Acf7 Is a Hair-Bundle Antecedent, Positioned to Integrate

ACF7 IS A HAIR-BUNDLE ANTECEDENT, POSITIONED TO INTEGRATE

CUTICULAR PLATE ACTIN AND SOMATIC TUBULIN

PATRICK J. ANTONELLIS

Submitted in partial fulfillment of the requirements

For the degree of Master of Science

Thesis Advisor: Dr. Brian M. McDermott

Department of Biology

CASE WESTERN RESERVE UNIVERSITY

May, 2013

CASE WESTERN RESERVE UNIVERSITY

SCHOOL OF GRADUATE STUDIES

We hereby approve the thesis/dissertation of

Patrick J. Antonellis ______

Master of Science candidate for the ______degree *.

Roy E. Ritzmann (signed)______(chair of the committee)

Brian M. McDermott ______

Stephen E. Haynesworth ______

Claudia Mizutani ______

______

3/22/2013 (date) ______

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

TABLE OF CONTENTS

LIST OF FIGURES……………………………………………………………………….2

LIST OF ABREVIATIONS………………………………………………………………3

ABSTRACT…………...…………………………………………………………………..4

INTRODUCTION………………………………………………………………………...5

Hearing and Deafness and the Ear…………………………………...... 5

The Hair Cell………………………………………………………………………8

Cytoskeleton dynamics…………………………………………………………..12

Important microtubule and actin interacting proteins……………………………14

MATERIALS AND METHODS……………………………………………………...... 19

RESULTS AND CONCLUSIONS……………………………………………………...26

ACF7 Results…………………………………………………………………….26

ACF7 Conclusions……………………………………………………………….41

Class IX Myosins Results………………………………..…………..…………..46

Class IX Myosins Conclusions……..…………………………………..………..49

REFERENCES………………………………………………………………………….52

LIST OF FIGURES

FIGURE 1…………………………………………………………………………………6

FIGURE 2………………………………………………………………………..………10

FIGURE 3………………………………………………………………………………..27

FIGURE 4………………………………………………………………………………..29

FIGURE 5………………………………………………………………………………..31

FIGURE 6………………………………………………………………………………..33

FIGURE 7………………………………………………………………………………..35

FIGURE 8………………………………………………………………………………..37

FIGURE 9………………………………………………………………………………..40

FIGURE 10………………………………………………………………………………42

FIGURE 11………………………………………………………………………………46

FIGURE 12………………………………………………………………………………48

TABLE 1…………………………………………………………………………………..8

LIST OF ABBREVIATIONS

ACF7, actin cross-linking family protein 7; MACF1, microtubule actin cross-linking factor 1; MYO9A, myosin9a; MYO9B, myosin 9b; F-actin, filamentous actin; G-actin, globular actin; CH, calponin homology; GAR, GAS2-related; GSR, glycine-serine- arginine; GSK3b, glycogen synthase kinase 3b; GAPs, GTPase activating proteins,

RhoGAP, Rho GTPase-activating protein; Arp2/3, actin-related proteins 2+3; PCDH15, protocadherin 15; RPKM, reads per kb per million total mapped reads; dpf, days- postfertilization; P2, postnatal day 2; TH, tail homology; MAPs, microtubule-associated proteins; +Tips, microtubule plus end tracking proteins

Acf7 is a Hair-Bundle Antecedent, Positioned to Integrate Cuticular Plate Actin and

Somatic Tubulin

Abstract

PATRICK J. ANTONELLIS

The hair cell of the inner ear converts mechanical stimuli, such as sound waves, into electrical impulses which are then sent to and interpreted by the brain. Proper organization of the actin and microtubule cytoskeletons is important for establishing the distinct morphology of the hair cell, which is crucial for its function. Here we identify the genes ACF7, MYO9A, and MYO9B are expressed in hair cells, and may be important for hair cell development and function. Through zebrafish transgenesis and indirect labeling of mouse hair cells we show that ACF7 is positioned to integrate the actin and microtubule networks in hair cells. In addition, electron micrographs demonstrate microtubules inserting into the cuticular plate, the ends of which are decorated with proteins that directly link to the F-actin of the cuticular plate. ACF7, which possesses both actin and microtubule binding domains, is a prime candidate to be this linker.

Introduction

Hearing, deafness, and the ear

Hearing and balance are complex senses that are extremely important for interpreting and interacting with the world. In the United States, approximately 36 million adults report some degree of hearing loss (National Institute on Deafness and

Other Communication Disorders). There are a variety of causes of hearing loss such as age, exposure to loud noises, illness, physical trauma, exposure to ototoxic compounds, as well as genetic factors.

For proper hearing and balance, the forces produced by sound waves, gravity, and head movements must be converted into information which can be sent to and interpreted by the brain. The organ primarily responsible for this process is the ear. The ear is split into 3 parts, the outer, middle, and inner ear (Figure 1A). The outer ear, also known as the pinna, funnels sound waves from the external environment to the middle ear. The middle ear is composed of the tympanic membrane, or ear drum, as well as three bones connected in series known as the malleus, incus, and stapes. When the sound waves collected by the outer ear hit the tympanic membrane it vibrates, which in turn vibrates the bones of the middle ear. This vibration is then passed on to the inner ear where it is processed into electrical impulses that are sent to the brain. The inner ear is composed of two parts, the cochlea and the vestibular system which function in hearing and balance respectively. The vibrations of the middle ear are sent to the cochlea while the vestibular system functions to detect angular and linear accelerations.

Figure 1. (A) The three regions of the ear; external, middle and inner ear. The middle ear is composed of the tympanic membrane and the ossicles (malleus, incus, and stapes). The inner ear consists of both the cochlea which functions in sound transduction and the vestibular labyrinth which detects angular and linear accelerations as well as gravitational forces. (B) A cross section of the cochlea reveals the organ of Corti which consists of three rows of outer hair cells and a row of inner hair cells sitting on top of the basilar membrane and overlayed by the tectorial membrane. Adapted from (Schwander et al., 2010). ©Schwander et al., 2010. Originally published in The Journal of Cell Biology. doi: 10.1083/jcb.201001138. The cochlea is a spiral shaped bony structure filled with fluid. The basilar membrane runs the length of the cochlea and vibrates in response to the stimulus provided by the bones of the middle ear and these vibrations are propagated from the base to the apex of the cochlea. A sensory epithelium known as the organ of Corti sits on top of the basilar membrane (Figure 1B). The sensory cell of the organ of Corti is the hair cell, which is named due to the hair-like projection at its apical surface. There are four rows of hair cells in the organ of Corti, three rows of outer hair cells and one row of inner hair cells. The inner hair cells are the major sensory output of the cochlea, while the outer hair cells are part of a mechanical amplifier system that amplifies the signal to the inner hair cells. Overlying the hair cells is the tectorial membrane. The oscillation of the basilar membrane causes the hair cells to move, this movement causes the bundles of the hair cells to be deflected by the techtorial membrane. It is this action which depolarizes the sensory hair cells.

There are two major components to the vestibular system, the semicircular canals and otolith organs. As in the cochlea, the sensory cell of the vestibular system is the hair cell. The semicircular canals function to detect rotational force and therefore there are three semicircular canals in the vestibular system, one for each plane of rotation. Each canal is filled with fluid which moves within the canal in response to angular head movements. The movement of fluid within the semicircular canal pushes on the cupula, stimulating the hair cells within. The otolith organs are responsible for detecting linear accelerations and are named because of the crystal otoconia they contain. The otolithes rest on a gel layer which overlies a sensory patch of hair cells. Linear accelerations cause the otolithes to move which leads to stimulation of the underlying hair cells.

The hair cell is the major sensory cell of the inner ear. Therefore, it is not surprising that many forms of deafness are caused by mutations in genes expressed in hair cells (Table 1). In addition, more than 90 percent of hearing loss occurs when either hair cells or auditory nerve cells are destroyed (National Institute on Deafness and Other

Communication Disorders). Therefore, to fully understand how hearing happens and what causes certain hearing defects, it is important to functionally and morphologically characterize the normal developmental processes of the hair cell.

Table 1. Partial list of genes expressed in the hair cell that are involved in deafness. Isolated forms of deafness are characterized by either autosomal dominant (DFNA) or autosomal recessive (DFNB) mode of inheritance. Adapted from (Petit and Richardson, 2009)

Gene Protein Forms of human deafness MYO1A Myosin IA (motor protein) DFNA48 MYO1C Myosin-1c (motor protein) DFNA MYH9 Myosin IIA (motor protein) DFNA17; Fechtner syndrome MYO3A Myosin IIIA (motor protein) DFNB30 MYO6 Myosin VI (motor protein) DFNA22; DFNB37 cardiomyopathy syndrome MYO7A Myosin VIIA (motor protein) DFNB2; DFNA11; Usher syndrome 1B MYO15A Myosin XV (motor protein) DFNB3 ACTG1 g-actin (cytoskeletal protein) DFNA20/26 ESPN Espin (actin-bundling protein) DFNB36; DFNA RDX Radixin (actin-binding protein) DFNB24 USH1C Harmonin (PDZ domain–containing protein) DFNB18; Usher syndrome 1C USH1G Sans (cytoskeletal protein) Usher syndrome 1G DFNB31 Whirlin (PDZ domain–containing protein) DFNB31; Usher syndrome 2D CDH23 Cadherin-23 (integral membrane adhesion DFNB12; Usher syndrome 1D protein) PCDH15 Protocadherin-15 (cell adhesion protein) DFNB23; Usher syndrome 1F TMC1 TMC1 (transmembrane channel-like protein) DFNB7/11; DFNA36

The hair cell

Hair cells are the mechanotransducers of the inner ear, converting mechanical

stimulus to electrical impulses. In the cochlea the stimulus is vibration that originated as

sound waves, while in the vestibular system the stimulus is gravitational forces as well as

angular and linear accelerations. Regardless of the origin of the stimulus, the process of

mechanotransduction is the same in all hair cells. In order for hair cells to perform their

function as a mechanotransducer they possess precisely built and organized structures

8 critical for this process. However, many of the mechanisms required for establishing and maintaining the morphology of the hair cell remain unknown.

Hair cells are named because of their hair-like projections at their apical surface called hair bundles. The hair bundle is the mechanosensative organelle of the hair cell which is composed of actin-based projections called stereocilia (Figure 2). The stereocilia are stiff, actin-filled rods with a tapered base that acts as a flexion point

(Tilney and DeRosier, 1986). The core of each stereocilia is made of parallel bundles of filamentous actin (F-actin) that are unidirectionally aligned with their plus end at the tip and minus end at the base. Most of these actin filaments terminate in the taper region; however, a few from the core of the stereocilia penetrate deep into the cuticular plate to form rootlets which serve to anchor the stereocilia (Tilney et al., 1980). The cuticular plate is a dense F-actin meshwork at the apical surface of the cell, beneath the hair bundle, that is thought to provide a strong platform on which the bundle can pivot, however its exact role and importance is unknown (DeRosier and Tilney, 1989). The cuticular plate is connected to the soma of the cell by microtubules oriented parallel to the hair cell’s longitudinal axis (Jaeger et al., 1994). In addition, microtubules eminate from the basal body, which resides in the fonticulus, a region in the cuticular plate devoid of F-actin (Chacon-Heszele et al., 2012). It is not known whether direct physical linkages are required to connect the cuticular plate to the microtubule cytoskeleton.

Figure 2. A detailed schematic of the hair cell flanked by supporting cells. The hair bundle at the apical surface consists of both actin based stereocilia and a microtubule based kinocilium. The stereocilia are organized into rows of graded height, with the tallest row next to the kinocilium. Adjacent stereocilia are connected by tip links which open mechanically gated ion channels when the bundle is deflected in the direction of the kinocilium. Stereocilia become thinner at the taper region and a few long actin filaments enter the actin-based cuticular plate where they form rootlets. Once depolarized, neurotransmitter is released to afferent nerves which signal to the central nervous system. Efferent nerves provide information to the hair cell from the central nervous system.

A mature hair bundle has a distinct morphology with the stereocilia arranged in rows of graded height. The direction of the tallest row of stereocilia denotes the morphological axis of the bundle. Individual stereocilia within the bundle are connected together by a variety of extracellular linkages which help provide the bundle with rigidity and ensures that all stereocilia within the bundle move in unison (Bashtanov et al., 2004).

At the tip of stereocilia, tip links connect the tips of shorter stereocilia to the side of the adjacent taller stereocilia along the morphological access of the bundle (Pickles et al.,

1984). When the bundle is deflected, the tip links are tensed causing ion channels associated with the tip links to open, allowing positively charged ions such as Ca+ and K+ to enter and depolarize the cell (Corey and Hudspeth, 1983). Depolarization of the hair cell causes transmitter release to neurons which synapse along its basolateral surface.

From studies on chick hair cells, the developmental stages of the hair bundle are well described (Tilney et al., 1992). At the beginning of morphogenesis the kinocilium erupts from the center of the apical surface of the cell followed by the growth of short microvilli of equal height. The kinocilium then migrates to one side of the cell; this migration is the first sign of hair bundle polarity. The microvilli adjacent to the kinocilium then elongate to form the tallest row of stereocilia, followed by the subsequent shorter rows. These processes require the organization of both F-actin and microtubule based structures as well as coordinated movements between the two networks.

Identifying genes involved in building and shaping these structures is critical for understanding how the hair cell works.

Cytoskeleton Dynamics

Microtubules

Microtubules are filaments composed of thirteen protofilaments, with each protofillament being a linear strand of tubulin dimers that has a slow growing minus end and a fast growing plus end. Microtubules undergo “dynamic instability” which is a process of continual growth and collapse. Dynamic instability occurs because the more stable form of tubulin, GTP-tubulin, is added to the plus ends of microtubules. However, because of continual GTP hydrolysis, new GTP-tubulin must be added to the plus end regularly in order to maintain stability. Therefore, when microtubule growth stalls they are prone to collapse (Desai and Mitchison, 1997).

There are many different proteins that associate with microtubules and perform a variety of functions. Microtubule-associated proteins (MAPs) bind along more mature microtubules and provide stability. Another class of proteins, the microtubule plus end tracking proteins (+Tips), bind specifically to the plus ends of microtubules and help regulate microtubule plus end dynamics (Akhmanova and Hoogenraad, 2005). In addition, +Tips provide an attachment point to the cell cortex which is critical to processes such as vesicle delivery and signaling for cortical polarity. The kinesins are a family molecular motor proteins which travel along microtubules. Kinesins carry cargos, such as organelles, protein complexes, and mRNAs, which they transport throughout the cell (Wu et al., 2006).

Microtubules are important for the establishment of cell polarity which is a critical first step in important cellular processes such as morphogenesis and migration.

During the establishment of cell polarity microtubule growth becomes polarized.

Directional growth of microtubules is achieved by a variety of interacting proteins that stabilize microtubules, regulate microtubule dynamic instability, and capture and localize microtubules. The directional growth of microtubules causes the microtubule organizing center of the cell and the Golgi complex to become reoriented. This leads to microtubule driven cargo, such as vesicles, becoming asymmetrically distributed within the cell

(Siegrist and Doe, 2007).

The microtubule cytoskeleton is extremely important for the structure and function of the hair cell. Specifically, the microtubule based kinocilium is known to be critical for establishing hair bundle polarity as well as mechanotransduction. In addition, microtubules of the cell soma may be important for establishing basal-lateral polarity during morphogenesis.

Actin

Actin is the most abundant of any intracellular protein. F-actin is a linear polymer made of numerous monomers of globular actin (G-actin). G-actin binds ATP, and when in solution, G-actin can polymerize into F-actin involving the hydrolysis of

ATP to ADP. However, this is a dynamic event and F-actin can be depolymerized into

G-actin. When in a filament all G-actin monomers face the same direction and, therefore, F-actin has a polarity, with actin polymerization occurring at the plus end and depolymerization occurring at the minus end (Dominguez and Holmes, 2011).

Within the cell, actin filaments are organized and shaped by a number of interacting proteins. Actin nucleators are proteins that help build actin filaments by

13 increasing the rate at which polymerization occurs. Nucleators can help elongate existing filaments or induce filament branching (Campellone and Welch, 2010). Actin bundling proteins bind multiple F-actin filaments and bundle them into parallel actin bundles.

Also, similar to kinesins that travel along microtubules, myosins are molecular motors, which move along actin tracks. Most myosins form a homodimer which binds to F-actin and “walks” towards the plus end of the filament, often carrying cargo with them.

The major structure that defines the shape of a cell is the cortex, which is a dense actin network directly beneath the plasma membrane. In addition, filamentous actin is a primary component of cellular structures such as lamellipodia and stress fibers, which are necessary for cellular functions such as migration. The coordinated reorganization of the actin cytoskeleton is necessary for changes in polarity or cell movement. For example, actin polymerization is greatly increased at the leading edge of migrating cells (Theriot and Mitchison, 1991).

Important microtubule and actin interacting proteins

Spectraplakins

Spectraplakins are extremely large cytoskeletal crosslinking proteins which share similarities to both the spectrin and plakin superfamilies. Spectrin superfamily members crosslink F-actin and plakin superfamily members bind intermediate filaments but can also crosslink other cytoskeleton elements. The spectraplakins can interact with all three cytoskeleton components, microtubules, F-actin, and intermediate filaments. There are two spectraplakin genes in mammals, bullous pemphigoid antigen 1 (BPAG1) and actin cross-linking family protein 7 (ACF7) (also known as MACF1; microtubule actin cross-

14 linking factor 1), and both genes produce multiple splicing isoforms which can be almost

9000 amino acids in length (Roper et al., 2002). ACF7 contains an N-terminal actin binding domain, a spectrin domain, EF hand domains, and microtubule associating domains at the C-terminus. The actin binding domain is composed of two calponin- homology (CH) domains in tandem with each other. The spectrin domain consists of multiple spectrin repeats, with each repeat forming a bundle of 3 alpha-helices. In the C- terminus, EF hands bind calmodulin and the GAS2-related (GAR) and glycine-serine- arginine (GSR) domains can associate with microtubules, (Sonnenberg and Liem, 2007).

There is evidence of coordinated movements between actin and microtubule networks. It has been observed that microtubules grow along F-actin bundles in epithelial cells (Salmon et al., 2002) and microtubules stop growing when the actin network is perturbed in neural growth cones (Schaefer et al., 2002). However, it remains unknown if direct physical linkages between actin and microtubules are necessary for these processes. The ability of spectroplakin proteins to bind both F-actin and microtubules presents the possibility of spectroplakins being important integrators of these cytoskeleton components.

ACF7 has been demonstrated to be an essential regulator of microtubule dynamics. Underscoring its critical role as a cytoskeletal linker, mutations in Acf7 orthologs are embryonically lethal in both zebrafish and mice (Chen et al., 2006; Gupta et al., 2010; Kodama et al., 2003). In fibroblasts derived from Acf7 null mice, cytoplasmic microtubules show abnormal path finding and no longer track to the actin rich cortex as normal (Kodama et al., 2003). It was also noted that Acf7 null mice showed a similar phenotype to Wnt-3 knockout as well as Lrp5/6 double null mice, all of which exhibit

15 defects in primitive streak, node, and mesdoderm development, indicating a possible signaling role for ACF7as well (Chen et al., 2006). It was also shown that the microtubule-binding ability of ACF7 is regulated by glycogen synthase kinase 3b

(GSK3b), which is downstream of Wnt signaling (Wu et al., 2011). Hair cell development requires the coordinated reorganization of the actin and microtubule cytoskeletons, which is under the control of yet undescribed signaling pathways. ACF7 displays all of the characteristics of a protein integral to this process. Specifically, we hypothesize that ACF7 links the actin based cuticular plate to the microtubule cytoskeleton of the hair cell and may be involved in positioning of the kinocilium during development.

Myosins

Myosins are a family of actin-based molecular motors that were first described in muscle. There are 24 members of the myosin superfamily that drive a number of cellular functions such as cytokinesis, organelle transport, cell polarization and signal transduction. Myosin proteins can be divided into three domains, the myosin head domain, a neck domain, and a variable tail domain. The myosin head domain, the most conserved domain in the family, is the motor and site of actin binding of myosin proteins.

The motor head domain is powered by its ability to bind and hydrolyze ATP. The neck region binds regulatory myosin light chains. The tail domain is the interaction site for cargos carried by myosins, but it also varies widely between subtypes providing functional diversity to the superfamily. Myosins usually function as oligomers, with a typical dimer consisting of two head groups and a varying amount of light chains.

Myosins move by using the energy generated by ATP hydrolysis to induce

16 conformational changes in the head domains. This allows the head domains to “walk” along actin filaments, and this movement is directed towards the plus end of F-actin for the majority of myosins (Lodish, 2000).

There are seven myosins that are known to be involved in human deafness, myosin IA, myosin-Ic, myosin IIA, myosin IIIA, myosin VI, myosin VIIA, and myosin

XV (Petit and Richardson, 2009). Of these, the majority localize to the hair cell. Myosin

VIIa is one of the genes encoded by Usher Type I syndrome and a defect in this gene leads to malformed hair bundles (Petit and Richardson, 2009). Myosin VIIA has been shown to localize to the upper tip link density (UTLD) and is thought to be involved with establishing the resting tension of the tip link (Grati and Kachar, 2011). Myosin XVA localizes to the tips of stereocilia and when mutated, leads to shortened stereocilia

(Belyantseva et al., 2005). Myosin-1c has been shown to be a component of the adaptation-motor complex of hair cells which carries out slow adaptation and provides tension to sensitize the transduction channels (Gillespie and Cyr, 2004).

Class IX myosins were first discovered in a screen for unconventional myosins in mammals (Bement et al., 1994). Invertebrates have a single myosin IX gene while vertebrates have two, MYO9A and MYO9B. Class IX mysosins have a conserved myosin head domain, but differ from traditional myosins in three notable ways, an N- terminal extension of about 145 amino acids, a loop 2 insertion of 145 or more residues, as well as a Rho GTPase-activating protein (RhoGAP) domain in the tail (Coluccio,

2008). This RhoGAP domain enables class IX myosins to play a signaling role because of the ability of RhoGAPs to regulate RhoGTPases, small GTP-binding proteins

(Reinhard et al., 1995).

GTPases are molecular switches that exist in either an active GTP bound state or an inactive GDP bound state. GTPase activating proteins (GAPs) stimulate the intrinsic

GTPase activity of the protein, inducing it to cleave GTP to GDP, inactivating itself. In the active GTP bound state, Rho GTPases perform their regulatory functions by interacting with effector proteins. One such effector, Rho, is able to regulate actin cytoskeleton organization through the actin nucleators formin and the actin-related proteins 2+3 (arp2/3) complex (Jaffe and Hall, 2005). Myo9b has been shown to be important for the production of actin based projections and migration in macrophages by inhibiting the activity of the actin depolymerization factor cofilin (Hanley et al., 2010).

The ability of class IX myosins to interact with F-actin through their head domain and influence actin cytoskeleton dynamics through their RhoGAP domain makes them an ideal candidate to be playing a role in hair cell morphogenesis.

Materials and Methods

Animals

Transgenic strain Gt(macf1-citrine)ct68a/+ (Trinh le et al., 2011)

(http://www.fliptrap.org/static/index_new.html) or the wild-type strain Tübingen (Tü) were used in these investigations. Zebrafish were maintained and bred at 28°C. White leghorn chickens and FVB/NJ mice were kept according to standard procedures. All animals were kept with the approval of the Case Western Reserve University Institutional

Animal Care and Use Committee.

Hair-cell isolation and RNA-seq

Generation of cDNA from chick hair cells and RNA-seq was performed by Brian

McDermott. For hair-cell isolation, chicken hair cells were used because they survive for long periods after dissociation, a property that facilitates isolation of individual cells with a glass micropipette. White leghorn chickens less them 3 weeks of age were euthanized with CO2 and immediately decapitated. The inner ears were dissected and the sensory patches were removed and placed in dissociation chicken saline solution (154 mM NaCl,

6 mM KCl, 0.1 mM CaCl2, 8 mM glucose, 5 mM HEPES [pH=7.4]) with 0.1 mg per ml

Type XXIV endopeptidase (Sigma). After a 20 minute incubation, then trituration, hair cells were isolated by picking with a glass micropipette (McDermott et al., 2007). RNA from 200 hair cells was isolated, and cDNA was synthesized and amplified (Ovation

RNA-Seq System, NuGen). cDNA was subjected to massively parallel sequencing

(Ozsolak and Milos, 2011) using a HiSeq (Illumina) or a HiScan (Illumina).

Analysis of RNA-seq data was performed by Lana Pollock. From RNA-seq data, we calculated the single-end and paired-end read counts associated with MACF1 and

PCDH15. For single-end sequencing, there were 3,388 reads, per 36,352,275 total mapped reads, that aligned to the MACF1 locus. For PCDH15-assocated single-end sequencing, 512 sequences, per 36,352,275 total mapped reads, aligned to the PCDH15 locus. Therefore, RPKM (reads per kilobase per million total mapped reads) values are

6.02 associated with MACF1 and 2.09 associated with PCDH15. For paired-end sequencing, there were 13,020 reads, per 103,045,330 total mapped reads, that aligned to

MACF1 and 1,286 reads that aligned to PCDH15, revealing RPKM values of 8.16 and

1.84 associated with MACF1 and PCDH15, respectively.

RT-PCR

RT-PCR experiments were performed to confirm transcript expression of our genes of interest in the inner ear of both zebrafish and mice. Template cDNA was made from mouse maculae as well as whole zebrafish embyos, zebrafish maculae, and picked and purified zebrafish hair cells. For whole fish cDNA, RNA was extracted from approximately 20 wild type 7dpf embryos and treated with DNaseI to eliminate possible genomic DNA contamination (RNeasy Micro kit; Qiagen, Valencia, CA). Next, randomly primed first strand cDNA synthesis was performed (Super-Script III Reverse

Transcriptase; Invitrogen, Carlsbad, CA). For maculae, tissue was dissected from either adult FVB/NJ mice or adult zebrafish and used to generate cDNA as described above.

Hair cell cDNA from adult zebrafish was produced as previously described (McDermott et al., 2007).

Interexonic primers were designed that span multiple exons, allowing for differentiation between amplification from transcripts and the genomic DNA locus. For

Acf7, the primer pair used to amplify from mouse template cDNA was mus_macf1_1

Forward (5’- GAAGATTTCCTCTTAGAACTCAATAG -3’) and mus_macf1_1 Reverse

(5’- TGATCAAGACAACATCCTGTTTCTG -3’) and the pair used to amplify from zebrafish template cDNA was zf_macf1_8 Forward (5’-

CCCAGTCTCATTCTGGACACTGT -3’) and zf_macf1_8 Reverse (5’-

TTCCTCCAGCTTGTGTTGCCTCT -3’). For Myo9a, the primer pair used to amplify from mouse template cDNA was mus_myo9a_1 Forward (5’-

CAGTGACTCCGAAATGACATCACA-3’) and mus_myo9a_1 Reverse (5’-

CATCTTTTTCAGAAGAAACTCATCC -3’) and the primer pair used to amplify from zebrafish cDNA was zf_myo9a_4 Forward (5’- CTCAGAGAAGCTTCCTCTGCGG-3’) and zf_myo9a_4 Reverse (5’- CCTCCACCAGTTCTGAATTTGGC -3’). Successful amplification from zebrafish hair cell cDNA required nested PCR. Nested PCR primer pairs zf_myo9a_nested 4 Forward (5’- GTCTGGTGCTAAGACAACTTCGC-3’) and zf_myo9a_nested 4 Reverse (5’-TGTCTTACTCTGAGGAAGTTCTTTC -3’) were designed to sit just downstream and upstream of zf_myo9a_4 Forward and zf_myo9a_4

Reverse respectively. Following the first round of PCR using the zf_myo9a_4 Forward and zf_myo9a_4 Reverse primers, 1ul of product was used as template for nested PCR.

For all PCR amplifications, Ex Taq DNA polymerase was used (Ex Taq DNA

Polymerase; Takara Bio).

Primary amino acid sequence alignments

Sequence analysis software (MacVector, MacVector Inc.) was used to align the full-length Acf7 isoforms from mice (ENSMUSP00000095507), chickens

(ENSGALP00000005847), and zebrafish (ENSDARP00000054273). SMART was used to identify relevant protein domains (Letunic et al., 2012).

Fish labeling and imaging

Transgenic strain Gt(macf1-citrine)ct68a zebrafish at 5 dpf were immunolabeled with anti-acetylated tubulin at a 1:1000 dilution (6–11B-1; Sigma) and/or labeled with

Alexa Fluor 546 phalloidin (Invitrogen) at a 1:50 dilution according to standard procedures (Chou et al., 2011). Where appropriate, Alexa Fluor 405-labeled goat anti- mouse IgG at a 1:200 dilution (Invitrogen) was employed as secondary antibody. Images were acquired by confocal microscopy (Leica) with a 40 ×, 63 ×, or a 100 × objective and visualized with imaging software (Leica Confocal Software, Leica; Volocity;

PerkinElmer).

Zebrafish transgenesis

Zebrafish transgenesis was performed by Victoria Chou. To generate a zebrafish strain that transmits through the germ line, the plasmid pMT/SV/PV3b/beta- actin/mCherry at 100 ng per μl was co-injected with Tol2 RNA at 25 ng per μl (Balciunas et al., 2006) into embryos at the single-cell stage. The F0 and F1 generations were screened using a fluorescence stereoscope (Leica).

Immunolabeling of whole-mount, mouse, vestibular tissue

For each experiment 3 6-12 month old FVB/NJ mice were used. Maculae of mice were removed by dissection and fixed in 4% paraformaldehyde (PFA) in 1x phospho buffered saline (PBS) for 1 hour at room temperature. Following 3 10 minute washes in PBS, the tissue was permeabilized in 0.05% Triton X-100 in PBS for 2 hours at room temperature then blocked in 1% bovine serum albumin (BSA), 0.05% Triton X-100 in PBS for 2 hours at room temperature. Primary antibody diluted to the appropriate concentration in blocking solution (rabbit polyclonal to Myo9a @ 1:500 (Chieregatti et al., 1998); rabbit polyclonal to ACF7 @ 1:100 (Karakesisoglou et al., 2000); (Kodama et al., 2003); and/or mouse monoclonal to acetylated tubulin (T6793, Sigma Aldrich, St.

Louis, MO @ 1:50) was then added to the tissue and incubated overnight at 4°C.

Following one 30 minute wash in blocking solution, diluted secondary antibody (chicken anti-rabbit Alexa Flour 488 @ 1:200; goat anti-mouse Alexa Flour 405 @1:200;

Invitrogen) were added and F-actin was stained with Alexa Flour phalloiden 568

(Invitrogen) at a 1:50 dilution. After a final 30 minute wash in 1x PBS, the tissue was mounted on a microscope slide with vectashield mounting medium (Vectashield, Vector

Laboratories, Burlingame, CA). Imaging was performed on a laser-scanning confocal microscope (Leica) using a 40x or 63x objective.

Immunolabeling of mouse cochlear cryosections

Whole inner ears were dissected out of 6-12 month old FVB/NJ mice and processed as described previously (Whitlon et al., 2001). Briefly, ears were fixed in 4%

PFA in 1x PBS for 1 hour then decalcified in 10% EDTA for approximately 4 days at

4°C. Ears were then washed 3 times for 30 minutes in progressively increasing amounts of sucrose then incubated in embedding medium (OCT; Sakura Finetek, Torrence, CA)

23 overnight at 4°C. Ears were then placed into a mold, covered in OCT and frozen. The frozen molds were cut into 10um sections using a cryostat and the sections placed on a microscope slide. To immunolabel sections, they were postfixed with 4% PFA for 15 minutes, washed 3 times with PBS, and blocked with 5% goat serum and 0.1% Triton X-

100 in PBS for 1 hour. Sections were sequentially incubated with primary and secondary antibodies diluted in 5% goat serum as described above. Finally, sections were set in

Hard-Set Vectashield (Vector Laboratories) with a cover slip and imaging was carried out as described above.

Immunolabeling of cultured mouse cochlea

Cultures of mouse cochlea were prepared as previously described (Russell and

Richardson, 1987). First, dissection and culture media was prepared. Dissection medium consisted of 5mL HEPES (Sigma) and 50mL Hank’s Buffered Salt Solution (Invitrogen) while culture media contained 49.5mL Dulbecco’s Modified Eagle Medium (Invitrogen),

0.5mL N1 supplement (Sigma), and 250ul 10,000Unit/mL Penicillin G (Sigma). Postnatal day 2 FVB/NJ pups were cleaned with 70% ethanol and decapitated. The inner ear was then removed and placed in dissection medium. The cochlea was then removed from the bony labyrinth and placed into a coverslip-bottom dish (MatTek, Ashland, MA) coated with cell and tissue adhesive (Cell-Tak, BD Biosciences, San Jose, CA) and containing culture medium. Following overnight culture at 37°C, cochleas were fixed with 4% PFA and sequentially incubated with primary and secondary antibodies, mounted on slides, and imaged as described above using a 100x objective.

3-dimensional modeling

For 3-dimensional reconstructions, a laser-scanning confocal microscope was used to acquire a series of confocal sections, each 0.122 μM, through labeled tissue; the resulting z-stacks each represent the entire volume of the region of interest. Using volume-rendering software (Volocity; PerkinElmer), z-stacks were then converted into

3D models, which were used to create movies and still frames. In Figure 3 G-L, the green channel is amplified 2 ×. In addition, the rendering software was used to identify regions of overlap between the actin- and Acf7-associated signals and to generate a new channel to represent the overlap. To view only areas where overlap occurs, the non-overlapping signal was removed and the overlapping signal was amplified (Fig. 2G = 2.5 ×, Fig. 3G =

2.5 ×, and Fig. 3M = 8 ×) to enable clear visualization.

Results and Discussion

Acf7 Results

Acf7 expression is conserved in vertebrates

Acf7 was initially found to be present in a microarray study of zebrafish hair cell

RNA (McDermott et al., 2007). Next, to see if ACF7 is expressed in the hair cells of other vertebrates we determined the chicken hair cell transcriptome. Hair cells from the chicken ear were isolated, cDNA was generated, and massively parallel sequencing was performed by Brian McDermott (Ozsolak and Milos, 2011). Through analysis of this data by Lana Pollock, we found that ACF7 mRNA was highly represented (Figure 3A) when compared to other genes known to be specifically expressed in hair cells, such as protocadherin 15 (PCDH15) (Alagramam et al., 2001). We calculated the reads per kb per million total mapped reads (RPKM) for both ACF7 and PCDH15 associated reads.

Single–end read counts yielded RPKM values of 6.02 and 2.09 for ACF7 and PCDH15 associated signals, respectively. To better visualize the expression pattern of acf7 mRNA in zebrafish whole mount in situ hybridization on 5-days-postfertilization (dpf) larvae was performed by Tracy Chen. Labeling of probe appeared to be specific to the ear

(Figure 3B), this was confirmed by a magnified view of the otocyst which exhibits high acf7 expression (Figure 3C). To confirm the expression of Acf7, RT-PCR was performed on cDNA libraries prepared from both zebrafish and mouse maculae.

Transcripts were successfully amplified from zebrafish and mouse (Figure 3F, G), confirming that Acf7 is expressed in the sensory epithelia of both species.

Figure 3. ACF7 mRNA is expressed in hair cells or maculae of zebrafish, chickens, and mice. (A) Expression of ACF7 was detected in isolated chicken hair cells by RNA-seq. The depth of reads aligning to the chicken genome is visualized using the Integrated Genome Browser (http://bioviz.org/igb/) with splice junctions predicted using TopHat software shown in red. Exons of the corresponding genes identified using Ensembl are displayed as blue. (B) RNA in situ hybridization experiment on a 5-dpf, whole-mount larvae with antisense probe demonstrates that acf7 mRNA is present in the otocyst (arrowhead). (C) Image of a zebrafish otocyst under higher magnification demonstrates expression in hair cells of the anterior macula and in the posterior crista. (D and E) Acf7 sense RNA was used as a control. (F and G) RT- PCR analyses demonstrate that Acf7 mRNA is present in maculae of zebrafish and mice. (F) Lane 1, image of agarose gel demonstrates that the acf7 gene product is detected in macular cDNA of zebrafish with primer pairs, zf_macf1_8 Forward and zf_macf1_8 Reverse. Lane 2, no PCR product is detected in a reaction that contains water in lieu of macular cDNA. (G) In lanes 1, 3, and 5, Acf7 cDNA was amplified from cDNA produced from mouse vestibular tissue, primer pairs mus_macf1_1 Forward and mus_macf1_1 Reverse, mus_macf1_2 Forward and mus_macf1_2 Reverse, and mus_macf1_3 Forward and mus_macf1_3 Reverse, respectively, but not from reactions where water was used instead of cDNA (Lanes 2, 4, and 6).

Taken together these data suggest that Acf7 expression in hair cells is conserved amongst the vertebrates. Due to this finding we investigated whether the primary amino acid sequences of these species showed similarities (Figure 4). We found that the actin binding CH domains (Figure 4B), EF Hand domains (Figure 4C), and microtubule interacting GAS2 (Figure 4D) domains exhibit extremely high conservation. Because of the high conservation in these functional domains we hypothesized that ACF7 shows similar localization patterns in these species.

Figure 4. Comparison of the primary amino acid sequences of zebrafish, chicken, and mouse ACF7s. (A) Schematic of ACF7 protein with relevant domains. (B) The actin-binding domains CH1 and CH2 (yellow), (C) EF hand domains (green), and (D) the microtubule- interacting GAS2 domain (blue) exhibit a very high degree of amino acid sequence similarity, suggesting that ACF7’s ability to bind microtubules and F-actin is conserved among vertebrates.

Acf7 encircles, underlies, and interwoven into the cuticular plate of zebrafish hair cells

For these studies, we obtained a zebrafish strain which expresses a citrine coupled

Acf7 fusion protein driven by the endogenous promoter. This strain contains a FlipTrap vector inserted in the acf7 gene, allele Gt(macf1-citrine)ct68a, on chromosome 19 (Figure

5A) resulting in the expression of Acf7 fusion protein with Citrine protein located between the plakin and spectrin-repeat domains (Figure 5B) (Trinh le et al., 2011)

(http://www.fliptrap.org/static/index_new.html). We performed confocal microscopy on

5-dpf Gt(macf1-citrine)ct68a/+ embryos to determine the localization of Acf7 in zebrafish hair cells. In posterior lateral line neuromasts labeled with phalloidin, Acf7 surrounds the

F-actin based cuticular plate and the fonticulus, which contains the basal body and doesn’t label with phalloidin (Figure 5C-E). This same pattern is observed in the posterior macula (Figure 5I). Looking at the citrine fusion protein only you can clearly see Acf7’s distinctive localization pattern (Figure 5C). A 3-dimensional reconstruction of a series of confocal images shows that Acf7 extends further into the cell body than the actin of the cuticular plate (Figure 5F). Also, when the reconstruction is analyzed for overlapping Acf7 and F-actin associated signals, Acf7 is found within cuticular plates

(Figure 5G). From a lateral view of an anterior macular hair cell, Acf7 is not only around the cuticular plate but also extends underneath it (Figure 5H). Additionally, in fish labeled with an anti-acetylated tubulin antibody Acf7 appears to cap the microtubule cytoskeleton of hair cells (Figure 5J). These data demonstrate that Acf7 is positioned between the microtubule cytoskeleton, which is more basal, and the cuticular plate, which is more apical.

Figure 5. Localization pattern of Acf-7-Citrine fusion protein in relation to the cytoskeleton of zebrafish hair cells. (A) Genomic structure of the acf7 locus, containing 97 exons (red bars), and the point of insertion (orange arrow) for the Fliptrap vector in the Gt(macf1-citrine)ct68a strain. Relevant elements of the Fliptrap vector are depicted: TE = transposable element, SA = splice acceptor, and SD = splice donor. Black arrow designates direction of the read. Black bar illustrates the segment of chromosome that contains acf7 and the cognate nucleotide positions that delimit the gene are represented. (B) Schematic of Acf7-Citrine highlighting major functional domains. (C) Neuromast hair cells of the posterior lateral line expressing Acf7-Citrine (yellow) are labeled with phalloidin linked to an Alexa Fluor. In C, D, F, H, and I, red is phalloidin. This apical view demonstrates that the fusion protein encircles the cuticular plate and a point beneath the kinocilium devoid of actin, the fonticulus. (D) Enlarged view of C. (E) Localization of Acf7 in a single hair cell. * and ϕ denote the positions of the cuticular plate and fonticulus, respectively. (F) A 3-dimensional reconstruction, generated from a z-series of confocal images, of a neuromast positioned for a lateral view demonstrates that Acf7 is more concentrated towards the basal portions of the cuticular plates. Hair bundles are marked with arrowheads, and the regions of the cuticular plates are indicated with a bracket. In G, the volume of red-yellow overlap from image F is pseudocolored in green, and regions without overlap were removed, demonstrating that there is significant overlap between the cuticular plate and the Acf7-Citrine. (H) Anterior macula hair cell is decorated below the cuticular plate with Acf7- Citrine. (I) Posterior macula hair cells have fusion protein encircling both the cuticular plate and the fonticulus. (J) Indirectly labeled microtubules (red) of lateral crista hair cells demonstrate that Acf7-Citrine is towards the ends of the microtubules proximal to the cells’ apical surface. * denotes position of nucleus. Scale bars = 1 µm.

The localization pattern of ACF7 is conserved in mice

A polyclonal antibody to ACF7 was used in immunolabeling studies to determine

where ACF7 localizes in adult mouse vestibular hair cells. In whole mount vestibular

hair cells colabeled with phalloidin, ACF7 can be seen not only around the circumference

of the cuticular plate, but adjacent to it’s basal face as well (Figure 6A). From an apical

view, ACF7 surrounds the cuticular plate and the fonticulus, which lacks phalloidin

labeling (Figure 6B). In addition, a cross section of the apical surface of a hair cell

shows that ACF7 lines the interior wall of the fonticulus (Figure 6C). We next wanted

to determine where ACF7 localizes in respect to cellular tubulin. By colabeling

32 vestibular hair cells with anti-ACF7 and anti-acetylated tubulin and labeling F-actin with phalloidin, ACF7 is seen at the interface between the F-actin of the cuticular plate and the microtubule cytoskeleton of the cell (Figure 6D). A fluorescent intensity profile shows distinct peaks for the signals associated with each protein with ACF7 located in between

F-actin and somatic microtubules (Figure 6E). We also noticed faint but consistent labeling of ACF7 antibody within the cuticular plate, so we generated a 3-dimensional reconstruction from serial confocal sections. The reconstruction shows that ACF7 extends beneath the cuticular plate (Figure 6F), but when non-overlapping signal is removed from the reconstruction, ACF7 is shown to localize within the cuticular plate as well (Figure 6G). From these results, ACF7 demonstrates a localization pattern consistent to that exhibited by fusion protein expressing transgenic zebrafish.

Figure 6. ACF7 localization in mouse vestibular hair cells. Whole mount vestibular hair cells labeled with an antibody to ACF7 (green) and F-actin labeled with phalloidin (red) (A-D, F). (A) From this lateral view ACF7 can be seen directly at the base of the cuticular plate (bracket). (B) An apical view shows ACF7 surrounds the actin-based cuticular plate and forms a ring around the fonticulus, which does not label with phalloidin. The pattern of antibody labeling agrees with the observed localization of Acf7 fusion protein in the zebrafish. (C) Lateral view of a hair cell’s apical region demonstrates that ACF7 coats the fonticulus’s lumen (bracket). (D) A hair cell colabeled with antibodies to ACF7 (green) and acetylated tubulin (blue) and labeled with phalloidin (red) shows that ACF7 is position between the tubulin cytoskeleton and the F-actin of the cuticular plate. (E) Fluorescence intensity profile using the yellow line region of interest (ROI) in D, demonstrating that ACF7 is at the interface between the F-actin and tubulin networks. Intensity scale is linear, but the units are arbitrary. X-axis represents the length of the yellow line ROI. (F) 3-dimensional reconstruction of hair cell in A; yellow indicates volume of red and green overlap. Bracket indicates position of cuticular plate. (G) ACF7 and F-actin associated signal overlap from F with non-overlapping signals removed. Scale bars = 1 µm.

To determine the distribution of ACF7 in auditory hair cells the same ACF7 antibody was used to immunolabel cultured mouse cochlea from postnatal day 2 (P2) mice. To accurately determine ACF7 position, a series of confocal sections was used to generate 3-dimensional reconstructions. In auditory hair cells, ACF7 is present in all three rows of outer hair cells as well as the inner hair cells (Figure 7A, B). Additionally,

ACF7 localizes around the cuticular plate and the fonticulus, which is consistent with previous observations; however, in all auditory hair cells ACF7 appears to be more highly concentrated closer to the fonticulus (Figures 7A, D). An apical view of two inner hair cells clearly demonstrates this asymmetric distribution (Figure 7C, D). A lateral view of these cells reveals that ACF7 extends underneath the cuticular plate

(Figure 7E) and when non-overlapping signal is removed, ACF7 can be seen colocalizing with the F-actin of the cuticular plate (Figure 7F).

Figure 7. ACF7 localization in mouse auditory hair cells. (A–F) 3-dimentional reconstructions of cultured mouse cochlear hair cells from P2 mice labeled with a ACF7 antibody (green) and phalloidin (red). (A) From an apical view ACF7 surrounds the cuticular plate and the fonticulus in both inner and outer hair cells. (B) ACF7 channel only shows stronger ACF7 labeling near the fonticlus with appreciable levels of labeling in the cuticular plates. (C–F) An enlarged view of two inner hair cells. (C) Apical view showing ACF7 localization around the cuticular plate and the fonticulus. (D) Green channel only displays a similar pattern to that seen in B. (E) This lateral view demonstrates that ACF7 resides basal to the cuticular plate and also within the cuticular plate. Yellow indicates overlapping signal. (F) Colocalization between ACF7 and F- actin from E is shown in yellow with all non-overlapping signal removed, demonstrating that ACF7 overlaps with cuticular-plate actin and actin at the lateral cortical surface of the hair cell. Scale bars = 1 µm.

Acf7 is a hair bundle and cuticular plate antecedent

We next endeavored to determine at what stage of hair-cell development Acf7 is positioned near the apical surface. The neuromast organ of zebrafish replaces hair cells throughout the life of the animal. When we analyzed neuromasts of phalloidin labeled

Gt(macf1-citrine)ct68a/+ 5-dpf embryos, we noticed that some cells contained apically localized Acf7-Citrine but did not have cuticular plates or hair bundles (Figure 8D, E).

Because these labeled cells were positioned near hair cells within the rosette, we hypothesized that these cells were supporting cells that were differentiating into hair cells and that Acf7 antecedes cuticular plate and hair-bundle formation. To confirm these cells are presumptive hair cells and not supporting cells which harbor Acf7-Citrine, Victoria

Chou generated stable transgenic zebrafish that express β-actin fused to the fluorescent protein mCherry expressed under the control of the parvalbumin 3b promoter

(McDermott et al., 2010). The parvalbumin 3b promoter is active in both supporting cells that are transitioning to hair cells and hair cells themselves. This novel transgenic line Tg(Parval3b: β-actin-mCherry) effectively labels stereocilia and cuticular plates of the lateral-line system with the actin fusion protein, as confirmed by phalloidin labeling

(Figure 8A), allowing for observation of developing hair cells in live animals.

Figure 8. The transgenic line Tg(Parval3b:mCherry-β-actin) produces mCherry-β-actin to label the hair bundle and the cuticular plate. (A) Confocal image reveals a neuromast expressing mCherry-β-actin fusion protein (red). Blue and white arrowheads mark a hair bundle and a cuticular plate, respectively. Expression in hair cells is variable. (B) The actin of the neuromast was labeled with phalloidin coupled to the fluorophore Alexa 488 (green). (C) Co-localization of mCherry-β-actin and phalloidin was detected. Acf7 localization precedes hair-bundle and cuticular-plate formation. (D) Hair cells of a neuromast of the posterior lateral line, which express Acf7-Citrine (yellow), are labeled with phalloidin (red). E displays only the red channel of D. The hair cells have cuticular plates, which are oriented opposite to each other. Two cells (arrowheads) within the rosette lack significant actin at their apical surfaces, that is they lack hair bundles and cuticular plates, but contain robust levels of Acf7-Citrine (D). (F and G) Confocal images of a neuromast from an offspring of a Gt(macf1-citrine)ct68a/+ × Tg(Parval3b:mCherry-β-actin) cross 24 hrs after treatment with neomycin. mCherry-β-actin of the cuticular plate and hair bundle are red and Acf7-Citrine is yellow. G displays red channel only. Four hair cells have regenerated and contain both Acf7-Citrine and mCherry-β-actin at apical regions; however, there are cells within the rosette that contain Acf7-Citrine and no significant amounts of mCherry-β-actin (arrowheads). (H and I) 48 hrs after neomycin treatment, the cells that lacked apical localization of mCherry-β- actin developed cuticular plates with planar polarity, confirming their status as hair cells (arrowheads).

In order to determine if Acf7 predates the hair bundle and cuticular plate at the apical surface of hair cells a regeneration assay was performed by Kevin Chen and Brian

McDermott. Hair cells of the lateral-line system regenerate after exposure to ototoxic chemicals (Murakami et al., 2003), so by observing neuromasts after exposure to neomycin we could follow hair cell development en masse. First, the strains Gt(macf1- citrine)ct68a/+ and Tg(Parval3b:mCherry-β-actin) were crossed. At 7 dpf, neuromast hair cells of double transgenic animals were ablated with neomycin and live imaging was performed at both 24 and 48 hours after treatment. At 24 hours after exposure, a few hair cells regenerated and matured to have mCherry-β-actin labeled cuticular plates with a ring of Acf7-Citrine encircling each cuticular plate and each fonticulus (Fig. 8F, G); however a few cells within the rosette had Acf7-Citrine near their apical surface, but with no observable mCherry-β-actin in the same regions (Fig. 8F, G). By imaging the same neuromast after another 24 hours (48 hours post exposure), we were able to confirm that these were indeed supporting cells developing into hair cells. The cells that had only expression of Acf7-Citrine matured into hair cells with identifiable hair bundles and cuticular plates (Fig. 8H, I); these cells also retained Acf7-Citrine localization. These findings combined indicate that Acf7 precedes hair-bundle and cuticular-plate formation and is maintained in the mature hair cell near the apical region.

Electron tomography reveals microtubules insert into and are linked to the cuticular plate

Through work with a collaborating lab, the Auer group, we attempted to determine if microtubules of the cell body enter the cuticular plate and whether or not the

F-actin and microtubules are directly linked. Using transmission electron microscopy 2D imaging of ~100 nm thin resin-embedded zebrafish inner ear hair cells we are able to confirm that microtubules do enter the F-actin meshwork of the cuticular plate (Figure

9A, B, D). This novel finding also shows a gap in the F-actin surrounding a microtubule

(Figure 9D) and by using 3D tomographic reconstruction we demonstrate that there is a

~15-20 nm space between this microtubule and the actin meshwork (Figure 9E).

Tomography also shows a number of filamentous connections bridge this gap, providing direct linkage of the microtubule to the cuticular plate matrix (Figure 9E insert). Visual extraction of the features of interest by manual segmentation of the high contrast densities shows a microtubule from the cell body extends past two mitochondria and across the border of the cuticular plate where it is decorated by proteins which link it toF- actin (Fig 9F, G, H). Filaments between the microtubule and the actin meshwork are elucidated, and they may correspond to Acf7. These results provide the first well defined evidence of connections between microtubules and the actin meshwork in the cuticular plate.

Figure 9. Microtubules inserted into the cuticular plate are bridged to the actin meshwork via filamentous linkers. (A, B, D) 2D transmission electron micrographs of high-pressure frozen inner ear hair cells of 6-dpf zebrafish reveal microtubules terminating deep within the cuticular plate. (C, E) Slices ~1-nm thick through a tomographic 3D reconstruction corresponding to the same region depicted as a 2D projection view in panel B and D, respectively. Inset in E is an enlargement of depicted region of interest. (F-H) Manual segmentation and surface rendering of the densities of interest, with microtubules depicted in mint, actin meshwork in pink, and connecting filaments in red, revealing a ~15-20-nm-wide gap between the microtubule and the actin meshwork, which is bridged by filamentous connections. A boundary in blue was drawn to indicate the edge of the actin meshwork. Mitochondria adjacent to the cuticular plate are shown in yellow and green. (G) Surface rendering of the segmented volume overlaid onto a central section through the 3D reconstruction. H contains only the segmented volume from G. The scale bars are 500 nm in A-C, 200 nm in D-H, and C inset.

ACF7 Conclusions

Acf7 links the cuticular plate to the microtubule network and this connection is structurally important

Ultrastructural studies have demonstrated that microtubules insert into the cuticular plate to fasten the plate to the rest of the hair cell’s soma (Jaeger et al., 1994), and our results show there are physical links between the microtubules and the cuticular plate (Figure 9); however, the identity, properties, and regulation of such linker proteins have yet to be described.. Here we demonstrate a spectraplakin in hair cells, Acf7, is positioned between the F-actin of the cuticular plate and the microtubules of the soma

Figure 10. Models of ACF7 function in the mature hair cell. (A) (Top) Image of a hair cell. Cuticular plate (CP) or striated organelle (SO) are represented as a solid green structure or green lines, respectively. Magenta structure is the basal body, which resides in the fonticulus. Green structures on lateral plasma membrane are cross-sections of the circumferential band (CB). Schematics of ACF7 are blue. Model 1, ACF7 links F-actin of the circumferential band to a microtubule. Model 2, ACF7 crosslinks a microtubule that penetrates the cuticular plate with the F-actin of the cuticular plate. Model 3, a microtubule that emanates from the basal body is joined to F-actin of the cuticular plate by ACF7. Model 4, the basal body directly interacts with ACF7 and F-actin of the cuticular plate. Model 5, F-actin of the striated organelle is tethered to a microtubule by ACF7. (B) Schematic of ACF7.

and that this pattern is conserved in both zebrafish and mice. As Acf7 harbors both actin-

and microtubule-binding domains, this provides strong evidence to suggest that Acf7

serves as a major linchpin between microtubules and the F-actin of the cuticular plate

(Figure 10, Model 2). We also hypothesize that Acf7 may be interacting with various

cytoskeletal elements near the apical surface of hair cells in other ways (Figure 10). It

remains possible that Acf7 also links microtubules to the circumferential band (Figure

10, Model 1); this notion is supported by Acf7-actin co-labeling of the apical lateral wall

in cochlear hair cells Figure 7H-K). Given the robust labeling of Acf7 around the

fonticulus, it is likely that the microtubules emanating from the basal body are tethered to

the cuticular plate by Acf7 (Figure 10, Model 3); alternatively, Acf7 may directly

contact the basal body to link the basal body to the cuticular plate (Figure 10, Model 4).

Because the labeling drops below the cuticular plate, it is plausible that microtubules are

linked to the striated organelle actin, which does not label efficiently with phalloidin

(Vranceanu et al., 2012), by Acf7 (Figure 10, Model 5).

In addition to being a cytoskeletal linker, Acf7 may provide additional, structurally important characteristics to hair cells. Numerous spectrin repeats are thought to endow spectraplakins with elasticity, which would allow them to withstand mechanical forces

(Suozzi et al., 2012). In hair cells, Acf7 may help absorb and distribute the forces experienced during mechanotransduction or in overstimulation.

The significance of Acf7 in apical-basal polarity of hair cells

Our data demonstrates that Acf7 predates the hair bundle and cuticular plate at the apical surface of the presumptive hair cell in zebrafish. It stands to reason that establishing apical-basal polarity would occur fairly early on in hair cell morphogenesis and that this would need to occur before apical structures, such as the cuticular plate and bundle, can be built. We posit that Acf7 may be important for establishing or maintaining apical-basal polarity of hair cells. Microtubules are able to specify and maintain the site of cortical actin polarity in various cell types by delivering positional information to the cortex through proteins that associate with the plus-ends of microtubules and induce local actin polymerization (Siegrist and Doe, 2007). Acf7 binds to the growing end of microtubules and in Acf7 null endodermal cells microtubules display irregular growth trajectories and dynamic instability as well as an inability to tether to the cell cortex (Kodama et al., 2003).

In addition, it has been shown that Acf7 is essential for skin stem cell migration in response to wounding because of its ability to organize microtubules along F-actin, resulting in cell polarization (Wu et al., 2011). Therefore, Acf7 may play a role in targeting microtubules to the cortex for the establishment of hair cell apical-basal polarity. Once apical-basal polarity is established, Acf7 persists to provide a foundation for the cuticular plate.

Acf7 is potentially involved in establishing hair cell planar cell polarity

The organ of Corti is a highly polarized epithelium with most hair cells oriented along a single plane, and this polarization is intertwined with the migration of the kinocilium and basal body during hair cell development (Cotanche and Corwin, 1991); however, the mechanisms which guide this process have yet to be described. Our data shows that the pattern of Acf7 localization in hair cells is the same for both zebrafish and mouse vestibular hair cells. However in mouse auditory hair cells, Acf7 is asymmetrically distributed around the cuticular plate, with a higher concentration toward the fonticulus, a pattern that is suggestive of a protein involved in planar cell polarity.

Acf7’s capacity to bind both microtubules and actin, as well as Acf7’s asymmetric distribution in auditory hair cells, suggests this protein’s involvement in planar cell polarity. The microtubule binding capacity of ACF7 is controlled through phosphorylation by glycogen synthase kinases 3b (GSK3b), and GSK3b activity is inhibited by Wnt signaling (Wu et al., 2011). Wnt ligands are secreted proteins with known morphogenic activity. In a different system, ACF7 was shown to complex with

GSK3b, B-catenin, Axin, and APC and to play a role upstream of GSK3b in the Wnt signaling pathway (Chen et al., 2006). In vitro, outer hair cells fail to properly orient their bundles when exposed to Wnt7a or molecules that block Wnt receptor binding (Dabdoub and Kelley, 2005). Further evidence of a link between Wnt signaling and ACF7 is the resemblance in phenotype between Acf7, Wnt-3, as well as Lrp5/6 double-knockout mice, all of which exhibit defects in primitive streak, node, and mesdoderm development and die embryonically (Chen et al., 2006). Therefore, the spatial and temporal regulation of

ACF7 under the control of Wnt signaling may be responsible for positioning of the basal body and kinocilium in hair cells.

In future studies we plan on using genetic techniques to investigate Acf7’s role in the hair cell. The Gt(macf1-citrine)ct68a zebrafish strain reported on in this study also allows for knocking out acf7 in cell-specific lineages. In addition to the elements described previously, the FlipTrap vector also contains heterotypic lox sites as well as mCherry and polyA sequences oriented in the “reverse” direction. Under Cre-mediated recombination, the Citrine and splice donor sequence are excised and the mCherry and polyA signal are flipped into the forward orientation. This leads to a truncation, blocking addition of exons 3’ of the FlipTrap insertion. To knock out Acf7 specifically in hair cells, the Gt(macf1-citrine)ct68a line will be crossed with a transgenic line which expresses

Cre under the control of the parvalbumin 3b promoter. In addition, to knock out Acf7 in mouse hair cells we currently possess an Acf7 floxed line as well as an inducible Cre driver line. The Cre line has an allele which expresses Cre-estrogen receptor fusion protein under the hair cell specific Atoh1 promoter. The Cre fusion protein is activated by tamoxifen treatment. By crossing these two strains we will have both spatial and temporal control over the expression of Acf7 in mice.

From knockout studies in both zebrafish and mice we hope to determine the importance of Acf7 to hair cell development. Our results indicate that Acf7 predates apical structures, therefore we hypothesize that hair bundles and cuticular plates will fail to form in hair cells lacking ACF7 due to improper actin and microtubule organization at the apical surface. In addition, because ACF7 is both asymmetrically distributed in

45 mouse auditory hair cells and is regulated by Wnt signaling, we also hypothesize that planar cell polarity will be disrupted in hair cells lacking Acf7.

Class IX Myosins Results

Myo9a and myo9b are expressed in zebrafish hair cells. Myo9a is expressed in mouse maculae.

Myo9a was initially found to be present in a microarray study on RNA from an individually picked and purified population of mouse hair cells (McDermott, unpublished data). To determine if class IX myosins were also expressed in other vertebrates, Brian

McDermott isolated hair cells from the chicken ear, generated cDNA, and performed massively parallel sequencing and both Myo9A and Myo9B were determined to be expressed by Lana Pollock. Reverse transcription polymerase chain reaction

(RT-PCR) experiments were performed to confirm expression of these genes. We were able to amplify myo9a and myo9b transcripts from zebrafish hair cell cDNA

(Figure 11A, B). Also, we successfully

Figure 11. Myosin IX RT-PCR analysis. Myo9a and myo9b are expressed in zebrafish hair cells (A + B) and Myo9a is expressed in mouse maculae (C). The correct size of amplicons was confirmed by agarose gel electrophoresis for each reaction. (A) Gel from nested PCR for myo9a . Lane one is amplification from whole fish cDNA and lane 2 is amplification from hair cell cDNA. Lanes 3 and 4 are no template controls. (B) Lane 1 is myo9b amplification from zebrafish hair cell cDNA template and lane two is a no template control. In C, lane one is amplification of Myo9a from mouse maculae cDNA and lane two is a no template control.

46 amplified Myo9a transcripts from mouse maculae cDNA (Figure 11C). Therefore, we were able to confirm that myo9a and myo9b are expressed in zebrafish hair cells and

Myo9a is expressed in the vestibular sensory epithelium of mice.

MYO9A localizes to various structures in hair cells.

To determine where MYO9A localizes within hair cells, a polyclonal antibody to

MYO9A (Chieregatti et al., 1998) was used to perform immunolabeling studies in mouse hair cells. The localization pattern of MYO9A varies depending on both the type of hair cell and preparation used. In whole mount vestibular tissue, MYO9A localizes to a structure beneath the cuticular plate as well as the kinocilium (Figure 12A-C). A thick cable of MYO9A labeling can be observed directly under the phalloidin labeled cuticular plate projecting into the soma of the cell (Figure 12A). This unknown structure appears to be made of microtubules as it also colabels with an anti-acetylated tubulin antibody

(Figure 12B). MYO9A can also be seen in the kinocilium of the hair bundle, as the antibody labels the microtubule based kinocilium and not the F-actin based stereocilia

(Figure 12C). A different localization pattern is observed in vestibular hair cells from inner ear cryosections. In these samples MYO9A has a diffuse localization pattern throughout the hair bundle (Figure 12D). In cultured cochlea, MYO9A localizes to the region of the bundle where the kinocilium is expected to reside (Figure 12E), which is similar to the pattern seen in whole mount vestibular tissue. However, cochlear hair cells lack MYO9A labeling underneath the cuticular plate (data not shown). These conflicting results make it difficult to make any determinations on where exactly MYO9A localizes in hair cells. Additionally, its affinity for microtubule based structures is perplexing because class IX myosins lack any putative tubulin interacting domains.

Figure 12. Localization of Myo9a in mouse hair cells. (A–E) Confocal images of mouse hair cells labeled with a rabbit polyclonal antibody to myo9a (green) and Alexa Flour 568 which labels F-actin (red). Whole mount vestibular hair cells (A–C) reveals myo9a localizing to a structure underneath the cuticular plate (A + B) as well as the kinocilium (C). (A) Lateral view of a 3-dimensional reconstruction shows Myo9a labeling a structure projecting from under the cuticular plate into the cell soma. Colabeling with an antibody to acetylated tubulin (blue) shows this structure is made of tubulin (B) and confirms localization to the kinocilium (C). A lateral view of a vestibular hair cell from a cryosection of the inner ears shows myo9a localization to the bundle (D), something not observed in whole mount vestibular tissue. An apical view of cultured mouse cochlea shows the bundles of the three rows of outer hair cells, with myo9a localizing to the region the kinocilium is normally found (E).

Class IX Myosins Conclusions

Here we show that expression of class IX mysosins in hair cells appears to be conserved in vertebrates (Figure 11) which is consistent with an important role in either hair cell development or function. The function of class IX myosins in hair cells is probably related to the unique RhoGAP domain in their tail domain. This is because

RhoGAPs are known to be involved in the organization of the actin cytoskeleton through their regulation of actin nucleators such as Arp2/3 and formin (Jaffe and Hall, 2005).

The ability of class IX myosins to interact with F-actin, through their mysosin head domain, and effect actin polymerization, through their RhoGAP domain, lends to the idea that they may be important regulators essential for building the actin-based structures at the apical surface of hair cells during development. It has been previously shown that

Myo9b null macrophages lack lamellipodia, an actin based structure, and display an inability to migrate in response to a chemotactic gradient. In addition, it was shown that the active form of the actin depolymerization factor cofilin, which is inhibited by

RhoGAPs, was increased in these cells (Hanley et al., 2010). We hypothesize that class

IX myosins may help in the construction of the cuticular plate and stereocilia of hair cells by inducing actin polymerization and/or inhibiting actin depolymerization during development.

Our attempts to determine the localization of MYO9A in hair cells produced varying results depending on the type of hair cell and the preparation used (Figure 12).

In vestibular hair cells, both punctate and diffuse MYO9A labeling was observed in the bundle (Figure 12A, D). Localization of MYO9A in the bundle is not completely unexpected since several myosins, such as myosin VIIa, myosin XVA, and myosin-1c,

49 are located in the hair bundle (Petit and Richardson, 2009); however, the lack of a consistent labeling pattern makes it difficult to confirm MYO9A as a bundle protein.

MYO9A also localized to microtubule cables underneath the cuticular plate (Figure 12A,

B) and the kinocilium (Figure 12C, E), which is also microtubule based. While it has been reported that myosin-1c binds to microtubules in Dictyostelium discoideum during mitosis (Rump et al., 2011), this interaction was attributed to its tail homology (TH) 1 and TH2 domains, which MYO9A lacks. In fact, MYO9A lacks any putative microtubule interacting domains.

In order to resolve the conflicting results obtained from immunolabelling experiments we plan on expressing a fluorescently tagged Myo9a fusion protein in zebrafish hair cells. To do this we have constructed an expression vector containing

Myo9a cDNA in frame with monomeric EGFP under the control of the palvalbumin 3b promoter. This promoter will drive expression of the Myo9a fusion protein specifically in hair cells (McDermott et al., 2010). Currently this vector contains two point mutations which change the amino acids their respective codons code for. So before the vector can be microinjected these mutations need to be fixed. Once the vector is corrected we hope to gain greater insight into where Myo9a localizes in hair cells.

50 plate and somatic microtubules, potentially integrating these two networks. We also demonstrate that ACF7 localization to the apical surface of hair cells predates the cuticular plate, suggesting it plays a major role in morphogenesis. Finally, through electron tomography performed by collaborators, we identify linkages between a microtubule inserting into the cuticular plate and the surrounding F-actin. We hypothesize that ACF7 is a strong candidate to be this molecular linker. These results provide evidence that ACF7 is a key regulator of cytomatricies in the hair cell.

In conclusion, we have determined MYO9A and MYO9B as well as ACF7 are expressed in the hair cells of vertebrates. Because class IX myosins interact with F-actin and ACF7 can bind both actin and microtubules, we hypothesize that these proteins are important for establishment and maintenance of hair cell morphology. Our results show that ACF7 localizes to the apical surface of hair cells where it surrounds, underlies, and is within the cuticular plate. In addition, ACF7 lies between the F-actin based cuticular plate and somatic microtubules, potentially integrating these two networks. We also demonstrate that ACF7 localization to the apical surface of hair cells predates the cuticular plate, suggesting it plays a major role in morphogenesis. Finally, through electron tomography performed by collaborators, we identify linkages between a microtubule inserting into the cuticular plate and the surrounding F-actin. We hypothesize that ACF7 is a strong candidate to be this molecular linker. These results provide evidence that ACF7 is a key regulator of cytomatricies in the hair cell.

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