VISUALIZATION OF THE RIBBON SYNAPSE USING RIBEYE A-MCHERRY FUSION PROTEIN

by

Megan C. West

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

August, 2011

CASE WESTERN RESERVE UNIVERSITY

SCHOOL OF GRADUATE STUDIES

We hereby approve the thesis/dissertation of

Megan C. West

candidate for the Master’s of Science degree *.

(signed) Michael F. Benard (chair of the committee)

Heather Broihier

Brian M. McDermott

Roy E. Ritzmann

(date) June 10, 2011

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

TABLE OF CONTENTS

LIST OF FIGURES …………………………………………………………………………………………………………….4

ABSTRACT………………………………………………………………………………………………………………………5

INTRODUCTION………………………………………………………………………………………………………………6

Chemical and Electrical Synapses………………………………………………………………………..6

Conventional Chemical Presynaptic Active Zones……………………………………………….7

The Ribbon Synapse……………………………………………………………………………………………9

Ribeye protein………………………………………………………………………………………………….12

Hair Cells…………………………………………………………………………………………………………..13

Zebrafish as a Model Organism…………………………………………………………………………15

Zebrafish Inner Ear Development and Lateral Line Development……………………..17

Modes of Investigating the Ribbon Synapse……………………………………………………..19

MATERIALS AND METHODS………………………………………………………………………………………….20

RESULTS……………………………………………………………………………………………………………………….29

DISCUSSION………………………………………………………………………………………………………………….45

REFERENCES…………………………………………………………………………………………………………………51

3

LIST OF FIGURES

FIGURE 1………………………………………………………………………………………………………………………..7

FIGURE 2………………………………………………………………………………………………………………………..8

FIGURE 3………………………………………………………………………………………………………………………10

FIGURE 4………………………………………………………………………………………………………………………14

FIGURE 5………………………………………………………………………………………………………………………17

FIGURE 6………………………………………………………………………………………………………………………18

FIGURE 7………………………………………………………………………………………………………………………24

FIGURE 8………………………………………………………………………………………………………………………26

FIGURE 9………………………………………………………………………………………………………………………30

FIGURE 10…………………………………………………………………………………………………………………….33

FIGURE 11…………………………………………………………………………………………………………………….36

FIGURE 12…………………………………………………………………………………………………………………….37

FIGURE 13…………………………………………………………………………………………………………………….38

FIGURE 14…………………………………………………………………………………………………………………….39

FIGURE 15…………………………………………………………………………………………………………………….43

FIGURE 16…………………………………………………………………………………………………………………….44

TABLE 1………………………………………………………………………………………………………………………..21

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Visualization of the ribbon synapse using Ribeye a-mCherry fusion protein

Abstract

by

MEGAN C. WEST

Specialized presynaptic matrices known as ribbon synapses are found in sensory cells of the visual, auditory, and vestibular systems of vertebrates. Hair cells of the auditory system and the vestibular system rely on ribbon synapses to transmit signals to the brain. Due to the lack of tools to visualize the ribbon synapse, studying the ribbon has been challenging. We designed the construct, Tg(pvalb3b:ribeye a-mCherry), which allows for visualization of ribbon synapses in the hair cells of live or fixed zebrafish. We show that the Ribeye a-mCherry fusion protein expressed from this construct localizes to appropriate areas using immunolabeling and live animal imaging. These results demonstrate that Tg(pvalb3b:ribeye a-mCherry) is an effective tool to label ribbons. We utilize this tool to characterize mutant zebrafish and to investigate ribbon dynamics.

Our results show that Ribeye a-mCherry has the ability to be used in mutant zebrafish and in fluorescent recovery after photobleaching experiments.

5

Introduction

Chemical and Electrical Synapses. The nervous system is a complicated network comprised of an extremely large number of cells. The main signaling cell in the nervous system is known as a neuron. In order for the nervous system to operate as a functional unit, there must be communication between neurons. The synapse is the site of communication between neurons and can be either electrical or chemical. Most synapses in vertebrates are chemical synapses (Kandel and Siegelbaum, 2000). Electrical synapses use an ion current to transmit information from one cell to another; however, chemical synapses use chemical transmitters to convey information. Synapses are comprised of a presynaptic structure and a postsynaptic structure. The morphology of these structures depends on the type of synapse. At electrical synapses, continuity between the presynaptic and postsynaptic cells is established by gap junctions channels found in each cell. In chemical synapses there is no contact between the two cells and the space between them is known as the synaptic cleft (Figure 1). The presynaptic cell of a chemical synapse houses neurotransmitter-filled vesicles. These chemicals are exocytosed into the synaptic cleft and eventually bind to receptors found on the postsynaptic cell.

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Figure 1. Schematic of a chemical synapse. The presynaptic cell is filled with vesicles (blue circles) containing neurotransmitter (black dots). These vesicles dock at the plasma membrane, where they undergo vesicle 'priming'. When an electrical impulse (wavy white line) arrives in the presynaptic cell the vesicles fuse to the active zone plasma membrane and neurotransmitter is released into the synaptic cleft. Receptors on the postsynaptic cell detect the neurotransmitter, and trigger a postsynaptic electrical response. Reprinted with permission from Macmillan Publishers Ltd: Nature, Dobrunz and Garner, 2002. www.nature.com

Conventional Chemical Presynaptic Active Zones. Chemical synapses contain active zones which are areas of the plasma membrane specialized for the release of neurotransmitter. These areas are regulated through voltage gated Ca2+channels which, when open, cause a depolarization of the cell and allows synaptic vesicles to fuse with the presynaptic membrane and release neurotransmitter (Figure 1). The active zone plasma membrane is characterized by an electron-dense meshwork associated with the plasma membrane, the presence of synaptic vesicles, and its alignment opposite of neurotransmitter receptors on the postsynaptic cell. There are two main components of the active zone, the active zone plasma membrane and the cytomatrix at the active zone (CAZ).

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The CAZ is a complex structure which extends ~ 50 nm into the cytoplasm (Gray,

1963; Pfenninger et al., 1972; Phillips et al., 2001). Many proteins found at the CAZ have been identified and can be grouped into six categories: (1) proteins involved in synaptic vesicle fusion, including syntaxin and SNAP-25; (2) cytoskeletal proteins, such as actin, tubulin, and myosin; (3) scaffolding proteins, such as CASK; (4) voltage gated calcium channels; (5) cell adhesion molecules, including neurexins and cadherins and (6)

CAZ-specific proteins known as UNC13/Munc13 proteins, Rab3-interacting molecule

(RIMs), ELKS (ERC/CAST), and Bassoon and Piccolo/aczonin (Fejtova and Gundelfinger,

2006). These molecules work together to form the scaffold of the CAZ as well as organize the synaptic vesicle cycle (Figure 2). The CAZ is believed to have the important functions of positioning the neurotransmitter release site opposite the postsynaptic receptors and supplying the active zone with synaptic vesicles (Zhai et al., 2001).

Figure 2. Molecular organization of the CAZ. This schematic shows several molecules working together to anchor the CAZ and assist in the synaptic vesicle cycling. Synaptic vesicles (SV) are depicted as circles with neurotransmitter (black) present inside. Reprinted with permission from Springer, Schoch and Gundelfinger, 2006.

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Many studies have been conducted to understand the functions of CAZ-specific proteins. The UNC13/Munc13 proteins have been shown to be involved in synaptic vesicle ‘priming’, which prepares the vesicles for fusion with the plasma membrane

(Brose et al., 2000). It has also been suggested that these proteins may function in the regulation of presynaptic short-term plasticity (Rosenmund et al., 2002). RIMs are scaffolding proteins known to interact with many of the proteins found at the CAZ.

Studies have shown that RIMs also function in priming and short-term plasticity as well as long-term plasticity (Calakos et al., 2004). ELKS proteins are believed to have important interactions with CAZ specific proteins. These interactions may be required to localize RIMs to the active zone and a disruption of the interaction between ELKS and

Bassoon results in reduced neurotransmitter release (Ohtsuka et al., 2002; Takao-Rikitsu et al., 2004). These three protein families, UNC13/Munc13,RIMs, and ELKS, are conserved in C. elegans, Drosophila, and vertebrates (Fejtova and Gundelfinger, 2006)

In contrast, Bassoon and Piccolo are not believed to be conserved in C. elegans and

Drosophila but are found in vertebrates; these two proteins are the largest active-zone specific proteins (Fejtova and Gundelfinger, 2006). Piccolo and Bassoon are proposed to be involved in synaptic vesicle clustering (Mukherjee et al., 2010) and Bassoon has been shown to play an important part in the assembly and function of the CAZ (Chen et al.,

2011).

The Ribbon Synapse. Ribbon synapses are specialized CAZs found in sensory cells in the auditory, visual, and vestibular systems. The ribbon synapse is characterized by the appearance of an electron dense structure known as the ribbon or dense body, which is

9 surrounded by a halo of vesicles. These vesicles are tethered to the ribbon through fine filaments and are in close proximity to each other. A portion of vesicles are found in contact with the plasma membrane and are considered ‘docked’ vesicles and the others are considered ‘tethered’ vesicles. Classic ribbon synapses are found only in vertebrates, but similar structures known as T-bars are found in Drosophila. The size and shape of a ribbon synapse can vary based on the stage of development, species of animal, and cell type (Figure 3).

Figure 3. Schematics and electron micrographs of synaptic ribbons and similar structures found in salamander, frog, and fly. A, C, E, Schematic representations of the electron micrographs shown to their right. Green vesicles represent the docked, ribbon-associated pool and yellow vesicles represent the tethered pool. B, Electron micrograph of a salamander rod photoreceptor. A ribbon-like projection is seen surrounded by synaptic vesicles. D, Electron micrograph of a frog saccular hair cell displaying a spherical synaptic body. Vesicles (yellow arrow) are tethered to the electron dense ribbon (blue arrow). F,

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Electron micrograph showing a T-bar shaped “ribbon” in a fly photoreceptor terminal. The T-bar contains a platform (double arrow) and a pedestal (single arrow). Reprinted by permission from Sages Publications, LoGiudice and Matthews, 2009.

As mentioned previously, exocytosis at a conventional chemical synapse is triggered by the influx of Ca2+ through voltage-gated calcium channels. An action potential is necessary to depolarize the cell which results in the opening of these channels and is often considered to have an ‘all or none’ response. In contrast, ribbon synapses release the primary neurotransmitter, glutamate, tonically based on the graded changes of depolarization. Ribbon synapses have been shown to have many more vesicles than conventional synapses (von Gersdorff et al., 1996). Therefore, it is believed that the ribbon provides these areas with a large number of vesicles to sustain the activity of tonic release as well as a high rate of release during sustained depolarization. The exact details of ribbon function are unknown but there are several theories. One theory is the ribbon acts as a conveyer belt making a plethora of vesicles available for rapid release and replenishment (von Gersdorff, 2001). Another theory is that the ribbons allow for multivesicular release (Singer et al., 2004; Parsons and

Sterling, 2003).

Ribbon synapses contain many similar proteins to conventional chemical synapses, including Bassoon, Piccolo, and RIM. Knockout studies of Bassoon have shown free floating ribbons in photoreceptors of mice indicating that Bassoon may have anchoring properties in ribbon synapses (Dick et al., 2003). However, there are some key differences in the molecular composition of the ribbon synapses when compared to

11 conventional synapses. Synapsin I, known to cluster vesicles, at conventional synapses is not found in ribbon synapses. Rabphilins, which are involved in endocytosis, are also not present at ribbon synapses. Ribbon synapses contain L-type calcium channels versus

N-type calcium channels found at conventional synapses. Ribbon synapses also have a unique protein known as Ribeye.

Ribeye protein. Ribeye is a unique and major component of the ribbon synapse comprising approximately 64-69% of the mass of the ribbon (Zenisek et al., 2003;

Zenisek et al., 2004). Ribeye has a novel A-domain and a B-domain which is almost identical to C-terminal binding protein 2 (Ctbp2) (Schmitz et al., 2000). Studies have shown that Ribeye is a splice variant of Ctbp2 (Schmitz et. al, 2000; Piatigorsky, 2001;

Verger et. al, 2006). C-terminal binding proteins are found at both conventional and ribbon synapses. Both domains of Ribeye contain binding sites for the other domain as well as to their own domain. This allows for self-assembly of Ribeye into a large scaffold and could be the building blocks of the ribbon synapse (Magupalli et al., 2008). The B- domain is highly conserved among species, but the A-domain is highly divergent

(Schmitz et al., 2000; Wan et al., 2005; tom Dieck and Brandstätter, 2006). Bassoon has been shown to interact with Ribeye, and it is thought that this interaction may be required to anchor the ribbon at the active zone (Dick et al., 2003; tom Dieck et al.,

2005).

In the case of ribeye in teleosts, a large and diverse group of ray-finned fish, two homologs of the mammalian ribeye gene are found, known as ribeye a and ribeye b

(Wan et al., 2005). Due to a whole-genome duplication that is believed to have occurred

12 in fish there is often expression of two sets of genes (Meyer and Van de Peer, 2005). In situ hybridizations in zebrafish show the expression of ribeye a in the pineal gland, inner ear, and the photoreceptors and bipolar cells of the retina but ribeye b is expressed in only the inner ear and photoreceptors of the retina (Wan et al., 2005). The differential expression pattern of these two genes is not unexpected due to the theory that some genes retain their original function after a whole genome duplication, but others have the ability to take on a new function (Meyer and Van de Peer, 2005). In both, ribeye a and ribeye b, the B-domain is highly conserved, but the A-domain has only at 35% sequence identity (Wan et al., 2005). A smaller form of ribeye a has also been identified, called ribeye a2, and is believed to be a splice variant of ribeye a (Wan et al.,

2005).

Hair cells. The expression of ribeye protein in the inner ear has been shown to be in hair cells (Obholzer et al., 2008). Hair cells, surrounded by nonsensory supporting cells, are found in the inner ear and in the lateral line organ of some aquatic vertebrates. They are highly specialized mechanosensory cells, which are of critical importance for hearing and balance in vertebrates as well as detection of water movement in some aquatic vertebrates. Hair cells have the ability to convert mechanical stimuli into electrical signals which are passed on to the afferent neurons through exocytosed neurotransmitters and eventually to the brain. Many forms of deafness and hearing loss are due to mutated or damaged hair cells.

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Hair cells are characterized by the hair bundle, which extends from the apical surface of the cell. The hair bundle is comprised of numerous stereocilia, which form a staircase-like structure with the tallest stereocilia residing next to the kinocilium (Figure

4A). Each stereocilium has a tapered region and is anchored in the cuticular plate through a rootlet. This tapered region of the stereocilium allows for pivoting of the stereocilia. The hair bundle is the mechanical sensor of the cell and stimulation from sound waves, gravity, or water can cause the stereocilia to deflect towards the kinocilium, which stretches tip links connecting each stereocilia to one another (Figure

4B). The stretching of tip links causes ion channels located in the vicinity to open and an

A B

Figure 4. Schematic diagram of a hair cell and associated stereocilia. A, Schematic of a hair cell showing the hair bundle located at the apical surface of the cell containing stereocilia and a single kilocilium. The ribbon is shown as a dark circular structure located opposite the afferent neuron. B, Schematic of two adjacent stereocilia with a tip link joining the ion channel to the shorter stereocilium. The most recent model proposes the link is stretched by deflection of the stereocilia, and the channel is opened. Image source: Dr. Brian M. McDermott.

14 influx of ions into the cell occurs. The cell is depolarized due to the influx of potassium and calcium ions which, then, causes the ribbon synapse located at the basolateral membrane to release neurotransmitter onto the adjacent afferent neuron.

Zebrafish as a model organism. Zebrafish (Danio rerio) are emerging as a highly advantageous system in research and are particularly beneficial for studies of the hair cell. One significant advantage is the ease of efficient manipulation and observation of developmental processes in vivo. This is possible through the fish’s ability to produce a large number of externally fertilized and transparent eggs, transparent larvae at different developmental stages, and rapid maturation. Although zebrafish have been in use for many decades, there has been a significant increase in the use of zebrafish as a model organism since 1990, providing readily available resources, such as the sequencing of the genome (Lieschke and Currie, 2007). There has also been a substantial number of techniques developed, which can be used to genetically alter zebrafish, including overexpression experiments by microinjection of plasmid DNA, reverse genetics, and knockout experiments using zinc finger nucleases (ZFNs). In zebrafish, antisense oligonucleotides, known as morpholinos, are used to knockdown the level of protein expressed by binding to a specific mRNA sequence and blocking translation. When morpholinos directed towards Ribeye a were injected into embryos, the fish did not react to visual stimuli and displayed defects in photoreceptors and bipolar cells (Wan et al., 2005).

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Another important advantage of zebrafish is that they are a vertebrate organism and have many morphologically similar structures with other vertebrates, including humans. This similarity extends to the ear, which despite the lack of an outer ear and a middle ear, the zebrafish possess an inner ear comprised of three communication chambers, semicircular canals, and sensory patches (Figure 5A, C). These sensory patches contain hair cells and support cells which are associated with sensory neurons

(Figure 5D). Hair cells found in zebrafish are similar in structure and function to those in humans. In addition to the sensory patches found in the ear, zebrafish have neuromasts found in the anterior lateral line and posterior lateral line (Figure 5A). The lateral lines are important structures in aquatic vertebrates which allow for detection of water movement. This ability is vital for schooling, feeding, and survival (Dambly-Chaudière,

2003). Neuromasts on the lateral lines contain six different types of cells including hair cells, support cells, mantle cells, and sensory neurons (Figures 5B). Neuromasts are positioned along the body with a cupula that is directly exposed to the surrounding environment. The direct exposure of neuromasts to the environment facilitates manipulation of the hair cells. Defects in balance of zebrafish are easily detected through swimming behaviors (Nicolson et al., 1998). Larval fish can be tested for hearing function through use of the startle response (Nicolson, 2005; Colwill and Creton, 2011).

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Figure 5. A schematic diagram of the zebrafish acousticolateralis system. A, A zebrafish showing the ear and lateral line organ. The ear is located posterior to the eye. The two lateral lines are found surrounding the eye and ear as well as along the length of the body. The neuromasts of the lateral line systems are displayed as orange dots. The anterior lateral line is comprised of neuromasts found on the head, but the posterior lateral line begins posterior to the ear and continues down the length of the body. B, The organization of the neuromast organ. Hair cells are shown in orange. Support cells are in blue. Mantel cells are green. The cupula which is exposed to the exterior environment is pink and the neurons are purple. C, The left ear of a four-day-old zebrafish larva depicts the sensory maculae (light blue), cristae of the semicircular canals (green), and otoliths (gray). A red box delimits the anterior macula. D, The organization of the anterior macula. Hair cells are shown in orange while support cells are in blue. Image and figure legend source: Dr. Brian M. McDermott.

Zebrafish Inner Ear Development and Lateral Line Development. The formation of the zebrafish ear is relatively rapid with the first visible sign of its formation at 16 hours post fertilization (hpf) by the thickening of the ectoderm forming the otic placode (Haddon and Lewis, 1996). Shortly after this, the otic placode begins to transform into a ball shape and the ear lumen appears. The otic vesicle grows by increasing the size of its lumen and the formation of hair cells, support cells, and sensory neurons begins at around 20 hpf (Haddon and Lewis, 1996; Whitfield et al., 2002; Nicolson, 2005).

Neuroblasts delaminate from the otic vesicle and accumulate under the otic epithelium to form the first-order neurons of the auditory/vestibular (VIIIth) nerves (Haddon and

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Lewis, 1996; Nicolson, 2005). Cells that do not delaminate remain part of the otic vesicle to form the hair cells and support cells (Nicolson, 2005). The hair cells then come in contact with the otoliths and continue to increase their size and number. These first two sensory patches develop into the anterior and posterior maculae.

Beginning at approximately 48 hpf and continuing to 72 hpf is the development of the three cristae associated with the semicircular canals. The ear continues to develop and at about 5 days post fertilization (dpf) all major structures are present with approximately 20 hair cells per crista and 60 hair cell per macula (Haddon and Lewis,

1996) (Figure 6). Hair cells are continuously being generated during development of the fish. Although lagena and macula neglecta, which may have auditory and vestibular functions, are not developed until after the larval stage, it has been shown that zebrafish at 5 dpf have the same response to stimulus thresholds and frequency bandwidth as adult fish (Zeddies and Fay, 2005).

Figure 6. Image of the developing zebrafish inner ear. Images of ears from zebrafish at 3, 5, 10, 15, 17, 20 days postfertilization (dpf) and adult. aa, rostral ampulla; ac, rostral canal; cc, common crus; D, dorsal; ha,

18 horizontal ampulla; hc; horizontal canal; la, lagena; P, posterior; pc, posterior canal; s, saccule; u, utricle. Scale bar = 500 μm. Reprinted by permission from John Wiley and Sons, Whitfield et al., 2002.

Posterior lateral line development occurs through migrating primordiums which originate from the cephalic placode found just posterior to the otic placode (Metcalfe et al., 1985). Development of the posterior lateral line nerve also takes place at the cephalic placode and is dependent on the activity of neurogenin 1 (Andermann et al.,

2002). The first primordium contains approximately 120 cells and begins to migrate posteriorly at around 20 hpf (Kimmel et al., 1995). As the primordium moves posteriorly along the body of the fish, it deposits interneuromasts and is soon followed by a second primordium which deposits proneuromasts. These cells develop gradually from head to tail to form the neuromasts. There are approximately 8 neuromasts found in the posterior lateral line of a 5 dpf zebrafish. Adult zebrafish contain many more neuromasts and studies have shown that another primordium is present after embryonic development resulting in the increase of neuromasts along the posterior lateral line in adults (Ledent, 2002; Sapède et al., 2002).

Modes of Investigating the Ribbon Synapse. As mentioned previously, knockout experiments have been performed with Bassoon and this study revealed the importance of the CAZ-specific proteins in ribbon synapse function (Dick et al., 2003). Since ribeye proteins are a major component of the ribbon synapses and they have has been shown to localize to the ribbons with high specificity (Schmitz et al., 2000; Obholzer et al.,

2008). Several studies have used ribeye to understand the development and function of

19 the ribbon synapse (Wan et al., 2005; Zenisek et al., 2004; Magupalli et al., 2008; Sheets et al., 2011).

Several techniques have been used to visualize the ribbon synapse including the use of Nomarski optics (Flock and Jorgenson, 1974), immunolabeling directed towards

Ribeye b (Obholzer etal. 2008), electron microscopy (Lenzi et al., 2002), and the introduction of a fluorescent peptide with a binding affinity to RIBEYE into cells by patch pipette (Zenisek et al., 2003). These studies have shown important findings but, unfortunately, there are several shortcomings with these methods. The shortcomings include the use of a fixed tissue instead of a living specimen, manipulation of the cells which has time constraints and can cause damage, and the use of organisms that are not genetically tractable. Our aim in the following studies was to develop a way to examine the ribbon synapse in vivo, without invasive manipulations, in a genetically tractable system. We developed a method to create transgenic zebrafish that express fluorescently tagged Ribeye a protein, which can be used to visualize the locations of the synaptic ribbons in the ear and lateral line hair cells of live or fixed tissues. Using our method, we were able to examine ribbon movements in wild-type and mutant zebrafish as well as conduct fluorescent recovery after photobleaching (FRAP) experiments to look at the dynamics of Ribeye a in vivo.

Material and Methods

Fish. One wild-type line, Tübingen, two stable transgenic zebrafish lines, Ppv3b-4

(McDermott et al., 2010), HGn39D (Faucherre et al., 2009), and one mutant line for neurogenin-1 (ngn1) (Lòpez-Schier and Hudspeth, 2005) were used in these

20 experiments. They were maintained and bred at 28°C by standard procedures (Nüsslein-

Volhard and Dahm, 2002) and kept with the approval of the Case Western Reserve

University Institutional Animal Care and Use Committee.

Molecular biology. All restriction endonucleases used in these experiments were obtained from New England Biolabs (NEB). In reverse transcription–polymerase chain reaction (RT-PCR) procedures, randomly primed cDNA (Superscript III; Invitrogen) was produced from lagenar RNA of adult zebrafish (McDermott et al., 2007). Polymerase chain reaction (PCR) experiments were performed (Ex Taq DNA Polymerase; Takara Bio or Pfu DNA Polymerase; Stratagene) with the primer pairs listed in Table 1. Ligations were done with T4 ligase (Promega).

Table 1. Oligonucleotides Primer Name Primer Sequence (5’-3’)

5’ MCS-pBSIISK+ CTAGTTTGGATCCTTAATTAAGTTTAAACAGGCGCGCCTGCGGC CGCACGCGTCTTAAGAGATCTCCGC

3’ MCS-pBSIISK+ GGAGATCTCTTAAGACGCGTGCGGCCGCAGGCGCGCCTGTTTAA ACTTAATTAAGGATCCAAA

Bam Pv3b 1 AAGGATCCTTTGATTTCTTCATTTAAG

3’_no_G_Pv3b TTGGATCCACCCGGGATATTCAAACTGTTGAGAGAATAAAACA

5-X-RibeyeA ATCCCGGGACCATGTTGATCTCCAGTAAGCAGTTG

3-P-RibeyeA ATTAATTAAGGTATACATTTTGTCTTGCAGGCCG

RPG 8 GGACTGGGCATGGGTGACATTG

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RPG 4 CTT TACCTGCAGTTCCTCAGCAAT

5-p-mCh GGTTAATTAAAGGCATGGTGAGCAAGGGCGAGGAG

3-a-mCh TTGGCGCGCCTTACTTGTACAGCTCGTCCATGC

Construction of the Ribeye a-mCherry fusion protein expression vector. The first step in making the Ribeye a-mCherry expression plasmid, Tg(pvalb3b:ribeye a-mCherry), was to create pMT/SV/PV. A multiple cloning site (MCS) was generated by annealing two oligonucleotides, 5’ MCSpBSIISK+ and 3’ MCS-pBSIISK+ (Sambrook and Russell, 2001)

(Figure 7A). The product was then ligated into pBluescript II SK(+) (Stratagene), which had been digested with SpeI and SacII. The resulting construct was digested with NotI and AflII to insert a SV40 polyadenylation addition sequence, which had been excised from pEGFP-1 (Clontech) using NotI and AflII (Figure 7B). The SV40 polyadenylation addition sequence is found in many vectors to provide stability and allow proper cleavage and polyadenylation of RNA (Sheets et al., 1990). The multiple cloning site with the polyadenylation addition sequence was removed with SpeI and BglII; this digested product was then ligated with the pminiTol2/MCS vector (Balciunas et al.,

2006), which had been digested with the same enzymes, to create pMT/SV (Figure 7C).

The miniTol2 vector was chosen because it is a part of a high cargo-capacity transposon system which has been shown to increase efficiency of transgenesis in zebrafish

(Balciunas et al., 2006). The zebrafish parvalbumin 3b promoter, which drives expression in hair cells, was amplified from the Ppv3b-4 vector (McDermott et al., 2010) by a PCR reaction that introduced BamHI sites onto the product termini using primers Bam Pv3b

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1 and 3’_no_G_Pv3b. The promoter was subcloned into pCRII-TOPO (Invitrogen), and the resulting plasmid was digested with BamHI. The fragment containing the promoter was then ligated into BamHI-digested pMT/SV, resulting in pMT/SV/PV (Figure 7D).

After creating pMT/SV/PV, mCherry cDNA (Shaner et al., 2005) was inserted. We determined that mCherry would be best suited for our needs. It is a monomeric protein with high photostability and is suited to complement other fluorescent proteins such as mKO and mPlum, which have potential for use in other developmental studies (Shaner et al., 2005). Another aspect to consider was that many stable transgenic zebrafish lines we would like to use in future experiments have expression with green fluorescent protein (GFP). Considering the differences in excitations and emissions, using GFP and mCherry together would allow for effective co-labeling during imaging. PCR was used to add a PacI restriction site to one terminus of the mCherry cDNA, and an AscI site to the other, using the primers 5-p-mCh and 3-a-mCh; this product was inserted into pCR-

Blunt II-TOPO (Invitrogen). The resulting vector was digested with AscI and PacI to extract the modified mCherry cDNA; this cDNA was ligated into AscI- and PacI-digested pMT/SV/PV to form pMT/mCh (Figure 7E).

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Figure 7. Schematic of the construction of pMT/SV/PV and MT/mCh vectors. A, Two oligonucleotides were annealed together to create a multiple cloning site (MCS). The MCS was then inserted into pBSIISK+. B, The SV40 poly A addition sequence was isolated from pEGFP-1 and ligated into the pBSIISK+ containing the MCS. C, The MCS and SV40 poly A addition sequence was removed from pBSIISK+ and inserted into the miniTol2 vector. D, The parvalbumin 3b promoter (Pv3b) was isolated from Ppv3b-4 and inserted into the miniTol2 vector containing the MCS and SV40 poly A addition sequence to create pMT/SV/PV. E, mCherry was removed from pRSET-B and ligated into pMT/SV/PV. The resulting construct is known as MT/mCh. MCS (blue), SV40 poly A addition sequence (yellow), parvalbumin 3b promoter (orange), mCherry (red).

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The final step in creating the Ribeye a-mCherry fusion protein was to insert

Ribeye a. Primers were made to isolate smaller pieces of ribeye a cDNA (GenBank number: AY878349), which were ligated together to form the full length cDNA coding region. A total of ten primers were made surrounding five restriction enzymes sites, which could be used to piece the ribeye a cDNA a together (Figure 8A). The smallest pieces were attempted first (i.e. primer 1 with primer 10), and then longer pieces were attempted (i.e. primer 1 with primer 7). We wanted to obtain the longest pieces possible to decrease the chances of sequence mutations, induced by PCR, when forming full-length ribeye a cDNA.

We were able to obtain two pieces of ribeye a cDNA using primers 1 and 8 together or primers 4 and 6 together. Once we established which primers could be used to make full-length ribeye a cDNA, we had to make new primers toward the 5’ and 3’ ends of the cDNA in order to add the necessary restriction enzyme sites to insert ribeye a cDNA into pMT/SV/PV. The cDNA fragments X8 RP1 or 4P 3.3 were amplified from lagenar cDNA by PCR using the primer pairs 5 X-RibeyeA and RPG 8 or 3-P-RibeyeA and

RPG 4, respectively. The X8 RP1 and 4P 3.3 amplicons were ligated into pCR-Blunt II-

TOPO (Invitrogen) and pCRII (Invitrogen), respectively. Both of the resulting plasmids were digested with HindIII, and the X8 RP1 fragment was ligated into the pCRII vector that contained 4P 3.3. The X8 RP1-4P 3.3 fragment, which contained the ribeye a cDNA, was removed from pCRII using XmaI and PacI. This fragment was ligated into XmaI- and

PacI-digested pMT/mCh. The resulting construct encodes the fusion protein Ribeye a- mCherry (Figure 8B).

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Figure 8. Schematic of the construction of full length Ribeye a and Tg(pvalb3b:ribeye a-mCherry). A, Ten primers were made to Ribeye a (black lines with correlating numbers) surrounding restriction enzyme sites (purple lines). Primer pairs were used to amplify the longest pieces of Ribeye a possible. The pieces were ligated together and inserted into pCRII. B, Ribeye a (green, RB) was isolated from pCRII and inserted into the MT/mCh vector to form Tg(pvalb3b:ribeye a-mCherry)(RB/mCh). SV40 poly A addition sequence (yellow), parvalbumin 3b promoter (orange), mCherry (red).

Production of transgenic zebrafish that express mCherry or Ribeye a-mCherry in hair cells and skin cells. Tg(pvalb3b:ribeye a-mCherry) or pMT/mCh DNA at 250 ng/µl and

Tol2 transposase RNA at 25 ng/µl (Balciunas et al., 2006) in 0.1 M KCl were injected into embryos at the one-cell stage. Each of the wild-type, Ppv3b-4, HGn39D, and ngn-1 lines were injected with Tg(pvalb3b:ribeye a-mCherry) and only the wild-type line was injected with pMT/mCh. To visually monitor injections, phenol red was added to each injection solution to make the final concentration of the tracer 0.05%.

Immunolabeling. Zebrafish injected with Tg(pvalb3b:ribeye a-mCherry) DNA, which exhibited mosaic transgene expression in somatic cells, were collected at 5 and 28 dpf and fixed using 4% paraformaldehyde in phosphate-buffered saline (PBS) at 4°C overnight. The larvae were rinsed with PBS, permeabilized in 3% Triton X-100 in PBS

26 overnight at room temperature, bathed in blocking solution (5% goat serum in PBS) for four to six hours at room temperature, and labeled overnight at 4°C with Ribeye b antiserum (Obholzer et al., 2008) diluted 1:100 in blocking solution. Larvae were washed at room temperature with blocking solution five times over a period of six hours. The secondary antibody (Cascade Blue goat anti-rabbit IgG (H+L); Invitrogen) was diluted

1:200 in blocking solution and incubated overnight with larvae at 4°C. Next, the larvae were rinsed twice in blocking solution, for a period of one hour for each wash, and then stored in mounting medium (VectaShield; Vector Laboratory).

Confocal Imaging and fluorescent recovery after photobleaching (FRAP). For live animal imaging, larvae expressing Tg(pvalb3b:ribeye a-mCherry) or pMT/mCh at 4, 5 and/or 6 dpf were anesthetized with 610 μM solution of 3-aminobenzoic acid ethyl ester methanesulfonate (also known as tricaine) (Sigma) and then mounted in low- melting-point agarose (Sigma) (López-Schier and Hudspeth, 2005). Larvae were imaged using an inverted microscope (DM IRE2; Leica), with a 20× or a 40× objective lens, and a confocal laser scanner (TCS SP2; Leica) with Leica confocal software (LCS). Ribeye- mCherry and GFP were excited using the 594 nm of the Helium/Neon (He/Ne) laser at

49% laser intensity and 488 nm of an argon laser at 8% laser intensity, respectively. All data collected was at a resolution of 512 x 512 pixels at 12 bits/pixel.

Photobleaching was performed using the FRAP application in the Leica confocal software (LCS). Two regions of interest surrounding two separate areas of the cell were defined. One region of interest was photobleached for 20 frames with the 594 nm of

27 the HeNe laser at 65% and the 546 nm of the HeNe laser at 65%. Time-lapse recording images were taken manually to adjust for focal drift at the specific time intervals and averaged four times.

4-Di-2-ASP labeling of hair cells. To stain larval fish with 4-Di-2-ASP (Invitrogen), a 12 well culture dish was used. 1 well was filled with staining solution (3.2 ml fish water,

800 µl of 1mM 4-Di-2-ASP solution, 4 µl of 10% BSA solution) and 6 wells were filled with wash solution (48 ml fish water, 2 ml tricaine, 50µl of 10% BSA solution). The fish were placed in netwells and transferred to the staining solutions for 10 minutes. After staining, the fish in the netwells were moved through the 6 wash solutions for 4-6 minutes in each well. The labeled fish were then placed in regular fish water and imaged under a fluorescent stereoscope (Leica).

Genotyping. For genotyping possible adult ngn-1 mutants, fish were anesthetized using

610 µM solution of tricaine and a small portion of the caudal fin was clipped. This portion of the fin was placed in 50 µl lysis buffer (50mM Tris (pH 8.3), 100mM NaCl,

0.4% SDS, 5mM EDTA, 100 µg/ml proteinase K) and incubated at 55°C overnight. The following day, the fin clipping in lysis buffer was vortexed and diluted 1:10 in water. The solution was then placed at 95°C for 5 minutes to heat inactivate the proteinase K. 1 µl of the solution was used for PCR using Taq polymerase (NEB) and the primers ngn primer 1 (5’-GCACAACGTTAGGTATTCACTGTTTGC-3’) and ngn primer 2 (5’-

GCTAGCTTGCCAAACCTACAGGT-3’). The PCR products were run on 1% agarose gels.

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To confirm the larval fish used in the experiment were homozygous for the ngn-1 mutation, the fish were euthanized after imaging using ice and placed in lysis buffer overnight at 55°C. The following day, the fish and lysis buffer were vortexed and diluted

1/10 in water. The solution was then placed at 95°C for 5 minutes to heat inactivate the proteinase K. 1 µl of the solution was used for PCR using Taq polymerase (NEB) and the primers het/homo ngn primer 1 (5’-GCACAACGTTAGGTATTCACTGTTTGC-3’), het/homo primer 2 (5’-CCGTCATGAGAGCTGGTTAACTG-3’), and het/homo viral seq primer (5’-

GTTCCTTGGGAGGGTCTCCTC-3’). The PCR products were then run on a 2.5% agarose gel.

Results

Creation of the Ribeye a-mCherry fusion protein and the pMT/mCh vector. We have designed and implemented a method that enables the visualization of synaptic ribbons of hair cells in live or in fixed zebrafish for developmental studies. We set out to generate the DNA construct Tg(pvalb3b:ribeye a-mCherry) to direct the expression of the fusion protein Ribeye a-mCherry in hair cells. This vector contains the parvalbumin

3b promoter to permit expression in hair cells (McDermott et al., 2010) within the miniTol2 transposon system (Balciunas et al., 2006). RT-PCR was used to amplify the coding region of the ribeye a cDNA using RNA isolated from adult zebrafish maculae.

We chose Ribeye a based on its expression pattern in the inner ear (Wan et al., 2005).

The ribeye a cDNA, lacking its stop codon, was ligated to mCherry cDNA to encode

Ribeye a fused, at its C-terminus, to the fluorescent protein tag. We anticipated that

29 generating a Ribeye a-mCherry fusion protein would be an effective ribbon-labeling tool.

During construction of the Ribeye a-mCherry fusion protein, we generated a vector called pMT/mCh. We anticipated that the pMT/mCh vector should be functional and produce red fluorescent hair cells when injected into zebrafish embryos. This construct was injected into wild-type zebrafish embryos and screened at 4 dpf under a fluorescent stereoscope. Somatic transgenic zebrafish have mosaic expression of the mCherry protein using the parvalbumin 3b promoter and display red fluorescence throughout the whole hair cell (Figure 9A, B, C). Expression is seen in both the lateral line hair cells (Figure 9D) and hair cells of the anterior macula (Figure 9E). This shows that the vector, known as pMT/mCh, is capable of driving red fluorescence expression in hair cells and that the mCherry protein has been inserted in frame. This construct should also serve as a useful tool to investigate hair cell development.

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Figure 9. A 4 dpf zebrafish expressing MT/mCh in hair cells of the ear and the anterior lateral line under a 20x objective. A, A bright field image of the zebrafish head showing the eye (black circle) and otic vesicle; otoliths are seen as two white circles located posterior to the eye. B, The expression of mCherry in the anterior macula and anterior lateral line from pMT/mCh. C, An overlay of bright field and fluorescent images shows that expression is occurring in the otic vesicle and the lateral line. Arrowhead indicates expression in the neuromast of the anterior lateral line. Arrow points out expression in the anterior macula. D, A magnified image of the neuromast identified with the arrowhead from C. One transgenic hair cell, with a kinocilium clearly visible, is present in the foreground with two other MT/mCh expressing hair cells out of focus in the background. E, A magnified image of the anterior macula identified with the arrow from C. Several transgenic hair cells are present in close proximity to the otolith. Scale bar for A, B, and C is 150 µm, for D is 17.87 µm, and for E is 23.29 µm.

Generation of transgenic fish that express Ribeye a-mCherry. We expected

Tg(pvalb3b:ribeye a-mCherry) construct to allow for labeling of red fluorescent puntca, presumably ribbons, at the basal lateral region of the hair cell, opposite the afferent nerve endings (Figure 10A). The Tg(pvalb3b:ribeye a-mCherry) construct was injected into embryos of the Ppv3b-4 transgenic line, which stably expresses GFP in hair cells

(McDermott et al., 2010); this transgenic background was selected because it allows for the delimitation of hair-cell boundaries during imaging. Hair cells of 5- and 28-dpf larvae that expressed Ribeye a-mCherry were imaged using confocal laser-scanning microscopy. These doubly transgenic fish exhibited a normal startle response and typical swimming behaviors, indicating that there was no gross disruption of either hearing or vestibular function as a result of fusion protein expression (data not shown). In maculae

(Figure 10B), cristae (data not shown), or lateral line organs (Figure 10D), the hair cells that expressed this fusion protein showed robustly labeled puncta in close proximity to the basolateral membranes. A similar pattern has been observed in hair cells labeled with antiserum raised against Ribeye b (Obholzer et al., 2008). When doubly transgenic zebrafish expressing GFP and Ribeye a-mCherry in neuromast hair cells were labeled

31 with Ribeye b antiserum, the mCherry signal overlapped with the antibody-associated fluorescence (Figure 10E, F). In hair cells that expressed Ribeye a-mCherry, all ribbons recognized by the Ribeye b antiserum were also labeled with the fusion protein (number of ribbons, N = 77). Tg(pvalb3b:ribeye a-mCherry) was also injected into wild-type embryos and a similar result was obtained (data not shown). This suggests that the fusion protein is an effective tool for identifying the subcellular locations of the ribbons.

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Figure 10. Labeling synaptic ribbons in hair cells with Ribeye a-mCherry fusion protein. A, A schematic of the construct Tg(pvalb3b:ribeye a-mCherry) that was developed to express Ribeye a-mCherry in hair cells for the labeling of synaptic ribbons is shown. Flanking the promoter and cDNA are segments of the miniTol2 transposon (purple). A graphic of a hair cell that expresses Ribeye a-mCherry to label ribbons (red), which reside close to regions of plasma membrane that are contacted by afferent nerve endings (green), is displayed. B, A hair cell from an anterior macula of a zebrafish that was fixed at 5 dpf is shown. This hair cell expresses Ribeye a-mCherry fusion protein (red) and GFP (green), each under control of the parvalbumin 3b promoter (McDermott et al., 2010). GFP allows demarcation of the cell’s boundaries.

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Puncta that contain the Ribeye a-mCherry fusion protein are in close proximity to the basolateral surface. C, In a live larva, ribbons of a neuromast are labeled with Ribeye a-mCherry. An HGn39D doubly transgenic zebrafish at 5 dpf, expressing GFP in afferent neurons of the posterior lateral line and Ribeye a- mCherry fusion protein in associated hair cells, reveals that most of the labeled ribbons (red) are proximal to the nerve endings (green). D, Positions of Ribeye a-mCherry-labeled ribbons (red) within a live hair cell (green) of the posterior lateral line in a Ppv3b-4 transgenic zebrafish at 5 dpf are displayed. All labeled ribbons are juxtaposed to the basolateral surface. Arrowhead shows the positions of two ribbons, in a single hair cell, that reside in close proximity to each other. E and F, Images reveal overlapping fluorescent signals from Ribeye a-mCherry and those produced by immunolabeling with Ribeye b antiserum at the basolateral surfaces of lateral-line hair cells in zebrafish at 28 (E) and 5 dpf (F). Doubly transgenic zebrafish expressing GFP (green) and Ribeye a-mCherry (red) in neuromast hair cells were labeled with antiserum raised against Ribeye b (cyan). When the three colors overlap, they appear as off-white. In F, the cell on the left does not express the transgene, but its ribbon is immunolabeled. G, Ectopic expression of Ribeye a-mCherry in a doubly transgenic peridermal cell is shown. This cell is from a larva that was labeled at 5 dpf with Ribeye b antiserum (cyan), and it contains GFP (green) and Ribeye a-mCherry (red). Ribeye a- mCherry forms fluorescent puncta that are labeled with Ribeye b antiserum. Insets show enlarged images of immunolabeled structures that contain the fusion protein. All scale bars are 2 µm. Asterisks indicate the positions of the apical surfaces of the hair cells.

To determine whether the Ribeye a-mCherry-labeled ribbons were in close proximity to afferent nerve endings, we generated doubly transgenic zebrafish that mosaically express the fusion protein transgene in hair cells and consistently express

GFP in afferent neurons of the posterior lateral line. For this study, we used the HGn39D transgenic line (Faucherre et al., 2009). Afferent neurons carry information from hair cells towards the central nervous system, and their nerve endings contact regions of the plasma membranes of hair cells that are associated with synaptic ribbons. Live imaging using confocal laser-scanning microscopy of doubly transgenic animals revealed that the majority of ribbons were proximal to afferent nerve endings (Figure 10C). This finding further confirms that the Ribeye a-mCherry fusion protein labels ribbons, indicating that it will be an effective tool for characterizing synapse development.

Rat RIBEYE aggregates to form discrete sphere-like structures when it is expressed in COS-7 cells or in cells of the R28 retinal precursor cell line (Magupalli et al.,

34

2008). Because the version of the parvalbumin 3b promoter that we used drives expression in skin cells in addition to hair cells, we looked for sphere-like formations in the periderm of zebrafish that expressed the fusion protein using the Ppv3b-4 transgenic background. Skin cells that expressed Ribeye a-mCherry displayed fluorescent sphere-like structures (Figure 10G) that resembled those observed in cultured cells expressing tagged RIBEYE (Magupalli et al., 2008). This demonstrates that the zebrafish and the rat orthologous proteins form morphologically similar assemblages. Indirect immunofluorescence demonstrated that the Ribeye b antiserum recognized an epitope in peridermal cells that expressed Ribeye a-mCherry (Figure 10G), and that the antibody-associated signal overlapped with the emissions of the fusion protein. No fluorescent puncta were observed in peridermal cells of Ppv3b-4 transgenics, which did not express Ribeye a-mCherry, upon labeling with Ribeye b antiserum (data not shown). These results indicate that this antiserum recognizes both

Ribeye a and Ribeye b. This finding is not wholly unexpected because of the high amino acid sequence similarity of these proteins.

A small percentage of fluorescent structures (4.4% of fluorescent structures, N =

180) were found as massive aggregates (Figure 11); these formations may be attributed to excessive Ribeye a-mCherry fusion protein expression. This occasionally occurs in animals where transgene distribution is mosaic among cells, and the amount of transgenic protein can vary from cell to cell. When the massive aggregates were exposed to Ribeye b antiserum, they were also effectively immunolabeled (data not shown).

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Figure 11. A hair cell containing massive aggregates of Ribeye a-mCherry fusion protein. This cell is from the posterior macula of a zebrafish that was fixed at 5 dpf. The doubly transgenic hair cell expresses Ribeye a-mCherry fusion protein (red) and GFP (green). The scale bar is 2 m.

Movement of ribbons in hair cells and the characterization of ngn-1 mutant zebrafish.

When the hair cells of the lateral line expressing the Ribeye a-mCherry fusion protein were imaged under a confocal microscope, some ribbons appear to show movement. In an attempt to obtain a better understanding of the occurrence and the rate of the movement, we performed time-lapse microscopy on Ppv3b-4 transgenics expressing the

Ribeye a-mCherry fusion protein (Figure 12). Most ribbons remain stable, but approximately 5% of the ribbons have the ability to move at a rapid rate of 0.37 µm per second. This phenomenon has also been shown previously in axolotls, but the purpose of these movements remains unclear (Flock and Jorgenson, 2006). It should be noted that the observed ribbons in zebrafish appear to move not only along the plasma membrane but into the interior of the cell as well. There are three potential models of synaptic ribbon formation and the location of formation could attribute to the movement we have observed in the lateral line. We propose three models of ribbon formation (Figure 13). The first model predicts the ribbon complex is formed in the

36 cytosol. Upon contact of the afferent nerve to the hair cell, the complex is recruited to the basolateral membrane. The second model indicates that the ribbon complex is formed at the basolateral surface after contact has been established with the afferent nerve. This model is unlikely since the presence of ribbons is found in hair cells absent of afferent nerves (Flock and Jorgenson, 1974). The third model postulates that the ribbon complex is first established at the basolateral membrane, and then the afferent nerves contact the hair cell plasma membrane in the appropriate areas juxtaposed to the ribbon synapse. Based on the observed movements of the ribbons, we can speculate that the ribbons are synthesized in the cytosol and are mobile until associated indirectly with an afferent nerve ending.

Figure 12. Time-lapse confocal laser-scanning microscopy was used to visualize ribbon movement in a neuromast in the posterior lateral line of a 4 dpf doubly transgenic zebrafish. GFP (green) is expressed in the hair cells while Ribeye a-mCherry (red) is seen in the right hair cell. 7 images were taken every 30.8 seconds. They are ordered from left to right. Pairs of labeled ribbons (yellow arrow), very close together and very near the plasma membrane, remained stationary throughout imaging. In transition from frame one to two, either one ribbon (yellow arrowhead) divided, or another ribbon entered the optical slice, to result in two ribbons moving fast within the cell. Size bar in image 1 is 6.6 m.

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Figure 13. Three models for the development of the synaptic ribbon. A, Model one, the afferent nerve ending contacts the hair cell and then the synaptic ribbon, which was preformed in the cytosol, is recruited to the active zone plasma membrane. B, Model two, the afferent fiber contacts the hair cells at the basolateral surface and this stimulates the formation of the synaptic ribbon at the plasma membrane. C, Model three, the ribbons are positioned at the active zone prior to afferent contact and, hereafter, the nerve endings identify the active zone. Image source: Dr. Brian M. McDermott.

In an effort to show a use of Ribeye a-mCherry fusion protein and investigate our speculations, we injected ngn-1 mutant zebrafish, which lack the afferent nerves of the posterior lateral line, with Ribeye a-mCherry and observed the location and movement of the ribbons. We anticipated Ribeye a-mCherry could be used for characterization of mutant zebrafish and that without contact from the afferent nerve, these ribbon

38 synapses would not be stable along the basolateral membrane of neuromast hair cells and show robust movement. Mutant ngn-1 fish were obtained from the Zebrafish

International Resource Center (ZIRC). These fish have been shown to lack innervation of the hair cells of the posterior lateral line by afferent neurons. They also have an increase in the number of neuromasts located on the posterior lateral line (Lòpez-Schier and

Hudspeth, 2005). Before beginning the experiment, it was necessary to identify fish that were heterozygous for the ngn-1 mutation before breeding to permit production of homozygous mutant offspring. We performed genotyping on the adult fish through fin clippings and PCRs with primers designed to distinguish between the wild-type copy and the mutated version of the gene (Figure 14A). Heterozygous fish would show one band at 355 base pairs, and the wild-type fish would have no bands present.

Figure 14. Agarose gel images of genotyping results for ngn-1 mutants. A, PCR results of fin clips of adult zebrafish. Primers were used to distinguish between fish heterozygous for the ngn-1 mutation and wild- type fish. Heterozygous fish would produce a 355-base pair band on the gel while no band would be seen with the wild-type fish. Lanes 1, 3, and 5 are from heterozygous fish while lanes 2 and 4 are from wild- type fish. B, Genotyping of larval fish to identify wild-type, heterozygous, and homozygous fish using a different set of primers. Lanes 1, 2, and 3 were obtained using imaged zebrafish, which lacked a response

39 to touch and expressed GFP and Ribeye a-mCherry. Lanes 4 and 5 were from fish that did not respond to touch and did not express GFP and Ribeye a-mCherry. Lane 6 is from a wild-type control fish and lane 7 is from a heterozygousngn-1 mutant control fish. Products from the wild-type gene would show a band at 395 base pairs, heterozygous fish would show two bands at 395 base pairs and at 130 base pairs, and homozygous fish would show a single band at 130 base pairs. The fish corresponding to lanes 1 and 7 were heterozygous fish. The fish associated with lanes 2 and 6 were wild-type fish while fish to lanes 3, 4, and 5 were homozygous.

We injected Tg(pvalb3b:ribeye a-mCherry) and the Ppv3b-4 vector into the offspring of the ngn-1 heterozygous mutant fish. The Ppv3b-4 vector expresses GFP in hair cells and would allow for easy identification of hair cells along the posterior lateral line (McDermott et al., 2010). At 2 dpf, homozygous mutant ngn-1 fish are insensitive to physical touch (Golling et al., 2002). The protective envelope surrounding the embryo known as the chorion was removed from the injected embryos at 2 dpf, and the fish were touched with a transfer pipet. Fish that expressed no movement when touched were separated and allowed to develop for two more days. At 4 dpf, agar-imbedded fish were screened for expression of mCherry and GFP and viewed under the confocal microscope. The fish were then removed from the agar and allowed to develop for another day.

At 5 dpf, the imaged fish were labeled with 4-Di-2-ASP to look for an increase in the number of neuromasts along the posterior lateral line to confirm the homozygous mutant phenotype. Fluorescent dyes such as 4-Di-2-ASP can be added directly to the fish water and has the ability to enter the hair cells of neuromasts through the open mechanically gated transduction channels located at the tips of the stereocilia (Meyers et al., 2003). 4-Di-2-ASP diffuses throughout the cell, and the number of neuromasts

40 can be observed under a fluorescent microscope. An increase in neuromasts would further corroborate a homozygous mutant finding. The fish were then taken and euthanized to perform genotyping. A set of three primers were used to identify whether the fish were wild-type, heterozygous, or homozygous. These results would confirm the fish imaged under the confocal microscope were homozygous mutants and, therefore, lacked afferent innervation of the posterior lateral line hair cells (Figure 14B).

After ten rounds of injections, we were able to isolate 29 fish that were insensitive to touch at 2 dpf and had expression of both GFP and mCherry. This showed that the Ribeye a-mCherry fusion protein can be expressed in mutant zebrafish. After approximately four rounds of injections, we decided to abandon the 4-Di-2-ASP labeling because we felt it was an unnecessary step and could identify homozygous fish based on the touch response test and confirm it with by genotyping. Of the 29 fish obtained, 4 were wild type, 6 were heterozygous, 16 were homozygous, and 3 were unknown. 16 homozygous fish were imaged with only 4 fish showing Ribeye a-mCherry expression in the posterior lateral line (data not shown). Of the 4 fish that did show expression in the posterior lateral line, 2 had no movement of the ribbons while the other 2 did show movement. However, this movement did not appear abnormal when compared with movements in wild-type fish. Due to the lack of robust variations in ribbon movement, we could not make any conclusions about our speculations but we were able to show that the Ribeye a-mCherry fusion protein can be expressed in mutant zebrafish and could be used for characterization of mutant zebrafish.

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Determining the stability of Ribeye a using FRAP. Little is known about the dynamics of the molecules found in the CAZ but Bassoon has been shown to have a very low exchange rate between individual presynaptic sites at conventional synapses (Tsuriel et al., 2006). These experiments were tested using FRAP. Considering the anchoring properties of Bassoon along with the interaction between RIBEYE and Bassoon (Dick et al., 2003; tom Dieck et al., 2005), we hypothesized that it would be possible to use

Ribeye a-mCherry for FRAP studies which could be used to determine the dynamics of

Ribeye a in a live zebrafish. FRAP experiments consist of using high laser intensities to photobleach a fluorescent area and then monitoring the area for the return of fluorescence. The return of fluorescence is known as recovery and indicates non- bleached molecules have entered the photobleached area. Analysis of photobleaching the ribbons will be required to determine the exchange rate of Ribeye a. There are three possible outcomes of these FRAP experiments (Figure 15): No recovery (Model 1) would indicate that the molecules in the photobleached area are stable and immobile, while recovery could indicate exchange of molecules between the cytoplasm (Model 2) or neighboring ribbon synapses (Model 3). Further studies would have to be conducted to determine the origin of the unbleached fluorescent molecules entering the photobleached area to be able to distinguish between Model 2 and Model 3.

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Figure 15. Models of Ribeye a behavior and experimental outcomes for FRAP experiments. Models of Ribeye a behavior are shown on the top row. Experimental paradigms and outcomes of FRAP studies of hair cells expressing Ribeye a-mCherry are shown in rows 2-4. Bottom row shows the fluorescence recovery time course for photobleached ribbons for Models 1, 2, and 3. In FRAP experiments, if Model 2 or 3 is valid, plots will show recovery after photobleaching; plots will not show recovery if Model 1 is appropriate. Arrowheads show ribbons targeted for photobleaching or photoactivation. Straight arrows show ribbons where fluorescence measurements will be taken. Image and figure legend source: Dr. Brian M. McDermott

We conducted FRAP experiments using the Ribeye a-mCherry fusion protein in

Ppv3b-4 transgenic fish and examined the dynamics of Ribeye a in hair cells as well as in

43 the skin cells. Ppv3b-4 transgenic fish were injected with Tg(pvalb3b:ribeye a-mCherry) and screened for Ribeye a-mCherry fusion protein and GFP expression. Fish at 5 or 6 dpf that were positive for expression of both fluorescent proteins were imaged under the confocal microscope. Cells with at least two fluorescent puncta were chosen for FRAP studies. Regions of interest were formed around two areas of the cell and photobleaching occurred in one of the two regions of interest. The second region of interest served as a control. The puncta was photobleached to diminish the fluorescent intensity but not completely eliminate it. Due to the possible movement of the photobleached ribbon, we determined it was necessary to be able to view the photobleached ribbon in order to ensure the proper area is quantified for recovery.

Fluorescent recovery was monitored by collecting images before photobleaching, immediately after photobleaching, and specified time intervals after photobleaching.

Figure 16. Fluorescent recovery after photobleaching on zebrafish at 5 or 6 dpf. A, Skin cells expressing Ribeye a-mCherry. Arrow indicates the cell to be photobleached. B, Skin cells immediately after photobleaching the top portion of the cell depicted by the arrow in A. C, Skin cells 1 minute after photobleaching. D, Skin cells 60 minutes after photobleaching. E, Hair cell expressing Ribeye a-mCherry in a neuromast. The arrow indicates the puncta to be photobleached. F, Hair cells immediately after photobleaching the top puncta indicated by the arrow in E. G, Hair cells 1 minute after photobleaching. H,

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Hair cells 90 minutes after photobleaching. These fish express GFP in the skin and hair cells but the green channel was not shown to prevent interference of viewing the photobleaching and recovery.

During imaging it appeared in the hair cells there was very little fluorescent recovery in the ribbons while, surprisingly, there was a faster recovery rate in the skin cells. Skin cells begin to show recovery within 1 minute after photobleaching and appear to gain a significant amount of the fluorescence back after 60 minutes (Figure 16A, B, C,

D). Hair cells, on the other hand, show almost no recovery at 1 minute and only appear to show some recovery after 60 minutes (Figure 16E, F, G, H). Interestingly, movement has been seen in the photobleached ribbons of hair cells in the direction towards the neighboring control ribbon (Figure 16H). It remains unclear if these movements are a result of photobleaching, the natural movements of the ribbon, or a change in focal plane. These results do show though that Ribeye a-mCherry fusion protein has the ability to be photobleached.

Discussion

We have developed a method to visualize synaptic ribbons of the zebrafish ear and lateral line. After cloning the coding region of the ribeye a cDNA and ligating it to mCherry cDNA, we expressed the chimeric protein using a hair-cell promoter. Live or fixed hair cells that contained this fusion protein had fluorescent ribbons in close proximity to the basolateral surface (Figure 10B, D). When hair cells expressing Ribeye a-mCherry were immunolabeled with Ribeye b antiserum an overlap between fusion protein- and antibody-associated fluorescent signals was revealed (Figure 10E, F). In

45 addition, Ribeye a-mCherry-labeled ribbons localized proximal to afferent nerve endings

(Figure 10C). These findings indicate that Ribeye a-mCherry is an effective tool for labeling ribbons in zebrafish hair cells and will serve as a useful tool for real-time and fixed-tissue analyses. These experiments were performed in zebrafish mosaically expressing the Ribeye a-mCherry transgene. We would like to obtain a stable transgenic line which would consistently express Ribeye a-mCherry in all hair cells. Work is underway to search for a founder that transmits the transgene, with dozens of adult fish already screened. To date, we have yet to find a stable transgenic line but are optimistic that one will be found in the near future. A stable line expressing Ribeye a-mCherry in hair cells would eliminate the need for injections and increase the efficiency of developmental studies.

In the process of creating the vector for expressing the Ribeye a-mCherry fusion protein, we were able to generate two useful vectors. pMT/SV/PV can be used to express numerous proteins in the hair cells of zebrafish. cDNA encoding a protein and a fluorescent tag could be inserted in this vector using the MCS and be used for expression in hair cells as it has been shown previously with the expression of Fascin 2b fusion protein (Chou et al., 2011). We also generated a construct, pMT/mCh, which was shown to produce red fluorescent hair cells in zebrafish (Figure 9). This could be a useful way to provide a contrast during imaging of transgenic fish expressing GFP other areas of the hair cells or in immunolabeling studies. A stable transgenic line of fish expressing this construct would also be beneficial for developmental studies.

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In an effort to use our newly developed Ribeye a-mCherry tool and to gain an understanding of ribbon development, we sought to analyze the movements of ribbons in ngn-1 mutant fish. We had previously seen movement with the hair cells of transgenic fish and theorized that these ribbons could be searching for an appropriate area of the plasma membrane to associate with which would be juxtaposed to the afferent neuron

(Figure 12). This would correlate with the proposed model that ribbons are generated in the cytosol and are situated at the basolateral membrane after afferent neuron contact. ngn-1 mutant fish lack posterior lateral-line afferent innervations, and we believed that the ribbons in the hair cells of the neuromasts of the posterior lateral line would all be moving at a high rate of speed searching for the appropriate area of the plasma membrane to associate with. This result would indicate Model one (Figure 13A) is the correct model.

This was not the case in the four homozygous ngn-1 fish expressing Ribeye a- mCherry that we imaged. Two of the fish had stable ribbons at the basolateral surface, and the other two showed some movement but nothing that appeared uncharacteristic when compared to the movements in transgenic fish. Had Model one (Figure 13A) been correct, we would have expected to see ribbons moving about, perhaps arbitrarily, in the cell, or the ribbons would have been located in the interior regions of the cells waiting for afferent neuron contact for their recruitment to the basolateral membranes.

Based on the data obtained, we believe that recruitment of the ribbons to the basolateral membrane is independent of innervation. An increase in overall size would be needed to provide more evidence which would allow for elimination of the notion

47 that the ribbons are formed in the cytosol and recruited to the basolateral membrane after afferent nerve contact but we were able to show that Ribeye a-mCherry is able to be used for characterization of mutant zebrafish. We have recently obtained a Ribeye b-

GFP stable transgenic line, which expresses this protein in hair cells, from another laboratory and these fish give the potential to revisit this project (Sheets et al., 2011). It is possible to cross the heterozygous ngn-1 fish with the stable Ribeye b-GFP line to produce a homozygous ngn-1 fish expressing Ribeye b-GFP in all the hair cells and similar experiments could be performed. Recent studies have shown that reducing the amount of Ribeye in hair cells results in reduced afferent innervations providing support for the notion that ribbons are required for stabilizing contact sites with afferent neurons (Sheets et al., 2011)

Exchange of Ribeye a between ribbons and cytosolic pools. In an attempt to understand the exchange of molecules in cells and to exhibit another use of Ribeye a- mCherry, we performed FRAP experiments on the hair cells and skin cells of fish expressing Ribeye a-mCherry. Our results show that the Ribeye a-mCherry fusion protein is capable of being photobleached and can be used for FRAP experiments

(Figure 16). Based on the data obtained, we can theorize that skin cells could have a high exchange rate of Ribeye a and hair cells could have a low exchange rate. It is possible that when Ribeye a is ectopically expressed in skin cells it is primed to bind other Ribeye a binding partners and there is a fast exchange between sites similar to what is seen in Model 3 of Figure 15. Slow exchange rates in the hair cells could be due to the degradation of Ribeye a and the incorporation of newly synthesized proteins as

48 seen in Model 2 of Figure 15. It will be necessary to perform more FRAP experiments as well as analyze the data to draw conclusive results about the dynamics of Ribeye a.

After obtaining more FRAP data, it will be important to determine whether

Model 2 or Model 3 is occurring in the skin cells and hair cells. One way would be to use photoactivatable GFP (PAGFP) (Patterson and Lippincott-Schwartz, 2002). PAGFP is virtually invisible until activated with high intensity lasers. We would like to perform fluorescent recovery after photoactivation (FRAPA) experiments by creating another vector in which PAGFP is exchanged for mCherry in Tg(pvalb3b:ribeye a-mCherry). We would then proceed to create hair cells and skin cells that express both Ribeye a- mCherry and Ribeye a-PAGFP. Ribeye a-mCherry would be used to locate the ribbons since PAGFP is nearly invisible before activation. We would then photoactivate PAGFP and observe any changes in fluorescence. If fluorescence is observed in a non-activated ribbon this would indicate exchange between the ribbons and Model 3 would be the appropriate model (Figure 15).

Another method which could be used to distinguish exchange between synapses or newly synthesized proteins is called TimeSTAMP, time-specific tagging for the age measurement of proteins (Lin et al., 2008). This is a nontoxic method which utilizes epitope tags. This strategy uses a specific protease activity to remove an epitope tag bound to the protein of interest. During protein synthesis the epitope would be removed until the corresponding protease inhibitor was added. After adding the inhibitor it would become evident which proteins were synthesized at that time point.

49

This method has been shown to work in vivo and in live animals which is beneficial for our use in zebrafish. We would like to add the TimeSTAMP tag to Ribeye a to determine if the slow recovery rates seen in hair cells is due to the incorporation of newly synthesized proteins. If fluorescence is seen after the addition of the drug then Model 2 would be the accurate account of fluorescent recovery in ribbon synapses.

In summary, we have developed a method to visualized synaptic ribbons of the zebrafish ear and lateral line. We were successful in creating a Ribeye a-mCherry fusion protein under the expression of a hair cell promoter. We showed in live and in fixed hair cells that expression of Ribeye a-mCherry is located at the basolateral region, juxtaposed to afferent nerve endings. In addition, hair cells expressing Ribeye a- mCherry displayed an overlap with Ribeye b antiserum when immunolabeled. These findings indicate that Ribeye a-mCherry is an effective tool for labeling ribbons in live and fixed hair cells of zebrafish. We were able to showcase the use of this tool through the characterization of ngn-1 mutant zebrafish and in FRAP studies.

50

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