BIOCHEMICAL AND BIOPHYSICAL CHARACTERIZATION OF THE HAIR CELL’S - BUNDLING

By XU HAN

Submitted in partial fulfillment of the requirements For the degree of Master of Science

Thesis Advisor: Dr. Brian M. McDermott Jr.

Department of Biology CASE WESTERN RESERVE UNIVERSITY

May, 2014

CASE WESTERN RESERVE UNIVERSITY

SCHOOL OF GRADUATE STUDIES

We hereby approve the thesis/dissertation of Xu Han

candidate for the degree of Master of Science.

Committee Chair Roy Ritzmann

Committee Member Brian M. McDermott

Committee Member Vera Moiseenkova-Bell

Committee Member Emmitt Jolly

Date of Defense March 24th 2014

*We also certify that written approval has been obtained for any proprietary material contained therein

Table of Contents

Abstract ------5

Introduction ------6 Mechanism of hearing ------7 The structure of hair cell ------9 Actin ------15 Fascin ------20 Espin ------27

Material and Methods ------35 1. Material ------35 1.1 Chemical reagents ------35 1.1.1 Cell culture ------35 1.1.2 purification ------35 1.1.2.1 Maltose-Binding Protein (MBP)-fascin 2b ------35 1.1.2.2 Hexa polyhistidine-tagged (His-tag) espin 2b------35 1.1.3 Actin-binding and –bundling assay ------36 1.1.4 SDS-PAGE ------36 1.1.5 Ruby protein gel staining ------36 1.1.6 Negative staining TEM ------37 1.2 Buffers ------37 1.2.1 General buffers ------37 1.2.2 Cell culture ------37 1.2.3 Protein purification ------37 1.2.3.1 MBP-fascin 2b ------38 1.2.3.2 His-tag espin 2b ------38 1.2.4 Actin-binding assay and actin-bundling assay ------38 1.2.4.1 MBP-fascin 2b alone ------38 1.2.4.2 His-tag espin 2b alone ------39 1.2.4.3 MBP-fascin 2b and His-tag espin 2b ------39 1.2.5 SDS-PAGE ------39 1.2.6 Ruby protein gel staining ------39 2 Methods ------40 2.1 Cell culture ------40 2.2 Protein purification ------40 2.2.1 MBP-fascin 2b ------40 2.2.2 His-tag espin 2b ------41 2.3 Actin protein preparation ------42 2.4 Negative stain transmission electron microscopy ------42 2.4.1 Sample preparation ------42 2.4.2 Microscope alignment and data collection ------43 2.5 Cryo-electron tomography ------43 2.5.1 Sample preparation ------43 2.5.2 Microscope alignment, calibration, and data collection ------43

1 2.6 Actin-binding assay and actin-bundling assay ------44 2.7 SDS-PAGE and Ruby protein gel staining ------45 2.8 Data processing and statistical analysis ------45

Results ------46 Part 1 Fascin 2b ------46 Maltose-Binding Protein (MBP)-fascin 2b protein overexpression and confirmation of binding activity ------46 Confirmation of MBP-Fascin 2b’s actin-binding and –bundling activity------47 Confirmation of the morphology of the MBP-fascin 2b-actin bundle and examination of the actin-bundling ability of MBP-fascin 2b using negative staining transmission electron microscopy ------49 Cryo-electron tomography of the fascin 2b-actin bundle------53

Part 2 Espin 2b ------55 Purification of hexa polyhistidine-tagged espin 2b protein ------55 Confirmation of His-tag espin 2b actin-binding and –bundling activity------56 Observation of the morphology of His-tag espin 2b-actin bundle and examination of the actin-bundling ability of His-tag espin 2b using negative staining transmission electron microscopy------57 Cryo-electron tomography of the His-tag espin 2b-actin bundle ------62

Part 3 Fascin 2b and Espin 2b ------64 Comparison of the actin-bundling capacity of fascin 2b and espin 2b ------64 Observation of the morphology of actin bundles formed by both MBP-fascin 2b and His- tag espin 2b using negative staining transmission electron microscopy ------65 Examination of fascin 2b and espin 2b coordinated actin-bundling using negative staining TEM------67 Cryo-electron tomography of the fascin 2b-espin 2b-actin bundle ------76

Discussion ------77

Reference ------89

2 List of Figures

Figure 1 ------8 Figure 2 ------9 Figure 3 ------18 Figure 4 ------21 Figure 5 ------28 Figure 6 ------47 Figure 7 ------49 Figure 8 ------50 Figure 9 ------52 Figure 10 ------54 Figure 11 ------56 Figure 12 ------57 Figure 13 ------59 Figure 14 ------61 Figure 15 ------63 Figure 16 ------65 Figure 17 ------66 Figure 18-1 ------69 Figure 18-2 ------70 Figure 18-3 ------72 Figure 18-4 ------74 Figure 19 ------75 Figure 20 ------76

3 Acknowledgment

My deepest thanks goes to the people who made this research possible. Dr. Brian McDermott, who accepted me as his Master’s student two-and-a-half years ago and gave me this chance to explore my research topic independently, has always supported me, and encourages me with great patience. Dr. Vera Moiseenkova-Bell, who made the structural biology part of my thesis achievable, provided me with a priceless opportunity to get training with transmission electron microscopy and cryo-electron tomography, by which I found my research interest for life. I would also like to give my deepest thanks to Dr. Emmitt Jolly and Dr. Roy Ritzmann for their support as my committee members and guiding me through the process of the thesis defense.

I have been very fortunate to work with some incredible people around campus. Heather Holdaway, electron microscope manager of the CCMSB, gave me training on TEM and worked with me on cryo-tomography. Dr. Gustavo Gomez, my scientific guide in my first year, who started this project, passed on all his knowledge about this project selflessly and, with great patience, showed me everything step by step. He was a constant source of support and guided me in learning new techniques and introduced me to people who had knowledge in relevant fields and key equipment. Kevin Huynh has always helped me in troubleshooting the protein-related and structural biology-related questions. Tara Fox showed me the way to process tomography data using ETomo and UCSF Chimera step by step, which saved me at least two months if I tried to explore this by myself. I also owe thanks to Dr. Tingwei Mu, who is my biophysics consultant and the person who I could talk to about my project and my life. Summer Watterson trained me and gave me permission to use the Department of Biophysics facilities, including the FPLC, the ultracentrifuge, and the shaker. I would also like to thank Julia Brown, the coordinator of the graduate program in the Department of Biology, for her heartfelt help related to both studying and living in Cleveland. It also has been great honor for me to work with everyone in the McDermott lab. They are lots of fun, and I have always looked forward to coming to the lab and working. I would like to thank Zongwei Chen, Shih- Wei Chou, Carol Fernando, Phil Hwang, and Lana Pollock for their help, giving me suggestions, and for corrections on both my thesis and defense presentation.

My deepest gratitude goes to my parents who were hesitant to let me wander so far from home but at the same time supported me as I chased my dream. They provided me with endless love and reassurance. I would have never become the person I am without their teaching and guidance and would not be able to accomplish this task without them as my warmest harbor. Gustavo Gomez, whom I have been fortunate to meet across the Pacific Ocean, is always my solid support to cheer for my successes, to share the failures with me, and to provide me 24/7 care when I stay in the States without my family.

4

Biochemical and Biophysical Characterization of the Hair Cell’s

Actin-Bundling Proteins

Abstract

by

XU HAN

Hair cells, the mechanoreceptors of the inner ear, transduce both auditory and vestibular signals allowing us to detect sound and accelerations of the head. As an actin-based protruding structure, a stereocilium requires structural proteins, particularly actin and its- bundling proteins, in order to properly maintain the stereociliary bundle shape for mechanotransduction of sound. Fascin 2b and espin 2b are known actin cross-linkers in stereocilia. Actin-binding and actin-bundling assays were carried out to confirm their actin-binding and -bundling capacity. Negative staining transmission electron microscopy was used to determine the changes in the thicknesses of actin bundles with titrated fascin

2b, espin 2b, or different combinations of the two proteins. Cryo-electron tomography was also carried out to determine the 3-dimensional structures of the actin bundles formed by fascin 2b, espin 2b, or by both. My studies give insight into the actin core of the stereocilium, which is required for hearing in vertebrates.

5 Introduction

Among the five major senses, hearing is of great importance. According to a report from the Hearing Loss Association of America, hearing loss is a major public health issue, which ranks third place among the most common physical conditions after arthritis and heart disease. Based on statistical data provided by the National Institute on Deafness and

Other Communication Disorders (NIDCD), about 17% of American adults (36 million) reported hearing loss of some degree. One third of people suffer hearing loss by age 65.

Furthermore, 2 to 3 per 1000 children have hearing impairment. Hearing loss may be caused by many factors including age-related progressive hearing loss and acute hearing loss (AHL), which is often caused by genetic aspects or exposure to ototoxic drugs. The hair cells in the cochlea are particularly important to the process of hearing because of their role as the mechanical transducer. There are many factors that can cause hearing loss, and they can be related to hair cells in the cochlea, including hair cell dysfunction, structural alteration of hair cells, and protein mutation within hair cells. Similarly, vestibular hair cells in the semicircular canals in the inner ear are responsible for the maintenance of balance. Loss or damage to hair cells may result in loss of hearing, balance, or both.

6 Mechanism of hearing

Hearing is a multistep process. The external ear, including the auricle, acts as a reflector and helps to capture the sound waves effectively. Mechanical energy passes through the external auditory meatus leading to the vibration of the tympanum, which conducts the energy into the air-filled middle ear. Energy traverses the middle ear and causes the vibration of three tiny ossicles: the malleus (hammer), incus (anvil), and stapes (stirrup).

With the assistance of the three ossicles, the stimuli eventually move into the inner ear

(Figure. 1A). The cochlea, snail shell-shaped structure of the inner ear, has approximately 16,000 hair cells, which serve as mechanical transducers. Sound stimuli cause the vibration of the basilar membrane, which leads to the movement of the organ of

Corti and the overlying tectorial membrane. The up and down movements of the organ of

Corti and the tectorial membrane are accompanied by the back and forth shearing motion of the upper surface of the organ of Corti. The deflection of the protrusion from the apical surface of the hair cell, called the hair bundle, is caused by the back and forth motion of organ of Corti and initiates mechanoelectrical transduction. Transduction begins from the mechanotransduction ion channel at the tips of the stereocilia. The opening and closing of the mechanical transduction channels are regulated by tension in elastic structures, which are also called gating springs, located in hair bundle (Corey and Hudspeth, 1983).

Deflection of the hair bundle applies mechanical tension on the tip link and opens the mechanically sensitive ion channels, allowing the influx of cations from the endolymph bathing the hair bundle. Endolymph is the watery fluid in the membranous labyrinth of the inner ear. The inward ion flow causes the depolarization of the hair cell, which

7 transforms the mechanical stimuli to an electrical signal. The endocochlear potential

(EP), which is the positive voltage seen in the endolymphatic space of the cochlear, is generated by the stria vescularis, which also helps to maintain the ionic homeostasis within the endolymph. The influx of Ca2+ triggers the cell to release synaptic vesicles, including excitatory neurotransmitters, to the presynaptic active zones located at the basolateral membrane of the cell, where the dendrites of the spiral ganglion contact the hair cell. The spiral ganglion transmits the electrical signal from the cochlea to the central nervous system. The brain analyzes auditory information to create an internal representation of the external world (Kandel et al., 2012) (Figure. 1B).

Figure 1. Structure of the human ear. (A) The external ear focuses sound into the external auditory meatus. The sound waves then vibrate the tympanum and the energy then passes through the air-filled middle ear via three tiny linked bones. Vibration stimulates the hearing organ of the inner ear, the cochlea. (B) Schematic of the human organ of Corti on the basilar membrane of the cochlea in the inner ear. It contains hair cells as the receptor cells. There are two kinds of hair cells: one row of inner hair cells sitting on the modiolar side of the arch of the Corti and three rows of outer hair cells (Schwander et al., 2010).

8 The Structure of the Hair cell

The unique structure of the hair cell is the foundation for its ability to accomplish its function as a mechanical transducer. The cell body, or the soma, of the hair cell is columnar or flask-shaped, lacking both dendrites and an axon (Kandel et al., 2012)

(Figure. 2A). The hair cell connects to its surrounding nonsensory supporting cells, which have stubbles of microvilli projecting from their apical surface, by the tight adherent junctions around its apex (Corwin and Warchol, 1991; Tliney et al., 1992). The apical aspect of the hair cell is soaked in a special saline solution, the endolymph. The tight junctions of the sensory epithelium (Nunes et al., 2006) are responsible for separating the endolymph from the perilymph, which contacts the basolateral region of the hair cell (Kandel et al., 2012).

Figure 2. Structure of a vertebrate hair cell. (A) Illustrates a hair cell from a frog’s internal ear. The cylindrical hair cell is surrounded by supporting cells, and joined to them by a junctional complex. The hair bundle is a mechanically sensitive organelle that

9 protrudes from the cell’s apical surface. About 60 stereocilia with varying lengths are arranged in a staircase shape to form the hair bundle. A single kinocilium, which is an axonemal structure, stands at the bundle’s tallest edge. (B) A zoomed-in view of the molecular structure of the two adjacent stereocilia. The proteins are mentioned in the text (Schwander et al., 2010).

A thick interlinked actin filament meshwork, also known as the cuticular plate, is located at the apical part of the hair cell (Kandel et al., 2012). The cuticular plate contains actin as well as (Drenckhahn et al., 1982). The proteins are packed so dense in the cuticular plate that even the organelles as tiny as ribosomes are expelled from this region.

Long and relatively straight single actin filaments pass through the whole region

(DeRosier and Tilney 1989). Smaller actin fragments in between help to connect actin filaments with the same or opposite polarity (DeRosier and Tilney 1989). In 1982,

Hirokawa and Tilney first used the quick-freezing and deep-etching techniques to show the interaction between the actin filaments in the cuticular plate and the inner-surface of the plasma membrane of the chicken cochlea hair cells (Hirokawa and Tilney 1982). The connections at the end and the side of the actin filaments have branched termination, which reside at the cytoplasmic-surface of the plasma membrane. also appeared in the cuticular plate region, which connected this region to the axial in the hair cell (Jaeger et al., 1994; Antonellis et al., 2014).

One of the most important functions of the cuticular plate in the mature hair cell is to provide a foundation and anchor for stereocilia (Kandel et al., 2012). Stereocilia taper near their bases, as the protrusions join the cell body (Kandel et al., 2012). In each stereocilium, only a small portion of the actin filaments uninterruptedly pass through the constriction formed by the taper structure and bury themselves into the cuticular plate as

10 a densely-packed rootlet (Kandel et al., 2012). This thin cluster of is formed by filamentous actin (F-actin), only 18 to 30 filaments in the alligator lizard

(Tilney et al., 1980), vertically inserts into the cuticular plate, and anchors the stereocilium in the cuticular plate (Corwin and Warchol, 1991). , TRIO and

F-actin-binding protein (TRIOBP), and are concentrated around the rootlets and may help to stabilize them (Corwin and Warchol, 1991; Tilney et al., 1992; Furness et al.,

2008, Kitajiri et al., 2010).

The hair bundle, which is the protrusion from the apical surface of the hair cell, is formed by about sixty stereocilia (Kandel et al., 2012). The hair bundle is the receptor that receives mechanical stimuli. All of the hair bundles are oriented with the tallest stereocilium pointing to the opposite direction of the center of the cochlea (Kandel et al.,

2012). This polarity is essential for hair cells to accomplish their work: the hair bundles only deflect to the direction of the longest stereocilium and the shortest stereocilium, which increases the possibility for the hair cell to open the mechanotransduction channels and close them, respectively (Hudspeth and Corey, 1977). The hair bundle protrudes several micrometers from the cell surface. The tallest row of the stereocilia is located on one side of the hair cell, with each subsequent row decreasing in height. The whole hair bundle forms a staircase shape (Kandel et al., 2012).

One insight into the understanding of stereocilia is the discovery of filamentous connections, also known as the tip links (Corwin and Warchol, 1991). A tip link projects from the top of a shorter stereocilium, and attaches to the slightly higher shaft of the

11 closest longer stereocilium (Osborne et al., 1984; Pickles et al., 1984). Upon deflection of the hair bundle towards the tallest stereocilium, the tip links are thought to stretch open the mechanically-gated channels at the tips of the stereocilia (Kandel et al., 2012).

During the development of the immature hair cell, the hair bundles in the cochlea each include a single -based cilium, kinocilium, which localizes next to the tallest stereocilium. Extracellular filaments connect the kinocilium with the stereocilia to form a whole bundle. The kinocilium has a bulbous swelling at its tip. The kinocilium has an axoneme as its core, array of nine paired microtubules, and sometimes an extra central pair of microtubules (Kandel et al., 2012). The existence of the kinocilium is crucial for hair cell morphogenesis: before the morphogenesis of the hair bundle starts, the kinocilium is located in the center of the apical surface of the hair cell with 20-300 microvilli surrounding it. The kinocilium then moves to the periphery of the developing hair cell and determines hair bundle orientation. Meanwhile, surrounding microvilli begin elongation in order to form stereocilia. The positioning of the kinocilium is nonrandom and plays a critical role in the development of hair bundle polarity. The molecules involved in regulating kinocilium movement and bundle polarity are not clear (Kandel et al., 2012).

The actin filaments of the developing stereocilia initiate from the barbed end for growth

(Flock & Cheung 1977). Stereocilia can be thought of as highly specialized microvilli.

Similar to other actin-based protrusions such as filopodia and microvilli, stereocilia are formed by actin filaments with a single polarity involving all barbed (plus) ends at the

12 tips of the stereocilia (Tilney et al., 1992). Electron microscopy data indicates that the actin core of the stereocilium, which is covered by a tubular sheath of plasma membrane, is arranged in a paracrystalline array; helices are densely packed (Tilney et al., 1980).

Although derived from microvilli, stereocilia have several key differences. Firstly, stereocilia are generally longer than microvilli. Secondly, stereocilia taper near their bases. Third, most stereocilia are thicker than microvilli and contain more strands of F- actin. The majority of the length of the stereocilia keeps a relatively stable diameter, which is about 0.4 µm wide. At the basal insertion of the stereocilia, most of the actin filaments terminate near the plasma membrane. The number of the actin filaments in the bundle decreases from several hundred to only a couple of dozen, which decreases the diameter of stereocilium to ~0.1 µm. The taper is critical for the pivot movement around the basal insertion. This region is also the most vulnerable structure of the hair cell when exposed to a deafeningly loud noise (Kandel et al., 2012). Recent studies suggest that minus end–directed molecular motor 6 (MYO6) and the protein tyrosine phosphatase receptor Q (PTPRQ) regulate the formation and maintenance of the taper region. Malfunction of these ’s products cause the deafness related to the taper

(Hasson et al., 1997; Goodyear et al., 2003; Sakaguchi et al., 2008). Since the longest stereocilia can reach up to 100 µm in length (Silver et al., 1998), regulated protein transportation would be an elegant explanation to supply actin assembly regulators to the barbed ends of actin filaments. Myosin 15A (MYO15A) is one of the motor proteins which is thought to be involved in the regulation of stereociliary growth as well as cooperate with an adaptor protein, whirlin, in this process (Probst et al., 1998; Wang et al., 1998; Mburu et al., 2003). It has been proposed that MYO15A binds to whirlin as a

13 cargo and walks along the F-actin to the tip of the stereocilia (Rzadzinska et al., 2004;

Belyantseva et al., 2005; Delprat et al., 2005; Kikkawa et al., 2005; Schneider et al.,

2006, Belyantseva et al., 2005). In other tissues such as erythrocytes and neurons of outer hair cells (OHCs), whirlin can bind to some actin assembly-regulating proteins, whose isoforms are confirmed to be expressed in inner hair cells (IHCs) as well (Mburu et al.,

2006). Thus, MYO15 and whirlin may act similarly in both the outer and inner hair cells

(Marfatiz et al., 1995; Biederer and Sudhof, 2001). MYO7A is another motor protein that might be involved in the process of regulation of stereociliary growth. MYO7A mutations can cause hearing loss, defects of hair bundle morphology, as well as excessive elongation of stereocilia (Gibson et al., 1995; Liu et al., 1997; Weil et al., 1997). MYO7A may also regulate the transportation of proteins such as twinfilin-2, actin-capping protein, which can restrict actin assembly (Palmgren et al., 2001; Paavilainen et al., 2007;

Rzadzinska et al., 2009).

14 Actin

The F-actin in the hair bundle include β– and γ-actin, which are cross-linked by actin- binding/bundling proteins (ABPs), such as espin, fimbrin, plastin 2, plastin 3, GRXCR1,

TRIOBP, fascin 1, fascin 2, espin-like protein, and Xin- related protein 2 (Tilney et al.,

1989; Zine et al., 1995; Zheng et al., 2000; Daudet and Lebart, 2002; Li et al., 2004;

Kitajiri et al., 2010; Shin et al., 2010; Chou et al., 2011). These actin-binding proteins form cross-bridges to maintain and stabilize the paracrystalline arrays in order to keep each stereocilium in a rigid, cylindrical shape (Flock et al., 1977; Flock et al., 1981;

Flock et al., 1982; Tilney et al., 1983; Sobin and Flock 1983; Slepecky and Chamberlain

1985; Tilney et al., 1989). Villin, which appears with actin and actin-binding proteins in the cytoskeleton of the intestinal microvilli, does not exist in the stereocilia (Tilney et al.,

1989). Cytoskeletal actin and its binding and bundling proteins (cross-linkers) are important in maintaining the stereocilia structure, and thus also are critical for the hair bundle to complete its proper function as a receptor. These proteins are also called structural proteins. They are essential and critical to the hair cell, like bricks and concrete to a building.

In short, the stereociliary core is in a paracrystalline array (Figure. 3D and E), which is built from densely cross-linked actin filaments (Figure. 2B). Every stereocilium anchors into the cuticular plate through a tightly bundled rootlet. Mainly γ-actin-based filaments form the mesh network of the cuticular plate of the alligator lizard (DeRosier and Tilney,

1989). This is a different arrangement from the one in the stereociliary actin core formed

15 by both β– and γ- F-actin and the array of F-actin in the rootlet (DeRosier and Tilney,

1989; Hofer et al., 1997; Furness et al., 2005).

Actin and its bundling proteins are so indispensible for hearing that experimental data indicates 19 out of 57 genes, whose mutated alleles can cause , encode proteins that directly or indirectly interact with actin (Drummond et al., 2012).

There are six different in the mammalian and avian genome. Four of them are muscle-specific (ACTA1, ACTC1, ACTA2, ACTG2); while the other two encode cytoplasmic β- and γ-actin (ACTB, ACTG1). A syndrome, which includes photosensitivity and intellectual impairment in addition to the recurrent infections resulting from neutrophil dysfunction, has deafness as the most significant feature. This is caused by a mutated β-actin in humans (Nunoi et al., 1999; Procaccio et al., 2006;

Riviere et al., 2011). Mutated γ-actin trigger syndromic and nonsyndromic progressive hearing loss (van Wijk et al., 2003; Zhu et al., 2003; Rendtorff et al., 2006; Liu et al.,

2008; de Heer et al., 2009; Morin et al., 2009; Riviere et al., 2011). Although encoded by six genes, mammalian actin proteins are 90% identical to each other (Khaitlina, 2001). As two 375-amino acid proteins, β- and γ-actin only have 4 amino acids among their first 10 residues that are different from each other. Both of these are 100% conserved among mammals and birds (Sheterline et al., 1998). β- actin is the predominant actin in almost all tissue types in the body except in the brush border of intestinal epithelial cells and hair cells in the inner ear. In young chicken auditory hair cells (Hofer et al., 1997) and adult guinea pig cochlear sensory and supporting cells (Khaitlina, 2001; Furness et al., 2005),

β- and γ-actin are present in a 1:2 ratio; although, the relative amount of these two

16 proteins changes during development (Tilney et al., 1980; Hofer et al., 1997; Beyer et al.,

2000; Furness et al., 2005).

Actin is a 42-kDa protein existing in cells either in monomeric (globular or G-actin) or polymeric form (filamentous or F-actin) (Drummond et al., 2012) (Figure. 3A). Every actin monomer has two main domains, which have two subdomains (Schutt et al., 1993), with an ATP-binding pocket in the center of the protein (Sheterline et al., 1998). Under certain conditions, ATP-G-actins polymerize to form F-actin (Figure. 3B and C), which is used to build stable or dynamic cell structures. Actin filaments are helical and polar, with barbed (plus) and pointed (minus) ends. A new ATP-actin monomer is added on the barbed-end of an activated filament; meanwhile, an ADP-actin monomer is released from the pointed-end (Sheterline et al., 1998; Bugyi and Carlier, 2010). Actin filament assembly generates the force for the cells to directly move or change their shape

(Sheterline et al., 1998; Welch and Mullins, 2002; Pollard and Borisy, 2003; Ridley,

2011).

17

Figure 3. Actin in stereocilia. (A) A 3-dimensional structure of actin protein (Drummond et al., 2012). (B) Actin filament formed by actin monomers. (C) Actin filament under transmission electron microscope. (D) TEM image shows the parallel and tightly packed actin filaments form the core of the stereocilium (Mogensen et al., 2007). (E) and (F) The cross-sections of the stereocilium illustrate the hexagonal arrangement of the F-actin (Mogensen et al., 2007).

Understanding the different biochemical properties of β- and γ-actin is an active research field (Drummond et al., 2012). β- and γ-actins are different in their nucleotide exchange rate and ion-dependent polymerization speed (Bergeron et al., 2010). In the presence of calcium in vitro, purified β-actin polymerizes approximately twice as fast as γ-actin. This difference causes faster rates of filament nucleation and elongation. In addition, the rates

18 of phosphate release and depolymerization of actin monomers in β-actin-based filaments is faster than in γ-actin-based filaments. Interestingly, when calcium gets replaced by magnesium, these differences are no longer as significant, which indicates the kinetics may rely on the intracellular microenvironment (Bergeron et al., 2010).

In the inner ear, the loss of one cytoplasmic actin and the resulting hair cell degeneration cannot be compensated by the existence of the other actin proteins or the up-regulation of total actin level (Belyantseva et al., 2009; Perrin et al., 2010). Different from muscle actins, either β- or γ-actin has the ability to initially build a functional stereocilium.

However, both of them are each required to maintain the structure of stereocilia in the long term (Bergeron et al., 2010). This suggests that these two cytoplasmic actins have their own unique functions, at least in the inner ear (Drummond et al., 2012)

19 Fascin

Fascins are a group of actin cross-linking proteins, which have structural uniqueness and have been evolutionarily conserved. The actin bundles formed with the help of fascin are of critical importance to various sub-cellular structures and to cell morphology. Fascin mainly plays an important role in two actin-based structures: dynamic cortical cell protrusions and cytoplasmic bundles. Cortical structures, such as filopodia, spikes, lamellipodia ribs and the dendrites of dendritic cells, participate in cell-matrix adherence, cell-cell interactions, and cell migrations. Cytoplasmic microfilament bundles are involved in cellular architecture (Kureishy et al., 2002).

Fascin was first identified in the 1970’s. As a 55 kDa actin-binding protein, fascin was purified from an extract of the cytoplasm of sea urchin oocytes or coelomocytes (Bryan and Kane, 1978; Otto et al., 1979). This protein was denominated as fascin for its ability to pack F-actin into tight and stable bundles (Latin, fasiculus, bundle) (Otto et al., 1979).

Other studies indicated that there was a similar protein existing in sand dollar oocytes and starfish sperm. In the early 1990’s, cDNA sequences showed that echinoderm fascin was related to singed protein in drosophila and a 55-kDa actin-binding protein in humans. In addition, its homologues are widely spread in other vertebrate species (Paterson and

O’Hare, 1991; Bryan et al., 1993; Mosialos et al., 1994; Holthuis et al., 1994). Ten years ago, it was concluded that the fascin gene family exists in all vertebrates (Kureishy et al.,

2002).

20

Figure 4. Fascin protein (A) illustrations of members of the fascin family. There are three members in the fascin family: fascin 1, fascin 2 (retinal fascin), and fascin 3 (testis fascin). All of them contain four beta-trefoil domains and a putative PKC phosphorylation site in the first beta-trefoil domain. (B) and (C) are 3-dimensional reconstructions of fascin-1. Two actin-binding sites are pointed out. Actin-binding site 1 includes an N- and C-terminus and actin-binding site 2 is formed by part of the beta- trefoil domain 1 and part of the beta-trefoil domain 2 (Yang et al., 2012). (D) through (G)

21 show that fascin 2b is specifically located in zebrafish hair cell stereocilia (Chou et al., 2011).

The first fascin gene product cloned, fascin 1, is highly conserved in both vertebrates and invertebrates (Paterson and O’Hare, 1991; Bryan et al., 1993; Mosialos et al., 1994;

Holthuis et al., 1994; Duh et al., 1994; Edwards et al., 1995). The second fascin-related gene product, fascin 2, which possesses 56% amino acid identity to human fascin 1, was subcloned from human and bovine retinas, and was therefore named retina fascin

(Saishin et al., 1997; Tubb et al., 2000). The third fascin-related gene product, fascin 3, was cloned from human testis. The testis fascin only has 27% sequence identity to the human fascin 1, and 27% identity to retina fascin (Kureishy et al., 2002) (Figure. 4A).

Fascin 1 is expressed in most organized structures of vertebrates, especially in the brain, ovary, and testis (Mosialos et al., 1994; Holthuis et al., 1994; Edwards et al., 1995).

Fascin 2 is specifically expressed in inner and outer segments of photoreceptor cells in the retina (Saishin et al., 1997, Saishin et al., 2000). Because all of these segments are joined together by an F-actin-rich cilium (Chaitin et al., 1984), it is possible that fascin 2 plays a role in the morphological specification of the photoreceptor cell. Fascin 3 contributes to the spermatid development (Kollers et al., 2006) and specifically localizes to the elongated spermatid head (Tubb et al., 2002). In humans, the fascin 1 gene

(FSCN1) and fascin 2 gene (FSCN2) are located on different : FSCN1 maps to 7q22 (Duh et al., 1994), but FSCN2 maps to chromosome 17q25 (Tubb et al., 2000, Saishin et al., 2000). Both of these genes have a neighbor, which encodes a member of the actin family (Tubb et al., 2000). The conservation of these two fascin genes and their neighboring genes throughout vertebrates indicate that the fascin 1 and 2

22 genes might have arisen in the early vertebrate lineages before genomic-wide duplication

(Hashimoto et al., 2011). In contrast, FSCN3 has a unique sequence-conserved neighboring gene, and FSCN3 has very low sequence identity with FSCN1 and FSCN2.

Based on the analysis of FSCN3’s neighboring gene, FSCN3 is proposed to have arisen during early vertebrate evolution (Hashimoto et al., 2011).

The structure of the fascin 1 protein was first elucidated by a transmission electron microscope (TEM) rotary shadowing experiment. This study showed that purified FSCN1 product is a globular monomer (Yamashiro-Matsumura et al., 1985). Sequence pattern and protein structure alignment analysis predicted FSCN1 belongs to the β-trefoil protein group (Ponting et al., 2000), characterized by β-trefoil folds (Murzin et al., 1992). The β- trefoil domain is found in numerous functionally unrelated proteins (Murzin et al., 1992).

Structural alignment had also forecasted that FSCN1 has four β-trefoil domains with 1 and 3 being larger than 2 and 4 (Ponting et al., 2000). In 1999, Fedorov et al., reconstructed the FSCN1 crystal structure with a resolution of 2.9 Å (Sedeh et al., 1999).

This structure of FSCN1 also revealed that there are four β-trefoil domains, which form 2 lobes, corresponding to β-trefoil-1 and -2 as well as β-trefoil-3, and -4, respectively.

These two lobes form a 56° angle with each other, with potential for allosteric coupling between β-trefoil-1 and -3 (Hashimoto et al., 2011).

Multiple sequence alignments for fascin revealed the characteristics of the conserved sequence and also demonstrated the difference between fascin 1, 2 and 3 (Kureishy et al.,

2002). In all fascin proteins, the most highly conserved region is located between amino

23 acid residues 11 and 50, which contains a putative consensus motif for phosphorylation by protein kinase C (PKC) (Kureishy et al., 2002) (Figure. 4A). This putative phosphorylation site exists among all of the members of the fascin family. In fascin 1, experimental evidence well supports the importance of this motif as a regulatory site.

Residue Ser-39, an amino acid that is exposed on an external loop of the first β-trefoil fold, was identified as the main phosphorylation site by PKC (Kureishy et al., 2002).

Through mutational analysis, Ser-39 was shown to regulate actin-binding ability in vitro

(Ono et al., 1997) and form actin-based protrusions in matrix-adherent cells (Adams et al., 1999). In zebrafish, two paralogous orthologs of FSCN2, and their phosphorylation sites were identified (Lin-Jones et al., 2007). Fascin 2b, one of the paralogs of fascin 2, showed regulation of actin-binding and -bundling properties by mutation of the putative phosphorylation site (Chou et al., 2011) (Figure. 4D-G).

As a monomer, fascin has to contain at least two actin-binding sites in order to cross-link

F-actin to form an actin bundle (Bryan and Kane, 1978). Based on the results from limited proteolysis, one of the binding sites was deduced to be located between residues

227 to 493 (Edwards et al., 1995). The mapping for this binding site was strongly supported by in vivo data (Cant et al., 1996). The second actin-binding site was firstly assumed to be located between residues 29 and 42 in the first β-trefoil fold. This region is similar to the actin-binding site of mytistoylated alanine-rich C-kinase substrate

(MARCKs) (Mosialos et al., 1994). In 2012, the Huang group reconstructed a fine crystal structure of human fascin 1 protein with the resolution of 2.2 Å to uncover the actin- binding sites. All four β-trefoil domains are involved in actin-bundling activity. Residues

24 from the N and C termini form actin-binding site 1. This binding site includes the cleft formed by β-trefoil domain 1 and 4. The second actin-binding site is located spatially opposite to actin-binding site 1. Residues from β-trefoil domains 1 and 2 are included in this binding site. They also found a smaller surface area located at the bottom of the molecule, which contains residues from β-trefoil domain 3 that is a potential, third, actin- binding site or a part of the other two actin-binding sites that somehow contributes to the actin-bundling activity (Yang et al., 2012) (Figure. 4B and C).

Understanding of fascin’s function and regulation in the cell can be obtained by studying its ultrastructure and mechanical properties (Kureishy et al., 2002). A needle-like actin bundle can be formed just by simply mixing fascin and F-actin together in a proper salt environment (Bryan and Kane, 1978). Under the T.E.M., F-actin bundles formed with invertebrate fascin have unipolar F-actin filaments, which are tightly packed together with an 8 nm inter-filament distance (Bryan and Kane, 1978; Maekawa et al., 1982; Cant et al., 1994). A rigid hexagonal arrangement of actin filaments can be produced by fascin with an 11-nm periodic transverse banding pattern on the bundle. Depending on the species source of fascin, the distance between two banding patterns is slightly different

(Bryan and Kane, 1978; Maekawa et al., 1982; Edwards et al., 1995; Cant et al., 1994;

Edwards et al., 1995; Kureishy et al., 2002). The structures formed by F-actin and cross- linking proteins critically depend on the 3-dimensional conformation and the size of the cross-linker molecules (Matsudaira et al., 1994). Compared to the actin bundle formed by

α- and , very interestingly, filamentous bundles constructed by fascin show

25 tight arrangement and a highly ordered parallel F-actin bundle (Bryan and Kane, 1978;

Yamashiro-Matsumura et al., 1985).

Fascin 2 is specifically transcribed in the retina and localizes in the inner and outer segments of bovine photoreceptor cells (Tubb et al., 2000; Saishin et al., 1997; Saishin et al., 2000). Like fascin 1, fascin 2 cross-links F-actin to form actin bundles (Saishin et al.,

2000). In zebrafish and Xenopus, fascin 2 localizes to the inner segments of photoreceptor cells, longitudinal actin bundles, and calycal processes of rod inner segments (Hashimoto et al., 2011). Zebrafish have two fscn 2 paralogues, DrF2A and

DrF2B, which share 77% sequence identity. Fascin 2b showed more vigorous F-actin- binding and -bundling ability (Lin-Jones et al., 2007). In the zebrafish hair cell, fascin 2b mRNA is the predominant fascin 2 paralogue, which showed a higher level of expression than fascin 2a by in situ hybridization. Localizing specifically to zebrafish hair cell stereocilia, fascin 2b participates in forming actin bundles as a cross-linker (Chou et al.,

2011). However, a clearer image of the biological function of fascin 2 is still coming into focus.

26 Espin

Espin is a multifunctional actin-binding and -bundling protein, which is encoded by a single gene with multiple isoforms in the human. A short time after it was discovered, espin was found to have a high expression level in the parallel actin bundles (PAB) of hair cell stereocilia in the cochlea and vestibular system (Zheng et al., 2000) (Figure. 5B-

D). In mice and humans, espin mutation can cause hearing loss (Sekerková et al., 2006).

Besides hair cell stereocilia, espin was also identified in the microvillar PABs of other kinds of sensory cells, including taste receptor cells, solitary chemoreceptor cells, vomeronasal sensory neurons and Merkel cells (Sekerková et al., 2004; Sekerková et al.,

2005). Every domain in espin appears to have different functions and activities involved in regulating the organization, dynamic and signaling capabilities of PAB-containing specializations, which distinguishes espin from other actin-bundling proteins and makes them greatly adept to sensory cells. Espin is conserved throughout vertebrates, from pufferfish to human. Espin shows no sequence similarity to other actin-binding proteins except the forked protein in Drosophila (Tilney et al., 2005).

27

Figure 5. Espin protein. (A) Illustrated structure of the members in the espin family. All of the members contain an ABM domain, which helps binding and bundling F-actin. The ABM domain’s activity is Ca2+ resistant, which is significantly important for espin function in the hair cell. A WH2 domain also appears in all members of the espin family and helps to bind with G-actin. (B) through (D) show that espin is expressed specifically in the stereocilia of both inner and outer hair cells (Sekerková et al., 2006).

The first identified espin was a ~110 kDa actin filament-binding protein that is highly concentrated in the PAB of Sertoli cell ectoplasmic specializations (Bartles et al., 1996;

Chen et al., 1999). Espin got its name from this special intercellular junction: ectoplasmic specialization + -in. Very soon, a smaller ~30 kDa espin isoform was identified as a relatively minor, although high affinity, actin-bundling protein in the PAB of the brush border microvilli and renal proximal tubule (Bartles et al., 1998). Espins form their own actin-bundling protein family, all the members of which are encoded from a single gene.

The multiple isoforms vary in their size and ligand-binding sites (Sekerková et al., 2004;

Sekerková et al., 2006). Different translational start sites produce four major espin

28 isoforms from 110 kDa to 25 kDa, named espin 1 to 4, in order of decreasing size; additional variants produced by differential splicing are specified alphabetically

(Sekerková et al., 2004) (Figure. 5A). Although appearing in numerous actin-containing structures, espin is specifically concentrated in hair cell stereocilia and the microvilli in other sensory cells, where it neatly distributes along the cores of the parallel actin bundles

(Sekerková et al., 2004; Sekerková et al., 2005; Zheng et al., 2000).

During their formation, PABs of stereocilia are associated with espin (Sekerková et al.,

2006; Li et al., 2004). In the rat, throughout stereociliogenesis, the accumulation of espin increases noticeably, along with the elongation of the stereocilia and staircase shape formation (Sekerková et al., 2006; Li et al., 2004). It has been shown that this increasing accumulation of espin is involved in a complicated spatiotemporal pattern in which different espin isoforms are expressed at different times (Sekerková et al., 2006). This indicates that specific espin isoforms function preferentially in particular hair cell types or in discrete phases during stereociliary bundle formation (Sekerková et al., 2006;

Sekerková et al., 2006).

Near the time that people identified espin in hair cell stereocilia, the espin gene in mice

(Espn) was mapped at through in situ hybridization (Zheng et al., 2000).

This region in chromosome 4 contains an autosomal recessive deafness mutation – the

Jerker mutation – through classical linkage analysis (Sekerková et al., 2006). Six mutations that cause deafness have been identified within ESPN, which is located at chromosome 1p36.3 in the human (Naz et al 2004; Donaudy et al., 2006). Two recessive

29 mutations in ESPN1, 1988delAGAG and 2469del GTCA, associate with prelingual and profound sensorineural hearing loss, delayed independent ambulation, and vestibular areflexia (Naz et al 2004; Donaudy et al., 2006). ESPN mutations also appear in patients who do not have obvious vestibular defects but so suffer autosomal dominant hearing loss (Donaudy et al., 2006).

All espin isoforms include a 116-amino acid C-terminal actin-bundling module (ABM)

(Figure 5A). The ABM is sufficient and necessary for actin bundling (Chen et al., 1999;

Bartles et al., 1998) as well as the elongation of microvillar PAB (Loomis et al., 2003).

All of the espin isoforms also have a Wiskott-Aldrich syndrome protein homology 2

(WH2) domain (Figure 5A). This domain can bind to actin monomers and is essential in actin-bundle formation that is mediated by espin (Sekerková et al., 2004; Loomis et al.,

2003; Loomis et al., 2006). Espin isoforms can have a totally different N-terminus, which can include 8 -like repeats, an additional F-actin-binding site, or other various domains (Sekerková et al., 2004, Sekerková et al., 2003). This difference in the N- terminus is understandable – although encoded by the same gene, the transcription start- sites for each isoform are different (Sekerková et al., 2004). Also worth noting, these upstream motifs or binding sites may have the ability to associate to the ligands that play roles in membrane-cytoskeleton interaction, actin-cytoskeleton regulation, or signal transduction. Although in most of the cases, these ligands are still unknown (Sekerková et al., 2004, Bartles et al., 1996; Chen et al., 1999; Bartles et al., 1998).

30 The rich accumulation of espin in hair cell stereocilia and the microvilli of other sensory cells raise the question about the function of espin in these PAB-containing structures. In vitro and in transfected cell models, espin shows different biological characteristics compared to other actin-bundling proteins. The root of these characteristics comes from espin ABM and its vigorous F-actin-binding and -bundling activity (Sekerková et al.,

2006). In a co-sedimentation binding assay/actin-binding assay, espin 2B bound to the rabbit skeleton muscle actin filaments with a Kd of ~70 nM (Chen et al., 1999). This affinity is more than twice as much as other actin-bundling proteins. Espin has even higher affinity with actin filaments from non-muscle sources (Bartles et al., 1998). It can efficiently join the actin filaments to form parallel bundles. In the co-sedimentation bundling assay/actin-bundling assay, purified recombinant espin 2B can transform all single actin filaments to sedimentable bundles even when their molar ratios are as low as one espin protein to 20-50 actin monomers (Chen et al., 1999). This effective cross- linking is further confirmed by negative staining electron microscopy (Chen et al., 1999;

Bartles et al., 1998).

Espin’s F-actin-binding and -bundling abilities are not inhibited by Ca2+ (Sekerková et al., 2004; Chen et al., 1999; Bartles et al., 1998). This is different than some of the other actin-bundling proteins in stereocilia or microvilli of vertebrates, such as fimbrin and villin, as Ca2+ inhibits their activity (Bartles et al., 2000; Athman et al., 2002). Espin can support Ca2+-stable cross-links within the PAB. This is essential for espin-containing sensory cells because the cells have transient increases of local Ca2+ concentration to accomplish transduction. This is one possible explanation of why espins are so

31 concentrated in hair cell stereocilia and microvilli of the sensory cells (Sekerková et al.,

2006).

Using an analytical ultracentrifuge, Chen and Bartles found that recombinant espin exists as a monomer in solution (Chen et al., 1999; Bartles et al., 1998), which indicates that every espin has at least two actin-binding sites in order to accomplish the actin-bundling activity. Based on mutagenesis studies of espins, all espin isoforms have actin-bundling activity in their C-terminal 116-amino acid ABM (Bartles et al., 1998). Deletion from either end of the ABM can erase actin-bundling activity. So, one F-actin binding site is probably located at either end of the ABM (Bartles et al., 1998). These regions show a high level of amino acid sequence conservation.

Espin 1 and 2 isoforms have an additional F-actin-binding site (Chen et al., 1999). This additional binding site was found when espins still showed binding ability to actin filaments in vitro after peptide fragments were subtracted upstream of ABM. The Kd for this binding site is about ~1 µM, which is similar to other actin-bundling proteins (Chen et al., 1999). Using mutagenesis, the additional F-actin-binding site was found to be located on a 23-amino acid strech, which is immediately C-terminal to the N-terminal proline-rich peptide of espin 1 and 2 (Chen et al., 1999). It’s still unknown how this additional F-actin-binding site contributes to the property of F-actin binding and bundling in these isoforms (Chen et al., 1999).

32 Aside from the ABM, all espins have a highly conserved WH2 domain (Sekerková et al.,

2004; Loomis et al., 2003; Loomis et al., 2006) (Figure. 5A). The first 30 amino acids, including a 17 amino acid region, which contains an actin monomer-binding region and core, are coded by a WH2 domain-containing exon (Paunola et al., 2002). This domain is identical across vertebrates (Sekerková et al., 2006). The WH2 domain can bind to the

ATP-actin monomer both in vitro and in vivo, and exists in other actin cytoskeleton proteins (Loomis et al., 2003; Paunola et al., 2002). The 17 amino acids of the WH2 core are essential for this interaction (Sekerková et al., 2004; Loomis et al., 2003; Loomis et al., 2006). This binding between the WH2 domain and ATP-actin monomers is also observed in vivo, and the bound actin monomer can be transiently exchanged. Although this process is mediated by an unknown mechanism, at least espin can increase the local polymerizable actin concentration via binding actin monomers with the WH2 domain

(Sekerková et al., 2006).

Espin 2 and 4 isoforms can bind to PIP2-containing vesicles under physiological pH, ionic strength, and temperature (Sekerková et al., 2004) (Figure. 5A). All of the espins, except espin 4, have a proline-rich peptide (Figure. 5A). Espin 1 and 2 have two of these regions and espin 3 has one, which acts as a common mediator in the protein-protein interactions in signaling cascades and multi-protein scaffolds (Li et al., 2005).

Previous studies of actin and actin-binding proteins, such as espin and fascin, expanded the understanding of these proteins, including their structures, biochemical characteristics, and their functions. However, there are still numerous questions awaiting

33 answers. For example, why do hair cells express so many different kinds of actin-binding proteins that all have “similar” actin-bundling activities in physiological situations? Do their functions overlap with each other, and the cell keeps these functional copies as redundancies to prevent lethal mutations from occurring, or does each of these binding proteins have their own unique characteristics? How do they coordinate with each other to form the stable stereocilium and to maintain this architecture in the long term? What kinds of mechanisms regulate bundling-protein distribution in stereocilia? All of these questions are still unclear and the functions of structural proteins in stereocilia still need to come into focus.

In vitro experiments are effective in understanding the functions of proteins. These experimental methods generally isolate materials from their normal biological environment in order to enable a more detailed, accurate, and practical analysis. In my study, using the traditional track of the previous studies used to understand the function of espin, an actin-binding and -bundling assay was applied to illustrate the properties of fascin 2b. Negative staining and TEM was used to observe the detail of actin bundles formed by fascin 2b or espin 2b. These two actin-binding proteins, both of which are expressed in zebrafish hair cell stereocilia were combined with actin to observe actin- bundle morphology. Cryo-electron tomography was also used to reconstruct the 3- dimentianl structure of the actin bundle formed by fascin 2b, espin 2b or both. By this process, a better understanding of how fascin 2b and espin 2b coordinate with actin to form the actin core of the stereocilium.

34 Materials and Method

1. Materials:

1.1 Chemical reagents

1.1.1 Cell culture

LB Broth (Fisher Scientific Inc, Hampton, NH), Carbenicillin (Fisher Scientific Inc,

Hampton, NH), and isopropyl β-D-1-thiogalactopyranoside (IPTG) (Fisher Scientific Inc,

Hampton, NH) were used.

1.1.2 Protein purification

1.1.2.1 Maltose-Binding Protein (MBP)-fascin 2b

Protease inhibitor (Sigma-Aldrich, St. Louis, MO), dimethyl sulfoxide (DMSO) (Sigma-

Aldrich, St. Louis, MO), Tris base (Fisher Scientific Inc, Hampton, NH), 70% isopropanol (Fisher Scientific Inc, Hampton, NH), β-Mercaptoethanol (Sigma-Aldrich,

St. Louis, MO), NaCl (Fisher Scientific Inc, Hampton, NH), edetic acid (EDTA) (Fisher

Scientific Inc, Hampton, NH), D-maltose (Fisher Scientific Inc, Hampton, NH), and amylose resin (Bio-Rad Laboratories, Inc. Hercules, CA) were used.

1.1.2.2 Hexa polyhistidine-tagged (His-tag) espin 2b

Protease inhibitor (Sigma-Aldrich, St. Louis, MO), dimethyl sulfoxide (DMSO) (Sigma-

Aldrich, St. Louis, MO), Tris base (Fisher Scientific Inc, Hampton, NH), 70% isopropanol (Fisher Scientific Inc, Hampton, NH), β-Mercaptoethanol (Sigma-Aldrich,

St. Louis, MO), NaCl (Fisher Scientific Inc, Hampton, NH), KCl (Fisher Scientific Inc,

35 Hampton, NH), imidazole (Sigma-Aldrich, St. Louis, MO), glycerol (Fisher Scientific

Inc, Hampton, NH), and Ni-NTA resin (Qiagen, Venlo, Limburg) were used

1.1.3 Actin-binding and –bundling assay

CaCl2 (Fisher Scientific Inc, Hampton, NH), MgCl2 (Fisher Scientific Inc, Hampton,

NH), KCl (Fisher Scientific Inc, Hampton, NH), rabbit muscle actin (Cytoskeleton, Inc.

Nenver, CO), dithiothreitol (DTT) (Fisher Scientific Inc, Hampton, NH), imidazole-HCl

(Sigma-Aldrich, St. Louis, MO), and adenosine triphosphate (ATP) were used.

1.1.4 SDS-PAGE

Tris base (Fisher Scientific Inc, Hampton, NH), sodium dodecyl sulfate (SDS) (Fisher

Scientific Inc, Hampton, NH), glycine (Fisher Scientific Inc, Hampton, NH), N,N,N’,N’- tetramethylethylenediamine (TEMED) (Fisher Scientific Inc, Hampton, NH), ammonium persulfate (APS) (Fisher Scientific Inc, Hampton, NH), 40% degassed acrylamide/N,N’- methylenebisacrylamide (bisacrylamide) (Fisher Scientific Inc, Hampton, NH), laemmli sample buffer (Bio-Rad Laboratories, Inc., Hercules, CA), and β-Mercaptoethanol

(Sigma-Aldrich, St. Louis, MO) were used.

1.1.5 Ruby protein gel staining

Methanol (Fisher Scientific Inc, Hampton, NH), acetic acid (Fisher Scientific Inc,

Hampton, NH), and SYPRO Ruby Protein Gel Stain (Invitrogen, Carlsbad, CA) were used.

36 1.1.6 Negative stain TEM

Uranyl acetate (UrAc) (Electron Microscopy Sciences, Hatfield, PA) was used.

1.2 Buffers

1.2.1 General buffers

1 M NaOH: dissolve 40 g NaOH pellet in 1 L deionized water (DDI H2O)

0.1 M NaOH: add 99 ml deionized water into 1 ml 1 M NaOH

10% SDS: dissolve 10 g SDS in 90 ml deionized water with gentle stirring and bring to

100 ml with deionized water

0.1% SDS: add 99 ml deionized water into 1 ml 10% SDS

0.5 M Tris pH-6.8: dissolve 6 g Tris base in 60 ml deionized water and adjust to pH-6.8 with 6 N HCl bring total volume to 100 ml with deionized water

1.5 M Tris pH-8.8: dissolve 27.23 g Tris base in 80 ml deionized water and adjust to pH-

8.8 with 6 N HCl, bring total volume to 150 ml with deionized water

10% (w/v) APS: 100 mg ammonium persulfate was dissolve in 1 ml of deionized water

1.2.2 Cell culture

LB media: dissolve 25 g LB powder in 1 L DDI H2O, autoclaved

Carbenicillin: dissolve 225 mg carbenicillin in 13 ml deionized water, sterilized by passing through 0.22 µm filter

IPTG: dissolve 1.55 g carbenicillin in 13 ml deionized water, sterilized by passing through 0.22 µm filter

1.2.3 Protein purification

37 1.2.3.1 MBP-fascin 2b

Protease inhibitor: add 1 ml of dimethyl sulfoxide (DMSO) to the 5 ml size bottle, vortex for one minute, then add 4 ml of deionized water

Fascin 2b lysis buffer: 200 mM NaCl, 20 mM Tris pH-7.4, 1 mM EDTA pH-8.0, 10 mM β-mercaptoethanol

Fascin 2b column buffer: 200 mM NaCl, 20 mM Tris pH-7.4, 1 mM EDTA pH-8.0, 10 mM β-Mercaptoethanol

Fascin 2b elution buffer: 300 mM NaCl, 20 mM Tris pH-7.4, 1 mM EDTA pH-8.0, 200 mM D-maltose, 10 mM β-mercaptoethanol

1.2.3.2 His-tag espin 2b

Protease inhibitor: add 1 ml of dimethyl sulfoxide (DMSO) to the 5 ml size bottle, vortex for one minute, then add 4 ml of deionized water

Espin 2b lysis buffer: 50 mM Tris-HCl, 10 mM β-mercaptoethanol, pH-8.5

Espin 2b column buffer: 20 mM Tris-HCl, 5% (v/v) glycerol, 100 mM KCl, 300 mM

NaCl, 20 mM imidazole, 10 mM β-mercaptoethanol, pH-8.5

Espin 2b elution buffer: 20 mM Tris-HCl, 100 mM KCl, 300 mM NaCl, 250 mM imidazole, 10 mM β-Mercaptoethanol, pH-8.5

1.2.4 Actin-binding assay and actin-bundling assay

1.2.4.1 MBP-fascin 2b alone

G buffer: 5 mM Tris [pH-8.0], 0.2 mM CaCl2, 0.5 mM DTT, 0.2 mM ATP

F buffer: 500 mM KCl, 20 mM MgCl2, 10 mM ATP

38

1.2.4.2 His-tag espin 2b alone

G buffer: 5 mM Tris [pH-8.0], 0.2 mM CaCl2, 0.5 mM DTT, 0.2 mM ATP

F buffer: 0.1 M KCl, 2 mM MgCl2, 1 mM ATP, 1 mM NaN3, 10 mM imidazole-HCl pH-7.4

1.2.4.3 MBP-fascin 2b and His-tag espin 2b

G buffer: 5 mM Tris [pH-8.0], 0.2 mM CaCl2, 0.5 mM DTT, 0.2 mM ATP

F buffer: 0.1 M KCl, 2 mM MgCl2, 1 mM ATP, 1 mM NaN3, 10 mM imidazole-HCl pH-7.4, 3.6 mM DTT

1.2.5 SDS-PAGE

Stacking gel (10 ml): 9.8 ml DDI H2O, 2.5 ml 40% degassed acrylamide/bis, 2.5 ml 0.5

M Tris-HCl pH-6.8, 0.1 ml 10% w/v SDS, 50 µl 10% APS, 10 µl TEMED

Running gel (10 ml): 9.8 ml DDI H2O, 2.5 ml 40% degassed acrylamide/bis, 2.5 ml 1.5

M Tris-HCl pH-8.8, 0.1 ml 10% w/v SDS, 50 µl 10% APS, 5 µl TEMED

Running buffer (10 ×): 30.3 g Tris base, 144.0 g glycine, 10.0 g SDS. Dissolve and bring total volume up to 1,000 ml with deionized water.

Loading dye: mix 950 µl laemmli sample with 50 µl β-mercaptoethanol

1.2.6 Ruby protein gel staining

Fix solution: 50% methanol and 7% acetic acid

Washing solution: 10% methanol and 7% acetic acid

39 2. Methods:

2.1 Cell culture

After transformation of E. coli strain BL21(DE3) with a recombinant vector, the bacteria were grown overnight in 100 ml LB broth (with 100 µg/ml sterilized carbenicillin) at 37

°C at 225 rpm. This overnight culture was used to inoculate 12 liters of fresh LB medium and cells were grown at 37 °C with shaking at 225 rpm. When the culture OD600 reached

0.4, which is the mid-long phage, protein expression was induced by adding sterilized

IPTG (fresh, prepared less than an hour before use) to a final concentration of 0.5 mM.

The culture temperature was reduced to 25 °C following induction. After an additional

4.5 hours of cultivation, cells were harvested by centrifugation at 4000 rpm for 10 min at

4 °C. The bacterial pellet (~45 g) was collected in petri dishes (~15 g/dish) and put on dry ice for freezing. The cells were stored at -80 °C for future use.

2.2 Protein purification

2.2.1 MBP-fascin 2b

E. coli was resuspended in fascin 2b lysis buffer, stirring at 4 °C until the bacterial was fully mixed. Microfluidizer (M-110Y Microfluidizer Processor, Microfluidics,

Westwood, MA) was used to lyse the bacterial. The cell lysate went through the microfluidizer at 80 psi at 4 °C for complete lysis. The cell lysate was then centrifuged at

3,000 g for 20 min. The supernatant was combined with amylose resin suspension and rotated at 4 °C for 3 hours. The resin was pelleted by centrifugation at 3,000 g for 5 min and the supernatant, which contains unbound protein, was discarded. The resin was then washed by fascin 2b column buffer 12 times at 4 °C. Fascin 2b protein bound to the

40 amylose resin was eluted by fascin 2b elution buffer at 4 °C overnight. The elution buffer containing MBP-fascin 2b protein was collected and spun down at 100,000 g for 1 hour to discard protein aggregations. MBP-fascin 2b elution buffer was then concentrated using a 50K concentrator (Amicon Ultra-15 Centrifugal Filter Units, EMD Millipore,

Billerica, MA) and spinning at 3,000 g at 4 °C. The fusion protein was further purified by running sample through a Superdex 200 column (GE Healthcare Life Sciences,

Pittsburgh, PA) equilibrated with fascin 2b column buffer.

2.2.2 His-tag espin 2b

E. coli was resuspended in espin 2b lysis buffer, while stirring at 4 °C until the bacteria was fully mixed. A microfluidizer (M-110Y Microfluidizer Processor, Microfluidics,

Westwood, MA) was used to lyse the bacteria. The cell lysate went through the microfluidizer at 80 psi at 4 °C for a complete lysis. The cell lysate was then centrifuged at 3,000 g for 20 min. The supernatant was combined with Ni-NTA resin suspension and rotated at 4 °C overnight. The resin was pelleted by centrifugation at 3,000 g for 5 min and the supernatant, which contains unbound proteins, was discarded. The resin was then washed by espin 2b column buffer 12 times at 4 °C. Espin 2b protein bound to the Ni-

NTA resin was eluted by espin 2b elution buffer at 4 °C overnight. The elution buffer containing His-tag espin 2b protein was collected and spun down at 100,000 g for 1 hour to discard protein aggregations. His-tag espin 2b elution buffer was then concentrated a using 10K concentrator (Amicon Ultra-15 Centrifugal Filter Units, EMD Millipore,

Billerica, MA) and spun at 3,000 g at 4 °C. The fusion protein was further purified by a

41 Superdex 200 column (GE Healthcare Life Sciences, Pittsburgh, PA) equilibrated with espin 2b column buffer.

2.3 Actin protein preparation

Rabbit skeletal muscle actin came in powder, 1 mg/tube. 100 µl cold DDI H2O was added to stock powder to reconstitute the actin protein. The 100 µl actin stock solution was aliquoted, snap frozen in dry ice ethanol, and stocked in -80 °C freezer. The actin stock solution was dissolved on ice, then diluted in G buffer and incubated on ice for 1 hour at

<1 mg/ml to depolymerize actin oligomers that formed during storage. Actin protein was centrifuged at 14,000 g for 15 min at 4 °C, and the supernatant was used in experiments.

2.4 Negative stain transmission electron microscopy

2.4.1 Sample preparation

Actin alone, mixed with MBP-fascin 2b, mixed with His-tag espin 2b and/or mixed with both were incubated at room temperature in the presence of F buffer for 1 hour. 400 mesh holey carbon-coated copper grids were glow discharged at 25 mA for 1 min. A 3 µl sample was placed on the grid at room temperature for 30 s and extra liquid was blotted with filter paper. Then, 10 µl DDI H2O was used to wash the grid twice for 10 s each, and extra water was blotted with filter paper. The grid was further stained, twice, with 10 µl

2% uranyl acetate for 20 s and 1 min, respectively, and then extra staining solution was blotted with filter paper. The grids were dried for about 5 min at room temperature with a humidity of ~10% until completely dry before placed into a grid box.

42 2.4.2 Microscope alignment and data collection

Two-dimensional automated data collection for negative staining was performed on a FEI

Tecnai Spirit BioTwin transmission electron microscope (Eindhoven, The Netherlands) operated at 100 KeV with LaB6 filament, imaging done with Gatan US4000 UHS charge- coupled device camera (4k × 4k) (Warrensdale, PA). Images were recorded at 49,000 × or 60,000 × magnification at ~1.5 µm under focus.

2.5 Cryo-electron tomography

2.5.1 Sample preparation

Actin mixed with fascin 2b, espin 2b, or both were incubated at room temperature in F- buffer for 1 hour. Quantifoil R3.5/1 200 mesh copper grids were glow discharged at 25 mA for 1 min. 3 µl sample was placed on the grid. 1 µl colloidal gold was added to serve as fiduciary markers for electron tomography. Extra liquid was blotted once with

Whatman filter paper. Grids were then rapidly frozen by manual plunging into liquid ethane. All frozen grids were stored in liquid nitrogen.

2.5.2 Microscope alignment, calibration, and data collection

Digital micrographs of two-dimensional automated data collections of the actin-actin- binding protein complexes were acquired under low-dose conditions at liquid nitrogen temperature on a FEI Tecnai 200kV Field Emission Gun transmission electron microscope (Eindhoven, The Netherlands), imaging with TVIPS CMOS (Complementary

Metal Oxide Semiconductor) camera (4k × 4k) (Gauting, Germany). TF20 electron

43 microscope was operated at 120 KeV. TF20 microscope alignment and calibration were performed according to standard procedures. For one tilt series, a total of 60 images were collected at an absolute magnification of 29,000 ×, corresponding to a pixel size of

0.3665 Å on the molecular scale. The grid was tilted from -68° to +58° and an image was collected every 2°. The defocus values of the micrographs ranged from 3.5 to 4 µm.

2.6 Actin-binding assay and actin-bundling assay

Depolymerized actin monomers were mixed with F-buffer, incubated at room temperature for 1 hour, and spun down at 11,000 g for 20 min to get rid of tangled F- actin. Supernatant was collected and mixed with and without cross-linkers (1.67 µM

MBP-fascin 2b or 1 µM His-tag espin 2b). F buffer was added to reach a final salt concentration. All samples were incubated at room temperature for 1 hour. For the actin- binding assay, the samples were ultracentrifuged at 100,000 g for 40 min to separate F- actin bound with or without cross-linkers from actin monomer and excess cross-linkers.

Supernatants and pellets were collected separately and resuspended in loading dye for

SDS-PAGE. For the actin-bundling assay, the samples were centrifuged at 10,000 g for

15 min to separate actin bundles formed by cross-linkers and single strands of F-actin and excess cross-linkers. Supernatants and pellets were collected separately and resuspended in loading dye for SDS-PAGE. All samples were boiled for 5 min and vortexed for 30 s before SDS-PAGE.

44 2.7 SDS-PAGE and Ruby protein gel staining

15 µl molecular marker and 20 µl samples were subjected to SDS-PAGE. The gel was run at 135 V for ~45 min to 1 hour. Each gel was washed twice by Ruby staining fixing solution for 30 min, followed by staining by Ruby protein gel stain overnight, and then washed by Ruby staining washing solution for 30 min. Gel images were collected on a

Typhoon 9410 Variable Mode Imager (GE Healthcare Life Sciences, Pittsburgh, PA).

2.8 Data processing and statistical analysis

Negative staining data was converted from .dm3 format to .tiff format using Digital

Micrograph (DM) software loaded on a T12 microscope. All images were opened and analyzed using ImageJ. Tomography images were processed by using IMOD (the

Boulder Laboratory for 3-D EM of Cells, Colorado), which is a set of image processing, modeling, and display programs used for tomographic reconstruction of EM serial sections and optical sections. Molecular graphics and analyses were performed with the

UCSF Chimera package. Chimera is developed by the Resource for Biocomputing,

Visualization, and Informatics at the University of California, San Francisco (supported by NIGMS P41-GM103311). The data was represented in mean ± SEM.

45 Result

Part one –Fascin 2b

Maltose-Binding Protein (MBP)-fascin 2b protein overexpression and confirmation of binding activity

Maltose-Binding Protein (MBP)-fascin 2b was overexpressed in BL21 E.coli. Following the previous procedures (Chou et al., 2011), the protein was purified from the cell lysate by both amylose affinity chromatography and size exclusion chromatography (SEC).

Amylose affinity chromatography uses amylose beads, which have a high binding affinity for the 42 kDa maltose-binding protein (MBP) to separate proteins with an MBP tag from the cell lysate. Purifying MBP-fascin 2b by amylose affinity chromatography was not sufficient as the sample preparation gave a mixture of MBP-fascin 2b protein, aggregated proteins and debris (Figure 6B, left lane). Therefore, SEC was needed to further purify the protein. SEC is a chromatographic method in which molecules in solution are separated by their sizes or their molecular weights. SEC was able to separate the aggregated protein from the MBP-fascin 2b (Figure 6B, right lane). In addition, SEC can be used as a way to identify purified protein. The SDS-PAGE with Ruby staining confirmed MBP-fascin 2b purity and molecular weight.

The purified MBP-fascin 2b protein was confirmed by both SEC and SDS-PAGE

(Figure 6). The SEC chromatogram revealed a void peak and an approximate 97 kDa peak representing our fascin 2b protein (55 kDa) fused to the MBP tag (Figure 6A). The

46 approximate 200 kDa peak was speculated to be aggregated MBP-fascin 2b from the purification process. The result from the SDS-PAGE confirmed the purity of MBP-fascin

2b protein after SEC (Figure 6B). Only the fractions from the MBP-fascin 2b peak were used for further analysis.

Figure 6. Purification of MBP-fascin 2b. (A) The chromatogram from gel filtration shows the existence of MBP-fascin 2b protein. (B) 10% SDS-PAGE with Ruby protein gel staining shows MBP-fascin 2b band at 97 kDa. Fascin 2b fusion protein is a 55 kDa protein, fascin 2b, with a 42 kDa MBP tag.

Confirmation of MBP-Fascin 2b’s actin-binding and –bundling activity

In this study, the MBP tag wasn’t removed as it can be used to locate fascin 2b in the 3- dimensional reconstruction of bundled actin. Since we used MBP-fascin 2b protein for our studies, it was necessary to confirm that the actin-binding and –bundling ability of

MBP-fascin 2b is comparable to fascin 2b without the MBP tag. Thus, actin-binding and actin-bundling assays were used to confirm MBP-fascin 2b’s actin-binding and –bundling activity in vitro.

47 Fascin 2b has two actin-binding sites. The actin-binding ability refers to the binding of actin to either binding sites whereas the actin-bundling ability requires actin binding to both of the binding sites. Actin-binding and –bundling assays have long been used to show actin-binding and –bundling ability by cross-linkers.

From these assays, the F-buffer alone provided proper salt concentration for actin polymerization but cannot stimulate the formation of actin bundles (Figure 7, lanes 1-4).

When MBP-fascin 2b was incubated with actin monomer at molar ratio of 1:3 MBP- fascin 2b to actin, the MBP-fascin 2b can bind with F-actin and form actin bundles

(Figure 7, lanes 5-8). This result is in agreement with previous studies (Lin-Jones and

Burnside, 2007) of fascin 2b without the MBP tag and that fascin 2b can bind and bundle

F-actin in vitro. Therefore, our data suggested that the MBP tag had no effect on fascin

2b actin-binding and -bundling activity in vitro. However, the morphology of the actin bundle formed by MBP-Fascin 2b can be changed by MBP tag. Thus, other techniques such as transmission electron microscopy (TEM), can further confirm the MBP-fascin

2b-actin bundle morphology.

48

Figure 7. Confirmation of the reconstructed MBP-fascin 2b actin-binding and –bundling capacity using actin-binding assay and actin-bundling assay. Rabbit muscle actin (5.0 µM) with or without 1.67 µM of MBP-fascin 2b were used. After ultracentrifugation (U) or centrifugation (C), actin and MBP-fascin 2b in the supernatant (S) and in the pellet (P) were separated by SDS-PAGE with Ruby protein gel staining to show MBP-fascin 2b actin-binding ability and actin-bundling ability (lane 5-8) with control groups of actin alone (lane 1-4). Standard protein ladder is shown on the left (L).

Confirmation of the morphology of the MBP-fascin 2b-actin bundle and examination of the actin-bundling ability of MBP-fascin 2b using negative staining transmission electron microscopy

Fascin 2 has been known to bundle F-actin into tightly packed, parallel bundles in vitro.

Our purified fascin 2b contains an MBP tag that may disrupt the parallel actin bundles.

Since MBP tag may affect the morphology of the fascin 2b-actin bundle, the MBP-fascin

2b-actin complex was examined under negative stain TEM for fine morphology observation. F-actins were tightly packed to form actin bundles when incubated with

MBP-fascin 2b protein (Figure 8).

49

Figure 8. MBP-fascin 2b-actin bundle morphology. Negative staining TEM shows the morphology of the actin bundle formed by 5.0 µM actin with 15.0 µM MBP-fascin 2b. Each arrow points to a putative bound MBP-fascin 2b protein on the actin bundle. Scale bar: 100 nm

According to the previous study, fascin 2 can saturate actin with molecular ratio of 1:3 fascin 2b to actin (Saishin et al., 2000; Kureishy et al., 2002). 5 µM of actin was mixed with 0 µM, 0.5 µM, 1 µM, 2 µM, 5 µM, 6 µM, 10 µM, and 15 µM of purified MBP-fascin

2b. In the control group, without any MBP-fascin 2b present, F-actin existed in the solution as flexible single filaments, or more rarely tangled together (Figure. 9A and B).

A single actin filament has a diameter of 7 nm. This number was used to convert the average width of the actin bundle in nanometers to the number of actin filaments per bundle.

F-actin started to form actin bundles when incubated with 0.5 µM MBP-fascin 2b, although single filaments was still observed under the TEM and were a large portion of the whole population (Figure. 9C and D). F-actins can form thick bundles stably at higher concentrations of MBP-fascin 2b, with almost no single actin filaments present at

50 5 µM of MBP-fascin 2b (Figure. 9E-H). An MBP-fascin 2b titration was done to determine the average thickness of actin bundles formed by different MBP-fascin 2b concentrations. From 0 µM to 15 µM, the average thickness of actin bundles increased in a concentration-dependent manner (Figure 9I). The thickness of the actin bundle formed by MBP-fascin 2b approached a plateau when MBP-fascin 2b concentration was greater than 15 µM.

The filaments in the bundle were straight and completely parallel to each other. Previous studies have shown that fascin protein facilities actin bundling into a hexagonal array

(Yang et al., 2012). We found that this spatial arrangement was difficult to view using negative staining. The top surface of the actin bundle formed by MBP-fascin 2b was very flat, probably due to the negative staining compound uranyl acetate. The “belt” formed by fascin 2b was observed at high fascin 2b concentrations to form thick actin bundles.

Although negative staining TEM can provide morphologic information of the MBP- fascin 2b-actin bundle, the negative staining compound may have caused the flattening of the bundle and presented possible staining artifacts. To view the MBP-fascin 2b-actin bundle in the native spatial arrangement, cryo-electron tomography was used as an alternative method.

51

Figure 9. MBP-fascin 2b actin-bundling capacity. Increasing concentrations of MBP- fascin 2b were incubated with 5.0 µM F-actin and images were gathered using negative staining TEM. All bundles in the same group were counted and the thicknesses were measured. A and B, 5.0 µM actin protein alone. Based on 259 filaments from 100 images, the average thickness of this group is 8.48 ± 0.08 nm, which equals to 1 ± 0.01 strand per bundle. (A) Single actin filament in the absence of MBP-fascin 2b. (B) Distribution of the bundle thicknesses of the actin alone group. C and D, 5.0 µM actin with 0.5 µM MBP- fascin 2b. Based on 679 filaments from 132 images, the average thickness of this group is 18.39 ± 0.26 nm, which equals to 3 ± 0.04 strands per bundle. (C) Actin bundle formed

52 by 0.5 µM MBP-fascin 2b with 3-strand thickness. (D) Distribution of the thickness of the actin bundle formed by 0.5 µM MPB-fascin 2b. E and F, 5.0 µM actin with 5.0 µM MBP-fascin 2b. Based on 565 actin bundles from 77 images, the average thickness is 58.56 ± 0.72 nm. This equals to 8 ± 0.10 strands per bundle. (E) Actin bundle formed by 5.0 µM MBP-fascin 2b with an 8-strand thickness. (F) Distribution of the thickness of the actin bundle formed by 5.0 µM MPB-fascin 2b. G and H, 5.0 µM actin with 15.0 µM MBP-fascin 2b. Based on 674 actin bundles from 159 images, the average thickness is 76.66 ±0.96 nm, which is equal to 11 ± 0.14 strands per bundle. (G) Actin bundle formed by 15.0 µM MBP-fascin 2b with 11-strand thickness. (H) Distribution of the thickness of the actin bundle formed by 15.0 µM MPB-fascin 2b. Scale bars: 100 nm. (I) Plot of the mean of the thicknesses of the actin bundles formed by different concentration of MBP- fascin 2b. Each point is the mean ± SEM.

Cryo-electron tomography of the fascin 2b-actin bundle

Cryo-electron tomography was used to analyze the three-dimensional bundles formed by

MBP-fascin 2b and F-actin in the native spatial arrangement in vitro (Figure 10). To date, I have imaged partially bundled F-actin strands that can be observed covered by ice

(Figure 10).

53

Figure 10. Raw data taken by cryo-EM shows some single actin filaments that are only partially bundled by MBP-fascin 2b. Scale bar: 200 nm

54 Part two – Espin 2b

Purification of hexa polyhistidine-tagged espin 2b protein

Hexa polyhistidine-tagged (His-tag) espin 2b was overexpressed in BL21 E. coli cells.

According to a previous purification protocol (Bartles et al., 1998), the His-tag espin 2b is isolated and purified from cell lysate by Ni-NTA affinity chromatography then SEC.

Affinity chromatography followed by SEC can help separate the His-tag espin 2b from aggregated protein and debris during the purification (Figure. 11B left lane and right lane). The chromatogram can also help to identify His-tag espin 2b. SDS-PAGE confirmed the purity and molecular weight of the His-tag espin 2b as well.

The purified His-tag espin 2b was confirmed by both SEC and SDS-PAGE (Figure 11).

The SEC chromatogram showed a void peak and an approximate 30 kDa peak representing the His-tag espin 2b. The 70 kDa peak was speculated to be aggregated His- tag espin 2b. The result from the SDS-PAGE confirmed the purity of His-tag espin 2b after SEC (Figure 11B).

55

Figure 11. Confirmation of the purified His-tag espin 2b. (A) The chromatogram from gel filtration shows the existence of His-tag espin protein. (B) 10% SDS-PAGE with Ruby protein gel staining shows a His-tag espin 2b band at 30 kDa.

Confirmation of His-tag espin 2b actin-binding and –bundling activity

Previous studies showed that espin 2b has actin-binding and –bundling ability in vitro.

Similar to MBP-fascin 2b, the polyhistidine tag was not cleaved from the protein. Thus, it is essential to use the actin-binding assay and the actin-bundling assay to confirm His-tag espin 2b actin-binding and –bundling ability in vitro.

Again, the F-buffer only provided proper salt concentration and enough energy (ATP) for actin monomers to polymerize. There was no ability of this buffer to form actin bundles by itself (Figure 12 lane 1-4). When His-tag espin 2b was incubated together with actin monomers it was able to bind to F-actin and form actin bundles in vitro at a molar ratio of

1:5 His-tag espin 2b to actin (Figure 12 lane 5-8). Thus, polyhistidine tag had no effect on espin 2b actin-binding and -bundling capacity.

56

Figure 12. Confirmation of the reconstructed His-tag espin 2b actin-binding and – bundling ability using actin-binding assay and actin-bundling assay. Rabbit muscle actin (5.0 µM) with or without 1 µM of His-tag espin 2b were used. After ultracentrifugation or centrifugation, actin protein and His-tag espin 2b in the supernatant (S) and in the pellet (P) were separated by SDS-PAGE with Ruby protein gel staining to show His-tag espin 2b actin-binding ability and actin-bundling ability (lane 5-8) with control groups of actin alone (lane 1-4). Standard protein ladder (L).

Observation of the morphology of His-tag espin 2b-actin bundle and examination of the actin-bundling ability of His-tag espin 2b using negative staining transmission electron microscopy

Actin–espin 2b complex was examined under negative-stain electron microscopy for a detailed morphology observation (Figure 13). Using espin 2b as a cross-linker, F-actins formed bundles with several distinguishing characteristics compared to the bundles formed by fascin 2b. Firstly, the espin 2b-actin bundles can reach up to 20 microns in length, which is longer compared to the length of fascin 2b-actin bundles that is around several microns. Secondly, the actin bundles formed by espin 2b are more isolated and less tangled (Figure 13). Fascin 2b-bundles are easier to tangle with each other (Figure

9E), unless fascin 2b’s concentration is very high and the bundles are very thick. A closer

57 examination of the His-tag espin 2b-actin bundles shows two different arrangements. F- actin bundles are more twisted with intersecting filaments at lower concentration of His- tag espin 2b (molecular ratio 1:5 His-tag espin 2b to actin; Figure 13A). However, the arrangement of the actin bundles formed by His-tag espin 2b is parallel to each other in the bundle at relatively high His-tag espin 2b concentration groups (molecular ratio of 2:1

His-tag espin 2b to actin; Figure 13B). Almost no criss-crossing was observed. Similar to the MBP-fascin 2b, the negative staining compound can cause flattening of the bundle.

Therefore, cryo-electron tomography was needed to view the His-tag espin 2b-actin bundle in the native spatial arrangement.

58

Figure 13. His-tag espin 2b-actin bundle morphology. Negative staining TEM shows the two different morphologies of the actin bundle formed by His-tag espin 2b. (A) Actin bundle formed by 5.0 µM actin and 1.0 µM His-tag espin 2b. (B) Actin bundle formed by 5.0 µM actin and 10.0 µM His-tag espin 2b. Scale bar: 100nm

5 µM of actin mixed with 0 µM, 1 µM, 2 µM, 5 µM, 6 µM, and 10 µM of His-tag espin

2b. The result is shown in Figure 14. In the control group, without any cross-linker, F- actin strands are present in the solution as curved single filaments, or in some rare situations, tangled together (Figure. 14 A and B).

1 µM His-tag espin 2b with F-actin can form thick actin bundles (Figure 14C and D).

There is a robust increase of the average bundle diameter that occurs when only 1 µM more of His-tag espin 2b (Figure. 14 E and F) is added. However, with further added

His-tag espin 2b protein an increase in the average bundle thickness did not occur to a significant degree (Figure G and H).

59 Figure 14M shows the average thickness of actin bundles formed by titrating His-tag espin 2b from 0 µM to 10 µM. In general, His-tag espin 2b is a very vigorous actin- bundler. From 1 µM to 10 µM, although the average thickness of actin bundles increases when adding more cross-linker proteins, there is a clear plateau in this plot. The inflexion seems around 2 µM. All groups whose concentrations are higher than this point, no matter how much more espin 2b protein is added, the average thickness of the actin bundles only increases a small amount. The dramatic increase of actin bundle thickness from the 1 µM group to the 2 µM group does not repeat at higher concentrations.

Negative staining transmission electron microscopy can reveal much information about the morphology and thickness of the actin bundles formed by His-tag espin 2b. More structural details of the His-tag espin 2b–actin bundle in native solution were approached using cryo-electron tomography.

60

Figure 14. His-tag espin 2b’s actin-bundling capacity. Different concentrations of His- tag espin 2b were incubated with 5.0 µM F-actin and images were taken of samples using negative staining and TEM. All bundles in the same group were counted and the thicknesses were measured. A and B, 5.0 µM actin alone. Based on 259 filaments from 100 images, the average thickness of this group is 8.48 ± 0.08 nm, which equals to 1 ± 0.01 strand per bundle. (A) Single actin filament in the absence of His-tag espin 2b. (B) Distribution of bundle thicknesses in the actin alone group. C and D, 5.0 µM actin with

61 1.0 µM His-tag espin 2b. Based on 524 actin bundles from 72 images, the average thickness of the actin bundles is 42.45 ± 0.48 nm, which is equal to 6 ± 0.07 strands per bundle. (C) Actin bundle formed by 1.0 µM His-tag espin 2b with a 6-strand thickness. (D) Distribution of bundle thicknesses formed by 1.0 µM His-tag espin 2b. E and F, 5.0 µM actin with 2.0 µM His-tag espin 2b. Based on 482 actin bundles from 100 images, the average thickness of the actin bundles was 84.70 ± 1.66 nm, which equals 12 ± 0.24 strands per bundle. (E) Actin bundle formed by 2.0 µM His-tag espin 2b has a 12-strand thickness. (F) Distribution of the thicknesses of the actin bundles formed by 2.0 µM His- tag espin 2b. G and H, 5.0 µM actin with 10.0 µM His-tag espin 2b. There are 664 actin bundles counted from 99 images. The average thickness of these actin bundles is 97.76 ± 1.47 nm. This equals 14 ± 0.21 strands per bundle. (G) Actin bundle formed by 10.0 µM His-tag espin 2b has a 14-strand thickness. (H) Distribution of the thicknesses of the actin bundles formed by 10.0 µM His-tag espin 2b. Scale bars: 100 nm. I, Plot of mean bundle thicknesses formed by different concentrations of His-tag espin 2b. Each point is a mean ± SEM.

Cryo-electron tomography of the His-tag espin 2b-actin bundle

Cryo-electron tomography was used to analyze the three-dimensional bundles formed by

His-tag espin 2b and F-actin in the native spatial arrangement in vitro (Figure 15). F- actins are packed to form an actin bundle with espin 2b as a cross-linker. As a monomer, in order to accomplish actin bundling, espin 2b has two actin-binding sites to cross-link two actin filaments. The resolution of the reconstruction is about 33.1 Å (Erickson,

2009). Unfortunately, because of the resolution limitations of cryo-electron tomography, it is very challenging to identify a 30 kDa His-tag espin 2b protein in the between of actin bundles, and the detail of the His-tag espin 2b structure could not be reconstructed. A higher resolution structure of espin 2b using cryo-electron microscopy may answer this question.

62

Figure 15. 3-dimensional reconstruction of a His-tag espin 2b-actin bundle using cryo- electron microscopy. (A), Raw data taken by cryo-EM. (B), 3-dimensional reconstruction of the His-tag espin 2b-actin bundle. (C) and (D), zoomed and slightly rotated view of the 3-dimensional reconstruction. Arrows show the decoration, which is the His-tag espin 2b protein, in the between of the actin bundles. The resolution of the tomogram is about 33.1 Å

63 Part three – Fascin 2b and Espin 2b

Comparison of the actin-bundling capacity of fascin 2b and espin 2b

To develop an understanding of how fascin 2b and espin coordinate in stereocilia, I compared actin bundles formed by both of these proteins. By evaluating the average thicknesses of the actin bundles, from TEM images, formed at different concentrations of

MBP-fascin 2b or His-tag espin 2b, Figure 16 shows the comparison of the actin- bundling power of these two proteins. MBP-fascin 2b and His-tag espin 2b have different

F-actin dissociation constants, which impacts bundling. First, the thickness of the actin bundle formed by MBP-fascin 2b can increase in a concentration-dependent manner and reach its plateau when MBP-fascin 2b concentration is greater than 15 µM (data not shown in this plot); however, His-tag espin 2b actin-bundling ability reaches a plateau when His-tag espin 2b concentration is only about 2 µM. Second, His-tag espin 2b is a more vigorous actin-bundling protein. In every group containing His-tag espin 2b, no matter if the His-tag espin 2b bundling ability reaches its plateau or not, the average thickness of the actin bundle formed by His-tag espin 2b is higher than the one formed by

MBP-fascin 2b under the same concentration conditions (Figure 16).

64

Figure 16. Comparison of MBP-fascin 2b and His-tag espin 2b actin-bundling capacities. Plot shows the mean thicknesses of the actin bundles formed by different concentrations of MBP-fascin 2b or His-tag espin 2b. Each point is mean ± SEM.

Observation of the morphology of actin bundles formed by both MBP- fascin 2b and His-tag espin 2b using negative staining transmission electron microscopy

Since stereocilia contain both fascin 2b and espin, I set out to determine if both proteins coordinate in actin bundling. Figure 17 shows the actin bundle formed by both MBP- fascin 2b and His-tag espin 2b. The bundle sample was examined under electron microscope for detailed morphology observation. With both the His-tag espin 2b and

MBP-fascin 2b as cross-linkers (molecular ratio of 1:1:5 MBP-fascin 2b, His-tag espin 2b to actin), F-actins can form bundles. Interestingly, the bundle formed by these two actin- bundling proteins has the characteristics that are the same as the bundles that are formed by each cross-linker separately. In some part of the bundle, F-actins are packed tight and

65 straight. F-actins in this region are parallel to each other in the bundle, which looks similar to the actin bundle formed by MBP-fascin 2b only (red rectangle in Figure 17). In some other parts of the bundle, F-actins are more twisted and packed relatively loose in the bundle, which is similar to the actin bundle formed by low- concentrations of His-tag espin 2b itself (blue rectangle in Figure 17).

However, these observations cannot be the final evidence to confirm the structure of the

MBP-fascin 2b - His-tag espin 2b- actin bundle because of the artificial flattening effect caused by the negative staining compound. The structure in native solution needs to be reconstructed using cryo-electron tomography.

Figure 17. MBP-fascin 2b-His-tag espin 2b-actin bundle morphology. Negative staining transmission electron microscopy shows the morphology of the actin bundle formed by both MBP-fascin 2b and His-tag espin 2b. The actin bundle formed by 5.0 µM actin, 1.0 µM MBP-fascin 2b and 1.0 µM His-tag espin 2b. Red box indicates the region that looks

66 similar to the actin bundle formed by MBP-fascin 2b only; blue box indicates the region that looks similar to the actin bundle formed by only a low-concentration of His-tag espin 2b. Scale bar: 100 nm.

Examination of fascin 2b and espin 2b coordinated actin-bundling using negative staining TEM

The coordination of MBP-fascin 2b and His-tag espin 2b was also studied by measuring the thicknesses of the actin bundles formed by different combination of MBP-fascin 2b and His-tag espin 2b, and then to compare with the data from MBP-fascin 2b alone groups and the one from His-tag espin 2b alone groups. The first combination was designed to construct the situation that both MBP-fascin 2b and His-tag espin 2b were in low concentration when incubated with F-actin. 5 µM of actin was mixed with 1 µM

MBP-fascin 2b and 1 µM His-tag espin 2b. A 2 µM MBP-fascin 2b alone group and a 2

µM His-tag espin 2b alone group were each used as controls for comparison with bundling of these two proteins together. Both the average thickness and the distribution of the thickness of the actin bundle formed by 1 µM MBP-fascin 2b and 1 µM His-tag espin 2b fall between the two control groups (Figure. 18-1 A-D).

The second combination was designed to build a situation where the concentration of

MBP-fascin 2b was lower than the concentration of His-tag espin 2b. 5 µM of actin incubated with 1 µM MBP-fascin 2b and 5 µM His-tag espin 2b. 6 µM MBP-fascin 2b alone and 6 µM His-tag espin 2b alone were used as controls. Both the average bundle thickness and distribution by 1 µM MBP-fascin 2b and 5 µM His-tag espin 2b is between

67 the two controls, but very close to the MBP-fascin 6 µM control group. Compared to the

6 µM His-tag espin 2b group, the mixed group has less thick bundles. (Figure 18-2 A-D).

68

Figure 18-1. Coordination of MBP-fascin 2b and His-tag espin 2b in forming an actin bundle. Different combinations of MBP-fascin 2b and His-tag espin 2b were incubated with 5.0 µM F-actin and images were taken of samples using negative staining transmission electron microscopy. All bundles in the same group were counted and the thicknesses were measured. A and B, 5.0 µM actin with 1.0 µM MBP-fascin 2b and 1.0 µM His-tag espin 2b. 540 actin bundles were counted from 125 images. The average bundle thickness is 62.86 ± 0.92 nm, which is equal to 9 ± 0.13 strands per bundle. (A) Actin bundle formed by 1.0 µM MBP-fascin 2b and 1.0 µM His-tag espin 2b with a 9- strand thickness. (B) Distribution of bundle thicknesses of actin bundles formed by 1.0 µM MBP-fascin 2b and 1.0 µM His-tag espin 2b. C and D, Comparing the average thicknesses (C) and the distribution of the thicknesses (D) of the actin bundles formed by 1.0 µM MBP-fascin 2b and 1.0 µM His-tag espin 2b with the two control groups, 2.0 µM MBP-fascin 2b and 2.0 µM His-tag espin 2b, respectively. Each column in C is mean ± SEM.

69

Figure 18-2. Coordination of MBP-fascin 2b and His-tag espin 2b in forming an actin bundle. A and B, 5.0 µM actin with 1.0 µM MBP-fascin 2b and 5.0 µM His-tag espin 2b. 167 actin bundles were counted from 60 images. The average bundle thickness is 63.18 ± 1.38 nm, which is equal to 9 ± 0.20 strands per bundle. (A) Actin bundle formed by 1.0 µM MBP-fascin 2b and 5.0 µM His-tag espin 2b with a 9-strand-thickness. (B) Distribution of the bundle thickness of the actin bundle formed by 1.0 µM MBP-fascin 2b and 5.0 µM His-tag espin 2b. C and D, Comparing the average thicknesses (C) and the distribution of the thicknesses (D) of the actin bundles formed by 1.0 µM MBP-fascin 2b and 5.0 µM His-tag espin 2b with two control groups, 6.0 µM MBP-fascin 2b and 6.0 µM His-tag espin 2b, respectively. Each column in C is mean ± SEM.

70 The third combination was designed to construct a situation where the concentration of

MBP-fascin 2b was higher than the concentration of His-tag espin 2b. 5 µM of actin is mixed with 5 µM MBP-fascin 2b and 1 µM His-tag espin 2b. 6 µM MBP-fascin 2b alone and 6 µM His-tag espin 2b alone are introduced as controls for comparison of the bundling ability of these two proteins. The average thickness of the bundle formed by 5

µM MBP-fascin 2b and 1 µM His-tag espin 2b is almost the same as the average thickness of the actin bundles formed by 6 µM His-tag espin by itself. This is much thicker than the bundle formed by only MBP-fascin 2b, which suggests there might be some coordination between His-tag espin 2b and MBP-fascin 2b to augment MBP-fascin

2b’s ability, although activated MBP-fascin 2b actin-bundling ability is still not thought to be able to create bundles of average thicknesses larger than 6 µM His-tag espin 2b alone. Comparing the bundle thickness distribution of the mixed group (5 µM MBP- fascin 2b and 1 µM His-tag espin 2b) to the 2 control groups, it is apparent that the distribution of the mixed group is very similar to 6 µM His-tag espin 2b alone, which indicates that 5 µM MBP-fascin 2b acts synergistically with 1 µM His-tag espin 2b to almost replaced the activity of 5 µM His-tag espin 2b. Many of the actin bundles in the mixed group were very thick, which indicates that in some microenvironments in the solution, His-tag espin 2b and MBP-fascin 2b have the ability to build very large (about

39 strands per bundle) actin bundles together (Figure 18-3 A-D).

71

Figure 18-3. Coordination of MBP-fascin 2b and His-tag espin 2b in forming an actin bundle. A and B, 5.0 µM actin with 5.0 µM MBP-fascin 2b and 1.0 µM His-tag espin 2b. 701 actin bundles were counted from 101 images. The average bundle thickness is 93.05 ± 1.69 nm, which is equal to 13 ± 0.24 strands per bundle. (A) Actin bundle formed by 5.0 µM MBP-fascin 2b and 1.0 µM His-tag espin 2b with a 13-strand-thickness. (B) Distribution of bundle thickness formed by 5.0 µM MBP-fascin 2b and 1.0 µM His-tag espin 2b. C and D, Comparing the average thicknesses (C) and the distribution of the thicknesses (D) of the actin bundles formed by 5.0 µM MBP-fascin 2b and 1.0 µM His- tag espin 2b with the two control groups, 6.0 µM MBP-fascin 2b and 6.0 µM His-tag espin 2b, respectively. Each column in C is mean ± SEM.

72 The fourth combination was designed to build the situation that both MBP-fascin 2b and

His-tag espin 2b were in high concentration when incubated with F-actin. 5 µM of actin was incubated with 5 µM MBP-fascin 2b and 5 µM His-tag espin 2b. 10 µM MBP-fascin

2b alone and 10 µM His-tag espin 2b were used as controls for comparison of the bundling capacity of these two proteins. Although the average thickness of the actin bundle in the mixed group is almost the same as the 10 µM His-tag espin 2b group, the distribution of the actin bundles in this group has a very interesting feature that contains the characteristics of the two control groups: the distribution of the percentage of the thinner bundles is very similar to that of 10 µM MBP-fascin 2b; while, the distribution of the percentage of the thicker bundles is almost the same as that of 10 µM His-tag espin 2b

(Figure 18-4 A-D).

73

Figure 18-4 Coordination of MBP-fascin 2b and His-tag espin 2b in forming an actin bundle. A and B, 5.0 µM actin with 5.0 µM MBP-fascin 2b and 5.0 µM His-tag espin 2b. 614 actin bundles were counted from 125 images. The average bundle thickness is 93.95 ± 1.97 nm, which is equal to 13 ± 0.28 strands per bundle. (A) Actin bundle formed by 5.0 µM MBP-fascin 2b and 5.0 µM His-tag espin 2b with a 14-strand- thickness. (B) Distribution of bundle thickness formed by 5.0 µM MBP-fascin 2b and 5.0 µM His-tag espin 2b. C and D, Comparing the average thicknesses (C) and the distribution of the thicknesses (D) of the actin bundles formed by 5.0 µM MBP-fascin 2b and 5.0 µM His- tag espin 2b with the two control groups, 10.0 µM MBP-fascin 2b and 10.0 µM His-tag espin 2b, respectively. Each column in C is mean ± SEM.

74

Figure 19. Actin bundle formed by MBP-fascin 2b and His-tag espin 2b. 5.0 µM actin incubated with 5.0 µM MBP-fascin 2b and 1.0 µM His-tag espin 2b. Occasionally large actin bundles were identified, including the above bundle with up to 39 strands. Scale bar: 100 nm

Among all combinations of proteins at different ratios, compared with the control groups, the average thickness of the actin bundles formed by two cross-linkers together doesn’t exceed the average of the actin bundle thickness formed by the two cross-linkers separately. However, occasionally, actin bundles formed by both fascin 2b and espin 2b with a thickness of up to 39 strands can be found in groups with both bundling proteis when screening whole grids (Figure 19). This thickness is higher than the thickness of the actin bundle formed by either fascin 2b or espin 2b alone. Although, this is very rare; it occurs.

75 Cryo-electron tomography of the fascin 2b-espin 2b-actin bundle

Cryo-electron tomography was used to analyse the 3-dimensional actin bundles formed by MBP-fascin 2b and His-tag espin 2b together in vitro. Figure 20 shows actin bundles formed by both fascin 2b and espin 2b. The raw data directly taken from the microscope and the 3-D reconstruction of the actin bundle are both shown in Figure 20. The signal from the raw data is somehow not strong enough for the UCSF Chimera software to separate it from the background noise. Another tilting series is needed to be collected to repeat the 3-D reconstruction.

Figure 20. 3-dimensional reconstruction of an actin bundle formed by 1 µM MBP-fascin 2b and 1 µM His-tag espin 2b using cryo-electron microscopy. (A), Raw data taken by cryo-EM. (B), 3-dimensional reconstruction of the actin bundle formed by 1 µM MBP- fascin 2b and 1 µM His-tag espin 2b.

76 Discussion

According to reports from the World Health Organization (WHO), more than 5% of the world’s population, about 360 million people, have disabling hearing loss: 328 million of whom are adults and 32 million are children. About one third of the seniors who are over

65 years old are suffering hearing loss. The majority of these people are living in low- and middle-income countries, such as South Asia, and sub-Saharan Africa. Many factors can lead to disabling hearing loss, which is defined as hearing loss greater than 40 dB in the better hearing ear in adults and 30 dB in children. The causes include hereditary factors, which can lead to hearing loss being present at or soon after birth, and acquired causes that can lead to hearing loss at any age. The acquired causes of hearing loss include infectious diseases, such as meningitis, measles and mumps, usage of ototoxic drugs at any age that can impact the inner ear, such as some antibiotics, and exposure to excessive noise and aging. The majority of these causes of hearing loss are often related to effects on the hair cells in the inner ear, including hair cell dysfunction, structural alteration, and functional protein mutation. The function of the hair cell is associated with its unique structure: the precise structure of the hair cell is necessary for its function as a mechanical transducer.

Compared to other cell types from different tissues, hair cells have an exclusive shape and get their name from their staircase shaped hair bundles that are located on the apical surface of the cell and formed by stereocilia. The stereocilia are the actin-based column- shaped protrusions that taper at their bases. Actin is highly concentrated in a stereocilium.

77 Multiple actin-binding proteins are also localized to hair cell stereocilia to assist with the packing of F-actin tightly and paralleled in to form actin bundles in a hexagonal arrangement. This fact indicates the important roles of actin and its binding proteins in forming and maintaining the proper structure of stereocilia, which is essential to the function of hair cells.

As an actin cross-linker that is specifically localized to zebrafish hair cell stereocilia, fascin 2b might have such a significant actin-bundling potential that it can enable F-actin to form uniquely large stereociliary actin bundles. This potential, on the one hand, may come from its own actin-binding and/or actin-bundling ability. On the other hand, the coordination between fascin 2b with other actin-binding proteins in the stereocilia, such as espin 2b may also lead to this potential on building oversize actin bundles. In this thesis, a series of experiments were designed to first prove that fascin 2b and espin 2b actin-binding ability and actin-bundling abilities separately. In addition, the morphologies of the actin bundles formed by the two actin cross-linkers are compared under the transmission electron microscope (TEM) and their three-dimensional structures analyzed using cryo-electron tomography. Moreover, in vitro imaging and structural approaches are also applied to study the possible coordination that happens between these two actin- binding proteins in constructing actin bundles.

The in vitro actin-binding assay and the actin-bundling assay have demonstrated that under a specific ionic environment, maltose-Binding Protein (MBP)-fascin 2b has the ability to bind with F-actin as well as form actin bundles. This result is consistent with

78 previous studies (Chou et al., 2011; Yang et al., 2012) and demonstrates MBP-fascin 2b actin-binding and -bundling capacity. The MBP tag has no effect on interrupting fascin

2b actin-binding and –bundling ability. Examining the morphology of the actin bundle formed by purified MBP-fascin 2b protein using negative stain TEM revealed that F- actins are bundled tightly and in a parallel arrangement. Morphologically, the images show that the F-actin strands remain straight and less twisted in the bundle formed by

MBP-fascin 2b. Using negative stain TEM, the “stripes” formed by bound MBP-fascin

2b on the surface of the actin bundles are easier and more frequently observed under at higher MBP-fascin 2b concentrations. Adding the MBP tag (42 kDa) helped to more easily localize 97 kDa MBP-fascin 2b than 55 kDa fascin 2b alone. The MBP-fascin 2b- actin bundles from the negative stain TEM images overall are very flat. This may be due to the flattening and the artificial effect caused by the negative staining compound.

Different concentrations of MBP-fascin 2b have been incubated with F-actin to examine the actin-bundling ability of MBP-fascin 2b. Systematic negative stain TEM shows the thickness of the MBP-fascin 2b actin bundle increases in a concentration-dependent manner. It has been shown that saturation for bundling of fascin 2 occurs at a stoichiometry of one fascin 2 molecule to three actin monomers (Saishin et al., 2000).

With a stoichiometry of 1:10, MBP-fascin 2b to actin, the majority of bundles have a thickness of 2-4 filaments. Because F-actin is not completely saturated by the MBP- fascin 2b, single actin filaments can still be observed under the TEM (Figure 9D). When increasing the concentration of MBP-fascin 2b, thicker actin bundles can be formed stably. When the MBP-fascin 2b concentration is higher than its saturation concentration, not only almost no single actin filament can be observed, but also F-actins prefer to form

79 thicker bundles with MBP-fascin 2b. The thickness of the actin bundle formed by MBP- fascin 2b reaches its plateau when the MBP-fascin 2b concentration is greater than 15 µM

(Figure 9I). In order to get rid of the artificial effect caused by the negative staining compound and to view the MBP-fascin 2b-actin bundles in a native solution, a series of tilted 2-D images at cryogenic temperatures of the sample was collected to reconstruct the 3-D structure of the MBP-fascin 2b-actin bundle. Unfortunately, so far only some single F-actins that are partially bundled can be observed covered by ice. This might be because the adhesive force of the MBP-fascin 2b binding with F-actin is not strong enough. The stretch applied on the sample when blocking the solution from the grid may cause the MBP-fascin 2b-actin bundle to fall apart. There are several more grids that can be checked in the near future.

As another actin-binding protein that is expressed in the hair cell, espin 2b has been identified earlier than fascin 2b and is well studied (Bartles et al., 1998). In this thesis, espin 2b is introduced into the experimental system to demonstrate a classical actin- binding and –bundling protein with a high affinity (Kd = 150 nM to skeletal muscle F- actin and 50 nM to nonmuscle F-actin) (Bartles et al., 1998) and efficiency to associate with actin protein, and to show its actin-binding and -bundling ability. Experiments for espin 2b were carried out to compare the difference between the espin 2b and fascin 2b actin-binding and –bundling ability. Overexpressed in E. coli and purified following the protocol in a previous paper (Bartle et al., 1998) with some optimization, purified hexa polyhistidine-tagged (His-tag) espin 2b has vigorous actin-binding and –bundling ability in vitro. The actin-binding assay and the actin-bundling assay demonstrate the effective

80 association between F-actin and His-tag espin 2b in a proper ionic environment using biochemical approaches. Observing the morphology of the actin bundles formed by purified His-tag espin 2b under negative stain TEM, it has several significant differences compared to the actin bundles formed by MBP-fascin 2b. First, the actin bundles formed by His-tag espin 2b is lengthier compared to the one formed by MBP-fascin 2b.

Occasionally, the length of the His-tag espin 2b-actin bundle can even reach about 20 to

30 micron. The MBP-fascin 2b-actin bundles, on the other hand, are more frequently found as thick sticks with shorter lengths, with about only several microns. Second, His- tag espin 2b-actin bundles are more isolated and less tangled with each other, while

MBP-fascin 2b-actin bundles are more likely to be tangle together, except the bundles that are very short and/or very thick. These two differences may indicate that there is a high probability that these two actin-binding proteins, fascin 2b and espin 2b, associate or interact with actin monomers or even F-actins differently. Third, espin 2b has significantly more twisting ability when it interacts with F-actin (Figure 13A). Fourth, different from the MBP-fascin 2b-actin bundles, which have a unique morphology, the negative stain TEM images show that His-tag espin 2b has the ability to form two different kinds of actin bundles based on different concentrations (Figure 13). When the espin 2b concentration is relatively low, the F-actins in the His-tag espin 2b- actin bundles are not completely paralleled with each other within the whole bundle.

Sometimes they criss-cross in the bundle. When the espin 2b concentration is relatively high, the F-actin can be packed parallel to each other and much less criss-crossing occurs in the bundle. Fifth, unlike the thickness of the MBP-fascin 2b-actin bundle, which reaches its plateau when MBP-fascin 2b concentration is greater than 15 µM, the

81 thickness of the His-tag espin 2b-actin bundle reaches its plateau only when His-tag espin

2b concentration is 2 µM.

In order to study the way in which they work together in constructing actin bundles, the two actin-binding proteins are then incubated together with F-actin in vitro. As two proteins that both localize in zebrafish hair cell stereocilia, there are several ways that fascin 2b and espin 2b may coordinate. First, they can compete with each other in binding

F-actin. The existence of one will inhibit the activity of the other. Second, they may work together to generate much thicker bundles than either of them alone. Third, they may work independently. The existence of one doesn’t interrupt the activity of the other one.

It has been proven that fascin 1 and human espin can form much thicker bundles together compared to either of these two proteins alone in vitro (Chaessens et al., 2008). Since fascin 1 and fascin 2b are both belong to the fascin family and have 56% sequence identity (Saishin et al., 1997), the characteristic of fascin 1 may provide some clue to understand the characteristic of fascin 2b. Therefore, it is interesting to study how different isoforms of fascin and espin proteins coordinate with each other, and to try to see if we can generate an actin bundle as big as the stereocilium in vitro.

Based on the observation of the morphology of the actin bundles formed by the two actin-binding proteins using regular negative stain TEM, both the characteristics of the

MBP-fascin 2b-actin bundles and His-tag espin 2b-actin bundles can be identified in the actin bundles formed by these two actin-binding proteins together. In part of these actin bundles, the F-actins are packed tighter and more parallel (Figure 17). This is very

82 similar to the morphology of the actin bundle formed by MBP-fascin 2b. In another part of the actin bundle, the F-actins are arranged in a looser, less paralleled, and more twisted way. This morphology more closely resembles the actin bundles formed by low- concentration His-tag espin 2b. This indicates that when both actin-binding proteins exist in the solution to interact with actin protein, it is possible that MBP-fascin 2b and His-tag espin 2b start to build smaller motifs separately to form small bundles first. Later on, these small bundles start to associate with each other to eventually form the bigger actin bundles. One explanation of this is that His-tag espin 2b has higher affinity for F-actins, so although both proteins were added into the solution and start interaction with actin at the same time, His-tag espin 2b may a have greater chance to occupy more actin-binding sites than MBP-fascin 2b at the very beginning. MBP-fascin 2b may have a different binding site on actin compared to the His-tag espin 2b-binding site on actin. But when

His-tag espin 2b occupy the actin-binding sites on one part of the actin and start to form the actin bundle in that region, spatially, it may be hard for MBP-fascin 2b to fit into its own binding site around the same area. In order to bind to the actin filament, MBP-fascin

2b may have to find the other actin-binding sites that are far away from the motif formed by His-tag espin 2b, and start to interact with F-actin to form actin bundle. This may be the reason why a bundle formed by the two proteins together can have both morphologies, which are similar to the bundle morphologies formed by each protein separately.

Comparing the average thickness of the actin bundles formed by two cross-linkers together with the thickness of the bundles formed by MBP-fascin 2b or His-tag espin 2b

83 separately, it seems that there may be weak cooperation between MBP-fascin 2b and His- tag espin 2b: His-tag espin 2b can slightly increase MBP-fascin 2b actin-bundling ability, better than MBP-fascin 2b by itself, but not as strong as His-tag espin 2b alone. In general, the coordination between the two cross-linkers is minimal, since the average thickness of the actin bundles formed by two cross-linkers together didn’t exceed the average of the actin bundle thickness formed by the two cross-linkers alone. However, in the in vitro microenvironment, cooperation may occur because actin bundles formed by both fascin 2b and espin 2b with the thickness of 39 strands can be found (Figure 19).

This thickness is higher than the thickness of the actin bundles formed by either fascin 2b or espin 2b alone, which indicates that there may be some coordination between fascin 2b and espin 2b. However, this coordination hasn’t been observed directly using cryo- electron tomography or another method.

In this thesis, cryo-electron tomography is used to reconstruct the 3-D structure of the actin bundles formed by different two actin-binding proteins separately or by both of them together in order to compare these actin bundles in detail and study their structural differences. Cryo-electron tomography is an electron cryo microscopy technology.

Herein, tomography is used to construct a 3-Dstructure of a sample from a series of tilted

2-D images at cryogenic temperatures (Frank et al., 2006). An obvious advantage for the cryo-electron tomography is that this technique can provide high-resolution 3-

Dreconstruction of biological material in its native state – by imaging hydrated specimens of biological and synthetic origin allowing the study in a state of preservation that is close to native (Nudelman et al., 2010). Chemical fixation, dehydration, and staining in regular

84 negative stain TEM can cause structural disruption, such as flattening of the actin bundle mentioned earlier in this thesis, perturbing the native state. Using cryo-electron tomography, specimens are examined in a frozen state, after use of rapid freezing methods that avoid formation of ice crystals that could damage the structure (Frank et al.,

2006).

In tomography, a 3-D reconstruction is generated by back-projecting a series of images that are recorded in the TEM as the specimen is tilted over a wide range (Frank et al.

2006). For virtually frozen samples, a “low-dose” imaging protocol is used that avoids unnecessary electron irradiation of the sample, which is easily damaged by even a moderate electron dose (Rath et al. 1997; Marko et al. 1999).

Cryo-electron tomography is also a powerful tool to help cryo-electron microscopy to overcome its limitations. In the cryo-TEM, the collected images are actually the 2-D projections of the 3-D object. This leads to an inherent limitation of this technology that the overlapping of multiple features cannot be discerned. Cryo-electron tomography can help to overcome this limitation because in this technique, images are taken at different tilt angles and then reconstructed into the 3-D object in order to reveal the detail information of the morphology and the structure or the 3-Dspatial arrangement of the macromolecules (Nudelman et al., 2010). Cryo-electron microscopy is a priceless tool to connect the spatial structural organization with the function or the activity of the macromolecular complex at the nanometer scale.

85 As a relatively new technology, first presented in the U.S. in 1997 (Goodsell, 2001), cryo-electron tomography still has some disadvantages that need to be improved. The resolution of cryo-electron tomography is relatively low compared to other traditional structural biology tools, such as NMR, and crystallization. The resolution limitation is due to the damage to the specimen caused by electron irradiation (Marko et al., 2006). It can only achieve a 30-100 Å in resolution; therefor, it is almost impossible to determine the detail of the secondary structures. However, just as all other technologies, cryo- electron tomography needs time for improvement. Resolution can be increased if more tilt images can be recorded without increasing the electron dose (McEwen et al.

2002). This can be facilitated by using a better TEM and camera to provide higher contrast. In the near future, cryo-electron tomography will be a great tool in helping biologists to reconstruct 3-D structures of macromolecules with higher resolution.

In conclusion, MBP-fascin 2b is a very active actin-binding and –bundling protein in vitro. The thickness of the actin bundle formed by MBP-fascin 2b increases in a concentration-dependent manner and reaches its plateau when MBP-fascin 2b concentration is higher than 15 µM (Figure 9I). Compared to MBP-fascin 2b, His-tag espin 2b has a more vigorous actin-bundling activity. The same concentration of His-tag espin 2b can build much thicker actin-bundles than MBP-fascin 2b. On the other hand,

His-tag espin 2b actin-bundling activity gets saturated earlier than MPB-fascin 2b. The average thickness of His-tag espin 2b-actin bundle does not increase much when His-tag espin 2b concentration is higher than 2 µM. There might be some coordination between fascin 2b and espin 2b when actin bundles are formed in vitro, but this effect is small.

86 This is different than previous studies of fascin 1 and human espin, which showed that two proteins can build much thicker actin bundles than the one formed by either of them alone (Chaessens et al., 2008). There are several possible explanations for this conflict.

First, this may be due to the difference among the isoforms in the same protein family: fascin 1 and fascin 2 share only 56% identity. The 44% difference in sequence may lead to the ortholog variance in cobundling with espin. Second, the components in or the pH of the reaction buffer may be crucial to the coordination between actin and fascin 2b or actin and espin 2b. Third, the MBP tag may block the association between these two cross-linkers or fascin 2b interaction with actin.

The purpose of this study was to determine if actin-binding proteins coordinate with each other in vitro and give some understanding to the in vivo situation, although this is only the first small step for this long journey to eventually apply the idea and bring the potential benefits to humans. This study also brings out more questions about how actin- binding proteins localize to hair cell stereocilia coordinate with each other in vivo. The very limited cooperation between MBP-fascin 2b and His-tag espin 2b in vitro can not fully prove that there is no cooperation between them in vivo, because many other factors may contribute in vivo. Also, because there are many other actin-binding proteins localized to hair cell stereocilia, such as fimbrin, it is possible that although fascin 2b and espin 2b cooperation is not significant, those proteins have more significant coordination with other actin-binding proteins. These questions are waiting for more in vivo and in vitro studies to discover the answers. In addition, the newest TEM in the CCMSB building could be used to collect a series of tilted images and provide a higher contrast, 3-

87 D reconstruction of the actin bundle formed by MBP-fascin 2b and/or His-tag espin 2b with higher resolution. More structural information about the actin bundle will be provided with the better models with higher resolution. It is important to measure the inter-filament distance and the inter-cross-linker distance of the MBP-fascin 2b-actin bundle, His-tag espin 2b-actin bundle, and the actin bundle formed by the two proteins together. Finally, in the future, we could develop an understanding of the distribution of these two cross-linkers on the same actin bundle by comparing the inter-filament distance and inter-cross-linker distance with the distances measured from actin bundles formed by only one or the other cross-linker.

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