Characterization of the Inherent Electrophysiology of Zebrafish Hair Cells and the Effect of Mutations in Met Channel Candidate Genes

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CHARACTERIZATION OF THE INHERENT ELECTROPHYSIOLOGY OF ZEBRAFISH HAIR CELLS AND THE EFFECT OF MUTATIONS IN MET CHANNEL CANDIDATE GENES

By KAYLA JEANNE KINDIG

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

Thesis Advisor: Dr. Brian McDermott Jr.

Department of Biology CASE WESTERN RESERVE UNIVERSITY

May 2019

CASE WESTERN RESERVE UNIVERSITY SCHOOL OF GRADUATE STUDIES

We hereby approve the thesis/dissertation of Kayla Kindig candidate for the degree of Master of Science*.

Committee Chair Nicole Crown

Committee Member Brian McDermott

Committee Member Ruben Stepanyan

Committee Member Hillel Chiel

Committee Member Susan Burden-Gulley

Date of Defense March 22nd, 2019

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

Table of Contents

List of Tables...... 3

List of Figures...... 4

Acknowledgments...... 5

Abstract...... 6

Introduction...... 7 The Human Ear...... 7 Sensory Hair Cells...... 13 The Mechanotransduction Channel...... 16 Properties of the MET Channel...... 16 Candidate Channel Proteins TMC1 and TMC2...... 17 Other Potential MET Channel Proteins...... 21 Important Components of the MET Complex...... 23 Zebrafish and the Lateral Line System...... 26

Materials and Methods...... 32 Zebrafish Breeding...... 32 Lateral Line Microphonic Potential Recordings...... 32 Inner Ear Microphonic Potential Recordings...... 33 Statistics and Software...... 34

Results...... 35 Hair Cell Directionality of Wildtype PrimI Neuromasts...... 35 Tmcs in the Inner Ear and Lateral Line...... 37 Tmc1...... 37 Tmc2a and Tmc2b...... 38 Tmc1, Tmc2a, and Tmc2b...... 42

Discussion...... 48 Channel Composition May Vary Between Hair Cells of the Lateral Line...... 48 TMCs May Combine to Form the MET Complex...... 52

Appendix...... 57

References...... 60

List of Tables Table 1...... 43

List of Figures Figure 1...... 8 Figure 2...... 11 Figure 3...... 11 Figure 4...... 12 Figure 5...... 18 Figure 6...... 24 Figure 7...... 29 Figure 8...... 30 Figure 9...... 31 Figure 10...... 36 Figure 11...... 40 Figure 12...... 41 Figure 13...... 43 Figure 14...... 44 Figure 15...... 45 Figure 16...... 46 Figure 17...... 47 Figure 18...... 57 Figure 19...... 58 Figure 20...... 59 Figure 21...... 60 Figure 22...... 61

Acknowledgements I would foremost like to thank Robin Woods-Davis for her patience in training me and for answering my many questions. Acknowledgment and sincere appreciation is necessary for every member of the McDermott lab, especially those who generated the mutant fish I used in my research. Of course, I would like to thank Dr. Brian McDermott for allowing me to work in his lab and Dr. Ruben Stepanyan for allowing me to use his

equipment, as well as his guidance with electrophysiology. I also need to thank my mom

and my sister Felica for all their efforts in maintaining my sanity. An extra big thanks to

my dear friend Alex Grossman, who proofread this document and gave me great

feedback.

Characterization of the Inherent Electrophysiology of Zebrafish Hair Cells and the Effect of Mutations in MET Channel Candidate Genes

KAYLA JEANNE KINDIG

Abstract

Mechanotransduction is vital for the senses of hearing and balance.

Mechanotransduction occurs when a physical stimulus causes mechanically-gated channels of a sensory cell to open, allowing ions to enter the cell, thus converting a mechanical signal into an electrical one. We know that the mechanoelectrical transduction (MET) channel of sensory hair cells of the inner ear is located at the tips of actin-based stereocilia, but the identity of the pore-forming protein of the channel is unknown. It is also uncertain whether the proteins composing the channel are constant, or if they vary between hair cells based on differences in physiological requirements. In this thesis, I measure stimulus-evoked microphonic potentials of zebrafish hair cells to first determine the electrophysiological response amplitude of wildtype lateral line neuromasts, and then I use this method to examine how the mutation of certain hair cell genes affects mechanotransduction of the lateral line and inner ear. I find evidence to suggest that the MET channel components vary between hair cells, and that the proteins at the pore of the channel differ between mammals and zebrafish. This information may allow us to better understand hair cell tuning at the level of the MET complex and how the proteins necessary for mechanotransduction vary among vertebrates.

Introduction

The Human Ear

The inner ear is the site of sound detection, without which our sense of hearing

would not be possible. The human ear is divided into three parts: the outer, middle, and

inner ear. Sound waves are funneled by the auricle and external meatus of the outer ear

and vibrate the tympanum of the middle ear, which transfers the vibration to the ear

bones—the malleus, incus, and stapes. Movement of the stapes is transferred through the

oval window of the cochlea in the inner ear (Figure 1a). The cochlea is a spiral structure composed of three fluid-filled canals called the scala vestibuli, scala media, and scala tympani. When the stapes moves back and forth in response to sound, it creates a pressure difference that propagates through the scala vestibuli and scala tympani, and this moves the basilar membrane. The up and down motion of the basilar membrane, combined with shearing of the tectorial membrane and vibration of the endolymph, stimulates hair cells in the organ of Corti (Figure 1b). The organ of Corti has two major subsets of hair cells, inner (IHCs) and outer (OHCs) (Figure 2). OHCs are primarily responsible for active amplification of sound via somatic contraction (Brownell et al.

1989) and/or force generated by the stereociliary bundle (Jaramillo et al. 1993). IHCs detect the vibrations of sound and transduce an electrical signal to the many afferent fibers with which they form synapses. These afferent fibers come from spiral ganglion neurons, projections of the eighth cranial nerve. The electrical signal is communicated to the cochlear nuclei of the brainstem and eventually to the auditory cortex through a number of intervening parts of the brainstem (Kandel 2013). Each IHC connects with many afferents, whereas one afferent may connect to several OHCs. Efferent neurons

7 from the superior olivary complex also synapse directly with OHCs to regulate their activity (Kandel 2013; Maison et al. 2003).

Figure 1. The human ear. a) Cross-sectional illustration of a human head showing the outer, middle, and inner ear. The foot of the stapes is like a piston that pushes the fluid of the scala vestibuli through the oval window (obscured). b) Cross-section of one turn of the cochlea showing the organ of Corti, where ~16,000 hair cells responsible for hearing reside. The scala media is filled with the potassium rich fluid called endolymph. When the stapes pushes and pulls through the oval window, compression and rarefaction moves the basilar membrane downward and upward, respectively.

The cochlea is arranged such that higher frequency sounds are detected at the

narrow base, where the basilar membrane is tighter, and low frequency sounds are

detected at the wider apex, where the basilar membrane is more flaccid (Békésy 1960).

Sound waves vibrate the basilar membrane proportionally to the intensity of sound, and

the amplitude of the vibration peaks at the position on the membrane corresponding to that sound frequency. However, frequency tuning involves more than just this passive mechanism. The afferent fibers also contribute to frequency tuning along this tonotopic

map, which ranges from approximately 20 Hz to 20 kHz in humans, as they have a

characteristic frequency that is determined by their position in the cochlear spiral (Geisler

et al. 1974). The firing rate of these neurons can relay not only the frequency of the

stimulus, but also its intensity. Afferent neurons encode sound intensity by increasing

their firing rate with an increase in sound pressure level (SPL) (Evans 1972; Geisler et al.

1974). Afferents across a gradient of low to high sensitivity, with high and low tonic firing rates, connect to a single IHC, and the integration of responses from these neurons can encode stimulus intensity (Sachs & Abbas 1973). Sound frequency and intensity are linked in that our hearing is most sensitive to frequencies of 1-4 kHz, but higher and lower frequencies can be detected at greater SPL (Heffner & Heffner 2007). Force generated from OHCs enhances hearing sensitivity, lowering detection thresholds within our frequency range by augmenting membrane displacement (Brownell 1990; Ryan &

Dallos 1975).

In addition to hearing, our sense of balance relies on the inner ear. There are three semicircular canals, and they contain vestibular hair cells that are responsible for detecting angular acceleration (Figure 3). These hair cells are arranged into epithelial

structures called cristae, housed in ampullae near the base of the canals, and connected by a gelatinous cupula. When the head is rotated, the cupula is displaced by inertia of the

endolymph. This deflects the hair cells of the cristae, which are oriented such that hair cells in one ear are excited while those in the paired canal of the other ear are inhibited, conveying directionality of the rotation (Kandel 2013). There are two types of vestibular hair cells—type 1 and type 2. Visually, they can be distinguished by their shape and the way in which they synapse with the afferent neuron. Type 1 hair cells have a flask-like shape and are innervated by cup-like calyx synaptic terminals, whereas type II hair cells are more rectangular and innervated by smaller bouton terminals (Rusch et al. 1998). In addition to the semicircular canals, there are also two flat patches of vestibular hair cells called maculae that lie between the semicircular canals and the cochlea (Figure 4). The two maculae are oriented at 90 degrees to one another and detect linear acceleration. The hair cells of the maculae lie beneath calcium carbonate crystals called otoconia embedded in gelatinous matrix. Together, the otoconia and the maculae of these two patches are called the utricle and saccule. The otoconia shifts with linear movement, and this motion relative to the hair cells causes their deflection. Each macula is organized such that any movement within its plane will activate some cells and inhibit others, and this combination of activation and inhibition can encode directionality of movement. As with the cochlea, vestibular hair cells synapse with projections of the eighth cranial nerve

(Gray 1997).

Figure 2. The organ of Corti. Magnified view of the organ of Corti pictured in Fig. 1b. When sound waves enter the ear, the basilar membrane vibrates. The vibration of the endolymph and the shearing of the tectorial membrane deflects the stereocilia of the IHCs and OHCs, respectively. Movement of the basilar membrane upward causes the tectorial membrane to push hair cells in the excitatory direction, whereas downward movement causes shearing in the inhibitory direction. At least 90% of spiral ganglion neurons connect to IHCs (Kandel 2013), a sign of their critical role in sound detection.

Figure 3. A human semicircular canal. Gray region (left) is the region of the horizontal canal transected and isolated (right). Cross section has also been rotated 90º for clarity. The hair cells of the crista are surrounded by a gelatinous cupula. Movement of the cupula stimulates the sensory hair cells. All hair cells within a crista are oriented in the same direction.

It is estimated that approximately 460 million people currently suffer from debilitating hearing loss (World Health Organization 2018). There are many potential causes of hearing loss. Sensorineural hearing loss, which is caused by damage to hair cells or the cochlear nerve, can result from noise exposure, bacterial or viral infections, drugs, head trauma, genetics, or aging (Mills & Going 1982). Unlike many other cell types, human hair cells do not divide, so any cell death is permanent (Kandel 2013).

Consequently, everyone loses some hearing acuity over time, especially in the high

frequency range (Gray 1997). However, some people are born with mutations that cause

hearing loss or deafness at a young age. There are two main types of congenital deafness:

nonsyndromic and syndromic. Nonsyndromic deafness occurs without any other

symptoms, whereas people with syndromic deafness present with other symptoms

besides hearing loss (Shearer et al. 2017). One prominent example of syndromic deafness is Usher syndrome, in which profound sensorineural deafness is accompanied by vestibular dysfunction and often retinopathy (Usher Syndrome 2019). Although hearing loss in itself is a serious public health concern that affects nearly everyone, the existence of syndromic deafness underscores the importance of identifying the genes involved in hearing loss, because they can often act across multiple sensory systems that utilize similar processes for different tasks. This is especially true for the sense of balance, since the cells responsible are the same as those used for hearing.

Figure 4. The human maculae. a) The utricle and saccule are patches of sensory hair cells positioned between the cochlea and semicircular canals. b) Diagram showing orientation of hair cells in the saccule (purple) and utricle (orange). The head of the arrow indicates the position of the kinocilium. P: posterior, I: inferior, A: anterior, L: lateral. c) Drawing of hair cells in each macula. Kinocilia in the utricle (orange) are directed toward the center line, or striola, whereas kinocilia in the saccule (purple) are directed away from the striola. Linear movement of the head or body will cause the otoconia to move relative to the hair cells, which will activate some cells while inhibiting others.

Sensory Hair Cells

Hair cells are highly polarized cells that are specialized for the task of

mechanotransduction. Hair cells have an apical surface lined with actin-based projections called stereocilia, arranged by graded heights in a staircase-like fashion. Next to the tallest row of stereocilia is a single microtubule-based kinocilium that degrades in mammalian auditory hair cells after birth, the function of which may be to permit mechanosensitivity in hair cells before they have fully developed (Kindt et al. 2012).

Stereocilia connect to the hair cell via a flexible tapered segment called the fonticulus that integrates with a meshwork of actin called the cuticular plate. At the tips of stereocilia are non-selective, mechanically-gated cation channels (Lumpkin & Hudspeth 1995) as well as small linkages composed of homodimers of the calcium adhesion proteins cadherin 23 and protocadherin 15 (Alagramam et al. 2011). When sound waves perturb the fluid of the scala media and shear the tectorial membrane, stereocilia bend at their tapered bases, and a deflection towards the tallest stereocilium causes tension in the tip links that pulls the shorter stereocilia and open the mechanically-gated channels at the tips. When these mechanoelectrical transduction (MET) channels open, cations enter the cell passively due to the high concentration of potassium ions in the endolymph relative to the inside of the

cell. This influx depolarizes the cell from its resting potential of approximately -50 mV,

and the receptor potential activates voltage-gated calcium channels on the basolateral

surface of the cell (Figure 5). An influx of calcium allows the fusion of glutamate-filled

vesicles with the basal membrane, mediated by the synaptic ribbon. The neurotransmitter

is received by the afferent neuron where it creates EPSPs that will trigger an action

potential, allowing the signal to propagate to the brain. Calcium-dependent potassium

channels, voltage-sensitive K+ channels, and Ca2+ pumps on the basolateral surface of the

hair cell remove K+ and Ca2+ to repolarize the cell and re-establish the concentration gradient for another cycle of mechanotransduction (Kandel 2013). A portion of the MET

channels are open at rest, so a deflection toward the shortest stereocilia will result in the

closing of these channels and thus a slight hyperpolarization of the cell, which will

reduce the spontaneous firing rate of the afferent neurons (Lumpkin & Hudspeth 1995).

Each hair cell is tuned to a specific frequency of sound along the tonotopic gradient of the cochlea. Tuning may be determined by a number of factors, such as the stereocilia length, the flexibility of the bundle, or electrical properties of the cell. Changes in stereocilia length and number of stereocilia per bundle are seen in both mammalian and non-mammalian hair cells, with a decrease in tallest row height and increase in average number of stereocilia from the low to high frequency end of the basilar membrane (Roth & Bruns 1992; Tilney & Saunders 1983). Rigidity of the bundle is directly proportional to the number of stereocilia (Howard & Ashmore 1986) and inversely proportional to the square root of their length (Crawford & Fettiplace 1985).

Stiffness can also be altered by the quantity of actin per stereocilium (Howard &

Ashmore 1986; Tilney et al. 1980). Width of stereocilia, and therefore the quantity of actin filaments, also increases from low to high frequency hair cells in the ear of chicks

(Tilney & Saunders 1983). Thus, stiffness of the hair cell bundle mimics that of the basilar membrane along the tonotopic gradient, as hair cells tuned to low frequencies exhibit multiple bundle characteristics that make them more flexible, whereas hair cells tuned to high frequencies have less flexible bundles. Another method of frequency tuning observed in non-mammals is electrical resonance, when differences in basolateral K+ or

Ca2+ channel number or protein composition alter a cell’s sensitivity to voltage or

calcium, which can change how easily the cell is repolarized and how quickly it can

communicate a signal (Fettiplace & Fuchs 1999). Tuning could also occur at the level of

the MET channel; systematic decreases in MET channel activation and adaptation time

constants from the low to high frequency end of the cochlea, perhaps caused by changes

in the channel proteins, can act as an intrinsic bandpass filter for the hair cell (Ricci et al.

2005). It has been proposed that a different number of TMC proteins could be incorporated into the MET complex depending on the hair cell’s position in the cochlear spiral (Pan et al. 2013; Beurg et al. 2018). Variation in MET composition is consistent with a nearly two-fold increase in OHC single channel conductance from the apex to the base of the cochlea (Beurg et al. 2014), and a decrease in OHC Ca2+ permeability from

the apex to the base before postnatal day 6 in mice (Kim & Fettiplace 2013). The

conductance shift may also be due to the exchange of neutral residues in the MET

channel pore for negatively charged residues, or by association with an increasing

number of negatively charged accessory proteins (Fettiplace & Kim 2014).

A known functional property of hair cells and the MET current is adaptation.

Adaptation is when the MET current from a hair cell becomes attenuated during a

prolonged stimulus, but it the channel is still sensitive to greater deflection. This process

is known to be calcium dependent, as the MET response is mitigated faster and to a

greater degree when more Ca2+ is added to the extracellular medium (Eatock et al. 1987) and adaptation can be eliminated with high intracellular concentrations of the calcium chelator BAPTA (Corns et al. 2014). During “slow” adaptation, which has a time constant of approximately 45 ms in the mouse utricle (Vollrath & Eatock 1999), it is

thought that the myosin motors associated with tip links adjust their vertical position

downward to reduce the amount of tension generated from the prolonged deflection

amplitude, causing fewer channels to be open unless deflected further (Hudspeth &

Gillespie 1994). The positional adjustment of myosin is likely dependent on calcium

either through calmodulin binding or a direct effect of calcium on the ATPase cycle

(Hudspeth & Gillespie 1994). There is also another mechanism of adaptation, called “fast

adaptation”, in which Ca2+ is thought to change the conductance of the MET channel by

binding to an intracellular regulatory site on the protein (Fettiplace & Ricci 2003).

Alternatively, fast adaptation may occur by the binding of Ca2+ to a nearby structural

component, which could reduce tension in the membrane and therefore reduce channel

activation (Stepanyan & Frolenkov 2009). Fast adaptation occurs on a timescale of less

than a millisecond in the OHCs of rats (Ricci et al. 2005).

The Mechanotransduction Complex

Properties of the MET Channel

It is currently uncertain which proteins form the pore of the MET channel, but the

channel does have a number of known properties. MET channels activate quickly when

mechanically stimulated, with a response latency of only a few microseconds in rats

(Ricci et al. 2005). As previously mentioned, the MET channel localizes to the tips of stereocilia (Lumpkin & Hudspeth 1995). There are likely 1-2 channels atop each stereocilium (Beurg et al. 2006). The channels are functional, according to dye uptake

assays, by postnatal day 0 in basal OHCs (Lelli et al. 2009) and by embryonic day 16.5 in

vestibular hair cells (Géléoc & Holt 2003) of mice, indicating that gene expression of the

necessary components should occur before that time. Potassium ions, which are highly

concentrated in the endolymph, are the cations primarily responsible for the MET current

(Hudspeth 1989), though the channel is a nonselective cation channel in vitro (Corey &

Hudspeth 1979). As discussed earlier, conductance of the channel varies along the

tonotopic gradient of the cochlea (Beurg et al. 2014), as does its calcium permeability in young OHCs (Kim & Fettiplace 2013). The single channel conductance in auditory hair cells has been measured to be between 67 and 320 pS in mice, and it appears to be lower on average for vestibular hair cells (Fettiplace & Kim 2014). The pore is thought to be larger on the extracellular side and contain an internal binding site for Ca2+ (Pan et al.

2012). The resting open probability of the channel is inversely related to Ca2+

concentration, likely due to the binding of Ca2+ in the pore or to an intracellular site

during fast adaptation (Fettiplace & Kim 2014). The pore diameter has been estimated at

a maximum of 1.7 nm at its widest point due to the channel’s ability to be blocked by

curare (Farris et al. 2004). The MET channel can also be blocked by aminoglycoside

antibiotics such as dihydrostreptomycin (Marcotti et al. 2005) and permeated by the florescent dye FM1-43 (Gale et al. 2001).

Channel Candidate Proteins TMC1 and TMC2

Transmembrane channel-like (TMC) protein 1 was first discovered when investigators found its gene mutated in cases of dominant and recessive inherited deafness (Kurima et al. 2002). Mutation or deletion of homologs of this gene in mice

yields animals that are deaf or have a greatly reduced ability to hear (Vreugde et al.

2002). TMC1 has been localized to the tips of stereocilia and has been shown to interact

with protocadherin 15 (Maeda et al. 2014). Point mutations in TMC1 greatly reduce the

17 single-channel conductance and Ca2+ permeability of the MET channel (Corey & Holt

2016), and mice lacking functional TMC1 will lose MET currents completely after postnatal day 10, when TMC2 is no longer expressed (Kim & Fettiplace 2013). It is currently thought that TMC1 has 10 transmembrane domains and assembles as a dimer

(Pan et al. 2018), though previously it was predicted to have 6 transmembrane domains

(Labay et al. 2010). Tmc1 mRNA can be detected by postnatal day 3 in the hair cells of mice, and expression is maintained into adulthood (Kawashima et al. 2011).

Figure 5. A simplified inner ear hair cell. Deflection of the stereocilia toward the kinocilium pulls on tip links and opens the MET channels, allowing K+ and some Ca2+ to enter the cell and depolarize it. The depolarization activates voltage-gated Ca2+ channels, which allow glutamate to be released to the afferent nerve fiber. Efferent neurons hyperpolarize OHCs when activated, and this is thought to disrupt electrical resonance tuning and desensitize the cells when their amplification ability is unnecessary (Art et al. 1984). In addition to tip links, there are side-links, ankle-links, and top-links that stabilize the stereocilia bundle (Goodyear et al. 2005). Tight junctions with surrounding cells help separate the K+-rich endolymph of the scala media from the Na+-rich perilymph that surrounds the nerve fibers.

The TMC family has eight members in total, all with 6-10 predicted

transmembrane domains (Kurima et al. 2003; Keresztes et al. 2003). TMC2 is another

member of the family that was found to be associated with hearing, though mutations in

the Tmc2 gene alone have never been found to cause deafness in humans (Kawashima et

al. 2015). TMC2 also localizes to the tips of stereocilia in mice and associates with

protocadherin 15 (Maeda et al. 2014). TMC2 is able to substitute for TMC1 in its absence and partially compensate for loss of function in cochlear hair cells until it is no longer expressed (Kurima et al. 2015). Tmc1-/- Tmc2-/- mice do not have detectable MET

currents in cochlear or vestibular hair cells at any time point, even though tip links appear

to be intact, and these mice have both hearing loss and vestibular dysfunction

(Kawashima et al. 2011). Overexpression of either TMC1 or TMC2 with adeno-

associated viral vectors can partially rescue the lack of MET current in double mutants

(Askew et al. 2015). It has been proposed that TMC1 and TMC2 may both form the MET

channel as a heteromeric complex (Pan et al. 2013), and while they are capable of

binding and forming a heterodimer (Pan et al. 2018), florescent labeling has shown

surprisingly little overlap between the localization of the two proteins in the cochlea

(Kurima et al. 2015). Even though Tmc2 expression appears to stop after postnatal day 10

in the mouse cochlea (Kurima et al. 2015), TMC2 can be detected in the utricle of mice

until at least postnatal day 25 (Kawashima et al. 2011; Nakanishi et al. 2018).

Additionally, Tmc1+/+ Tmc2-/- mice show a nearly 10-fold reduction in MET currents of

the utricle and have noticeable vestibular defects, while there is no significant difference

in macroscopic MET current of Tmc1+/+ Tmc2-/- OHCs and those of wildtype mice, nor is

their hearing impaired (Kawashima et al. 2011). However, knockout of TMC2 in mice

leads to decreased Ca2+ permeability of OHC MET channels across the entire tonotopic

gradient before postnatal day 6 (Kim & Fettiplace 2013), so TMC2 may assist in the

development of OHCs by supplying them with transiently elevated levels of Ca2+, as

there is evidence that Ca2+ influx is required to establish proper stereocilia morphology

(Vélez-Ortega et al. 2017). The prolonged presence of TMC2 in the utricle and the effect

of its mutation on utricular MET currents indicates that perhaps mammalian vestibular hair cells rely more on TMC2 while cochlear hair cells rely more on TMC1, which could explain why Tmc1 mutations that cause deafness in humans do not necessarily cause vestibular defects (Kurima et al. 2002; Makishima et al. 2004; der Heer et al. 2011).

Regardless, there appears to be some degree of functional redundancy between the two proteins.

Whether TMC1 or TMC2 are auxiliary or actually form the pore of the MET channel is unknown. Invertebrate TMC1 orthologues have roles in ion regulation or

mechanosensitivity that suggest the protein could be an ion channel, such as alkaline

sensation in nociceptive neurons of C. elegans (Wang et al. 2016) and food texture

sensation in Drosophila (Zhang et al. 2016). In C. elegans, changes in resting membrane

potential with manipulation of TMC-1 suggest a possible role as a leak channel (Yue et

al. 2018). Some topology predictions for TMC1 suggest a re-entrant loop between the 4th

and 5th transmembrane domains that could form a pore (Labay et al. 2010), which would

make the structure similar to known voltage-gated K+ channel KCNA1 (Cunnningham &

Muller 2018). If TMC1 has ten transmembrane domains, this makes it similar in topology to the TMEM16 ion channel family, whose structure has been characterized (Brunner et al. 2014). In this case, the pore region is thought to be formed by transmembrane

domains 4-7 (Pan et al. 2018). Since the crystal structure of TMC1 has yet to be resolved to corroborate the existence of a channel pore, it is still possible that the protein could be acting as an accessory. Beethoven mice, with a mutation in Tmc1 that adds a positive charge near the 4th transmembrane domain, appear to have increased Ca2+ dependent

block in IHCs (Pan et al. 2013) and a decreased sensitivity to the MET channel blocker

dihydrostreptomycin in OHCs (Corns et al. 2016). Since these are presumably both

properties of the MET pore, it seems plausible that TMC1 could be at the pore of the

channel. Recent evidence showing a roughly 20 mV change in MET channel reversal

potential with chemical alteration of TMC1 cysteine residues, which can be prevented

with application of a pore blocker or negative stereocilia deflection, further suggests that it is at the pore (Pan et al. 2018). However, TMC1 has not yet been shown to confer mechanosensitivity to non-mechanosensitive cells or a lipid bilayer. TMC1 and TMC2 remain in the endoplasmic reticulum of heterologous cells, even when other known components of the MET complex are present (Giese et al. 2017). Until it is discovered how to insert and assemble TMC1 properly into a membrane, we will not know if it is sufficient for hair cell mechanosensitivity.

Other Potential MET Channel Proteins

Another protein unrelated to the TMCs that has been considered as a possible pore-forming subunit of the MET channel is lipoma high mobility group IC fusion

partner-like 5 (LHFPL5). As with TMC1, mutations in LHFPL5 can cause deafness in

humans and mice (Kalay et al. 2006). LHFP expression has been shown to start when the

MET response starts in mice, around postnatal day 0, and the protein localizes to the tips

of stereocilia where it binds with both protocadherin 15 and TMIE (Xiong et al. 2012).

Removal of LHFPL5 leads to a significant reduction in macroscopic MET current, but also a reduction in the number of tiplinks, which could be the actual cause of the reduced current (Xiong et al. 2012). However, single channel conductance is also affected, as well as channel activation and adaptation (Beurg et al. 2015). LHFPL5 alone or together with protocadherin 15 does not confer mechanosensitivity to a heterologous cell (Zhao et al.

2014), and so it cannot be definitively determined as a mechanosensitive ion channel.

LHFPL5 is still present in stereocilia when Tmc1 and Tmc2 are knocked out, but TMC1 is no longer present at the tips of stereocilia when LHFPL5 is knocked out (Beurg et al.

2015), suggesting that LHFPL5 may play a role in transporting TMC1 to the tips of stereocilia. Thus, the primary impact of mutations in LHFPL5 may be through its effect on TMC1, though it has yet to be shown if the two proteins interact (Beurg et al. 2015).

An additional candidate for the MET channel pore is the transmembrane inner ear

(TMIE) protein. Dysfunctional TMIE also causes deafness in humans (Naz et al. 2002) and mice (Mitchem et al. 2002), as well as zebrafish (Gleason et al. 2009).

Immunolabeling of TMIE shows that it localizes to the tips of stereocilia, although when its expression begins is unclear (Zhao et al. 2014). On its own, TMIE can bind to one splice variant of protocadherin 15, and it can bind to the two other main splice variants when complexed with LHFPL5 (Zhao et al. 2014). In Tmie-null mice, there are no detectable MET currents, while the localization of protocadherin 15, TMC1/2, and

LHFPL5 appears unaffected (Zhao et al. 2014). The MET currents can be rescued by exogenous expression of TMIE (Zhao et al. 2014). It is unclear if the studied mutations of Tmie result in improper localization of the truncated protein, which could potentially confound its effects on the MET current if it plays a regulatory role in the complex

(Cunningham & Muller 2018). Notably, TMIE cannot confer mechanosensitivity to a heterologous cell, even in the presence of TMC1/2, LHFPL5, or protocadherin 15 (Zhao et al. 2014). Assuming that one of the aforementioned proteins forms the pore of the

MET channel, a lack of conferred mechanosensitivity by any of them implies that one of the components was not properly localized, or that another factor required for correct assembly of the complex has yet to be identified. If TMIE is not the pore of the MET channel, then it is possible that its function is to link the channel to protocadherin 15

(Cunningham & Muller 2018).

Important Components of the MET Complex

Other proteins function as part of the MET complex and can cause hearing loss if removed or impaired (Figure 6), though they are not considered to be part of the channel proper. As established previously, tip links composed of homodimers of protocadherin 15 and cadherin 23 connect adjacent stereocilia and transmit tension to the cell membrane, opening MET channels (Kazmierczak et al. 2007; Alagramam et al. 2011). Removal of tip links via a calcium chelator will result in loss of the conventional MET current

(Indzhykulian et al. 2013) and mutation of the gene for protocadherin 15 can cause deafness in humans (Lelli et al. 2010). Pulling on tip links directly is enough to open the

MET channel without deflection of the bundle (Basu et al. 2016), but it is currently unknown if protocadherin 15 directly connects to the MET channel or if it merely activates the channel through deformation of the lipid bilayer. Myosin-7a, HARMONIN, and SANS, which are all implicated in Usher syndrome type I, reside around the insertion point of cadherin 23 and are thought to mediate the resting tension of the tiplink (Grati &

Kachar 2011). Near the tip of the shorter stereocilia, myosin-15a, WHIRLIN, and

CLRN1 appear to be involved in development of the hair bundle (Stepanyan & Frolenkov

2009: Delprat et al. 2005; Geng et al. 2012; Gopal et al. 2015). Myosin-15a may also be necessary for fast adaptation in IHCs (Stepanyan & Frolenkov 2009).

Figure 6. Current understanding of the MET complex. Several proteins work together to coordinate mechanotransduction at the tips of actin-based stereocilia. See text for a description of each component. Note that Pan et al. predicts 10 transmembrane domains for TMC1 (2018). Myosin-1c may also localize near myosin-7a (Holt et al. 2002). Figure modeled after Cunningham & Muller 2018.

The role of calcium and integrin binding family member 2 (CIB2) in the MET

complex is somewhat unclear. The protein is part of a family thought to mediate calcium

signaling (Gentry et al. 2005). In vitro, CIB2 interacts with WHIRLIN, myosin-15a,

TMC1, and TMC2 (Giese et al. 2017). A mutation in Cib2 that is associated with human

nonsyndromic deafness eliminates the MET current (Giese et al. 2017). Deafness-causing

mutations in Cib2 appear to reduce interaction between CIB2 and TMC1/2, yet the

position of both TMCs, as well as myosin 7a, HARMONIN, and protocadherin 15 seem

to be unaffected in Cib2 mutants (Giese et al. 2017). CIB2 is also insufficient to insert

TMC1/2 in the cell membrane of heterologous cells (Giese et al. 2017). Shorter

stereocilia appear to overgrow in two different Cib2 mutants (Giese et al. 2017),

suggesting its function may be to limit shorter stereocilia growth, permitting the height

gradient that allows normal mechanotransduction to occur.

Transmembrane O-methyltransferase (TOMT) is required for TMC1 and TMC2

to properly localize to the tips of stereocilia in zebrafish (Erickson et al. 2017) and mice

(Cunningham et al. 2017). However, presence of TOMT alone is not enough for TMC1/2

to integrate into the membrane of heterologous cells, indicating that another protein is

probably necessary (Cunningham et al. 2017; Erickson et al. 2017). Immunostaining places TOMT in the cytoplasm of the cell body (Ahmed et al. 2008) as opposed to the stereocilia tips. Mutations in TOMT can cause deafness in mice and humans (Du et al.

2008), as well as zebrafish (Erickson et al. 2017).

PIP2 is a phospholipid in the membrane of hair cells that can affect seemingly

inherent properties of the MET channel. PIP2 is a known component of frog (Hirono et

al. 2004), rat (Effertz et al. 2017), and mouse stereocilia (Goodyear et al. 2008).

Pharmacological removal of PIP2 from the hair cells of the frog sacculus results in

reduced adaptation and decreased macroscopic MET current (Hirono et al. 2004). There

are also a number of other MET channel properties that are affected by PIP2 removal,

such as single channel conductance, time course of channel activation, and resting open

probability (Effertz et al. 2017). Additionally, removal of PIP2 changes the reversal

potential of the MET channel (Effertz et al. 2017), similarly to cysteine modifications of

TMC1 (Pan et al. 2018). PIP2 likely influences the rigidity of the lipid bilayer, which can

affect the degree of tension experienced by the channel when the membrane is deformed.

The Lateral Line System and Zebrafish

Zebrafish (Danio rerio) have sensory organs called neuromasts on the sides of their bodies, arranged in a line called the lateral line. Each neuromast is composed of several sensory hair cells that face opposite to one another in approximately equal numbers (Figure 7a). This morphological feature causes a phenomena known as the “2f response” when taking electrophysiological recordings with sinusoidal stimuli (Corey &

Hudspeth 1983), since the excitatory direction for one subset of hair cells is the inhibitory direction of the other (Figure 7b). The lateral line is split into the anterior lateral line, on the head, and the posterior lateral line on the tail (Figure 8). Neuromasts of the anterior lateral line are deposited between 34 and 72 hours post fertilization (hpf) (Raible & Kruse

2000). Within the posterior lateral line, there are two populations that can be differentiated by the groups of migrating primordial cells (Prim) from which they originate. PrimI neuromasts are deposited first, around 20 hpf, and they contain hair cells that face anteriorly and posteriorly, while PrimII neuromasts begin to be deposited

around 48 hpf and they contain hair cells that face dorsally and ventrally (Pujol-Martí &

López-Schier 2013; López-Schier et al. 2004). Neuromasts closer to the head are deposited first, as the primordium originates from cephalic placodes just behind the ear and migrates caudally (Ghysen & Dambly-Chaudière 2004). The two populations of hair cells within a neuromast are innervated by separate afferents, which connect to the reticulospinal mauthner neuron in the hindbrain (Ghysen & Dambly-Chaudière 2004).

Each hair cell in a neuromast is surrounded by a ring of supporting cells, and the hair cell stereocilia are enveloped by a gelatinous mass called the cupula. Neuromasts are used by fish to detect water flow (Montgomery et al. 2000), enabling them to sense predators and perform rheotaxis (Olszewski et al. 2012). Although there is evidence to suggest that unique neuromasts are tuned to slightly different frequencies of motion via modification of cupula morphology (McHenry & van Netten 2007), they do not appear to be arranged tonotopically like the hair cells of the mammalian cochlea (Trump & McHenry 2008).

Neuromasts are, however, arranged somatotopically— the organization of afferent neurons reflects neuromast position on the body (Alexandre & Ghysen 1999).

Zebrafish also have inner ears that contain hair cells. Unlike humans, they do not have a cochlea, instead having flat patches of hair cells called maculae beneath ear bones or otoliths. There are three otoliths and three associated macula: the utricle, the saccule, and the lagena. The utricular and saccular otoliths are visible at 19.5 hpf, and the first macular hair cells are visible around 24 hpf (Haddon & Lewis 1996). The utricle or anterior macula primarily detects linear acceleration (Riley & Moorman 2000), whereas the saccule or posterior macula primarily detects sound (Abbas & Whitfield 2010; Yao et al. 2016; Inoue et al. 2013). Although the two maculae appear distinct, they arise from

the same epithelial sheet and are not physically separated until the fish is an adult

(Haddon & Lewis 1996). The lagena detects both vestibular and auditory stimuli, but

does not develop until after the other maculae, at least 11 days post fertilization (dpf ) (Lu

& DeSmidt 2013). Zebrafish also have three semi-circular canals with cristae that detect

angular acceleration, which become visible around 60 hpf (Haddon & Lewis 1996).

Similarly to the lateral line, the utricle and saccule have hair cells that face

opposing directions, although they are arranged in a radial fashion instead of at precisely

180 degrees to one another. The saccular macula has distinct anterior and posterior

regions; the mean orientation of cells in the anterior of the saccule is approximately

dorsal or ventral, whereas the mean orientation of cells in the posterior region is

approximately anterior or posterior (Yao et al. 2016; Inoue et al. 2013). The utricular

macula appears to contain hair cells that are mostly in a lateral or medial orientation

(Inoue et al. 2013) (Figure 9). This difference in polarity allows the characteristic 2f

microphonics response to be observed with inner ear recordings. The polarity of the

zebrafish maculae is also somewhat analogous to that of the human maculae, which have

hair cells of various orientations. Unlike humans, zebrafish continue to add hair cells to

their ears for most of their lives, though this addition seems to stabilize after about 10 months, at which point they have roughly 3,500 saccular hair cells (Higgs et al. 2001).

When vibrations enter the ear, the difference in inertia between the otolith and the hair cell bundle allows for the detection of sound or linear motion (Abbas & Whitfield

2010). The division of auditory and vestibular stimuli between the two maculae of zebrafish larva can be attributed to a difference in otolith mass as opposed to properties of the hair cells themselves, with saccular otoliths having a larger size (Inoue et al. 2013).

Experimental estimates place the range of zebrafish hearing from 100 to 4,000 Hz (Higgs

et al. 2001), though others estimate an upward limit of 12,000 Hz (Wang et al. 2015).

Presumably, larval zebrafish are unable to hear higher frequency sounds until the growth

of the Weberian ossicles, which do not fully form until around 20 dpf (Yao et al. 2016;

Monroe et al. 2016), but some argue that only hearing sensitivity and not frequency range

change with age (Higgs et al. 2001; Wang et al. 2015). Hair cells of the maculae are

innervated by projections of the VIIIth nerve, which connect to the ipsilateral mauthner

cell (Tanimoto et al. 2009). Zebrafish hair cells and the associated circuitry are fully

functional by 4 dpf, as they have an acoustic startle response and will orient themselves

to be upright by this age (Kimmel et al. 1974). Sound-evoked microphonic potentials can

be detected from the ear at 40 hpf and appear mature by 55 hpf (Tanimoto et al. 2009).

Figure 7. Hair cells of a lateral line neuromast. a) Lateral view of a neuromast. Hair cells of the two opposing directions are innervated by separate afferent nerve fibers. While kinocilia degrade in mammals after birth, they persist into adulthood for zebrafish. b) Two simplified hair cells isolated from a neuromast. Top: lateral view. As a consequence of opposing polarity, stimulation of the neuromast in either direction along its axis of sensitivity excites one subset of hair cells while inhibiting the other. Bottom: top-down view. The kinocilium appears dark due to its density, allowing for a determination of cell polarity when visualizing cells from the top.

Figure 8. Mechanosensory map of the zebrafish lateral line at ~6 dpf. Diagram showing the abbreviated names and approximate position of major lateral line neuromasts. Neuromasts in blue have hair cells oriented along the A-P axis. Neuromasts in orange have hair cells oriented along the D-V axis. The ganglion for the anterior lateral line (green) is positioned anterior to the ear (grey), whereas the posterior lateral line ganglion (red) is posterior to the ear. Anterior lateral line is composed of several distinct lines, such as the infraorbital (IO), mandibular (M), otic (O), and opercular (OP) lines.

Zebrafish are a useful model organism for a number of reasons. They produce

many embryos, up to 200 per clutch, allowing for a large pool of samples with siblings to

use as controls. Adults have external fertilization, allowing for easy genetic manipulation

of embryos at the one-cell stage. The embryos develop relatively quickly, hatching at

approximately 3-4 dpf and reaching sexual maturity around 3 months post-fertilization,

which allows experiments to be planned and executed faster than with mice. Most

importantly, zebrafish are transparent up to 1 month post fertilization, making their cells

easy to visualize under a microscope. For the purpose of studying mechanotransduction,

they are quite useful in that hair cells line the exterior of their bodies, and hair cells of the

ear are relatively easy to access without dissection. Zebrafish hair cells are functionally similar to those of mammals with regard to their basolateral current profiles (Olt &

Marcotti 2014; Haden et al. 2013) and ototoxic drug susceptibility (Owens et al. 2008).

Thus, zebrafish are a valuable tool in the study of hearing.

Zebrafish are also useful for studying genes related to deafness. There is

approximately 70% similarity between the zebrafish and human reference genomes

(Howe et al. 2013). Due to an evolutionary event called the teleost-specific genome

duplication, many human genes have multiple zebrafish orthologs (Howe et al. 2013). It

is possible that duplicate genes could have evolved to have slightly different functions in

zebrafish, as appears to be the case for Tmc2a and Tmc2b, and this may complicate

attempts to extrapolate human function from results in zebrafish. However, roughly 47%

of human genes have just one zebrafish ortholog (Howe et al. 2013), and one such

example is Tmc1. In addition to their sequence similarity with humans, there are several

mutations known to cause human deafness that will cause deafness if applied to zebrafish

orthologs (Pickett & Raible 2019). There are over a dozen zebrafish models of human

hereditary deafness (Pickett & Raible 2019), indicating that the function of many

hearing-related genes is conserved.

Figure 9. Ear of a larval zebrafish at ~7 dpf. Diagram showing the position of otoliths and cristae in the otic vesicle of a laterally-mounted fish. Each of the three cristae is associated with a semicircular canal, but these have been omitted for clarity. Bundles of the cristae are depicted as grey lines. Beneath the otoliths lie the hair cells of the maculae. Grey arrowheads show the approximate mean polarity of hair cells in that macula, with circles used to indicate the z-plane, based on data from Inoue et al. 2013. The saccule has hair cells that primarily face the anterior and posterior, as well as the dorsal and ventral side of the animal. The utricle has hair cells that face laterally and medially. All hair cells within a crista face the same direction.

Materials and Methods

Zebrafish Strains

Zebrafish strains were bred and cared for in the Case Western Reserve Zebrafish

Facility following standard protocols (Nusslein-Volhard, et al. 2002). Fish included in the following studies have mutations in tmc1 (ENSDARG00000056386), tmc2a

(ENSDARG00000033104), or tmc2b (ENSDARG00000030311). Wildtype fish are

Tübingen (Tü). Fish with sa807 mutation in tmc1 were obtained from the zebrafish stock center.

Lateral Line Microphonic Potential Recordings

Recordings were made from fish 5-7 dpf, anesthetized at room temperature

(~22ºC) with MS-222 (Sigma-Aldrich, St. Louis, MO) in a bath solution consisting of 12 mM Nacl, 2mM KCl, 10 mM HEPES, 2mM CaCl2, and 0.7 mM NaH2PO4, adjusted to pH of ~7.3. Larvae were restrained using strands of dental floss in a dish containing the bath solution. Heart rate and blood flow were visually monitored to assess larva viability before, during, and after experiments. The fish were visualized under an upright Olympus

BX1WI microscope using a 4x 0.1NA and a 100x 1 NA objective. All neuromast images were collected with a Grasshopper3 CMOS camera (Point Grey, Richmond, BC,

Canada). jClamp software (SciSoft, Joseph Santos-Sacchi, Yale, New Haven, CT) was used to generate either a sinusoidal stimulus of 50 Hz or sustained deflections for 200 ms, delivered as a fluid jet from a borosilicate glass pipette. The diameter of the stimulus pipette was approximately 7 µm and was placed about 50 µm away from the neuromast, parallel to the neuromast’s axis of best sensitivity. A HSPC-1 (ALA Scientific

instruments, Farmingdale, NY) was used to control the fluid jet. The recording electrode was placed inside a borosilicate glass pipette of approximately 1 µm tip diameter, which was placed near the apical surface of the neuromast. Resistance of the recording pipette was 3–6 MΩ when filled with the bath solution. Placement of pipettes was controlled by

a set of micromanipulators (MPC-325; Sutter Instrument). Microphonic potentials were recorded by jClamp in current clamp mode and amplified using a PC-505B amplifier

(Warner Instruments, Hamden, CT), SIM983 scaling amplifier, (Stanford Research,

Sunnyvale, CA), and PCI-6221 digitizer (National Instruments, Austin, Texas). All

sinusoidal recordings were low pass filtered at 200 Hz, and each trace shown and used in

analysis is an average of at least 500 deflections. Step recordings were low pass filtered at

500 Hz, and each step trace is an average of at least 400 deflections.

Inner Ear Microphonic Potential Recordings

Fish 5-10 dpf were anesthetized as described above and mounted in low-melt

agarose (Thermo Fisher Scientific, Waltham, MA) on their lateral side. Larvae were

visualized using the same microscope as for lateral line microphonics with a 4x 0.1NA

objective and 40x 0.6 NA objective. The recording pipette was inserted into the otic

vesicle, in proximity to both sensory maculae. The piezo pipette was created by heat

polishing a recording pipette using a microforge CPM-2 (ALA Scientific Instruments,

Farmingdale, NY) until the tip was sealed and smooth and had a diameter of ~7 µm. The

sinusoidal command stimulus of 200 Hz was generated by jClamp and power amplified

(ENV 800, Piezosystem Jena, Jena, Germany). The stimulus was delivered by a piezo

pipette placed on the surface of the skin, approximately 50 µm away from the utricular

otolith, which was controlled by a piezo actuator (PA 4/12, Piezosystem Jena, Jena,

Germany). Resultant traces were averaged together and then low pass filtered at 2000 Hz.

Each trace shown and used in analysis is an average of at least 500 deflections.

Statistics and software

All statistical analyses were performed using GraphPad Prism 7 or RStudio. Data are reported as mean ± SEM. Statistical tests used to compare groups include Mann-

Whitney test, Student’s t-Test, Kruskal-Wallis test, or ANOVA with Holm-Sidak post hoc testing.

Results

Hair Cell Directionality of Wildtype PrimI Neuromasts

Neuromasts are composed of hair cells that are oriented 180º to one another,

making it possible to record from both populations of hair cells almost simultaneously by

using a stimulus that alternates in the positive and negative directions along this axis. We

wanted to know if anterior-facing (A) hair cells and posterior-facing (P) hair cells of

PrimI neuromasts of the lateral line had different microphonic potential amplitudes. We

designed the protocol to control for two potentially confounding factors, 1) the possibility

that one component of the stimulus, either the positive or negative deflection, is stronger

than the other, and 2) the possibility that recording from some region of a neuromast will

bias the response towards one hair cell type due to its proximity to the recording

electrode. We stimulated fish from the anterior side, then rotated the fish 180º and

recorded with the stimulus pipette on the posterior side (Figure 10a). For both of these

conditions, we took recordings with the electrode in the dorsal and ventral position, and

then in either the anterior or posterior position, whichever was opposite to the side with

the stimulus pipette (Figure 10a, b; 11c). After several trials, we discovered a bias

towards A-facing hair cells when the recording pipette was placed on the anterior side of

the neuromast, and a bias towards P-facing hair cells when the recording pipette was

placed on the posterior side (Figure 11b). Mean microphonic amplitude for A-facing cells when recording from the anterior was 14.95 ± 0.73 µV, whereas the mean microphonic amplitude for A-facing cells when recording from the posterior was 7.32 ±

0.31 µV (n = 8). Similarly, the mean microphonic amplitude for P-facing hair cells when recording from the anterior was 8.87 ± 0.49 µV, while the mean microphonic amplitude

35 for P-facing cells when recording from the posterior was 15.46 ± 1.08 µV (n = 8).

Configurations with the pipette on the dorsal or ventral side of the neuromast yielded recordings that were not significantly different for hair cells of the same polarity (Figure

11b). These results suggest that the position of the recording pipette could be a confounding factor when determining true differences in response amplitude, but only for the anterior and posterior recording positions. Consequently, all subsequent recordings were made from the dorsal and ventral side of the neuromast only.

Figure 10. PrimI wildtype directionality experiment setup. a) Schematic of experimental configurations: top-down view of neuromast with a fluid jet stimulus pipette and a recording pipette, shown in the three possible recording positions simultaneously. Stimuli are delivered parallel to the neuromast’s axis of best sensitivity, which can be determined by the position of the kinocilia (grey dots). Blue hair cells are P-facing and are depolarized when deflected towards the anterior of the animal; yellow hair cells are A- facing and depolarize when deflected posteriorly. Recordings were taken from the dorsal, ventral, and posterior side of the neuromast (Configuration I), and then the fish is rotated 180º (Configuration II) so that recordings may be repeated with the stimulus pipette on the other side. Due to the layout of the electrophysiology rig, the recording pipette cannot be placed on the same side of the neuromast as the stimulus pipette. The pipettes in the diagram and their distance from the neuromast are not to scale. b) DIC image at 100x magnification of a PLL neuromast with pipettes in proper recording position. Top panel: plane of focus for stimulus pipette, aligned with the tip of the bundle. Bottom panel: plane of focus for recording pipette, near the surface of the skin. Scale bar = 5 μm.

When comparing only recordings from the dorsal and ventral side of the neuromast, there was a difference between A- and P-facing hair cells in overall amplitude

(Figure 12a). To make sure that this difference was not simply due to an unequal number

of hair cells for each orientation, responses were averaged by the number of A- or P- facing hair cells in that neuromast to get the mean response per hair cell. P-facing hair cells had a slightly larger mean response amplitude than A-facing hair cells (Figure 12a- c) when comparing responses from the same neuromast. The mean P-facing amplitude was 1.35 ± 0.02 µV per hair cell, while the mean A-facing amplitude was 1.12 ± 0.01 µV per hair cell (n = 21). To determine if the magnitude of the discrepancy could be due to the duration of stimulation, we performed the same experiments but with longer, sustained stimuli. We found that the mean difference was indeed greater when sustained deflections were used, with P-facing hair cells still demonstrating the greater potential amplitude (Figure 12d-f). P-facing hair cells had a mean amplitude of 1.44 ± 0.03 µV per hair cell, whereas A-facing hair cells had a mean amplitude of 1.13 ± 0.02 µV per hair cell (n = 9). There was also a semi-consistent difference in response shape; P-facing hair cells appeared to have a greater steady state potential for the duration of the stimulus

(Figure 12d), but it is unclear if this is due to an actual difference between the two hair cells or simply a larger signal to noise ratio for P-facing hair cells that makes the plateau more visible (Figure 21).

Tmcs in the Inner Ear and Lateral Line

Tmc1

Since the mammalian TMC1 protein is considered essential for hearing and is a prime candidate for the pore of the MET channel, we decided to investigate its role in zebrafish hearing by recording inner ear microphonic potentials (Figure 13) of tmc1

mutants. Other members of the lab generated two different tmc1 mutants: tmc1 TLN1, which was generated via transcription activator-like effector nucleases (TALEN), and

tmc1 CR11, which was generated via clustered regularly interspaced short palindromic

repeats (CRISPR). We also acquired a third tmc1 mutant, tmc1 sa807, generated via targeting induced local lesions in genomes (TILLING) (Stemple 2004). All mutations and nomenclature are summarized in table 1. Each mutation is thought to cause a premature stop codon and truncate the protein, reducing its functionality. We tested the ear microphonics of three different mutations to make sure that any lack of phenotype was not due to residual functionality in the truncated protein. For all three mutants, the microphonic potentials from the maculae of the inner ear were comparable for homozygous mutants and wildtype or heterozygous controls (Figure 14a-c). Mean microphonic potential amplitudes were 72.22 ± 7.08 µV for tmc1 cwr5 (n = 8) and 76.48 ±

4.84 uV for their heterozygous siblings (n = 8), 90.98 ± 8.80 µV for tmc1 sa807 (n = 7) and

91.20 ± 9.14 µV for their wildtype or heterozygous siblings (n = 7), and 69.49 ± 7.41 µV

for tmc1 cwr4 (n = 5) and 75.47 ± 6.44 µV for their wildtype or heterozygous siblings (n =

6). There was also no significant difference between each of the three unique mutants in

terms of the mean amplitude of their ear microphonics response (Figure 14d).

Tmc2a and Tmc2b

Unlike mammals, zebrafish have two homologs of TMC2—Tmc2a and Tmc2b.

Since TMC2 can compensate for a loss of TMC1 in early mammalian development, we

wanted to see if Tmc2a or Tmc2b played a role in mechanotransduction in zebrafish. We

first looked at the lateral line. We previously found that most lateral line hair cells of

tmc2b mutant fish no longer had a microphonics response (Chou et al. 2017). However,

some hair cells maintained a low-level response, and some were unaffected. The tmc2b requirement of different hair cells was related to the direction they face and the neuromast in which they reside. All PrimI neuromast hair cells that retained their functionality were posterior-facing (Chou et al. 2017). To see if residual function of

PrimI neuromasts could be due to tmc2a, we tested the lateral line microphonics response of tmc2b cwr2 tmc2a cwr3 double mutant fish. We found that residual function of posterior- facing hair cells was eliminated in these mutants, as there were no detectable microphonic potentials for 4/4 fish (Figure 15a-b). Mean response amplitude from wildtype or heterozygous siblings was 9.56 ± 0.75 µV (n = 4).

We wanted to know if tmc2a and tmc2b played a similar role in the ear, so we tested the ear microphonics response of double mutants. We found that 7/10 fish had no response (Figure 15c), and the remaining fish had a very low response compared to double heterozygous siblings (mean ± SEM = 0.424 ± 0.07 μV for homozygotes vs.

96.13 ± 3.55 μV for heterozygotes, n = 12) (Figure 15d). To test if hair cell function was diminishing over time, we performed the same experiments on younger fish. The results at 5 dpf were comparable to those at 7-10 dpf, with 4/4 double homozygous mutant fish having no microphonics response (Figure 22). We tested tmc2a and tmc2b single

mutants individually, and we found that both tmc2a and tmc2b single mutants have a

detectable inner ear microphonics response (Chou et al. 2017) (Figure 16a), though it

may be somewhat ameliorated (Figure 16b). To get tmc2a single mutants, we bred tmc1

cwr4/+ tmc2a cwr6/+ fish with tmc2a cwr3/+ fish, resulting in some offspring with the genotype

tmc1 cwr4/+ tmc2a cwr3/cwr6. Single homozygous mutants of tmc2a had a mean microphonic

39 potential amplitude of 32.09 ± 1.47 μV (n = 6), whereas their heterozygous and wildtype siblings had a mean microphonic potential amplitude of 115.62 ± 2.73 μV (n = 19).

Figure 11. Recording pipette position affects microphonic potential amplitude. a) DIC image of the neuromast on which subsequent illustrations in c) are based, 100x. Scale bar = 10 µm. b) Mean potential amplitude ± SEM of A and P-facing hair cells from each of the six recording possibilities. First letter indicates hair cell population, either A-facing (yellow) or P-facing (blue). Second and third letters indicate stimulus direction—i.e. AP means pipette is positioned on anterior side and a positive deflection pushes hair cells toward the posterior. Last letter indicates recording position. ****P < 0.0001, one-way ANOVA, all groups. ***P = 0.0003, one-way ANOVA, all A-facing groups. *P = 0.0236, one-way ANOVA, all P- facing groups. **P = 0.0078, Wilcoxon test, A-PAA vs A-APP. *P = 0.0234, Wilcoxon test, P-APP vs. P- PAA. c) Illustration of the 6 recording possibilities and corresponding traces from the sample neuromast shown in a). Note bottom row where P > A when pipette is placed on posterior side (left), whereas A > P when pipette is placed on anterior side (right), even though both hair cell types are present in equal number.

Figure 12. Difference in microphonic potential amplitude of P and A-facing hair cells of PrimI posterior lateral line (PLL) neuromasts. a) Representative microphonic traces from an A-P oriented neuromast with fluid jet pipette delivering a sinusoidal stimulus, which excites hair cells of opposing orientations sequentially. Top trace corresponds to Configuration I in figure 10a, bottom trace corresponds to Configuration II in figure 10a. Recordings were taken from either the D or V position only. b) Mean microphonic response of A-facing (yellow) and P-facing (blue) hair cells to sinusoidal stimuli, divided by

41 the number of hair cells (HC) facing that direction. Bars are mean ± SEM. A-facing amplitude = 1.123 ± 0.014 μV/HC, P-facing amplitude = 1.353 ± 0.024 μV/HC; mean of differences = 0.2301 ± 0.0936 μV/HC (n = 21). *P = 0.0233, paired t-test. c) Mean microphonic amplitude per hair cell in response to sinusoidal stimuli, as pictured in b), with means from each neuromast shown individually. d) Sample microphonic traces from an A-P oriented neuromast with fluid jet pipette delivering steps of sustained deflection. Top trace corresponds to configuration I in figure 10a, bottom trace corresponds to configuration II in figure 10a. Recordings were taken from either the D or V position only. e) Mean microphonic response of A- facing (yellow) and P-facing (blue) hair cells to step stimuli, divided by the number of hair cells facing that direction. Bars are mean ± SEM. A-facing amplitude = 1.131 ± 0.025 μV/HC, P-facing amplitude = 1.436 ± 0.032 μV/HC; mean of differences = 0.3052 ± 0.0948 μV/HC (n =9). *P = 0.0123, paired t-test. f) Mean microphonic amplitude per hair cell in response to step stimuli, as pictured in e), with means from each neuromast shown individually.

Tmc1, Tmc2a, and Tmc2b

Although tmc2b cwr2 tmc2a cwr3 fish have no ear microphonics response, they do have an inconsistent acoustic startle response, and some hair cells of the inner ear still take up FM1-43 dye, which enters cells through functional MET channels. Members of our lab generated fish with mutations in tmc1, tmc2a, and tmc2b using multiplex genome editing (Minkenburg et al. 2017) to see if the homozygous fish would demonstrate an inability to hear for all of these assays. tmc2b cwr8 tmc1 cwr4 tmc2a cwr6 triple mutants had no lateral line microphonics response from the IO4 neuromast (n = 6) (Figure 17a, b) and no detectable ear microphonics response from the maculae (n = 10) (Figure 17c, d).

The heterozygous and wildtype controls exhibited a normal response amplitude of 6.75 ±

0.45 µV (n = 5) for the IO4 neuromast and 82.20 ± 3.37 μV (n = 10) for the inner ear.

Other members of our lab discovered that the triple homozygotes do not have an acoustic startle response, nor do their hair cells take up dye through MET channels, and thus multiple assays suggest that they are unable to hear or detect water motion.

Table 1. Summary of genotypes.

Gene Allele Exon Region Genotype Protein DNA Missing Change tmc1 cwr4 exon7 CRISPR11 tmc1 cwr4 568aa -8bp (990aa) cwr5 exon5 TALEN1 tmc1 cwr5 668aa -1bp

sa807 exon19 tmc1 tmc1 sa807 82aa C>T tmc2a cwr6 exon9 CRISPR4 tmc2a cwr6 553aa -3bp, +2bp (916aa) cwr3 exon9 CRISPR4 tmc2a cwr3 553aa -2bp tmc2b cwr8 exon6 CRISPR2 tmc2b cwr8 615aa -5bp (892aa) cwr2 exon6 CRISPR3 tmc2b cwr2 615aa -5bp

Figure 13. Ear microphonics recording setup. a) DIC image of larval zebrafish mounted on lateral side, 4x magnification. Piezo (stimulus) pipette is on the left, recording pipette is on the right. Square shows region magnified in b). b) Inner ear, 40x magnification. Ovular shapes are the otoliths, which cover the hair cells of the maculae. The recording pipette is positioned between the anterior and posterior maculae. White outline shows relative position of piezo pipette on the x and y axes. The piezo pipette is angled at ~15º with respect to the petri dish, and moves back and forth in the dorsal and ventral direction during stimulation (arrows).

Figure 14. Inner ear microphonic potentials of tmc1 mutant fish resemble wildtype. Representative microphonics traces for a) tmc1 cwr5, b) tmc1 sa807, and c) tmc1 cwr4. d) Graph of mean potential amplitude ± SEM for each mutant; tmc1 cwr5 = 72.22 ± 7.08 μV (n = 8), heterozygous siblings = 76.48 ± 4.84 μV (n = 8) (P = 0.3823); tmc1 sa807 = 90.98 ± 8.80 μV (n = 7), wildtype or heterozygous siblings = 91.20 ± 9.14 μV (n = 7) (P = 0.9015); tmc1 cwr4 = 69.49 ± 7.41 (n = 5), wildtype or heterozygous siblings = 75.47 ± 6.44 (n = 6) (P = 0.9307). P = 0.9710, one-way ANOVA. Pairwise significance determined by Mann-Whitney test.

Figure 15. tmc2a tmc2b double mutants have no detectable microphonic potentials from lateral line or ear. a) Representative traces from lateral line of tmc2b cwr2 tmc2a cwr3 mutant fish and controls. There was no response from PrimI neuromasts of double homozygous mutants. b) Mean ± SEM of lateral line microphonic potentials for tmc2b cwr2 tmc2a cwr3 and controls. Heterozygous and siblings = 9.56 ± 0.75 µV (n = 4); double mutants = 0 ± 0 uV (n = 4). *P = 0.0286, Mann-Whitney test. c) Representative traces from maculae of tmc2b cwr2 tmc2a cwr3 mutant fish and controls. d) Mean ± SEM of ear microphonic potentials for tmc2b cwr2 tmc2a cwr3 and controls. tmc2b cwr2 tmc2a cwr3 = 0.424 ± 0.07 µV (n = 10), heterozygotes = 96.13 ± 3.55 μV (n = 12). ****P < 0.0001, Mann-Whitney test.

Figure 16. tmc2a mutant fish exhibit reduced inner ear microphonic potentials. a) Representative microphonic potential traces from maculae of tmc2a cwr3/cwr6 fish and controls. b) Mean potential amplitude ± SEM of tmc2a cwr3/cwr6 and controls. tmc1 cwr4/+ tmc2a cwr3/cwr6 = 32.09 ± 1.47 μV (n = 6); heterozygous and wildtype siblings = 115.62 ± 2.73 μV (n = 19). ****P < 0.0001, Mann-Whitney test. Both tmc1 tmc2a single and double heterozygous siblings were included as controls, as no significant difference was observed between these groups (P = 0.7209).

Figure 17. Fish without functional tmc1, tmc2a, or tmc2b cannot hear or detect water motion. a) Representative traces from IO4 neuromasts of triple mutant and control fish. b) Mean ± SEM of lateral line microphonic potentials from triple mutants and controls. tmc2b cwr8 tmc1 cwr4 tmc2a cwr6 = 0 ± 0 μV (n = 6), wildtype or heterozygous siblings = 6.75 ± 0.45 μV (n = 5). **P = 0.0022, Mann–Whitney test. c) Representative traces from maculae of triple mutant and control fish. d) Mean ± SEM of ear microphonic potentials for triple mutants and controls. tmc2b cwr8 tmc1 cwr4 tmc2a cwr6 = 0 ± 0 μV (n = 10), wildtype controls = 82.20 ± 3.37 μV (n = 10). ****P < 0.0001, Mann-Whitney test.

Discussion

Channel Composition May Vary Between Hair Cells of the Lateral Line

We demonstrated a difference in microphonic potential amplitude between hair cells of the lateral line that face opposing directions. Before anterior and posterior recording positions were omitted, the primary difference in amplitude seemed to be based on the number of hair cells of that polarity in proximity to the pipette. This result is consistent with what others have suggested (Olt et al. 2016). However, we continued experiments using the dorsal and ventral recording position to see if an effect could still be observed. We found that P-facing hair cells have a higher mean amplitude than A- facing hair cells. This indicates some physical difference between the two hair cell types.

A potential difference between the two subpopulations could be rigidity or the average length of the bundles. Testing the stiffness of the stereocilia would be difficult, due to their encapsulation by the cupula and their small size, but differences in rigidity could correlate with greater concentration of actin or increased utilization of actin bundling proteins, which could potentially be labeled fluorescently and quantified. Mammalian hair cells utilize the actin bundling proteins fascin-2 and espin (Shin et al. 2010; Zheng et al. 2000), and zebrafish inner ear hair cells possess analogs of these proteins, whose concentrations appear to correlate with stereocilia length (Chou et al. 2011). For hair cells with otherwise equivalent properties, longer stereocilia should correspond to greater activation at lower frequencies (Kandel 2013), so it is possible that one subset of hair cells could have longer stereocilia and be optimized to a different frequency of motion than the other. This seems unlikely, however, when considering that sustained deflection of a lower frequency still results in a lower A-facing amplitude (Figure 12d-f).

Additionally, the primary determinant of characteristic frequency between hair cells of different neuromasts, which varies by less than 100 Hz, appears to be cupula height

(Trump & McHenry 2008), but hair cells within a neuromast share the same cupula.

Stereocilia heights have been measured for the lateral line, but measurements from the two polarities are usually not distinguished or compared, perhaps due to the relatively small length of the stereocilia, which is about 1.2 µm on average (Schuler et al. 2013).

Morphants that have reductions in mean stereocilia length as small as 0.2 µm experience swimming defects (Schuler et al. 2013), suggesting that there might be a narrow range of functional stereocilia lengths for the lateral line, and if so, height would not vary markedly. Regardless, it may be worth performing the same set of microphonic potential recordings at different stimulation frequencies to see if the ratio of A-facing to P-facing responses stays consistent, as this could point to a difference in bundle morphology.

Another possible difference between the two cell polarities could be electrical resonance; if A-facing and P-facing hair cells had differences in basolateral calcium or potassium channels, this would also cause them to have different characteristic frequencies and thus different levels of sensitivity to the same stimulus. This too could be tested by varying the frequency of microphonic stimulation, but would require an intracellular recording technique such as patch clamping to distinguish from other methods of frequency tuning. Tuning via electrical resonance has been established in the ears of goldfish (Sugihara & Furukawa 1989) and toadfish (Steinacker & Romero 1991), but not in a lateral line system. It is unclear if tuning the lateral line in this way would be functionally beneficial for the animal, since these hair cells detect low frequencies of motion in a limited range compared to those of the fish’s ear. The basolateral currents of

lateral line hair cells have been dissected via pharmacological antagonists and voltage

clamp, but A-facing and P-facing hair cells were both patched indiscriminately and not

compared (Olt et al. 2014).

One last explanation for the discrepancy between the two polarities could be that

the MET channel proteins are different between A-facing and P-facing hair cells. A change in protein composition would reasonably correspond to a change in conductance of the channel, which would cause the microphonic potential amplitudes to vary.

Different proteins could also create a change in the kinetic properties of the channel, although precise measurements of activation and adaptation time constants would need to be done with a larger sample size. A change in protein composition of the MET channel could allow one subset of hair cells to have higher conductance at all frequencies of motion instead of making each optimized to a different frequency, or it could also contribute to frequency tuning via changes in channel kinetics. The lateral line detects water velocity and acceleration (Kroese et al. 1992; Otieza et al. 2017) in addition to low-frequency vibration. Detection of water velocity by the lateral line appears to rely upon fluctuations in flow created by the fish’s body or upstream obstacles (Chagnaud et al. 2008a; 2008b), as a single hair cell will adapt quickly to a prolonged deflection, even if optimized for relatively low frequencies. The fish most likely integrates responses from many neuromasts to determine water velocity or acceleration, using the timing, degree, and position of neuromast activation to understand its surrounding environment. It is possible that small differences in MET channel activation and adaptation time constants between hair cells of a single neuromast could fine tune the temporal component in

50 estimating local water velocity, or that differences in overall conductance could affect the weighting of some hair cells in their contribution to the spatial component.

Whether the discrepancy in response between the two subpopulations of hair cells is due to an electrical or morphological factor, the existence of the discrepancy suggests a physiological need for a difference in mechanosensitivity between the two polarities. It makes intuitive sense that a fish would need to be less sensitive to deflections that move from anterior to posterior; zebrafish swim against a current during rheotaxis (Olszewski et al. 2012), and so this would be a consistent and non-threatening stimulus for them.

However, deflections that travel from the posterior to anterior could be created by a predator behind them, so it would make sense that cells that detect this motion might be designed with higher intrinsic sensitivity. Additionally, in measuring water flow fluctuations around the body of a goldfish, it was found that water flowing from the anterior to the posterior had more fluctuations than a flow traveling from the posterior to anterior (Chagnaud et al. 2008b), likely due to asymmetry in the fish’s body shape. If this is true of zebrafish as well, it might require P-facing hair cells, which depolarize when deflected towards the anterior, to be slightly more sensitive in order to sense water velocity to the same degree as their A-facing counterparts.

The hair cells of our own ears, through various means, are tuned to enhance our survival. We have a range of frequencies to which we are the most sensitive because they are the most salient in our natural environment, and this range is different from the optimal frequency range of other animals, e.g. mice, who are sensitive to higher frequency noises, since they communicate using high frequency sounds (Heffner &

Heffner 2007). Fish are organized differently from mammals, as their survival depends

51 upon detection of water velocity and acceleration in addition to the ability to hear sounds.

They do not have analogues of OHCs that can amplify slight water movements, so it makes sense that they might change the physical properties of their existing hair cells to meet their physiological needs. However, these changes are probably similar to molecular strategies mammals use for encoding stimuli, as we share a common vertebrate ancestor and it is more likely that we would reuse a motif rather than invent an entirely new one, and it is likely that they would conserve a method that works. It will take further experimentation to figure out the exact means utilized to tune mechanotransduction of the lateral line.

Tmcs May Combine to Form the MET Complex

Our results suggest that Tmc1 alone is not required for hearing in zebrafish, unlike what is observed in mammals. Neither Tmc2a nor Tmc2b on their own appear to be absolutely necessary for hearing or water motion detection, though Tmc2b seems to be more important in hair cells of the posterior lateral line, since PrimII neuromasts lose all function while less than half of PrimI hair cells respond, and neuromasts such as IO4 are unaffected (Chou et al. 2017). Single mutants of either tmc2a or tmc2b have a measurable ear microphonics response, though it appears to be lower than that of wildtype, at least for tmc2a. Mutation of both tmc2a and tmc2b eliminates the lateral line and almost all of the inner ear microphonics response, and the fish seem at least partially deaf by startle assays and dye injection. Mutation of tmc1, tmc2a, and tmc2b simultaneously results not only in the elimination of the lateral line and inner ear microphonics response, but the fish do not startle and their hair cells do not take up FM1-43 dye, indicating that the fish

are completely deaf and unable to sense water flow. These results indicate that Tmc2a

and Tmc2b work together to allow for zebrafish mechanotransduction, and that Tmc1 may have only an auxiliary function, perhaps allowing for low levels of residual hair cell

activity in the tmc2a tmc2b double mutants. The redundancy of Tmcs is almost reversed

with respect to mammals, for whom lack of TMC1 can be partially supplemented by

TMC2 to allow for residual function before P10 (Kurima et al. 2015), but who seem to

experience no reduction in cochlear MET current when TMC2 is eliminated (Kawashima

et al. 2015). It seems that larval zebrafish are more similar to embryonic mammals with

respect to proteins utilized for mechanotransduction, and so advances made in zebrafish

mechanotransduction may be most applicable to developing mammals. Data potentially

in agreement with this show that zebrafish lateral line and inner ear hair cells have a

complement of basolateral potassium and calcium currents that resemble immature

mammalian vestibular and auditory systems (Olt et al. 2014). However, it is worth noting

that mature mechanotransduction currents can be observed in the mammalian utricle

(Geleoc & Holt 2003) and OHCs (Lelli et al. 2009) even before the hair cells have fully

matured (Rusch et al. 1998; Marcotti & Kros 1999). Although zebrafish maculae are

responsible for both hearing and vestibular function, it is also possible that their macular

hair cells are more similar to mammalian vestibular hair cells with respect to

mechanotransduction, as these hair cells seem to be more reliant on TMC2 (Kawashima

et al. 2011).

One relevant question for the data presented surrounds what may cause the

discrepancy between the microphonics data and other hearing assays for the tmc2a tmc2b

double mutants. If these fish have a partial startle response and partial dye uptake, why

53 do they not exhibit a larger electrical response from the ear? One possibility is that hair cells retaining functionality are confined at a site in the maculae too far from the recording pipette to be detected. This seems plausible given that microphonic potential recordings from neuromasts can be biased toward the hair cells most proximal to the electrode and therefore biased against the most distal hair cells (Figure 11b; 18; 19).

Most, if not all, of the inner ear microphonics response should be from the posterior/saccular macula, but moving the recording electrode further from the posterior macula does not appear to significantly affect the amplitude of the microphonics response as long as the pipette is still in the otic vesicle (Yao et al. 2016; Tanimoto et al. 2011).

However, those results were obtained using wildtype fish at ~7 dpf, possessing approximately 80 saccular hair cells (Yao et al. 2016; Haddon & Lewis 1996). The tmc2a tmc2b mutants may have few enough functional hair cells close to the otolith’s edge that the signal to noise ratio is too low for extracellular potentials to be recorded. Only a few hair cells should be required to initiate a startle response, whereas many more will be required for an extracellular electrode to pick up ionic flux near their apical surface. One experiment that should be done is to increase the intensity of the stimulus for tmc2a tmc2b double mutants, as microphonic potential amplitude can increase with greater displacements (Lu & DeSmidt 2013).

The Tmc results may be related to the observation that wildtype P-facing and A- facing lateral line hair cells have different microphonic potential amplitudes. For tmc2b mutants, the only PrimI PLL hair cells that still show a microphonics response are the posterior facing hair cells (Chou et al. 2017), and tmc2a tmc2b double mutants have no response from any hair cells. This suggests that P-facing hair cells could have MET

channels that incorporate both Tmc2a and Tmc2b, whereas A-facing hair cells utilize

only Tmc2b. If we assume that it is more important for fish to be able to detect a stimulus

that flows from posterior to anterior, it makes sense as an evolutionary strategy to use two

unique proteins in the MET channel, so that if one is rendered useless by a random

mutation, the other protein may still compensate to some degree. Also, the incorporation

of Tmc2a and Tmc2b could explain why P-facing hair cells have a higher microphonic

potential amplitude. Perhaps Tmc2b, or the Tmc2a-Tmc2b complex, has a higher

conductance than Tmc2a alone. There should at least be some change to the biophysical

properties of the channel when another distinct protein is added, even if the conductance

is somehow unchanged. Other aspects that might change include activation and

adaptation time constants. Studies that directly measure single channel conductance or

the time course of MET currents in the presence or absence of these two proteins should

be done to know what effect their association might have.

Studying the TMC/Tmc proteins directly is unfortunately complicated by our

current inability to insert them in the membrane of a heterologous cell, a lack of resolved

structure that indicates a definitive pore region, and the difficulty of recording single

channel currents from a stereocilium that is less than a micron in diameter. In lieu of

these major advances, it would be helpful to record lateral line microphonic potentials

from tmc1 and tmc2a single mutants, as this would let us know if tmc1 or tmc2a single

mutants have reduced amplitudes from any hair cells of the lateral line. Similarly, inner

ear microphonic amplitudes should be more rigorously measured for tmc2b single

mutants to determine if they respond at reduced levels, as is seen with tmc2a mutants.

The results of these experiments would shed light on the association between these three

Tmc proteins and how it varies across the mechanosensitive map of the larval zebrafish.

This will help reveal how hair cells with different physiological roles are tuned and differentiated at the level of MET channels, which could occur in mammalian ears as well.

Appendix

Figure 18. Distance from recording pipette to closest hair cell for sample neuromast. DIC images of neuromast shown in figure 11 with recording pipette in the proper position for each configuration. First and second letter indicate the position of the stimulus pipette. AP: stimulus pipette is on the anterior side, positive deflection pushes toward the posterior of the fish; PA: stimulus pipette is on the posterior side, positive deflection pushes toward the anterior of the fish. Last letter indicates position of recording pipette. D: dorsal, V: ventral, A: anterior, P: posterior. Gray scale bar = 10 µm. Yellow and blue bars indicate the approximate distance from the center tip of the recording pipette to the center of the nearest A-facing or P- facing hair cell, respectively.

Figure 19. Distance from recording pipette to nearest A-facing and P-facing hair cell, as ratio of A:P. For each recording configuration, the distance from the center tip of the recording pipette to the center of the nearest A-facing and P-facing hair cell was measured. Bars show mean ± SEM of the A:P distance ratio. APD = 1.13 ± 0.05 (n = 10), APV = 1.11 ± 0.05 (n = 9), APP = 2.32 ± 0.11 (n = 6), PAD = 1.05 ± 0.02 (n = 12), PAV = 1.08 ± 0.03 (n = 13), PAA = 0.44 ± 0.01 (n = 6). **P = 0.0016, ****P < 0.0001. Significance determined by Mann-Whitney test.

Figure 20. Number of anterior or posterior-facing hair cells per PrimI neuromast for wildtype fish 5- 7 dpf. a) Hair cell numbers for fish given sinusoidal stimuli. Number of A-facing hair cells = 7.10 ± 0.08, number of P-facing hair cells = 7.19 ± 0.08 (n = 21) (P = 0.4100) b) Hair cell numbers for fish given step stimuli. Number of A-facing hair cells = 5.1 ± 0.1, number of P-facing hair cells = 5.8 ± 0.2 (n = 21) (P = 0.8710). Significance determined by Mann-Whitney test.

Figure 21. Average of all step stimulus traces. When the microphonic potentials are averaged over 7,000 deflections for each stimulation direction, the steady state response becomes more visible.

Figure 22. tmc2b tmc2a double mutant microphonics response at 5 dpf. a) Representative traces from tmc2b cwr2 tmc2a cwr3 fish and tmc2b cwr2/+ tmc2a cwr3 siblings. b) Mean microphonic amplitudes for tmc2b tmc2a mutants and controls. Mean microphonics response from mutants = 0 ± 0 µV (n = 4), mean microphonic amplitude from tmc2b cwr2/+ tmc2a cwr3 siblings = 49.09 ± 5.46 µV (n = 4).

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