A systems-level approach reveals new gene regulatory modules in the developing ear

RESEARCH ARTICLE

A systems-level approach reveals new gene regulatory modules in

the developing ear

Jingchen Chen

*, Monica Tambalo

*, Meyer Barembaum

*, Ramya Ranganathan

*, Marcos Simões-Costa

Marianne E. Bronner

and Andrea Streit

‡

ABSTRACT

The inner ear is a complex vertebrate sense organ, yet it arises from

a simple epithelium, the otic placode. Specification towards otic

fate requires diverse signals and transcriptional inputs that act

sequentially and/or in parallel. Using the chick embryo, we uncover

novel genes in the gene regulatory network underlying otic

commitment and reveal dynamic changes in gene expression.

Functional analysis of selected transcription factors reveals the

genetic hierarchy underlying the transition from progenitor to

committed precursor, integrating known and novel molecular

players. Our results not only characterize the otic transcriptome in

unprecedented detail, but also identify new gene interactions

responsible for inner ear development and for the segregation of

the otic lineage from epibranchial progenitors. By recapitulating the

embryonic programme, the genes and genetic sub-circuits

discovered here might be useful for reprogramming naïve cells

towards otic identity to restore hearing loss.

KEY WORDS: Auditory system, Cell fate, Chick, Embryo, Hearing,

Placode, Transcription factor

INTRODUCTION

In vertebrates, the entire inner ear arises from the otic placode, a

simple epithelium next to the hindbrain, which invaginates to form

the otic vesicle. The vesicle undergoes extensive morphogenesis,

ultimately giving rise to the adult inner ear, an organ of exquisite

complexity comprising distinct sensory, non-sensory and neuronal

cell types of the auditory and vestibular apparatus. In humans,

congenital hearing defects are often due to mutations in

developmental genes. Thus, a mechanistic understanding of ear

development not only provides insight into the molecular control of

ear formation, but could also provide information relevant to the

aetiology of human sensory disorders.

Specification towards otic fate occurs early in development and

requires diverse signals and transcriptional inputs that act sequentially

and/or in parallel. This process is initiated when sensory precursors in

the pre-placodal region (PPR) become specified as otic-epibranchial

progenitors (OEPs) under the influence of fibroblast growth factor

(FGF) signalling (Fig. 1A; Ladher et al., 2000; Maroon et al., 2002;

Martin and Groves, 2006; Nechiporuk et al., 2007; Nikaido et al.,

2007; Phillips et al., 2001; Sun et al., 2007; Urness et al., 2010;

Wright and Mansour, 2003). The OEP state is characterized by

transcription factors like

Foxi1/3

(Khatri et al., 2014; Ohyama and

Groves, 2004; Solomon et al., 2003), Dlx genes (Brown et al., 2005;

Solomon and Fritz, 2002),

Pax2/8

(Christophorou et al., 2010; Freter

et al., 2012; Hans et al., 2004; Mackereth et al., 2005) and

Spalt4

(Barembaum and Bronner-Fraser, 2007, 2010; Schlosser, 2006).

After this step, Wnt and Notch pathways cooperate to promote otic

and repress epibranchial character (Fig. 1B; Freter et al., 2008;

Jayasena et al., 2008; Park and Saint-Jeannet, 2008; Shida et al.,

2015). However, the transcriptional networks that control each step

and the distinct differentiation programmes for otic and epibranchial

cells are very poorly understood.

Here, we examine the active transcriptome of the developing ear

from sensory progenitor to the overtly recognizable placode stage.

Major changes occur as cells transit from a progenitor state to

become OEPs, highlighting this as the most crucial step during otic

induction. Time course analysis reveals previously unknown steps

of otic commitment, defined by unique sets of transcription factors,

and functional analysis not only reveals new downstream targets of

known otic transcription factors, but also allows us to construct the

first otic gene regulatory network (GRN) and predict connections

therein. Its hierarchical organization reveals how, starting from a few

factors initiated by otic induction, information is propagated

through the network using positive feedback and feed-forward

loops to stabilize otic identity and generate diversity by segregating

otic and epibranchial fates.

RESULTS

New genes in otic placode development

The progressive commitment of ectodermal cells towards otic

identity occurs gradually, via a series of regulatory interactions that

are not well understood. In avian embryos, otic specification begins

around the 5-somite stage (ss) and by the 10ss, the otic ectoderm is

already committed to its fate and to form an otic vesicle (Adam

et al., 1998; Groves and Bronner-Fraser, 2000). To examine the

steps leading up to this cell fate decision, we chose three time points

for genome-wide transcriptome analysis, corresponding to the

stages when (1) cells become specified as OEPs (5-6ss; Fig. 1A), (2)

the placode acquires its characteristic thickened morphology (8-9ss)

and (3) cells become committed to an otic fate (11-12ss; Fig. 1B).

To identify otic-enriched genes, we compared the otic

transcriptome with that of whole embryos (3ss). Several hundred

genes are enriched more than 1.5-fold at each stage (5-6ss: 1202

transcripts; 8-9ss: 1079 transcripts; 11-12ss: 1315 transcripts;

Fig. 1C,D; Table S2). This analysis recovers many known otic

transcription factors (23/27; e.g.

Pax2

Gata3

Gbx2

Foxg1

Eya1

and

Soho1

; Fig. S1A; Fig. S2). Functional annotation of otic-

Received 20 December 2016; Accepted 24 February 2017

Department of Craniofacial Development and Stem Cell Biology, King

’

s College

London, London SE1 9RT, UK.

Division of Biology and Biological Engineering,

California Institute of Technology, Pasadena, CA 91125, USA.

*These authors contributed equally to this work

‡

Author for correspondence (andrea.streit@kcl.ac.uk)

A.S., 0000-0001-7664-7917

This is an Open Access article distributed under the terms of the Creative Commons Attribution

License (http://creativecommons.org/licenses/by/3.0), which permits unrestricted use,

distribution and reproduction in any medium provided that the original work is properly attributed.

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Development (2017) 144, 1531-1543 doi:10.1242/dev.148494

DEVELOPMENT

enriched transcripts (Fig. 1F-H) reveals progressive commitment to

otic identity:

‘

epithelium development

’

is the most represented gene

ontology (GO) term in OEPs (5-6ss; Fig. 1F), but this rapidly

changes at 8-9ss and 11-12ss, when

‘

inner ear development

’

becomes the prominent term (Fig. 1G,H). Interrogating disease

association databases reveals that transcripts enriched at the

commitment stage are associated with hearing loss (Fig. 1E).

In total, this analysis identified 135 potential transcriptional

regulators, of which 112 are novel with respect to the otic placode.

To verify that they do represent otic-enriched transcripts, we

assessed their expression using complementary methods. Twenty-

three factors are indeed expressed in the otic placode according to

the gene expression database GEISHA (http://geisha.arizona.edu/

geisha/; Fig. S2). We found ten additional factors enriched in

placode tissue as assessed by qPCR from dissected otic-

epibranchial domains (Fig. S1B,C). Likewise, of 52 transcripts

tested by NanoString 46 are present in the otic placode at 11-12ss

with mean count >300 (Fig. S1D). For further validation, we

performed

in situ

hybridization of 39 additional factors (Fig. 2;

Figs S2, S3). This confirmed that the majority (34/39) is present,

although not necessarily restricted to the otic placode. In summary,

the transcriptome analysis identifies many genes not previously

associated with ear development.

Dynamic changes of gene expression in the developing otic

placode

To capture changes in gene expression as otic cells mature, we used

complementary approaches:

in situ

hybridization, transcriptome

changes over time and hierarchical clustering. This allowed us to

define synexpression groups and uncover distinct transcriptional

states during otic placode maturation.

Analysis of transcription factor expression by

in situ

hybridization

Lmx1a

Sox13

and

Zbtb16

are expressed in OEPs at 5-6ss and their

expression persists in the otic placode until at least 12ss (Fig. 2A-F;

Fig. S2). In contrast,

Zfhx3

Rere

and

Tcf4

(

Tcf7l2

) only become

Fig. 1. Otic-enriched transcripts.

(A,B) Diagrams showing the location of OEPs at 5ss (A,A

′

, graded pink-blue) and the otic and epibranchial placodes at 11-

12ss (B,B

′

; otic: purple; epibranchial: blue). OEPs are induced by mesoderm-derived FGFs (green in A

′

). Later, FGFs activate Wnt ligands in the neural tube,

which cooperate with Notch to promote otic identity (B

′

), while FGFs and BMPs from the endoderm promote epibranchial fate. (C,D) RNAseq was performed

on dissected otic placodes from 5-6ss, 8-9ss and 11-12ss and otic-enriched transcripts enriched were identified by comparison to the whole embryo (3ss); see

also Tables S1 and S2. (C) Genes enriched in the otic placode at 5-6ss (blue; fold-change >1.5). (D) Venn diagram showing the number of otic-enriched genes at

5-6ss, 8-9ss and 11-12ss and their intersection. (E) Disease-association of otic-enriched genes. (F-H) Biological processes and signalling pathways over-

represented in otic-enriched genes at each stage showing at most the five top over-represented terms for which

<0.01 (Fisher

’

s Exact test) for each category.

Epi, epibranchial domain; OEPD, otic-epibranchial progenitor domain.

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DEVELOPMENT

prominent in the otic territory from 10ss onwards (Fig. 2G-L; other

factors:

Arid3

Atn1

Bach2

Klf8

Prep2

Tead3

Znf384

; Fig. S3I,J,

O-AD; Fig. S2), whereas

Nr2f2

Vgll2

and

Klf7

are confined to the

epibranchial region (Fig. 2O-R; Fig. S3K,L).

Prdm1

and

Tfap2e

transcripts change rapidly: they are broadly expressed at 5ss but then

become restricted to the epibranchial territory after 9ss with

Tfap2e

also present in neural crest cells (Fig. 2M,N; Fig. S3M,N). In

addition, several transcriptional regulators surround the otic placode

at 11-13ss (Fig. S3AE-AJ), whereas others are widely expressed in

the ectoderm including the otic territory (Fig. S3A-H; Fig. S2) or

also present in neural crest cells (Fig. S3AJ-AN). We summarize the

temporal and spatial expression of 95 known and new transcripts in

Fig. S2.

Dramatic transcriptome changes accompany OEP specification

To highlight the main changes that occur at key steps of otic

development, we performed pairwise comparisons of the otic

transcriptome at consecutive stages: we compared (1) the PPR at

0ss with OEPs at 5-6ss; (2) OEPs at 5-6ss with the otic placode at

8-9ss; and finally (3) otic placodes at 8-9ss and at 11-12ss

(Fig. 3A,D,H; Table S3). The most dramatic change occurs as cells

transit from a sensory progenitor state in the PPR to specified

OEPs, with 1569 transcripts being upregulated and 1733

downregulated at 5-6ss (Fig. 3A). Thereafter, changes occur

more gradually (Fig. 3D,H). Transcripts associated with GO terms

related to the acquisition of anterior character such as

‘

eye,

pituitary gland, nose, forebrain and diencephalon

’

(

Pax6

Otx1/2

Mafa

Hesx1

Pax3

Dlx5

; Table S3) are significantly under-

represented at the OEP stage (Fig. 3B,B

′

), consistent with the

earlier suggestion that repression of anterior fate is an important

step for otic induction (Bailey et al., 2006; Lleras-Forero and

Streit, 2012).

Hierarchical clustering identifies distinct TF synexpression groups

during otic commitment

Hierarchical clustering of all transcription factors that are either

enriched in the otic placode compared with the whole embryo

(Fig. 1) or differentially expressed over time (Fig. 3) reveals five

major clusters denominated transcription factor cluster 1-5 (TFC1-

5; Fig. 4A; Table S4). These clusters show distinct temporal profiles

(Fig. 4A-F) and generally confirm our

in situ

hybridization data. For

example, transcripts in TFC1 and TFC2 increase over time and these

clusters include

Lmx1a

Zbtb16

Rere

and

Tcf4

(Fig. 2; Fig. S1B,C).

Combining these three approaches allows us to define distinct

regulatory states as OEPs become committed to an otic fate. A

number of PPR genes are reduced during OEP induction (

Dlx5/6

Irx1

Foxi3

Gbx2

; Fig. 4B; Fig. S2; see also Khatri et al., 2014),

whereas a small group of transcripts [

Irx5

Lmx1a

cMyb

(Betancur

et al., 2011),

Prdm1

Sall4

(Barembaum and Bronner-Fraser, 2007),

Sox13

Zbtb16

and

Znf385c

] becomes upregulated together with

Pax2

and

Etv4

(Figs 2, 4; Figs S2, S3).

Sox10

expression is initiated

around 10ss together with

Foxg1

and

Dlx3

(Betancur et al., 2011;

Khudyakov and Bronner-Fraser, 2009; Yang et al., 2013), and

Prdm1

becomes restricted to the epibranchial territory (Fig. 2),

where it is co-expressed with

Pax2

Foxi2

and

Sox3

(Abu-Elmagd

et al., 2001; Freter et al., 2008; Groves and Bronner-Fraser, 2000).

Thus, already at the 10ss stage, otic and epibranchial progenitors

begin to segregate and become molecularly distinct. As cells

become committed to otic identity many new transcription factors

start to be expressed (Figs 2, 4; Figs S2, S3) and the otic and

epibranchial fates continue to diverge. In summary, our time course

analysis reveals distinct regulatory states as cells acquire otic

identity. Substantial transcriptome rearrangements occur within a

brief period of only 6-8 h (from 1ss to 5ss) during the first step of

otic induction: anterior character is inhibited and cells are specified

Fig. 2. Expression of transcription factors in

theoticplacode.

(A-R)

Lmx1a

(A,B),

Sox13

(C,D)

and

Zbtb16

(E,F) are expressed in OEPs and in

the otic placode (OP).

Rere

(G,H),

Tcf4

(I,J) and

Zfhx3

(K,L) expression starts at placode

stages, whereas

Prdm1

is expressed in OEPs (M)

but later restricted to the epibranchial territory (Epi)

(N). At 12-13ss,

Nr2f2

(O,P) and

Vgll2

(Q,R) are absent from the otic placode, but present

in epibranchial cells and the ventral ectoderm

(

Vgll2

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DEVELOPMENT

as OEPs. Our analysis defines new factors that characterize a

transcriptional state characteristic for OEPs at 4-5ss. As

development proceeds, known ear-specific transcripts become

more prominent, as do genes associated with hearing impairment,

suggesting that our data might harbour new candidate deafness

genes.

Pathway analysis suggests potential novel regulators of otic

placode formation

Signals from the surrounding tissues induce and pattern the otic

placode. Although the role of FGF, Notch and Wnt pathways is well

established (Abello et al., 2010; Freter et al., 2008; Ladher et al.,

2000; Ohyama et al., 2006; Park and Saint-Jeannet, 2008; Phillips

et al., 2004; Urness et al., 2010), it is likely that other signals are also

involved. To explore this possibility, we used hierarchical clustering

of all otic-enriched genes (from Fig. 1) together with differentially

expressed genes from stage-wise comparisons (from Fig. 3) to

generate six major clusters (denominated C1-C6; Fig. S4).

Following pathway enrichment analysis for each cluster

(Fig. S4A), we extracted the components of each significantly

enriched or depleted pathway (Fig. 3C,F,G,J; Fig. S4B-D).

First, we evaluated pathways known to mediate otic development.

As expected, Notch signalling components are present throughout

placode formation and are over-represented in OEPs and in the

placode (Fig. 3C; Fig. S4A,D) with

Lfng

expression increasing

sharply as the placode forms and

Deltex2

and -

rising gradually.

Wnt signalling components are highly enriched in cluster C2

(Fig. S4A,C) with the Wnt receptors

Fzd1-3

and mediators

Lef1

and

Tcf7l2

increasing steadily. In contrast, the Wnt antagonist

Sfrp2

drops sharply at 5-6ss. Components of the non-canonical Wnt

pathway, such as

Wnt5a

, rise gradually together with

Rac1

and

Jun

suggesting a role in placode assembly and morphogenesis. These

findings are consistent with known changes in signalling events

during otic commitment and therefore confirm the usefulness of this

Fig. 3. Temporal changes in otic gene expression.

Pairwise comparison of the otic transcriptome at consecutive developmental stages: 5-6ss compared with

0ss (PPR; A-C), 8-9ss compared with 5-6ss (D-G) and 11-12ss compared with 8-9ss (H-J); see also Table S3. (A,D,H) Differentially expressed genes with a fold

change >1.5; blue indicates upregulated transcripts; orange indicates downregulated transcripts. (B,E,I) Gene ontology analysis of up- and downregulated genes

showing the five top over-represented biological processes or signalling pathway (

<0.01; Fisher

’

s Exact test). There is no significant association for the

downregulated genes shown in H. (B

′

) At 5-6ss terms related to anterior structures are significantly under-represented relative to 0ss. (C,F,G,J) Changes of

transcripts associated with signalling pathways over the entire time course. Asterisk indicates that the gene expression level is indicated by the

-axis on the right.

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DEVELOPMENT

approach to predict the potential new pathways regulating otic

development.

Next, we investigated whether new pathways emerge from this

analysis. As OEPs become specified, components of the steroid

biosynthesis pathway are markedly upregulated (Fig. 3E,G),

whereas TGF

signalling components drop sharply (Fig. 3I,J).

Spliceosome components (Fig. S4A,B, cluster C2) peak at 5-6ss

and 8-9ss. Consistent with this, spliceosomal defects are known to

cause craniofacial disorders, some of which are associated with

hearing loss (Lehalle et al., 2015). As a morphological placode

forms, focal adhesion-related components become increasingly

enriched, suggesting a role in placode assembly (Fig. 3E,F). These

observations point to signals and pathways not previously

associated with ear formation to explore in the future.

Fig. 4. Clusters of otic transcription

factors.

(A) Otic transcription factors from

the enrichment and time course analysis

cluster into five clusters (TFC1-5) based

on the row z-score of fold change relative

to the PPR at 0ss. (B-F) Expression level

of the top 50% transcription factors in

each cluster. Line in the top right of each

cluster represents the overall expression

profiles across the three time points. See

also Table S4.

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DEVELOPMENT

Regulatory relationships reveal distinct transcriptional

modules during otic commitment

Our time course analysis of gene expression predicts a

transcriptional hierarchy during otic induction. To begin to test

this hierarchy, we selected three transcription factors, Etv4, Pax2

and Lmx1a, for perturbation experiments for the following reasons.

The transition from sensory progenitors to OEPs is mediated by the

FGF pathway (Ladher et al., 2000; Maroon et al., 2002; Martin and

Groves, 2006; Nechiporuk et al., 2007; Nikaido et al., 2007; Phillips

et al., 2001; Sun et al., 2007; Urness et al., 2010; Wright and

Mansour, 2003). Accordingly, the FGF mediator

Etv4

is expressed

in OEPs (Lunn et al., 2007) and upregulated 7.5-fold from 1ss to

5-6ss (Table S3). Only ten transcription factors are strongly initiated

at OEP stages (6- to 235-fold), with

Pax2

being the top factor

(235-fold; Table S3), but

Lmx1a

is one of the few genes (6.3-fold;

Table S3) exclusively expressed in otic, but not epibranchial cells

(Fig. 2A,B). To explore the gene network downstream of these

factors, we knocked down their expression by electroporation of

antisense morpholino oligonucleotides at 1-2ss (MOs; Barembaum

and Bronner-Fraser, 2010; Betancur et al., 2011; Christophorou

et al., 2010). Experiments were assessed by NanoString nCounter

at 10-12ss using a total of 216 probes including 70 otic genes

(mostly transcription factors), markers for placode progenitors,

other placodes, the neural plate, neural crest cells and non-neural

ectoderm. This analysis provides a large-scale view of

transcriptional changes in a single experiment enriching

previously published data, which generally assessed a few genes

at a time. In addition, selected transcripts were also assessed by

RT-qPCR and/or

in situ

hybridization (Fig. 7; Fig. S6; Table S5).

A gene was considered to be activated or repressed when its

expression was reduced or enhanced after knockdown, respectively

[NanoString: normalized mean count >300, ±1.2-fold change,

adjusted

-value (

-adj)<0.1; RT-qPCR: ±1.5-fold change,

<0.05;

in situ

hybridization: absence or reduction of signal in

electroporated cells]. These data allow us to add functional links

between Etv4, Pax2, Lmx1a and other transcription factors in the

otic GRN (Fig. 6), although cis-regulatory analysis will be required

to distinguish between direct and indirect interactions. Although our

experiments do not determine precisely when these interactions take

place, we can infer this from our expression data, which show the

onset of target genes.

Etv4 and Pax2 control the onset of OEP factors

Etv4 knockdown leads to a reduction of

Pax2

expression (Fig. 5B-B

′′

;

Fig. S6A,B) confirming a requirement of FGF activity for

Pax2

expression. In addition, Etv4 activates other early OEP transcripts

(

Irx5

Prdm1

Zbtb16

Sall4

Sox8

; Fig. 5A-F; Barembaum and

Bronner-Fraser, 2007; Yang et al., 2013), and is also required for

genes present at placode stages (

Lef1

Lmx1b

Sox10

Tcf4

; Table S5).

In contrast, Etv4 represses some PPR genes (

Six1

Eya2

), the OEP

factor

Znf385c

and late otic placode transcripts (

Tead3

Arid3

Sall1

;

Fig. S6A,B; Table S5).

Many Etv4 targets are also regulated by Pax2 (OEP transcripts:

Irx5

Prdm1

Zbtb16

Sall4

; placode genes:

Lef1

Lmx1b

Sox10

Tcf4

; Fig. 5G,I-J

′′

; Fig. S6A,C; Table S5). In addition, Pax2

activates the Etv4-independent OEP genes

Lmx1a

(Fig. 5H-H

′′

) and

Sox13

, the placode transcripts

Eya1

Meis1

Zfhx3

and

Znf521

and

maintains the PPR factor

Six1

, while repressing the posterior PPR

genes

Foxi3

and

Gbx2

, which are normally cleared from the placode

as it matures, the trigeminal marker

Pax3

(Wakamatsu, 2011), late

onset otic genes (

Foxg1

Dlx3

) and

Kfl7

, which is later expressed in

the epibranchial region (Fig. S6A,C). Pax2 is also required for the

epibranchial-specific factor

Vgll2

(Fig. 5K-K

′′

). Electroporation of

control morpholinos does not affect

otic gene expression (Fig. 5N-P

′′

Thus,

many inner ear transcription factors depend on Etv4 and/or Pax2

activity placing these factors at the top of the otic hierarchy.

Lmx1a and Pax2: a positive feedback loop that maintains OEP factors

and represses alternative fates?

Exploring Lmx1a function, we find that

Pax2

depends on Lmx1a

input:

Pax2

expression is reduced when Lmx1a is knocked down

(Fig. 5L-M

′

; Fig. S6A). Thus, they mutually regulate each other in

a positive feedback loop and have common targets: both are

required for

Sox13

and

Zbtb16

expression (Fig. 6A). In addition,

Lmx1a is necessary for

Foxg1

and

Gbx2

expression. Like Pax2,

Lmx1a also suppresses several transcripts (Fig. 5L; Fig. 6A),

among them the PPR genes

Six1

and

Foxi3

and the lens/olfactory

factor

Pax6

. Thus, together both transc

ription factors appear to

promote OEP, but might also participate in the repression of

alternative fates.

DISCUSSION

Commitment to otic fate is initiated by the specification of OEPs

from the posterior part of the pre-placodal region, followed by the

acquisition of columnar placode morphology and, finally, placode

invagination to form the otic vesicle. During this process, the otic

territory is exposed to signals from surrounding tissues, and as

morphogenetic events shape the embryo, its extrinsic environment

changes constantly. As a result, ectodermal cells first initiate a

transcriptional programme unique to otic progenitors and then form a

patterned otic vesicle. Here, we have greatly expanded this

transcriptional repertoire by identifying more than 100 factors that

might be crucial for placode development, and are also new candidate

genes for hearing impairment. Exploring the dynamic changes in

gene expression over time together with perturbation analysis of

selected transcription factors and integrating data from the literature

(Fig. S5) allows us to propose the first GRN for otic commitment.

A transcriptional mechanism for OEP specification

To establish the first GRN that describes how sensory progenitor

cells are committed to the inner ear lineage, we used a strategy that

measures changes of all otic transcription factors after experimental

perturbation using partial knockdown of three transcription

factors (Fig. 7). This GRN has the deep structure characteristic of

embryonic networks (Davidson, 2010), revealing the hierarchical

organization of the process that drives otic commitment. Within this

network, distinct transcriptional modules can be identified that

gradually establish otic identity.

The posterior PPR module

For the otic placode to develop, ectodermal cells must first go

through a PPR state (Martin and Groves, 2006), which is identified

by Six and Eya family members and by

Irx1

and

Dlx5/6

. Six1 is an

important PPR determinant (Ahrens and Schlosser, 2005;

Brugmann et al., 2004; Christophorou et al., 2009) and Irx1, Dlx5

and Foxi3 are known to regulate its expression (Glavic et al., 2004;

Khatri et al., 2014; Pieper et al., 2012; Sato et al., 2010; Woda et al.,

2003). In the posterior PPR,

Six1

Eya2

Gbx2

and

Foxi3

are crucial

for otic placode formation (Brugmann et al., 2004; Christophorou

et al., 2009; Solomon et al., 2003; Steventon et al., 2012). Gbx2

restricts the expression of

Otx2

to the anterior PPR (Steventon et al.,

2012), whereas Foxi1/3 and the Six1/Eya2 complex regulate each

other in a positive feedback loop (Khatri et al., 2014), perhaps to

maintain posterior PPR identity. Together, all four proteins provide

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DEVELOPMENT

crucial input for the OEP transcription factor

Pax2

(Christophorou

et al., 2010; Hans et al., 2004, 2007; Solomon et al., 2003, 2012):

loss of Foxi1 function in fish and repression of Gbx2 and Six1

targets genes in

Xenopus

and chick, respectively, lead to the absence

Pax2

(Fig. 6B; Fig. 7A). However, none of these transcription

factors is sufficient to induce Pax2 in non-otic cells, suggesting that

additional input is required.

The OEP module

It is well established that FGF signalling induces OEPs from the

posterior PPR (Ladher et al., 2000, 2005; Maroon et al., 2002;

Martin and Groves, 2006; Nechiporuk et al., 2007; Nikaido et al.,

2007; Phillips et al., 2001; Sun et al., 2007; Urness et al., 2010) and

we show that the FGF target Etv4 is crucial for this process.

Etv4

upregulated as posterior PPR cells transit to OEPs and is required for

Fig. 5. Regulation of otic transcription factors by Pax2, Etv4 and Lmx1a.

Target-specific morpholinos were electroporated at 0-1ss and changes in gene

expression were assessed by NanoString with three biological replicates, each of which containing five pieces of otic placode (A,G; see also Table S5),

in situ

hybridization (B-B

′′

,C-C

′′

,D-F,H

′

-K

′′

,M,M

′

) or RT-PCR with two biological replicates each containing five pieces of otic placode (L). (A) Etv4 knockdown analysed

by NanoString. Green indicates downregulated genes, red indicates upregulated genes. Open triangles represent data points that have a value beyond the axis

limit. (B-F)

In situ

hybridization after Etv4 knockdown for the genes indicated in each panel. A reduction of

Pax2

(8/12; B

′

Zbtb16

(6/6; C

′

);

Prdm1

(7/11; D),

Tcf4

(8/10; E) and

Vgll2

(4/5; F) is observed. Asterisks indicate the electroporated side. B and C show morpholino fluorescence of the embryos shown in B

′

and C

′

respectively; B

′′

and C

′′

show sections through the embryos shown in B

′

and C

′

, respectively, at the level marked by the horizontal lines. (D-F) Sections of embryos

electroporated with Etv4 morpholino. (G-K

′′

) Pax2 knockdown analysed by NanoString (G). Green indicates downregulated genes, red indicates upregulated

genes. Open triangles represent data points that have a value beyond the axis limit. (H-K

′′

)

In situ

hybridization after Pax2 knockdown for the genes indicated in

each panel. Asterisks indicate the electroporated side.

Lmx1a

(4/4; H

′

Zbtb16

(4/4; I

′

Prdm1

(3/4; J

′

) and

Vgll2

(4/4; K

′

) are reduced. H-K show morpholino

fluorescence of the embryos shown in H

′

-K

′

′′

-K

′′

show sections through the embryos shown in H

′

-K

′

, respectively, at the level marked by the horizontal lines.

(L) Lmx1a knockdown analysed by RT-qPCR. The results are presented as fold change ±s.d. and two-tailed Student

’

-test was used to calculate

-value.

(M) Morpholino fluorescence of the embryo shown in M

′

(3/4). (N-P

′′

In situ

hybridization after control morpholino electroporation for the genes indicated in the

panels; N

′

-P

′

sho

w sections of the embryos in N-P, respectively, at the level marked by the horizontal lines.

In situ

hybridization for each gene was performed on

four embryos electroporated with control morpholino.

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DEVELOPMENT

Pax2

expression. Thus, a typical

‘

AND

’

gate controls

Pax2

OEPs: its expression requires dual input from the posterior PPR

factors Six1, Foxi3 and Gbx2 and from the FGF mediator Etv4

(Fig. 6B; Fig. 7A).

Our data suggest that Etv4 and Pax2 work in concert to promote

otic identity and place them at the top of the transcriptional

hierarchy for otic commitment (Fig. 7A). Together, they rapidly

activate an OEP module consisting of

Sox8

Lmx1a

Zbtb16

Sox13

Prdm1

Sall4

Znf385c

and

Irx5

. The OEP module also contains the

Pax2- and Etv4-independent factor

cMyb

(Betancur et al., 2011);

however, its upstream regulators are currently unknown. As Etv4

and Pax2 share many targets, two scenarios could explain their

mode of action. Etv4 might only regulate

Pax2

, which in turn

controls other targets in a linear hierarchy (Fig. 6C; for simplicity,

we depict this possibility in Fig. 7). However, it is equally possible

that Etv4 and Pax2 act in a feed-forward loop, in which Etv4 is

required for

Pax2

and both together control the expression of

downstream targets (Fig. 6D) as is indeed the case for

Sall4

(Barembaum and Bronner-Fraser, 2007).

Although these and previous data implicate Pax2 as a key factor

for otic specification and proliferation in chick (Christophorou et al.,

2010; Freter et al., 2012), the mouse otic placode still forms in the

absence of Pax2 (Burton et al., 2004; Favor et al., 1996; Torres et al.,

1996) or Pax2 and Pax8 function (Bouchard et al., 2010). The fact

that sauropsids have lost the

Pax8

gene (Christophorou et al., 2010;

Freter et al., 2012) could explain the more prominent role of Pax2 in

the otic GRN.

The newly identified OEP factors might play an important role in

‘

locking

’

cells in an OEP transcriptional state, where they are

competent to respond to signals committing them towards otic and

epibranchial fate. It has previously been suggested that continued

FGF activity inhibits otic placode formation, while promoting

epibranchial cells (Freter et al., 2008). Indeed, when chick OEPs are

cultured in isolation they maintain otic identity from 5-6ss onwards

and even generate neurons in the absence of additional signalling

(Adamska et al., 2001; Groves and Bronner-Fraser, 2000). These

findings suggest that in an OEP state ear precursors are FGF

independent and we propose that the OEP module could be

instrumental to maintain cell identity. Like Pax2 and Lmx1a, other

OEP transcription factors might act in positive feedback loops to

maintain OEP gene expression and repress alternative fates.

Two distinct steps segregate otic and epibranchial

progenitors

Our temporal analysis reveals two steps during the segregation of

otic and epibranchial fates (Fig. 7A,B). Under continued FGF

signalling, OEPs differentiate into epibranchial cells (Freter et al.,

2008; Sun et al., 2007). This is consistent with our finding that Etv4

is required for epibranchial transcripts, as is Pax2. Dependent on the

cellular context, Pax2 is likely to use different partners to impart cell

fate (Kamachi and Kondoh, 2013; Stolt and Wegner, 2010): SoxE

group members are prominent in otic cells, but they are absent from

epibranchial placodes, where SoxB group members could represent

major Pax2 partners (Ishii et al., 2001; Wood and Episkopou, 1999).

In the otic lineage, expression of a small set of transcription factors

(

Sox10

Foxg1

and

Dlx3

; Fig. 7B) is initiated downstream of Pax2

and the OEP module; for example, the Sox10 enhancer is directly

controlled by Pax2, Sox8 and cMyb (Betancur et al., 2011), whereas

epibranchial cells begin to initiate a different transcriptional

programme.

In a second step, a large number of otic transcripts begin to be

expressed, among them several that depend on canonical Wnt

Fig. 6. Regulatory modules during OEP specification.

All diagrams summarize data from the literature and from this study (for details see text). (A) Lmx1a and

Pax2 mutually activate each other, control common targets and appear to repress alternative fates (see text for details). (B)

Pax2

is controlled by the posterior

PPR factors Six1, Eya2, Foxi3 and Gbx2, as well as by the FGF mediator Etv4. (C,D) Etv4 and Pax2 could act in a linear pathway (C) or in a feed-forward

loop (D) to control other OEP genes (see text for details).

Irx5

is regulated by both Etv4 and Pax2; however, because

Pax2

is also regulated by Etv4 for simplicity

the network in Fig. 7 assumes that Etv4 regulates

Irx5

via Pax2: Etv4

→

Pax2

→

Irx5.

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DEVELOPMENT

Fig. 7. Gene regulatory network incorporating new functional data.

(A-C) Signalling inputs, gene expression changes and regulatory relationships at

the three different stages. Regulator links are based on data from the literature (Fig. S5) and our perturbation experiments (Fig. 5; Fig. S6A-C). Direct interactions

confirmed from literature are indicated with blue diamonds and bold lines. As enhancers for most genes are currently unknown, the network assumes the

simplest interactions depending on perturbation data (see also Fig. 6). Epi, epibranchial domain; OEPD, otic-epibranchial progenitor domain; OEPs, otic

epibranchial progenitors; OP, otic placode; pEpi, pre-epibranchial domain.

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DEVELOPMENT

signalling from the neural tube (Freter et al., 2008; Ohyama et al.,

2006) (Fig. 7C). Interestingly, FGF signalling, either directly or

indirectly, promotes the Wnt mediators

Lef1

and

Tcf4

, suggesting

that these upstream events might prime OEP cells for Wnt

signalling. In contrast, under the influence of bone morphogenetic

protein (BMP) signalling (Begbie et al., 1999) epibranchial

progenitors continue to diverge from otic cells and the two fates

become firmly established. In summary, the temporal resolution of

our analysis highlights the complexity of cell fate decisions and

reveals new transcriptional states as sensory progenitor acquire ear

and epibranchial identity.

The OEP module: a molecular circuit re-deployed in other

organs?

Our analysis has defined a new transcriptional circuit, operating

downstream of Pax2 and FGF/Etv4, which we propose is important

for the specification of otic-epibranchial precursors. In addition to

the ear, members of the OEP module are also co-expressed in the

developing kidney and limb, including

Zbtb16

(Cook et al., 1995),

Lmx1

(Fernandez-Teran et al., 1997),

Prdm1

(Ha and Riddle, 2003)

as well as

Sox8

(Chimal-Monroy et al., 2003) and

Sox13

(Kido

et al., 1998; Wang et al., 2006)

Members of the Six and Eya

families lie upstream of the OEP module, and together with Pax

proteins are part of the fly retinal determination network (Salzer and

Kumar, 2009), which in vertebrates also regulates the formation of

other organs including the ear and kidney. Interestingly, the

homologues of Zbtb16, Lmx1a and Prdm1 participate in fly eye

formation although their relationship to the retinal determination

network is unknown (Ko et al., 2006; Maeng et al., 2012; Wang

et al., 2006). It is therefore tempting to speculate that the OEP

module is part of an ancient sub-circuit that is re-deployed as a unit

to govern cell fate decisions in different species and organs. The

OEP module could provide an initial link between the top upstream

regulators (Six/Eya and Pax proteins) that propagates information to

the next level of the network.

Uncovering new candidate deafness genes

Although much progress has been made recently to identify the

genetic causes for hearing impairment in humans, many causative

genes remain to be uncovered. In particular, mutations in

developmental genes are associated with human deafness: for

example Six1 or Eya1 mutations are known to cause Branchio-Oto-

Renal Syndrome, an autosomal dominant disorder (Ruf et al.,

2004). Functional annotation of our transcriptome data reveals a

significant association with hearing loss, in particular for transcripts

enriched in the committed otic placode (11-12ss; Fig. 1E). Indeed,

the number of otic placode-enriched genes that fall into known

human deafness loci is significantly higher than expected for a set of

random genes (data not shown). These findings suggest that our data

might harbour a number of novel candidates for congenital

malformations of the ear and for hearing impairment.

In summary, integrating data from the literature (Fig. S5) and our

new analysis has allowed us to establish the first gene regulatory

network (Fig. 7) that models the early stages of ear development. By

identifying key otic genes and the linkages between transcription

factors at different levels of the network, we define new distinct

regulatory states as cells acquire otic identity as well as the

regulatory loops that stabilize cell fate decisions. In the future it will

be important to identify the cis-regulatory elements controlling otic

gene expression to determine direct and indirect interactions. The

information gleaned from our analysis of otic transcriptional

programmes will be crucial for future experiments aiming to

reprogramme cells after loss of hearing and/or balance for the

purpose of repair and regeneration.

MATERIALS AND METHODS

Tissue dissection and embryo experiments

Fertilized hens

’

eggs (Winter Farm, Herts) were incubated in a humidified

incubator at 38°C to reach the appropriate stage. For dissection, embryos

were isolated from the egg and pinned out ventral side up on a resin plate in

Tyrode

’

s saline. The endoderm and mesoderm layers were removed using a

G-31 syringe needle in the presence of a small volume of 10 mg/ml dispase

before dissecting the future otic ectoderm based on the region expressing

Pax2

(Streit, 2002), rostral to the first somite and adjacent to future hindbrain

rhombomeres 4-6.

For

in situ

hybridization, embryos were isolated in PBS, fixed in 4%

paraformaldehyde in PBS and processed as described previously (Streit and

Stern, 2001). Digoxigenin-labelled antisense RNA probes were synthesized

with T7, T3 or SP6 RNA polymerase (Roche) as appropriate from expressed

sequence tags, cloned fragments or previously published plasmids (Table S6).

For morpholino knockdown experiments, electroporation at 0-1ss was

performed either unilaterally with control and experimental morpholino, or

in the same embryo with control and experimental morpholino on different

sides.

Pax2

and

Etv4

morpholinos were validated previously (Betancur

et al., 2011; Christophorou et al., 2010); as controls, standard control

morpholinos (5

′

-CCTCTTACCTCAGTTACAATTTATA-3

′

) were used.

For

Lmx1a

, a splicing-blocking morpholino (5

′

-ACCCCCAGTGTCCCC-

ATACCTTCCT-3

′

) targeting the exon 4-intron 4 junction was used and

deletion of exon 4 was confirmed by RT-PCR followed by sequencing the

PCR product. Changes in gene expression were assessed by whole-mount

in situ

hybridization or from five to ten dissected otic placodes by RT-qPCR

or NanoString. For dissection, the non-electroporated or control morpholino

electroporated side and the first somite is used as a guide.

RNA isolation, library construction and sequencing

About 100 placodes were collected from each stage for RNAseq library

preparation. Immediately after dissection, tissues were lysed in lysis buffer

(Ambion) and RNA was isolated using the RNAqueous-Micro Kit

(Ambion). Libraries were prepared with TrueSeq RNA Sample

Preparation V2 kit (Illumina) and sequenced with Illumina HiSeq 2000

(Illumina) to 1×50 cycles or HiSeq 2500 to 2×100 cycles.

Sequence alignment

Reads ware aligned with TopHat2 (v2.0.7) to Ensembl chick genome

Galgal4.71 (Kim et al., 2013). Transcripts for individual samples were

assembled with Cufflinks (v2.1.1) (Trapnell et al., 2010), with combined

Ensembl gene annotation (Galgal4.71.gtf) and Refseq acquired from UCSC

table browser as a guide. All assemblies were merged to obtain a merged

annotation file, which was passed to Cuffdiff (v2.1.1) to obtain normalized

RPKM value for each gene, and to easyRNAseq (v2.1.0) to retrieve the

count table for all genes (Delhomme et al., 2012). All sequencing data were

deposited in Gene Expression Omnibus (GEO) under accession number

GSE69185.

Identification and analysis of differentially expressed genes

DEseq (v1.16.0) was used to identify enriched otic genes relative to the

whole embryo (Anders and Huber, 2010) based on the count table generated

above. A gene was considered to be expressed in the otic region when the

normalized RPKM was >4, and the number of normalized counts was >300.

Owing to a lack of biological replicates,

‘

Blind

’

mode was used in DEseq,

which treats the two samples to be compared as replicates to estimate the

variance of gene expression. This approach assumes that most genes are not

differentially expressed (as would be the case in biological replicates), thus

generally overestimating variance, because the variance among samples

from different conditions is usually larger than among biological replicates.

Therefore, this method produces very conservative results with small

numbers of differentially expressed genes (Anders and Huber, 2010).

Using

-adj<0.1 and a fold change of >1.5, only a fraction of known otic

genes (14/37) is recovered (Table S1). To maximize the discovery of otic

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DEVELOPMENT

placode-enriched genes, all genes with a fold change of >1.5 were included

as candidates for further analysis. This analysis recovers 27/37 known otic

genes. Validation by

in situ

hybridization was used as a secondary screen to

ensure that the gene list did indeed represent otic placode-enriched

transcripts. Indeed, of all 39 factors screened by

in situ

hybridization only

five are not expressed in the placode, validating this as a useful strategy to

identify genes enriched in otic cells.

Gene ontology analysis for differentially expressed genes was performed

with DAVID (DAVID Bioinformatics Resources 6.7, http://david.abcc.

ncifcrf.gov/) (Huang da et al., 2009a,b). Disease-related enrichment was

analysed with Webgestalt (Wang et al., 2013; Zhang et al., 2005).

Partitioning of otic genes into different clusters

Both enriched otic genes and differentially expressed transcription factors

from pairwise analysis (except downregulated genes at 5-6ss relative to 0ss)

were used for hierarchical cluster analysis. Fold change of these factors at

5-6ss, 8-9ss and 11-12ss relative to 0ss was transformed into row z-score

with heatmap.2 and corresponding heatmaps were generated using gplot

within R (R Core Team, 2012; Warnes, 2012). Otic genes were clustered in

the same way to generate the clusters in Fig. S4.

Quantitative RT-PCR and NanoString nCounter analysis

RNA from dissected otic tissue and the whole embryo was isolated using the

RNAqueous-Micro Kit (Ambion) and reverse transcribed. Primers for target

genes were designed with PrimerQuest (IDT). qPCR was performed in

technical triplicates using Rotor-Gene Q (Qiagen) with SYBR green master

mix (Roche). The

ΔΔ

Ct method was used to calculate the fold change, which

was expressed as FC=2

∼

(

−

ΔΔ

Ct)

(Livak and Schmittgen, 2001).

Gapdh

Hprt

and

Rplp1

were used as reference genes to calculate the fold change.

RT-qPCR to validate the otic-enriched genes was performed from a single

biological replicate, and RT-qPCR for morpholino experiments used two

biological replicates; the

-value generated from a two-tailed Student

’

-test

was used to evaluate the statistical significance.

NanoString analysis was performed in triplicate per experimental

condition with the nCounter Analysis System using a customized probe

set of 216 genes. Five to ten tissues were lysed in lysis buffer (Ambion) and

total RNA was hybridized with capture and reporter probes according to the

nCounter Gene Expression Assay manual. The results were analysed using

the raw count with DEseq2 (Love et al., 2014). A transcript was considered

dysregulated when the mean count was >300, up- or downregulated by at

least |log2foldchange|>log2(1.2) and the

-adj<0.1. The cut-off of mean

count 300 is empirical based on the observation that genes not expressed or

very weakly expressed in the otic region had a count of less than 300.

GRN construction

GRN construction was performed manually using BioTapestry. Different

regions of the network were defined according to the known biology of OEP

induction, segregation of otic and epibranchial domains and the signalling

input from adjacent tissues taking into account the new transcriptional states

identified in this study and data from the literature (Fig. S5). Genes were

allocated to each region based on their temporal and spatial expression

pattern (published or determined in this study). Interactions were plotted

according to published data from different species as summarized in Fig. 5;

these largely depend on the analysis of mutants or knockdown experiments

using a few genes as otic markers and, with few exceptions, lack enhancer

information. The model presented in Fig. 7 incorporates the data from the

current study, which determined changes in gene expression after MO

knockdown of three transcription factors by RT-qPCR, NanoString

nCounter and/or whole-mount

in situ

hybridization. For some transcripts

all three methods were used, whereas for others only one or two approaches

were employed (Fig. S6). Occasionally, we observed a discrepancy between

the three detection methods; in this case, an interaction was defined if

expression change was detected by

in situ

hybridization, or if two methods

provided the same result. NanoString nCounter evaluation allows the

analysis of hundreds of genes in the same sample, thus providing a global

view of gene expression changes.

To establish links between upstream regulators and their downstream

targets, unless otherwise stated we assumed the most parsimonious

pathway: e.g. if gene 1 regulates gene 2 and 3, and gene 2 regulates gene

3 we assumed the simplest explanation of gene 1 gene 2 gene 3.

Acknowledgements

We thank Ewa Kolano, Annabelle Scott, Chia-Li Liao and Hilary McPhail for

excellent technical assistance and C. D. Stern for critical reading of the manuscript.

Competing interests

The authors declare no competing or financial interests.

Author contributions

A.S. designed the experiments; J.C. and M.T. collected otic tissues; R.R. performed

RNAseq for PPR tissue; J.C. analysed RNAseq data; J.C. and M.T. performed most

functional experiments and analysed all data together with A.S.; M.B. contributed to

the knockdown experiments; M.S.-C. contributed to RNAseq experiments; J.C., A.S.

and M.E.B. wrote the manuscript.

Funding

This study was funded by grants from the Biotechnology and Biological Sciences

Research Council (BB/I021647/1), Deafness Research UK (513:KCL:AS) and the

National Institute on Deafness and Other Communication Disorders (DC011577).

Deposited in PMC for immediate release.

Data availability

All sequencing data have been deposited in Gene Expression Omnibus under

accession number GSE69185 (https://www.ncbi.nlm.nih.gov/geo/query/acc.cgi?

acc=GSE69185).

Supplementary information

Supplementary information available online at

http://dev.biologists.org/lookup/doi/10.1242/dev.148494.supplemental

References

Abello

́

, G., Khatri, S., Radosevic, M., Scotting, P. J., Gira

́

ldez, F. and Alsina, B.

(2010). Independent regulation of Sox3 and Lmx1b by FGF and BMP signaling

influences the neurogenic and non-neurogenic domains in the chick otic placode.

Dev. Biol.

339

, 166-178.

Abu-Elmagd, M., Ishii, Y., Cheung, M., Rex, M., Le Roue

̈

dec, D. and Scotting,

P. J.

(2001). cSox3 expression and neurogenesis in the epibranchial placodes.

Dev. Biol.

237

, 258-269.

Adam, J., Myat, A., Le Roux, I., Eddison, M., Henrique, D., Ish-Horowicz, D. and

Lewis, J.

(1998). Cell fate choices and the expression of Notch, Delta and Serrate

homologues in the chick inner ear: parallels with Drosophila sense-organ

development.

Development

125

, 4645-4654.

Adamska, M., Herbrand, H., Adamski, M., Kru

̈

ger, M., Braun, T. and Bober, E.

(2001). FGFs control the patterning of the inner ear but are not able to induce the

full ear program.

Mech. Dev.

109

, 303-313.

Ahrens, K. and Schlosser, G.

(2005). Tissues and signals involved in the induction

of placodal Six1 expression in Xenopus laevis.

Dev. Biol.

288

, 40-59.

Anders, S. and Huber, W.

(2010). Differential expression analysis for sequence

count data.

Genome Biol.

, R106.

Bailey, A. P., Bhattacharyya, S., Bronner-Fraser, M. and Streit, A.

(2006). Lens

specification isthe ground state of all sensory placodes, from which FGF promotes

olfactory identity.

Dev. Cell

, 505-517.

Barembaum, M. and Bronner-Fraser, M.

(2007). Spalt4 mediates invagination and

otic placode gene expression in cranial ectoderm.

Development

134

, 3805-3814.

Barembaum, M. and Bronner-Fraser, M.

(2010). Pax2 and Pea3 synergize to

activate a novel regulatory enhancer for spalt4 in the developing ear.

Dev. Biol.

340

, 222-231.

Begbie, J., Brunet, J. F., Rubenstein, J. L. and Graham, A.

(1999). Induction of

the epibranchial placodes.

Development

126

, 895-902.

Betancur, P., Sauka-Spengler, T. and Bronner, M.

(2011). A Sox10 enhancer

element common to the otic placode and neural crest is activated by tissue-

specific paralogs.

Development

138

, 3689-3698.

Bouchard, M., de Caprona, D., Busslinger, M., Xu, P. and Fritzsch, B.

(2010).

Pax2 and Pax8 cooperate in mouse inner ear morphogenesis and innervation.

BMC Dev. Biol.

, 89.

Brown, S. T., Wang, J. and Groves, A. K.

(2005). Dlx gene expression during chick

inner ear development.

J. Comp. Neurol.

483

, 48-65.

Brugmann, S. A., Pandur, P. D., Kenyon, K. L., Pignoni, F. and Moody, S. A.

(2004). Six1 promotes a placodal fate within the lateral neurogenic ectoderm by

functioning as both a transcriptional activator and repressor.

Development

131

5871-5881.

Burton, Q., Cole, L. K., Mulheisen, M., Chang, W. and Wu, D. K.

(2004). The role

of Pax2 in mouse inner ear development.

Dev. Biol.

272

, 161-175.

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