-R3 domain...... 53

21. FPLC elution of purified Zq on Superdex 75 suggests smooth muscle titin Zq domain forms a dimer ...... 54

22. Proposed Model for titin Zq domain interaction with the α-actinin R2-R3 central rod domain...... 55

vii

ABSTRACT

The striated muscle Z-line is a complex network of proteins in which N-terminal domains of titin interact with the actin filament cross-linker α-actinin, and other proteins to establish and maintain the structural integrity of the sarcomere. Actin-myosin II filament-based contractile structures in smooth muscle and nonmuscle cells also contain the actin filament-crosslinking protein α-actinin. We previously discovered a titin isoform - originally named smitin, hereafter called sm-titin - in smooth muscle cells suggesting that similar interactions may exist in the α- actinin rich dense bodies and dense plaques that act as the smooth muscle equivalent of the Z- line. I have found that purified native smooth muscle α-actinin binds with nanomolar affinity to sm-titin in sm-titin-myosin coassemblies in vitro. Smooth muscle α-actinin also interacts with striated muscle titin. In contrast to striated muscle α-actinin interaction with titin and sm-titin,

which is significantly enhanced by PIP2, smooth muscle α-actinin interacts with sm-titin and titin

equally well in the presence and absence of PIP2. Using expressed regions of smooth muscle α- actinin, I have demonstrated sm-titin-binding sites in the smooth muscle α-actinin R2-R3 spectrin-like repeat rod domain and a C-terminal domain formed by cryptic EF hand structures. These sm-titin-binding sites are highly homologous to the titin-binding sites of striated muscle α- actinin. This suggests that sm-titin contains at least some of the domains found in titin that are known to bind to the α-actinin sites. In striated muscle Z-disks, titin N-terminal Z-repeat domains interact with the α-actinin EF hand region and the titin Zq domain interacts with the α-actinin central rod. RT-PCR analysis of RNA from various smooth muscle sources and western blot analysis with an N-terminal region-specific antibody presented here reveals that sm-titin contains the α-actinin-binding Z repeats Zr1, Zr2, Zr3, and Zr7 and the Zq domain encoded by the titin gene. I investigated whether the sm-titin Zq domain interacts with R2 and R3 spectrin repeat-like domain loops that lie in proximity on the surface of the smooth muscle α-actinin central rod. I found alanine and

viii

phosphomimetic mutations of α-actinin R2 and R3 loop residues decreased binding to expressed sm-titin Zq domain in GST-pull-down and solid phase binding assays. Likewise, surface plasmon resonance experiments revealed that alanine mutagenesis of a Zq domain region with high propensity to form α-helix decreased binding to the α-actinin R2-R3 region. Additionally, the Zq peptide migrates on FPLC size exclusion chromatography as an apparent dimer. I present a model of how the sm-titin Zq domain, which may form an anti-parallel homodimer, could interact with mirror image sites formed by R2 and R3 loops in the smooth muscle α-actinin central rod domain. Taken together, our results suggest that direct interaction between α-actinin and titin or titin isoforms is a common feature of actin-myosin II contractile structures in striated muscle and smooth muscle cells and that the molecular bases for α-actinin interaction with these proteins are similar, although regulation of these interactions may differ according to tissue.

CHAPTER 1 GENERAL INTRODUCTION

Striated muscle sarcomere organization The skeletal muscle cell contractile system consists of myofibrils composed of linearly arranged sarcomeres. The sarcomere is the structural functional unit of muscle contraction in both skeletal and cardiac muscle. The sarcomere structure has four major regions—the Z-line, I band, A band and M-line, with one sarcomere being defined as stretching from one Z-line to the next. Each sarcomere contains two prominent types of filaments: thick filaments composed of molecular motor myosin II and thin filaments containing actin, which together provide the molecular interactions that drive filament sliding. Actin thin filaments are anchored in the Z-line via the actin filament cross-linker α- actinin and extend into the I-band. The myosin II thick filaments, which form the A band, are anchored to the M-line via its interaction with myomesin. Near the center of the sarcomere, thin filaments interdigitate with the thick filaments in the AI zone, where the thick and thin filaments slide past each other while not changing in length. Two additional proteins, titin and nebulin, also contribute to the sarcomere structure and stability. Nebulin filaments associate with actin and regulate the length of the thin filaments by acting as a ruler (McElhinny, Kazmierski, Labeit, & Gregorio, 2003). Titin molecules, the third major set of filaments in the vertebrate striated muscle sarcomere, act as an elastic scaffold which centers the A-band in the middle of the muscle sarcomere and maintains the resting tension that allows muscles to snap back if overextended (Tskhovrebova & Makedonov, 2004). Muscle titin Titin is expressed as several isoforms, most of which are extremely large proteins. The complete sequence of the human titin gene (363 exons) and of several of the differentially 1

spliced titin gene transcripts expressed in vertebrate striated muscles are known (Bang et al., 2001). The longest striated muscle titin isoforms (~1.0 μm contour length molecules composed of ~28,000 amino acids) are elastic and span each half sarcomere, with anchorage points in the Z-disk and M-line and on the myosin filaments.

Figure 1. Three-filament model of the sarcomere and domain architecture of human soleus titin according to Labeit and Komerer (1995). The main section of I-band titin is made up of two structurally distinct regions, stretches of tandem-Ig modules and PEVK domain. The A-band portion of titin is composed of a super-repeat region composed of alternating Ig modules and FN III domains. The N-terminal region of titin molecule is anchored to the Z-line through interactions with the ‘Z-repeats’ and ‘Zq-domain’. The C-terminus interacts with M-line proteins to maintain its anchorage to the myosin thick filament. A unique kinase domain is located between the myosin binding and M-line binding domains.

Titin’s long structure contains multiple copies of five identified structural motifs, fibronectin III domains (FN III), immunoglobulin C2-like domains (IgC2), PEVK domains, Z- repeat domains and a kinase domain (Fig. 1). The amino terminal 80 kDa of each titin spans the entire Z-line structure of the sarcomere and interacts with α-actinin, T-cap/telethonin and actin, 2

whereas the carboxy-terminal region interacts with myosin, C-protein and M protein. By spanning the sarcomere and interacting with the thick filaments, the titin network maintains the position of each thick filament in the center of the sarcomere during contraction. Elasticity of the titin molecules also provides tension that resists overstretch of the passive sarcomere through reversible unfolding of the titin I-band PEVK region (Gregorio, Granzier, Sorimachi, & Labeit, 1999; Keller, III, 1997). It has been known for some time that titin filaments play important roles in sarcomere assembly, stability, and passive tension (Granzier & Labeit, 2004; Keller, III, 1995; Trinick, 1994; Tskhovrebova & Trinick, 2004). More recent investigations of titin elasticity and interactions with a variety of proteins confirm titin also plays roles in stretch sensing, protein turnover, and gene regulation (Lange et al., 2005; Tskhovrebova & Trinick, 2005).

Non-muscle titin isoforms Several non-muscle titin isoforms or c-titins have been found in the brush border cytoskeleton of chicken intestinal epithelial cells and human blood platelets. They have been shown to interact directly with myosin and α-actinin (Eilertsen, Kazmierski, & Keller, III, 1997) and have molecular composition originating from the striated muscle titin gene (Cavnar, Olenych, & Keller, III, 2007). Additionally, the existence of titin isoforms in smooth muscle has recently been confirmed (Kim & Keller, III, 2002; Labeit et al., 2006). Originally found in chicken smooth muscle cells, smooth muscle titin or smitin (now known as sm-titin), has been found to be a component of the smooth muscle contractile apparatus. Its molecular morphology also resembles that of striated titin and its function seems to parallel that of striated titin in its ability to organize myosin filaments and interact with α-actinin (Kim et al., 2002).

α-Actinin structure α-Actinin is a modular protein encoded by a family of genes. The protein products of these gene families consist of two calponin-homology domains at the N-terminus, which has the ability to bind actin, followed by four spectrin repeats in the central rod domain and four Calmodulin (CaM)-like EF-hands near the C-terminus. The CaM-like EF-hands have diverged to the point of losing their Ca2+ binding ability in striated and smooth muscle α-actinins and retaining it in a nonmuscle α-actinin (Dixson, Forstner, & Garcia, 2003). This serves to segregate

α-actinins into two distinct groups, muscle α-actinins with nonfunctional CaM-like EF-hands and nonmuscle α-actinins with functional CaM-like EF-hands. All α-actinins exist as a rod- shaped homodimer with a subunit molecular mass of 94-103 kDa. The antiparallel orientation of the units enables Ca2+ binding to the CaM-like EF-hands to inhibit nonmuscle α-actinin binding to and cross-linking actin. α-Actinins interact with CapZ, vinculin, zyxin and cell surface receptors in focal adhesions and cell-cell contacts such as the NMDA or integrin receptors, as well as with many signaling factors along the actin cytoskeleton. Interaction with the sarcomeric protein ruler titin localizes α-actinin to the Z disk region and specifies the number of crosslinks between overlapping actin filaments in myofibrillogenesis.

Titin binding to α-actinin In the Z-disk, interaction of two types of titin N-terminal domains with two different sites on α-actinin provides a key structural linkage required for the assembly and structural integrity of the vertebrate striated muscle sarcomere (Young, Ferguson, Banuelos, & Gautel, 1998). Disruption of this structural linkage by expression of a dominant-negative titin fragment causes sarcomere breakdown (Ayoob, Turnacioglu, Mittal, Sanger, & Sanger, 2000). One type of titin- α-actinin interaction involves binding any of seven differentially-spliced 45-residue titin Z- repeat (Zr) domains to a cryptic EF-hand motif in the C-terminal domains on each end of the α- actinin antiparallel dimer (Fig. 2A). The C-terminal end of α-actinin binds to the first (Zr1) and last (Zr7) titin repeats with nanomolar affinity. It binds to the other Z-repeats with millimolar affinities (Atkinson et al., 2000). The first and last repeats seem to be of greater importance, because they are found in all long striated muscle titins. The number of Z-repeats in titin varies between species and muscle types. This variation arises from alternative splicing and is correlated with the thickness of the Z- disk. A solution structure determination of this interaction revealed how the α-helical region of the titin Zr7 domain lies in a groove of the cryptic EF-hand motif of striated muscle α-actinin. Recently, Young and Gautel found a region of α-actinin to be a Z repeat-like pseudo-ligand (Young & Gautel, 2000). Interaction of this region of one α-actinin monomer with the C- terminal titin-binding site of the anti-parallel α-actinin blocks titin binding. This blocking is

relieved by binding of phosphatidylinositol-bisphosphate to the actin-binding domain of α- actinin (Fig. 2B, C).

Figure 2. Striated muscle titin interaction with α-actinin (A) Published model of striated muscle titin interaction with α-actinin in the Z-disk. Antiparallel α-actinin homodimers crosslink titin molecules through interactions between titin Z-repeat domains and C-terminal α-actinin domains and through an interaction between the titin Zq domain (red ball) and the central rod of α-actinin. (B, C) Model for the regulation of titin Z-repeat binding in α-actinin by phospholipids. (B) The closed or inactive state of the molecule when the EF3/4 region of the CaM-like domain interacts with a region between the ABD and R1 of the opposite subunit. Thus, binding of titin Z-repeats to EF34 is prevented. (C) Binding of PIP2 to the ABD induces a conformational change that switches on titin binding by making the EF3/4 region available for binding to Z- repeats (ZR7). Modified from Young and Gautel (Young et al., 1998)

The second type of interaction occurs between the striated muscle titin Zq domain (Fig. 2A, red ball) and the second and third (R2-R3) spectrin-like triple helical domains in the striated muscle α-actinin rod. The structure of this interaction remains very poorly characterized, in part because the entire R2-R3 region appears to be required for binding.

Smooth muscle cell contractile apparatus Smooth muscle cells (SMCs) use a highly organized actin-myosin II based contractile apparatus to generate contractile force for a variety of vascular and visceral smooth muscle activities; however, the contractile apparatus structural organization remains poorly understood. The current models suggest the smooth muscle cell contains thousands of actomyosin contractile 5

units, housed in a sarcomere-like arrangement maintained by α-actinin in the dense bodies and dense plaques. One contractile unit is defined as two dense bodies with actin (thin) filaments attached, and a myosin sidepolar (thick) filament lying between the parallel thin filaments (Small, 1995). In the smooth muscle cells, these contractile units are oriented obliquely relative to the longitudinal axis of the cell, so when stimulated to contract, uniform shortening occurs throughout the cell (Fig. 3).

Figure 3. Schematic rendition of the organization of the cytoskeleton and contractile apparatus of the smooth muscle cell proposed by JV Small (1995). Intermediate filaments (If) and nonmuscle actin filaments (n-m act) compose the ‘membrane’ cytoskeleton and the obliquely oriented muscle actin (m. act) and myosin filaments (my) would compose the ‘contractile’ cytoskeleton. Inset demonstrates the components of a smooth muscle contractile unit. Note its obliquely oriented actin filaments ending either at the dense body (db) or dense plaque (dpl) (if membrane terminal).

Small expanded the general model by reporting the existence of a membrane skeleton, which is attached to the cytoskeleton and provides the interface between the contractile machinery on the inside of the cell and the extracellular matrix on the outside to which force is transmitted. He proposed that the smooth muscle structural lattice has two distinguishable parts: the `cytoskeleton', the structural framework that surrounds and supports the contractile apparatus in the body of the cell, and the `membrane skeleton', the anchorage for the cytoskeleton and contractile apparatus at the cell surface (Fig. 3). The complete 'cytoskeleton' contains two major components: first, a complement of actin filaments that link the cytoplasmic dense bodies at equispaced intervals in longitudinal fibrils;

and second, a network of desmin intermediate filaments that co-distributes with the cytoskeletal actin (Small & Gimona, 1998). The actin filaments of the contractile apparatus are presumed to interface with the cytoskeleton at the cytoplasmic dense bodies and with the longitudinal rib-like arrays of dense plaques of the membrane skeleton that couple to the extracellular matrix. The ‘membrane skeleton’ of smooth muscle is distinguished by the presence of either dense plaques or caveolae. These are adhesion sites that are thought to be equivalent to the terminal Z-line in skeletal muscle and which provide anchorage for the actin filaments and form rib-like arrays all over the cell surface (Fig. 3). More recent investigations, which focused on smooth muscle as a bundle instead of a single cell, found no evidence for obliquely oriented contractile units or sidepolar myosin filaments short enough to be compatible with earlier models (Ali, Pare, & Seow, 2005). Moreover, Seow’s group believes the myosin filaments span the entire distance from dense body to dense body (or dense plaque if at terminal Z-line) and that contractile units are arranged in parallel rather than being obliquely oriented (Fig. 4).

Figure 4. Purposed model of parallel contractile units in tissue syncytium by Seow (2005). Schematic depiction of contractile unit arrangement in a bundle of smooth muscle cells viewed in a rectangular window. Note the contractile units are parallel the longitudinal axis and in some cases bind to the nucleus. (Ali et al., 2005)

They believe the parallel arrangement is accomplished by attachment of actin filaments to the nuclear envelope, making the nucleus a force transmitting structure (Fig. 4). Taken in context of a tissue syncytium, the nuclear attachment would provide enough force to account for uniform cell shortening. In addition, a thick filament spanning the whole contractile unit length (dense body to dense body) would decrease the amount of overlap between the thick and thin filaments in a manner that is exactly proportional to the distance between the dense bodies during cell shortening or contraction (Kuo & Seow, 2004).

Smooth muscle cell phenotypic plasticity Normal smooth muscle cells exhibit a remarkable degree of plasticity in being able to convert between a contractile state and a proliferative or 'synthetic' state in response to wound or atherosclerotic lesions (Halayko & Solway, 2001). In the contractile state, smooth muscle cells use the contractile apparatus containing large sidepolar filaments of smooth muscle myosin to produce contractile force for activities such as airway or arterial constriction (Somlyo, 1997). Synthetic smooth muscle cells migrate and divide to repair tissue damage. Under pathophysiological or carcinogenic conditions, synthetic state cells can form atherosclerotic plaques, airway or arterial wall thickening and metastatic tumors. Smooth muscle cells converting from the contractile state to proliferative state reorganize their contractile proteins and down regulate expression of specific proteins such as caldesmon, beta-tropomyosin and α-1 integrin; these proteins are known molecular markers that distinguish between the two states (Halayko et al., 2001). Reorganization also appears to involve disassembly of the large, sidepolar myosin filaments of the contractile system and reassembly of the myosin into stress fiber-like structures, similar to those found in many types of motile cells.

Potential role of smooth muscle titin Our discovery of the titin isoform sm-titin revealed an important new component of the smooth muscle contractile apparatus. Based on current knowledge of the numerous functions of titin in striated muscle sarcomeres, it is reasonable to predict that sm-titin plays major roles in the organization and physiology of the smooth muscle contractile apparatus. From the molecular composition and biochemical data we have obtained thus far, it has become even more evident

that sm-titin plays a functional role similar to that of striated muscle titin in organizing assembly and regulating stability of the dynamic contractile apparatus in SMCs through interactions with smooth muscle α-actinin, myosin II and other yet to be identified proteins (Kim et al., 2002; Chi, Olenych, Kim, & Keller, III, 2005). Smooth muscle titin also may play a major role in smooth muscle cell plasticity, because remarkably sm-titin and myosin filaments form macromolecular arrays resembling both the contractile apparatus and stress fiber-like structure when coassembled in vitro under different conditions (Kim et al., 2002). Therefore, a transition of sm-titin–myosin interaction from the organization of large sidepolar filaments in the contractile apparatus of the contractile phenotype to the stress fiber-like arrangement of small bipolar filaments may play a role in the change of the cell to the synthetic phenotype.

Sm-titin-α-actinin interaction Recently, we have determined that sm-titin interacts directly with α-actinin from smooth muscle, which could provide a structural linkage that is functionally similar to the linkage provided by titin in striated muscle sarcomeres. Therefore it is reasonable to predict a combination of sm-titin interactions with α-actinin may play a role in organization of the smooth muscle phenotype. This dissertation is focused on understanding the molecular basis for this interaction. Elucidating the exact molecular specificities of the sm-titin-α-actinin interaction could contribute to the long-term goal of understanding plasticity in smooth muscle cells and could provide crucial insight into the understanding of the smooth muscle cell contractile apparatus.

CHAPTER 2 SMOOTH MUSCLE α-ACTININ INTERACTION WITH SM-TITIN

Introduction

Vertebrate smooth muscle cells use an actin-myosin II-based contractile system to produce force for a variety of vascular and visceral smooth muscle activities, including gut peristalsis and blood pressure regulation. Although the major components of the smooth muscle cell contractile apparatus are known, the molecular basis for its organization remains poorly understood. Current models propose that the smooth muscle cell contractile apparatus shares certain structural and functional features with the striated muscle sarcomere. As in muscle sarcomeres, the smooth muscle contractile apparatus generates contractile force by myosin filament interaction with antiparallel actin filaments that are anchored through interaction with α-actinin to dense bodies, which appear to be the functional equivalents of Z-lines in sarcomeres (Cooke, 1983; Small, 1985; Bagby, 1986; Small, 1995; Small et al., 1998). In the striated muscle sarcomere, a third filament system based on the long, fibrous protein titin plays several structural roles, which include organizing assembly of new sarcomeres, maintaining the central position of the myosin filaments in contracting sarcomeres, and passively resisting overstretch of relaxed sarcomeres (Keller, III, 1995; Clark, McElhinny, Beckerle, & Gregorio, 2002; Granzier, Labeit, Wu, & Labeit, 2002; Tskhovrebova & Trinick, 2003). The functions of titin all depend on anchorage of its N-terminus to the Z-line through interactions with α-actinin, T-cap/telethonin, and actin. Two types of titin N-terminal domains interact directly with α-actinin: multiple ~45 residue titin Z-repeat domains interact with an α-actinin C- terminal domain and the titin Zq domain interacts with an α-actinin rod domain (Ohtsuka, Yajima, Maruyama, & Kimura, 1997; Young et al., 1998; Atkinson et al., 2000; Joseph et al., 10

2001). Overexpression of titin fragments containing one or more Z-repeats disassembles existing myofibrils and prevents formation of new myofibrils in cardiac muscle cells (Turnacioglu, Mittal, Dabiri, Sanger, & Sanger, 1997; Ayoob et al., 2000), demonstrating the importance of the α-actinin-titin linkage for sarcomere structure and stability. α-Actinin localization in smooth muscle dense bodies positions it where it could play a similar structural role in the smooth muscle contractile apparatus. Vertebrate α-actinins are encoded by a small family of genes (Virel & Backman, 2004). Despite some variation in sequence, all vertebrate α-actinin proteins are anti-parallel homodimers, in which each peptide has an N-terminal calponin-homology actin-binding domain, followed by four spectrin-like triple-helical repeats comprising a central rod domain, and four calmodulin (CaM)-like EF-hand structures near the C-terminus (Virel et al., 2004). Ca2+ binds to EF-hands in nonmuscle α-actinin, regulating its interaction with actin. Sequence differences in striated and smooth muscle α-actinin EF hand-like domains prevent Ca2+ binding (Virel et al., 2004). The two EF-hand-like domains closest to the C-terminus of striated muscle α-actinin (Act-EF34) form the site that interacts with titin Z-repeats (Atkinson et al., 2000; Atkinson et al., 2001; Joseph et al., 2001). Structural analysis has revealed that a helical region of the titin Z-7 repeat binds into the groove formed by the semi-open configuration of the Act-EF34 structure (Atkinson et al., 2001). In striated muscle α-actinin, an intramolecular interaction regulates this interaction with titin. Within the α-actinin anti-parallel homodimer, a Z repeat-like pseudoligand sequence located in the linker region between the N-terminal actin-binding domain and the first triple-helical repeat domain of each peptide binds to the Act-EF34 of the anti-parallel peptide blocking its interaction with titin (Young et al., 2000). Binding of phosphatidylinositol- bisphosphate (PIP2) to a site in the second Calponin-Homology (CH) domain of the N-terminal actin-binding region of α-actinin displaces the pseudoligand from the Act-EF34 domain, opening it for interaction with titin. Titin-like proteins that interact with myosin filaments in vitro exist in vertebrate nonmuscle and smooth muscle cells (Eilertsen & Keller, III, 1992; Eilertsen et al., 1992; Eilertsen, Kazmierski, & Keller, III, 1994; Kim et al., 2002). The titin-like protein isolated from intestinal epithelial cell brush borders interacts with α-actinin in vitro (Eilertsen et al., 1997). 11

This report presents evidence that the smooth muscle titin-like protein, sm-titin, also interacts with α-actinin and that the smooth muscle α-actinin domains mediating this interaction with sm- titin are similar to those in striated muscle α-actinin that interact with titin.

Materials and Methods

Native protein purification α-Actinin was purified from fresh chicken gizzards by a published procedure (Wenegieme, Babitch, & Naren, 1994), with an additional final purification by HPLC Mono Q chromatography. Smooth muscle sm-titin and myosin were isolated from chicken gizzard smooth muscle, and titin and skeletal α-actinin were isolated from cardiac chicken heart muscle, all by published protocols (Langer & Pepe, 1980; Kim et al., 2002; Pan, Damodaran, & Greaser, 1994).

Sm‐titin-myosin coassembly assay Sm-titin-myosin preparations were coassembled into previously characterized coassembly structures (Kim et al., 2002) by dialysis using Pierce Slide-Α-Lyzers (MWCO

10kDa)for 24 hrs into coassembly buffer (10 mM Imidazole, 2 mM MgCl2, 1 mM EDTA, 0.1 mM EGTA, 0.2 mM DTT, and 50 mM KCl, pH 7.0), at 4oC. Triton X-100 (2%) was included in the coassembly buffer in experiments containing GST-fusion proteins to aid solubility. The coassembly structures were pelleted by centrifugation at 12,000 x g for 1 minute in a microfuge. For analysis, the resulting pellets were resuspended in 1X SDS-PAGE sample buffer. Samples of the supernatants were made by mixing with a concentrated sample buffer stock. All samples were heated to 90 oC for 1 minute prior to electrophoresis. Electrophoresis was performed on high porosity 4-20% SDS-polyacrylamide gradient separating gels (Eilertsen et al., 1992; Kim et al., 2002). Polypeptide bands were visualized by staining with Coomassie Brilliant Blue R-250.

Native α-actinin interaction with sm-titin-myosin coassemblies Various concentrations of intact purified α-actinin were mixed with sm-titin and myosin and dialyzed overnight into coassembly buffer. Coassembly aggregates were pelleted by centrifugation at 12,000 x g for 1 minute, and the supernatant was removed. Samples of the supernatant and pellet fractions were made by resuspending/mixing with SDS sample buffer and heating to 90 oC for 1 minute. The amount of α-actinin bound to the sm-titin-myosin coassemblies was determined by densitometry (Gel Doc system, Quantity One software; Bio- Rad) of the Coomassie blue-stained protein SDS gels of the pellet fraction. The dissociation constant for α-actinin interaction with the sm-titin-myosin coassemblies was determined from the relationship between bound and free α-actinin plotted on a fitted curve (two-parameter fit, Sigma Plot). The mean dissociation constant was calculated from the dissociation constants determined for three separate experiments.

Native α‐actinin proteolytic fragment binding assay Gizzard α-actinin (1 mg/ml) in a buffer containing 40 mM ammonium acetate and 1.0 mM CaCl2 (pH 8.0) was treated with thermolysin (enzyme to substrate ratio of 1:25 (w/w) at 37oC for 2 hours, generating proteolytic fragments of α-actinin (Pavalko & Burridge, 1991). The 53 kDa and 27 kDa fragments were further purified by HPLC Mono Q chromatography and then independently added to mixtures of purified sm-titin and myosin or an equivalent amount of buffer, as a control. The mixtures were dialyzed overnight at 4 oC in coassembly buffer. The coassembled proteins were pelleted by centrifugation and analyzed by SDS-PAGE as described above.

GST-fusion protein binding and competition with native α-actinin binding Clones of smooth muscle α-actinin were generously donated by D.R. Critchley, Department of Biochemistry, University of Leicester, UK (ACTN1-ABD, ACTN1-R1-R4, and ACTN1-R3-814). Other clones generated by PCR spanned regions of the C-terminal domain (ACTN1-815-C and ACTN1-847-C) and various regions of the rod domain (ACTN1-R1-R3, ACTN1-R2-R3,). These clones were inserted into the pGEX-2T vector and expressed as GST- fusion proteins in BL21 (DE3) strain. The expressed GST-fusion proteins were purified using 13

Pierce GST Fusion Protein Purification Kit according to manufacturer’s recommendations. Expressed fragments were then dialyzed into coassembly buffer containing 2% Triton X-100, in the presence or absence of sm-titin and myosin. All samples were centrifuged at 12,000 x g for 1 minute. The pellets and supernatants were fractionated by SDS-PAGE as described above. In order to assess competition of GST-fusion protein competition with native α-actinin for binding to sm-titin-myosin coassemblies, sm-titin and myosin were coassembled in the presence of various combinations of the GST-fusion proteins GST-ACTN1-815-C and GST- ACTN1-R2-R3 and purified smooth muscle α-actinin in coassembly buffer containing 2% Triton X-100.

Far western overlay A sample of the sm-titin-myosin preparation used for coassembly studies was subjected to SDS-PAGE and electroblotted to nitrocellulose as previously described (Kim et al., 2002). The blot was blocked overnight with 5% (w/v) non-fat dry milk in basic buffer (20mM HEPES o (pH 7.5), 50 mM KCl, 10 mM MgCl2, 1 mM dithiothreitol, 0.1% Nonidet P-40) at 4 C, then incubated with expressed fragment GST-ACTN1-815-C at 4oC in basic buffer containing 1% (w/v) non-fat dry milk overnight. The blot was washed three times in phosphate-buffered saline containing 0.2 % Triton X-100 for 10 minutes and washed again with phosphate-buffered saline containing 0.2% Triton X-100 and 100 mM KCl. The blot was incubated with an anti-GST antibody (1:1000, Affinity BioReagents, Golden, CO) in incubation buffer (10% (w/v) nonfat dry milk, 1XPBS, 0.3% (v/v) Tween-20) for 1.5 hours and washed three times with 0.3% (v/v) 1XPBS-Tween-20 followed by secondary antibody (1:10,000, Sigma, St. Louis, MO) conjugated to horseradish peroxidase in incubation buffer for 1 hour. The chemiluminescent signal was developed using Amersham Biosciences ECL Plus western blotting detection system according to manufacturer’s recommendations.

Solid-phase protein-binding assay Microtiter plate wells were coated with purified sm-titin-myosin in 0.5 M KCl, 1 mM DTT, 10 mM Tris, pH 7.5 (Buffer A) at 4ºC overnight. After washing with 100 mM KCl, 3 mM

MgCl2, 10 mM Tris-HCl, pH 8.0 containing 0.5% Tween-20 (Buffer B), the wells were blocked

with Buffer B containing 1% BSA at room temperature for 4 hours. Control wells lacking sm- titin-myosin were blocked similarly with BSA. After three washes with Buffer B, the plates were incubated with α-actinin fragments; ABD-GST, R1-R3-GST, R2-R3-GST, R3-814-GST, 846-C-GST or GST alone in Buffer A containing 2% Triton X-100 for 3-4 hours at 4ºC. The plates were then washed with Buffer B and incubated with anti-GST antibody (1:500, Affinity BioReagents, Golden, CO) in Buffer B containing 0.1% BSA for 2 hrs at room temperature. Following washes with Buffer B, the secondary antibody conjugated to alkaline phosphatase (1:2500; Sigma A-3687) was added to the plates and incubated for 60 mins. After a final wash with buffer B and the plates were incubated for 10 mins in a substrate containing p-nitrophenyl phosphate (1mg/ml) for color development. The A405nm of each assay well was measured with a

µQuant instrument (BIO-TEK Instruments, Inc.) The A405nm for fragment binding to control wells was subtracted from the values for the fragment binding in wells containing sm-titin- myosin. The mean value and standard error was calculated from three separate experiments.

Effect of PIP2 on smooth and cardiac muscle α-actinin interactions with sm-titin and titin sm-Titin-myosin, cardiac titin, and smooth and cardiac muscle α-actinins were mixed in

various combinations and dialyzed into coassembly buffer. The effect of PIP2 (25 µM/ml) on α- actinin binding to sm-titin-myosin coassemblies was assayed with L-α-phosphatidylinositol-4, 5-

diphosphate (Sigma, St. Louis, MO) present in different micelle sizes. PIP2 micelles were prepared in 0.5% Triton X-100 by bath sonication for 15 minutes (Steimle, Hoffert, Adey, &

Craig, 1999). The PIP2 micelles were added to mixtures of α-actinin and coassemblies. Sm- titin-myosin coassemblies were pelleted at 12,000 x g for 1 minute. Cardiac titin and associated α-actinins were pelleted by centrifugation in a Beckman airfuge at 100,000 x g for 2 hours. The pellets and supernatants were separated by SDS-PAGE as described above. The affect (fold

difference) of PIP2 on α-actinin binding to the sm-titin-myosin coassemblies and cardiac titin was determined by densitometry (Gel Doc system, Quantity One software; Bio-Rad) of the Coomassie blue-stained protein in SDS gels of the pellet fraction in the presence and absence of

PIP2. The mean fold differences were calculated from the fold difference determined for three separate experiments.

Results

α-Actinin-sm-titin binding in vitro α-Actinin interaction with sm-titin was tested in vitro using cosedimentation assays with purified smooth muscle α-actinin and coassemblies of smooth muscle sm-titin and myosin II. Coassemblies of sm-titin and myosin II were used for these experiments because of the extreme difficulty of purifying sm-titin free of myosin II and the relative ease of pelleting sm-titin-myosin coassemblies for the cosedimentation assays. α-Actinin present during coassembly of sm-titin and myosin cosedimented with the sm-titin-myosin aggregates (Fig. 5, A). Varying the concentration of α-actinin present in the assay from 0.5-1.4 μM during the coassembly of sm- titin and myosin yielded little difference in the amount of α-actinin cosedimented with the sm- titin-myosin coassemblies. Significantly less α-actinin cosedimented with the sm-titin-myosin coassemblies when the total α-actinin concentration was <0.5 μM. The relationship between the amounts of α-actinin in the pellet as determined by densitometry of Coomassie-blue-stained SDS gels and the concentration of α-actinin present during the coassembly is consistent with that expected for α-actinin binding to a saturable number of distinct sites in the sm-titin-myosin coassemblies (Fig. 5, B). Three independent experiments yielded similar results with a mean dissociation constant of 272 ± 64(SE) nM α-actinin. Far Western gel overlay analysis confirmed that the α-actinin interacted with sm-titin and not the myosin in the sedimented coassemblies. For this assay, the proteins in the sm-titin- myosin preparation used for the coassembly cosedimentation assay were separated by SDS- PAGE and electroblotted to nitrocellulose. α-Actinin incubated with the strip of nitrocellulose bound only to the sm-titin band and failed to interact with the myosin heavy chain on the blot even though the amount of myosin heavy chain on the blot far exceeded the amount of sm-titin (Fig. 6).

The native α-actinin rod domain interacts with sm-titin To determine the domain or domains of α-actinin that mediate the interaction with sm- titin, two major proteolytic fragments were obtained by digestion of α-actinin with thermolysin.

These are referred to as the 53 kDa triple helical rod domain, which covers most of the four spectrin-like repeats, and the 27 kDa head domain comprising most of the N-terminal actin- binding domain. These α-actinin domains were tested for binding to sm-titin-myosin coassemblies (Fig. 7, lanes 1-12). Both native α-actinin and the 53 kDa triple helical region of α-actinin independently cosedimented with sm-titin-myosin coassemblies, suggesting that the smooth muscle α-actinin triple helical rod domain contains at least one sm-titin-binding site (Fig. 7, lanes 1, 5). The 27 kDa actin-binding domain failed to cosediment with the sm-titin-myosin coassemblies (Fig. 7, lane 9). Neither proteolytic fragment pelleted in the absence of the sm- titin-myosin coassemblies (Fig. 7, lanes 3, 7, 11).

α-Actinin rod and C-terminal domains bind sm-titin GST-fusion protein fragments spanning α-actinin were expressed to map the rod sm-titin- interacting domain at higher resolution and to determine whether the C-terminal region of α- actinin not present in the 27 kDa and 53 kDa domains also binds to sm-titin. The GST-fusion proteins expressed covered the N-terminus actin-binding domain (GST-ACTN1-ABD), the rod domains spanning spectrin-like repeats 1-3 (GST-ACTN1-R1-R3) and spectrin-like repeats R2- R3 (GST-ACTN1-R2-R3), the region containing the spectrin-like repeats 3 and 4 to the beginning of the C-terminal domain (GST-ACTN1-R3-814), the complete C-terminal domain (GST-ACTN1-815-C) and a truncated portion of the C-terminus (GST-ACTN1-847-C) (Fig. 8A). These GST-fusion proteins were expressed, purified to homogeneity (Fig. 8B), and individually added to sm-titin-myosin coassembly reactions. Two experimental approaches were used to assay binding of these fragments to sm-titin – cosedimentation and a solid-state binding assay. For the coassembly cosedimentation assay, the coassembly conditions were modified by the inclusion of 2% Triton X-100, which enhanced the solubility of the GST-fusion proteins and minimized pelleting of the proteins in the absence of the sm-titin-myosin coassemblies. The presence of the GST-α-actinin fragment in the cosedimentation pellet was determined by SDS-PAGE. Only the GST-fusion proteins containing spectrin-like repeats R2-R3 and/or residues 815-C-terminus of α-actinin cosedimented with the sm-titin-myosin coassemblies (Fig. 8, D, lane1; E, lane 1; G, lane 1). Fusion proteins lacking those regions failed to interact with the sm-titin-myosin coassemblies. 17

For the solid-state binding assay, the GST-fusion proteins were tested for interaction with sm-titin in sm-titin-myosin preparations immobilized in microtiter plates (Fig. 9). As in the cosedimentation assay, the α-actinin fragments containing the R2-R3 rod domain and the 815-C- terminus domain bound significantly better than the fragments not containing these regions of α- actinin.

The α-actinin R2-R3 spectrin-like repeat and C-terminus domains out compete native α- actinin for sm-titin binding In order to confirm that the R2-R3 spectrin-like repeat and C-terminus domains represent the sites mediating α-actinin interaction with sm-titin, GST-ACTN1-R2-R3 and GST-ACTN1- 815-C were tested independently and together for ability to compete with native α-actinin for interaction with sm-titin (Fig. 10). Independently, each of the two expressed domains failed to compete with the native α-actinin for sm-titin binding, even at great molar excess (Fig. 10 A, C; lanes 4). Together, however, the GST-ACTN1-R2-R3 and GST-ACTN1-815-C domains displaced the native α-actinin from binding to sm-titin; virtually all of the native α-actinin remained in the supernatant in the cosedimentation assay (Fig. 10, D; lane 2).

Effects of PIP2 on interactions of α-actinins with cardiac muscle titin and smooth muscle sm-titin A cosedimentation assay using purified cardiac striated muscle α-actinin and titin confirmed that α-actinin interacts with native titin and that PIP2 enhances the interaction of these striated muscle proteins 2.2 ± 0.2 (SE, n=3) fold (Fig. 11, A; lanes 2 and 4, and 6 C). Interestingly, the cardiac muscle α-actinin interacted with the sm-titin-myosin coassemblies, and

PIP2 also enhanced this interaction by 2.05 ±0.13 (SE, n=3) fold (Fig. 11, B; lanes 6 and 8, and 6

C). In contrast, PIP2 failed to enhance and even slightly inhibited the interaction of purified smooth muscle α-actinin with the sm-titin-myosin coassemblies (Fig. 11, A; lanes 6 and 8, and 6

C) or with the cardiac muscle titin (Fig. 11, B; lanes 2 and 4, and 6 C). Differences in PIP2 sensitivity suggest that the molecular mechanism regulating smooth muscle α-actinin-sm-titin interaction differs from that regulating the extremely stable striated muscle α-actinin-titin interaction. 18

Discussion

Although the overall organization of the smooth muscle cell is complex and remains poorly understood, organization of the smooth muscle contractile apparatus is thought to have parallels with that of the striated muscle sarcomere. In both striated and smooth muscle contractile systems, myosin filaments produce force on oppositely oriented actin filaments that are anchored to one of a pair of functionally similar α-actinin-containing structures - the sarcomere Z-disk or the smooth muscle dense body (or dense plaque) - to pull the structures closer together. In striated muscles, a third filament system composed of the protein titin establishes and maintains sarcomere organization and integrity. Our discovery of the titin-like protein sm-titin in the contractile apparatus of smooth muscle cells raised the possibility that it plays roles similar to those of titin in the striated muscle sarcomere. The work presented here supports that notion by demonstrating a high degree of similarity in smooth muscle α-actinin- sm-titin and striated muscle α-actinin-titin interactions, with one significant difference. Our data indicate that smooth muscle α-actinin interacts with sm-titin with nanomolar affinity and has two sm-titin-binding sites in molecular locations similar to those of the two titin- binding sites in striated muscle α-actinin. We found that, as in striated muscle α-actinin, one of the smooth muscle α-actinin sm-titin-binding sites is in the R2-R3 triple helical repeat region. This region of striated muscle α-actinin interacts with the Zq domain of titin (Young et al., 1998). The molecular basis for interaction of either the striated or smooth muscle α-actinin rod domains with titin or sm-titin remains unknown because neither binding site has been mapped at higher resolution. It is interesting to speculate that the loop regions of the R2 and R3 triple helical domains that are located in proximity to each other in models of both the striated and smooth muscle α-actinin rods may participate in the interaction with titin and sm-titin (Ylanne, Scheffzek, Young, & Saraste, 2001; Liu, Taylor, & Taylor, 2004). The other titin/sm-titin-binding site is located at the C-terminal end of both striated muscle and smooth muscle α-actinins. This site in both types of α-actinin is composed of two cryptic EF-hand domains, neither of which binds Ca2+ in the native protein. The solution structure of a complex between the striated muscle C-terminal EF hand domain (Act EF34) and the titin Z7 repeat has revealed the pattern of intermolecular connectivities (Joseph et al., 2001). 19

All thirteen of the striated muscle α-actinin residues contacting the titin Z7 domain are perfectly conserved in smooth muscle α-actinins from chicken to man. This sequence similarity along with our data demonstrating that striated muscle α-actinin interacts with sm-titin and smooth muscle α-actinin interacts with striated muscle titin suggests that the molecular basis for the C- terminal end of smooth muscle α-actinin interacting with sm-titin is similar to that for striated muscle interaction with titin. Interaction of both striated and smooth muscle α-actinins with both titin and sm-titin also suggests that sm-titin contains α-actinin-binding motifs similar to both the titin Z repeat and Zq sequences. Confirmation of whether sm-titin contains such sequences awaits availability of sm-titin sequence. Regardless of the molecular basis for interaction, the smooth muscle α-actinin C-terminal and rod domains appear to interact independently with sm-titin. Neither domain alone effectively competes for binding of intact α-actinin to sm-titin. The presence of both domains is required to out compete native α-actinin for binding to sm-titin in the coassembly cosedimentation assays. This is consistent with an existing model of striated muscle α-actinin with striated muscle titin (Young et al., 1998). The importance of two different and independent titin-binding and sm-titin-binding sites on α-actinin molecules remains to be determined but may be related to the intricacies of regulating assembly of the striated muscle sarcomere and smooth muscle contractile apparatus. Despite the apparent similarities in interaction of the striated and smooth muscle α- actinin with titin and sm-titin, our data indicate that there is a distinct difference in PIP2 regulation. PIP2 enhances the interaction of striated muscle α-actinin with both titin and sm-titin but not the interaction of smooth muscle α-actinin with either sm-titin or titin. A model for how

PIP2 regulates striated muscle α-actinin interaction with titin proposes that intramolecular interaction of a Z-repeat pseudoligand domain located in the sequence connecting the CH2-actin- binding and R1-triple helical domain in one peptide with the C-terminal titin-binding domain on the antiparallel peptide blocks it interaction with titin Z repeats (Young et al., 2000). Binding of

PIP2 to a site in the CH2 domain relieves this autoinhibition by displacing the pseudoligand from the C-terminal titin-binding site (Young et al., 2000).

Why PIP2 has little effect on binding of native smooth muscle α-actinin to sm-titin or titin remains unclear. Direct comparison of the striated and smooth muscle α-actinin sequences yields few clues. The PIP2-binding and proposed pseudoligand sequences are essentially identical in vertebrate striated and smooth muscle α-actinins. Indeed, PIP2 binds to smooth muscle α-actinin and therefore could regulate smooth muscle α-actinin-sm-titin interaction via a similar mechanism (Fukami, Sawada, Endo, & Takenawa, 1996). One clue becomes apparent, however, from analysis of a recent structural model of smooth muscle α-actinin (Liu et al., 2004). In this model, the two ends of the smooth muscle α-actinin have different structures; neither of the two C-terminal sm-titin-binding domains is positioned well to interact with the homologous smooth muscle α-actinin titin Z-7 repeat pseudoligand domain without significant distortion of the protein structure. Perhaps minor sequence variations between striated and smooth muscle α-actinins impart different structural constraints that allow interaction between the striated muscle α-actinin C-terminal and pseudoligand domains but prevent a similar interaction in smooth muscle α-actinin. Structural constraints in smooth muscle α-actinin therefore could maintain the C-terminal domain unblocked for sm-titin or titin interaction even in the absence of PIP2. This raises the possibility that there is an alternative mechanism for regulating interaction between smooth muscle α-actinin and sm-titin. Smooth muscle cell phenotypic and mechanical plasticity involves contractile apparatus reorganization. It is reasonable to assume that regulation of smooth muscle α-actinin-sm-titin interaction plays important roles in both types of plasticities.

Figure 5. Affinity of smooth muscle α-actinin for sm-titin-myosin coassemblies. Various known concentrations (given above lanes) of native α-actinin were dialyzed with sm-titin and myosin into a solution containing 50 mM KCl. Coassembly structures were pelleted by centrifugation. Resulting supernatant and pellet fractions were subjected to SDS-PAGE (A). The amount of α-actinin bound to the sm-titin-myosin coassemblies was determined by densitometry of the Coomassie blue-stained gels. The dissociation constant for α-actinin interaction with the sm-titin-myosin coassemblies was determined from the relationship between bound and free α-actinin plotted using a fitted curve (B, Sigma Plot two-parameter fit). The results of one experiment that yielded a dissociation constant of 375 nM are shown (A and B). A mean dissociation constant of 272 ± 64 nM (SE) was calculated from the dissociation constants determined for three separate experiments.

Figure 6. α-Actinin binds only to sm-titin in far western overlay analysis. Proteins in the sm-titin-myosin preparation used for the coassembly cosedimentation assay were separated by SDS-PAGE and electroblotted to nitrocellulose. The strip of nitrocellulose was incubated in the presence of GST-ACTN1-815-C and probed with an antibody for GST. The α-actinin bound only to the region of the blot containing sm-titin and failed to interact with the much greater amount of myosin heavy chain or other contaminating proteins, which appear to be proteolytic fragments of myosin that are enriched by the blotting procedure. SmT, sm-titin; MHC, myosin heavy chain, α-A, intact α-actinin, MLC, myosin light chains.

Figure 7. Assay of α-actinin actin-binding and triple-helical rod domain interactions with sm-titin. Native intact α-actinin and α-actinin actin-binding (27 kDa) and triple-helical rod (53 kDa) domains generated by thermolysin partial proteolysis of isolated native protein were dialyzed alone or mixed with sm-titin and myosin under coassembly conditions and centrifuged. The pellet and supernatant fractions were subjected to SDS-PAGE and the gels stained with Coomassie blue. Lanes 1 and 2, pellet and supernatant of sm-titin-myosin coassemblies containing intact α-actinin. Lanes 3 and 4, pellet and supernatant of control sample of intact α- actinin alone. Lanes 5 and 6, pellet and supernatant of sm-titin-myosin coassemblies containing the triple-helical (53kD) rod region of α-actinin. Lanes 7 and 8, pellet and supernatant of control sample of the triple-helical (53kD) rod region of α-actinin alone. Lanes 9 and 10, pellet and supernatant of sm-titin-myosin coassemblies containing the actin-binding (27kD) region of α- actinin. Lanes 11 and 12, pellet and supernatant of control sample of the actin-binding (27kD) region of α-actinin alone. Lanes 13 and 14, pellet and supernatant of control sample of coassembled sm-titin-myosin without α-actinin or fragments. SmT, sm-titin; MHC, myosin heavy chain; α-A, intact α-actinin; 53 kDa, triple helical rod region of α-actinin, 27 kDa, actin- binding region of α-actinin.

Figure 8. Sites of α-actinin interaction with sm-titin mapped using expressed GST-α- actinin fragments. GST fusion fragments encompassing all regions of the full length α-actinin were expressed and purified to homogeneity (A, B). The GST-α-actinin fusion fragments were dialyzed alone or mixed with sm-titin and myosin, dialyzed under coassembly conditions, and centrifuged. The pellet and supernatant fractions were subjected to SDS-PAGE and the gels stained with Coomassie blue (C-I). Shown are pellet and supernatant fractions of sm-titin- myosin coassemblies containing GST-ACTN1-ABD (C, Lanes 1 and 2), GST-ACTN1-R1-R3 (D, Lanes 1 and 2), GST-ACTN1-R2-R3 (E, Lanes 1 and 2), GST-ACTN1-R3-814 (F, Lanes 1 and 2), GST-ACTN1-815-C (G, Lanes 1 and 2), GST-ACTN1-847-C (H, Lanes 1 and 2), or a GST control (I, Lanes 1 and 2). Also shown are pellet and supernatant fractions of control samples containing only GST-ACTN1-ABD (C, Lanes 3 and 4), GST-ACTN1-R1-R3 (D, Lanes 3 and 4), GST-ACTN1-R2-R3 (E, Lanes 3 and 4), GST-ACTN1-R3-814 (F, Lanes 3 and 4), GST-ACTN1-815-C (G, Lanes 3 and 4), GST-ACTN1-847-C (H, Lanes 3 and 4), or GST (I, Lanes 3 and 4).

Figure 9. Sites of α-actinin interaction with sm-titin mapped using a solid-phase protein- binding assay. GST-fusion fragments of smooth muscle α-actinin and GST alone were added to microtiter wells containing sm-titin and myosin. Anti-GST antibody and a secondary antibody conjugated to alkaline phosphatase were used to detect the binding of the GST-fusion fragments. Shown is the mean A405nm (±SE, n=3) for each of the following fragments: (A) ABD-GST; A405nm 0.046 ± 0.006, (B) R1-R3-GST; A405nm 0.150± 0.02, (C) R2-R3-GST; A405nm 0.075± 0.012, (D) R3-814-C-GST; A405nm 0.013 ± 0.003, (E) 815-C-GST; A405nm 0.070 ± 0.003, (F) 846-C-GST; A405nm 0.030 ± 0.011, (G) GST alone; A405nm 0.002 ± 0.009.

Figure 10. Both the α-actinin R2-R3 triple-helical repeat and C-terminus domains are necessary to out compete native α-actinin for sm-titin binding. GST-ACTN1-R2-R3, GST- ACTN1-815-C, native α-actinin, and GST alone, each were dialyzed into 50 mM KCl coassembly buffer with or without sm-titin and myosin and centrifuged. Coassembly structures and control samples of fusion proteins alone were pelleted by centrifugation (A-D). Shown are pellet and supernatant fractions of sm-titin-myosin coassemblies containing native α-actinin (A- C, lanes 2 and 3, and D, lanes 4 and 5), native α-actinin and GST-ACTN1-815-C (A, lanes 4 and 5), native α-actinin and GST (B, lanes 4 and 5), native α-actinin and GST-ACTN1-R2-R3 (C, lanes 4 and 5), or native α-actinin, GST-ACTN1-815-C, and GST-ACTN1-R2-R3 (D, lanes 2 and 3). Also shown are pellet and supernatant fractions of control samples containing only native α-actinin (A-D, lanes 6 and 7), GST-ACTN1-815-C (A, lanes 8 and 9; D, lanes 10 and 11), GST (B, lanes 8 and 9), or GST-ACTN1-R2-R3 (C-D, lanes 8 and 9). Chicken pectoralis muscle extract standard (A-D, first lane). SmT, sm-titin; MHC, myosin heavy chain; α-A, native α-actinin.

Figure 11. Interaction of smooth muscle and striated muscle α-actinins with sm-titin and titin -PIP2 dependence. Intact cardiac and smooth muscle α-actinins were coassembled separately with sm-titin and myosin and cardiac titin in the presence and absence of PIP2. Coassembly structures and control samples of native α-actinin and isolated sm-titin and titin were pelleted by centrifugation (A-B, D-E). Striated muscle α-actinin bound to titin (A, lane 2) and to coassemblies of smooth muscle sm-titin-myosin (B, lane 6). These interactions were enhanced by PIP2 (A, lane 4 and B, lane 8) 2.2± 0.1 (SE, n=3) and 2.02± 0.13 (SE, n=3) fold, respectively (C). Smooth muscle α-actinin bound to titin and coassemblies of smooth muscle sm-titin and myosin in the absence of PIP2 (A, lane 6 and B, lane 2) and were not enhanced by PIP2 (C). The mean fold differences for smooth muscle and cardiac α-actinins binding to titin and sm-titin were calculated from the fold differences determined from three separate binding experiments. Chicken pectoralis muscle extract standard (A-D, lane 1). SmT, sm-titin; MHC, myosin heavy chain, α-A, α-actinin.

CHAPTER 3 SMOOTH MUSCLE TITIN ZQ DOMAIN INTERACTION WITH THE SMOOTH MUSCLE α-ACTININ CENTRAL ROD

Introduction

Smooth muscle cells use an extensively organized but poorly understood actin-myosin contractile apparatus to generate force for physiological requirements such as blood pressure regulation, airway constriction, and gut peristalsis. Dynamic changes in the organization of this contractile apparatus support a remarkable degree of smooth muscle cell mechanical and phenotypic plasticity in response to physiological cues (Gerthoffer & Gunst, 2001; Seow, Pratusevich, & Ford, 2000). Mechanical plasticity enables a smooth muscle cell to adapt to chronic or transient changes in cell length or load by reorganizing its contractile apparatus to maintain maximal force generating capability (Ford, Seow, & Pratusevich, 1994; Seow et al., 2000). Phenotypic plasticity permits a smooth muscle cell to convert reversibly from a ‘contractile’ state, in which it can generate force for contraction, to a ‘synthetic’ state, in which it can proliferate and migrate to participate in angiogenesis and wound healing or pathological formation of atherosclerotic plaques and thickening of arterial walls (Nakamura, Isoyama, Watanabe, Katoh, & Sawai, 1998; Newby & Zaltsman, 2000; Orford, Selwyn, Ganz, Popma, & Rogers, 2000; Zanellato et al., 1990). Conversion between the contractile and synthetic phenotypic states involves changes in the expression and organization of smooth muscle contractile apparatus proteins (Halayko et al., 2001). Some of these changes have been defined, but most of the underlying structural and molecular mechanisms supporting smooth muscle contractile apparatus organization and its plasticity have yet to be elucidated.

Although there are significant structural and stability differences, certain properties of the smooth muscle contractile apparatus organization resemble those of the better-understood striated muscle sarcomere (Herrera et al., 2005) Both striated and smooth muscle contractile systems contain antiparallel arrays of actin filaments, each of which anchors to one of a pair of α-actinin-containing functionally homologous structures - the striated muscle sarcomere Z-disk or the smooth muscle dense body. The myosin filaments – large bipolar filaments in striated muscles and large sidepolar filaments in smooth muscle - are positioned between the ends of the anti-parallel actin filaments, where both types of myosins use similar crossbridge cycles of force production on actin filaments to pull the Z-disks and dense bodies closer together. In striated muscles, a third major set of filaments, each of which is a molecule of the protein titin, organizes and stabilizes the sarcomere structure. Complete sequences of the human titin gene and of several differentially spliced titin gene transcripts expressed in vertebrate striated muscles are known (Bang et al., 2001). The longest striated muscle titin isoforms (~1.0 μm contour length molecules composed of ~28,000 amino acids) are elastic and span each half sarcomere, with anchorage points in the Z-disk and M-line and on the myosin filaments. Titin filaments play important roles in sarcomere assembly, stability, and passive tension, as well as in stretch sensing, protein turnover, and gene regulation (Lange et al., 2005; Tskhovrebova et al., 2005; Linke, 2007) through interactions with other proteins including α-actinin. We found a long isoform of the protein titin (originally named smitin, hereafter called sm-titin) in the smooth muscle contractile apparatus (Kim et al., 2002). We demonstrated that this sm-titin interacts in vitro with smooth muscle myosin (Kim et al., 2002) and smooth muscle α-actinin (Chi et al., 2005). The existence of titin isoforms in smooth muscle recently has been confirmed by others (Labeit et al., 2006). Smooth muscle α-actinin is one of several isoforms of α-actinin encoded by a family of genes, which are differentially expressed in different tissues (Dixson et al., 2003) All α-actinin monomers are composed of an actin-binding domain at the N-terminus, four spectrin-like triple helical repeat regions (R1-4) in the central rod domain, and four C-terminal end cryptic calmodulin (CaM)-like EF-hands (EF1-4), which in striated and smooth muscle isoforms fail to bind Ca2+. All α-actinins exist as rod-shaped anti-parallel homodimers (Fig. 2B).

In the Z-disk, interaction with the sarcomeric protein titin localizes α-actinin to the Z disk region and specifies the number of crosslinks between overlapping actin filaments. Two types of titin N-terminal domains interact with two different sites on α-actinin to provide this key structural linkage, which is required for the assembly and structural integrity of the vertebrate striated muscle sarcomere (Young et al., 1998). Disruption of this interaction by expression of a dominant-negative titin fragment causes sarcomere breakdown (Ayoob et al., 2000). One type of titin-α-actinin interaction involves binding any of seven differentially-spliced 45-residue titin Z- repeat (Zr) domains to the EF34 set of cryptic EF-hand motifs in the C-terminal domains on each end of the α-actinin antiparallel dimer (Young et al., 1998)(Fig. 2A). A solution structure determination of this interaction revealed how the α-helical region of the titin Zr7 domain lies in a groove formed by the EF34 domain in striated muscle α-actinin (Atkinson et al., 2000). We have confirmed that the sm-titin Zr7 domain interacts with similar cryptic EF hands near the C- terminus of smooth muscle α-actinin, but regulation of this interaction appears to be different in striated and smooth muscles (Chi et al., 2005). A second type of interaction occurs between the striated muscle titin Zq domain and the second and third (R2-R3) spectrin-like triple helical repeats in the striated muscle α-actinin rod (Young et al., 1998)(Fig. 2A). The molecular basis for this interaction remains very poorly characterized, in part because the entire R2-R3 region appears to be required for binding. Here, we report characterization and molecular mapping of a similar interaction between the sm-titin Zq domain and the R2-R3 region of the smooth muscle α-actinin rod. Based on evidence presented, we propose a model for how Zq domain interaction with loop regions of both R2 and R3 and with itself may contribute to the sm-titin-α-actinin linkage in smooth muscle.

Materials and Methods

Smooth muscle RT-PCR analysis RT-PCR analysis of commercially available human carotid artery, bladder, and uterus RNAs (Stratagene) and TRIZOL-isolated RNA from the A7r5 rat aorta smooth muscle cell line was used to determine whether a smooth muscle titin isoform contained known α-actinin-

binding domains and the sequence of that region. Primers complementary to human and rat titin gene sequence encoding this region were designed using PrimerQuest (Integrated DNA Technologies). First strand cDNA was synthesized using SuperScript III (Invitrogen) and primers complementary to the sense strand of exons 20 and 28. Subsequent PCR amplification was done using forward and reverse primer pairs covering sequence in the 5’ end of human titin exons 1, 8, 20 (forward) and the 3’ end of exons 14, 20, 28 (reverse). The standard PCR protocol used included 35 cycles of denaturation for 30 seconds at 94oC, annealing for 30 seconds at 0.5oC below the lowest primer calculated melting temperature and extension for 1 minute per kb of possible product sequence at 68oC. The length of products was assessed compared to standards on an agarose gel and purified using a gel extraction kit (QIAGEN). Purified products were sequenced in both directions using the PCR amplification primers and analyzed for exon- exon junctions to insure mRNA origin. Forward primers used were from exon 1 - 5'CGTTTCAGAAGCAACCTTGGGCTT, exon 8 - 5’ACTGCTGTGCACATCCAACCTGCT, and exon 20 - 5’ACCTGCCGCGCCTTACTTTATTAC. The reverse primers used were from exon 14 - 5'TGACTGCTTTAGGGACAACGTGGG, exon 20 - 5'CCTGCTTGTTCCTCTGTGAGGCTA, and exon 28 - 5’TACCAGTTGACTTTGGGCTGAGGGT.

Fusion protein expression and mutagenesis The human sm-titin Zq domain was amplified from human carotid RNA using primers designed against the 5’ end of titin exon 15 and the 3’ end of titin exon 16, cloned into pET-15b

vector (Invitrogen), and expressed as a C-terminal His6- tagged fusion protein in BL-21 (DE3) E. coli. After Ni-NTA agarose (Pierce, Rockford, Ill.) affinity chromatography, HPLC size exclusion chromatography through Superdex 75 (GE Healthcare Bio-Sciences Corp., Piscataway, NJ) run at a rate of 0.5 ml/min was used to remove minor contaminants. The human smooth muscle α-actinin spectrin-like repeat domain (ACTN1-R2-R3-GST) was cloned and expressed as a GST-fusion protein, as previously described (Chi et al., 2005). Alanine and

phosphomimetic mutations of α-actinin R2-R3 loop (K421A, Y423A, T425A, T427D, N586A, T590A,

N591A) and sm-titin Zq domain (S43A-Q53A-K64A) residues were obtained by site-directed mutagenesis (QuikChange Site-Directed Mutagenesis Kit, Stratagene).

For surface plasmon resonance experiments, purified ACTN1-R2-R3-GST was cleaved with thrombin (10 U/1mg, Sigma) according to manufacturer’s recommendations and passed over immobilized glutathione agarose (Pierce, Rockford, Ill.) in order to isolate the ACTN1-R2-

R3 domain. WT-Zq and the S43A-Q53A-K64A Zq triple mutant were purified to homogeneity and dialyzed extensively into HBS-EP+ Running Buffer (Biacore, BR-1008-26). Protein concentrations were calculated using predicted extinction coefficients (ProtParam site at ExPASy) and absorbance at 280 nm obtained from a full wavelength scan using a Varian Cary 300 UV/VIS spectrophotometer.

Gel electrophoresis and western blots of native sm-titin Chicken striated pectoralis muscle and pig aorta and uterus tissues were obtained from local slaughterhouses. The tissues were transported to the lab on ice, minced, quick-frozen in liquid nitrogen, and pulverized into powder. The powder was extracted using 1X SDS-sample buffer. The SDS-extract was separated on highly porous 4-20% gradient SDS-polyacrylamide, as described previously (Eilertsen et al., 1992). The gel was electroblotted onto nitrocellulose using semi-dry blot transfer system (Bio-Rad Laboratories, Hercules, CA) and stained using Ponceau S. The blots were blocked in 5.0% non-fat dry milk for 1-2 hours at room temperature. The anti- TKZ rabbit polyclonal primary antibody raised in the T. Keller lab against an expressed titin region containing the Z-repeats Zr1, Zr2, Zr3, Zr7 and the Zq domain, described previously (Cavnar et al., 2007), was applied to the membrane and incubated for 1-2 hours at room temperature, followed by multiple washes in TBS-Tween 20, and incubated with a secondary antibody conjugated to horseradish peroxidase for 1 hour. The chemiluminescent signal was developed using Amersham Biosciences ECL Plus western blotting detection system according to the manufacturer’s recommendations.

Immunolocalization of sm-titin in cryosections and cultured cells A segment of porcine aorta obtained from a slaughterhouse was placed into ice-cold cryoprotectant solution containing sucrose-phosphate and fixed overnight in a solution containing freshly prepared 3.7% formaldehyde and 0.1% Triton-X-100. The frozen aorta was cross-sectioned at a thickness of 10 μm in a cryostat. The sections were collected on 2% gelatin-

coated slides and the cryosection slides were incubated with anti-TKZ polyclonal antibody (diluted 1:100), followed by Alexa Fluor 488-coupled secondary antibody (diluted 1:500), mounted with ProLong ® Gold mounting medium (Invitrogen Corp., Carlsbad, CA), and viewed with a Zeiss LSM 510 (Zeiss, Inc., Thornwood, NY). The A7r5 rat aorta and human coronary artery smooth muscle cells were grown on Nafion®-coated coverslips for 3-4 days (Salloum, Olenych, Keller, & Schlenoff, 2005). The cells were fixed with 3.7% formaldehyde in 1xPBS for 15 minutes and then permeabilized with 0.2% Triton X-100-PBS for 15 minutes at room temperature. To minimize non-specific binding, the coverslips were blocked in 10% Goat Serum, 0.05% TX-100-PBS for 60 minutes before incubation at 37oC for 1hr each in the anti-TKZ polyclonal antibody primary (diluted 1:100) and Alexa Fluor 488-conjugated (diluted 1:300) secondary antibodies. The A7r5 cells were double labeled with phalloidin (rhodamine) and the anti-TKZ polyclonal antibody. The human coronary artery cells were labeled with only the anti-TKZ polyclonal antibody. These coverslips were observed with a Leica TCS SP2 AOBS laser confocal microscope.

ACTN1-R2-R3-GST binding assays ACNT1-R2-R3-GST and mutations were purified to homogeneity and immobilized onto glutathione-agarose beads (1mg/ml). Purified sm-titin Zq domain (0.5 mg/ml) was dialyzed into interaction buffer containing 100 mM KCl and 0.2% Triton X-100 and mixed with the ACNT1- R2-R3-GST beads overnight at 4°C. The beads were pelleted by centrifugation, washed twice with interaction buffer, and eluted with 1XSDS-PAGE sample buffer. The amount of sm-titin Zq binding was assessed by SDS-PAGE of the elutant. All pull-down experiments were done in duplicate. Solid-phase protein-binding assay using microtiter plates was done as previously described (Chi et al., 2005). For these experiments, microtiter plate wells were coated with expressed sm-titin Zq domain and incubated with ACTN1-R2-R2-GST in 100 mM KCl, 0.2% Triton X-100. The amount of ACTN1-R2-R2-GST bound was assessed with anti-GST antibody (1:1000, Affinity BioReagents, Golden, CO). The mean value and standard error were calculated from three separate experiments.

Surface plasmon resonance 2000 Response Units (RU) of ACTN1-R2-R3 domain in 10 mM acetate pH 4.5 was immobilized on a Series S Senor chip CM5 (Biacore Life Sciences, Piscataway, NJ) using amine

coupling chemistry. The sm-titin Zq domain (WT-ZQ) or mutant ZQ domain (S43A-Q53A-K64A- ZQ) was dialyzed extensively into HBS-EP+ Running Buffer (Biacore, BR-1008-26) and flowed over the chip at a rate of 10 μl/min (300 seconds contact time) and regenerated with running buffer containing 1M NaCl (30 μl/min). To determine steady state affinity, binding of various

concentrations (0-6.4 μM) of WT-ZQ or S43A-Q53A-K64A-Zq were assayed. Binding constants were extrapolated from response (RU) versus time (seconds) values using Biacore T-100 analysis software. For relative ranking studies, WT-ZQ and S43A-Q53A-K64A-ZQ were included in the same experiment and binding was compared directly from the resulting sensograms. All sensograms were double zeroed with values of the running buffer alone and from a reference cell containing a blank chip containing no ACTN1-R2-R3 domain. All experiments were done in duplicate using a Biacore T-100 system housed in the Florida State University College of Medicine Biomedical Proteomics Facility.

Graphic modeling and sequence alignment The Visual Molecular Dynamic (VMD) viewer (University of Illinois at Urbana- Champaign) was used for visual modeling of the mutations in the loops of the α-actinin R2-R3 domain and modeling of the ZQ domain binding to this region (PDB 1SJJ). Sequence similarity and alignment were produced using ClustalW (EMBL-EBI) and Jalview (Barton Group, University of Dundee).

Results

Smooth muscle RNA contains a titin transcript encoding Z-repeat and Zq α-actinin- binding domains RT-PCR analysis of commercially available human carotid artery, uterus, and bladder total RNAs yielded cDNAs with sequences matching those of exons from the exon 1-28 region of the only known human titin gene. All cDNAs obtained lacked intron sequence and crossed

predicted exon-exon boundaries, proving they arose from mRNA transcripts and not contaminating genomic DNA. The three smooth muscle RNA sources each yielded a single cDNA product (~2100 bp) from a pair of primers designed to exon 1 and exon 14 of the titin gene that was significantly smaller than the predicted size of the product if all exon sequences were present. Sequence analysis verified the exclusion of exons 11-13, with all products bridging the exon boundary between exons 10 and 14 (Fig. 12). The excluded exons 11-13 encode the Z repeats 4-6. The cDNA products obtained therefore encode the Z1 and Z2 Ig domains, Zis1 unique sequence, and the α-actinin binding domains known as the Z-repeats Zr1, Zr2, Zr3, Zr7 and Zq domain. The cDNA products obtained using primers designed to exons 8 and exon 20 contained the sequences of all known intervening exons (Fig. 12B). We found the same pattern of exon usage by similar RT-PCR analysis of total RNA isolated from rat aortic smooth muscle A7r5 cells grown in culture.

Western blot demonstrates presence of sm-titin protein in smooth muscle To confirm the expression of titin containing the α-actinin-binding region in adult smooth muscle tissues, we raised a rabbit polyclonal antibody (anti-TKZ) against the region encoded by cloned exons 8-14 expressed as a GST fusion protein in bacteria. Western Blot analysis using the anti-TKZ antibody demonstrated a reactive band in crude extracts of porcine aorta and stomach smooth muscle (Fig. 13B, lanes 2, 3) that migrated at a rate similar to that of chicken skeletal muscle titin (Fig. 13B, lane 1). Lack of reactivity with any of the other bands in the crude extracts demonstrated the high titin specificity of the anti-TKZ antibody. Reactivity of the anti-TKZ antibody, which was raised against an expressed fragment encoded by titin gene sequence, with the protein previously identified by the lab as smitin confirms smitin as a smooth muscle isoform of titin.

Sm-titin immunolocalization Immunofluorescence localization using the anti-TKZ antibody demonstrated the presence of sm-titin in porcine aorta and cultured smooth muscle cells (Fig. 14, A-C). In intact porcine aorta, anti-TKZ labeled the smooth muscle layer of the tissue and not the elastic filaments lining the aorta (Fig. 14A). In cultured smooth muscle cells, the anti-TKZ antibody stained in a punctate pattern along the stress fiber-like structures.

Loops in the R2-R3 domain mediate sm-titin binding In reviewing a recent model of smooth muscle α-actinin structure, it became apparent that loops from the spectrin-repeat R2 and R3 domains could constitute the unknown binding site for the sm-titin Zq domain. The structural model predicts an R2 domain loop (residues 421-425) and an R3 domain loop (residues 586-591), which are 145 residues apart in the primary sequence, lie in proximity on the surface of the α-actinin monomer rod and that the sites from both monomers lie on the same side of the dimer (Fig. 15). We used pull-down experiments with alanine mutations of several R2 loop residues

(K421A, Y424A, and T425A) and R3 loop residues (N586A, T590A, and N591A), a phosphomimetic

mutation of an R2 loop threonine residue (T427D), and a double mutant of both R2 and R3

residues (T425A/N586A) to investigate whether the predicted α-actinin R2 and R3 loops contribute to the site that binds the titin Zq domain. For the GST-pull-down experiments, the R2-R3 domain region constructs were expressed as a GST fusion proteins and the titin Zq region was expressed as a HIS-tagged fragment. Glutathione-coupled beads efficiently pelleted the GST-R2-R3 domain alone (Fig. 16A, lane 6). A saturable amount of the Zq domain co-pelleted when bound to the wild type R2-R3 (Fig. 16A, lane 5), but none pelleted in the absence of the R2-R3 domain under the same conditions (Fig. 16B, lane 4). Alanine mutations of several residues from both the R2 loop domain and the R3 loop domain significantly decreased Zq binding indicated by the amount of Zq co-pelleted with the R2-R3 region (Fig. 16C). A solid phase binding assay in which the amounts of the R2-R3 domain wild type and mutant fragments bound to wild type titin Zq domain immobilized in the wells were determined confirmed the results of the pull-down assay. All of the R2-R3 domain mutants exhibited a significant decrease in binding compared to the wild type binding, with those exhibiting the least

interaction in the pull-down assay also yielding the least binding in the solid phase assay (Fig.

17). It is interesting to note that alanine mutation of both the R2-T425 and the R3-N586, which the model predicts to be in proximity, has significantly less effect on binding than mutation of each residue alone. Taken together, these results support the hypothesis that loops from the R2 and R3 domains combine to form a binding site for the titin Zq domain. Moreover, the effect of the phosphomimetic mutation of the R2 domain T427 residue (T427D), which is part of a predicted Casein Kinase II phosphorylation site (TLSE), raises the possibility that phosphorylation may regulate this region of sm-titin-α-actinin interaction, although evidence for phosphorylation of the rod region of α-actinin in vivo remains lacking.

A helical region in the sm-titin Zq domain is important for α-actinin binding We used surface plasmon resonance to quantify sm-titin Zq domain interaction with wild type smooth muscle α-actinin R2-R3 fragment and to identify the region of sm-titin Zq that interacts with α-actinin. Wild type sm-titin Zq domain bound to immobilized α-actinin R2-R3 domain with a Kd of 344±3 nM (Fig. 18B). Site-directed alanine mutagenesis of Zq domain residues narrowed the region of Zq that interacts with the R2-R3 domain to a highly conserved mostly helical region. The titin Zq domain fragment used for all the α-actinin R2-R3 binding studies described contains 135 amino acids encoded by sequence in titin gene exons 15 and 16. A region of this sequence is highly conserved across a wide variety of vertebrate species (Fig. 19). Several secondary structure prediction programs applied to the entire 135 residue human sequence predict various amounts of extended β structure (14-40%), random coil (49-56%), and α-helix (11-30%), but all support a high propensity for α-helix formation (14-100%) in the highly conserved region of the titin Zq sequence. Predicting that a highly conserved α-helical Zq region (aa42-64 in Fig. 19) may bind in the groove between the α-actinin R2 and R3 domain loops, we tested the effect of alanine

mutation of three well-conserved residues in the predicted Zq domain. The triple S43A-Q53A-

K64A Zq mutant bound very poorly to immobilized α-actinin R2-R3 domain when compared to wild type Zq in relative rank studies (Fig. 20). For these experiments, equal concentrations of the wild type and mutant Zq domains were tested, but only one wild type concentration that exhibits 39

binding in the range of that of the highest mutant concentrations is shown in the sensogram. Subsequent steady state affinity experiments using the Zq triple mutant failed to reach saturation even at the highest Zq domain concentration (data not shown). These results suggest that the highly conserved helical region in the Zq domain may bind to the conserved surface loops from the R2 and R3 spectrin repeat-like domains. A gel filtration step in the protocol routinely used to purify each expressed fragment revealed one possible difference between the wild type Zq and mutant Zq domains. The wild type Zq domain migrated through the Superdex 75 column as a single peak at a rate greater than twice that (34-44 kDa) predicted for a protein with the predicted 17 kDa molecular weight, when compared to the migration of six molecular weight standards from 13.2 kDa to 66.4 kDa run under the same condition (Fig.21A and B). This raised the possibility that the wild type Zq domain forms a dimer in solution. In contrast, the triple mutant migrated as two distinct peaks. The leading peak migrated at a rate similar to that of the wild type fragment (34-44 kDa), whereas the trailing peak migrated at the rate (17-25 kDa) predicted for a monomer of the predicted size (17 kDa), suggesting that mutation of the three residues partially disrupts the dimer and creates a pool of monomers during purification (Fig. 21A and C).

Discussion

We previously mapped smooth muscle α-actinin binding sites for native sm-titin to the central rod R2-R3 spectrin-like repeats and C-terminal domains, sites that are analogous to those in striated muscle α-actinin that bind respectively to striated muscle titin Zq and Z-repeat domains (Chi et al., 2005). Whether Z-repeat and Zq domains are present in any adult smooth muscle titin isoforms recently was brought into doubt, however, by an investigation that revealed the presence of smooth muscle transcripts containing sequence from only 80-90 titin gene exons and excluding the Z-repeat- and Zq-encoding exons (Labeit et al., 2006). We used RT-PCR analysis and antibody detection to confirm the presence of these domains in a long adult smooth muscle titin isoform. RT-PCR analysis of human carotid artery RNA yielded products encoding Z-repeats Zr1, Zr2, Zr3, Zr7, and the Zq domain. Other products from this RNA encoded the largest exon of the myosin-binding domain, the C-terminal kinase domain, as well as uniquely

spliced portions of the PEVK domain. Similar products were obtained from commercially available human bladder and uterus RNA and RNA isolated from cultured rat aortic smooth muscle A7r5 cells (data not shown). Moreover, an antibody (anti-TKZ) raised against this region, which reacts with striated muscle titin, also reacted in Western blots with the very high molecular protein originally characterized by us as smitin and yielded strong immunofluorescence signal from the smooth muscle cells in cryosections of adult smooth muscle. This anti-TKZ antibody also revealed epitope localization in a punctate pattern along the stress fiber-like structures in the contractile apparatus of A7r5 cells, which were induced to become contractile by culture on a polyelectrolyte substrate (manuscript in preparation). Inspiration for this investigation arose from visual analysis of a smooth muscle α-actinin structural model (Liu et al., 2004), which revealed loops from the R2 and R3 spectrin-like domains lie in proximity to each other on the rod surface, despite being separated by ~150 residues in the primary sequence. Moreover, the model indicates that identical R2-R3 loop pairs form a two-fold axis of symmetry on the same side of the α-actinin antiparallel homodimer rod domain. Based on this model, we targeted certain R2-R3 loops residues for alanine mutagenesis to determine effects on Zq domain binding. We found that single alanine mutations of residues in the R2-R3 domain loops region significantly reduced sm-titin Zq domain binding in GST- pulldown experiments and solid phase binding assays. Moreover, a phosphomimetic mutation of the R2 domain T427 residue (T427D) also decreased binding (Fig. 16C), raising the possibility that phosphorylation may regulate sm-titin-α-actinin interaction. Although such phosphorylation has yet to be demonstrated in vivo, it may occur only transiently during structural rearrangement of the contractile apparatus for mechanical or phenotypic plasticity of the smooth muscle cell. The specific region of the Zq domain that interacts with the R2-R3 loops was suggested by existing deletion analysis of cardiac titin and cardiac α-actinin constructs (Young et al., 1998; Young et al., 2000) and analysis presented here demonstrating a high degree of sequence conservation across various vertebrate titins in a region predicted to have high α-helical propensity. Using surface plasmon resonance, we confirmed that a triple alanine mutation of this region severely decreased its affinity for immobilized α-actinin R2-R3 rod. Various algorithms predict that the region encompassed by the triple mutation (SIAGSAIATLQKELSATSSAQK) forms an amphipathic helix or perhaps a broken helix, which might be optimal for lying in the

groove between the α-actinin R2 and R3 loops. Considering that in sm-titin the Zr7 domain, which is more N-terminal than the Zq domain, can interact with the α-actinin C-terminal EF34, it is reasonable to predict that the N-terminal end of the interacting Zq sequence is oriented toward the T427-T591 end of the R2-R3 groove on the same monomer. The proximity on the same side of the rod and the two-fold symmetry of the R2-R3 loop alignments supports the possibility that Zq helical regions lying in the R2-R3 loop grooves might interact with each other in an anti-parallel orientation. Interestingly, while purifying the bacterially expressed wild type Zq domain with size-exclusion FPLC chromatography, we found it eluted as if it had twice the anticipated molecular weight, perhaps indicating dimerization. In contrast, the triple mutant yielded two elution peaks, the apparent dimer peak and a slower peak migrating at the rate predicted for a 17 kDa monomer. Confirmation of anti-parallel interaction of helices in this region of the Zq domain will require additional experimentation. Based on these results and our predicted orientation of the Zq domain interaction, we present a model of sm-titin Zq domain interaction with the smooth muscle α-actinin rod domain in which Zq domains from two sm-titins homodimerize to promote and support the protein complex (Fig. 22). If correct, this would suggest that a single α-actinin molecule could crosslink two sm-titin molecules in the smooth muscle dense body independently of other possible crosslinking proteins, which have yet to be identified in the smooth muscle contractile apparatus.

Figure 12. Smooth muscle titin RT-PCR products. (A) The diagram of the 28 exons comprising the 5' end of the human titin gene [Modified from Bang et al. 2001] illustrates exons (colored) included in RT-PCR products obtained from three commercially available RNA sources; human carotid artery, uterus, and bladder RNA and RNA from cultured rat aorta smooth muscle A7r5 cells. The green boxes indicate exons encoding the four Z-repeat and Zq domains that bind to α-actinin. The red boxes encode Ig domains. The blue boxes encode unique domains. The purple area indicates exons excluded from all products. The arrows (not to scale) designate positions of the exon 1, 8, and 20 forward (F) primers and the exon 14, 20, and 28 reverse (R) primers used for the RT-PCR analysis. (B) The relationship between the possible product PCR size if all exon sequences are included and the actual PCR product size obtained reveals the exclusion of the Z-repeat 4, 5, and 6-encoding exons 11, 12, and 13 covered by the purple box.

Figure 13. Anti-TKZ reactivity against a high molecular weight band in smooth muscle extracts. The Anti-TKZ rabbit polyclonal antibody was raised against a bacterially-expressed fragment composed of the sm-titin α-actinin-binding region (Z-repeats Zr1, Zr2, Zr3, Zr7, and the Zq domain). (A) SDS extracts of chicken pectoralis striated muscle used as a standard for migration of titin (lane 1) and two porcine smooth muscles (lanes 2 and 3) were fractionated by SDS-PAGE, electroblotted to nitrocellulose, and stained with India ink. T, titin; MHC, myosin heavy chain. (B) Western Blot analysis of a duplicate blot from the same gel using anti-TKZ antibody reveals a band with a migration rate similar to that of chicken striated muscle titin (lane 1) in the smooth muscle extracts (lanes 2 and 3).

Figure 14: Localization of smooth muscle titin in cryosections of smooth muscle and cultured smooth muscle cells. (A) In cross-sections (10 μm thick) of adult porcine aorta, anti- TKZ labels cells in the smooth muscle layer and not the elastic filaments lining the aorta. (A; 10x, B; 20x, C; 40x) Nuclei are labeled with DAPI (blue) Bars, 50 μm. (B) Anti-TKZ antibody staining of human coronary artery cells that were grown for 4 days on a contractile phenotype- inducing polyelectrolyte multilayer (Nafion)-coated coverslip reveals a punctate pattern of labeling along the stress fiber-like structures of the smooth muscle cell. (C) Anti-TKZ antibody stains the contractile apparatus of rat aorta smooth muscle A7r5 cells cultured on Nafion-coated coverslips in a punctate pattern. Bars, 20 μm.

Figure 15. Model representation of the smooth muscle α-actinin central rod spectrin repeat-like R2 and R3 domains and key loop residues. A structural model of the smooth muscle α-actinin R2 and R3 domains in the central rod [modified from Liu et al. 2004] shows positions of the R2 and R3 loop residues mutated for sm-titin Zq binding interaction experiments. One monomer peptide of the antiparallel α-actinin homodimer is shown in blue. The antiparallel monomer is shown in red.

Figure 16. Sm-titin Zq domain binding to wild type and mutated smooth muscle α-actinin R2-R3 domains in vitro. (A-C) Wild type and mutant smooth muscle α-actinin R2-R3 domains expressed as GST fusion proteins and immobilized on glutathione beads were incubated with an expressed smooth muscle Zq domain fragment in buffer containing 100 mM KCl and 0.2% triton X-100. (A) Shown is a representative SDS gel illustrating unbound Zq domain in the supernatant and washes of the pelleted beads (lanes 2-4), Zq bound to the wild type R2-R3 domain on the beads after the washes (lane 5), and the binding or the R2-R3 fragment alone to the beads. Lane 1 contains chicken pectoralis muscle extract used as a molecular weight standard. (B) A control experiment demonstrates that the Zq domain fails to bind to blocked beads lacking the α-actinin R2-R3 domain. (C) A gel showing the amount of bound Zq (only the Zq region of the gel is shown) in the washed pellet fractions from experiments in which R2-R3 domains with the specified mutations were bound to the beads reveals variable effects of the mutations on Zq binding. The mutations tested include the R2 loop mutations K421A and T425A, the R3 loop mutations N586A, T590A, N591A, the R2-R3 double mutant T425A/N586A, and the phosphomimetic mutation T427D of a putative CK2 phosphorylation site. The +/- signs indicate the lanes in which Zq binding was detectable (+) or undetectable (-) on the gel.

Figure 17. Solid Phase Binding Assay confirms a decrease in binding in mutated R2-R3 Loops. Microtiter wells coated with the expressed smooth muscle titin Zq domain were incubated with Wild type and mutant α-actinin R2-R3 GST fusion proteins or GST alone. The presence of GST was detected with an anti-GST antibody, followed by a secondary antibody conjugated to alkaline phosphatase, and incubation with substrate for color development. The A405 nm mean ± SEM for each mutant was calculated from three separate experiments.

Figure 18. Sm-titin Zq domain binds to α-actinin R2-R3 domain with nanomolar affinity. (A) The surface plasmon resonance responses were recorded for various wild type sm-titin Zq concentrations (0-6400 nM) flowed at 10 μl/min over wild type α-actinin R2-R3 domain that was immobilized on a Biacore CM5 Sensor Chip. (B) Shown is a representative binding isotherm generated using sensogram plateau RU values from one experiment. The mean Kd determined from two experiments is Kd=344 ± 3 (SE) nM.

Figure 19. Predicted sm-titin Zq domain contains highly conserved residues. Multispecies alignment of the sm-titin Zq domain predicted from the titin gene sequences of representative

vertebrates shows highly conserved regions within predicted Zq domains. Mutated S43A, Q53A, and K64A residues are highlighted (green, red) and were chosen for their highly conserved nature across all or most species. Numbering reflects residue position in the primary sequence encoded by human titin gene exons 15 and 16.

Figure 20. Mutations in the sm-titin Zq domain significantly decrease affinity for the α- actinin R2-R3 domain. A surface plasma resonance relative ranking experiment was used to

compare the binding of wild type Zq domain (WT-Zq) domain and the S43A-Q53A-K64A-Zq triple mutant (Sqk-Zq-a). An expanded region of the sensogram generated from binding of one wild type concentration (156 nM) and two triple mutant concentrations (625 and 1250 nM) demonstrates a significantly (more than 8 fold) weaker binding of the mutants compared to wild type.

Figure 21. FPLC elution of purified Zq on Superdex 75 suggests smooth muscle titin Zq domain forms a dimer (A) Superimposed Superdex 75 FPLC elution profiles of purified wild type Zq (WT-Zq) and the S43A-Q53A-K64A-Zq triple mutant (sqk-Zq-a) demonstrates that the WT-Zq migrates through the column as a single peak (A and B) at a rate predicted for a 30-35 kDa protein and the triple mutant migrates as two peaks (A and C), the slower of which migrates with a rate predicted for a 17-18 kDa protein. The 20 kDa and 15 kDa indicate the protein markers in the molecular weight standard mix (Benmark Prestained, Invitrogen). The slight shift in peak fractions between the scan (A) and gels (B and C) reflects the volume in the column system between the scan and fraction collection sites. Superdex 75 column was standardized with BSA, Carbonic Anhydrase, and RNase A.

Figure 22. Proposed Model for titin Zq domain interaction with the α-actinin R2-R3 central rod domain. We hypothesize that loops in the R2 and R3 spectrin repeat-like domains in each of the anti-parallel monomers (blue and red), which are modeled to lie in proximity on the surface of the central rod and are highly conserved between smooth and striated muscle α- actinins, constitute the binding site for the 26 residue helical region of the Zq domain (yellow). Proximity of Zq domains interacting with the R2-R3 domain sites on each monomer may allow interaction of the helical regions of two Zq domains, which may promote stability of the complex.

CHAPTER 4 SUMMARY AND CONCLUSION

Although the overall organization of the smooth muscle cell is complex and remains poorly understood, organization of the smooth muscle contractile apparatus is thought to be similar to that of the striated muscle sarcomere. In both striated and smooth muscle contractile systems, myosin filaments produce force on oppositely oriented actin filaments that are anchored to one of a pair of functionally similar α-actinin-containing structures - the sarcomere Z-disk or the smooth muscle dense body (or dense plaque) - to pull the structures closer together. In striated muscles, a third filament system composed of the protein titin establishes and maintains sarcomere organization and integrity. We first discovered sm-titin (previously named smitin) as an extremely high molecular weight polypeptide (~2 MDa) in a high ionic strength-ATP extract of chicken gizzard smooth muscle (Kim et al., 2002). Sm-titin is a long (700-800 nm), fibrous molecule with a single globular head. The molecular morphology of sm-titin is similar to that of striated muscle titin and what we found previously for the cytoskeletal titin-like protein c-titin. The length of sm-titin is somewhat shorter than that of muscle titin, in accord with its apparent lower molecular weight. The initial discovery raised the possibility that sm-titin plays roles in the smooth muscle contractile apparatus similar to those of titin in the striated muscle sarcomere. The work presented here supports that notion by demonstrating a high degree of similarity in smooth muscle α-actinin-sm-titin and striated muscle α-actinin-titin interactions. Data presented in Chapter 2 demonstrate that smooth muscle α-actinin interacts with sm-titin with nanomolar affinity and has two sm-titin-binding sites in molecular locations similar to those of the two titin-binding sites in striated muscle α-actinin. We found, as in the striated muscle α-

actinin, one of the smooth muscle α-actinin sm-titin-binding sites is located at the C-terminal end of both striated muscle and smooth muscle α-actinins. PIP2 enhances the interaction of striated muscle α-actinin with both titin and sm-titin but not the interaction of smooth muscle α-actinin with either sm-titin or titin. This raises the possibility that there is an alternative mechanism for regulating interaction between smooth muscle α-actinin and sm-titin. The other titin/sm-titin-binding site is in the R2-R3 triple helical repeat region. This region of striated muscle α-actinin interacts with the Zq domain of titin (Young et al., 1998). Prior to the work presented here, the molecular basis for interaction of either the striated or smooth muscle α-actinin rod domains with titin or sm-titin remained unknown, because neither binding site had been mapped at higher resolution. Lack of sm-titin sequence availability had been a major impediment in previous attempts to understand better the roles of sm-titin in smooth muscle cells. For example, whether sm-titin contained any of the α-actinin-binding Z-repeat or Zq domains was not known. For the investigation presented in Chapter 3, we used RT-PCR analysis and antibody detection to confirm the presence of these domains in adult smooth muscle sm-titin. RT-PCR analysis of four different smooth RNAs yielded products encoding Z-repeats Zr1, Zr2, Zr3, Zr7, and the Zq domain. Moreover, an antibody (anti-TKZ) raised against this region reacted in Western blots with a very high molecular protein. The antibody also yielded strong immunofluorescence signal from the smooth muscle cells in cryosections of adult smooth muscle and revealed epitope localization in a punctate pattern along the stress fiber-like fibers in A7r5 cells and human coronary artery cells. In studies not presented here, we also found that approximately 70% of the 208 distinct peptide masses of trypsin-digested pig stomach sm-titin resolved by Fourier transform ion cyclotron resonance mass spectrometry (FT-ICR MS) match masses of predicted human titin gene-encoded peptides (Keller Lab unpublished observations). In order to further investigate the Zq domain binding to the α-actinin R2-R3 domain, we utilized a smooth muscle α-actinin structural model (Liu et al., 2004). Visual analysis of this model suggested that loops for the R2 and R3 domains of α-actinin might provide the binding site for the sm-titin Zq domain. To test this hypothesis, we introduced single alanine mutations of residues in the R2-R3 domain loops region and found they significantly reduced sm-titin Zq domain binding in GST-pulldown experiments and solid phase binding assays. Moreover, a 57

phosphomimetic mutation of the R2 domain T427 residue (T427D) also decreases binding, raising the possibility that phosphorylation may regulate sm-titin-α-actinin interaction. Using surface plasmon resonance, we found a region within the reported Zq domain that is responsible for the interaction with the α-actinin R2-R3 region. A triple alanine mutation of this region severely decreased its affinity for the immobilized α-actinin R2-R3 region. Additionally, purifying bacterially-expressed wild-type Zq domain with size-exclusion chromatography produced an elution velocity predictive of a protein greater than twice the molecular weight, supporting the possibility that the Zq domain dimerizes even in the absence of the α-actinin R2-R3 rod domain. Whereas, purification of the triple mutant eluted both the apparent dimer and monomer peaks. This suggests the sm-titin Zq domain is a dimer and residues within the predicted helical portion of Zq domain mediates the sm-titin-Zq-α-actinin interaction. Based on these results, we have presented a model of sm-titin Zq domain interaction with the smooth muscle α-actinin rod domain in which Zq domains from two sm-titins homodimerize to promote and support the protein complex while oriented toward the T427-T591 end of the R2-R3 groove on the same α- actinin monomer and each sm-titin Zr7 domain interacts with the α-actinin C-terminal EF34. If correct, this would suggest that a single α-actinin molecule alone could crosslink two sm-titin molecules in the smooth muscle dense body independent of other possible interactions of sm-titin with itself or other unknown crosslinking proteins. It is also interesting to speculate that the binding of a single molecule of titin to the α-actinin C-terminal and rod domains on each α-actinin monomer could determine the orientation of these two domains with respect to each other. If so, titin binding to both the C-terminal and rod sites may lock an α-actinin molecule into a conformation that constrains the mobility of the actin-binding domains to opposite orientations on the two ends of the α-actinin molecule. This would effectively orient crosslinked actin filaments in antiparallel directions. Taken together, the evidence presented in this study not only provides substantial support for the conclusion that the titin gene encodes sm-titin but also hints at its potentially vital role in smooth muscle cells.

Smooth muscle titin in the contractile unit The morphology of sm-titin combined with its known interactions fit nicely into the current models of the smooth muscle cell, where the myosin sidepolar thick filament is sandwiched between two antipolar actin thin filaments anchored to two dense bodies or dense plaques, ranging 1.8 – 2.2 μm apart (Hodgkinson, Newman, Marston, & Severs, 1995). Through electron micrographs of rotary replicated molecules of sm-titin, we estimate the size to be 750- 900 nm. We also know sm-titin associates strongly with smooth muscle myosin and α-actinin in vitro and colocalizes well with both in vivo (Kim et al., 2002). This orientates the N-terminal of sm-titin to the α-actinin enriched dense bodies (or plaques) and stretches its C-terminal domain to the myosin thick filament. We estimate the myosin to sm-titin molecular ratio in side-polar coassemblies to be 85:1. Assuming each sidepolar filament contains no more than 170 myosins, this would be most consistent with a ratio of ~2 sm-titins molecules per thick filament or contractile unit. The length of the sm-titin tail domain ( 900 nm) appears to be on average a little shorter than the tail domains of the striated muscle titin (~1,000 nm long), however, if the myosin thick filament was unusually long, as found in smooth muscle cells (1,600 nm), this would allow the sm-titin to physically reach the thick filament. However, a thick filament the same size as the contractile unit as postulated by the “parallel syncytium” model would require sm-titin to encode an extremely large myosin binding domain (90%), which is highly unlikely. It is also interesting to speculate on the nuclear envelope/dense plaque junction, where the thin filament anchors to the nucleus because this would also localize sm-titin to the periphery of the nucleus (Zastrow, Flaherty, Benian, & Wilson, 2006) and suggest an unknown signaling role for sm-titin.

Smooth muscle titin in phenotypic plasticity Reorganization due to phenotypic or mechanical plasticity appears to involve disassembly of the large, sidepolar myosin filaments and antiparallel actin filaments of the contractile system and reassembly of the myosin and actin into stress fiber-like structures, similar to those found in many types of motile cells. We have demonstrated the ability of sm-titin to organize small bipolar filaments of smooth muscle myosin into linear arrays similar to those found in cytoskeletal structures such as stress fibers (Kim et al., 2002). Transition of sm-titin– myosin interaction from the organization of large sidepolar filaments in the contractile apparatus

of the contractile phenotype to the stress fiber-like arrangement of small bipolar filaments may play a role in the change of the cell to the synthetic phenotype. Alternatively, sm-titin organization of small bipolar filaments of smooth muscle myosin may play a role in formation of the contractile apparatus during smooth muscle development. Sm-titin is likely to be involved in one of the primary events in adding contractile units. Its interaction with flanking dense bodies in a contractile unit would create the initial framework needed to quickly add thick and thin filaments to a stretching cell. This is similar to what has been described for skeletal muscle titin in myofibrillogenesis, where titin acts as a cytoskeletal scaffold or “molecular blueprint” to coordinate contractile unit assembly (Begum, Song, Rienzie, & Ragolia, 1998). Therefore, a combination of sm-titin interactions with myosin, actin, and α-actinin may play a role in the change of the cell to the synthetic phenotype (Kim et al., 2002). Overall, these investigations provide the groundwork necessary for pinpointing other mechanistic aspects of sm-titin function that contribute to physiological properties of smooth muscle cells.

REFERENCES

Ali, F., Pare, P. D., & Seow, C. Y. (2005). Models of contractile units and their assembly in smooth muscle. Can.J.Physiol Pharmacol., 83, 825-831.

Atkinson, R. A., Joseph, C., Dal, P. F., Birolo, L., Stier, G., Pucci, P. et al. (2000). Binding of alpha-actinin to titin: implications for Z-disk assembly. Biochemistry, 39, 5255-5264.

Atkinson, R. A., Joseph, C., Kelly, G., Muskett, F. W., Frenkiel, T. A., Nietlispach, D. et al. (2001). Ca2+-independent binding of an EF-hand domain to a novel motif in the alpha-actinin- titin complex. Nat.Struct.Biol., 8, 853-857.

Ayoob, J. C., Turnacioglu, K. K., Mittal, B., Sanger, J. M., & Sanger, J. W. (2000). Targeting of cardiac muscle titin fragments to the Z-bands and dense bodies of living muscle and non-muscle cells. Cell Motil.Cytoskeleton, 45, 67-82.

Bagby, R. (1986). Toward a comprehensive three-dimensional model of the contractile system of vertebrate smooth muscle cells. Int.Rev.Cytol., 105, 67-128.

Bang, M. L., Centner, T., Fornoff, F., Geach, A. J., Gotthardt, M., McNabb, M. et al. (2001). The complete gene sequence of titin, expression of an unusual approximately 700-kDa titin isoform, and its interaction with obscurin identify a novel Z-line to I-band linking system. Circ.Res., 89, 1065-1072.

Begum, N., Song, Y., Rienzie, J., & Ragolia, L. (1998). Vascular smooth muscle cell growth and insulin regulation of mitogen-activated protein kinase in hypertension. Am.J.Physiol, 275, C42- C49.

Cavnar, P. J., Olenych, S. G., & Keller, T. C., III (2007). Molecular identification and localization of cellular titin, a novel titin isoform in the fibroblast stress fiber. Cell Motil.Cytoskeleton, 64, 418-433.

Chi, R. J., Olenych, S. G., Kim, K., & Keller, T. C., III (2005). Smooth muscle alpha-actinin interaction with smitin. Int.J.Biochem.Cell Biol., 37, 1470-1482.

Clark, K. A., McElhinny, A. S., Beckerle, M. C., & Gregorio, C. C. (2002). Striated muscle cytoarchitecture: an intricate web of form and function. Annu.Rev.Cell Dev.Biol., 18, 637-706.

Cooke, P. (1983). Organization of contractile fibers in smooth muscle. Cell Muscle Motil., 3, 57- 77.

Dixson, J. D., Forstner, M. J., & Garcia, D. M. (2003). The alpha-actinin gene family: a revised classification. J.Mol.Evol., 56, 1-10.

Eilertsen, K. J., Kazmierski, S. T., & Keller, T. C., III (1994). Cellular titin localization in stress fibers and interaction with myosin II filaments in vitro. J.Cell Biol., 126, 1201-1210.

Eilertsen, K. J., Kazmierski, S. T., & Keller, T. C., III (1997). Interaction of alpha-actinin with cellular titin. Eur.J.Cell Biol., 74, 361-364.

Eilertsen, K. J. & Keller, T. C., III (1992). Identification and characterization of two huge protein components of the brush border cytoskeleton: evidence for a cellular isoform of titin. J.Cell Biol., 119, 549-557.

Ford, L. E., Seow, C. Y., & Pratusevich, V. R. (1994). Plasticity in smooth muscle, a hypothesis. Can.J.Physiol Pharmacol., 72, 1320-1324.

Fukami, K., Sawada, N., Endo, T., & Takenawa, T. (1996). Identification of a phosphatidylinositol 4,5-bisphosphate-binding site in chicken skeletal muscle alpha-actinin. J.Biol.Chem., 271, 2646-2650.

Gerthoffer, W. T. & Gunst, S. J. (2001). Invited review: focal adhesion and small heat shock proteins in the regulation of actin remodeling and contractility in smooth muscle. J.Appl.Physiol, 91, 963-972.

Granzier, H., Labeit, D., Wu, Y., & Labeit, S. (2002). Titin as a modular spring: emerging mechanisms for elasticity control by titin in cardiac physiology and pathophysiology. J.Muscle Res.Cell Motil., 23, 457-471.

Granzier, H. L. & Labeit, S. (2004). The giant protein titin: a major player in myocardial mechanics, signaling, and disease. Circ.Res., 94, 284-295.

Gregorio, C. C., Granzier, H., Sorimachi, H., & Labeit, S. (1999). Muscle assembly: a titanic achievement? Curr.Opin.Cell Biol., 11, 18-25.

Halayko, A. J. & Solway, J. (2001). Molecular mechanisms of phenotypic plasticity in smooth muscle cells. J.Appl.Physiol, 90, 358-368.

Herrera, A. M., McParland, B. E., Bienkowska, A., Tait, R., Pare, P. D., & Seow, C. Y. (2005). 'Sarcomeres' of smooth muscle: functional characteristics and ultrastructural evidence. J.Cell Sci., 118, 2381-2392.

Hodgkinson, J. L., Newman, T. M., Marston, S. B., & Severs, N. J. (1995). The structure of the contractile apparatus in ultrarapidly frozen smooth muscle: freeze-fracture, deep-etch, and freeze-substitution studies. J.Struct.Biol., 114, 93-104. 62

Joseph, C., Stier, G., O'Brien, R., Politou, A. S., Atkinson, R. A., Bianco, A. et al. (2001). A structural characterization of the interactions between titin Z-repeats and the alpha-actinin C- terminal domain. Biochemistry, 40, 4957-4965.

Keller, T. C., III (1995). Structure and function of titin and nebulin. Curr.Opin.Cell Biol., 7, 32- 38.

Keller, T. C., III (1997). Muscle structure. Molecular bungees. Nature, 387, 233-235.

Kim, K. & Keller, T. C., III (2002). Smitin, a novel smooth muscle titin-like protein, interacts with myosin filaments in vivo and in vitro. J.Cell Biol., 156, 101-111.

Kuo, K. H. & Seow, C. Y. (2004). Contractile filament architecture and force transmission in swine airway smooth muscle. J.Cell Sci., 117, 1503-1511.

Labeit, S., Lahmers, S., Burkart, C., Fong, C., McNabb, M., Witt, S. et al. (2006). Expression of distinct classes of titin isoforms in striated and smooth muscles by alternative splicing, and their conserved interaction with filamins. J.Mol.Biol., 362, 664-681.

Lange, S., Xiang, F., Yakovenko, A., Vihola, A., Hackman, P., Rostkova, E. et al. (2005). The kinase domain of titin controls muscle gene expression and protein turnover. Science, 308, 1599- 1603.

Langer, B. G. & Pepe, F. A. (1980). New, rapid methods for purifying alpha-actinin from chicken gizzard and chicken pectoral muscle. J.Biol.Chem., 255, 5429-5434.

Linke, W. A. (2007). Sense and stretchability: The role of titin and titin-associated proteins in myocardial stress-sensing and mechanical dysfunction. Cardiovasc.Res..

Liu, J., Taylor, D. W., & Taylor, K. A. (2004). A 3-D reconstruction of smooth muscle alpha- actinin by CryoEm reveals two different conformations at the actin-binding region. J.Mol.Biol., 338, 115-125.

McElhinny, A. S., Kazmierski, S. T., Labeit, S., & Gregorio, C. C. (2003). Nebulin: the nebulous, multifunctional giant of striated muscle. Trends Cardiovasc.Med., 13, 195-201.

Nakamura, A., Isoyama, S., Watanabe, T., Katoh, M., & Sawai, T. (1998). Heterogeneous smooth muscle cell population derived from small and larger arteries. Microvasc.Res., 55, 14-28.

Newby, A. C. & Zaltsman, A. B. (2000). Molecular mechanisms in intimal hyperplasia. J.Pathol., 190, 300-309.

Ohtsuka, H., Yajima, H., Maruyama, K., & Kimura, S. (1997). Binding of the N-terminal 63 kDa portion of connectin/titin to alpha-actinin as revealed by the yeast two-hybrid system. FEBS Lett., 401, 65-67.

Orford, J. L., Selwyn, A. P., Ganz, P., Popma, J. J., & Rogers, C. (2000). The comparative pathobiology of atherosclerosis and restenosis. Am.J.Cardiol., 86, 6H-11H.

Pan, K. M., Damodaran, S., & Greaser, M. L. (1994). Isolation and characterization of titin T1 from bovine cardiac muscle. Biochemistry, 33, 8255-8261.

Pavalko, F. M. & Burridge, K. (1991). Disruption of the actin cytoskeleton after microinjection of proteolytic fragments of alpha-actinin. J.Cell Biol., 114, 481-491.

Salloum, D. S., Olenych, S. G., Keller, T. C., & Schlenoff, J. B. (2005). Vascular smooth muscle cells on polyelectrolyte multilayers: hydrophobicity-directed adhesion and growth. Biomacromolecules, 6, 161-167.

Seow, C. Y., Pratusevich, V. R., & Ford, L. E. (2000). Series-to-parallel transition in the filament lattice of airway smooth muscle. J.Appl.Physiol, 89, 869-876.

Small, J. V. (1985). Geometry of actin-membrane attachments in the smooth muscle cell: the localisations of vinculin and alpha-actinin. EMBO J., 4, 45-49.

Small, J. V. (1995). Structure-function relationships in smooth muscle: the missing links. Bioessays, 17, 785-792.

Small, J. V. & Gimona, M. (1998). The cytoskeleton of the vertebrate smooth muscle cell. Acta Physiol Scand., 164, 341-348.

Somlyo, A. P. (1997). Signal transduction. Rhomantic interludes raise blood pressure. Nature, 389, 908-9, 911.

Trinick, J. (1994). Titin and nebulin: protein rulers in muscle? Trends Biochem.Sci., 19, 405-409.

Tskhovrebova, L. & Trinick, J. (2003). Titin: properties and family relationships. Nat.Rev.Mol.Cell Biol., 4, 679-689.

Tskhovrebova, L. & Trinick, J. (2004). Properties of titin immunoglobulin and fibronectin-3 domains. J.Biol.Chem., 279, 46351-46354.

Tskhovrebova, L. & Trinick, J. (2005). Muscle disease: a giant feels the strain. Nat.Med., 11, 478-479.

Tskhovrebova, L. B. & Makedonov, G. P. (2004). [On the mechanism of adaptive response in human cells]. Radiats.Biol.Radioecol., 44, 657-661.

Turnacioglu, K. K., Mittal, B., Dabiri, G. A., Sanger, J. M., & Sanger, J. W. (1997). An N- terminal fragment of titin coupled to green fluorescent protein localizes to the Z-bands in living muscle cells: overexpression leads to myofibril disassembly. Mol.Biol.Cell, 8, 705-717.

Virel, A. & Backman, L. (2004). Molecular evolution and structure of alpha-actinin. Mol.Biol.Evol., 21, 1024-1031.

Wenegieme, E. F., Babitch, J. A., & Naren, A. P. (1994). Cation binding to chicken gizzard alpha-actinin. Biochim.Biophys.Acta, 1205, 308-316.

Ylanne, J., Scheffzek, K., Young, P., & Saraste, M. (2001). Crystal structure of the alpha-actinin rod reveals an extensive torsional twist. Structure., 9, 597-604.

Young, P., Ferguson, C., Banuelos, S., & Gautel, M. (1998). Molecular structure of the sarcomeric Z-disk: two types of titin interactions lead to an asymmetrical sorting of alpha- actinin. EMBO J., 17, 1614-1624.

Young, P. & Gautel, M. (2000). The interaction of titin and alpha-actinin is controlled by a phospholipid-regulated intramolecular pseudoligand mechanism. EMBO J., 19, 6331-6340.

Zanellato, A. M., Borrione, A. C., Tonello, M., Scannapieco, G., Pauletto, P., & Sartore, S. (1990). Myosin isoform expression and smooth muscle cell heterogeneity in normal and atherosclerotic rabbit aorta. Arteriosclerosis, 10, 996-1009.

Zastrow, M. S., Flaherty, D. B., Benian, G. M., & Wilson, K. L. (2006). Nuclear titin interacts with A- and B-type lamins in vitro and in vivo. J.Cell Sci., 119, 239-249.

BIOGRAPHICAL SKETCH

Education

2001-2007 Doctor of Philosophy, Florida State University 1996-2000 Bachelor of Science, Florida State University, Tallahassee, FL 1991-1996 High School Diploma, The Benjamin School, North Palm Beach, FL

Teaching Experience

2003-2007 Graduate Teaching Instructor, BSC1005, General Biology Lecture 2004 Graduate Teaching Assistant, PCB 3134, Cell Structure and Function Lecture 2004 Graduate Teaching Assistant, PCB 4023L, Molecular Biology Lab 2001-2002 Graduate Teaching Assistant, BSC1005L, General Biology Lab

Research Experience

2001-2007 Graduate Research Associate Smooth muscle titin interactions with alpha-actinin Dr. Thomas C.S Keller III, Florida State University

2001 James R. Fisher Summer Fellowship Molecular investigation of a smooth muscle cell organizing protein American Cancer Society Dr. Thomas C.S Keller III, Florida State University

Publications Chi, R. J., Olenych, S. G., Kim, K., and Keller, T. C., III Smooth muscle alpha-actinin interaction with smitin Int. J. Biochem. Cell Biol. 37, 1470-1482., 2005 Chi, R. J. and Keller, T.C.S II. Smitin-myosin II coassembly arrays in vitro. In: Cell Biology Protocols, J.Robin Harris, ed., Wiley Press, Hoboken, NJ, pp. 369-371. 2005. Chi, R. J. Simon, A..R. and Keller, T.C.S II. Smooth muscle titin Zq domain interaction with smooth muscle alpha-actinin. (Submitted to Int. J. Biochem. Cell Biol)

Abstracts Kim,K; Chi, R; Keller and TCS Keller III. Smitin, a titin-like component of the smooth muscle contractile apparatus Mol Biol Cell, 12:864 Suppl. S NOV 2001

Chi, RJ; Kim, K; Olenych, SG, TCS Keller III. Smitin, a titin-like component of the smooth muscle contractile apparatus, interacts with alpha-actinin Mol Biol Cell, 13: 220 NOV 2002

Chi, RJ; Olenych, SG, TCS Keller III. Interactions of alpha-actinin with titin and titin-like proteins Mol Biol Cell, 14: 1722 NOV 2003

Chi, RJ; Olenych, SG, TCS Keller III. Mapping smooth muscle alpha-actinin smitin binding sites ASCB Supplemental CD, 2004

Chi, RJ; Olenych, SG, TCS Keller III. Interaction of Smitin Zq Domain with Alpha-acitnin Central Rod Spectrin Repeat-like R2-R3 Loops. ASCB Supplemental CD, 2005

Chi, RJ; Simon AR, Olenych, SG, TCS Keller III. Smitin Interactions with Smooth Muscle Contractile Apparatus Proteins ASCB Supplemental CD, 2006

Honors and Awards

2001 James R. Fisher Summer Fellowship, Molecular investigation of a smooth muscle cell organizing protein, American Cancer Society

2005 Outstanding Teaching Assistant Award, Florida State University

2005 Brenda Weems-Bennison University Memorial Scholarship Recipient, FSU

2005 Graduate Student Publication Award, Florida State University

2006 FSU Dissertation Grant Award Recipient, Florida State University

2005-2007 Predoctoral Fellowship, American Heart Association, FL-Puerto Rico Affiliate

Professional Societies

2001-Present Member of the American Society for Cell Biology