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Long-term maintenance of developmental enhancers enables hair cell regeneration in the zebrafish inner ear

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Long-term maintenance of developmental enhancers enables hair cell regeneration in the zebrafish inner ear

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Content Long-term Maintenance of Developmental Enhancers Enables
Hair Cell Regeneration in the Zebrafish Inner Ear
By
Tuo Shi
A Dissertation Presented to the
FACULTY OF THE USC GRADUATE SCHOOL
UNIVERSITY OF SOUTHERN CALIFORNIA
In Partial Fulfillment of the
Requirements for the Degree
DOCTOR OF PHILOSOPHY
(DEVELOPMENT, STEM CELLS, AND REGENERATIVE MEDICINE)
May 2025
Copyright 2025 Tuo Shi

ii
Acknowledgements
This manuscript is dedicated to Dr. Neil Segil, without whom this thesis project would
not have existed. Neil encouraged me to continue to pursue my interest in
understanding of hair cell regeneration and offered his mentorship even during his
grueling battle with cancer. His selflessness and care for young minds became what I
strived for when I mentored my own students. This project also would not be so fruitful
without the mentorship of Dr. Ksenia Gnedeva and Dr. Gage Crump. I’m so grateful for
Ksenia to take over for Neil in his passing as a mentor and an expert in inner ear
biology. Her insight and support have been crucial in the successful and timely
completion of this dissertation project. I’m also thankful for Gage for allowing me to
pursue my own interest and project in his lab, starting a brand new inner ear project in
his lab without even any second thoughts. To both Gage and Ksenia, I also really
appreciate the independence and trust that they gave me to drive this dissertation
project along, which really shaped me into the scientist I am today. In addition to the
science, both of them also really cared for and supported other aspects of my Ph.D.
journey. Without their words of encouragement during my self-doubt and difficult times,
their support for my mental health and dealing with burnout, their advice on scientific
career and job searching, I would have had a much more difficult experience getting
where I am right now. I’m forever grateful to the amazing mentors I had during this
journey – I could not have asked for better support along the way.
I would also like to thank the amazing scientists and friends I met along the way. My
collaborators Marielle Beaulieu and Dr. David Raible at University of Washington, it was

iii
such a fun collaboration and valuable exercise in integrating and comparing crossspecies transcriptome data. Thank you Leah Kim and Juan Llamas for helping out with
the mouse experiments and also providing moral support during the writing of our
manuscript. Thank you Dr. Thomas Lozito also for providing lizards for my project. I
would like to give special thanks to Dr. Peter Fabian, who was the best mentor anyone
could have asked for. He made science and every day in the Crump lab so much fun.
Thank you Dr. Litao Tao, Dr. Vincent Yu, Dr. Emily Wang, Dr. Talon Trecek, Dr. Olivia
Chen and Ted Tseng for helping me get started with bioinformatics analysis. Thank you
Dr. Kelsey Elliott for taking your time to help me whenever I needed help, even though
you were busy yourself. Thank you Dr. D’Juan Farmer and Dr. Mathi Thiruppathy for
spending hours and hours discussing my project with me and showing your interest in
what I was doing. Special shout out to Oluchi Ofoegbu, who also spent hours and hours
chatting with me, but about non-science related matters, without which I probably could
have graduated earlier. Her presence and friendship made the last few years of my
Ph.D. so much more enjoyable and less stressful, so it was all worth it! Another special
shout out to Gio Kim, who was the best high school student, or student, period, that I
had. I wish you the best in pursuing your future goals at Johns Hopkins University!
Last but not least, I’d like to thank my committee chair Dr. Oliver Bell and his advice and
supporting during my committee meetings. Thank you Megan Matsutani for maintaining
our zebrafish and putting up with my last minute order requests. Thank you Bernadette
Masinsin for helping out with FACS. Thank you NIDCD for funding me through my entire
Ph.D.

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Table of Contents
Acknowledgements...................................................................................................................... ii
List of Figures .............................................................................................................................. vi
Abstract .................................................................................................................................... ix
Chapter 1. Introduction................................................................................................................ 1
1.1 Auditory and vestibular organs in the inner ear....................................................... 1
1.2 Permanent hair cell loss in mammals and hearing loss......................................... 2
1.3 Development and regeneration of inner ear hair cells ........................................... 2
1.4 Zebrafish lateral line neuromasts .............................................................................. 5
1.5 The role of epigenetics in hair cell regeneration ..................................................... 6
Chapter 2. Single-cell transcriptomic profiling of the zebrafish inner ear reveals
molecularly distinct hair cell and supporting cell subtypes................................. 9
2.1 Background ................................................................................................................... 9
2.2 Zebrafish inner ear hair cells and supporting cells are transcriptionally
distinct from those of the lateral line ....................................................................... 12
2.3 Single-cell RNA-seq reveals distinct hair cell and supporting cell populations
in the juvenile and adult inner ear of zebrafish ..................................................... 16
2.4 Developmental trajectories in the zebrafish inner ear.......................................... 22
2.5 Distinct supporting cell types in the cristae versus maculae............................... 25
2.6 Distinct types of hair cells in the zebrafish inner ear ............................................ 28
2.7 Global homology of striolar and extrastriolar hair cells between fish and mice 34
2.8 Discussion ................................................................................................................... 37
Chapter 3. Long-range Atoh1 enhancers maintain competency for hair cell
regeneration in the inner ear................................................................................. 43
3.1 Background ................................................................................................................. 43
3.2 Single-cell profiling of the zebrafish inner ear identifies distinct classes of
atoh1a enhancers...................................................................................................... 44
3.3 Differential activity of atoh1a enhancers in hair cells and supporting cells....... 47
3.4 Notch-dependent repression of Class 2 enhancer activity in supporting cells
through the atoh1a promoter ................................................................................... 51
3.5 Differential requirements of Class 1 and 2 enhancers for hair cell
development in the inner ear and lateral line ........................................................ 53
3.6 Selective requirement of Class 2 enhancers for inner ear hair cell
regeneration ............................................................................................................... 57
3.7 Cross-species activity of Class 2 enhancers ......................................................... 63
3.8 Potential role of SOX transcription factors in Class 2 enhancer activity ........... 68

v
3.9 Discussion ................................................................................................................... 74
Chapter 4. Conclusions............................................................................................................. 81
4.1 Zebrafish lateral line versus inner ear as a model to study hair cell
development and regeneration................................................................................ 81
4.2 Challenges to using the zebrafish inner ear .......................................................... 82
4.3 Multitude of epigenetic barriers to hair cell regeneration..................................... 83
4.4 Reestablishment of Class 1 enhancers in zebrafish supporting cells................ 84
Materials & Methods .................................................................................................................. 86
References ................................................................................................................................ 109

vi
List of Figures
Figure 1. Anatomy of the human ear............................................................................... 1
Figure 2. Hair and supporting cell patterning through Notch-mediated lateral inhibition . 4
Figure 3. Anatomy of zebrafish and mouse inner ears.................................................. 10
Figure 4. Molecularly distinct cell types between the zebrafish inner ear and lateral
line.................................................................................................................. 13
Figure 5. Gene modules for embryonic to larval inner ear and lateral line dataset........ 14
Figure 6. Selection of otic sensory cells from snRNA-seq dataset ................................ 15
Figure 7. scRNA-seq of 12 mpf zebrafish inner ear captures sensory hair cells and
supporting cells as well as non-sensory supporting cells ............................... 17
Figure 8. Cell subtypes in the zebrafish inner ear end organs ...................................... 18
Figure 9. Hair cell and supporting cell marker expression in the integrated
scRNA-seq dataset ........................................................................................ 19
Figure 10. Putative progenitor marker expression in individual progenitor and
supporting cell clusters................................................................................. 20
Figure 11. Gene modules for integrated inner ear sensory patch dataset..................... 21
Figure 12. Pseudotime analysis reveals developmental trajectories in the zebrafish
inner ear....................................................................................................... 23
Figure 13. Pseudotime analysis of cristae hair and supporting cells in the zebrafish
inner ear....................................................................................................... 24
Figure 14. dla labels putative hair cell progenitors in the cristae and maculae.............. 25
Figure 15. Distinct markers separate macula and crista supporting cells...................... 26
Figure 16. zpld1a and tectb are primarily expressed in supporting cells ....................... 27
Figure 17. Gene expression differences between lateral line and inner ear hair cells... 28

vii
Figure 18. cabp1b+and cabp2b+label hair cells in distinct regions of sensory end
organs .......................................................................................................... 30
Figure 19. Distinct markers separate macula and crista hair cells ................................ 31
Figure 20. skor2 and loxhd1b label subsets of hair cells in utricle or saccule ............... 31
Figure 21. Zebrafish cabp2b+domain shares features with the mouse striolar region .. 32
Figure 22. Striola marker pvalb9 is expressed in all inner ear sensory end organs ...... 33
Figure 23. Inner ear hair cell subtypes differentially express mechanosensory
apparatus genes .......................................................................................... 33
Figure 24. Inner ear hair cell subtypes differentially express voltage-gated calcium
and potassium channel genes...................................................................... 34
Figure 25. SAMap analysis reveals conserved gene expression patterns between
mouse and zebrafish hair cell types ............................................................. 36
Figure 26. SAMap analysis of mouse utricle versus zebrafish macular and lateral line
cells.............................................................................................................. 37
Figure 27. Inner ear enhancer architecture of the zebrafish atoh1a locus..................... 45
Figure 28. Single-cell analysis of the zebrafish inner ear .............................................. 46
Figure 29. Cell type-specific reporter activities of zebrafish atoh1a enhancers............. 49
Figure 30. Additional analysis of atoh1a enhancer transgenic lines.............................. 50
Figure 31. Notch inhibition of eh7 enhancer activity through the endogenous atoh1a
promoter....................................................................................................... 52
Figure 32. Requirements of Class 1 and 2 atoh1a enhancers in HC development ....... 54
Figure 33. Additional analysis of atoh1a enhancer deletion alleles ............................... 56
Figure 34. Requirements of Class 1 and 2 atoh1a enhancers in HC regeneration ....... 58

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Figure 35. atoh1a and lfng co-localization in the uninjured and regenerating utricle
and evidence for SC transdifferentiation ...................................................... 62
Figure 36. Lizard saccule multiome............................................................................... 64
Figure 37. Comparative atoh1a enhancer architecture and activity across species...... 66
Figure 38. Enhancer testing controls in the mouse cochlea .......................................... 67
Figure 39. Global analysis of Class 2-like enhancers in zebrafish and mouse.............. 70
Figure 40. Analysis of Class 1 and 2 peaks in mouse sensory progenitors, design of
SOX motif deletion, and gene pairs with Class 1 and 2 peaks..................... 73

ix
Abstract
The loss of inner ear mechanosensory hair cells is permanent in mammals, which is a
major contributing factor to deafness in humans. Conversely, non-mammalian
vertebrates such as zebrafish can replenish hair cells throughout life from the
transdifferentiation of neighboring supporting cells. The understanding of hair cell
development and regeneration in zebrafish has largely been investigated through the
lateral line neuromasts due to their surface localization for drug treatment and imaging
access. However, through single-cell transcriptomic profiling of the zebrafish inner ear,
we find a closer resemblance of zebrafish inner ear hair and supporting cells to their
mammalian counterparts than those of neuromast origin. Additionally, we uncover a
group of hair cell gene enhancers required for hair cell development and regeneration
specifically in the inner ear. These enhancers remain accessible in adult zebrafish inner
ear supporting cells, potentially enabling their life-long ability to transdifferentiate into
hair cells upon damage. Conversely, a syntenic group of enhancers found in mouse
cochlear supporting cells becomes permanently silenced postnatally, correlating to their
loss of transdifferentiation potential. Cross-species enhancer testing also reveals that
while both mouse and zebrafish versions of a sequence-conserved enhancer can
continuously drive fluorescent reporter activities in juvenile zebrafish supporting cells,
they become quickly silenced in postnatal mouse cochlea, regardless of their species of
origin. This suggests that differences in upstream regulators in mouse versus zebrafish
supporting cells are responsible for their contrasting abilities to maintain these
regenerative enhancers and allow for hair cell regeneration.

1
Figure 1. Anatomy of the human ear
Chapter 1. Introduction
1.1 Auditory and vestibular organs in the inner ear
The inner ear is a sensory organ responsible for maintaining balance and detecting
sound. The vestibular system, consisting of three semicircular canals, the utricle, and
the saccule, is where linear and rotational head movement is detected. The organ of
Corti residing within the snail-shaped cochlea, on the other hand, is responsible for
capturing auditory inputs. All of these sensory organs are comprised of two major cell
types: the
mechanosensory
hair cells and their
surrounding
supporting cells.
Both vestibular and
auditory inputs can
deflect hair bundles
on the apical
surface of
hair cells, opening
up
Figure 1. Anatomy of the human ear. (A) The human ear consists of the outer ear and ear
canal, middle ear ossicle bones, and the inner ear sensory organs. (B) Cross section of the
cochlea shows fluid filled scala vestibuli, scala media, and scala tympani, and the hearing
organ, the organ of Corti. (C) Patterning of the organ of Corti shows three rows of outer hair
cells and one row of inner hair cells, surrounded by different supporting cell subtypes. (D) A
typical hair cell has hair bundle on the apical surface of the cell, which can be deflected in
response to sensory inputs. Modified from Dror et al. 2010.

2
mechanotransduction channels on the cell surface and allowing an influx of K+. This
results in depolarization of hair cells and sensory input through the vestibular or spiral
ganglion to the central nervous system (Figure 1) (Durrant and Lovrinic, 1984; Purves et
al., 2001a; Purves et al., 2001b).
1.2 Permanent hair cell loss in mammals and hearing loss
Age-related hearing loss is one of the most common disabilities faced by the aging
population in the United States, and can be contributed to the irreversible damage to the
mechanosensory hair cells that reside within the inner ear cochlea (Corwin and
Cotanche, 1988; Hoffman et al., 2017). In mice, even though damaged cochlear hair
cells can be replaced through the transdifferentiation of neighboring supporting cells at
neonatal stage, this regenerative ability of supporting cells is quickly lost within the
postnatal first week (Doetzlhofer et al., 2009; Kelly et al., 2012; Maass et al., 2015;
White et al., 2006; Zhao et al., 2011). In contrast, supporting cells of adult nonmammalian vertebrate animals retain the ability to transdifferentiate into hair cells
following hair cell injury (Cafaro et al., 2007; Corwin and Cotanche, 1988; Monroe et al.,
2015). This has sparked great interests in understanding the fundamentals of hair cell
development, as well as the mechanism behind lifelong hair cell regeneration in nonmammalian vertebrates, with the hope that we can one day enable hair cell
regeneration in adult humans and reverse hearing loss.
1.3 Development and regeneration of inner ear hair cells

3
During development, the inner ear arises from a specialized region of head ectoderm
called the otic placode, which invaginates into the head to form the otic vesicle, with its
dorsal region becoming the vestibular system while the ventral region becoming the
cochlea (Anniko and Wikström, 1984; Douarin, 1986; Morsli et al., 1998). In these
sensory organs, hair and supporting cells are derived from a common prosensory
progenitor population that expresses the transcription factor Sox2 (Doetzlhofer et al.,
2006; Kempfle et al., 2016; Kiernan et al., 2005). In addition to the specification of
prosensory domain via SOX2, another two HMG domain transcription factor family
members, SOX4 and SOX11, are also required to establish the hair cell competence of
prosensory progenitors (Wang et al., 2023). Although how each progenitor is selected to
become hair versus supporting cell is still largely unknown, it has been established that
the transcription factor ATOH1 plays a critical role in committing the progenitors towards
a hair cell fate (Bermingham et al., 1999; Chen et al., 2002; Pan et al., 2011; Woods et
al., 2004). Initially during development, a low wave of Atoh1 expression is detected
across prosensory progenitor cells prior to the onset of hair cell differentiation (Cai and
Groves, 2015; Cai et al., 2013; Chen et al., 2002). In some of these progenitors, Atoh1
expression becomes further upregulated, leading to the expression of various other hair
cell-related genes, including another required hair cell lineage transcription factor,
Pou4f3 (Helms et al., 2000; Masuda et al., 2011; Yu et al., 2021), and Notch ligands
Dll1 and Jag2 (Kiernan et al., 2006; Lanford et al., 1999). The expression of Notch
ligands in hair cells activates Notch signaling and the expression of Hes and Hey
transcription factors in surrounding progenitors via cell surface Notch receptors. HES
and HEY factors can then repress and silence Atoh1 expression in these surrounding

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Figure 2. Hair and supporting cell patterning through Notch-mediated lateral inhibition
cells by binding to the Atoh1 promoter, diverting them towards a supporting cell fate
(Figure 2) (Abdolazimi et al., 2016; Doetzlhofer et al., 2009; Zheng et al., 2000).
In addition to its role in development, Atoh1 also plays an integral part to hair cell
regeneration in non-mammalian vertebrates. During hair cell regeneration, Atoh1 is
among one of the first genes to be upregulated in supporting cells of regenerative
species such as the chick and zebrafish, leading to a cascade of events that allow
supporting cells to transdifferentiate into hair cells (Baek et al., 2022; Cafaro et al.,
2007; Lewis et al., 2012; Ma et al., 2008a; Monroe et al., 2015). On the other hand,
mammalian postnatal cochlear supporting cells lost the ability to re-establish various
conditions necessary to convert to a hair cell fate, including the re-initiation of Atoh1
expression (Figure 2) (Groves, 2010). As a result, hair cell loss and the subsequent
hearing impairment are irreversible in mammals. Even though ATOH1 plays a critical
role during hair cell development and regeneration, efforts using it to force postnatal
Figure 2. Hair and supporting cell patterning through Notch-mediated lateral inhibition
Cartoon illustration shows the fate specification of hair cells from progenitors via Atoh1
upregulation. This in turn initiates Notch signaling in surrounding cells and inhibits Atoh1
expression in these cells, diverting them towards a supporting cell fate. In response to hair
cell damage, zebrafish supporting cells can initiate the expression of atoh1a and
transdifferentiate into hair cells, whereas mouse supporting cells no longer have the ability to
upregulate Atoh1 expressing, blocking hair cell regeneration.
rogenitors

ge, noise,
ototo ic
drugs, etc.

Atoh1

5
mammalian supporting cells to transdifferentiate remain largely unsuccessful without the
help of other hair cell-lineage transcription factors (Iyer et al., 2022; Kelly et al., 2012;
Liu et al., 2012; Liu et al., 2024; Maass et al., 2015; McGovern et al., 2024).
Nonetheless, understanding how Atoh1 is transcriptionally regulated, especially how its
expression is initiated in prosensory progenitors and non-mammalian supporting cells,
may reveal common mechanisms utilized by these two cell types to establish hair cell
competence.
1.4 Zebrafish lateral line neuromasts
As a highly regenerative species, zebrafish has been utilized to study the regeneration
of many cell types and tissues, including hair cells. In fish species, hair and supporting
cells are also found in the neuromasts of the lateral line system, where they serve to
detect the motion of water. Due to its superficial location that aids in vivo drug treatment
and imaging, the zebrafish lateral line system has been widely used to study the
mechanisms of hair cell development, patterning, and regeneration (Baek et al., 2022;
Harris et al., 2003; López-Schier and Hudspeth, 2006; Ma et al., 2008a). During
development, the lateral line neuromasts are deposited along the body of a fish by a
migrating primordium as prepatterned rosettes. Within each of these rosettes, atoh1a
expression is already restricted to the hair cell precursors located at the center, which
divide and give rise to a pair of hair cells with opposite polarities (Itoh and Chitnis, 2001;
López-Schier and Hudspeth, 2006; Nechiporuk and Raible, 2008). This is unlike the
seemingly random upregulation of Atoh1 and formation of hair cells from bipotent
sensory progenitors in the inner ear (Cai et al., 2013; Chen et al., 2002; Driver et al.,

6
2013). Moreover, hair cell regeneration in the lateral line also follows cell division of
atoh1a-expressing hair cell progenitors, which are replenished by the proliferation and
asymmetric division of a subpopulation of neuromast supporting cells (Mackenzie and
Raible, 2012; Romero-Carvajal et al., 2015). This is again different from inner ear hair
cell regeneration, where hair cells are replaced by direct transdifferention of supporting
cells without necessarily going through cell division (Cafaro et al., 2007; Roberson et al.,
2004; Shi et al., In press). Although inner ear and lateral line sensory development and
regeneration share several key features, it is not completely understood whether deeper
molecular differences in the two organs may underlie these different modes of hair cell
formation, and the extent to which findings using the lateral line neuromasts will
translate to mammalian inner ear biology. We therefore in this thesis project sought to
compare the molecular composition of the zebrafish inner ear versus lateral line
neuromasts, and found that the zebrafish inner ear more closely resembles its
mammalian counterpart.
1.5 The role of epigenetics in hair cell regeneration
Even though the postnatal mouse cochlea can no longer regenerate damaged hair
cells, it still serves as a valuable model to understand how the ability for supporting cells
to transdifferentiate into hair cells is lost during supporting cell maturation. During tissue
regeneration, lineage-related cells are often found to switch their identity and activate
the cell fate of which they are replacing (Lee et al., 2023; Thorel et al., 2010). It has
been proposed that the transdifferentiation events during regeneration are a result of
retained epigenetic memory during the development of close-related cell types, and the

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silencing of lineage-related enhancers during cell maturation plays a critical role in the
loss of regenerative capacity in adult mammals, as they can no longer drive the
expression of genes required for cell fate specification (Nicetto and Zaret, 2019; Ong
and Corces, 2012; VandenBosch et al., 2020).
Consistent with this idea, the Segil lab has found that the progressive epigenetic
silencing of hair cell gene-related promoters and enhancers during supporting cell
maturation can explain the loss of transdifferentiation potential of supporting cells. This
is achieved through the removal of permissive histone modifications as well as de novo
hypermethylation of hair cell enhancers. More specifically, in neonatal supporting cells,
hair cell enhancers are initially maintained at a bivalent or poised state and associated
with both the permissive H3K4me1 and repressive H3K27me3 histone modifications. At
this stage, supporting cells still retain the ability to activate these hair cell enhancers in
response to hair cell injury, likely due to the role of H3K4me1 in poising enhancers for
activation (Creyghton et al., 2010). However, by postnatal day 6, these hair cell
enhancers lose their H3K4me1 mark, likely through the activity of the histone
demethylase LSD1, resulting in the decommissioning of hair cell enhancers. In the
meantime, de novo DNA methylation at CpG dinucleotides in hair cell enhancers also
occurs in maturing supporting cells, further silencing hair cell enhancers and restricting
supporting cells in a terminally differentiated cell state. Lastly, both the loss of H3K4me1
and DNA hypermethylation at hair cell enhancers also coincide with the loss of
chromatin accessibility of hair cell enhancers in supporting cells. This further limits hair
cell enhancers from being utilized in postnatal mouse supporting cells to upregulate hair

8
cell-related genes and transdifferentiate into hair cells (Nguyen et al., 2023; Tao et al.,
2021).
As zebrafish inner ear supporting cells can upregulate hair cell genes in response to
injury, we sought to profile the chromatin landscape in zebrafish supporting cells near
hair cell genes, and identified a unique class of supporting cell-specific hair cell
enhancers that correspond to an inner ear-specific potential to upregulate hair cell
genes. These enhancers are maintained in zebrafish but not mouse supporting cells
through adulthood, revealing the epigenetic basis of hair cell regeneration in the
zebrafish inner ear.

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Chapter 2. Single-cell transcriptomic profiling of the
zebrafish inner ear reveals molecularly distinct hair cell and
supporting cell subtypes
(This chapter contains content taken from Shi et al. 2023, in collaboration with Marielle
Beaulieau, Lauren Saunders, Cole Trapnell, and David Raible from University of
Washington, and Peter Fabian, now at Masaryk University)
2.1 Background
Zebrafish and mammals share several inner ear sensory organs. Three semicircular
canals with sensory end organs called cristae sense angular rotation of the head. Two
additional sensory end organs detect linear acceleration and gravity: the utricular and
saccular macula each with an associated otolith crystal (Figure 3). Fish lack a specific
auditory structure such as the mammalian cochlea and instead sense sound through
the saccule, utricle, and a third otolith organ, the lagena. Although historically the utricle
was thought to be for vestibular function and the saccule and lagena analogous to the
cochlea for sound detection, there is now substantial evidence for all three otolith end
organs being used for sound detection with diverse specializations across fishes
(Popper and Fay, 1993). Zebrafish exhibit behavioral responses to sound frequencies
between 100 and 1200 Hz (Bhandiwad et al., 2013; Zeddies and Fay, 2005), and neural
responses up to 4000 Hz (Poulsen et al., 2021). In larval zebrafish, both saccule and
utricle hair cells respond to vibration stimuli, with the utricle responding to relatively

10
Figure 3. Anatomy of zebrafish and mouse inner ears
lower frequencies than the saccule, as well as additive effects when both are stimulated
(Favre-Bulle et al., 2020; Yao et al., 2016).
Within the mammalian utricle and saccule, there are both morphological and spatial
differences between hair cells (Eatock and Songer, 2011; Lysakowski and Goldberg,
2004). Hair cells are broadly classified by their morphology and innervation, with Type I
hair cells having calyx synapses surrounding the hair cell body and Type II hair cells
having bouton synapses. Both Type I and Type II cells can be found within the central
region of the macular organs known as the striola and in the surrounding extrastriolar
zones. Although the role of spatial segregation into striolar versus extrastriolar zones
Figure 3. Anatomy of zebrafish and mouse inner ears. (A) Illustration of the lateral line
system of a 5 dpf zebrafish. Blue circles represent individual neuromasts located on the
body of the fish. Boxed region indicates location of the ear. (B) Enlarged diagram of the 5
dpf zebrafish ear showing cristae (red) and macular (blue) sensory organs. (C,D) Illustrations
of adult zebrafish and mouse inner ears showing homologous end organs in the semicircular
canal crista ampullaris (red) and macula otolith organs (blue). Light green and dark green
represent unique end organs of the lagena in zebrafish and cochlea in mice. (E) Illustration
of the mouse utricle showing striolar and extrastriolar regions of the sensory organ. Arrows
represent hair cell planar polarity within the sensory organ and red dashed line represents
the line of polarity reversal within the striola. ac: anterior crista, c: cochlea, l: lagena, lc:
lateral crista, o: otolith, pc: posterior crista, s: saccule, u: utricle.

11
has not been fully elucidated, hair cells across these regions vary in morphology,
electrophysiology, and synaptic structure (Desai et al., 2005). The striola is
characterized by hair cells with taller ciliary bundles and encompasses a line of polarity
reversal where hair cells change their stereocilia orientation (Figure 3E). Whereas
distinct Type I and Type II hair cells, and in particular the calyx synapses typical of Type
I cells, have not been identified in the maculae of fishes, afferent innervation with some
calyx-like properties has been reported in goldfish cristae (Lanford and Popper, 1996).
Spatial heterogeneity in the maculae, including those of zebrafish, has also been
previously noted (Chang et al., 1992; Liu et al., 2022; Platt, 1993). However, the
homologies of cells at the cellular and molecular levels have remained unknown.
Recent single-cell and single-nucleus RNA-sequencing efforts have generated a wealth
of transcriptomic data from hair cells in several model systems, facilitating more direct
comparison of cell types and gene regulatory networks between species. Although
single-cell transcriptomic data have recently been published for the zebrafish inner ear
(Jimenez et al., 2022; Qian et al., 2022), the diversity of hair cell and supporting cell
subtypes has not been thoroughly analyzed. In order to better understand the
diversification of cell types in the zebrafish inner ear, and their relationships to those in
mammals, here we perform single-cell and single-nucleus RNA sequencing of the
zebrafish inner ear from embryonic through adult stages. We find that hair and
supporting cells from the zebrafish inner ear and lateral line are transcriptionally distinct,
and that hair and supporting cells differ between the cristae and maculae. All of these
distinct cell types are present during larval development and are maintained into

12
adulthood. In situ hybridization reveals that these hair cell subtypes occupy distinct
spatial domains within the utricle, saccule, and lagena, and computational comparison
of hair cell types reveals homology with striolar and extrastriolar hair cell types in
mammals. These findings point to an origin of striolar and extrastriolar hair cell types in
at least the last common ancestor of fish and mammals.
2.2 Zebrafish inner ear hair cells and supporting cells are transcriptionally distinct
from those of the lateral line
To assess differences between inner ear and lateral line cells, we analyzed a subset of
cells from a large single-nucleus RNA-seq dataset of whole zebrafish at embryonic and
larval stages (36–96 hours post-fertilization (hpf)), which was prepared by singlenucleus combinatorial inde ing and sequencing (‘sci-Seq’; Saunders et al., 2022).
Within an initial dataset of 1.25 million cells from 1233 embryos spanning 18 timepoints
between 18 and 96 hr (see Saunders et al., 2022 for more detail), a total of 16,517 inner
ear and lateral line cells were isolated, combined, and re-processed using Monocle 3
(Figure 4A–B). Initially, otic vesicle and lateral line cell clusters were identified by eya1
expression (Sahly et al., 1999) in combination with the following known marker genes.
Inner ear nonsensory cells were identified by expression of the transcription factor gene
sox10 (Dutton et al., 2009) in combination with inner ear supporting cell genes (stm,
otog, otogl, otomp, tecta, and oc90; Figure 4C) (Kalka et al., 2019; Petko et al., 2008;
Söllner et al., 2003; Stooke-Vaughan et al., 2015). Lateral line nonsensory cells were
identified by expression of known markers fat1b, tfap2a, tnfsf10l3, lef1, cxcr4b, fgfr1a,
and hmx3a (Figure 4D) (Feng and Xu, 2010; Haas and Gilmour, 2006; Lee et al., 2016;

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Figure 4. Molecularly distinct cell types between the zebrafish inner ear and lateral line
McGraw et al., 2011; Steiner et al., 2014; Thomas and Raible, 2019). We identified hair
cells by expression of the pan-hair cell genes otofb, cdh23, pcdh15a, ush1c, myo7aa,
slc17a8, and cacna1da (Figure 4E) (Chatterjee et al., 2015; Ernest et al., 2000;
Obholzer et al., 2008; Phillips et al., 2011; Seiler et al., 2005; Sheets et al., 2012;
Söllner et al., 2004). To distinguish between inner ear and lateral line hair cells, we
Figure 4. Molecularly distinct cell types between the zebrafish inner ear and lateral
line. Ear and lateral line cells were selected from a whole-embryo single-nucleus RNA-seq
dataset from animals between 18 and 96 hpf using known marker genes for hair cells and
supporting cells. (A–B) UMAP projection of inner ear and lateral line cells grouped by (A)
developmental timepoint and (B) broad cell type: ear nonsensory SC (red), lateral line
nonsensory SC (green), ear HC (blue), and lateral line HC (yellow). Clusters in (B)
correspond to columns of following gene expression plots. Widely accepted marker genes
for (C) inner ear nonsensory cells, (D) lateral line nonsensory cells, and (E) hair cells show
enriched expression in the corresponding clusters from B, confirming their identity. (F)
Expression of previously identified marker genes for inner ear or lateral line hair cells was
used to identify hair cell origin.

14
Figure 5. Gene modules for embryonic to larval inner ear and lateral line dataset queried expression
of previously
described markers
for inner ear (gpx2,
kifl, strc, and lhfpl5a)
and lateral line
(strc1, lhfpl5b, and
s100t) (Erickson and
Nicolson, 2015;
Erickson et al.,
2019). Although
many of these
markers are at low
abundance, these
populations are
marked distinctly by
strc and s100t
(Figure 4F). We
used Monocle3 to
identify differentially
expressed genes
and to generate
Figure 5. Gene modules for embryonic to larval inner ear and
lateral line dataset. Gene modules calculated in Monocle 3 for the
embryonic to larval inner ear and lateral line dataset displayed as (A)
a heatmap of module gene enrichment by cluster where red
indicates higher enrichment and blue indicates de-enrichment and
(B) module expression across the UMAP for the dataset.

15
Figure 6. Selection of otic sensory cells from snRNA-seq dataset
modules of co-expressed genes (Figure 5).
Both hair cells and nonsensory supporting cells from the inner ear and lateral line
formed distinct clusters, with nonsensory cells from the two mechanosensory organs
showing greater distinction than hair cells (Figure 4B and 2.4A). To confirm the relative
differences between inner ear and lateral line hair cells and nonsensory cells, PartitionFigure 6. Selection of otic sensory cells from snRNA-seq dataset. (A) Clustering
of 18 hpf to 96 hpf dataset to illustrate cell subtypes. PAGA analysis of this dataset
shows strong connectivity among ear nonsensory cells and among lateral line
nonsensory cells, but weak interconnectivity between these two groups. (B) Feature
plots show expression of the supporting cell marker lfng, and markers of structural
otic cells matn4 and col2a1a. (C) UMAP of sensory patch cells from 36 to 96hpf are
clustered without structural and early otic vesicle cells. PAGA analysis again shows
strong connectivity within hair cells and supporting cell groups and weak connectivity
between lateral line and inner ear supporting cells.

16
based Graph Abstraction (PAGA) analysis was used to measure the connectivity of
clusters (Wolf et al., 2019). PAGA analysis revealed strong connectivity within inner ear
supporting cell clusters and within lateral line supporting cell clusters but little
connectivity between them (Figure 6A).
The inner ear nonsensory cluster includes structural cells forming the otic capsule,
identified by expression of the extracellular matrix protein-encoding genes collagen type
2 a1a (col2a1a) and matrilin 4 (matn4) (Xu et al., 2018), as well as sensory supporting
cells expressing lfng (Figure 3D; Figure 6B). Inner ear and lateral line supporting cells
remain as distinct clusters even when structural matn4+ cells are excluded from
analysis (Figure 6C). Thus, both hair cells and supporting cells have distinct gene
expression profiles between the inner ear and lateral line at embryonic and larval
stages.
2.3 Single-cell RNA-seq reveals distinct hair cell and supporting cell populations
in the juvenile and adult inner ear of zebrafish
To identify distinct subtypes of inner ear hair cells and supporting cells from larval
through adult stages, we first re-analyzed single-cell RNA sequencing (scRNA-seq)
datasets from larval stages (72 and 120 hpf) (Fabian et al., 2022), in which otic placode
cells and their descendants were labeled with Sox10:Cre to induce recombination of an
ubiquitous ubb:LOXP-EGFP-STOP-LOXP-mCherry transgene (Kague et al., 2012). We
also performed additional scRNA-seq using these transgenic lines by dissecting ears
from juvenile (14 days post-fertilization (dpf)), and adult (12 months post-fertilization

17
Figure 7. scRNA-seq of 12 mpf zebrafish inner ear captures sensory hair cells and supporting cells as well as non-sensory
supporting cells
(mpf)) animals. Following cell dissociation and fluorescence-activated cell sorting
(FACS) to purify mCherry + cells, we constructed scRNA-seq libraries using 10x
Chromium technology. For all datasets, hair cells and supporting cells were identified for
further analysis based on the expression of hair cell markers myo6b and strc and
supporting cell markers stm and lfng; structural cells were removed from further analysis
based on expression of matn4 and col2a1a (Figure 7). Using Seurat, we integrated this
Figure 7. scRNA-seq of 12 mpf zebrafish inner ear captures sensory hair cells and
supporting cells as well as non-sensory supporting cells. (A) Clustering of 12 mpf
dataset to illustrate cell types in the adult zebrafish inner ear. (B–I) Feature plots of 12 mpf
zebrafish scRNA-seq dataset alone showing expression of hair cell markers (B) myo6b and
(C) strc, pan-supporting cell marker (D) stm, sensory supporting cell markers (E) lfng and
(F) hey1, and pan-otic marker (G) otomp, and non-sensory supporting cell markers (H)
matn4 and (I) col2a1a.

18
Figure 8. Cell subtypes in the zebrafish inner ear end organs
dataset with the sci-Seq embryonic and larval dataset (36–96 hpf) (Figure 8A and B).
The combined dataset comprises 3246 inner ear cells separated into 10 groups based
Figure 8. Cell subtypes in the zebrafish inner ear end organs. (A–D) Integration and
analysis of single-cell RNAseq data generated by sci-Seq (sci) or 10x Chromium sequencing
(10x) for inner ear hair cells and supporting cells from embryonic (sci), larval (sci,10x), and
adult (10x) stages. UMAP projection of cells are grouped by (A) dataset of origin and (B)
timepoint. (C) Unsupervised clustering divides cells into 10 clusters that were grouped into 9
cell subtypes. (D) Feature plots showing hair cell marker myo6b, nascent hair cell marker
dla, supporting cell marker lfng, and putative progenitor marker fgfr2 expression in the
integrated dataset. (E) Differentially expressed genes across the 10 cell clusters.

19
Figure 9. Hair cell and supporting cell marker expression in the integrated scRNA-seq
dataset
on unsupervised clustering, with differentially expressed genes for each cluster shown
in Figure 8E. We identified six clusters of hair cells based on shared expression of
myo6b, strc, lhfpl5a, and gfi1aa (Yu et al., 2020), a nascent hair cell cluster based on
expression of atoh1a (Millimaki et al., 2007) and the Notch ligand dla (Riley et al.,
1999), and two clusters of supporting cells based on expression of lfng and stm (Figure
8C and D and Figure 9). An additional putative progenitor cluster (cluster 0), enriched
for cells from embryonic
stages, is characterized by
expression of genes such
as fgfr2 (Rohs et al., 2013),
fat1a (Down et al., 2005),
igsf3, and pard3bb (Figure
10). Although these marker
genes are differentially
expressed in the putative
progenitor cluster, some of
them (e.g. fat1a and
pard3bb) retain a lower
expression level in
supporting cell populations
(Figure 10). This is further Figure 9. Hair cell and supporting cell marker expression
in the integrated scRNA-seq dataset. Feature plots of
integrated zebrafish scRNA-seq datasets showing
expression of nascent hair cell marker (A) atoh1a, inner ear
hair cell markers (B) strc, (C) gfi1aa, and (D) lhfpl5a, and
pan-supporting cell marker (E) stm.

20
Figure 10. Putative progenitor marker expression in individual progenitor and supporting cell clusters

21
Figure 11. Gene modules for integrated inner ear sensory patch dataset
demonstrated by gene
modules of these
clusters (Figure 11),
where the progenitor
signature module
genes (Module 1) are
expressed in lower
levels in the
supporting cell
clusters. This
Figure 10. Putative progenitor marker expression in individual progenitor and
supporting cell clusters. (A) Combined and individual UMAP projections of putative
bipotent progenitor cluster (cluster 0), macular supporting cell cluster (cluster 6), and cristae
supporting cell cluster (cluster 7) from the integrated zebrafish inner ear dataset. (B–E)
Feature plots show expression of putative progenitor genes in the integrated dataset, as well
as in individual clusters of 0, 6, and 7. (F) Violin plots showing differential gene expression of
fgfr2, igsf3, fat1a, and pard3bb among clusters 0, 6, and 7. Wilco on rank sum test, *: p ≤
0.05, ***: p ≤ 1e-3, ****: p ≤ 1e-4.
Figure 11. Gene
modules for
integrated inner ear
sensory patch
dataset. Gene
modules calculated in
Monocle 3 for the
integrated inner ear
sensory patch dataset
displayed as (A) a
heatmap of module
gene enrichment by
cluster where red
indicates higher
enrichment and blue
indicates deenrichment and (B)
module expression
across the UMAP for
the dataset.

22
transcriptional relatedness between progenitors and supporting cells may underlie the
role of supporting cells as a resident stem cell population during zebrafish hair cell
regeneration.
2.4 Developmental trajectories in the zebrafish inner ear
To understand potential lineage relationships between clusters, we performed
pseudotime trajectory analysis using Monocle3. We anchored the pseudotime projection
at the putative progenitor cell cluster. Analysis revealed two major trajectories toward
hair cell and supporting cell clusters for both maculae and cristae (Figure 12A and B
and Figure 13), with distinct patterns of gene expression along each trajectory. We find
that average gene expression of the putative progenitor (Cluster 0) markers follow two
patterns: decreasing along both hair cell and supporting cell trajectories (fgfr2 and igsf3)
and decreasing only along the hair cell trajectory (fat1a and pard3bb) (Figure 12C and
D, Figure 13B and C). The hair cell trajectory progresses first through a stage marked
by expression of dla and then atoh1a (Cluster 2, Figure 12E, Figure 13D). Concurrent
with decreasing expression of nascent hair cell genes, we observe increasing
expression of mature hair cell genes gfi1aa and myo6b (Figure 12F, Figure 13E). Along
the supporting cell trajectory we observed upregulation of supporting cell-specific
markers, including stm and lfng (Figure 12G, Figure 13F). These bifurcating lineage
trajectories from Cluster 0 (Figure 12A) to hair and supporting cell clusters are
consistent with the identification of Cluster 0 as a population of bipotent progenitors
regulated by Notch signaling during early development (Haddon et al., 1998; Riley et al.,

23
Figure 12. Pseudotime analysis reveals developmental trajectories in the zebrafish inner ear
1999). To localize these developmental stages in vivo, we examined dla expression by
Figure 12. Pseudotime analysis reveals developmental trajectories in the zebrafish
inner ear. (A,B) Pseudotime analysis of macular cells showing simulated developmental
trajectories of a putative bipotent progenitor population into hair cell and supporting cell
clusters. (C,D) Changes in putative progenitor markers along (C) hair cell and (D) supporting
cell trajectories. fat1a and pard3bb only decrease along the hair cell trajectory, while fgfr2
and igsf3 decrease along both hair cell and supporting cell trajectories. (E) Transient
expression of early hair cell genes dla and atoh1a along hair cell trajectories. (F) Increases in
gene expression levels of gfi1aa and myo6b along hair cell trajectories. (G) Increases in stm
and lfng along supporting cell trajectories.

24
Figure 13. Pseudotime analysis of cristae hair and supporting cells in the zebrafish inner ear
in
situ
Figure 13. Pseudotime analysis of cristae hair and supporting cells in the zebrafish
inner ear. (A) Pseudotime analysis showing simulated developmental trajectories of a
putative bipotent progenitor population into both cristae hair and supporting cell clusters.
(B,C) Changes in putative progenitor markers along (B) hair cell and (C) supporting cell
trajectories. fat1a and pard3bb only decrease along the hair cell trajectory, while fgfr2 and
igsf3 decrease along both hair cell and supporting cell trajectories. (D) Transient
expression of early hair cell genes dla and atoh1a along hair cell trajectories. (E) Increases
in gene expression levels of gfi1aa and myo6b along hair cell trajectories. (F) Increases in
stm and lfng along supporting cell trajectories.

25
Figure 14. dla labels putative hair cell progenitors in the cristae and
maculae
hybridization (Figure 14). We find that dla is expressed in supporting cells adjacent to
myo6:GFP hair cells in both cristae and maculae, consistent with peripheral addition of
new cells at the margins of the sensory patches.
2.5 Distinct supporting cell types in the cristae versus maculae
Supporting cells comprise two major clusters that can be distinguished by expression of
tectb and zpld1a among other genes (Figure 8C). The tectb gene encodes Tectorin
beta, a component of the tectorial membrane associated with cochlear hair cells in
mammals (Goodyear et al., 2017), and a component of otoliths in zebrafish (Kalka et al.,
2019). The zpld1a gene, encoding Zona-pellucida-like domain containing protein 1 a, is
expressed in the cristae in fish (Dernedde et al., 2014; Yang et al., 2011) and mouse
(Vijayakumar et al., 2019). Using fluorescent in situ hybridization, we find that tectb is
Figure 14. dla labels putative hair
cell progenitors in the cristae
and maculae. HCR in situ
hybridization of 5 dpf zebrafish.
Maximum intensity projections of
(A) posterior crista (lateral view),
(B) utricle (dorsal view), and (C)
saccule (lateral view) showing dla
expression in a subset of support
cells (arrowheads) peripheral to
myo6b+hair cells. Scale bars = 10
μm.

26
Figure 15. Distinct markers separate macula and crista supporting cells
expressed in the macular organs but not cristae, and zpld1a is expressed in cristae but
not maculae (Figure 15C and D). Neither were detected in lateral line neuromasts
(Figure 15C and D), showing they are inner ear-specific genes. Both tectb and zpld1a
are expressed primarily in supporting cells, as they show little overlap in expression with
the hair cell marker myo6b:GFP, similar to expression of the supporting cell marker lfng
(Figure 15B–D and Figure 16). These results demonstrate the presence of distinct
supporting cell subtypes for the maculae and cristae.
Figure 15. Distinct markers separate macula and crista supporting cells. (A) Feature
plots showing expression of macula supporting cell marker tectb and crista supporting cell
marker zpld1a. (B–D) HCR in situ hybridization in myo6b:GFP transgenic animals. Each
set of images shown represents a projection of one z-stack split into cristae (lateral) and
macula (medial) slices. Lateral line neuromasts positioned over the ear are visible in lateral
slices. Expression pattern for (B) the pan-supporting cell marker lfng, (C) macula-specific
marker tectb, and (D) crista-specific marker zpld1a in 5 dpf myo6b:GFP fish. Each set of
images shown represents a projection of one z-stack split into cristae (lateral) and macula
(medial) slices. ac: anterior crista, lc: lateral crista, nm: neuromast, pc: posterior crista, u:
utricle, s: saccule. Scale bars = 20 μm.

27
Figure 16. zpld1a and tectb are primarily expressed in supporting cells
Figure 16. zpld1a and tectb are primarily expressed in supporting cells. HCR in situ
hybridization of 5 dpf myo6b:GFP zebrafish. (A–B) Confocal slices through (A) anterior
crista and (B) lateral crista (lateral view) show localization of cabp5b in hair cells and
zpld1a in supporting cells. (C) Slice through utricle (dorsal view) shows cabp2b expression
in hair cells and tectb expression primarily in the surrounding supporting cells. (D) Slices
through saccule (lateral view) at the level of hair cell bodies (top row) and supporting cell
bodies (bottom row). cabp2b is primarily expressed in hair cells and tectb is primarily
e pressed in supporting cells. Scale bars = 10 μm.

28
Figure 17. Gene expression differences between lateral line and inner ear hair cells
2.6 Distinct types of hair cells in the zebrafish inner ear
While inner ear and lateral line hair cells share many structural and functional features,
we sought to determine if these cells also have distinct molecular signatures. We
compared published datasets of lateral line hair cells (Baek et al., 2022; Kozak et al.,
2020; Ohta et al., 2020) to our data, restricting analysis to datasets generated by 10x
Chromium preparation to avoid technical batch effects across studies. Using
Scanorama for alignments (Hie et al., 2019), hair cells from the inner ear and lateral line
form distinct clusters, with a number of differentially expressed genes (Figure 17),
including the known markers for lateral line (s100t) and inner ear (strc) (Figure 4). This
analysis suggests that inner ear hair cells of the maculae and cristae are more similar to
each other than to lateral line hair cells.
Figure 17. Gene expression differences between lateral line and inner ear hair cells.
(A) UMAP of our 12 mpf hair cell dataset integrated by Scanorama with published lateral
line hair cell datasets. Lateral line hair cells cluster separately from inner ear hair cells. (B)
Differential gene expression analysis identifies novel marker genes specific to either lateral
line or inner ear hair cells.

29
Within the maculae and cristae, we find that hair cells can be subdivided into two major
groups (clusters 1 and 3 versus cluster 4). These clusters are distinguished by
differential expression of a number of genes including two calcium binding protein
genes, cabp1b and cabp2b (Di Donato et al., 2013) (Figure 8E). Hair cell cluster 5 has a
mixed identity with co-expression of a number of genes shared between these two
groups, including cabp1b and cabp2b.
We next tested the in vivo expression of genes in each cluster using in situ
hybridization, choosing cabp1b and cabp2b as representative markers for each cluster
(Figure 18A). In the larval cristae, utricle, and saccule, cabp1b and cabp2b mark
myo6b+hair cells in largely non-overlapping zones (Figure 18B–D). By adult stages,
complementary domains of cabp1b+and cabp2b+hair cells become clearly apparent
(Figure 18E–K). In the adult utricle, a central crescent of cabp2b+; myo6b+hair cells is
surrounded by a broad domain of cabp1b+; myo6b+hair cells. In the saccule and
lagena, a late developing sensory organ, central cabp2b+; myo6b+hair cells are
surrounded by peripheral cabp1b+; myo6b+hair cells. We also find several genes that
are specific for hair cells in the cristae, utricle, or saccule (Figure 19A). These include
the calcium binding protein gene cabp5b in the cristae, the transcription factor skor2 in
the utricle, and the deafness gene loxhd1b in the saccule (Figure 19B–D, Figure 20).

30
Figure 18. cabp1b+and cabp2b+label hair cells in distinct regions of sensory end organs
The domain organization of hair cells in the adult macular organs resembles that of
striolar and extrastriolar hair cells in the mammalian utricle. We therefore examined
expression of pvalb9, the zebrafish ortholog of the mouse striolar hair cell marker Ocm
(Hoffman et al., 2018; Jiang et al., 2017) (Figure 21, Figure 22). In the larval utricle, we
observe near complete overlap of pvalb9 with cabp2b (Figure 21B–D). In the adult
utricle, there is substantial overlap of pvalb9 with cabp2b expression (except for a thin
Figure 18. cabp1b+and cabp2b+label hair cells in distinct regions of sensory end
organs. (A) Feature plots showing differential expression of cabp1b and cabp2b among
crista and macula hair cells. (B–D) HCR in situ projections of individual sensory patches
from 5 dpf myo6b:GFP fish showing differential spatial expression patterns of cabp1b and
cabp2b. (B) cabp1b is expressed at the ends of the cristae, while cabp2b is expressed
centrally. Anterior crista is shown. (C) In the utricle, cabp1b is expressed medially and
cabp2b is expressed laterally. (D) In the saccule, cabp1b is expressed in peripheral cells at
the dorsal and ventral edges of the organ. cabp2b is expressed centrally. Scale bars for
HCR images = 10 μm. (E) Cartoon illustrations of the zebrafish utricle, saccule, and
lagena, and the expression patterns of cabp1b (yellow) and cabp2b (magenta) within each
sensory patch. (F–H) Wholemount RNAScope confocal images of adult inner ear organs
showing peripheral expression pattern of cabp1b (n=3) in the adult zebrafish (F) utricle, (G)
saccule, and (H) lagena. (I–K) Whole-mount RNAScope confocal images showing central
expression pattern of cabp2b (n=4) in the adult zebrafish (I) utricle, (J) saccule, and (K)
lagena. Scale bars for RN Scope images = 25 μm.

31
Figure 19. Distinct markers separate macula and crista hair cells
Figure 20. skor2 and loxhd1b label subsets of hair cells in
utricle or saccule
strip of pvalb9+; cabp2b- cells), and little
overlap with cabp1b expression (Figure
Figure 19. Distinct markers separate macula and crista hair cells. (A) Feature plots
showing marker genes enriched in organ-specific subsets of inner ear hair cells: cabp5b,
skor2, and loxhd1b. (B–D) HCR in situs in 5 dpf myo6b:GFP fish show expression of (B)
cabp5b in crista but not macula hair cells, (C) skor2 in the utricle only, and (D) loxhd1b in
the saccule, as well as lateral line neuromast hair cells. Each set of images represents an
orthogonal projection of one z-stack split into cristae (lateral) and macular (medial) slices.
ac: anterior crista, lc: lateral crista, nm: neuromast, pc: posterior crista, s: saccule, u:
utricle. Scale bar = 20 μm.
Figure 20. skor2 and loxhd1b label
subsets of hair cells in utricle or
saccule. HCR in situ hybridization of 5
dpf zebrafish. (A) Maximum intensity
projection of utricle (dorsal view) showing
skor2 expression in medially located
myo6b:GFP+ hair cells. (B) Maximum
intensity projection of saccule (lateral
view) showing loxhd1b expression in a
peripheral subset of hair cells. Scale bars
= 10 μm.

32
Figure 21. Zebrafish cabp2b+domain shares features with the mouse striolar region
21F and G). In addition, anti-Spectrin staining of hair bundles reveals a line of polarity
reversal within the cabp2b+domain of the utricle (Figure 21H, I), consistent with polarity
reversal occurring within the striolar domains of mammalian macular organs (Li et al.,
2008). Cluster 1/3 (cabp1b+) and Cluster 4 (cabp2b+) populations also differentially
express genes related to stereocilia tip link and mechanotransduction channel
components (Figure 23) and various calcium and potassium channels (Figure 24). We
also note that the utricle marker skor2 labels primarily extrastriolar hair cells within this
end organ, with loxhd1b labeling striolar hair cells within the saccule. These findings
suggest that zebrafish Cluster 4 (cabp2b+) and Cluster 1/3 (cabp1b+) hair cells largely
correspond to striolar and extrastriolar hair cells, respectively, with distinct
mechanotransduction and synaptic properties.
Figure 21. Zebrafish cabp2b+domain shares features with the mouse striolar region.
(A) Feature plot shows enrichment for the striola marker pvalb9 in cabp2b-expressing
striolar cells. (B–D) HCR in situs in 5 dpf myo6b:GFP fish shows pvalb9 and cabp2b coe pression in the utricle. Scale bar = 10 μm. (E) Cartoon illustration of overlapping
expression of pvalb9 (white) and cabp2b (magenta) that coincides with the line of hair cell
polarity reversal. (F, G) Whole-mount RNAScope confocal images of adult zebrafish
utricles showing expression of pvalb9 relative to (F) cabp1b (n=3) and (G) cabp2b (n=4).
Scale bar = 25 μm. (H,I) Whole-mount RNAScope RNA and protein co-detection assay
showing co-localization of cabp2b expression (RNA) and the hair cell line of polarity
reversal indicated by Spectrin (protein) staining (n=3). Scale bar = 25 μm. rrows denote
hair cell polarity and dotted line outlines line of polarity reversal.

33
Figure 22. Striola marker pvalb9 is expressed in all inner
ear sensory end organs
Figure 23. Inner ear hair cell subtypes differentially express mechanosensory apparatus genes
Figure 22. Striola marker pvalb9 is
expressed in all inner ear sensory
end organs. HCR in situ hybridization
of 5 dpf zebrafish. (A) Maximum
intensity projection of saccule (lateral
view) shows pvalb9 expression in
centrally located myo6b:GFP+ hair
cells. (B) Slice through the anterior
crista shows pvalb9 expression in a
subset of crista hair cells. Scale bars =
10 μm.
Figure 23. Inner ear hair cell subtypes differentially express mechanosensory
apparatus genes. Feature plots for mechanosensory transduction genes from the
integrated zebrafish scRNA-seq dataset

34
Figure 24. Inner ear hair cell subtypes differentially express voltage-gated calcium and potassium channel genes
2.7 Global homology of striolar and extrastriolar hair cells between fish and mice
To further probe similarities between zebrafish Cluster 4 (cabp2b+) and Cluster 1/3
(cabp1b+) hair cells versus striolar and extrastriolar hair cells in mammals, we utilized
the Self-Assembling Manifold mapping (SAMap) algorithm (Musser et al., 2021;
Tarashansky et al., 2021) to compare cell types across distant species. A strength of
this algorithm is that it compares not only homologous gene pairs but also close
paralogs, which is especially useful considering the extensive paralog switching
observed between vertebrate clades (Postlethwait, 2007), as well as the extra round of
Figure 24. Inner ear hair cell subtypes differentially express voltage-gated calcium
and potassium channel genes. Feature plots for ion channel genes from the integrated
zebrafish scRNA-seq dataset

35
genome duplication in the teleost lineage leading to zebrafish. When comparing adult
zebrafish maculae with the postnatal mouse utricle (Jan et al., 2021), we find the
highest alignment score between supporting cells (Figure 25A). Consistent with the
spatial domains revealed by our in situ gene expression analysis, we find that mouse
striolar Type I hair cells exclusively map to zebrafish Cluster 4 (cabp2b+) hair cells, and
mouse extrastriolar Type I and Type II hair cells predominantly to zebrafish Cluster 1/3
(cabp1b+) hair cells. In contrast, zebrafish lateral line hair cells (Lush et al., 2019) align
exclusively to mouse extrastriolar and not striolar hair cells (Figure 26). The small
degree of mapping of mouse extrastriolar Type I hair cells to zebrafish Cluster 4
(cabp2b+) hair cells suggests that zebrafish Cluster 4 (cabp2b+) hair cells may have
more of a Type I identity than Cluster 1/3 (cabp1b+) cells in general. Gene pairs driving
the homology alignment include striolar markers Ocm, Loxhd1, and Atp2b2 for zebrafish
Cluster 4 (cabp2b+) hair cells, and mouse extrastriolar markers Tmc1, Atoh1, and Jag2
for zebrafish Cluster 1/3 (cabp1b+) hair cells. Thus, zebrafish Cluster 4 (cabp2b+)
macular hair cells are closely related to striolar cells of the mouse utricle, with zebrafish
lateral line and Cluster 1/3 (cabp1b+) macular hair cells more closely related to mouse
extrastriolar hair cells.
A recent single-cell study revealed distinct central versus peripheral hair cell
subpopulations in postnatal mouse cristae, reminiscent of the striolar and extrastriolar
populations in the maculae (Wilkerson et al., 2021). As our zebrafish cristae hair cells
also separate into distinct clusters, Cluster 9 (cabp1b+) and Cluster 8 (cabp2b+) (Figure
18A and B), we performed SAMap analysis between the crista cell populations of the

36
Figure 25. SAMap analysis reveals conserved gene expression patterns between mouse and zebrafish hair cell types
two species to investigate cell type homology. Similar to what we observed for the
utricle, zebrafish centrally located Cluster 8 crista hair cells predominantly map to
mouse central crista hair cells, and zebrafish peripherally located Cluster 9 crista hair
cells exclusively map to mouse peripheral crista hair cells (Figure 25B). Conserved
types of spatially segregated hair cells therefore exist in both the maculae and cristae of
zebrafish and mouse.
Figure 25. SAMap analysis reveals conserved gene expression patterns between
mouse and zebrafish hair cell types. (A–B) Sankey plot showing the SAMap mapping
scores (0–1) that indicate transcriptome relatedness between (A) mouse utricular and
zebrafish macular single-cell clusters and (B) mouse and zebrafish cristae single-cell
clusters. A mapping score of 0 indicates no evolutionary correlation in transcriptome while
a mapping score of 1 indicates perfect correlation. Correlations below 0.15 were not
plotted.

37
Figure 26. SAMap analysis of mouse utricle versus zebrafish macular and lateral line cells
2.8 Discussion
Our single-cell transcriptomic profiling of the embryonic to adult zebrafish inner ear
reveals a diversity of hair cell and supporting cell subtypes that differ from those of the
lateral line. As much of our knowledge about zebrafish hair cell regeneration comes
from studies of the lateral line, understanding similarities and differences between the
lateral line and inner ear has the potential to uncover mechanisms underlying the
distinct regenerative capacity of inner ear hair cell subtypes. Recent tools to
Figure 26. SAMap analysis of mouse utricle versus zebrafish macular and lateral line
cells. (A–B) Sankey plot showing the SAMap mapping scores (0–1) that indicate
transcriptome relatedness between mouse utricular and integrated zebrafish macular and
lateral line single-cell clusters. (A) Zebrafish 12 mpf macular HCs integrated with 5 dpf
lateral line HCs. (B) Zebrafish 3–5 dpf macular HCs integrated with 5 dpf lateral line HCs. A
mapping score of 0 indicates no evolutionary correlation in transcriptome, and a mapping
score of 1 indicates perfect correlation. Correlations below 0.2 were not plotted.

38
systematically damage inner ear hair cells in zebrafish (Jimenez et al., 2021) should
enable such types of comparative studies.
We identify hair cells and supporting cells specific for maculae versus cristae, as well as
two spatially segregated types of zebrafish inner ear hair cells with similarities to
mammalian striolar and extrastriolar hair cells. These molecular signatures are
conserved across larval and adult stages. However, consistent with other recent work
(Jimenez et al., 2022; Qian et al., 2022), we were not able to resolve distinct clusters of
hair cells or supporting cells corresponding to the distinct types of maculae: i.e. utricle,
saccule, and lagena.
The division of auditory and vestibular function across the otolith organs in zebrafish
remains somewhat unclear. The saccule is thought to act as the primary auditory organ
of larval zebrafish, as the utricle is not necessary for sound detection above low
frequencies (Yao et al., 2016). In the zebrafish adult, excess sound exposure can
damage the saccule, while damage to the utricle is unknown (Schuck and Smith, 2009).
Conversely, the utricle is critical for larval vestibular function, while input from the
saccule is unnecessary (Riley and Moorman, 2000). However, there is contrasting
evidence for overlap in function of both saccule and utricle for sound detection in larvae
(Favre-Bulle et al., 2020; Poulsen et al., 2021). Currently we are not able to identify
clearly distinct hair cell types in the utricle compared to the saccule that might reflect
functional differences; whether such genetic signatures exist remains an important
question that will require further in-depth analysis. It is interesting to note that

39
mammalian vestibular end organs are also capable of responding to high-frequency
sound stimuli (reviewed in Curthoys, 2017), suggesting that sound detection by hair
cells may not be linked to a distinct end organ-specific molecular signature.
Our study supports zebrafish possessing distinct types of striolar and extrastriolar hair
cells in the maculae and cristae, with molecular differences between these subtypes
implying different physiological properties. In the zebrafish utricle, vibration is
preferentially transduced by striolar cells while static tilt is received by extrastriolar cells
(Tanimoto et al., 2022). Consistent with use of a s100s-hs:tdTomato transgene to mark
striolar cells in this previous study, s100s is a highly specific marker for our striolar hair
cell cluster (Figure 8E). We also find zebrafish striolar and extrastriolar hair cell
subtypes express distinct combinations of ion channel genes and mechanotransduction
components, consistent with previous reports of distinct current profiles in central versus
peripheral hair cells in the zebrafish utricle, saccule, and lagena (Haden et al., 2013; Olt
et al., 2014), as well as spatial differences in ciliary bundle morphology and synaptic
innervation in the larval zebrafish utricle (Liu et al., 2022). The distinct spatial
distribution, channel expression, and hair bundle morphologies in these hair cells
resembles the known spatial, electrophysiological, and hair bundle compositional
differences seen in the striolar versus extrastriolar hair cells in the amniote vestibular
end organs (Holt et al., 2007; Kharkovets et al., 2000; Lapeyre et al., 1992; Meredith
and Rennie, 2016; Moravec and Peterson, 2004; Rüsch et al., 1998; Xue and Peterson,
2006).

40
In each of the zebrafish end organs, striolar and extrastriolar hair cells can be defined
by differential expression of calcium binding proteins, in particular cabp1b versus
cabp2b. As these calcium binding proteins closely interact with synaptic calcium
channels (Cui et al., 2007; Picher et al., 2017) with potential functionally different
consequences (Yang et al., 2018), their differential expression may confer unique
electrophysiological properties to each cell type. Mutations in human CABP2 associated
with the autosomal recessive locus DFNB93 result in hearing loss (Picher et al., 2017;
Schrauwen et al., 2012), underlining its functional importance. Even though we
chose cabp1b and cabp2b as characteristic markers for zebrafish extrastriolar and
striolar regions, it is worth noting that Cabp2, but not Cabp1, is expressed in all mouse
postnatal utricular hair cells with differentially higher expression in the striola (Jan et al.,
2021). Of note, lateral line hair cells express higher levels of cabp2b than cabp1b (Lush
et al., 2019), despite our analysis suggesting that they are more closely related to
extrastriolar hair cells. These observations emphasize the importance of examining
global patterns of gene expression rather than individual markers when assigning
homology of cell types.
By contrast, we found no clear homology of zebrafish inner ear hair cells with
mammalian Type I and Type II hair cells. The lack of molecular signatures
corresponding to Type I hair cells is consistent with previous reports that one of their
major features, calyx synapses, are absent from macular organs in fishes (Lysakowski
and Goldberg, 2004, but see Lanford and Popper, 1996 for evidence of calyx synapses
in goldfish cristae). These findings suggest that the diversification of inner ear hair cells

41
into Type I and Type II cells may have largely emerged after the evolutionary split of
ray-finned fishes from the lineage leading to mammals.
We recognize that identifying cell type homology across tissues and species through
molecular analysis has several potential caveats. Although we have collected
transcriptomic data from the zebrafish inner ear from a wide range of developmental
stages, we are limited by the fact that the publicly available datasets for zebrafish lateral
line and mouse utricle and cristae are restricted to immature stages. Thus, cell maturity
could be a confounder in our analyses. However, when we limited the comparison of
lateral line hair cells and postnatal mouse vestibular hair cells to 3–5 dpf inner ear hair
cells, we see similar alignments as when we used our 12 mpf data (Figure 2.24). In
addition, we collected fewer supporting cells from adult zebrafish than expected,
skewing cell type representation towards hair cells (Figure 8C). Thus, additional
optimization may be needed to further interrogate the cell subtypes within zebrafish
inner ear supporting cell populations.
Nonetheless, our integrated dataset reveals distinct molecular characteristics of hair
cells and supporting cells in the zebrafish inner ear sensory organs, with conservation of
these patterns from larval stages to adults. Although not discussed in detail here, our
data include additional cell populations of the zebrafish inner ear that express
extracellular matrix-associated genes important for otic capsule structure and ion
channel-associated genes associated with fluid regulation. These data form a resource
that can be further explored to inform molecular aspects of hair cell electrophysiology,

42
mechanotransduction, sound versus motion detection, maintenance of inner ear
structure and ionic balance, and inner ear-specific hair cell regeneration.

43
Chapter 3. Long-range Atoh1 enhancers maintain
competency for hair cell regeneration in the inner ear
(This chapter contains content taken from Shi et al. 2024, in collaboration with Leah
Kim, Juan Llamas, Xizi Wang, and Thomas Lozito from University of Southern
California, and Peter Fabian, now at Masaryk University)
3.1 Background
Since Atoh1 plays a crucial role in the development of hair cells, and it is one of the first
genes to be upregulated in supporting cells during hair cell regeneration, I decided to
focus my work on understanding how zebrafish is able to initiate the expression of
atoh1a during hair cell development and regeneration. Previous studies of Atoh1
regulation have largely focused on enhancer elements accessible and active in hair
cells where Atoh1 is expressed (Helms et al., 2000; Luo et al., 2022). In particular,
these studies identified autoregulatory enhancers that are bound by ATOH1 to amplify
its expression in developing hair cells. Deletion of these Atoh1 autoregulatory
enhancers in mice consequently resulted in loss of hair cells in the cochlea due to
severe but not complete reduction of Atoh1 expression (Luo et al., 2022). These
findings point to the presence of another group of yet to be identified enhancers
responsible for Atoh1 initiation.
Here we performed comparative single-cell profiling of gene expression and accessible
chromatin in the inner ears of zebrafish and the green anole lizard and compared this to
published datasets of the mouse cochlea. In so doing, we identified a syntenic group of
long-range enhancers distal to the Atoh1 autoregulatory enhancers that become

44
accessible and active in progenitor cells before the Atoh1 autoregulatory enhancers.
These long-range enhancers then rapidly closed in developing hair cells, in contrast to
the Atoh1 autoregulatory enhancers that maintained accessibility. In addition, these
long-range Atoh1 enhancers maintained accessibility in adult supporting cells of the
regenerative zebrafish and lizard inner ears but not in the non-regenerative postnatal
mouse cochlea. Deletion of these enhancers from the zebrafish genome resulted in a
severe reduction in atoh1a expression and hair cell number during development and
regeneration of the inner ear but not the lateral line. Moreover, we identified putative
enhancers associated with a number of other prosensory and hair cell genes in
zebrafish and mouse with similar properties to the long-range Atoh1 enhancers. Global
motif enrichment analysis of these enhancers revealed SOX motifs as commonly
enriched in both species, and mutation of predicted SOX binding sites in a sequenceconserved Atoh1 enhancer destroyed its activity in zebrafish supporting cells. Our study
thus reveals a distinct class of enhancer that maintains the competency of adult
zebrafish supporting cells to initiate expression of atoh1a and transdifferentiate into hair
cells following injury.
3.2 Single-cell profiling of the zebrafish inner ear identifies distinct classes of
atoh1a enhancers
To examine chromatin accessibility in the juvenile and adult zebrafish inner ear, we
permanently labeled otic placode lineage cells with Sox10:Cre (Kague et al., 2012);
ubb:LOXP-EGFP-LOXP-mCherry (Mosimann et al., 2011), dissected and dissociated
inner ears into single-cell suspensions, and performed fluorescence-activated cell
sorting (FACS) to enrich for otic cells. We then isolated nuclei and used the 10X

45
Figure 27. Inner ear enhancer architecture of the zebrafish atoh1a locus
Genomics platform and Illumina next-generation sequencing to make a single-nuclei
ATACseq dataset from 14 days post-fertilization (dpf) juvenile fish and a combined
single-nuclei RNAseq and ATACseq dataset from 12 months post-fertilization (mpf)
adult zebrafish (Figure 27A). In both datasets, we identified hair cell and supporting cell
clusters based on imputed (14 dpf) or actual (12 mpf) expression of previously
described hair cell markers (strc and lhfpl5a), and crista (zpld1a) and macular (tectb)
supporting cell markers (Shi et al., 2023) (Figure 28A-C). We then subclustered hair
cells and supporting cells and identified distinct subtypes based on markers identified in
a previous study (Shi et al., 2023) (Figure 27B). cabp5b- macular hair cells could be
Figure 27. Inner ear enhancer architecture of the zebrafish atoh1a locus. (A) Workflow
used to construct single-nuclei ATAC-seq (14 dpf) and multiome (12 mpf) libraries of
zebrafish inner ear cells labeled by Sox10:Cre-mediated genetic recombination. (B)
UMAPs of 14 dpf and 12 mpf libraries following reclustering of HCs and SCs. (C) Feature
plot showing atoh1a expression in HCs but not SCs at 12 mpf. (D,E) Coverage plots
showing chromatin accessibility downstream of zebrafish atoh1a in 14 dpf (D) and 12 mpf
(E) snATAC-seq datasets. Class 1 enhancers (eh1-4, yellow) are differentially accessible in
HCs. Class 2 enhancers (eh5, eh7-9, purple) are differentially accessible in SCs. eh6 is
accessible in both HCs and SCs and shaded in green.

46
Figure 28. Single-cell analysis of the zebrafish inner ear
divided into striolar, extrastriolar, and mixed hair cells, based on cabp1b and cabp2b
expression, and cabp5b+ cristae hair cells could be divided into cabp1b+ peripheral and
cabp2b+ central hair cells (Figure 28D,E). We also uncovered dla+ young hair cells in
the 12 mpf dataset (Figure 28E).

47
At the atoh1a locus, we identified several putative enhancers downstream of the coding
region, which we divided into two classes (Figure 27D,E). Class 1 enhancers (eh1-4)
have selective accessibility in hair cell compared to supporting cell clusters and contain
predicted ATOH1 binding motifs. As eh1 is sequence-conserved with a well-known
autoregulatory enhancer of mouse Atoh1 (Helms et al., 2000; Luo et al., 2022), Class 1
enhancers are likely also autoregulatory in zebrafish. Class 2 enhancers (eh5, eh7-9)
are located further downstream of Class 1 enhancers, have selective accessibility in
supporting cells compared to hair cells, and lack predicted ATOH1 binding motifs.
Enhancer eh6 is accessible in both hair cells and supporting cells. We observed the
distribution of Class 1 versus Class 2 peaks is largely consistent within the single-cell
clusters of hair cell and supporting cell subtypes, with some variation of Class 1
accessibility in hair cell clusters that may reflect their maturity. For the rest of the study,
we therefore focus on pseudo-bulk chromatin landscapes of the atoh1a locus in
aggregated hair cell and supporting cell clusters (Figure 29A).
3.3 Differential activity of atoh1a enhancers in hair cells and supporting cells
Figure 28. Single-cell analysis of the zebrafish inner ear. (A) UMAPs of single-nuclei
14 dpf ATAC-seq and 12 mpf multiomic ATAC-seq and RNA-seq libraries of dissected
inner ears show HC and SC clusters among all recovered cells. (B) Feature plots show
imputed gene activities of the HC markers strc and lhfpl5a and SC markers zpld1a and
tectb used to identify clusters in the 14 dpf dataset. (C) Feature plots show expression of
the HC markers strc and lhfpl5a and SC markers zpld1a and tectb used to identify clusters
in the 12 mpf dataset. (D) Feature plots show imputed gene activities of the HC subtype
markers cabp1b, cabp2b, and cabp5b and SC subtype markers zpld1a and tectb used to
identify HC and SC subtype clusters in the 14 dpf dataset. (E) Feature plots show
expression of the HC subtype markers cabp1b, cabp2b, and cabp5b, young HC marker
dla, and SC subtype markers zpld1a and tectb used to identify HC and SC subtype
clusters in the 12 mpf dataset.

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As atoh1a expression is largely absent from adult supporting cells (Figure 27C), the
continued accessibility of eh6-9 in adult supporting cells (Figure 27E) could indicate
their involvement in either actively repressing atoh1a expression or allowing supporting
cells to re-express atoh1a after hair cell damage. To assess their in vivo activity, we
used Tol2 transgenesis (Kwan et al., 2007) to test the ability of putative Class 1 and 2
enhancers to drive mCherry or GFP expression, respectively, when combined with a
minimal E1b promoter. Examination of several independent founder lines for each
revealed that all nine atoh1a enhancers were able to drive reporter activity in the inner
ear and/or lateral line (Figure 29B,C and Figure 30A). Consistent with their differential
accessibility, Class 1 enhancers (eh1-4) drove hair cell-specific mCherry reporter
expression (Figure 29B and Figure 30B), Class 2 enhancers (eh5 and eh7-9) drove
supporting cell-specific GFP expression (Figure 29C and Figure 30D), and eh6, which is
accessible in both hair cells and supporting cells, drove GFP expression in both (Figure
29C and Figure 30C). We confirmed hair cell expression of eh1 by co-localization with
myo6b:GFP (Figure 30B), and mutually exclusive expression of hair cell-specific
eh2:mCherry and supporting cell-specific eh7:GFP (Figure 30C). We also confirmed
dual hair cell and supporting cell activity of eh6 by partial co-localization with hair cellspecific eh1:mCherry (Figure 30D). Of the Class 2 enhancers, eh5 drove GFP
expression only in neuromast supporting cells, and eh7 and eh9 drove expression only
in inner ear supporting cells, with eh8 showing stronger expression in inner ear versus
neuromast supporting cells. While Class 1 enhancers also drove strong expression in
the brain (eh1) and intestine (eh1, eh3, eh4) (Figure 30A), tissues known to express

49
Figure 29. Cell type-specific reporter activities of zebrafish atoh1a enhancers
atoh1a (Farah et al., 2000; Yang et al., 2001), we did not detect Class 2 enhancer
activities in those tissues.

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Figure 30. Additional analysis of atoh1a enhancer transgenic lines
Figure 29. Cell type-specific reporter activities of zebrafish atoh1a enhancers. (A)
Genome browser tracks show the pseudo-bulk chromatin accessibility downstream of
zebrafish atoh1a in aggregated HCs or SCs at 14 dpf. Highlighted are HC-specific Class 1
enhancers (1-4, yellow), SC-specific Class 2 enhancers (5, 7-9, purple), and commonly
accessible eh6 enhancer (green). (B,C) Individual enhancers were combined with the
minimal E1b promoter and used to drive expression of mCherry (eh1-4) or GFP (eh5-9) in
stable transgenic zebrafish. Single-z images show activity in the neuromasts of the lateral
line and the cristae and utricles of the inner ear at 3 dpf, with DIC channel for eh1 image
providing context. We observed consistent expression patterns in at least 5 embryos each
from 2 or more independent transgenic founders for each line, except for 1 founder each
for eh2 and eh8. Scale bars = 50 μm.

51
3.4 Notch-dependent repression of Class 2 enhancer activity in supporting cells
through the atoh1a promoter
The ability of Class 2 enhancers to drive reporter expression in supporting cells was
unexpected given the lack of atoh1a expression in these cells. As Atoh1 expression is
normally suppressed in mouse supporting cells through binding of Notch-dependent
HES and HEY transcription factors to the sequence-conserved Atoh1 promoter
(Abdolazimi et al., 2016), we asked whether transgenic activity of Class 2 enhancers in
supporting cells could be due to use of the minimal E1b promoter that lacks such Notch
inhibition (Figure 31A). Consistently, in multiple independent transgenic zebrafish lines
in which the Class 2 enhancer eh7 was paired with the endogenous atoh1a promoter to
drive GFP expression (eh7_Patoh1a:GFP), we observed lack of GFP in supporting cells
(Figure 31B). However, blocking Notch signaling by treatment with 10 µM of the γsecretase inhibitor DBZ resulted in re-expression of GFP in supporting cells of the crista
and utricle within 48 hours (Figure 31B). As a control, we verified that the atoh1a
promoter alone did not drive any reporter expression with or without DBZ (Figure 31C).
We also found that expression of Class 1 enhancer eh2 in hair cells was unaffected by
pairing with the atoh1a promoter with or without DBZ treatment (Figure 31D), although
we did not test whether ectopic activation of Notch signaling could silence it. These
Figure 30. Additional analysis of atoh1a enhancer transgenic lines. (A) Low
magnification views of the 5 dpf zebrafish head and anterior trunk show transgenic
expression driven by atoh1a enhancers (mCherry for eh1-4, GFP for eh5-9) paired with the
E1b minimal promoter. Arrow indicates brain activity and arrowheads indicate intestine
activity. We observed consistent expression patterns in at least 5 embryos each from 2 or
more independent transgenic founders for each line, except for 1 founder each for eh2 and
eh8. (B-D) Single-z images of 3 dpf zebrafish neuromasts, cristae, and utricles show colocalization of eh1:mCherry with HC-specific myo6b:GFP (B), mutually exclusive activity of
HC-specific eh2:mCherry and SC-specific eh7:GFP (C), and partial co-localization of HCspecific eh1:mCherry with dual HC/SC-expressed eh6:GFP (D). We observed consistent
expression patterns in at least 5 embryos each from 2 or more independent crosses of
zebrafish carrying each transgene, Scale bars are 50 μm.

52
Figure 31. Notch inhibition of eh7 enhancer activity through the endogenous atoh1a promoter
results reveal that, despite their accessibility, the activity of eh7, and potentially other
Class 2 enhancers, is normally repressed in supporting cells by the action of Notch
signaling on the atoh1a promoter.
Figure 31. Notch inhibition of eh7 enhancer activity through the endogenous atoh1a
promoter. (A) Diagrams of the eh7 reporter constructs show SC activity when paired with
the minimal E1b promoter and silencing when paired with the endogenous atoh1a
promoter. Previous work has shown that Notch silences the endogenous atoh1a promoter
by inducing expression of Hes/Hey factors that directly bind to the promoter. Inhibition of
Notch signaling with DBZ reactivates the eh7-endogenous promoter construct. (B-D)
Confocal sections of the crista and utricle show reactivation of eh7_Patoh1a:GFP (blue)
following 10 uM DBZ treatment (B), lack of activity of the endogenous atoh1a promoter
when not paired with an enhancer (C), and HC activity of the eh2 enhancer when paired
with the endogenous atoh1a promoter (mCherry) with or without DBZ treatment (D). We
observed consistent expression patterns in at least 5 embryos each from 2 or more
independent transgenic founders for eh7_Patoh1a:GFP and no_eh_Patoh1a:GFP, and 1
founder for eh2_Patoh1a:mCh. Fluorescence images are shown with and without DIC
channels for conte t. Scale bars = 50 μm.

53
3.5 Differential requirements of Class 1 and 2 enhancers for hair cell development
in the inner ear and lateral line
To interrogate enhancer requirements, we used CRISPR/Cas9 genome editing to delete
atoh1a enhancers individually or in combination from the zebrafish genome (Figure
32A). For combinatorial deletions, we targeted the entire regions spanning the
enhancers rather than each enhancer individually. We then performed RNAScope to
detect and quantify atoh1a expression levels and phalloidin staining to assess hair cell
numbers at 5 dpf. We observed no changes in inner ear hair cell numbers in individual
Δeh6 and Δeh9 deletions or combinatorial Δeh1-4, Δeh7-8, and Δeh7-9 deletions
(Figure 32A and Figure 33H-J). In contrast, we observed distinct hair cell defects in
homozygous animals where all hair cell-accessible Class 1 (Δeh1-6) or all supporting
cell-accessible Class 2 (Δeh5-9) enhancers were deleted (Figure 32).
Homozygous Δeh1-6 zebrafish lack all Class 1 enhancers (eh1-4), the lateral linespecific supporting cell enhancer eh5, and enhancer eh6 with dual activity in hair cells
and supporting cells. In Δeh1-6 fish, we observed a mild reduction of atoh1a expression
and hair cell numbers in the inner ear cristae and utricle, and a more severe loss of
atoh1a expression and hair cell number in lateral line neuromasts (Figure 32B-E). The
preferential effect of eh1-6 deletion on lateral line versus inner ear hair cells could
reflect removal of the lateral line-specific eh5 enhancer and/or greater dependency of
lateral line hair cell formation on the atoh1a autoregulatory function of Class 1
enhancers (Luo et al., 2022).

54
Figure 32. Requirements of Class 1 and 2 atoh1a enhancers in HC development

55
Reciprocally, in homozygous Δeh5-9 zebrafish, which lack all Class 2 enhancers
accessible in supporting cells, we observed a near complete loss of atoh1a expression
and a severe reduction of hair cells in the inner ear, but only mild effects on lateral line
neuromast hair cells (Figure 32B-E). Further supporting a unique role of eh5-9 in inner
ear hair cell development, Δeh5-9 zebrafish exhibited a somersaulting swimming
pattern associated with vestibular dysfunction (Baeza-Loya and Raible, 2023), with
failure to inflate swim bladders and death around 10 dpf. In contrast, Δeh1-6 zebrafish
displayed no vestibular defects and lived to adulthood, though we did not test for other
potential behavioral changes associated with the loss of neuromast hair cells. Neither
homozygous Δeh1-6 nor Δeh5-9 animals had a complete loss of atoh1a expression or
hair cells as seen in atoh1a null mutants (Hewitt et al., 2024) (Figure 33C,F,G),
suggesting potential redundancy between Class 1 and Class 2 enhancers and/or roles
of other unidentified enhancers. These results point to preferential requirements of
Class 2 supporting cell-specific enhancers for inner ear hair cell development,
Figure 32. Requirements of Class 1 and 2 atoh1a enhancers in HC development. (A)
Scheme of atoh1a enhancer deletions relative to pseudo-bulk chromatin accessibility at 14
dpf. Black lines indicate enhancer deletions that generated no phenotypes. (B) Triple
RNAScope in situ hybridization for atoh1a and markers of HCs (myo6b) and SCs (lfng)
with nuclei labeled by DAPI in white. Individual channels to the right highlight preferential
loss of atoh1a e pression in the neuromasts of homozygous Δeh1-6 animals and in the
cristae and utricles of homozygous Δeh5-9 animals at 5 dpf. (C) Confocal projections of
inner ears stained with halloidin show selective reductions of neuromast HCs in Δeh1-
6/Δeh1-6 fish and utricle and crista HCs in Δeh5-9/Δeh5-9 fish at 5 dpf. Note that for the
utricle the smaller extrastriolar HCs are more affected. (D) Violin plots show quantification
of atoh1a puncta per atoh1a-expressing cell in different sensory organs of the indicated
genotypes. A minimum of 10 cells from 3 or more animals per group were scored. Means
and standard errors of the mean are shown. Statistical tests used Tukey’s HSD multiple
pairwise test. (E) Violin plot show quantification of HC number in different sensory organs
of the indicated genotypes. A minimum of 3 animals were scored per group. Means and
standard errors of the mean are shown. Statistical tests used Tukey’s HSD multiple
pairwise test. Scale bars are 50 μm.

56
Figure 33. Additional analysis of atoh1a enhancer deletion alleles
consistent with the selective activity of Class 2 enhancers eh7-9 in the inner ear (Fig.
3.3C).

57
3.6 Selective requirement of Class 2 enhancers for inner ear hair cell regeneration
As some hair cells still form in Δeh5-9 animals, we were able to test whether Class 2
enhancers are also required for hair cell regeneration. Homozygous Δeh5-9 animals did
not survive past 10 dpf, limiting the timeframe we could test for regeneration defects to
late larval stages. To ablate hair cells, we crossed the Δeh5-9 allele onto the
myo6b:hDTR line (Jimenez et al., 2021) and injected 1-2 nL of 1 ng/µL Diphtheria Toxin
(DT) into otic vesicles at 4 dpf. By 1 day post-injection (dpi), heterozygous Δeh5-9 fish
exhibited somersaulting behavior, and we confirmed by phalloidin staining that hair cells
in the neuromasts and utricle were ablated (Figure 34A). hair cell ablation in the cristae
was inconsistent using this method and not reported here. We then allowed hair cells to
regenerate over the next 3 days (1-4 dpi/5-8 dpf) and sacrificed a portion of the injected
zebrafish every day to detect atoh1a expression and visualize hair cells.
Figure 33. Additional analysis of atoh1a enhancer deletion alleles. (A-C) Triple
RNAScope in situ hybridization for atoh1a and markers of HCs (myo6b) and SCs (lfng)
with nuclei labeled by DAPI in white. To the right are the atoh1a channel alone and
Phalloidin staining of HCs. At 5 dpf, atoh1a expression and HCs are completely lost in
homozygous atoh1amRuby null animals(4) (C) compared to heterozygous Δeh1-6 (A) and
Δeh5-9 (B) controls. (D-G) Violin plots show quantification of atoh1a puncta per atoh1aexpressing cell (D,F) and HC numbers (E,G) for the indicated sensory organs and
genotypes shown. For each group, a minimum of 8 cells were counted from 3 animals per
group. Means and standard errors of the mean are shown. Statistical tests used Tukey’s
HSD multiple pairwise test. (H) Top panels shown Phalloidin staining of HCs in indicated
sensory organs and bottom panel is triple RNAScope in situ hybridization of utricle for
atoh1a and markers of HCs (myo6b) and SCs (lfng) with nuclei labeled by DAPI in white.
No defects were seen in homozygous Δeh1-4 and Δeh7-9 animals. (I,J) Violin plots show
quantification of atoh1a puncta per atoh1a-expressing cell (I) and HC numbers (J) for the
indicated sensory organs and genotypes shown. A minimum of 3 animals were counted for
each group. Means and standard errors of the mean are shown. Statistical tests used
Tukey’s HSD multiple pairwise test. We did not observe any statistical differences between
genotypes. Scale bars are 50 μm.

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Figure 34. Requirements of Class 1 and 2 atoh1a enhancers in HC regeneration

59
In heterozygous Δeh5-9 zebrafish, we found near complete regeneration of lateral line
hair cells and partial regeneration of hair cells in the utricle at 4 dpi (Figure 34A,H,I),
reflecting previous reports of slower regeneration in the inner ear compared to the
lateral line (Beaulieu et al., 2024; Ma et al., 2008b; Schuck and Smith, 2009). In both
the lateral line and inner ear, atoh1a was strongly upregulated at 3 dpi (Figure 34C). In
the regenerating inner ear, we observed broad upregulation of atoh1a in the lfng+
supporting cell layer (Figure 35A-B), indicative of supporting cells initiating a hair cell
fate. To more directly test for supporting cell transdifferentiation in the zebrafish inner
ear, we identified a supporting cell-specific accessible region near the supporting cellenriched gene kcnq5a and used it to drive expression of a photoconvertible nuclear
EOS fluorescent protein in stable transgenic line kcnq5a_p1:nlsEOS (Figure 35C,D).
We crossed this transgenic line to supporting cell- and hair cell-specific reporter lines to
Figure 34. Requirements of Class 1 and 2 atoh1a enhancers in HC regeneration.
(A,B) Phalloidin staining shows HCs of the neuromast and utricle at daily intervals
following injection of DT into the otic vesicles of myo6b:DTR; +/Δeh5-9 (A) and
myo6b:DTR; Δeh5-9/Δeh5-9 (B) zebrafish at 4 dpf. Utricle HCs selectively fail to
regenerate in Δeh5-9/Δeh5-9 fish. For the utricle, the HCs are below the strongly stained
oval-shaped otic vesicle. (C,D) Images correspond to animals in (A,B) 3 days after DT
injection. Triple RNAScope in situ hybridization for atoh1a and markers of HCs (myo6b)
and SCs (lfng) with nuclei labeled by DAPI in white. Channels to the right highlight
selective failure to recover atoh1a e pression in the utricles of Δeh5-9/Δeh5-9 animals,
and merged atoh1a/lfng channels highlight double-positive atoh1a/lfng cells in the
regenerating utricle of +/Δeh5-9 controls. (E) Single-z images show reactivation of
eh7_Patoh1a:GFP (blue) in utricle 2 days after DT injection compared to PBS-injected
controls. Dashed circles indicate otoliths for context. (F) Phalloidin staining shows HCs of
the neuromast and utricle at 1, 3, and 4 days following injection of DT into the otic vesicles
at 4 dpf. Compared to myo6b:DTR; +/Δeh1-6 controls, myo6b:DTR; Δeh1-6/Δeh1-6
zebrafish show a partial reduction in HC regeneration. (G) Triple RNAScope in situ
hybridization for atoh1a and markers of HCs (myo6b) and SCs (lfng) with nuclei labeled by
DAPI in white. Channels to the right show largely normal atoh1a expression and
atoh1a/lfng colocalization in the regenerating utricles of both genotypes at 3 days post-DT
injection. (H-J) Quantification of HC numbers at daily intervals following DT injection for the
indicated genotypes. For each group, a minimum of 3 animals were scored. Means and
standard errors of the mean are shown. Scale bars are 25 μm ( ,B) and 50 μm (C-G).

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confirm the pattern of EOS expression. In 4 dpf offspring, we observed co-localization of
kcnq5a_p1:nlsEOS with the supporting cell-specific eh7:GFP but not the hair cellspecific myo6b:GFP in the cristae, confirming supporting cell-specific EOS expression
(Figure 35E,F). We then crossed kcnq5a_p1:nlsEOS and myo6b:DTR; myo6b:GFP
animals to directly assess whether supporting cells in the zebrafish ear directly convert
to hair cells as was described for other species (Roberson et al., 2004; White et al.,
2006). We used photoconversion to label kcnq5a_p1:nlsEOS+ cristae supporting cells
red at 4 dpf and assessed their contribution to new hair cells three days after hair cell
ablation. Compared to control injection of kcnq5a_p1:nlsEOS; myo6b:DTR; myo6b:GFP
fish with PBS, injection with DT at 4 dpf to partially ablate cristae hair cells resulted in
kcnq5a_p1:nlsEOS-labeled myo6b:GFP+ hair cells by 7 dpf (Figure 35G,H). These
findings indicate that at least a subset of inner ear hair cells arise directly from
supporting cells labeled by class 2 atoh1a enhancers during regeneration.
Next, we assessed the requirement of class 2 atoh1a enhancers in inner ear hair cell
regeneration. In contrast to fish with heterozygous loss of eh5-9 or homozygous loss of
eh6, eh7-8, or eh9 enhancers, hair cell regeneration and atoh1a expression were
completely blocked at 4 dpi in the utricles of homozygous Δeh5-9 zebrafish lacking all
class 2 enhancers (Figure 34B,D,I and Figure 35I,K). Highlighting inner ear-specific
requirement of Class 2 enhancers, neuromast hair cells regenerated to a similar extent
in homozygous Δeh5-9 zebrafish versus heterozygous controls (Figure 34A,B,H),
although we did observe a moderate reduction in atoh1a expression in homozygous
Δeh5-9 fish (Figure 34D and Figure 35J). We also found that hair cell ablation

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reactivated expression of eh7_Patoh1a:GFP, which was silenced during homeostasis,
in utricle supporting cells within 2 dpi, coincident with upregulation of atoh1a expression
(Figure 34E). Together, these findings show that Class 2 enhancers, required for normal
hair cell development, are redeployed during regeneration to promote the
transdifferentiation of supporting cells into hair cells specifically in the inner ear.
Using the same myo6b:hDTR approach, we also examined the extent to which deletion
of Class 1 enhancers (eh1-6) impacted hair cell regeneration in the inner ear at larval
stages. s Δeh1-6 fish lack neuromast hair cells at 4 dpf, we were unable to assess the
effects on lateral line hair cell regeneration at these same stages. Unlike the failed
regeneration seen in Δeh5-9 zebrafish, we observed only a modest delay of hair cell
regeneration in the utricles of homozygous Δeh1-6 fish compared to heterozygous
siblings (Figure 34F,J), accompanied by partial reduction of atoh1a upregulation (Figure
34G and Figure 35I). These animals recovered hair cell number to functional levels and
resumed normal swimming patterns 7-10 days after DT injection, further highlighting a
preferential requirement for Class 2 over Class 1 enhancers for hair cell regeneration in
the inner ear.

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Figure 35. atoh1a and lfng co-localization in the uninjured and regenerating utricle and evidence for SC transdifferentiation

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3.7 Cross-species activity of Class 2 enhancers
To determine whether Atoh1 Class 2 enhancer activity correlates with regenerative
potential across species, we performed single-cell multiome of the adult green anole
Figure 35. atoh1a and lfng co-localization in the uninjured and regenerating utricle
and evidence for SC transdifferentiation. (A) Triple RNAScope in situ hybridization for
atoh1a and markers of HCs (myo6b) and SCs (lfng) with nuclei labeled by DAPI in white.
To the right are merged myo6b/atoh1a/lfng channels and individual myo6b, lfng, and
atoh1a channels to highlight increased atoh1a expression in SCs during regeneration (3
days post-DT injection at 4 dpf) versus control (PBS-injected). Dashed lines separate
myo6b-high top HC layer from the lfng-high bottom SC layer). (B) Violin plot shows
quantification of the percentage of lfng+/myo6b- SCs per section that co-express atoh1a 3
days following HC ablation by DT injection versus PBS-injected control. For each group, a
minimum of 9 sections from 3 animals were scored. Means and standard errors of the
mean are shown. Statistical tests used t test. (C) Dot plot shows expression of kcnq5a in
12 mpf inner ear single-cell clusters. (D) Genome browser tracks show the pseudo-bulk
chromatin accessibility of the kcnq5a locus in aggregated HCs or SCs at 12 mpf. p1
(shaded in green) indicates the region used to make the kncq5a_p1-E1b:nlsEOS line. (EF) Single-z images of 4 dpf zebrafish cristae show co-localization of photoconverted
kcnq5a_p1-E1b:nlsEOS with SC-specific eh7:GFP (E), and mutually exclusive activity of
HC-specific myo6b:GFP and photoconverted kncq5a_p1-E1b:nlsEOS prior to HC ablation
(F). To the right are the GFP and EOS channels alone. We observed consistent
expression patterns in at least 5 embryos each from 2 or more independent crosses of
zebrafish carrying each transgene. (G) Single-z images of 7 dpf zebrafish cristae show
short-term lineage trace of photoconverted kcnq5a_p1-E1b:nlsEOS in relation to
myo6b:GFP marked HCs, 3 days after DT injection versus PBS-injected control.
Arrowheads indicate cells with double positive nuclear EOS and cytoplasmic GFP
expression. To the right are the GFP and EOS channels alone. (H) Violin plot shows
quantification of cells co-expressing photoconverted nuclear EOS+ and cytoplasmic GFP+
3 days following partial cristae HC ablation by DT injection versus PBS-injected control,
compared to immediately before injections. For each group, a minimum of 3 animals were
scored. Means and standard errors of the mean are shown. Statistical tests used Tukey’s
HSD multiple pairwise test. (I) Violin plot shows quantification of atoh1a puncta per atoh1aexpressing cell in utricle sections in the indicated genotypes 3 days after DT injection. For
each group, a minimum of 8 cells from at least 3 animals were scored. Means and
standard errors of the mean are shown. Statistical tests used Tukey’s HSD multiple
pairwise test. (J) Quantification of atoh1a puncta per atoh1a-expressing cell in neuromast
sections in the indicated genotypes 3 days after DT injection. For each group, a minimum
of 10 cells were scored from at least 3 animals. Means and standard errors of the mean
are shown. Statistical tests used t test. (K) Quantification of HC numbers at daily intervals
following DT injection for the indicated genotypes. For each group, a minimum of 3 animals
were scored. Means and standard errors of the mean are shown. (L) Genome browser
tracks show pseudo-bulk chromatin accessibility downstream of atoh1a in 12 mpf adult
zebrafish inner ear HCs and SCs, compared to pseudo-bulk chromatin accessibility in a
mixed cell population from the uninjured adult (6-10 mpf) zebrafish utricle and in the
regenerating utricle 4, 5, and 7 days post-ablation in the myo6b:DTR model(23). Scale
bars are 50 μm.

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Figure 36. Lizard saccule multiome
lizard inner ear saccule and re-analyzed published mouse cochlear bulk ATAC-seq
datasets (Tao et al., 2021). Marker expression identified distinct hair cell and supporting
cell clusters in the anole saccule (Figure 36A,B). In anole and mouse, we observed
three regions (EH1-3/Eh1-3) corresponding to Class 1 enhancers selectively accessible
in hair cells. Anole EH1 is also sequence-conserved with zebrafish eh1 and mouse Eh1.
In both the adult anole and neonatal (P1) mouse cochlea, we also observed long-range
Class 2-like Atoh1 enhancers with accessibility in supporting cells but not hair cells that
were syntenic to those of the zebrafish atoh1a locus (Figure 37A-C), with anole EH4
sequence-conserved with zebrafish eh7 and mouse Eh4. Similar to zebrafish, anole
Class 2 enhancers were accessible in adult supporting cells. In contrast, mouse Class 2
enhancers were no longer accessible in cochlear supporting cells at P6 and P21,
corresponding to loss of supporting cell transdifferentiation potential at these stages
(Cox et al., 2014; Kelly et al., 2012; Liu et al., 2012; Maass et al., 2015). Further
supporting our hypothesis that Class 2 are utilized to initiate Atoh1 expression, we
Figure 36. Lizard saccule multiome. (A) UMAP of the combined snATAC- and snRNAseq (multiome) library from the adult green anole saccule shows HC and SC clusters
among all recovered cells. (B) Feature plots show marker genes used to identify the HC
(MYO7A, MYO6) and SC (LFNG, HEY2) clusters.

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found that they were accessible and co-enriched for the active enhancer histone mark
H3K27ac in embryonic day 13.5 (E13.5) mouse bipotent prosensory progenitors
(Gnedeva et al., 2020; Wang et al., 2023) (Figure 37D). These findings reveal a
common architecture of the Atoh1 locus across vertebrates, with Class 2 enhancers
maintaining accessibility in adult supporting cells only in regenerative species.
To determine whether species-specific maintenance of Class 2 enhancer accessibility
can be explained by sequence differences, we tested the activities of the conserved
zebrafish eh7 and corresponding mouse Eh4 enhancers in the zebrafish and mouse
inner ear. In zebrafish, we found that mouse Eh4, when coupled with the minimal E1b
promoter, drove GFP expression in supporting cells of the utricle at 3, 5, and 21 dpf in a
pattern identical to that of zebrafish eh7 (Figure 37E,F). Reciprocally, we tested
zebrafish eh7 or mouse Eh4 for their ability to drive GFP expression in the mouse
cochlea when paired with the minimal b-globin promoter. We packaged reporter
constructs into an adeno-associated virus (AAV) serotype AAV-ie-K558R that has been
shown to broadly infect inner ear cell types (Tao et al., 2022) and delivered them via the
posterior semicircular canal (Isgrig and Chien, 2018) to P0 mice. Consistent with their
patterns of accessibility, both eh7 and Eh4 were active in SOX2+ supporting cells and
not MYO7A+ hair cells of the cochlea at P2 (Figure 37G,H and Figure 38), when
cochlear supporting cells still have the capacity for transdifferentiation (Cox et al., 2014;
White et al., 2006). However, by P14 we found loss of reporter activity of the mouse
Eh4 and zebrafish eh7 in cochlear supporting cells (Figure 37I,J). These findings show
that it is the distinct regulatory environment within the zebrafish inner ear versus the

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Figure 37. Comparative atoh1a enhancer architecture and activity across species
mouse cochlea, and not species-specific sequence differences in eh7/Eh4, that dictates
the maintenance of Class 2 enhancer accessibility in supporting cells.

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Figure 38. Enhancer testing controls in the mouse cochlea
Figure 37. Comparative atoh1a enhancer architecture and activity across species.
(A-C) Genome browser tracks show pseudo-bulk chromatin accessibility in inner ear HCs
and SCs of various stages at the zebrafish atoh1a (A), lizard ATOH1 (B), and mouse
Atoh1 (C) loci. ATACseq datasets are from the entire zebrafish inner ear, lizard saccule,
and mouse cochlea(39). Syntenic groups of HC-specific Class 1 enhancers are shaded
yellow, and SC-specific Class 2 enhancers are shaded purple. Zebrafish also have
neuromast SC-specific eh5 (magenta) and dual HC/SC eh6 (green). Sequence-conserved
enhancers are labeled * and #. Note maintenance of Class 2 enhancer accessibility in 12
mpf zebrafish and adult lizard SCs but not in P6 and P21 mouse SCs. (D) Genome
browser tracks show chromatin accessibility and H3K27ac CUT&RUN peaks downstream
of mouse Atoh1 locus in E13.5 cochlear prosensory progenitors(42, 43). (E,F) Single-z
images show GFP reporter activities (blue) of mouse Eh4 (E) and zebrafish eh7 (F)
enhancers paired with the E1b minimal promoter in the zebrafish utricle from larval to
juvenile stages. Dashed circles indicate otoliths for context. (G-J) Images of the mid-region
cochlea from P2 (G,H) or P14 (I,J) mice that were infected at P0 with Mm-Eh4-PHbb:GFP
(G,I) or Dr-eh7-PHbb:GFP (H,J) AAVs. Only SC layers are shown. SCs are labeled by antiSOX2 antibody, and infected SCs are labeled by anti-RFP antibody. Each enhancer was
injected to five P0 pups and repeated three times. The transduction efficiency varied
among the injected animals. The mouse inner ear sensory organs with good transduction
efficiency were used for analysis and similar reporter activity was observed in each. Scale
bars are 50 μm.

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3.8 Potential role of SOX transcription factors in Class 2 enhancer activity
In order to understand how zebrafish inner ear supporting cells retain a permissive
regulatory environment for hair cell transdifferentiation in response to injury, and how
this may differ in the mouse cochlea, we sought to identify a broader set of Class 2-like
enhancers and examine them for common transcription factor binding motifs. First, we
determined a set of early hair cell genes expressed in developing E17.5 mouse hair
cells and absent in P6 supporting cells, where Atoh1 Class 2 enhancers are closed (Tao
et al., 2021). We then identified their associated ATACseq peaks within 200 kb up or
downstream of their transcription start sites in P1 supporting cells, where Atoh1 Class 2
enhancers are still open. In zebrafish, we identified orthologous early hair cell genes
and searched for ATACseq peaks within +/-200 kb of their transcription start sites in 14
dpf supporting cells. This analysis revealed 6208 Class 2-like peaks associated with
1143 early hair cell genes in mouse, and 5834 Class-2-like peaks associated with
orthologous early hair cell genes in zebrafish (Figure 39A,B and Figure 40A,B).
Mirroring Atoh1/atoh1a Class 2 enhancer behavior, 82.5% of Class 2-like peaks (5124)
lost accessibility in supporting cells (log2FC < -0.5) from P1 to P6 in mouse, yet 79.9%
Figure 38. Enhancer testing controls in the mouse cochlea. (A) Images of the midregion cochlea from P2 mice that were infected at P0 with enhancer-less PHbb:GFP
negative control AAV. Only SC layers are shown. SCs are labeled by anti-SOX2 antibody,
and infected SCs are labeled by anti-RFP antibody. Each enhancer was injected to five P0
pups and repeated three times. The transduction efficiency varied among the injected
animals. The mouse inner ear sensory organs with good transduction efficiency were used
for analysis. The enhancer reporter activity was consistent between the sensory organs
analyzed. (B-D) Images of the mid-region cochlea from P2 mice that were infected at P0
with Mm-Eh1-PHbb:GFP (B), Mm-Eh4-PHbb:GFP (C), and Dr-eh7-PHbb:GFP (D) AAVs. Only
the HC layers labeled by anti-MYO7A antibody are shown. Note HC-specific activities of
Mm-Eh1 compared to lack of Mm-Eh4 or Dr-eh7 activities in HCs. Each enhancer was
injected to five P0 pups and repeated three times. The transduction efficiency varied
among the injected animals. The mouse inner ear sensory organs with good transduction
efficiency were used for analysis. The enhancer reporter activity was consistent between
the sensory organs analyzed. Scale bar = 50 μm.

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of Class 2-like peaks (4662) maintained accessibility (log2FC > -0.5) in supporting cells
from 14 dpf to 12 mpf in zebrafish. Examples of genes with Class 2-like peaks in mouse
and zebrafish include Six1/six1b and Eya1/eya1 (Figure 40C), prosensory genes
thought to act upstream of Atoh1/atoh1a in hair cell formation (Ahmed et al., 2012;
Bricaud and Collazo, 2006). In contrast, Class 2-like peaks were not observed for
downstream targets of ATOH1 necessary for hair cell maturation such as
Pou4f3/pou4f3 and Gfi1/gfi1ab (Jen et al., 2022; Masuda et al., 2011; Yu et al., 2021)
(Figure 40D).

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Figure 39. Global analysis of Class 2-like enhancers in zebrafish and mouse

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We next performed Homer de novo motif enrichment for both mouse and zebrafish
Class 2-like peaks. In each species, the top motif was SOX, and we also uncovered
common EBF and TEAD motifs (Figure 39C,D). Although we performed the analysis in
neonatal supporting cells, the motif enrichment profile for Class 2-like peaks resembled
that previously described for accessible chromatin regions of E13.5 bipotent prosensory
progenitors (Wang et al., 2023; Wilkerson et al., 2019). These findings suggest that
Class 2-like enhancers may be broadly utilized in adult zebrafish supporting cells to
maintain a progenitor-like chromatin landscape for transdifferentiation into hair cells
after damage, with loss of Class 2-like enhancer accessibility in the postnatal mouse
cochlea resulting in a lack of competency for supporting cell transdifferentiation.
As SOX motifs were predicted in over 60% of Class 2-like peaks, including within the
conserved regions of the eh7/Eh4 Atoh1 enhancer pair, we tested whether SOX binding
is required for zebrafish eh7 activity. We destroyed all predicted SOX binding sites in
eh7 by either deleting or making point mutations in key nucleotides of SOX recognition
sequences (Figure 40E). Either method resulted in loss of the ability of eh7 to drive GFP
Figure 39. Global analysis of Class 2-like enhancers in zebrafish and mouse. (A,B)
Heatmaps show differentially accessible chromatin within 200kb of mouse(39) (A) and
zebrafish (B) HC genes. Class 1 peaks are selectively accessible in P1 HCs and SCs
versus P6 SCs in mouse, and in 14 dpf HCs versus 14 dpf and 12 mpf SCs in zebrafish.
Class 2 peaks are selectively accessible in P1 SCs versus P1 HCs and P6 SCs in mouse,
and in 14 dpf and 12 mpf SCs versus 14 dpf HCs in zebrafish. (C,D) Homer de novo motif
predictions for mouse (C) and zebrafish (D) Class 2-like peaks. Note common identification
of SOX, EBF, and TEAD in top 6 motifs for each species. (E) Single-z images of 5 dpf
zebrafish with the wild-type version of eh7 and E1b minimal promoter driving GFP
expression (blue) in SCs, or version of eh7 in which all SOX sites have been deleted (delsox) or mutated (mut-sox). Note loss of GFP expression in SCs of utricle when SOX sites
are destroyed. DIC channels provide conte t. Scale bars are 50 μm.

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expression in the crista and utricle in zebrafish transgenic lines (Figure 39E). These
findings support SOX factors having a key role in the activity of the Class 2 eh7
enhancer.

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Figure 40. Analysis of Class 1 and 2 peaks in mouse sensory progenitors, design of SOX motif deletion, and gene pairs with
Class 1 and 2 peaks

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3.9 Discussion
Through cross-species profiling of the Atoh1 chromatin landscape, we discovered two
distinct classes of enhancers that regulate atoh1a expression during hair cell
development and regeneration. Whereas Class 1 enhancers have been shown
previously to have an autoregulatory function in amplifying Atoh1 expression in hair
cells, we also uncovered a more distal set of Class 2 enhancers that were accessible in
supporting cells of the neonatal mouse and adult regenerative species. Despite their
lack of accessibility in hair cells, we showed that these enhancers were necessary for
the initiation of atoh1a expression during hair cell development and regeneration of the
zebrafish inner ear but not lateral line. We further showed that similar Class 2-like
enhancers likely exist for many other key prosensory/early hair cell genes, with Class 2-
like enhancers maintaining accessibility in supporting cells of adult zebrafish but not
postnatal mice. Our data suggest that the ability of supporting cells to maintain
accessibility of Class 2-like enhancers into adulthood in regenerative species such as
zebrafish and lizards may facilitate their transdifferentiation into hair cells upon injury.
3.9.1 Requirement of Class 2 enhancers for atoh1a initiation in the zebrafish inner
ear
Figure 40. Analysis of Class 1 and 2 peaks in mouse sensory progenitors, design of
SOX motif deletion, and gene pairs with Class 1 and 2 peaks. (A) Heatmap showing
accessibility of Class 1 and Class 2 peaks within 200kb of HC genes in E13.5 cochlear
prosensory progenitors(16). (B) Venn diagram showing number of HC genes with only
Class 1 peaks, only Class 2 peaks, or both. (C,D) Genome Browser tracks show
accessibility of Class 1 (yellow) and Class 2 (purple) peaks near mouse and zebrafish
gene pairs in E13.5 progenitors(16) (PG), P1 HCs, and P1 and P6 SCs(15) (mouse); and
14 dpf HCs and 14 dpf and 12 mpf SCs (zebrafish). (E) Schematics show position of
predicted SOX binding sites (red) within the zebrafish eh7 enhancer and their deletion or
mutation in eh7_del-sox-E1b:GFP and eh7_mut-sox-E1b:GFP transgenic constructs.

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We discovered a distinct set of enhancers that are not accessible in hair cells where
atoh1a is expressed, but rather in developmental progenitors that give rise to hair cells
and later in adult supporting cells that can transdifferentiate into hair cells after damage.
Enhancers associated with the development of a cell type are often defined as being
differentially accessible in that cell type (Heinz et al., 2010; Nott et al., 2019; Rajderkar
et al., 2024; Tao et al., 2021), similar to the previously identified Atoh1 autoregulatory
(Class 1) enhancers (Helms et al., 2000; Luo et al., 2022). However, our data suggest
that Class 2 enhancers are only transiently required for atoh1a initiation and then rapidly
lose accessibility as hair cells form, in contrast to Class 1 autoregulatory enhancers that
then amplify and maintain atoh1a expression during hair cell differentiation. This may
explain why enhancers initiating Atoh1 expression were previously missed, and likely
reflects a bias when searching for the earliest enhancers that initiate gene expression
during developmental and regenerative transitions.
Whereas we did not capture early sensory progenitors in our zebrafish single-cell
analyses, reanalysis of embryonic mouse datasets indicate that Class 2 enhancers
become accessible and active in E13.5 bipotent prosensory progenitors, where
chromatin accessibility related to competence for hair cell specification is established
(Wang et al., 2023). Along with the lack of atoh1a e pression in homozygous Δeh5-9
zebrafish deleted for all Class 2 enhancers, these results suggest Class 2 enhancers
may be responsible for establishing atoh1a expression in prosensory progenitors prior
to hair cell fate specification. Additionally, when paired with the endogenous atoh1a
promoter, Class 2 enhancers were silent in supporting cells yet could be activated when

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Notch signaling was inhibited or hair cells ablated. This is consistent with our hypothesis
that Class 2 enhancers poise supporting cells to initiate Atoh1 expression for hair cell
transdifferentiation in zebrafish and potentially other regenerative species. Our results
also highlight important caveats when using minimal versus endogenous promoters for
in vivo enhancer testing.
3.9.2 Distinct enhancer requirements for inner ear versus lateral line development
and regeneration
hair cell development in both the zebrafish inner ear and lateral line requires atoh1a
(Millimaki et al., 2007; Sarrazin et al., 2006). Whereas inner ear hair cells have a
preferential requirement for Class 2 enhancers during development and regeneration,
lateral line hair cell formation preferentially require Class 1 autoregulatory enhancers. In
Class 2 enhancer deletion fish (Δeh5-9), we removed three enhancers with preferential
activity in inner ear supporting cells (eh7-9) and one with selective activity in lateral line
supporting cells (eh5). While the milder effects on lateral line hair cell development and
regeneration in Δeh5-9 fish could be due to redundant lateral line Class 2 enhancers
outside the deletion region, stronger effects on lateral line hair cells upon Class 1
enhancer deletion (Δeh1-6) suggest that molecular and cellular differences between the
inner ear and lateral line may underlie distinct enhancer dependencies. Consistently,
our previous single-cell RNA expression analyses revealed distinct molecular signatures
of inner ear and lateral line supporting cells (Shi et al., 2023). In the developing inner
ear, hair cells arise from transient prosensory progenitors, equally potent to differentiate
as hair cells or supporting cells, through initially stochastic upregulation of Atoh1 (Cai et

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al., 2013; Chen et al., 2002; Driver et al., 2013). In contrast in the lateral line, hair cells
are born in pairs from the cell division of specified hair cell progenitors (Itoh and Chitnis,
2001; López-Schier and Hudspeth, 2006; Nechiporuk and Raible, 2008). In addition,
several decades of studies have shown that hair cell regeneration in the inner ears of
vertebrate species largely occurs through transdifferentiation of mature supporting cells
with or without cell division (Roberson et al., 2004; White et al., 2006). In line with a
recent study showing inner ear hair cell regeneration is also uncoupled from cell division
in the zebrafish inner ear (Beaulieu et al., 2024), our lineage tracing demonstrated direct
transdifferentiation of supporting cells in the zebrafish cristae. In contrast, lateral line
hair cell regeneration is dependent on proliferation of a specific population of hair cell
progenitors (Mackenzie and Raible, 2012; Romero-Carvajal et al., 2015). One possibility
is that during development and regeneration of the lateral line, hair cell progenitors
maintain a low level of atoh1a expression, and due to their more rapid turnover have a
greater dependency on Class 1 enhancer autoregulation to efficiently boost atoh1a
expression. In the inner ear, induction of atoh1a expression from a prosensory
progenitor during development, or a mature supporting cell during regeneration, may
require de novo atoh1a expression through Class 2 initiation enhancers. These findings
underscore the importance of studying hair cell development and regeneration in
diverse mechanosensory organs.
3.9.3 Maintenance of Class 2 enhancer accessibility in supporting cells of
regenerative species

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We found that the spatial arrangements of Class 1 and 2 Atoh1 enhancers are
conserved between zebrafish, lizard, and mouse, with two of these enhancers
sequence-conserved. Whereas Class 2 enhancers remain accessible in adult inner ear
supporting cells of zebrafish and the green anole lizard, they lose accessibility in
supporting cells of the mouse cochlea shortly after birth. The selective ability of
zebrafish and lizard supporting cells to maintain Class 2 enhancer accessibility may
therefore contribute to the lifelong ability of supporting cells to upregulate Atoh1 and
transdifferentiate into hair cells during regeneration. Consistently, we observed failure of
inner ear hair cell regeneration following deletion of all Class 2 enhancers in zebrafish.
We also reanalyzed previously published snATAC-seq data for the regeneration of adult
zebrafish utricles (Jimenez et al., 2022) and did not identify any additional peaks gaining
accessibility around the atoh1a locus during zebrafish hair cell regeneration (Figure
35L), supporting Class 2 enhancers being the main regenerative enhancers in the inner
ear.
Differences in Atoh1 regulation alone are unlikely to fully explain regenerative
differences between zebrafish and mouse as previous studies forcing Atoh1 expression
in postnatal mouse supporting cells have been unsuccessful in inducing hair cell
transdifferentiation (Kelly et al., 2012; Liu et al., 2012). Indeed, our global analysis of
chromatin accessibility in mouse and zebrafish revealed that many other genes
necessary for early hair cell fate specification have Class 2-like enhancers selectively
accessible in supporting cells. These genes included several transcription factors
involved in prosensory specification and hair cell competence (Ahmed et al., 2012;

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Bricaud and Collazo, 2006; Wang et al., 2023), such as Six1, Eya1, and Sox4/11 family
members (Figure 40). As for Atoh1, Class 2-like enhancers as a group maintain
accessibility in zebrafish supporting cells throughout life but are silenced in the postnatal
mouse cochlea. A recent single-cell study of the regenerating zebrafish inner ear
revealed that supporting cells transiently pass through a progenitor-like state during
their transdifferentiation into hair cells (Jimenez et al., 2022). It is therefore possible
that, in addition to re-initiating atoh1a expression, Class 2-like enhancers more broadly
promote a progenitor-like state in zebrafish supporting cells during inner ear
regeneration by activating prosensory genes necessary for hair cell specification
following injury.
An unanswered question is why zebrafish but not mouse Class 2-like enhancers can
maintain accessibility in supporting cells through later stages. By identifying transient
Class 2-like enhancers in the neonatal mouse and comparing to those of zebrafish, we
identified several transcription factor binding motifs in common, with the top motif being
SOX. Both the zebrafish and mouse versions of the eh7/Eh4 Class 2 enhancer contain
conserved predicted SOX sites, and mutation of these sites destroyed enhancer activity
in zebrafish transgenic assays. In addition to SOX, we also found enrichment of SIX
motifs in mouse and fish Class 2 enhancers, consistent with known roles of SOX and
SIX factors in orchestrating inner ear hair cell regeneration in zebrafish (Jimenez et al.,
2022). Whereas a sox2 enhancer has recently been shown to be required for timely hair
cell regeneration in zebrafish (Jimenez et al., 2022), continued expression of Sox2 in
postnatal supporting cells of mouse argues against it being a differential factor

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regulating Class 2 enhancer accessibility in zebrafish (Wang et al., 2023). Instead,
SoxC transcription factors (Sox4 and Sox11) have been shown to be required for hair
cell competence in mouse prosensory progenitors and are rapidly downregulated in
postnatal mouse supporting cells (Wang et al., 2023). Further, Eh4 and Eh5 Class 2
enhancers of Atoh1 are directly bound by Sox4, and their accessibility in E13.5
prosensory progenitors is lost when Sox4 and Sox11 are deleted in the otic lineage
(Wang et al., 2023). It is therefore possible that continued expression of Sox4/11 family
members in zebrafish but not mouse supporting cells may allow continued competence
for hair cell generation. Alternatively or in addition, Class 2-like enhancers may be
selectively silenced in postnatal mouse supporting cells. For example, EBF was the
second motif for mouse and fourth for zebrafish, and recent studies found a repressive
role of Ebf1 in hair cell formation in the developing mouse cochlea (Kagoshima et al.,
2024; Powers et al., 2024). Future studies will be required to determine which
transcriptional activators and/or repressors differentially regulate the activity of Class 2
enhancers and regenerative capacity across vertebrates.

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Chapter 4. Conclusions
4.1 Zebrafish lateral line versus inner ear as a model to study hair cell development
and regeneration
As mentioned in Chapters 2 and 3, we identified both transcriptomic signature and
distinct epigenetic mechanisms in the zebrafish inner ear. We further demonstrated that
the transcriptomes of zebrafish inner ear hair cell subtypes aligned better to those of
their mammalian counterparts than those of the lateral line hair cells. Although we did
not test whether deletion of Class 2 Atoh1 enhancers in mice would result in similar
defects seen in the zebrafish inner ear, the deletion of Class 1 enhancers did not
eliminate Atoh1 expression nor hair cells in the mouse cochlea, suggesting the
existence of independent enhancers that would be able to initiate Atoh1 expression,
likely to be Class 2 enhancers (Luo et al. 2022). Yet, the deletion of Class 1 enhancers
seemed to have a much more severe phenotype to cochlear hair cell development than
what we observed in the zebrafish inner ear with Class 2 deletion, and it seemingly
resembled the lateral line phenotype in zebrafish when Class 1 enhancers were
deleted. This could be a result of the precise timing required for the development of the
cochlea (Groves et al., 2013), where the upregulation of Atoh1 via autoregulatory Class
1 enhancers may need to happen within a precise window for hair cells to develop
normally. In contrast, in the utricles of the zebrafish with Class 1 enhancer deletion,
Atoh1a level can still slowly accumulate through the activity of Class 2 enhancers and
allow a delayed hair cell development. This will need to be tested in the mouse utricle
as Luo et al. did not demonstrate the effect of Class 1 enhancer deletion in the mouse
utricle. All being said, it is worth further investigating and comparing both the zebrafish

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inner ear and lateral line to the mouse inner ear, and to determine whether one or the
other sensory organ in the zebrafish is more similar to mammalian hair cell biology and
will yield more translatable results.
4.2 Challenges to using the zebrafish inner ear
Using the zebrafish inner ear as a model to study hair cell biology is not without its own
challenges. First of all, due to its internal location, the inner ear is less accessible for
drug treatment. Though Jimenez et al. demonstrated that they were able to ablate hair
cells in myo6b:hDTR fish by simply adding diphtheria toxin (DT) to larval zebrafish
water, we had to resort to injecting DT directly into the inner ear to see any hair cell
ablation. This creates a significant amount of learning experience and labor in order to
execute a hair cell ablation experiment. Luckily, another inner ear hair cell ablation
model exists using capsaicin to kill hair cells ectopically expressing the rat TRPV1
receptor (Beaulieu et al. 2024). Nonetheless, the lack of access to drug treatment in the
zebrafish inner ear may hinder many other research projects unrelated to hair cell
ablation. Along the same line, the deeper location of the inner ear creates difficulty in
confocal imaging as well, especially in juvenile and adult zebrafish and for fluorescent
reporters with weak activities.
The limited availability of transgenic tools to specifically label cells of the inner ear also
creates difficulties to isolate a pure inner ear cell population for NGS experiments
without contamination from the lateral line neuromasts, as many of the transgenic lines
available label both the lateral line and the inner ear. One of the efforts of this study was

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to identify inner ear supporting cell-specific enhancers to drive the expression of
fluorescent reporter proteins. Even though we identified a kcnq5a enhancer that had
inner ear-specific activity, it was either restricted to the cristae supporting cells or very
weakly active in the utricle depending on the allele, which could be the result of random
integration of the transgene into the zebrafish genome via Tol2 transgenesis.
Additionally, in our single-cell datasets, we were only able to identify a few supporting
cells compared to the large number of hair cells, suggesting that our cell dissociation
and isolation protocols may need to be further optimized to recover a more
representative population of supporting cells.
4.3 Multitude of epigenetic barriers to hair cell regeneration
In this study, we only investigated the chromatin accessibility aspect of the epigenetic
control of hair cell genes in inner ear supporting cells of the zebrafish. We found that
zebrafish supporting cells were able to maintain the accessibility of Class 2 enhancers,
which were established during development and required for hair cell regeneration. In
contrast, Class 2 enhancers become inaccessible in mouse cochlear supporting cells by
P6, correlating to their loss of transdifferentiation potential. This suggests that the ability
to maintain Class 2 hair cell enhancer accessibility could explain why zebrafish but not
mouse supporting cells can transdifferentiate into hair cells upon injury. We further
hypothesized that this long-term maintenance of Class 2 enhancers is a result of
continued expression and binding of certain transcription factors in the zebrafish but not
mouse supporting cells, as it has been shown that the binding of transcription factors to
DNA can positively regulate chromatin accessibility (Friman et al., 2019; Lamparter et

84
al., 2017). Such transcription factors can be identified in a future meta-analysis
comparing the transcriptomes of mouse versus zebrafish supporting cells.
However, it is unlikely that the binding of transcription factors and the maintenance of
accessible chromatin are the only reasons why zebrafish supporting cells are able to
regenerate hair cells. It has been shown that multiple layers of epigenetic silencing,
including removal of permissive histone marks and encroachment of DNA methylation
to hair cell enhancers, are also involved in the permanent silencing of hair cell genes in
postnatal cochlear supporting cells (Tao et al. 2021; Nguyen et al. 2023). However, it is
not clear what the histone modification and DNA methylation statuses are like for Class
2 enhancers in zebrafish supporting cells due to technical challenges related to ChIP
and CUT&RUN sequencing, as well as whole genome bisulfite sequencing, using the
limited cell population we can collect. Based on the fact that Class 2 enhancers remain
competent to drive fluorescent reporter activity in supporting cells until at least juvenile
stages in zebrafish, we hypothesize that they are associated with the active histone
mark H3K27ac and are hypomethylated at CpG dinucleotides within the enhancers.
4.4 Reestablishment of Class 1 enhancers in zebrafish supporting cells
While Class 2 enhancers remain accessible in zebrafish supporting cells, Class 1 hair
cell enhancers become mostly silenced, similar to what is observed in postnatal mouse
supporting cells (Tao et al. 2021). This suggests that zebrafish supporting cells may
need to establish Class 1 enhancers de novo during transdifferentiation. In addition, we
found that several transcription factors downstream of Atoh1, including Pou4f3 and Gfi1

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(Jen et al., 2022; Masuda et al., 2011; Yu et al., 2021), are only associated with Class 1
enhancers, suggesting the expression of these hair cell transcription factors may not be
immediately required for supporting cells to start the transdifferentiation process, but are
more related to hair cell maturation. If this is the case, then how are Class 1 enhancers
reestablished in transdifferentiating supporting cells? Does it involve certain pioneer
factors that are regulated by Class 2 enhancers? Will allowing these pioneer factors to
express via activating Class 2 enhancers in mammalian supporting cells also
spontaneously reestablish Class 1 enhancers? Or will this still not be sufficient as
zebrafish and mouse Class 1 enhancers maybe have different statuses regarding
histone modification and DNA methylation that allow zebrafish but not mouse Class 1
enhancers to reopen during regeneration? This series of questions will need to be
answered in a future study to better understand the epigenetic basis of hair cell gene
regulation in the zebrafish inner ear and how it differs from the non-regenerative mouse
cochlea.

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Materials & Methods
Materials & methods for Chapter 2:
Zebrafish lines
The Institutional Animal Care and Use Committees of the University of Southern
California (Protocol 20771) and University of Washington (Protocol 2997-01) approved
all animal experiments. Experiments were performed on zebrafish (Danio rerio) of AB or
mixed AB/Tubingen background. For adult stages, mixed sexes of animals were used
for constructing single-cell libraries, as well as RNAScope experiments. Published lines
include Tg(Mmu.Sox10-Mmu.Fos:Cre)zf384,(Kague et al., 2012); Tg(-3.5ubb:LOXPEGFP-LOXP-mCherry)cz1701Tg,(Mosimann et al., 2011); and
Tg(myosin6b:GFP)w186
,(Xiao et al., 2005).
In situ hybridization and RNAScope
Hybridization chain reaction in situ hybridizations(Choi et al., 2018) (Molecular
Instruments, HCR v3.0, PMID: 29945988) were performed on 5 dpf myo6b:GFP larvae
as directed for whole-mount zebrafish embryos and larvae(Choi et al., 2016) (PMID:
27702788). Briefly, embryos were treated with 1-phenyl 2-thiourea (PTU) beginning at
24 hpf. At 5 dpf, larvae were fixed in 4% PFA overnight at 4°C. Larvae were washed
with PBS and then stored in MeOH at -20°C until use. Larvae were rehydrated using a
gradation of MeOH and PBST washes. Larvae were treated with proteinase K for 25
min and post-fixed with 4% PFA for 20 min at room temperature. For the detection
phase, larvae were pre-hybridized with probe hybridization buffer for 30 min at 37°C,
then incubated with probes overnight at 37°C. Larvae were washed with 5X SSCT to

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remove probes. For the amplification stage, larvae were pre-incubated with amplification
buffer for 30 min at room temperature while hairpin solution was prepared. Larvae were
incubated with hairpins overnight in the dark at room temperature. Hairpins were
removed by washing with 5X SSCT. Larvae were treated with DAPI and stored at 4°C
until imaging. All HCR in situ patterns were confirmed in at least 3 independent animals.
RNAScope samples were prepared by fixation in 4% paraformaldehyde either at room
temperature for 2 hours or at 4 °C overnight. Adult inner ears were dissected and
dehydrated in methanol for storage. RNAScope probes were synthesized by Advanced
Cell Diagnostics; Channel 1 probe myo6b, Channel 2 probe pvalb9, and Channel 3
probes cabp1b and cabp2b. Whole inner ear tissues were processed through the
RN Scope Fluorescent Multiple V2 ssay according to manufacturer’s protocols with
the ACD HybEZ Hybridization oven. All RNAScope patterns were confirmed in at least 3
independent animals.
Imaging
Confocal images of whole-mount RNAScope samples were captured on a Zeiss
LSM800 microscope (Zeiss, Oberkochen, Germany) using ZEN software. HCR-FISH
imaging was performed on a Zeiss LSM880 microscope (Zeiss, Oberkochen, Germany)
with Airyscan capability. Whole larvae were mounted between coverslips sealed with
high vacuum silicone grease (Dow Corning) to prevent evaporation. Z-stacks were
taken through the ear at intervals of 1.23 μm using a 10X objective or through individual

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inner ear organs at an interval of 0.32 μm using a 20X objective. 3D iryscan
processing was performed at standard strength settings using Zen Blue software.
Single-cell preparation and analysis
scRNA-seq library preparation and alignment
For 14 dpf animals, heads from converted Sox10:Cre; ubb:LOXP-EGFP-LOXP-mCherry
fish were decapitated at the level of the pectoral fin with eyes and brains removed. For
12 mpf animals, utricle, saccule, and lagena were extracted from converted Sox10:Cre;
ubb:LOXP-EGFP-LOXP-mCherry fish after brains and otolith crystals were removed.
Dissected heads and otic sensory patches were then incubated in fresh Ringer’s
solution for 5–10 min, followed by mechanical and enzymatic dissociation by pipetting
every 5 min in protease solution (0.25% trypsin (Life Technologies, 15090-046), 1 mM
EDT , and 400 mg/mL Collagenase D (Sigma, 11088882001) in BS) and incubated at
28.5 °C for 20–30 min or until full dissociation. Reaction was stopped by adding 6× stop
solution (6 mM CaCl2 and 30% fetal bovine serum (FBS) in PBS). Cells were pelleted
(376 × g, 5 min, 4 °C) and resuspended in suspension media (1% FBS, 0.8 mM CaCl2,
50 U/mL penicillin, and 0.05 mg/mL streptomycin (Sigma-Aldrich, St. Louis, MO) in
phenol red-free Leibovitz’s L15 medium (Life Technologies)) twice. Final volumes of
500 μL resuspended cells were placed on ice and fluorescence-activated cell sorted
(FACS) to isolate live cells that excluded the nuclear stain DAPI; see Supplemental Fig.
for gating strategy. For scRNAseq library construction, barcoded single-cell cDNA
libraries were synthesized using 10X Genomics Chromium Single Cell 3′ Library and
Gel Bead Kit v.3.1 (14 dpf) or Single Cell Multiome ATAC + Gene Expression kit (12

89
mpf, T C data not shown) per the manufacturer’s instructions. Libraries were
sequenced on Illumina NextSeq or HiSeq machines at a depth of at least 1,000,000
reads per cell for each library. Read2 was extended from 98 cycles, per the
manufacturer’s instructions, to 126 cycles for higher coverage. Cellranger v6.0.0 (10X
Genomics) was used for alignment against GRCz11 (built with GRCz11.fa and
GRCz11.104.gtf) and gene-by-cell count matrices were generated with default
parameters.
Data processing of scRNA-seq
Count matrices of inner ear and lateral line cells from embryonic and larval timepoints
(18-96 hpf) were analyzed using the R package Monocle3 (v1.0.0) (Cao et al., 2019;
Qiu et al., 2017; Trapnell et al., 2014). Matrices were processed using the standard
Monocle3 workflow (preprocess_cds, detect_genes, estimate_size_factors,
reduce_dimension(umap.min_dist = 0.2, umap.n_neighbors = 25L)). This cds was
converted to a Seurat object for integration with 10X Chromium sequencing data using
SeuratWrappers. The count matrices of scRNA-seq data (14 dpf and 12 mpf) were
analyzed by R package Seurat (v4.1.0) (Hao et al., 2021). Cells of neural crest origins
were removed bioinformatically based on our previous study (Fabian et al., 2022)
(ref,Fabian). The matrices were normalized (NormalizeData) and integrated with
normalized scRNA-seq data from the embryonic and larval time points according to
package instruction (FindVariableFeatures, SelectIntegrationFeatures,
FindIntegrationAnchors, IntegrateData; nfeatures = 3000). The integrated matrices were
then scaled (ScaleData) and dimensionally reduced to 30 principal components. The

90
data were then subjected to neighbor finding (FindNeighbors, k = 20) and clustering
(FindClusters, resolution = 0.5), and then visualized through UM with 30 principal
components as input. After data integration and processing, RNA raw counts from all
matrices were normalized and scaled according to package instructions to determine
gene expression for all sequenced genes, as the integrated dataset only contained
selected features for data integration.
Mouse utricle scRNA-seq data (Jan et al., 2021) was downloaded from NCBI Gene
Expression Omnibus (GSE155966). The count matrix was analyzed by R package
Seurat (v4.1.0). Matrices were normalized (NormalizeData) and scaled for the top 2000
variable genes (FindVariableFeatures and ScaleData). The scaled matrices were
dimensionally reduced to 15 principal components. The data were then subjected to
neighbor finding (FindNeighbors, k = 20) and clustering (FindClusters, resolution = 1)
and visualized through UMAP with 15 principal components as input. Hair and
supporting cells were bioinformatically selected based on expression of hair and
supporting cell markers Myo6 and Lfng, respectively. Hair cells were further
subcategorized into striola type I hair cells by co-expression of striola marker Ocm and
type I marker Spp, extrastriola type I hair cells by expression of Spp without Ocm, and
extrastriola type II hair cells by expression of Anxa4 without Ocm.
Mouse crista scRNA-seq data (Wilkerson et al., 2021) was downloaded from NCBI
Gene Expression Omnibus (GSE168901). The count matrix was analyzed by R
package Seurat (v4.1.0). Matrices were normalized (NormalizeData) and scaled for the
top 2000 variable genes (FindVariableFeatures and ScaleData). The scaled matrices

91
were dimensionally reduced to 15 principal components. The data were then subjected
to neighbor finding (FindNeighbors, k = 20) and clustering (FindClusters, resolution = 1)
and visualized through UMAP with 15 principal components as input. Hair and
supporting cells were bioinformatically selected based on expression of hair and
supporting cell markers Pou4f3 and Sparcl1, respectively. Hair cells were further
subcategorized into central hair cells by expression of Ocm and peripheral hair cells by
expression of Anxa4.
Pseudotime analysis
We used the R package Monocle3 (v1.0.1) to predict the pseudotemporal relationships
within the integrated scRNA-seq dataset of sensory patches from 36 hpf to 12 mpf. Cell
paths were predicted by the learn_graph function of Monocle3. We set the origin of the
cell paths based on the enriched distribution of 36 to 48 hpf cells. Hair (all macular hair
cells, clusters 0-5) and supporting (macular supporting cells clusters 0 and 6) cell paths
were selected separately (choose_cells) to plot hair and supporting cell marker
expression along pseudotime (plot_genes_in_pseudotime).
Differential gene expression
We utilized Monocle3 package’s differential gene e pression function for identification of
differentially expressed genes among the different cell types with a q-value of 0.01 or
smaller, using a negative binomial distribution based generalized linear model. To
compare inner ear hair cells to lateral line hair cells, we used the following datasets from
GEO: GSE144827 (Kozak et al., 2020) , GSE152859 (Ohta et al., 2020), and

92
GSE196211 (Baek et al., 2022). Hair cells were selected from datasets by expression of
otofb and integrated along with our 10x Chromium dataset with Scanorama (Hie et al.,
2019).
SAMap analysis for cell type homology
We used the python package SAMap (v1.0.2) (Tarashansky et al., 2021) to correlate
gene expression patterns and determine cell type homology between mouse utricle
(GSE155966)(Jan et al., 2021) or crista (GSE168901)(Wilkerson et al., 2021) hair and
supporting cells and our 12 mpf zebrafish inner ear scRNA-seq data. Zebrafish lateral
line hair cell sc-RNA data (GSE123241)(Lush et al., 2019) was integrated with our 12
mpf inner ear data using Seurat in order to compare to mouse. First, a reciprocal
BLAST result of the mouse and zebrafish proteomes was obtained by performing blastp
(protein-protein BLAST, NCBI) in both directions using in-frame translated peptide
sequences of zebrafish and mouse transcriptome, available from ensembl
(Danio_rerio.GRCz11.pep.all.fa and Mus_musculus.GRCm38.pep.all.fa). The
generated maps were then used for the SAMap algorithm. Raw count matrices of
zebrafish and mouse scRNA-seq Seurat objects with annotated cell types were
converted to h5ad format using SeuratDisk package (v0.0.0.9020) and loaded into
Python 3.8.3. Raw data were then processed and integrated by SAMap. Mapping
scores between cell types of different species were then calculated by
get_mapping_scores and visualized by sankey_plot. Gene pairs driving cell type
homology were identified by GenePairFinder.

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Materials & methods for Chapter 3:
Experimental model and subject details
Zebrafish We raised zebrafish in a vivarium under standard conditions at 28.5°C with
health and water conditions monitored daily. Heterozygous mutant adult fish were raised
in similar conditions and showed no visible defects, with homozygous mutant embryos
produced by breeding for experimental analysis. Published mutant and transgenic lines
include: Tg(Mmu.Sox10-Mmu.Fos:Cre)zf384,
(Kague et al., 2012); Tg(–3.5ubb:LOXPEGFP-STOP-LOXP-mCherry)cz1701,
(Mosimann et al., 2011);
Tg(myosin6b:GFP)w186,
(Hailey et al., 2017); atoh1aw271Tg,
(Hewitt et al., 2024) and
Tg(myo6b:Hsa.HBEGF,myl7:EGFP)hg148Tg,
(Jimenez et al., 2021). Transgenic lines
generated in this study include Tg(atoh1a_eh1-Mmu_E1b:mCherry)el950; el951;
Tg(atoh1a_eh2-Mmu_E1b:mCherry)el952; Tg(atoh1a_eh3-Mmu_E1b:mCherry)el953 ;el954;
Tg(atoh1a_eh4-Mmu_E1b:mCherry)el955; el956; Tg(atoh1a_eh5-Mmu_E1b:GFP)el957; el958;
el959; Tg(atoh1a_eh6-Mmu_E1b:GFP)el960; el961; Tg(atoh1a_eh7-Mmu_E1b:GFP)el962; el963;
Tg(atoh1a_eh8-Mmu_E1b:GFP)el964; Tg(atoh1a_eh9-Mmu_E1b:GFP)el965; el966;
Tg(atoh1a:GFP)el967; el968; el969; Tg(atoh1a_eh2-atoh1a:mCherry)el970; Tg(atoh1a_eh7-
atoh1a:GFP)el972; el973; Tg(Mmu_Atoh1_Eh4-Mmu_E1b:GFP)el974; el975; Tg(atoh1a_eh7_
del-sox-Mmu_E1b:GFP)el976; Tg(atoh1a_eh7_mut-sox-Mmu_E1b:GFP)el977; el978 ;el979;
Tg(kcnq5a_p1-Mmu_E1b:nlsEOS)el1033
. Enhancer deletion alleles generated in this study
include atoh1ael980; el981; el982 (Δe -4); atoh1ael983 (Δe -6); atoh1ael984 (Δe 5-9);
atoh1ael985; el986 (Δe 6); atoh1ael987 (Δe 7-8); atoh1ael988 (Δe 9); and atoh1ael989 (Δe 7-
9). All transgenic and mutant lines generated in this study are on a Tubingen background.
Developmental stages and numbers of embryos used are described for each experiment.

94
As zebrafish develop gender differences after most of the stages studied here, the gender
of embryos was undetermined. For 12 mpf zebrafish used for single-cell multiome library
construction, both male and female zebrafish were sampled. To genotype atoh1a
enhancer deletion alleles, both external and internal primers were used to PCR amply
genomic DNA. Since external primers for each enhancer deletion allele flank large
genomic regions, wild type yielded expected products with internal primers but no product
with external primers. Heterozygous enhancer deletion zebrafish were identified by
presence of both internal and external PCR products, while homozygous animals were
identified by the absence of internal PCR products but presence of external PCR products.
External PCR products for enhancer deletion alleles were subsequently sent for Sanger
sequencing to confirm enhancer deletion.
Mice All experiments were conducted in accordance with the policies of the Institutional
Animal Care and Use Committees of the Keck School of Medicine at the University of
Southern California. Wild-type mice of CD1 or FVB/NJ genetic backgrounds were
obtained from Charles River and the Jackson laboratory, respectively. Animals were
housed with free access to food and water in 12-hr light/dark cycle. Neonatal animals
were anesthetized on ice following decapitation, and CO2-induced euthanasia followed
by cervical dislocation was used on young-adult mice prior to tissue collection.
Lizards Green anole lizards were housed in metal mesh cages under 12-h light/12-h dark
schedules using 50 W spot basking heat lamps and UVB lamps during light hours. Lizards
were maintained at 65% humidity with temperatures of 24–26 °C during light hours and

95
18.5–21 °C during dark hours. Cages were misted with water five times per week, and
lizards were fed ½-inch crickets dusted in calcium supplement three times per week. Prior
to tissue collection, lizards were euthanized according to the protocol outlined in Conroy
et al., 2009, JAALAS (Conroy et al., 2009): 1. Intracoelomic injection of 25-50 ul pHneutralized 1% MS222. 2. After loss of righting reflex, another intracoelomic injection of
25-50 ul 50% MS222. 3. Decapitation. 4. Double pithing of brains/spinal cords.
Zebrafish transgenesis
To test putative enhancers for zebrafish atoh1a (eh1-9), kcnq5a p1, Mouse Atoh1 (Eh4),
and eh7 lacking SOX binding sites, IDT gBlocks containing corresponding accessible
chromatin regions were cloned into a plasmid containing Mmu.E1b minimal promoter,
mCherry or GFP fluorescent reporter, and cryaa:Cerulean as a co-selection marker,
flanked by Tol2 transposase sequences (Tol2kit(Kwan et al., 2007)), using In-Fusion
cloning. Mmu.E1b promoter was replaced by gBlock containing the atoh1a endogenous
promoter using restriction digestion followed by In-Fusion cloning to construct
atoh1a_eh7-Patoh1a:GFP. To generate zebrafish lines carrying fluorescent reporter
transgenes, donor plasmids (final concentration of 20 ng/μl) were co-injected with Tol2
transposase RN (final concentration of 30 ng/μl) into one-cell-stage embryos. In most
instances, two to three independent founders were identified per gene construct. Reporter
activities observed in at least three embryos of each allele were reported in this study.
Enhancer deletion

96
All CRISPR guides were designed using CHOPCHOP
(https://chopchop.cbu.uib.no/)(Labun et al., 2019) and synthesized as crRNAs from
Integrated DNA Technologies (IDT). Each crRNA and tracrRNA (IDT, 1072532) were
diluted to 100 μM in IDTE buffer (IDT, 11-01-02-02). Equal amount of each crRNA was
then mixed with tracrRNA in a nuclease-free duplex buffer (IDT, 11-01-03-01) to a final
concentration of 36 ng/μl for the crRN and 67 ng/μl for the tracrRN . The mi ture was
then heated to 95°C for 5 min and allowed to cool slowly to RT. To prepare Cas9 protein,
we diluted IDT Alt-R™ S.p. Cas9 Nuclease V3 (1081058) to 1 μg/μl. Prior to injection, we
mi ed 2 μl duple ed left gRN and 2 μl duple ed right gRN with 1 μl of diluted Cas9
protein and incubated the mixture at 37°C for 10 min. The mixture was then injected into
one-cell-stage zebrafish embryos. We identified 3 founders for Δe -4, 1 founder for
Δe -6, 1 founder for Δe 5-9, and 1 founder for Δe 7-9 from at least 100 injected animals
for each deletion type. We also identified 2 founders for Δe 6, 1 founder for Δe 7-8, and
1 founder for Δe 9 from 12, 20, and 18 injected animals, respectively. Each enhancer
deletion was verified by genotyping and subsequent sequencing of the PCR product.
Sectioning
For paraffin sections, we fixed genotyped larval zebrafish in 4% PFA/1X PBS at 4°C
overnight followed by dehydration through EtOH series (two 1X PBS, one 30% EtOH, one
50% EtOH, one 70% EtOH, one 95% EtOH, and two 100% EtOH for 20 min each at RT).
Prior to paraffin embedding, samples were infused with Hemo-De (xylene substitute,
Electron Microscopy Sciences 23412–01) through one 50% EtOH/50% Hemo-De, one
25% EtOH/75% Hemo-De, and one 100% Hemo-De wash of 15 min each at RT. Samples

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were then placed at 65°C in 50% Hemo-De/50% paraffin for one hour, followed by 100%
paraffin overnight. After embedding in paraffin, blocks were left overnight at RT to harden.
We cut blocks into 6 μm sections on a Shandon Finesse Me+ microtome (cat. no.
77500102) with High-Profile Disposable Surgipath DB80 HS Blades (VWR, 10015-022),
and collected sections on Apex Superior Adhesive slides (Leica Microsystems, cat. no.
3800080).
RNA in situ hybridization
RNAScope probes were synthesized by Advanced Cell Diagnostics, with myo6b on
Channel 1, atoh1a on Channel 2, and lfng on Channel 3. PFA-fixed paraffin-embedded
sections were deparaffinized, and RNAScope Fluorescent Multiplex v2 Assay was
performed according to manufacturer’s protocol ( dvanced Cell Diagnostics, protocol
323100-USM) with optimization. Specific modifications to the RNAscope protocol
included using a 4 min steaming step during target retrieval and a 30 min protease step
using the provided Protease Plus reagent. Probes were visualized using TSA Fluorescein
(1:750, Akoya, NEL701A001KT), TSA Cy3 (1:750, Akoya, NEL744001KT), and TSA Cy5
(1:750, Akoya, NEL745001KT). Slides were then mounted with DAPI Fluoromount-G
Mounting Solution (Southern Biotech, 0100-20). Expression level of atoh1a indicated by
RNAScope puncta was quantified using Fiji108 following manufacturer’s recommendation
(ACD, SOP 45-006 Technical Note).
Phalloidin Staining

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To visualize HCs, genotyped larval zebrafish were fixed in 4% PFA/1X PBS at RT for 2
hours, followed by one 10 min wash in 1X PBS and 30 min in 1X PBS/1% Triton-X. We
then stained zebrafish with 1:40 dilution of methonolic Phalloidin stock (Alexa Fluor 647
Phalloidin, Invitrogen A22287) in 1X PBS for 2 hours at RT. Samples were rinsed several
times with PBS before imaging. We opted for a same-day phalloidin staining to ensure
that the otolith crystals in the otic vesicles did not dissolve overnight, which served as a
landmark for the utricle, especially in cases where HCs were ablated. HC number was
quantified in the sensory organs of at least three animals per genotype and condition
using Fiji(Schindelin et al., 2012).
Drug treatments
To access the involvement of Notch signaling in the activation of Class 2 enhancers, we
treated Tg(atoh1a:GFP)el967; el968; el969; Tg(atoh1a_eh2-atoh1a:mCherry)el970; and
Tg(atoh1a_eh7-atoh1a:GFP)el972; el973 larval zebrafish with 10 μM DBZ (Tocris, 4489) or
DMSO control in embryo media starting at 4 dpf. Animals were in DBZ for up to 48 hours,
with DBZ refreshed every 24 hours. DBZ-treated animals were anesthetized and
subjected to confocal imaging at 24 or 48 hours post-treatment. To access the effect of
Δe -6 and Δe 5-9 on HC regeneration, we bred alleles atoh1ael983 and atoh1ael984 on
the Tg(myo6b:Hsa.HBEGF,myl7:EGFP)hg148Tg,
(Jimenez et al., 2021) background in order
to ablate HCs. As we were unable to ablate HCs by submerging animals in Diphtheria
Toxin (DT, Sigma-Aldrich, 322326) as originally described(Jimenez et al., 2021), we
instead injected DT directly into the otic vesicles of larval zebrafish. Briefly, a 1 ng/μl
solution was prepared by serial dilution in BS from a 1 mg/μl stock and phenol red was

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added to the final solution to guide injection. We anesthetized 4 dpf larval zebrafish
obtained from in-crossing atoh1ael983/+; Tg(myo6b:Hsa.HBEGF,myl7:EGFP)hg148Tg or
atoh1ael984/+; Tg(myo6b:Hsa.HBEGF,myl7:EGFP)hg148Tg animals and stabilized them on a
petri dish lid by removing as much water as possible. We then used the same setup as
embryo injection and used a borosilicate glass needle to inject diluted DT solution or PBS
control into the otic vesicle. Injected animals were immediately transferred to fresh
embryo media to recover. Feeding and water change was done daily post-injection.
Immediately after injection and every 24 hours thereafter, a portion of the injected fish
were euthanized and genotyped for either phalloidin staining or paraffin embedding.
AAV production and delivery
AAVie-K558 capsid plasmid, generously provided by Dr. Guisheng Zhong, was described
previously(Tao et al., 2022) and used to package the enhancer testing constructs. To
purify AAVs, CsCl-gradient centrifugation followed by virus isolation and dialysis was
performed (SBPD viral core). For inner ear delivery, 2 uL of AAV (1012 VG/mL) was
injected into the posterior semicircular canal of P0 mice anesthetized with ice(Isgrig and
Chien, 2018). Prior to surgery, hypothermic anesthesia was judged based on lack of
response to a firm toe pinch and Buprenorphine XR (MWI Veterinary Supply Co; 3.25
mg/kg) was administered subcutaneously for pain control. Sterilized forceps and scissors
were used to bisect the skin, subcutaneous fat, and muscle layer to expose the posterior
semicircular canal. Virus was injected free-hand over ~40 s using 10 uL Hamilton syringe
(Hamilton Company, #7653-01) outfitted with a 34-gauge needle (Hamilton Company,
#207434). The surgical site was sealed at the level of the skin with Vetbond (3M, #1469SB)

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and animals were placed on a 37°C heating pad for 30 min for full recovery before being
re-introduced to their mother. The injected animals were monitored twice daily for three
days post-surgery. After the third day, the injected animals were monitored daily until the
collection day.
Immunostaining
For whole-mount neonatal mouse organ preparations, the cochlea and utricles were
dissected from the AAV-injected animals and fixed in 4% paraformaldehyde for 10 min.
The organs were blocked for 4 hr at RT or overnight at 4°C with buffer containing 2 0mM
Tris-Buffered Saline (Bio-Rad), 0.1% Tween-20 (Sigma-Aldrich), 5% normal donkey
serum (Jackson Immuno Research), and 0.01% Sodium Azide (Sigma-Aldrich). The
primary antibodies were diluted in the same blocking buffer and incubated overnight at
4°C. The primary antibodies used include goat anti-Sox2 (R&D Systems AF2018), rabbit
anti-Myo7a (Proteus Bioscience 25-6790), rat anti-RFP (Proteintech 5f8-150), and
chicken anti-GFP (GeneTex GTX13970). Samples were washed with 20 mM TBS
supplemented with 0.1% Tween-20 (Sigma-Aldrich) (TBST) 3 times, 5 min each.
Secondary antibodies conjugated to Alexa Fluorophore dyes (Abcam) were diluted in the
same blocking buffer and incubated for 2 hr at RT or overnight at 4°C. For whole-mount
juvenile and adult organ preparations, the otic capsules from the AAV-injected animals
were dissected and fixed in 4% paraformaldehyde for 30 min at RT. To improve the
accessibility to the sensory organs, the bone surrounding the anterior semicircular canal,
utricle, and the apex of the cochlea was opened with forceps. After performing
immunostaining as described above, the otic capsules were decalcified with 0.5 mM

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EDTA (pH 8.0) overnight at 4°C. Then the utricle and cochlea were micro-dissected for
whole-mount imaging.
Imaging
All images in this study were captured on a Zeiss LSM800 confocal microscope using
ZEN software. For photoconversion, larval zebrafish were transferred to 12-well dish and
exposed to a handheld UV light for 15 min prior to imaging and/or DT injection. All images
shown are maximum intensity projections, unless otherwise specified. All imaging settings
were modified consistently across samples during imaging and during image processing
in Adobe Photoshop for each experiment.
Single nuclei isolation
Sox10:Cre; ubb:LOXP-EGFP-LOXP-mCherry zebrafish were screened for conversion of
GFP to mCherry in the cranial neural crest cell population and otic placode at 5 dpf under
a fluorescent dissection microscope. We then raised these animals to 14 dpf or 12 mpf
and sacrificed them for dissection and dissociation. We decapitated 14 dpf animals (n=40)
at the level of the pectoral fin and extracted the otic vesicles with some of the surrounding
tissues still attached. For 12 mpf animals (n=6, 27-31 mm, males and females equally
sampled), we extracted and pooled their utricle, saccules, and lagenas and removed
otolith crystals. For green anole lizards, we dissected and isolated the saccules from adult
lizard heads (n=7, between 2.5 and 3 cm at the point of nose to just after dewlap) and
removed otoconia crystals. The dissected tissues were incubated in fresh Ringer’s
solution for 5–10 min, followed by mechanical and enzymatic dissociation by pipetting

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every 5 min in protease solution (0.25% trypsin (Life Technologies, 15090–046), 1 mM
EDT , and 400 mg/mL Collagenase D (Sigma, 11088882001) in BS) and incubated at
28.5°C for 20–30 min or until full dissociation. Reaction was stopped by adding 6×stop
solution (6 mM CaCl2 and 30% fetal bovine serum (FBS) in PBS). Cells were pelleted
(376 × g, 5 min, 4°C) and resuspended in suspension media (1% FBS, 0.8 mM CaCl2,
50 U/mL penicillin, and 0.05 mg/mL streptomycin (Sigma-Aldrich, St. Louis, MO) in phenol
red-free Leibovitz’s L15 medium (Gibco, 11415064)) twice. Final volumes of 500 μL
resuspended cells were placed on ice and fluorescence-activated cell sorted (FACS) to
isolate live mCherry+ cells (zebrafish) that excluded the nuclear stain DAPI, and DAPIonly lizard cells into 0.04% BSA/ PBS at 4°C. Nuclei isolation was performed per
manufacturer’s instructions (10X Genomics, low cell input protocol CG000169), with
optimization for zebrafish sensory organs. FACS-sorted cells were pelleted for 15 min
(300 rcf at 4°C) and incubated with lysis buffer for 45-50 s on ice. Isolated nuclei were
washed by Wash buffer and Nuclei buffer and checked for nuclear integrity with a
fluorescence confocal microscope with DAPI staining before use in library construction.
snATAC and multiomic library construction, sequencing, and alignment
For the 14 dpf sample, a snATACseq library was constructed using Chromium Next GEM
Single Cell T C Library and Gel Bead Kit v1.1 per manufacturer’s instructions (10
Genomics, Cat# 1000176, protocol CG000209) with no modifications. For the 12 mpf
zebrafish inner ear and adult lizard samples, a multiomic library of snATACseq and
snRNAseq from the same barcoded nuclei was constructed using Chromium Next GEM
Single Cell Multiome T C + Gene E pression Reagent Bundle per manufacturer’s

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instructions (10x Genomics, Cat# 1000285, protocol CG000338) with no modifications.
Quality control of libraries was assessed with the 4200 TapeStation system and Qubit
dsDNA HS assay kit. Libraries were sequenced on Illumina HiSeq platform (discontinued
by manufacturer as of date of publication). For sequencing snATACseq libraries, both
Read1 and Read2 were extended to 60 cycles, and for sequencing snRNAseq libraries,
Read2 was extended to 102 cycles for longer coverage. Sequencing reads were aligned
to customized genomes. Zebrafish genome was built with GRCz11.fa, GRCz11.104.gtf
(ensembl), and JASPAR2020.pfm. Green anole lizard genome was built with
AnoCar2.0v2.fa, AnoCar2.0v2.109.gtf (ensembl), and JASPAR2020.pfm. Alignment and
peak calling were performed by Cell Ranger ATAC v2.0 for 14 dpf snATACseq reads and
Cell Ranger RC v2.0.0 for the multiomic library per manufacturer’s instructions to
generate peak-by-cell and gene-by-cell count matrices.
snATACseq data analysis
snATACseq data were processed using Seurat v5.0.1(Hao et al., 2024) and Signac
v1.12.0(Stuart et al., 2021) packages in Rstudio following standard workflow with
optimization. Cell Ranger ATAC output count matrices were used to create Seurat object
using “CreateChromatin ssay” and “CreateSeuratObject” functions. Quality control was
performed by setting the thresholds of accessible region counts (peak_region_fragments)
between 1000 and 50000, fraction of all fragments that fall within ATACseq peaks
(pct_reads_in_peaks) > 30, ratio of mononucleosome to nucleosome-free region
(nucleosome_signal) < 2, and enrichment at TSS (TSS.enrichment) > 2. Normalization
and linear dimension reduction by LSI were performed by “RunTFIDF”, and “RunSVD”

104
functions. To cluster cells, “FindNeighbors” (2:30 dimensions of LSI was used, with the
first dimension of LSI excluded due to its high correlation with sequencing depth),
“RunUM ” (2:30 dimensions of LSI was used), and “FindClusters” (resolution = 1)
functions. Predicted gene activity based on chromatin accessibility near each gene was
constructed with “Gene ctivity” and “Create ssayObject” functions. We then visualized
the gene activities for HC and SC markers and determined their corresponding clusters
in the dataset.
Multiomics data analysis
Multiomics data were processed by Seurat v5.0.1(Hao et al., 2024) and Signac
v1.12.0(Stuart et al., 2021) packages in Rstudio following standard workflow with
optimization. Cell Ranger ARC output count matrices in H5 format of individual libraries
were used to create Seurat objects by “CreateSeuratObject” and “CreateChromatin ssay”
functions. Quality control was performed by setting the thresholds of accessible region
counts (nCount_ATAC) between 100 and 50000, transcript counts (nCount_RNA)
between 100 and 10000. Normalization and linear dimension reduction by LSI were
performed by “RunTFIDF”, and “RunSVD” functions. RNA data (gene-by-cell count
matrices) were normalized and scaled, and the top 3000 most common genes were
selected by “SCTransform” function for dimension reduction by C (“Run C ” function).
To cluster cells with ATAC and RNA information in combination,
“FindMultiModalNeighbors” (2:50 dimensions of LSI and 1:50 dimensions of C were
used, with the first dimension of LSI excluded due to its high correlation with sequencing
depth), “RunUM ” (nn.name = “weighted.nn”), and “FindClusters” (resolution = 1,

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graph.name=”wsnn”, algorithm = 3) functions. Based on the gene e pression of HC and
SC markers, we identified their corresponding clusters in the multiomics datasets.
Visualization of chromatin accessibility
For zebrafish and lizard single-cell datasets, cell barcodes associated with either HCs or
SCs were exported into a csv file. Using subset-bam v1.1.0
(https://github.com/10XGenomics/subset-bam), we extracted sequencing reads related
to these cell barcodes from the possorted_bam.bam file of each sample. The resulting
bam files for 14 dpf and 12 mpf zebrafish and adult lizard HCs and SCs were processed
and normalized as pseudobulk ATACseq data into bigwig coverage files using
deepTools(Ramírez et al., 2014) “bamCoverage” function (--normalizeUsing CPM). We
used bam files associated with previously published mouse bulk ATACseq and histone
CUT&RUN data(Gnedeva et al., 2020; Tao et al., 2021; Wang et al., 2023) and similarly
created bigwig coverage files. Bigwig coverage tracks were then loaded into IGV genome
browser(Robinson et al., 2011) to visualize open chromatin peaks near genes of interest.
Identification of conserved enhancers
Genomic coordinates of each peak downstream of Atoh1/atoh1a were used to extract
DNA sequences of each peak using UCSC genome browser (https://genome.ucsc.edu/)
“get DN ” function. We then performed pair-wise alignment of mouse and lizard enhancer
sequences to zebrafish enhancers using NCBI nucleotide BLAST
(https://blast.ncbi.nlm.nih.gov/Blast.cgi), with the program selected to optimize for
somewhat similar sequences (blastn).

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Transcription factor motif prediction within single enhancers
We used CIS-BP Database(Weirauch et al., 2014) and predicted transcription factor
binding sites within each of the Atoh1/atoh1a enhancers of interest. The motif model of
“ WMs – LogOdds” was used with a default threshold of 8.
Differential gene expression
Previously published bulk RNAseq data from E17 HCs and P6 SCs(Tao et al., 2021) were
used to determine genes differentially enriched in developing HCs. Genes with a base
mean count over 50, RPKM greater than 1, log2 fold change greater than 1 (or smaller
than −2) and adjusted p-value less than 0.01, were considered as differentially expressed.
Using “multi-species comparisons” filter from Ensembl Biomart
(http://www.ensembl.org/biomart/martview), we further identified the orthologous genes
(both a and b versions) of these mouse genes in the zebrafish genome
Differentially accessible peaks near HC genes
Using the R package DiffBind(Ross-Innes et al., 2012) and previously published HC and
SC chromatin accessibility data(Tao et al., 2021), we determined a list of open chromatin
regions that were differentially accessible in P1 HCs compared to P6 SCs (log2FC > 0.25).
We used P6 SCs as it has been shown that most HC gene-associated chromatin is closed
by this stage(Tao et al., 2021). We also obtained a list of differentially accessible peaks
in P1 SCs compared to P1 HCs (log2FC<-0.25), when progenitor-associated peaks are
open in SCs but not HCs (Fig. 1E-F). Using the “distanceToNearest” function from the

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GenomicRanges R package(Lawrence et al., 2013), we associated and filtered these
differential peaks for those that are within 200kb up or downstream of HC genes identified
by comparing E17 HCs and P6 SCs. We then exported the genomic coordinates of these
peaks as bed files for downstream analysis. In zebrafish, we used “FindMarkers” function
in Signac(Stuart et al., 2021) to identify differentially accessible peaks between 14 dpf HC
and SC clusters with the threshold set at log2FC > 0.25 or < -0.25. We then used the
“ClosestFeature” function to associate and filter these differential peaks for those that are
within 200kb of the zebrafish homologs of mouse HC genes. We then exported the
genomic coordinates of these peaks as bed files for downstream analysis.
Visualization of differentially accessible peaks in HCs and SCs
Using deepTools(Ramírez et al., 2014) functions “computeMatri ” followed by
“plotHeatmap”, we plotted the chromatin accessibility from HCs and SCs at different
developmental timepoints around the genomic coordinates determined in the above step
(--regionsFileName “differential peaks in HCs”.bed “differential peaks in SCs”.bed –
scoreFileName “HC T C”.bigwig “SC T C from different timepoints”.bigwig). This
allowed us to visualize how the chromatin accessibility changed in mouse and zebrafish
SCs over time for the peaks that are differentially accessible in either HCs or SCs.
Log2FCs of Class 2 peak accessibility between P1 and P6 SCs in mouse or 14 dpf and
12 mpf SCs in fish were determined using “bigwigCompare” function. verage log2FC of
each peak was scored in a +/- 500 bp window centered at the peak center.
Homer Motif Enrichment Analysis

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To predict transcription factors that can bind to Class 2 peaks in mouse and zebrafish
SCs, we inputted the genomic coordinates of differentially accessible peaks in SCs
described above and ran Homer motif analysis(Heinz et al., 2010) using the function
“findMotifsGenome.pl” following standard instructions (size – given).
Quantification and statistical analyses
Quantification of HC numbers across different atoh1a alleles was performed by counting
hair bundles labeled by phalloidin staining in serial confocal sections of each sensory
organ using Fiji software(Schindelin et al., 2012). Unpaired t test or Tukey HSD multiple
pairwise test were used to compare the difference in HC numbers between wild types
and heterozygous or homozygous atoh1a enhancer deletion alleles, or between
atoh1amRuby/mRuby animals and homozygous atoh1a enhancer deletion alleles.
Quantification of atoh1a e pression level was done following manufacturer’s protocol
(ACD, SOP 45-006 Technical Note) to count number of signal dots within each of the
atoh1a-expressing cells on an RNAScope section. Number of atoh1a dots per cell was
tallied for all atoh1a-expressing cells across all sections imaged for that particular
genotype. Differences in number of atoh1a dots per cell between different genotypes
were compared using unpaired t test or Tukey HSD multiple pairwise test.

109
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Abstract (if available)

Abstract The loss of inner ear mechanosensory hair cells is permanent in mammals, which is a major contributing factor to deafness in humans. Conversely, non-mammalian vertebrates such as zebrafish can replenish hair cells throughout life from the transdifferentiation of neighboring supporting cells. The understanding of hair cell development and regeneration in zebrafish has largely been investigated through the lateral line neuromasts due to their surface localization for drug treatment and imaging access. However, through single-cell transcriptomic profiling of the zebrafish inner ear, we find a closer resemblance of zebrafish inner ear hair and supporting cells to their mammalian counterparts than those of neuromast origin. Additionally, we uncover a group of hair cell gene enhancers required for hair cell development and regeneration specifically in the inner ear. These enhancers remain accessible in adult zebrafish inner ear supporting cells, potentially enabling their life-long ability to transdifferentiate into hair cells upon damage. Conversely, a syntenic group of enhancers found in mouse cochlear supporting cells becomes permanently silenced postnatally, correlating to their loss of transdifferentiation potential. Cross-species enhancer testing also reveals that while both mouse and zebrafish versions of a sequence-conserved enhancer can continuously drive fluorescent reporter activities in juvenile zebrafish supporting cells, they become quickly silenced in postnatal mouse cochlea, regardless of their species of origin. This suggests that differences in upstream regulators in mouse versus zebrafish supporting cells are responsible for their contrasting abilities to maintain these regenerative enhancers and allow for hair cell regeneration.

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Asset Metadata

Creator Shi, Tuo (author)

Core Title Long-term maintenance of developmental enhancers enables hair cell regeneration in the zebrafish inner ear

School Keck School of Medicine

Degree Doctor of Philosophy

Degree Program Development,Stem Cells and Regenerative Medicine

Degree Conferral Date 2025-05

Publication Date 02/03/2025

Defense Date 02/28/2025

Publisher Los Angeles, California (original), University of Southern California (original), University of Southern California. Libraries (digital)

Tag enhancer,hair cell,inner ear,OAI-PMH Harvest,Regeneration,zebrafish

Format theses (aat)

Language English

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Advisor Bell, Oliver (committee chair), Crump, Gage (committee member), Gnedeva, Ksenia (committee member)

Creator Email tuoshi@usc.edu,tuoshi2.0@gmail.com

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