Localization Study of Supervillin in Zebrafish Hair Cells

LOCALIZATION STUDY OF SUPERVILLIN IN ZEBRAFISH HAIR CELLS USING IMMUNO-FLUORESCENCE ASSAY & IDENTIFICATION OF SMALL MOLECULES THAT IMPACT THE INNERVATION OF THE LATERAL LINE SYSTEM OF DEVELOPING ZEBRAFISH

NILAY GUPTA

Submitted in partial fulfilment of the requirements for the degree of

Masters of Science

Department of Biology

CASE WESTERN RESERVE UNIVERSITY

MAY 2016

CASE WESTERN RESERVE UNIVERSITY

SCHOOL OF GRADUATE STUDIES

We hereby approve the thesis of

Nilay Gupta

Candidate for the degree of Master of Science.

Committee Chair

Hillel Chiel, Ph.D.

Committee Member/Research Advisor

Brian M. McDermott, Ph.D.

Committee Member

Stephen Haynesworth, Ph.D.

Committee Member

Susan Burden-Gulley, Ph.D.

Date of Defense

March 18th, 2016

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

Dedicated to my little brother, Kush

Table of Contents Contents Page number List of Figures 7

List of Abbreviations 9

Acknowledgements 13

Abstract 15

Background The Sense of Hearing 17 Information

The Hair Cells 19

Hearing in Zebrafish 21

Advantages of using zebrafish as a model 23 organism

Part I - Localization study of supervillin in zebrafish hair cells using 25 immuno-fluorescence assay

Introduction Hair cell cuticular plate 26

Known proteins that constitute cuticular plate 29

Supervillin 31

Supervillin in the vertebrate hair cells 33

Materials and Zebrafish strains and husbandry 37 Methods

Generation of zebrafish Svila antibody 37

Zebrafish whole-mount immunofluorescence 38

Imaging under confocal microscope 38

Results Supervillin localization in GFP-fascin 2b 39 transgenic zebrafish

Supervillin localization in Gt(macf1a-citrine)ct68a/+ 40 transgenic zebrafish

Discussion 42

Part II – Identification of small molecules that impact the innervation of 47 the lateral line system of developing zebrafish

Introduction Lateral line system in zebrafish 48

Development of the posterior lateral line (PLL) 52 system in zebrafish

Innervation of the lateral line system in zebrafish 54

The HGn39D transgenic zebrafish 56

In vivo chemical screening in zebrafish 58

Materials and Zebrafish strain and husbandry 60 Methods Chemical library 60

Dechorionation of zebrafish embryos 61

Primary chemical screen and rescreen protocol 61

Imaging and phenotype analysis 62

Results Phenotype 1 68

Phenotype 2 69

Phenotype 3 71

Phenotype 4 72

Phenotype 5 73

Phenotype 6 75

Phenotype 7 76

Phenotype 8 76

Phenotype 9 77

Phenotype 10 78

Phenotype 11 79

Phenotype 12 80

Discussion 83

Appendix 88

References 110

List of Figures Figure Description Page Number number Figure 1 The structure of the human ear 18

Figure 2 Schematic of a hair cell 20

Figure 3 Ear and lateral line system in the larval zebrafish 23

Figure 4 Presence of linker proteins in the cuticular plate 28

Figure 5 Schematic showing supervillin functional domains 32

Figure 6 Images from whole mount in situ hybridization of 4 dpf 34 zebrafish

Figure 7 Supervillin localization in the mouse vestibular and 36 cochlear hair cells as observed under confocal microscope

Figure 8 Confocal images of a 4 dpf GFP-fascin 2b transgenic 39 zebrafish immunolabeled with anti-Svila

Figure 9 Confocal images of a 4 dpf Gt(macf1a-citrine)ct68a/+ 41 transgenic zebrafish immunolabeled with anti-Svila

Figure 10 Schematic for supervillin localization in the organ of Corti 44 and models for its function

Figure 11 Schematic for supervillin localization in the zebrafish 45 macula

Figure 12 Schematic showing zebrafish lateral line system and 50 individual neuromast

Figure 13 Schematic to depict planar cell polarity and mirror 51 symmetry in the hair cells of a nueromast

Figure 14 Posterior lateral line primordium migration in a Zebrabow 54 transgenic line

Figure 15 Schematic depicting the ALL and PLL ganglia 55

Figure 16 Expression of EGFP by the HGn39D transgenic line of 57 zebrafish in the afferent neurons of the lateral line system

Figure 17 Schematic to show the design of the chemical screen 63

Figure 18 Phenotype 1 - Phenotype with poor afferent neuronal 69 development

Figure 19 Phenotype 2 - Phenotype with hair cell-like structures at the 70 end of dendritic arbors

Figure 20 Confocal micrographs of the phenotype exhibiting the hair 71 cell-like structures on the dendritic arbors

Figure 21 Phenotype 3 - Phenotype with the wavy sensory afferent 72 axon in the lateral line system

Figure 22 Phenotype 4 - Phenotype wherein the sensory afferent axon 73 in the lateral line system terminates prematurely

Figure 23 Phenotype 5 - Phenotype where the sensory afferent axon 74 in the posterior lateral line system appears to be thinner

Figure 24 Phenotype 6 - Phenotype where the sensory afferent axon 75 in the posterior lateral line system is wavy and ends prematurely Figure 25 Phenotype 7 - Phenotype where the zebrafish embryos 76 show abnormal morphology

Figure 26 Phenotype 8 - Phenotype in HGn39D zebrafish embryos 77 where they show GFP expression/green fluorescence in the somites Figure 27 Phenotype 9 - Phenotype wherein zebrafish embryos 78 exhibit an abnormal morphology of the heart

Figure 28 Phenotype 10 - Phenotype showing fluorescent cell-like 79 structures in zebrafish circulation

Figure 29 Phenotype 11 - Phenotype showing fluorescent patches on 80 the body surface

Figure 30 Bar-graph depicting the number of hits obtained in this 82 small molecule screen

Figure 31 Schematic showing different approaches for target 84 identification after a small molecule screen

Table 1 ‘Hits’ obtained from the small molecule screen on the 64 HGn39D transgenic zebrafish and their phenotypes

List of Abbreviations ac anterior crista

ACF7 actin crosslinking factor 7

ALL anterior lateral line

AP anterior-posterior axis bp base pairs

CaCl2 calcium chloride cDNA complementary DNA

Cntnap2a contactin associated protein-like 2/Caspr2

CP cuticular plate

CWRU Case Western Reserve University

Da Dalton

DC Deiters’ cells

DMSO dimethylsulfoxide

DNA deoxyribonucleic acid

Dpf days post fertilization

EB embryo buffer

EGFP enhanced green fluorescent protein

F-actin filamentous actin

FCH FER-CIP4 homology

FCHSD1 FCH domain and double SH3 domains containing protein 1

FDA US food and drug administration

G-actin globular actin

GEF GTP exchange factor

GFP green fluorescent protein

GST glutathione S-transferase

GTP guanosine triphosphate h hours

HNK-1 human natural killer 1 hpf hours post fertilization

HTS high throughput screening

IACUC Institutional Animal Care and Use Committee

IgG gamma immunoglobulin

IHC inner hair cells

IPC inner phalangeal cells

KCl potassium chloride kDa kilodalton

KH2PO4 monopotassium phosphate lc lateral crista

MDBK Madin–Darby bovine kidney

MgSO4 magnesium sulfate mM millimolar

Mm millimeter

MOA mechanism of action mRNA messenger RNA

Na2HPO4 disodium phosphate

NaCl sodium chloride

NaHCO3 sodium bicarbonate

ngn1 neurogenin-1

NIDCD National Institute on Deafness and Other Communication Disorders

NLS nuclear localization signal oC degree centigrade

OHC outer hair cells

OP outer pillar cells

PBS phosphate buffered saline pc posterior crista

PCDH15 protocadherin 15

PCP planar cell polarity

PCR polymerase chain reaction

PLL posterior lateral line

PTU 1-phenyl 2-thiourea

RNA ribonucleic acid

RNA-seq high throughput RNA sequencing

RPKM reads per kilobase per million mapped reads

RT-PCR reverse transcription PCR

SAR structure-activity relationship

SDF1 stromal-derived factor 1 sm saccular macula

SNX9 sorting Nexin 9

SVIL supervillin

TALEN transcription activator-like effector nuclease

um utricular macula

XIRP2 Xin-actin binding repeat containing 2

l microliter

m micrometer

Acknowledgements

I owe these research projects and this thesis to my research advisor, Dr. Brian M.

McDermott. I cannot thank him enough for his constant guidance and support. He plays a big role in shaping me as a researcher and has always been encouraging during the hard times. Apart from research, he has also encouraged me to reignite my interest in art, which has helped me to develop skills as an illustrator.

My deep sense of gratitude goes to my thesis committee members, Dr. Stephen

Haynesworth, and Dr. Susan Burden-Gulley who have been excellent teachers and have helped me develop a deeper understanding of cell and developmental biology. Dr.

Burden-Gulley’s guidance during independent research course and her valuable suggestions have truly helped me shape this project. I would also like to thank Dr. Hillel

Chiel for his guidelines pertaining to defense and for serving as my CGA representative.

Words are inadequate to thank all my lab members who continue to make a great work atmosphere in the lab. I specially want to thank Carol Fernando for teaching me the basics about the zebrafish and helping me maintain them, without which this project would not have been possible. I owe my success in the supervillin project to Dr. Lana

M. Pollock, who has been a great mentor, teacher, and a friend. Thanks to Robin Wood

Davis for always providing an honest opinion, and for numerous texts during this thesis- writing time, which kept me going. I also extend my thanks to other lab mates, including

Shih-Wei (Victoria) Chou, Nicholas Sarn, Shaoyuan (Sara) Zhu, Kevin Chen, Wenbo

Chen, Bona Ko, Tracy Chen, and Karthik Mohanarangan.

I would also like to thank my friends outside of the lab including everyone from

International Student Fellowship. I thank Sharoon Hanook for being an elder brother

and my family away from home. Thank you Megan Stonebraker for being the nicest friend who always takes my side; Soumili Chatterjee, Wayne Gibson, Richard Nii Larte

Lartey, Thomas Atta-Fosu, Sneha Bandi, Nandini Puttarudraiah, and Bok Chew for making my study abroad a memorable experience. A special thanks to Robert

McMurray and his dog Lulu, who has been exceptionally helpful as a stress-reliever during this time.

I am truly thankful to God for His blessings. I am indeed lucky to have parents who have always shown trust in me and have stood by me through thick and thin. My little brother has been my constant motivation during this time and it is his strength that has ultimately helped me to successfully complete my masters. It is their support and blessings without which anything including this project would have been a distant reality.

Working on these projects and writing this thesis has by no means been a solo effort. I would like to thank everyone who has contributed directly or indirectly towards this project.

Localization Study of Supervillin in Zebrafish Hair Cells Using Immuno-fluorescence Assay & Identification of Small Molecules that Impact the Innervation of the Lateral Line System of Developing Zebrafish

Abstract NILAY GUPTA

The zebrafish has emerged as a powerful model organism over the last two decades due to its closer evolutionary relationship to humans than invertebrates, easy maintenance, and high fecundity. These qualities of zebrafish have been exploited in this project to address two main questions. First, to localize supervillin in zebrafish hair cells. Hair cells are responsible for hearing and balance in zebrafish as well as humans. Actin is a cytoskeletal protein, which along with actin-associating proteins, provides the structural framework for stereocilia and the cuticular plate in the hair cells. The cuticular plate provides a strong foundation for stereocilia but is poorly understood in terms of protein composition and its development. Supervillin is one of the actin- associating proteins. In this study, we examined zebrafish hair cells using immunofluorescence assay to localize supervillin. We have demonstrated that supervillin localized specifically to the hair cell cuticular plate in the transgenic lines of zebrafish, which can be indicative of its role in shaping and maintaining the integrity of this mysterious organelle.

In the second study, the transgenic HGn39D zebrafish line is used to design a small molecule screen to test the effects of the chemicals on the lateral line afferent neurons.

Afferent innervation in the lateral line of the zebrafish serves a model for peripheral nervous system and this preliminary screen can help in identifying the chemicals that

can be used to address problems such as peripheral nerve damage. The HGn39D fish express GFP in the afferent neurons of the lateral line system, including anterior lateral line (ALL) and posterior lateral line (PLL) ganglia. Migration of growth cones of afferent neurons start around 24 hpf, and we used this time point to dechorionate the embryos and subject them to 1,040 different small molecules from a chemical library.

A small molecule that altered the phenotype of the HGn39D zebrafish was considered a ‘hit’. A total of 63 ‘hits’ were confirmed after rounds of phenotypic screening. These

‘hits’ were grouped into 12 broad categories, based on the phenotype displayed, which ranged from almost no afferent neuronal development to hair cell-like structures on the dendritic arbors. Studies leading to the identification of the mechanism of action of the

‘hits’ that exhibit a particular phenotype might also help in better understanding of sensory processing of lateral line system in zebrafish and its implication with respect to hearing research.

Background Information

The Sense of Hearing

Hearing is one of the five traditional senses in humans. It is defined as perception of the sound vibrations through the ear by the brain. The perception of sound relies on mechanosensation since it depends on the mechanical waves, or the sound vibrations, that are perceived by the ear and are transmitted to the brain through a nerve impulse.

Unfortunately, 2 to 3 out of every 1000 children are born with some degree of deafness in either one or both ears in the USA, according to the National Institute on Deafness and Other Communication Disorders (NIDCD). Hearing loss can either be conductive hearing loss, wherein the deafness is due to damage to the outer or middle ear, or it can be sensorineural hearing loss which happens due to damage in the inner ear and to the hair cells. Although not life-threatening, hearing disorders can cause serious outcomes, such as impaired speech development in childhood, and social depravation in adults, which in the worst cases can lead to psychological disorders (Principles of Neural

Science 5th Ed. Kandel et al).

Sound stimuli are received by the outer ear, pinna, which serves as an antenna for the sound waves. These waves then travel to the middle ear through a tubular structure that is open only at one end and on the other end is the tympanum, or the eardrum. The tympanum is responsible for transmitting sound waves to the inner ear with the help of three ossicles – malleus, incus, and stapes. These ossicles are connected to each other to form an ossicular chain, which ends at the oval window of the bony, fluid-filled cochlea. The cochlea is a snail-shaped organ that is responsible for the sense of hearing in mammals and forms the auditory system (Keen, J.A., 1939). The organ of Corti is located in the cochlea and contains an array of hair cells, which are the ultimate

receptors of sound stimuli. The sound waves that vibrate the tympanum subsequently vibrate the basilar membrane inside the cochlea, which in turn causes deflection of the stereocilia of the hair cells and transmission of the sound signal. The inner ear also contains the vestibular system, which is comprised of three semi-circular canals that help mammals in spatial orientation and balance (Figure 1).

Figure 1 – The structure of the human ear. The human ear is divided into three parts – outer, middle and inner ear. The outer and middle ear are responsible for transferring the sound waves to the inner ear, which is comprised of auditory system (cochlea) for hearing, and the vestibular system (semicircular canals) for balance. The cochlea contains the organ of Corti which is comprised of hair cells – the cells that confer to us the sense of hearing. Image from Principles of Neural Science 4th Ed. Kandel et al. (A. J. Hudspeth).

The Hair Cells

Hair cells in humans and other mammals are found both in the auditory system as well as the vestibular system. The hair cells have been named so because of the bundle of

‘hair’ like structures called stereocilia that are present on the apical surface of these cells. Each hair cell has about 30 – 300 actin-based stereocilia that project into the K+ ion-rich endolymph (Corwin, J. T. et al., 1991). In addition to stereocilia, hair cells also have one microtubule-based projection called the kinocilium, which is the only true cilium found in the hair cell. It is believed that the kinocilium plays a role in establishing the polarity of the hair cell. Stereocilia are graded according to their height and form a staircase like pattern which is essential to their function. Each stereocilium is connected to the adjacent stereocilium via a tip link, which are attached to the gated-ion channels

(Furness, D. N. et al., 1985). Towards the base, stereocilia taper to form rootlets and insert themselves into another actin-based organelle – the cuticular plate (CP). The cuticular plate is believed to be responsible for forming a firm foundation for the stereocilia (hair bundle) so that they can endure the mechanical stress caused by the deflections due to sound waves (Figure 2). Each hair cell is also tightly connected to the adjacent supporting cells via tight junctions and adherens junctions (Hirokawa, N. et al., 1982).

The cochlear canal in the auditory system of mammals is divided into three compartments – scala tympani, scala vestibuli and scala media (Keen, J.A., 1939). Scala media contains the organ of Corti that sits on the basilar membrane and is overlaid by the tectorial membrane. Movement of the basilar membrane against the tectorial membrane deflects the hair bundle, and the gated ion channels on the stereocilia are pulled open to allow the influx of K+ ions into the hair cells, causing the hair cell to depolarize (Corey, D. P. et al., 1979). This electric signal is transmitted to afferent

neurons through the neurotransmitter, glutamate, and is perceived by the brain as a sound signal.

Figure 2 – Schematic of a hair cell. Hair cells are the receptors of sound in the inner ear of vertebrates. Actin-based stereocilia are rooted in a dense meshwork of actin called the cuticular plate. Every hair cell is surrounded by supporting cells and is innervated by afferent neurons. The sound is detected by the hair cell when a deflection of stereocilia causes depolarization of the cell; this signal is transferred to the brain through the afferent neurons. Image from Principles of Neural Science 5th Ed. Kandel et al.

Hearing in zebrafish

We might not think of hearing as an anthropomorphic characteristic, but evolutionary biologists have proven that the auditory functions in the vertebrates evolved as early as in hagfish, lampreys, bony fishes and amphibians (Jorgensen, J. M. et al., 1998). Birds and mammals have the most advanced auditory function and one of the most elaborate inner ear structures (Fay R. R. et al., 2000). In acousticolateralis theory, it is believed that the inner ear has been evolved as a variation of the lateral line system of the fish

(Ayers H., 1892).

Zebrafish exhibit an auditory system that is widely shared among the vertebrates in their functions and functional relationships. Zebrafish and other fishes have a lateral line system, which also works on the similar principle as hearing but has different functions such as predator avoidance, prey-finding, schooling and rheotaxia

(Engelmann, J. et al., 2000). The fundamental transducing unit in all the vertebrate ears and fish lateral lines is the hair cell, which has the same overall morphology from the most primitive vertebrates to humans (Jorgensen, J. M. et al., 1998). In addition to that, the deafness-related genes are conserved between zebrafish and humans (Whitfield, T.

T., 2002).

Development of zebrafish ear starts as an ectodermal thickening in the form of the otic placode on either side of the head at around 16 hours post-fertilization (hpf).

Collectively, five patches of sensory epithelium arise from this placode – two maculae

– utricular macula (um) and saccular macula (sm) – and three cristae – anterior crista

(ac), lateral crista (lc) and posterior crista (pc) (Figure 3). The ear also contains three semi-circular canals made up of projections from epithelial tissue over the three cristae and two calcium carbonate crystals called otolith that overlay the maculae (Nicolson,

T., 2005). These structures detect changes in pressure, gravity, flow and movement of the fish and pass it to the hair cells in the sensory epithelium for mechanosensation

(Whitfield, T. T. et al., 2002). In addition to the ear, zebrafish also have sensory patches on their skin along the lateral line system, which are functionally similar to the ear

(Figure 3). The sensory patches on the lateral line are a group of hair cells clustered together in what is called a neuromast. The anterior lateral line (ALL) system is comprised of the neuromasts on the head of the fish. The posterior lateral line (PLL), which extends from trunk to tail in the zebrafish larvae contain around 5 lateral line neuromasts and 2-3 terminal neuromasts on the caudal fin (Raible, D. W. et al., 2000).

The hair cells in the neuromasts are virtually identical to the hair cells found in the ear of the zebrafish.

Figure 3 – Ear and lateral line system in the larval zebrafish. A. Schematic showing the ear of a larval zebrafish. Cristae are shown in green – ac, lc and pc for anterior, lateral and posterior cristae and, maculae are shown in red – am and pm for anterior and posterior macula; ao and po are the anterior and posterior otoliths. B. Lateral line system in zebrafish with neuromasts in blue and red dots shown for anterior and posterior lateral line respectively.

Advantages of using the zebrafish as a model organism

As mentioned in the previous section, the auditory system has been generally conserved within vertebrates. Moreover, hair cells are morphologically similar in zebrafish and humans. Although humans do not have the hair cell regenerative capacity as zebrafish, they are still very much suited for hearing research because zebrafish larvae are optically transparent which makes it possible to observe the development of hair cells

without any need for dissections, unlike mammalian model systems like mice. In addition to this, zebrafish have ex utero fertilization which facilitates the study of the development of ear from the very beginning. The lateral line system also has clusters of hair cells on the surface of the fish which can be easily manipulated and studied using techniques such as electrophysiology and microphonics. Zebrafish also have rapid development and a short generation time (2 - 4 months), and they are easy to handle and maintain compared to other animal models. The zebrafish genome has been sequenced and they have 12,719 genes in common with mice and humans (Howe, K. et al., 2013). A well-annotated zebrafish genome enables useful and relatively easier gene manipulation and genetic screens. They have become an animal model of choice in the fields of research in muscular dystrophy, tuberculosis, schizophrenia and even cancer. Zebrafish can produce around 150-300 eggs at a time, making them one of the most suitable models for high-throughput drug screens as well.

In this study, the quality of optical transparency during early development has been exploited to study the localization of a protein, supervillin, in the hair cells of the larval zebrafish using fluorescence microscopy. In addition to this, the high brood number in zebrafish has also facilitated a drug screen to examine the effects of various small molecules on the afferent neurons of developing transgenic HGn39D zebrafish.

Part I

Localization study of supervillin in zebrafish hair

cells using immuno-fluorescence assay

Introduction

Hair cells are the fundamental receptors of sound stimuli. These specialized cells are considered an excellent example of cytological micro-architecture. The apical surface of the hair cell contains rows of graded actin-based stereocilia, which upon deflection cause the transduction of sound stimuli (Hudspeth, A. J., 2005). The stereocilia are composed of parallel F-actin filaments (Tilney, L. G. et al., 1980) which are held together with the help of various proteins such as fimbrin (Slepecky, N. et al., 1985), espin (Sekerková, G. et al., 2006), and fascin 2b (Chou, S. W. et al., 2011). Each stereocilium has uniform thickness but narrows towards the base into rootlets, which are composed of densely packed actin filaments that anchor each stereocilium into the actin-based cuticular plate (CP). The rootlets are necessary to provide flexibility to the stereocilia during deflection (Furness, D. N. et al., 2008). When the stereocilia are deflected, tip links on the stereocilia pull open the gated channels that allow for influx of ions and subsequent transduction of sound stimuli (Assad, J. A. et al., 1991). The cuticular plate is believed to be the foundation that anchors the hair bundle and provides strength to overcome the mechanical stress caused due to deflections (Kachar, B. et al.,

1997). There are various speculations about the functions and development of the cuticular plate but very little is known about the development and maintenance of this actin-based organelle.

Hair cell cuticular plate

The apical surface of the hair cell is crowned with a bunch of specialized microvilli called stereocilia. Just beneath the stereocilia, and in the most apical region of the cytoplasm in the hair cell, lies the cuticular plate. The cuticular plate is mainly made up

of a meshwork of F-actin filaments. The core actin filaments of the stereocilia enter the cuticular plate in the form of rootlets (Pollock, L. M. et al., 2015) (Figure 2). Since stereocilia are specialized microvilli, the cuticular plate is also considered a specialized version of the terminal web (Hull, B. E. et al., 1979). Actin filaments in the microvilli

(such as those found in the brush border epithelium of the intestine) are rooted into the meshwork of actin and intermediate filaments called the terminal web. However, the cuticular plate has been found not to contain any intermediate filaments (Tilney, M. S. et al., 1989). Another actin-based structure on the apicolateral surface of the hair cells is the circumferential belt, which is made up of parallel actin filaments running along the lateral edges of the cell. This band of actin filaments lies adjacent to the adherens junctions and is responsible for cell-cell adhesion with the supporting cells (Hirokawa,

N. et al., 1982). The circumferential belt however, does not make any actin-based contact with the cuticular plate and the region between the cuticular plate and the circumferential belt is mostly actin-free (Kachar, B. et al., 1997).

Nobutaka Hirokawa and Lewis G. Tilney performed a study in 1982 where they found several linker proteins in the chick hair cell cuticular plate using quick freezing/etching, rotary shadowing and electron microscopy (Hirokawa, N. et al., 1982). These linker proteins were found to be linking actin filaments in the rootlets to one another, actin filaments in the rootlets to the actin filaments in the cuticular plate, actin filaments in the cuticular plate to one another, actin filaments to the underlying microtubules, and actin filaments to the apical plasma membrane (Figure 4) (these branched connecting units were called “crow’s foot”) (Hirokawa, N. et al., 1982). The results from these studies prove the presence of some linker proteins in the cuticular plate, but their precise identity still needs to be discovered (Arima, T. et al., 1987).

Figure 4 – Presence of linker proteins in the cuticular plate. A. Cartoon of a hair cell showing cuticular plate with various linker proteins (colored rectangles) connecting actin filaments to one another in the rootlets (red), actin filaments to the plasma membrane (blue), actin filaments to one another in the cuticular plate (orange), actin filaments from cuticular plate to the filaments of the rootlets (yellow) and actin filaments to the microtubules (pink). B. Cross section of a chick hair cell cuticular plate as observed under an electron microscope showing rootlets and the linker proteins (arrows) (Hirokawa, N. et al., 1982).

Known proteins that constitute the cuticular plate

The cuticular plate is composed of a meshwork of actin filaments that are made up of both -actin and -actin (Perrin, B. J. et al., 2010). Actin filaments in the stereociliary rootlets interact with tropomyosin, an actin-binding protein, which may confer the stability to the rootlets within the cuticular plate (Slepecky, N. et al., 1985). Another protein -actinin localizes to the apical region of the cuticular plate and between the stereociliary rootlets. -actinin is believed to crosslink and organize actin filaments in the cuticular plate to give rigidity (Wagner, O. et al., 1999). It might also play a role in linking the actin filaments that make up the circumferential belt (Drenckhahn, D. et al.,

1982). A spectrin-related protein called fodrin is found to localize in the cuticular plate and in the infracuticular structure called the striated organelle (Dememes, D. et al.,

1992). Fodrin is believed to cross-link actin filaments in the cuticular plate to provide rigidity and also link actin to the proteins associated with the plasma membrane.

Fimbrin was found to be present in the stereocilia and the cuticular plate using immunofluorescent localization studies, but the function of the cuticular plate is still unknown (Slepecky, N. et al., 1985).

Myosin-VI is found in the hair cell cytoplasm but is concentrated mainly in the pericuticular necklace (actin-free zone around the CP) and the cuticular plate (Hasson,

T. et al., 1997); it is believed to link the apical plasma membrane to the actin filaments of the cuticular plate. Myosin-VIIa is also located to the cuticular plate, pericuticular necklace, and also in the stereociliary bundle (Hasson, T. et al., 1997). Absence of this protein causes disorganized stereocilia and a structurally abnormal cuticular plate. In addition to that, mutations in the gene encoding myosin-VIIa – MYO7A, causes Usher syndrome type-1b and non-syndromic deafness in humans (Weil, D. et al., 1995).

Harmonin b localizes in the stereocilia and the pericuticular necklace, and might be

indirectly involved in F-actin bundling and interactions with myosin-VIIa (Boeda, B. et al., 2002).

Profilin is found to co-localize with actin in the hair cell cuticular plate (Slepecky, N.

B. et al., 1992). This protein has been known to bind with globular actin (G-actin) and prevent its polymerization into actin filaments (Carlsson, L. et al., 1977). However, it also interacts with another protein called diaphanous, which promotes actin polymerization (Lynch, E. D. et al., 1997). Protocadherin-15 is found in the stereociliary tip links, stereocilia, and near the cuticular plate (Ahmed, Z. M. et al.,

2006). Mutations in the gene that encodes for this protein have shown to cause hearing and balancing defects in the mice (Alagramam, K. N. et al., 2001). XIRP2 (Xin-actin binding repeat containing 2) is found in the hair bundle, cuticular plate, and the circumferential actin belt of the sensory hair cells (Francis, S. P. et al., 2015). Loss of

XIRP2 function may cause hair bundle degeneration and may compromise the epithelial barrier since it is located in the circumferential belt region. FCHSD1 (FCH (FER-CIP4 homology) domain and double SH3 domains containing protein 1) is mainly found in the cuticular plate of the mouse cochlear hair cells (Cao, H. et al., 2013). It interacts with SNX9 (Sorting Nexin 9) and promotes F-actin polymerization. FCHSD1 along with SNX9 is also believed to modulate the F-actin assembly and maintenance in the cuticular plate.

Acf7 (Actin crosslinking factor 7) is another protein that is interwoven into the cuticular plate and encircles the fonticulus – a region where the kinocilium inserts in the hair cell.

It also circumscribes and underlies the cuticular plate in the zebrafish hair cell. Acf7 connects actin filaments in the cuticular plate to the microtubules and might also play a role in hair cell polarity because Acf7 precedes hair bundle and cuticular plate formation in the hair cells (Antonellis, P. J. et al., 2014).

Supervillin

Previous studies in the lab (by Lana M. Pollock) predicted and identified various candidate proteins that interact with actin using the mRNA information from the hair cell transcriptome (McDermott Jr., B. M. et al., 2007). One of those candidate proteins is an actin-binding protein called supervillin (SVIL) or p205. The name ‘supervillin’ was proposed since it is the largest member of the villin/gelsolin superfamily

(Pestonjamasp, K. N. et al., 1997) weighing 205 kDa. The protein sequence is comprised of a conserved COOH terminal which is similar to other members of the villin/gelsolin family and an amino terminus that has multiple potential nuclear localization signals (NLSs) (Figure 5). The NLS part of supervillin is involved in various specific nuclear functions including binding to androgen receptors, thereby translocating it to the nucleus and enhancing its transactivation (Ting, H. J. et al., 2002).

Supervillin was also found to co-localize with LSD1-bound (lysine-specific demethylase 1) receptors where it mediates histone H3K9me2 demethylation in the SH-

SY5Y cells. Interaction of supervillin with LSD1 and its role as a cofactor was found to be important in neuronal maturation since supervillin controls LSD1 mediated H3K9 demethylation (Laurent, B. et al., 2015).

SVIL was first isolated from bovine neutrophils where it was found to be tightly associated with actin near the plasma membrane. It is also proposed to be involved in cell adherence and cell-cell junctions since it showed increased expression in the high cellular density of Madin–Darby bovine kidney (MDBK) cells and low expression in isolated cells (Pestonjamasp, K. N. et al., 1997). Human supervillin is strikingly similar to its bovine homolog with respect to nuclear localization signals and F-actin binding domains. Expression of this protein in humans is mainly found in muscle cells, bone marrow, thyroid and salivary glands (Pope, R. K. et al., 1998).

Supervillin tightly associates with actin filaments adjacent to the plasma membrane and its over-expression has been shown to dramatically increase the amounts of F-actin and vinculin in the cell. It has also been shown that supervillin promotes the F-actin bundling, and recruitment of F-actin filaments to the plasma membrane and cell nucleus

(Wulfkuhle, J. D. et al., 1999). The amino terminus of supervillin has one domain that interacts with myosin IIA and three domains that interact with F-actin (Chen, Y. et al.,

2003); it also has the dominant nuclear targeting signal (Wulfkuhle, J. D. et al., 1999)

(Figure 5).

Figure 5 – Schematic showing supervillin functional domains. The N-terminus contains one myosin IIA binding domain (blue) and three F-actin-binding domains (red). The dominant nuclear localization signal (NLS) is shown in green. The carboxy terminus has gelsolin repeats and comprises of a villin/gelsolin homology domain. The amino acid numbers have been mentioned on the scale starting from the amino terminus.

Archvillin (~250 kDa) is an isoform of supervillin which is formed as a result of alternative splicing and is mainly found in the costameres of the muscle cells, where it might be one of the first proteins to assemble during myogenesis and contribute to the myogenic membrane structure (Oh, S. W. et al., 2003). Supervillin is also implicated in various carcinoma cell lines including HeLa cervical carcinoma, lung carcinoma and

adenocarcinoma (Pestonjamasp, K. N. et al., 1997). Supervillin also affects the tumor suppressor protein, p53, by inversely regulating its levels in the cells. Therefore, supervillin increases the cell survival by suppressing the p53 protein (Fang, Z. et al.,

2013). Supervillin has recently been found to interact with Rac1-GEF Trio which might be involved in the regulation of Rac/Rho pathways during cell spreading (Son, K. et al., 2015).

Current studies in our lab focus on the location and function of supervillin in the hair cells of zebrafish and mice and its possible role in the hair cell development and maintenance.

Supervillin in the vertebrate hair cells

(Experiments conducted by Lana M. Pollock for her doctoral thesis)

RNA-seq analysis was done on chicken hair cells to examine the transcriptome and identify the candidate proteins that might interact or associate with actin. SVIL mRNA was found to have a higher RPKM (reads per kilobase per million mapped reads) value as compared to the genes that have already been shown to be expressed in the hair cells, like protocadherin 15 (PCDH15) (Alagramam, K. N. et al., 2001). This information was used to further investigate the expression of supervillin in vertebrate hair cells.

RT-PCR was done using RNA from zebrafish and mouse hair cells to confirm the expression of supervillin. Due to a genome duplication event in a teleost ancestor during evolution, zebrafish possess two or more paralogs of mammalian genes (Robinson-

Rechavi, M. et al., 2001). Previous studies in our lab identified four supervillin genes in zebrafish – svila, svilb, svilc, and svild. Gene svila (mRNA accession number

XM_009306580.1) was present on chromosome 12 and had 56.94% identity with

human SVIL (mRNA accession number XM_011519633.1). Gene svilb (mRNA accession number XM_009298867.1) was present on chromosome 2 and had 53.22% identity with human SVIL. Gene svilc (mRNA accession number XM_001344915.5) was present on chromosome 3 and had 41.14% identity with human SVIL. Gene svild

(mRNA accession number XM_009306541.1) was present on chromosome 12 and had

43.16% identity with human SVIL. All four supervillin paralogs mentioned above were found to be expressed in zebrafish whole maculae using RT-PCR, out of which svila and svilc were detected in zebrafish hair cell cDNA. Paralogs svila (XM_009306580.1) and svilc (XM_001344915.5) were found to have 64% identity. Svil was detected in mouse hair cell cDNA as well. To further test the svila and svilc expression, mRNA in situ hybridization was performed on 4 dpf (days post-fertilization) zebrafish embryos.

Both paralogs were found to be expressed in the anterior macula of the zebrafish ear

(Figure 6) (hybridization experiments conducted by Xi Chen).

Figure 6 – Images from whole mount in situ hybridization of 4 dpf zebrafish. Anti- sense probes against svila and svilc mRNAs were used. A to C – expression of svila is detected in the anterior macula (AM) of the zebrafish ear (orange arrowhead in A). D to F – expression of svilc is also confirmed in the anterior macula of the zebrafish ear (orange arrowhead in D). Experiments performed by Xi Chen.

Subsequent studies were done to find the exact localization of supervillin in the mouse vestibular and cochlear hair cells using immunolabelling. In mouse vestibular hair cells, supervillin was found to localize specifically in the hair cell cuticular plate (Figure 7).

In mouse cochlear hair cells, supervillin localized to the apicolateral margins of the supporting cells (Deiters’ cells, outer pillar cells, and inner phalangeal cells) (Gulley,

R. L. et al., 1976) in addition to the similar localization in the hair cell cuticular plate

(Figure 7).

The apicolateral localization of supervillin in the supporting cells was further studied using antibodies against the adherens junction marker – -catenin along with supervillin immunolabelling. It was found that supervillin localized on the inside of the supporting cells towards the periphery (Figure 7), which is a region where the circumferential F- actin belt was found (Tilney, L. G. et al., 1980). This observation aligns with previously studied characteristics of supervillin in other cultured cell types, where supervillin promotes F-actin binding and targeting to the focal adhesions (Wulfkuhle, J. D. et al.,

1999). Interestingly, the circumferential pattern of supervillin localization in the supporting cells was not found in the mouse vestibular sensory epithelia.

In this study, I aim to investigate the localization of supervillin (Svila) in zebrafish hair cells using immunolabelling assays and transgenic lines of zebrafish.

Figure 7 – Supervillin localization in the mouse vestibular and cochlear hair cells as observed under a confocal microscope. A and B show SVIL localization in mouse vestibular hair cells. A shows the lateral view of a single hair cell showing supervillin in green co-localizing with the actin in the cuticular plate. B shows supervillin in the cuticular plate from a top-down view; fonticulus (arrowhead) and hair bundles (arrow) have also been noted. C and D show supervillin (green) localization in the cochlear hair cells of organ of Corti. SVIL is found to localize in the cuticular plates of inner hair cells (IHC) and the outer hair cells (OHC1, OHC2, and OHC3). It is also found in the apicolateral margins of the supporting cells such as Deiters’ cells (DC1 and DC2), outer pillar cells (OP), and inner phalangeal cells (IPC). E and F show the exact localization of supervillin (green) in the supporting cells towards the circumferential margins and adjacent to the adherens junctions that have been labelled with antibodies against -catenin (blue), a specific adherens junction marker. Scale bars, 2 m. Experiments performed and imaging by Lana M. Pollock.

Materials and Methods

Zebrafish strains and husbandry

Wild type strain, Tübingen, of zebrafish (Danio rerio) was used along with two stable transgenic lines, Gt(macf1a-citrine)ct68a/+ (Trinh, L. A. et al., 2011) and GFP-fascin 2b

(Chou, S. W. et al., 2011) for all the experiments. Zebrafish were maintained and bred at 28oC using standard procedures (Nüsslein-Volhard, C. et al., 2002). Zebrafish strains were kept with the approval of the Case Western Reserve University (CWRU)

Institutional Animal Care and Use Committee (IACUC) (protocol number 2013-0031).

All experimental protocols were approved by the IACUC at CWRU.

Generation of zebrafish Svila antibody

(By Lana M. Pollock and Proteintech, Inc, USA)

Since commercial zebrafish Svila antibody is not available, a novel rabbit polyclonal antibody was generated against 364-723 amino acids of zebrafish Svila. The corresponding svila gene sequence was amplified by PCR using adult zebrafish maculae cDNA and primers Zf_svila_antigen_F 5ʹ-

AACCCGGGCAAAGCTCCATGGTGAGAGAGCAGGCCAGAG-3ʹ and

Zf_svila_antigen_R 5ʹ-

AAGAATTCTCATACTCGCTCAAGCTGTTGGCTTTCCACAACTTCATTTCCC-

3ʹ. The resultant PCR product was inserted into pCR8/GW/TOPO (Life Technologies,

USA), which was then digested using XmaI and EcoRI. After digestion, the svila fragment was isolated and cloned into pGEX-3X, which was used to express glutathione S-transferase (GST) fusion protein. The cognate protein was then used for the immunization and affinity purification of the novel antibodies (Proteintech, Inc,

USA)

Zebrafish whole-mount immunofluorescence

Zebrafish were bred under standard conditions and the eggs were maintained in fish water with 0.1% methylene blue. The embryos were procured at 4 dpf and fixed in ice- cold fixative, Cytoskelfix (Cytoskeleton Inc., USA) for 10 minutes. The embryos were given four washes of five minutes duration each with 1X PBS. They were subsequently permeabilized with 1.5% TritonX-100 (Sigma, USA) in 5% normal goat serum for one hour. The embryos were then incubated in 5% normal goat serum for blocking overnight on a rotor. The primary antibody, anti-Svila (1:200) was then added to the embryos and incubated overnight on a rotor. After subsequent washes with 5% goat serum, secondary antibody conjugated to a fluorophore, Alexa Fluor 633 goat anti- rabbit IgG (H+L) (1:200) (Invitrogen, USA) was added and incubated overnight on a rotor. Gt(macf1a-citrine)ct68a/+ zebrafish was also labelled with Alexa Fluor 546 phalloidin (Invitrogen, USA) at 1:50 dilution. The embryos were finally washed five times with 5% normal goat serum and stored at 4oC until imaging.

Imaging under confocal microscope

Prepared samples of immunolabeled zebrafish embryos were mounted in Vectashield

(Vector Laboratories, USA) and imaged under a Leica SP8 confocal microscope using a 40X objective and the images were acquired on Leica Confocal Software, Leica,

Germany.

Results

Supervillin localization in GFP-fascin 2b transgenic zebrafish

Supervillin localization in the cuticular plate of mouse hair cells called into question if the same pattern of localization is observed in vertebrates other than mammals. This could provide insights into whether the protein is evolutionarily conserved. Therefore, a novel polyclonal antibody was generated in rabbit (by Lana M. Pollock) against a

321-amino-acid-long N-terminal fragment of zebrafish Svila.

To determine the localization of supervillin relative to the hair bundle, GFP-fascin 2b transgenic zebrafish were used. GFP-fascin 2b fish express fascin 2b fused to GFP.

Fascin 2b localizes specifically in the stereociliary hair bundle. From immunolabelling experiments on this strain of fish at 4 dpf, it was found that supervillin (Svila) localizes just below the hair bundle in the cuticular plate region (Figure 8).

Figure 8 – Confocal images of a 4 dpf GFP-fascin 2b (red) transgenic zebrafish immunolabeled with anti-Svila (green) showing hair cells in anterior crista (A). Svila is found to be located in the cuticular plate region just below the hair bundles (arrow). The arrowheads mark the hair bundles that are bend or deflected and the asterisk indicates the cuticular plate. B shows a lateral view of a magnified single immunolabeled hair cell in GFP-fascin 2b zebrafish. Scale bars, 2 m.

Supervillin localization in Gt(macf1a-citrine)ct68a/+ transgenic zebrafish

To confirm the localization of supervillin to the cuticular plate in zebrafish hair cells, immunolabelling was performed on Gt(macf1a-citrine)ct68a/+ zebrafish (Trinh, L. A. et al., 2011). Gt(macf1a-citrine)ct68a/+ transgenic zebrafish expresses Acf7a-citrine fusion protein in the hair cells that circumscribes, underlies and enmeshes the cuticular plate.

It is also found to localize around the fonticulus, which is an F-actin free region of the

CP, where the kinocilium inserts (Antonellis, P. J. et al., 2014).

In 4 dpf Gt(macf1a-citrine)ct68a/+ zebrafish, supervillin was found to localize in the cuticular plate and throughout the region that is circumscribed by Acf7a-citrine (Figure

9). The localization was also compared with the phalloidin labelling of the cuticular plate in Gt(macf1a-citrine)ct68a/+ zebrafish, which further confirmed that Svila indeed localizes to the cuticular plate of zebrafish hair cells (Figure 9).

Figure 9 – Confocal images of a 4 dpf Gt(macf1a-citrine)ct68a/+ transgenic zebrafish immunolabeled with anti-Svila (green) in the posterior macula hair cells (A). Acf7a- citrine fusion protein shown in red. B shows phalloidin (green) labelling under similar conditions. Svila localizes in the region circumscribed by Acf7a-citrine in the hair cell cuticular plate (asterisk). C shows a top-down view of a single magnified zebrafish hair cell. Scale bars, 2 m.

Discussion

Mechanosensory hair cells are one of the most specialized cells in vertebrates and their structure is essential to their proper functioning. They are the fundamental units of sound perception as well as balance. The stereociliary hair bundle and the underlying cuticular plate are the structural components of the hair cell, which play a pivotal role in its functioning. Both stereocilia and the cuticular plate are actin-based structures.

Stereociliary deflections are important for sound stimuli to be transduced by a hair cell, and the cuticular plate is essential to hold the hair bundle in place and serve as a strong foundation against the mechanical stress due to the sound waves.

The cuticular plate is composed of a meshwork of F-actin which is held together by other unknown linker proteins (Hirokawa, N. et al., 1982). The composition and precise function of the cuticular plate is poorly understood. Characterization and identification of different molecules and their mechanisms in shaping the cuticular plate is a primary question that needs to be addressed to improve our understanding of this fascinating organelle in hearing. Therefore, to address this question, our lab identified genes that are expressed in the hair cells and might encode for the proteins which interact with F- actin. One of the important candidate proteins we identified was supervillin (SVIL).

Our lab aimed to confirm the presence of supervillin in the vertebrate hair cell and its exact localization. RT-PCR studies confirmed the presence of svil in mouse and zebrafish hair cells. Zebrafish have four supervillin paralogs, out of which, expression of svila and svilc was confirmed in the zebrafish ear using mRNA in situ hybridization.

The exact localization of SVIL in mouse hair cells was determined using immunolabelling. Supervillin was found to localize specifically to the cuticular plate

but not the hair bundle. It was also found to localize in the apicolateral margins of supporting cells such as Deiters’ cells (Figure 10).

The localization of supervillin (svila) was also determined in zebrafish hair cells using immunolabelling. Using transgenic lines GFP-fascin 2b and Gt(macf1a-citrine)ct68a/+, it was confirmed that supervillin indeed localizes specifically to the cuticular plate of the hair cells. This finding proved that the localization of supervillin is the same in non- mammalian animal models, therefore the protein might be evolutionarily conserved.

However, supervillin localization in the supporting cells’ apicolateral margins was only observed in the mouse models, and not in zebrafish suggesting the evolutionary differences (Figure 11).

Actin-binding properties of supervillin (Wulfkuhle, J. D. et al., 1999) and its specific localization to the cuticular plate in both mouse and zebrafish models leads to the conclusion that it might be responsible for cross-linking actin filaments in the CP to give it its structure and rigidity (Figure 10). Interestingly, supervillin also localizes near the circumferential belt and the cell-cell junctions of the supporting cells in the organ of Corti. It is therefore proposed that supervillin might play a role to bundle and stabilize the actin filaments in these junctions. These cellular junctions are important to maintain the integrity of the reticular lamina and to provide strength against the mechanical stress due to sound waves (Bahloul, A. et al., 2009). Moreover, since circumferential F-actin belts in the supporting cells are believed to be involved in scar formation when a hair cell is lost (Hordichok, A. J. et al., 2007), we hypothesize that supervillin might be involved in interacting with F-actin filaments during this process of cytoskeletal rearrangement.

Figure 10 – Schematic for supervillin localization in the organ of Corti and models for its function. A single hair cell (hair bundle not shown) and its supporting cell. Supervillin (red) has been shown to localize in the hair cell CP and the circumferential band (blue) of the supporting cell (Deiters’ cell). B shows a schematic of organ of Corti and the reticular lamina comprising of apical surface of outer and inner hair cells (hair bundles not shown) and head plates of supporting cells (DCs, IPCs and OPs), with supervillin localization shown in green.

Figure 11 – Schematic for supervillin localization in the zebrafish macula in a Gt(macf1a-citrine)ct68a/+ transgenic fish. Acf7a-citrine expression has been shown in red around the fonticulus as well as the cuticular plate. Supervillin localization in the cuticular plate has been shown in green. Supervillin is not found to be localized in the zebrafish supporting cells suggesting its evolutionary differences as compared to localization in mouse models.

With the information described above, localization of supervillin to the hair cell cuticular plate has been shown in vertebrates. However, its precise function and its role in the development of hair cells is unknown. Knocking out supervillin in mouse models has been unsuccessful because supervillin is involved in other vital processes and knocking it out is lethal. However, since zebrafish have four paralogs of supervillin, this disadvantage can potentially be overcome by knocking out hair cell specific paralogs.

Our lab has generated a zebrafish line with an 8-bp deletion in exon 6 of svila using

TALEN (transcription activator-like effector nuclease). However, preliminary analysis in this mutant fish have not shown any defects in the hair cells. To confirm the complete absence of Svila in the hair cells, techniques such as Western blotting need to be done.

It might also be possible that the normal phenotype was rescued either by the alternative

splice variant of Svila or some other paralogs, and therefore the mutant zebrafish did not show any expected defects.

Further studies in the lab aim to answer the above-mentioned questions as well as to determine the domains of Svila that might be involved in the hair cells and their supporting cells. Analysis of this protein may help us understand the structure of the

CP and its role in hair bundle development and hearing.

Part II

Identification of small molecules that impact the innervation of the lateral line system of developing

zebrafish

Introduction

The optical transparency of larval zebrafish, combined with its ex utero and rapid development, small larval size, its diverse genetic toolkit, and the simplicity of the lateral line system, makes it one of the best models for in vivo small molecule screens.

Around two decades ago researchers realized the potential of zebrafish as a model organism for large-scale forward genetic screens (Eisen, J. S., 1996). Classic genetic screens have been fruitful but with certain drawbacks such as embryonic lethality due to mutations. Therefore, conditional approaches such as chemical screens are employed to bypass the early requirements of developmental processes. After the first chemical screen in 2000 (Peterson, R. T. et al., 2000), the use of zebrafish for similar screens has seen a steep rise in the field of drug discovery. In this particular study, I designed a chemical screen on a transgenic line of zebrafish (HGn39D) that specifically expresses

GFP in its lateral line afferent neurons and documented various phenotypes. The afferent innervation of the zebrafish lateral line serves a model for the peripheral nervous system and is utilized in this screen for the identification of the compounds that impact the peripheral nervous development and organization. These compounds might not only help us gain insights to defects such as peripheral nerve damage, but also help us gain a deeper understanding of the sensory perception by the lateral line system and its applications in hearing research.

Lateral line system in zebrafish

Superficial sensory organs are distributed over the body of all the vertebrates, for instance, mammals have Pacini corpuscles to sense pressure and Meissner corpuscles to sense light touch. Zebrafish, like other fish and amphibians, have a specialized

superficial lateral line system that senses the changes in the motion of water (Figure

12). It helps the fish in detection of prey, avoiding predators and obstacles, surface feeding, and social behaviours such as school swimming and sexual courtship

(Partridge, B. L. et al., 1979). The units of lateral line system responsible for its functions are sensory patches called neuromasts, which comprise of a cluster of mechanosensory hair cells (Figure 12).

Figure 12 – Schematic showing the zebrafish lateral line system and individual neuromast. A depicts a lateral view of a larval zebrafish showing individual neuromasts (green dots) on the anterior and posterior lateral line system. The anterior lateral line (ALL) and posterior lateral line (PLL) ganglia are shown in red. B depicts a lateral view of an individual neuromast, which comprises of clusters of mechanosensory hair cells (green), their supporting cells (orange) and mantle cells (red). Afferent nerve fibers innervating the hair cells are shown in black. Image B has been adapted from Chiu, L. L. et al., 2008.

Neuromasts have a core of hair cells, surrounded by supporting cells and mantle cells

(Figure 12, B). They are innervated by sensory neurons from each major branch of the lateral line, which localize in a cephalic ganglia (Metcalfe, W. K., 1985). There are two sets of hair cell populations in each neuromast – both populations equal in number and comprised of hair cells that have stereocilia oriented at 180o relative to each other

(Ghysen, A. et al., 2007). The neuromasts on the head form the ALL (anterior lateral line) system and the neuromasts in the trunk and the tail form the PLL (posterior lateral line) system. The ALL ganglion is located between the eye and the ear and PLL ganglion is located just posterior to the ear (Figure 12, A).

Hair cells in the neuromast show a characteristic planar cell polarity (PCP) wherein the kinocilium and the stereocilia are arranged in a specific fashion in both of the populations of the hair cells, which are oriented in the anterior-posterior (AP) axis. This

AP axis of morphological polarity of the stereocilia detect the direction of the excitability which enables the zebrafish to determine the vectorial component of a mechanical stimulus (López-Schier, H. et al., 2004). The two populations of the hair cells have opposite orientations and are equal in number (Figure 13). These two oppositely oriented hair cells arise from a single precursor cell, which develop to form two cells having mirror symmetry. Mirror symmetry between these two populations is perpendicular to the body axis (Figure 13).

Figure 13 – Schematic to depict planar cell polarity and mirror symmetry in the hair cells of a neuromast. A single neuromast has been magnified to show hair cells and their arrangement along the AP axis. The void in each hair cell represents the fonticulus, where the kinocilium inserts. It marks the location where the tallest row of stereocilia will be present, and the stereocilia will be graded from tall to short as we go further from the fonticulus. Two oppositely oriented populations of hair cells are shown and the axis has been depicted as the red dotted line. The hair cells anterior to the line are polarized posteriorly and hair cells posterior to the line are polarized anteriorly.

Development of the posterior lateral line (PLL) system in zebrafish

The onset of posterior lateral line development is marked by a migrating primordium

(Figure 14) that originates from the cephalic placodes and deposits the neuromasts as the lateral line develops and progresses towards the posterior end (Stone, L. S., 1922).

This primordium originates from a placode just posterior to the otic placode and follows a pre-existing pathway along the horizontal myoseptum (Metcalfe, W. K. et al., 1985).

Migration of the PLL primordium is a form of chemotaxis that entirely depends upon the interaction between the CXCR4 receptor and its ligand, stromal-derived factor 1

(SDF1) (David, N. B. et al., 2002). This particular ligand-receptor pair is also involved in other long distance migrations such as migration of germ cells, lymphocytes, and metastases (Ghysen, A. et al., 2004).

The migration of primordium begins at about 20 hpf (hours post fertilization) and completes at about 40 h. At the onset of migration, primordium consists of a group of cells measuring ~7 m each to form a broad stripe right under the epidermis. This primordium is around 4-5 cells wide and 20-25 cells long and usually extends over 2-3 consecutive somites (Figure 14). The migration starts by the end of somatogenesis in the zebrafish embryo (Kimmel, C. B. et al., 1995). Each cell moves independently of

other cells in the primordium, but it appears as a collective migration of a block of cells

(Haas, P. et al., 2006). The number of cells that make up the primordium vary over the course of lateral line development. Before depositing its first proneuromast (L1), the primordium consists of around 100 or more cells. This number depletes as the primordium progresses and starts depositing proneuromasts, which are comprised of around 20 cells each. Over the course of its migration, the primordium deposits seven to eight proneuromasts which develop into fully functional neuromasts in 6 hours

(Gompel, N. et al., 2001). The neuromasts are numbered from L1 to Ln (n = number of neuromasts) from anterior to posterior end, in order of their appearance. Proneuromasts

L1 to L5 are deposited at regular intervals on the horizontal myoseptum when a group of primordium cells slow down their own migration and eventually break off from the leading group of cells that continue to migrate at the same pace (Figure 14). The deposition of terminal proneuromasts is different as it occurs below the level of the myoseptum and two proneuromasts form simultaneously (Raible, D. W. et al., 2000).

Therefore, by 40 h, the primordium migration is complete in the PLL system and the larval zebrafish has ~5 neuromasts on the trunk (L1 to L5) and 2-3 terminal neuromasts on the caudal end (ter). Transition from these few neuromasts in the larval zebrafish to multiple neuromasts in the adult zebrafish was initially thought to be because of budding from existing embryonic neuromasts. However, it has been proven that new primordia are generated and they fill the gaps in somite borders until all of them have been occupied by one neuromast (Ledent, V., 2002).

Figure 14 – Posterior lateral line primordium migration in a Zebrabow transgenic line. The primordium has been marked by a dotted white line and the anterior- posterior axis has been shown in the first panel. The migration of primordium occurs towards the posterior end. A group of about 20 cells slow down and deposit a proneuromast at the trailing edge of the primordium. The primordium’s leading edge keeps migrating posteriorly leaving a trail of interneuromast cells in its wake, as seen in last panel. A cell marked with an asterisk has slowed down and has been displaced anteriorly as it moves out of the leading edge. A cell marked with a plus (+) sign becomes relatively immobile as it has been deposited in a proneuromast. Image adapted from Thomas, E. D. et al., 2015.

Innervation of the lateral line system in zebrafish

Hair cells in the neuromasts are innervated by the afferent neurons that arise from the lateral line ganglia. It has been shown in the amphibians that the growth cones of sensory neurons accompany the primordia as it starts migrating and that establishes a physical contact between the cranial ganglion and the neuromasts (Harrison, R. G.,

1903). In zebrafish, the early primordium is found adjacent to the developing sensory ganglion in the postotic region. Growth cones of PLL sensory neurons are found in the early primordium even before it starts migration. Growth cones of these sensory

neurons are later found within the primordium as it migrates along the horizontal myoseptum (Metcalfe, W. K. et al., 1985). Sensory axons have been found to be guided on their course of migration by the repulsive effect of sema3A expression in the dorsal and ventral side of the pathway, independent of primordium migration (Shoji, W. et al.,

1998). However, sensory neurons still follow the migration of the primordium and would stall if the primordium were immobilized (David, N. B. et al., 2002), proving that sensory axons are guided by the migrating primordium (Harrison, R. G., 1903).

This association for comigration might depend on the expression of various proteins such as HNK-1 by the PLL neurons and surface proteins such as Tag1 (Warren, J. T. et al., 1999). The lateral line sensory neurons coalesce in a cephalic ganglion, which projects the fibers into the rotrocaudal column of the hindbrain (Pujol-Martí, J. et al.,

2013) (Figure 15). Anterior lateral line (ALL) and posterior lateral line (PLL) have separate ganglia and both of them send central axons into the hindbrain (Figure 15).

The PLL ganglion forms due to the expression of a proneural gene called neurogenin-

1 (ngn1) (Andermann, P. et al., 2002) and inactivation of this gene prevents the formation of PLL ganglion completely; however, the neuromasts still develop independently of innervation.

Figure 15 – Schematic depicting the ALL and PLL ganglia in red and blue respectively along with their afferent projections in the same color projecting into the hindbrain and from the neuromasts (purple) (Wada, H. et al., 2015).

Glial cells ensheath the PLL nerve and are thought to originate from both the neural crest cells as well as the primordium (Ghysen, A. et al., 2004). The glial cells migrate along the sensory neurons and myelinate the PLL nerve and maintain its fasciculation

(Gilmour, D. T. et al., 2002).

The HGn39D transgenic zebrafish

Lateral line afferent neurons in the zebrafish can be visualized either by using molecular markers on a fixed sample or by injecting afferent fibers with neuronal tracers

(Gilmour, D. et al., 2004). However, these methods are not useful for long-term imaging in live specimen. In 2008, a group led by Koichi Kawakami performed a transposon mediated large-scale enhancer-trap screen and created 73 zebrafish lines that were insertional mutants (Nagayoshi, S. et al., 2008). One of the zebrafish transgenic lines generated through this insertional mutagenesis was HGn39D, which expresses EGFP (enhanced green fluorescent protein) in all the afferent neurons of the lateral line system (Faucherre, A. et al., 2009). It also expresses strong fluorescence in the lens of the eye (Figure 16). The transgene integration site in HGn39D is a single insertion on chromosome 24 within a locus that encodes for zebrafish homolog of contactin associated protein-like 2/Caspr2 (Cntnap2a) (Pujol-Martí, J. et al., 2012).

This protein belongs to the neurexin family of neuronal cell-adhesion proteins that are implicated in human disorders such as autism and epilepsy (Peñagarikano, O. et al.,

2011). In 2009, Faucherre, A. et al. proved that HGn39D enhancer-trap line of zebrafish specifically expresses EGFP in the anterior lateral line (ALL) ganglion, posterior lateral

(PLL) ganglion, axonal projections forming the rostrocaudal column in the hindbrain, afferent axonal fibers along the entire trunk of the zebrafish, and the dendritic arbors

with thin extensions which contact each hair cell in the neuromasts (Figure 16).

Collectively, this transgenic line expresses EGFP in the whole population of lateralis afferent neurons (Faucherre, A. et al., 2009).

Figure 16 – Expression of EGFP by the HGn39D transgenic line of zebrafish in the afferent neurons of the lateral line system. A shows GFP expression in a 4 dpf HGn39D zebrafish (Scale bar – 100 m) from the lateral view. GFP is expressed specifically in the ALL and PLL ganglia (dashed boxes), PLL axon throughout the trunk, and the dendritic arbors that contact each hair cell in a neuromast. White arrowhead points to the neuronal projection into the hindbrain and the yellow arrowhead points to the neuronal projections in the neuromasts (dendritic arbor). Image adapted from Faucherre, A. et al., 2009. B shows a schematic of HGn39D fish to highlight the organs with GFP expression; a single dendritic arbor has been zoomed in for clarity. Each neuronal extension from the arbor makes contact with an individual hair cell of the neuromast.

In vivo chemical screening in zebrafish

The potential of zebrafish as a model organism for in vivo chemical screening was not realized until the year 2000, prior to which the small molecule drug discovery was based on trial-and-error testing on cell lines and other less favorable animal models. In the year 2000, a study by Randall T. Peterson demonstrated how chemicals could be added simply to a 96-well plate containing larval zebrafish in their aquatic environment

(Peterson, R. T. et al., 2000). The plates can then be incubated to allow the absorption of chemicals into the fish, after which they can be screened visually or automatically under a microscope.

Zebrafish are vertebrates like humans, and despite having evolutionarily diverged around 450 million years ago, the similarity between their genes is very high (Barbazuk,

W. B. et al., 2000). Zebrafish have similar pharmacokinetics and efficacy as humans, which makes it a reasonable model for drug discovery. Zebrafish are also best suited for high-throughput phenotyping which is not possible with any other vertebrates (Zon,

L. I. et al., 2005). Zebrafish embryos are transparent which facilitates detection of functional and morphological changes without having to dissect the organism. The scale of the chemical screen using zebrafish embryos is very high because of the small embryo size (less than 1 mm) coupled with great number of eggs (~300) laid by each female at a time. In addition to this, the fertilization in zebrafish is ex utero, which makes it possible to study drug interactions and its effects in a developing embryo

(Taylor, K. L. et al., 2010). These advantages added together can explain why zebrafish are the model organisms of choice when it comes to high throughput screening (HTS)

(Delvecchio, C. et al., 2011). Several molecules that have been shown to be effective in the mammalian models owe their discovery to zebrafish chemical screens

(Rennekamp, A. J. et al., 2015).

There are certain considerations while using zebrafish for chemical screening, most importantly, the choice of chemical library which directs the molecular processes that are disrupted as well as identification of its mechanism of action (Wheeler, G. N.et al.,

2009). Most of the chemical screens performed on zebrafish till date have used biologically active compounds (Patel, D. V. et al., 1996). It is also important to consider the number of embryos per well in a HTS to ensure the phenotypic certainty which might be compromised if the wells are too crowded. The readout also plays an important role while designing a chemical screen because manual visualization of phenotypes might not be feasible and can be error-prone (Peal, D. S. et al., 2010). Taking these considerations into account, the last 15 years have seen a steep increase in use of zebrafish in the field of chemical screening and drug discovery (Rennekamp, A. J. et al., 2015). Chemical screens using zebrafish have led to important discoveries in fields ranging from cardiology (Burns, C. G. et al., 2005) to cancer research (Amatruda, J. F. et al., 2002).

In this study, I have designed a small-molecule screen on the HGn39D transgenic line of zebrafish to visually examine the phenotypes impacting the afferent neurons.

Transgenic lines of zebrafish account for 35% of the chemical screens, whereas 51% of screens use wild-type fish; the rest are performed on mutant fish (Rennekamp, A. J. et al., 2015). Previous studies by our group and other labs have focused on the lateral line system to screen drugs that might prevent or cause hearing loss (Ou, H. C. et al., 2010).

This study particularly focuses on the afferent innervation of the lateral line system and phenotypic effects of small molecules from the chemical library.

Materials and Methods

Zebrafish strain and husbandry

Transgenic zebrafish line HGn39D was maintained and bred at 28 oC using standard procedures (Nüsslein-Volhard, C. et al., 2002) in the laboratory fish facility. Eggs were collected, cleaned, and maintained in the embryo buffer (EB) [EB (pH 7.0 - 7.2) - 7.5 mM NaCl, 0.25 mM KCl, 0.5 mM MgSO4, 0.075 mM KH2PO4, 0.025 mM Na2HPO4,

o 0.5 mM CaCl2, and 0.35 mM NaHCO3] at 28 C in an incubator. An average of 70 embryos were kept in a standard petridish. The HGn39D transgenic line of zebrafish was generated by insertional mutagenesis using an enhancer-trap gene in a large-scale genetic screen by Kawakami lab (Nagayoshi, S. et al., 2008). This strain expresses

EGFP specifically in the afferent neurons of the lateral line system in the zebrafish

(Faucherre, A. et al., 2009). Zebrafish strains were kept with the approval of the Case

Western Reserve University (CWRU) Institutional Animal Care and Use Committee

(IACUC) (protocol number 2013-0031). All experimental protocols were approved by the IACUC at CWRU.

Chemical library

The chemical library comprised of 5,040 chemicals was obtained from the Drug

Discovery Center at University of Cincinnati. These chemicals were selected from a

50K Diversity Screening Library (Drug discovery core, University of Cincinnati) based on their unique structures that have been predicted using bioinformatics to have good oral bioavailability. The mean molecular weight of the chemicals is 344.5 Da. 1,040 chemicals were randomly selected from this chemical library for this study. The

chemicals were dissolved in dimethylsulfoxide (DMSO) to a concentration of 10 mM and then stored frozen in 96-well polypropylene plates at -80 oC.

Dechorionation of zebrafish embryos

The migration of the primordium along with the afferent neuronal growth cones begins approximately around 20 hpf. Therefore, it is important to subject embryos to the chemicals during this time to obtain optimum phenotypic outcome. However, the embryos at this stage are surrounded by an acellular envelope called the chorion. The effectiveness of chorion to serve as a barrier is not yet completely understood but in order to avoid the risk of generating false negatives, the chorion should be removed and the ‘naked’ embryos should be exposed directly to the chemicals (Henn, K. et al., 2011).

After procuring eggs at 24 hpf, enzyme-based dechorionation was carried out using 1X pronase (Sigma, USA) (protease from Streptomyces griseus) diluted in embryo buffer.

The embryos were incubated with pronase for about 5 minutes with occasional agitation, due to which the chorions start to come off. The embryos were then washed thoroughly multiple times with embryo buffer to remove excess pronase. The process of dechorionation is harsh on the young embryos and therefore the survival rate is considerably less than the embryos that have not been dechorionated.

Primary chemical screen and rescreen protocol

The dechorionated embryos at 24 hpf were transferred into the fresh embryo buffer.

Three undamaged embryos were then carefully transferred into each well of a glass- bottom 96-well plate (P96G-0-5-F; MatTek) along with 240 l of embryo buffer. Each

96-well chemical screen plate was comprised of 80 wells containing chemicals and 10 wells containing negative control. The chemicals from the chemical library (10 mM concentration) were diluted to a final concentration of 20 M using 0.2% DMSO. The negative control was comprised of 0.2% DMSO in embryo buffer. 60 l of the diluted chemicals (20 M) and negative control were then added to the chemical screen plate containing three embryos in each well. The final volume in each well of the glass- bottom chemical screen plate was 300 l (Figure 17). The embryos treated with chemicals in the 96-well glass-bottom plate were then incubated at 28 oC until they were 72 hpf. Occasional cleaning using a Pasteur pipette was done to get rid of any dead embryos.

Imaging and phenotype analysis

At 72 hpf, the embryos were anesthetized using 1X tricaine (3-aminobenzoic acid ethyl ester methanesulfonate) (Sigma, USA). Most of the liquid from the 96-well screening plate was drawn out prior to microscopic observation. Flushing using the pipette was done to orient the embryos on their lateral side to facilitate observation. DMI6000

(Leica, Germany), inverted fluorescent microscope was used for all imaging purposes to observe the phenotypes (Figure 17). Samples that showed altered phenotypes were considered possible ‘hits’. The ‘hits’ from the primary screen were subjected to additional rounds of screening to ensure reproducibility.

Results

The aim of the study is to identify small molecules that could alter the phenotype in the

HGn39D transgenic line of the zebrafish. A total of 1,040 chemicals were tested using the procedure described above and depicted below in figure 17.

Figure 17 – Schematic to show the design of the chemical screen. The transgenic HGn39D zebrafish are bred and their embryos are dechorionated (not shown). The chemicals from the chemical library are diluted to a final concentration of 20 M. In a glass-bottom 96-well screening plate, three dechorionated embryos are added along with the diluted chemical. The embryos are incubated with the chemical until they are 72 hpf, after which they are screened under a confocal microscope for phenotypic alterations. The chemicals that cause phenotypic alterations are considered potential ‘hits’ and are rescreened using the same procedure.

The chemicals were added to the screening plate when the embryos were 24 hpf and observations were made under the fluorescent microscope at 72 hpf. During this time the chemical was absorbed by the zebrafish embryos, which caused possible phenotypic alterations due to the pharmacological effects on the lateral line afferent neurons. A total of 63 hits were identified using this study after the primary screen and the rescreening. These 63 hits were chemicals that caused phenotypic alterations in the lateral line afferent neurons in HGn39D transgenic zebrafish. The phenotypes observed were grouped into 12 categories depending upon the type of alterations (Table 1).

Table 1 – ‘Hits’ obtained from the small molecule screen. A total of 63 ‘hits’ were obtained and they have been grouped into categories depending on their similar phenotype. The number of ‘hits’ exhibiting a particular phenotype has also been mentioned.

Number of ‘hits’ Serial Plate/well number for the Phenotype with a particular Number ‘hits’ phenotype 1. 1600000216/B5 2. 1600000216/G9 3. 1600000207/B5 4. 1600000243/F4 Severely 5. 1600000243/A10 underdeveloped lateral line afferent 6. 1600000200/A10 neurons; ALL and 7. 1600000205/C8 PLL ganglia seem disintegrated with a 14 8. 1600000232/D2 few scattered cells. 9. 1600000232/G4 Lateral line axon is either completely 10. 1600000202/B10 absent or 11. 1600000213/A5 undeveloped. 12. 1600000213/G5 13. 1600000213/F10 14. 1600000224/A10 15. 1600000216/B1 16. 1600000243/G8 Hair cell-like 17. 1600000200/F2 structures seen on the dendritic 18. 1600000200/H3 arbors. The afferent 19. 1600000203/B1 neurites that contact each hair 10 20. 1600000203/G9 cell in a neuromast 21. 1600000232/F2 appear to have hair cell-like, GFP 22. 1600000232/H6 expressing 23. 1600000224/D7 structures. 24. 1600000213/C10 25. 1600000205/A7 The afferent axon 26. 1600000205/G9 in the posterior lateral line appears 27. 1600000205/G10 to be wavy as compared to the 9 28. 1600000207/A1 normal axon that follows a straighter 29. 1600000232/E3 path during migration. 30. 1600000196/D5

31. 1600000213/B4

32. 1600000203/G4

33. 1600000203/D5

34. 1600000216/D3 The sensory 35. 1600000207/D6 afferent axon in the lateral line appears 36. 1600000200/D7 5 to terminate before 37. 1600000202/E6 reaching the base of the caudal fin. 38. 1600000224/B7 39. 1600000216/D1 40. 1600000205/F6 The sensory afferent axon 41. 1600000205/D4 fascicles in the PLL 6 42. 1600000243/G5 appear to be thinner than the normal 43. 1600000232/G5 fascicles. 44. 1600000224/D9

The afferent axon 45. 1600000216/A1 is wavy and terminates 2 prematurely before reaching the base 46. 1600000243/B3 of the caudal fin.

47. 1600000216/A8 The zebrafish exhibit an abnormal 48. 1600000216/A9 morphology such 4 as abnormal 49. 1600000243/A9 curvature in the body. 50. 1600000196/A6

51. 1600000216/G4 Bright fluorescence in the somites on 52. 1600000207/A3 3 the dorsal side of 53. 1600000196/F3 the zebrafish.

54. 1600000207/D5 The fish show an abnormal 55. 1600000207/A7 3 morphology of the 56. 1600000203/G5 heart.

Fluorescent cell- like structures were 57. 1600000200/G2 found circulating in 1 the body of the zebrafish

Fluorescent patches which appear to be GFP expressing 58. 1600000213/F7 1 cells are found on the entire body of the zebrafish.

59. 1600000235/A1 60. 1600000216/B9 The larval zebrafish 61. 1600000196/F5 lacked 5 pigmentation. 62. 1600000202/E10 63. 1600000213/H4

Phenotype 1

The embryos treated with chemicals from wells B5 and G9 from plate 1600000216; well B5 from plate 1600000207; wells F4 and A10 from plate 1600000243; well A10 from plate 1600000200; well C8 from plate 1600000205; wells D2 and G4 from plate

1600000232; well B10 from plate 1600000202; wells A5, G5, and F10 from plate

1600000213, and well A10 from plate 1600000224 showed similar phenotypes with severely underdeveloped afferent neurons of the lateral line. The ALL and PLL ganglia were either completely absent or were disintegrated with very few scattered cells. The lateral line axon was absent or underdeveloped which means there was no growth cone migration during development towards the caudal end of the fish (Figure 18). It has been shown that the development of the cranial sensory ganglia depends upon the expression of the proneural gene called neurogenin-1 (ngn1) and inactivation of this gene leads to a phenotype that lacks both the cranial ganglia (Andermann, P. et al.,

2002). Therefore, it might be possible that the ‘hits’ that produce a phenotype where the ALL and PLL ganglia are either absent or severely underdeveloped, are altering the expression of this proneural gene leading to a compromised phenotype.

Figure 18 – Phenotype with poor afferent neuronal development. The embryos have disintegrated ALL and PLL ganglia with scattered cells and almost no axonal migration towards the caudal end of the zebrafish. A (1600000232/D2), B (1600000216/G9), and C (1600000232/G4) show the observed phenotype wherein the cells of the ALL ganglion (yellow arrowhead) and PLL ganglion (red arrowhead) seem to be scattered. D shows the negative control with much more compact appearance of both the ganglia. Eye of the fish also expresses GFP in the HGn39D line of zebrafish (asterisk). Scale bar 100 m.

Phenotype 2

The embryos treated with chemicals from well B1 from plate 1600000216; well G8 from plate 1600000243; wells F2 and H3 from plate 1600000200; wells B1 and G9 from plate 1600000203; wells F2 and H6 from plate 1600000232; well D7 from plate

1600000224, and well C10 from plate 1600000213 exhibited a phenotype wherein the dendritic arbors ended in the hair cell-like structures. HGn39D specifically expresses

GFP in the afferent neurons of the lateral line system; however, it does not express GFP

in the hair cells (Faucherre, A. et al., 2009). In this phenotype, hair cell-like structures appear on the ends of afferent neurites that are supposed to innervate the hair cells in the neuromasts (Figure 19). A closer observation under a confocal microscope revealed that these cell-like structures appear very similar to the hair cells in the neuromast

(Figure 20); however, the hair cells in the HGn39D transgenic zebrafish do not show the GFP expression. Therefore, it might be possible that the ‘hits’ exhibiting this phenotype alter the promoter so that the GFP is expressed in the lateral line hair cells.

Figure 19 – Phenotype with hair cell-like structures at the end of dendritic arbors. HGn39D fish express GFP specifically in the afferent neurons of the lateral line system but these chemicals have altered the phenotype wherein hair cell like structures are visible along with afferent neurites (red arrowheads). ALL and PLL ganglia in A are marked with white arrowheads. A (1600000216/B1) and B (1600000203/G9) show a 10 ╳ view of the lateral line system whereas C (1600000200/F2) shows 20 ╳ magnification of a single altered phenotype showing hair cell-like structures. D shows the negative control with normal dendritic arbors marked with white arrows. Eye of the fish has been marked with an asterisk. Scale bar 100 m.

Figure 20 – Confocal micrographs of the phenotype showing the hair cell-like structures. A and B show the hair cell-like structures (marked with yellow arrowheads) as observed on the dendritic arbors of the posterior lateral line system of a 72 hpf HGn39D zebrafish treated with the chemical from the plate1600000224, well D7. These structures appear very similar to the hair cells in the neuromast because of their appearance and presence of what seems to be a nucleus (asterisk). Scale bar 3 m. Images acquired by Zongwei (Kevin) Chen.

Phenotype 3

The embryos treated with chemicals from wells A7, G9, and G10 from plate

1600000205; well A1 from plate 1600000207; well E3 from plate 1600000232; well

D5 from plate 1600000196; well B4 from plate 1600000213, and wells G4 and D5 from plate 1600000203 exhibit a phenotype wherein the afferent axon in the lateral line system is wavy as compared to a normal axon in an HGn39D phenotype (Figure 21).

In a normal HGn39D phenotype, sensory axons follow a straighter pathway because the axonal guidance is restricted by the repulsive effect of sema3A expression on the dorsal and ventral side of the constrained pathway (Shoji, W. et al., 1998). Therefore,

it might be possible that the ‘hits’ exhibiting this phenotype alter the sema3A expression so that the axonal guidance is less constrained leading to a wavy pathway of the lateral line afferent axon.

Figure 21 – Phenotype with the wavy sensory afferent axon in the lateral line system. A (1600000205/G9), B (1600000232/E3), and C (1600000203/D5) show sensory axon that seems to be kinked in certain places marked with yellow arrowheads. C shows a magnified view of an undulating afferent axon. The dendritic arbors of afferent neurites have been encircled. D shows the negative control with a much straighter axon. Scale bar 100 m.

Phenotype 4

The embryos treated with chemicals from well D3 from plate 1600000216; well D6 from plate 1600000207; well D7 from plate 1600000200; well E6 from plate

1600000202, and well B7 from plate 1600000224 show a phenotype where the sensory afferent axon of the PLL system ends prematurely before it reaches the base of the

caudal fin of the zebrafish. In a normal phenotype, the afferent axon terminates at the caudal tip forming dendritic arbors for the terminal neuromasts (Figure 22).

Figure 22 – Phenotype wherein the sensory afferent axon in the lateral line system terminates prematurely before reaching the base of the caudal fin of the fish. A (1600000216/D3), B (1600000207/D6), and C (1600000224/B7) show the sensory axon that has terminated before reaching the base of the caudal fin (end-point marked with a red arrowhead). The normal phenotype of the HGn39D transgenic fish has been shown as a negative control in D; the afferent axon terminates at the caudal tip where it forms sensory dendritic neurites for terminal neuromasts (shown in red ellipse). It can also be seen that the afferent axon takes a slightly ventral course when it is about to terminate. The lateral line dendritic arbors have been circled and the asterisk marks an additional out-of-focus zebrafish in the view. Scale bar 100 m.

Phenotype 5

The embryos treated with chemicals from well D1 from plate 1600000216; wells F6 and D4 from plate 1600000205; well G5 from plate 1600000243; well G5 from plate

1600000232, and well D9 from plate 1600000224 show a phenotype where the sensory axon fascicles in the posterior lateral line appear to be thinner than normal fascicles. In a normal phenotype, the sensory axons are fasciculated to give it a fuller appearance than observed in the phenotypes obtained from these hits (Figure 23).

Figure 23 – The phenotype where the sensory afferent fascicle in the posterior lateral line system appears to be thinner than the normal. A (1600000205/F6), B (1600000243/G5), and C (1600000232/G5) show the thin sensory afferent axon (marked with orange arrowheads). D shows the negative control where the afferent axon in the trunk of the fish seems to have normal thickness due to the axonal fibers fasciculation. The lateral line dendritic arbors have been circled and the yellow arrowhead in C marks the posterior lateral line ganglion. Scale bar 100 m.

Phenotype 6

The embryos treated with chemicals from well A1 from plate 1600000216, and well B3 from plate 1600000243 exhibit a phenotype which appears to be the combination of two phenotypes – the embryos have a wavy sensory afferent axon in the posterior lateral line which also terminates prematurely before reaching the base of the caudal fin of the zebrafish (Figure 24).

Figure 24 – Phenotype where the sensory afferent axon in the posterior lateral line system is wavy and ends prematurely before reaching the caudal tip of the zebrafish. A (1600000216/A1), B (1600000216/A1), and C (1600000243/B3) show the wavy afferent axon (marked with yellow arrowhead) and its premature termination (marked with a red arrow). D shows the negative control where the afferent axon has normal appearance and ends at the caudal tip forming afferent neurites for the terminal neuromasts. The lateral line dendritic arbors have been encircled and the asterisk marks the additional out-of-focus fish in the view. Scale bar 100 m.

In addition to the small-molecule hits that alter the phenotype by modifying the afferent neurons in transgenic HGn39D zebrafish lateral line, certain hits were found to affect the fish as a whole and not just the lateral line afferent neurons. These hits were not directed specifically at the afferent neurons but altered the normal phenotype elsewhere in the body of the zebrafish. These hits have been described in the following section.

Phenotype 7

The embryos treated with chemicals from well A8 and A9 from plate 1600000216; well

A9 from plate 1600000243, and well A6 from plate 1600000196 exhibit abnormal morphology as compared to the normal zebrafish of the same age (Figure 25).

Figure 25 – Phenotype where the zebrafish embryos show abnormal morphology. A (1600000196/A6) shows a zebrafish with an abnormally curved posterior end (yellow arrow). B shows the negative control wherein the zebrafish have a normally straight posterior end. The lateral line dendritic arbor has been encircled. Scale bar 100 m.

Phenotype 8

The embryos treated with chemical from well G4 from plate 1600000216; well A3 from plate 1600000207, and well F3 from plate 1600000196 exhibit bright fluorescence in their dorsally-located somites, also known as chevrons. This phenotype occasionally interfered with the observation of afferent neurons in HGn39D zebrafish (Figure 26).

Figure 26 – Phenotype in HGn39D zebrafish embryos where they show GFP expression or green fluorescence in the somites. A (1600000207/A3) shows an HGn39D zebrafish embryo with fluorescent somites (red arrowheads). B shows the negative control with normal expression of GFP specifically in the afferent neurons of lateral line system including ALL/PLL ganglia, and in the lens of the eye. Scale bar 100 m.

Phenotype 9

The embryos treated with chemicals from well D5 and A7 from plate 1600000207, and well G5 from plate 1600000203 exhibit an abnormal morphology of the heart. The heart of the zebrafish in these cases appeared more tubular than normal (Figure 27).

Figure 27 – Phenotype wherein zebrafish embryos exhibit an abnormal morphology of the heart. A (1600000207/D5) shows a zebrafish that has an abnormal tubular heart. B shows the negative control – a zebrafish which has a heart with normal morphology. The heart has been indicated with red arrows. The yellow arrowheads indicates the ALL and PLL ganglia, and the asterisk indicates the eye of the zebrafish. Scale bar 100 m.

Phenotype 10

The embryos treated with chemical from well G2 from plate 1600000200 exhibit a phenotype wherein certain fluorescent cell-like structures seemed to be circulating in the body of the zebrafish. These cells were only visible in the experimental fish and no such cells were visible in the negative control (Figure 28). Due to the transparency of the larval zebrafish, it was also possible to see these cell-like structures being pumped through the heart of the fish.

Figure 28 – Phenotype showing fluorescent cell-like structures in zebrafish circulation. A (1600000200/G2) shows a ventral view of a zebrafish that has fluorescent cell-like structures circulating in the body (seen as green dots and indicated with red arrowheads). B shows the negative control where no such fluorescent cell-like structures are seen. The yellow arrowheads indicate the ALL and PLL ganglia, and the asterisk indicates the eye of the zebrafish. Scale bar 100 m.

Phenotype 11

The embryos treated with chemical from well F7 from plate 1600000213 exhibit a phenotype where the entire body of the zebrafish larvae is covered with fluorescent patches which appear to be GFP expressing cells on the body surface of the zebrafish.

Under white light, the fish seem to have no pigmentation whereas under a fluorescent microscope, patches of fluorescence are seen on the body surface of the zebrafish

(Figure 29). The GFP negative fish do not show the same phenotype when treated with the same chemical. An HGn39D transgenic zebrafish which has not been treated with this chemical serves as a negative control and does not show such a pattern of fluorescence.

Figure 29 – Phenotype showing fluorescent patches on the body surface of the zebrafish which has been treated with the chemical (well F7, plate 1600000213). A, B, and C (1600000213/F7) show the body surface of the fish which seems to be covered in fluorescent patches (indicated with red arrowheads). D shows the negative control wherein no such fluorescent patches are observed on the body surface. White arrowheads indicate the ALL and PLL ganglia, and the asterisk indicates the eye of the zebrafish. Scale bar 100 m.

Phenotype 12

The embryos treated with chemical from well A1 from plate 1600000235; well B9 from plate 1600000216; well F5 from plate 1600000196; well E10 from plate 1600000202, and well H4 from plate 1600000213 exhibit a phenotype with no pigmentation.

Zebrafish embryos start producing melanophores from their neural crest cells as early as 24 hpf (Lister, J. A., 2002). At 24 hpf, the experimental fish in this particular small molecule screen are subjected to the chemicals. Therefore, it might be possible that these chemicals are inhibiting the formation of melanocytes and therefore the fish lack

pigmentation even after 72 hpf. An important chemical that has been already described in detail for its pigmentation-inhibiting quality is 1-phenyl 2-thiourea (PTU) (Karlsson,

J. et al., 2001). PTU facilitates imaging by making the embryos completely transparent and improving the signal during gene expression experiments or confocal microscopy.

PTU inhibits melanocyte formation by blocking the tyrosinase-dependent steps in the melanin pathway (Karlsson, J. et al., 2001). In this screen, the small molecule hits that produce this phenotype can also be further characterized in the same way to produce a consistent outcome of zebrafish lacking pigmentation.

This small molecule screen on the transgenic HGn39D zebrafish produced 63 potential hits and the phenotypes have been categorized into 12 broad categories. The number of potential hits that produce same phenotype have been grouped together and depicted below in the form of a bar-graph (Figure 30).

Figure 30 – Bar-graph depicting the number of hits obtained in this small molecule screen. The observed phenotypes can be grouped into 12 broad categories that have been listed on the X-axis. The number of hits that manifested the similar phenotype are shown in numbers on the Y-axis.

Discussion

Small molecule screens are a very powerful tool that can be used to understand complex biological systems and identify molecules that affect the development, cause a disease and, the mechanisms behind other biological processes. It also serves as the very first step towards drug discovery. Zebrafish have been historically used for various genetic screens but the last 15 years have seen a rise in exploiting zebrafish for small molecule screens (Rennekamp, A. J. et al., 2015). The first published small molecule screen was carried out by Randall Peterson in the year 2000, wherein he screened 1,100 compounds and observed phenotypes that affected the nervous system, cardiovascular system, and pigmentation (Peterson, R. T. et al., 2000).

Zebrafish exhibit a wide range of phenotypes which can be used to design small molecule screens that modify any of these phenotypes. However, studies to explore the pathway from the phenotype to the mechanism have been cited as one of the most challenging aspects of drug discovery (Rennekamp, A. J. et al., 2013). Once a phenotype has been identified by a small molecule screen, there are four main approaches to identify the targets and understand the mechanism of action (MOA) – comparison based on the structure of the chemical; comparison based on phenotypes exhibited by genetic mutations; comparison based on pharmacological phenocopy, and comparison based on compound affinity (Rennekamp, A. J. et al., 2013).

The most important step towards drug discovery after performing a small molecule screen is the target identification of the small molecule hits. Target identification can be approached using various methods such as structure-activity relationship (SAR) to find similar molecules with known mechanisms, studying previously known binding partners for the small molecule of interest, studying the similarity of chemotype to a

known genetic phenotype, affinity purification of the small molecule binding partners, and a systems biology approach to statistically compare large datasets to categorize them into functional units (Peal, D. S. et al., 2010) (Figure 31).

Figure 31 – Schematic showing different approaches for target identification after a small molecule screen. 1. The target can be compared with the known targets binding to the same binding partners. 2. SAR approach to identify similar molecules whose mechanism of action has already been discovered. 3. Identifying chemicals that give rise to similar phenotypes. 4. Affinity purification of the binding partners of the small molecules. 5. Using a systems biology approach to group the small molecules in similar functional units. Image from Peal, D. S. et al., 2010.

Another challenging field of study after a small-molecule screen is the identification of mechanism of action (MOA) for the potential ‘hits’. Identification of small molecule mechanism of action has led to important discoveries in biology such as opioid

receptors (Peterson, R. T. et al., 2011). The scale of a small molecule screen usually depends on the mechanism of action (MOA) studies. A screen performed in a well characterized area of biology can be relatively small and less time consuming whereas open-ended MOA studies can be done on more diverse chemical libraries (Peterson, R.

T. et al., 2011).

Around 35% of chemical screens that are carried out on zebrafish are performed on different transgenic lines of zebrafish. In this study, I designed a small molecule screen to be performed on a transgenic line of zebrafish called HGn39D, which expresses GFP specifically in the afferent neurons of the lateral line system (Faucherre, A. et al., 2009).

The aim of the study is to find out if the small molecules from the chemical library could induce changes in the phenotype of the HGn39D fish. This phenotypic study is important since the afferent neurons are responsible for relaying the signals from the neuromasts of the lateral line system to the hindbrain in the central nervous system via

ALL and PLL ganglia. Therefore, the altered phenotypes would subsequently modify the sensory-signal processing of the lateral line system. Additionally, this study can be combined with the previous studies in the lab (by Wenbo Chen) where certain chemicals were identified that induced the proliferation of the hair cells, providing insights to study the induction of hair cell formation in the vertebrates. Afferent innervation in the phenotypes that induce hair cell formation is important to address the effectiveness in using those small molecules for further studies to cure deafness caused due to the loss of hair cells. In addition to its application in the hearing research, the hits identified in this screen might also help us gain a better understanding of the mechanisms that govern the peripheral nerve damage, since the afferent neurons in the zebrafish lateral line system serve as a great model for the peripheral nervous system. Future directions in this small molecule screen would involve analysis of dose-response relationship of the

small molecule hits; studying the hits in zebrafish double transgenics to identify if the small molecule hits simultaneously alter more than one target; further characterization of the hits, and eventually identifying their mechanism of action (MOA) using one of the approaches mentioned above.

Small molecule screens in zebrafish are very useful but they involve certain limitations that we currently encounter. High throughput screening is currently only limited to larval zebrafish and age of the fish can be a major limitation especially in studies that involve age-related genes. However, there has been at least one chemical screen that has been performed on adult zebrafish which studied the fin regeneration (Oppedal, D. et al., 2010). Another limitation pertaining to the design of this chemical screen is the dechorionation of the embryos which reduce the survival rate of the embryos considerably. Apart from these drawbacks, the set of limitations pertaining to imaging exist because most phenotypic studies need manual observation under the microscope which is labor-intensive. Orientation of the fish in each well is also a major drawback while imaging since the phenotype can be best studied when all the fish are oriented in a similar fashion.

Despite these drawbacks, small molecule screens using zebrafish have seen a steep increase in the number of studies since it was first performed in the year 2000. The potential of chemical screen is evident in the fields of cardiology, cancer research, as well as hearing research (Taylor, K. L. et al., 2010). A chemical screen performed using

FDA-approved drugs identified certain compounds that could protect the hair cells from cancer-related drugs such as cisplatin (Vlasits, A. L. et al., 2012). Another screen was performed on the lateral line system of the zebrafish to identify small molecules that could cause and prevent hearing loss (Ou, H. C. et al., 2010). Therefore, small molecule screens holds a tremendous potential in the field of drug discovery and a successful

drug can be discovered after the identified hits in a small molecule screen are further studied in the mammalian models and then further passed on to clinical trials.

Appendix

A total of 63 ‘hits’ were confirmed in the small molecule screen and they were grouped into 12 categories based on their phenotype. This appendix lists the phenotypes exhibited by all the ‘hits’.

Phenotype 1

Severely underdeveloped afferent neurons of the lateral line system including the ALL

(yellow arrowhead) and the PLL (red arrowhead) ganglia.

1600000216 (B5)

1600000216 (G9)

1600000207 (B5)

1600000243 (F4)

1600000243 (A10)

1600000200 (A10)

1600000205 (C8)

1600000232 (D2)

1600000232 (G4)

1600000202 (B10)

1600000213 (A5)

1600000213 (G5)

1600000213 (F10)

1600000224 (A10)

Phenotype 2

Hair cell-like structures (red arrowheads) observed on the dendritic arbors (white arrowheads in the negative control) of the afferent neurons, which innervate the hair cells in the neuromasts.

1600000216 (B1)

1600000243 (G8)

1600000200 (F2)

1600000200 (H3)

1600000203 (B1)

1600000203 (G9)

1600000232 (F2)

1600000232 (H6)

1600000224 (D7)

1600000213 (C10)

Phenotype 3

The afferent axons in the posterior lateral line were observed to be wavy (red arrowheads) as compared to the normal axon (white arrowheads) in the negative control.

1600000205 (A7)

1600000205 (G9)

1600000205 (G10)

1600000207 (A1)

1600000232 (E3)

1600000196 (D5)

1600000213 (B4)

1600000203 (G4)

1600000203 (D5)

Phenotype 4

The afferent axons in the posterior lateral line were observed to end prematurely before reaching the tip of the caudal fin (end point marked with red arrowheads). Normally ending axons have been indicated using white arrowheads.

1600000216 (D3)

1600000207 (D6)

1600000200 (D7)

100

1600000202 (E6)

1600000224 (B7)

Phenotype 5

The afferent axon fascicles in the posterior lateral line appeared to be thinner (indicated by red arrowheads) than the normal phenotype (indicated by white arrowheads).

101

1600000216 (D1)

1600000205 (F6)

1600000205 (D4)

102

1600000243 (G5)

1600000232 (G5)

1600000224 (D9)

103

Phenotype 6

The afferent axons in the posterior lateral line appeared to be wavy (indicated by red arrowheads) as well as ended prematurely (yellow arrowheads); terminating before reaching the base of the caudal fin. The normal phenotype of the posterior lateral line sensory axon has been indicated by white arrowheads.

1600000216 (A1)

1600000243 (B3)

104

Phenotype 7

The embryos show an abnormal morphology such as curved body (red arrowheads).

1600000216 (A8)

1600000216 (A9)

1600000243 (A9)

105

1600000196 (A6)

Phenotype 8

The embryos exhibit bright fluorescence in their dorsally-located somites (red arrowheads).

1600000216 (G4)

106

1600000207 (A3)

1600000196 (F3)

107

Phenotype 9

The embryos exhibit an abnormal morphology of the heart (red arrowheads).

1600000207 (D5)

1600000207 (A7)

1600000203 (G5)

108

Phenotype 10

Certain fluorescent cell-like structures were observed circulating in the body of the zebrafish (region indicated by an asterisk).

1600000200 (G2)

Phenotype 11

Fluorescent patches appear on the entire body surface of the zebrafish (indicated by red arrowheads).

1600000213 (F7)

109

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