ABSTRACT
Morphogenesis of the inner ear epithelium requires coordinated deployment of several signaling pathways and disruptions cause abnormalities of hearing and/or balance. The FGFR2b ligands, FGF3 and FGF10, are expressed throughout otic development and are required individually for normal morphogenesis, but their prior and redundant roles in otic placode induction complicates investigation of subsequent combinatorial functions in morphogenesis. To interrogate these roles and identify new effectors of FGF3 and FGF10 signaling at the earliest stages of otic morphogenesis, we used conditional gene ablation after otic placode induction and temporal inhibition of signaling with a secreted, dominant-negative FGFR2b ectodomain. We show that both ligands are required continuously after otocyst formation for maintenance of the otic ganglion and patterning and proliferation of the epithelium, leading to normal morphogenesis of both the cochlear and vestibular domains. Furthermore, the first genomewide identification of proximal targets of FGFR2b signaling in the early otocyst reveals novel candidate genes for inner ear development and function.
INTRODUCTION
The membranous labyrinth of the mammalian inner ear is among the most complex examples of organ morphogenesis. An unremarkable patch of cranial ectoderm is transformed into a structurally intricate sensory apparatus with two functionally distinct compartments: the ventral cochlea and the dorsal vestibular system, responsible for the perception of sound and acceleration, respectively. Within these compartments an exquisitely patterned array of sensory, non-sensory and supporting cell types are poised to transduce auditory and vestibular stimuli through sensory ganglia to the brain. Proper morphogenesis of the labyrinth is essential for normal auditory and vestibular function as indicated by imaging studies of hearing loss patients (Kimura et al., 2018; Sennaroglu and Bajin, 2017). In light of the advent of cochlear implantation to treat hearing loss in cases of inner ear malformation (Isaiah et al., 2017), elucidating signals governing otic morphogenesis and appreciation of the spectrum of labyrinthine morphogenetic defects are necessary to advance treatment.
Amniote inner ear development initiates during neurulation when hindbrain proximal ectoderm is induced to thicken, forming the otic placode, the source of both the otic epithelium and the neurons of its sensory ganglia. Next, the placode invaginates, forming a cup that deepens and delaminates neuroblasts, before pinching off from the overlying ectoderm to form a spherical vesicle, the otocyst, that at embryonic day (E)9.5 in mouse is already patterned along the three anatomical axes. Otocyst morphogenesis initiates with dorsomedial budding to form the endolymphatic duct and sac (EDS) anlagen. Vestibular structures initiate by sequential dorsal and lateral evaginations of the epithelium to form vertical and horizontal pouches, which are further sculpted by epithelial fusion and resorption, generating the three orthogonal semicircular canals. The utricle and saccule form from anterior/central bulges, and the cochlear duct (CD) emerges as a ventral outgrowth. In mice, it undergoes progressive ventral extension and coiling, reaching 1.75 turns by E15.5, when gross morphogenesis is largely complete (Morsli et al., 1998; Sajan et al., 2007). Cell type differentiation and functional maturation however, continue until well after birth (reviewed in Alsina and Whitfield, 2017; Basch et al., 2016; Whitfield, 2015; Wu and Kelley, 2012).
Signals regulating otic placode induction and early otocyst patterning emanate from surrounding tissues and are understood in some detail (Ladher, 2017), but by otocyst stages, intrinsic signals are also produced and their roles in driving region-specific morphogenesis are less well understood. Extrinsic signals regulating dorsal morphogenesis include hindbrain WNTs and BMPs. SHH secreted from the ventral hindbrain and notochord initiates ventral morphogenesis. Crosstalk between these signals involves regulation of key region-specific transcription factors (Ohta and Schoenwolf, 2018). FGF signaling also plays critical roles in otic development and functions at multiple stages. A cascade of FGFs from endoderm, mesoderm and hindbrain is required for otic placode induction (Alvarez et al., 2003; Ladher et al., 2005; Wright and Mansour, 2003a; Zelarayan et al., 2007). In particular, Fgf3 and Fgf10, encoding ligands that signal through the same FGF receptor isoforms (FGFR1b and FGFR2b; Zhang et al., 2006), are required redundantly for otic placode induction, such that germline double null mutants (F3KO;F10KO) have no inner ear (Alvarez et al., 2003; Wright and Mansour, 2003a). Applications of FGFs and FGFR inhibitors to chick embryos revealed profound influences of FGFs on otic morphogenesis (Chang et al., 2004) and studies of individual mouse mutants revealed roles for Fgf3 and Fgf10 in morphogenesis. Mice lacking Fgf3 (F3KO) fail with variable penetrance and expressivity to form an EDS, and consequently have variable morphogenesis and dysfunction of both the cochlea and vestibule (Hatch et al., 2007; Mansour et al., 1993). Mice lacking Fgf10 (F10KO) fail to form posterior semicircular canals (PSCCs), and have milder deformations of the anterior and lateral canals. Fgf10 heterozygotes also exhibit PSCC reductions or agenesis (Pauley et al., 2003; Urness et al., 2015). The CD is also affected in Fgf10 null mutants, being shorter and narrower than that of heterozygous or wild-type mice due to loss of non-contiguous non-sensory domains (Urness et al., 2015). Fgfr2b null mutants form otocysts (Pirvola et al., 2000), however, they develop with severe cochlear and vestibular dysmorphology, suggesting that Fgf3 and Fgf10 could have additional and combinatorial roles during morphogenesis.
Here, we define the dynamic expression of Fgf3 and Fgf10 in the developing mouse otic epithelium and ganglion, and interrogate their functions after otic placode induction. We employ two complementary genetic strategies: otic placode lineage-restricted gene ablation and timed induction of a soluble dominant-negative FGFR2b ectodomain that acts rapidly as an extracellular ligand trap to block signaling. Together, our data show that Fgf3 and Fgf10 are not required in the placode lineage for otocyst formation, but are required subsequently for otocyst patterning, neuroblast maintenance, epithelial proliferation and both vestibular and cochlear morphogenesis. Furthermore, the first differential RNA-Seq analyses of otocysts revealed FGFR2b signaling targets that define novel candidates for genes involved in otic morphogenesis and function.
RESULTS
Fgf3 and Fgf10 are expressed dynamically during otocyst and ganglion formation, and cochlear morphogenesis
To determine Fgf3 and Fgf10 expression during otocyst formation and cochlear morphogenesis, we used RNA in situ hybridization (ISH). Before E9.0, both genes were exclusively periotic (data not shown). Consistent with previous studies (Schimmang, 2007; Wright and Mansour, 2003b), Fgf3 and Fgf10 transcripts were non-overlapping at the otic cup stage, with Fgf3 expressed in hindbrain adjacent to the cup (Fig. 1A), and Fgf10 expressed in the cup itself, exclusive of the dorsal and lateral-most regions (Fig. 1B). Once the otocyst closed, Fgf3 was diminished in the hindbrain and was first seen in the ventrolateral otocyst and in the forming otic ganglion (Fig. 1C), whereas Fgf10 was expressed in the ventral and medial otocyst (Fig. 1D). By E10.25-E11.25, Fgf3 was confined to the ventrolateral otocyst (Figs. 1E,G). At this stage, Fgf10 overlapped with and was more extensively expressed in the ventral otocyst than was Fgf3, and also began to be expressed strongly in the ganglion (Figs. 1F,H).
We confirmed overlap of Fgf3 and Fgf10 in the developing vestibular sensory tissues, with Fgf10 expression much stronger than Fgf3 (data not shown; see Pauley et al., 2003; Pirvola et al., 2000). Therefore, we focused on the developing CD. From E12.5-E16.5, we found Fgf3 in a progressively limited portion of the CD that appeared by E16.5 to flank the developing sensory organ of Corti (Figs. 1I,K,M,O). Fgf10 continued expression in a broader cochlear epithelial domain than Fgf3, resolving to Kölliker’s organ by E16.5, and was maintained at high levels in the cochlear ganglion (Figs. 1J,L,N,P). These observations suggested that after otic induction, Fgf3 and Fgf10 could have combinatorial roles in morphogenesis and ganglion development.
Epithelial Fgf3 and Fgf10 are not required for otocyst formation, but both are required for vestibular and cochlear morphogenesis
Since F3KO;F10KO embryos lack inner ears (Alvarez et al., 2003; Wright and Mansour, 2003a), we took a conditional approach to disrupt these genes individually and combinatorially after otic placode induction using Tg(Pax2-Cre), which is active in the otic placode lineage starting at E8.5 (Fig. 2A; Ohyama and Groves, 2004). As this lineage comprises both epithelium and ganglion, Tg(Pax2-Cre) recombines in both tissues (Figs. 2B,C). In contrast to the variable otic phenotypes observed in F3KO mutants (Hatch et al., 2007; Mansour et al., 1993), deletion of Fgf3 alone in the Pax2-Cre lineage (F3cKO) had no effect on otic morphogenesis at E15.5 (Figs. 2D,D’) or on CD histology at E18.5 (Figs. 2E,E’). Indeed, F3cKO animals survived in the expected numbers and had normal auditory thresholds and motor behavior (data not shown). In contrast, F10cKO ears showed both vestibular and cochlear abnormalities, including reduction or loss of the PSCC and variable shortening and narrowing of the CD (Figs. 2F,F’), reflecting loss of Reissner’s membrane (Figs. 2G,G’). These abnormalities were similar to those of F10KO ears (Urness et al., 2015) and, indeed, immunostaining of E18.5 F10cKO CDs was similar to F10KO CDs (data not shown). Notably, Fgf10-/c Cre-negative ears had mild PSCC shortening (Fig. 2F), but CD defects appeared only in F10cKO ears (Fig. 2F’).
Next, we evaluated morphogenesis after varying Fgf3 and Fgf10 allele dosage. At E9.5, embryos of all genotypes had otocysts starting to develop an EDS (Figs. 2H,H’-K,K’), showing that, as expected from the timing of CRE activity onset, otic placode induction occurred normally, even in the absence of both Fgfs (Fig. 2K’). In contrast, when analyzed at E15.5 by paintfilling, distinct defects in epithelial morphology were apparent in many different genotypes (Figs. 2L,L’-O,O’). Even Cre-negative ears (Figs. 2L-O) showed vestibular defects when heterozygous for the Fgf10 null allele (Figs. 2N,O). Not surprisingly, all Cre-positive ears had reduced or absent PSCCs, as Fgf10 is at least heterozygous in those cases (Figs. 2L’-O’). These ears formed an allelic series of increasing severity, with F3cHet;F10cHet ears showing only mild PSCC reductions (Fig. 2L’). This was exacerbated in F3cKO;F10cHet ears (Fig. 2M’). However, F3cHet/F10cKO ears lost virtually the entire SCC system, showed reductions in the saccule and utricle, and had a more extreme shortening and narrowing of the CD (Fig. 2N’) than did F10cKO ears (Fig. 2F’). Only the EDS appeared normal. Preliminary analyses of E18.5 F3cHet/F10cKO ears did not reveal any exacerbation of changes in cochlear marker genes analyzed previously in F10KO ears (data not shown, see Urness et al., 2015). Strikingly, conditional disruption of both Fgf3 and Fgf10 blocked both vestibular and cochlear development, leaving only a small spherical vesicle (Fig. 2O’). Histologic sections of E18.5 F3cHet;F10cHet and F3cKO;F10cHet CDs (Figs. 2P,P’,Q,Q’) were indistinguishable from those of F3cKO CDs (Fig. 2E,E’), whereas F3cHet;F10cKO CDs were very narrow and lacked Reissner’s membrane (Fig. 2R,R’), similar to those of F10cKO (Fig. 2G,G’) and F10KO CDs (Urness et al., 2015). Whether the extreme narrowing of the CD in F3cHet;F10cKO ears is consistently more severe than in other types of Fgf10 mutants is unknown. The F3cKO;F10cKO “ear” had an epithelium comprising a thin, non-sensory region and a thickened vestibular-like sensory region. Most notably, these mutants showed no evidence of cochlear or vestibular neurons (Figs. 2S,S’). These data show that both Fgf3 and Fgf10 are required in the Pax2-Cre lineage, not only for vestibular morphogenesis, but also for cochlear morphogenesis and otic gangliogenesis, with the role of Fgf3 being revealed only in the absence of Fgf10.
Fgf3 and Fgf10 are not required in the Pax2-Cre lineage for early otocyst proliferation, but are required for otocyst patterning and maintenance of otic neuroblasts
Since F3cKO;F10cKO embryos ultimately develop very small otic vesicles, and this was first apparent at E10.5-E11.5, we quantified mitotic cells in E10.5 otocyst sections by calculating the number of phosphohistone H3 (pHH3)-positive cells per otic epithelial area. Somewhat surprisingly, the difference between control and F3cKO;F10cKO vesicles was not significant (Fig. S1A-C), perhaps as a consequence of Fgf3 maintained in the hindbrain through at least E9.25 (Fig. 1A). Indeed, in the absence of Fgf10, Fgf3 is required in the Sox1Cre lineage (hindbrain) to form a normally sized otocyst (Fig. S1D-G’), but this source of Fgf3 is not affected by Pax2-Cre (Fig. 2A).
To assess otocyst patterning in conditional mutants, we conducted whole-mount ISH analyses of E9.5-E11.5 samples using probes to detect regionally expressed genes that are known targets of FGF3 and/or FGF10 signaling and/or are required for morphogenesis. To manage the number of samples analyzed, we omitted single conditional mutants and used only a single Cre-negative genotype (Fgf3-/c;Fgf10-/c). Stained embryos were sectioned through the otocyst or viewed as whole mounts. At E9.5 most genes tested, as exemplified by Sox9, were unaffected even in F3cKO;F10cKO ears (Figs. 3A1-A5). Other genes unaffected by loss both Fgf3 and Fgf10 alleles at this stage included Dusp6, Spry1, Foxg1, Has2, Gbx2, Hmx3, Sox2 and Pax2 (data not shown), all of which are lost in F3KO;F10KO ears by otic placode stages (Alvarez et al., 2003; Urness et al., 2010; Wright and Mansour, 2003a). However, the common FGF signaling target, Etv5, which has distinct ventromedial and dorsolateral domains in E9.5 control otocysts (Fig. 3B1, B2), showed differential localization in conditional mutants with only a single Fgf3 or Fgf10 allele remaining (Figs. 3B3,3B4), and expression was entirely absent from F3cKO;F10cKO ears (Fig. 3B5), demonstrating that epithelial FGF3/FGF10 signaling was disrupted within 24 hours of CRE activation. The only other gene affected at E9.5 was Tbx1, which had dorsolateral and posteroventral otocyst domains in all genotypes (Figs. 3C1–3C4) except F3cKO;F10cKO, which lost the dorsolateral domain (Fig. 3C5). By E10.5, the dorsolateral Tbx1 domain was lost from both F3cHet;F10cKO and F3cKO;F10cKO otocysts (Figs. 3D4,D5).
At E10.5, expression of several other genes was still unaffected, as exemplified by Gbx2 (Fig. 3E1-E5), which is expressed dorsomedially, is down-regulated at this stage in F3KO mutants (Hatch et al., 2007), and is required for vestibular morphogenesis (Lin et al., 2005). Other genes unaffected by loss of Fgf3 and Fgf10 at E10.5 included Hmx3, Spry2, Gli1, Id1, and Lfng (data not shown). In contrast, Sox2, Foxg1 and Bmp4, which are primarily lateral at E10.5, were all unaffected, except in F3cKO;F10cKO ears, where expression was extinguished (Figs. 3F5,G5,H5). Other genes lost from E10.5 F3cKO/F10cKO otocysts included Dusp6, Etv5, Etv4, and Spry2 (data not shown), all of which are known transcriptional targets of FGF signaling. Curiously, we found that Pax2, which at E10.5 is normally expressed medially (Fig. 3I1), was unchanged in all otocysts except F3cKO/F10cKO, where it was expanded (Figs. 3I1-I5).
By E11.5 the only tested genes still unaffected in F3cKO;F10cKO otocysts were Gli1, Hmx3, Lfng and Id1 (data not shown), so these are unlikely to be targets of FGF3/FGF10 signaling in the otocyst. Whether the affected genes are direct or indirect targets of FGF signaling at this stage could not be determined.
To assess the otic ganglion, we assayed Neurog1 and its target, Neurod1, at both E9.5 and E10.5. At E9.5, all genotypes exhibited similar expression in a ventrolateral epithelial domain and in delaminating neuroblasts (Figs. 3J1-J5,3L1-L5). In contrast, at E10.5 Neurog1 was strongly reduced and Neurod1 was virtually eliminated from the otic epithelium, and otic ganglion development was suppressed specifically in F3cKO/F10cKO otocysts (Figs. 3K5,M5). These data show that epithelial/ganglion expression of Fgf3 and Fgf10 are required for aspects of gene expression driving otic morphogenesis, particularly in lateral regions, and that they are also required for otic ganglion formation.
Doxycycline induction of a secreted FGFR2b ectodomain phenocopies Fgf3/Fgf10 double null mutants
FGF3 and FGF10 bind to and signal primarily through “b”-type FGF receptors (FGFR2b>FGFR1b, Zhang et al., 2006). To enable simultaneous and inducible inhibition of their signaling activity at any stage, we employed two alleles that together enable doxycycline (DOX)-inducible expression of a secreted, dominant-negative form of FGFR2b (dnFGFR2b), which serves as a ligand trap. Rosa26rtTA drives ubiquitous expression of the reverse tetracycline transactivator (Belteki et al., 2005) and Tg(tetO-dnFgfr2b) encodes a tetO-regulated and secreted FGFR2b ectodomain (Hokuto et al., 2003). This system is validated for temporally controlled inhibition of mammary gland, tooth, limb and lung development, which depend on various FGFR2b and FGFR1b ligands (Al Alam et al., 2015; Danopoulos et al., 2013; Parsa et al., 2010; Parsa et al., 2008).
To validate this system for inner ear studies we crossed the alleles together, fed DOX-chow to pregnant females from E5.5-E10.5 and observed gross embryonic phenotypes. Relative to Rosa26rtTA/+ embryos, which appeared normal (Fig. 4A), double heterozygotes had a short curly tail, lacked limb buds and had tiny otic vesicles (Fig. 4B), phenocopying F3KO;F10KO mutants. Double heterozygotes exposed to DOX from E5.5 or E6.5 to E11.5 showed only an otic remnant (Fig. S2) and they did not exhibit mid-hindbrain phenotypes characteristic of inhibition of ligands such as FGF8 and FGF17, that signal through “c”-type FGFRs (Chi et al., 2003; Sato and Joyner, 2009; Xu et al., 2000).
Secreted dnFGFR2b acts rapidly to inhibit signaling by FGFR2b ligands
To determine the timing of signaling inhibition we initiated dnFGFR2b expression by injecting DOX at different stages, providing DOX-chow for various intervals, and assaying for Etv5 expression by ISH. After only 4 hours of DOX starting at E9.5, Tg(tetO-dnFgfr2b)/+ embryos showed robust Etv5 expression in numerous sites of FGF signaling (Fig. 4C), including throughout most of the otic vesicle (Fig. 4C’). In contrast, double heterozygotes retained many sites of Etv5 expression, but lacked any otic Etv5 (Fig. 4D,D’). In addition, 6 hours of DOX starting at E8.25 caused a near ablation of Etv5 throughout double heterozygotes, including in the otic cup (Fig. S3A,A’,B,B’) and 4 hours of DOX starting at E10.25 significantly downregulated Etv5 in the dorsolateral quadrant of the otic vesicle (Fig. S3C,C’,D,D’). Thus, inhibition of FGFR2b ligands has a rapid onset in otic tissue, consistent with studies of dnFGFR2b induction in the limb (Danopoulos et al., 2013).
FGFR2b ligands are required continuously for otic morphogenesis
Next, we asked when FGFR2b ligands are required for otocyst morphogenesis. Starting between E8.5 and E13.5 we injected DOX into pregnant dams and provided DOX-chow continuously through E15.5 when the inner ears were paintfilled. We compared Rosa26rtTA/+ (control) to Rosa26rtTA/+;Tg(tetO-dnFgfr2b)/+ (experimental) samples. DOX exposure from E8.5-E15.5 inhibited inner ear development such that experimental embryos had no otic tissue to fill (n=6/6; data not shown). When DOX was started on E9.5, control ears appeared normal (Fig. 4E) and experimental samples showed two distinct phenotypes: either no detectable inner ear (n=6/10; not shown) or a structure resembling an EDS (n=4/10; Fig. 4F). Starting DOX on E10.5 blocked development of distinct SCCs from the vertical canal pouch, reduced the size of the saccule and utricle, and caused a dramatic shortening and narrowing of the CD (n=16/18; Fig. 4G). This phenotype resembled the most strongly affected F10KO mutants (Urness et al., 2015). DOX exposure from E11.5-E15.5 was compatible with SCC formation, though these appeared thinner than those of control ears (n=8/8; Fig. 4H). Development of the utricle, saccule and CD were somewhat variable, but not markedly better than in the E10.5-E15.5 samples. Inner ears exposed to DOX from E12.5-E15.5 had a much more normal morphology, but still, the SCCs and CD were narrow (n=8/8; Fig. 4I). Even experimental ears exposed to DOX from only E13.5-E15.5 showed the thin SCC defect, but the rest of the inner ear appeared grossly normal (n=8/8; Fig. 4J). These data show that FGFR2b ligands are required continuously during otic morphogenesis.
Transient activation of dnFGFR2b reveals critical periods for FGFR2b ligands in otic morphogenesis
To determine more precise intervals for FGFR2b ligand requirements in particular events of otic morphogenesis, we treated pregnant dams with different DOX pulses and examined E15.5 inner ears by paintfilling. As expected, all ears exposed to DOX for even the longest pulse (24 h) had normal morphology (Figs. 5A,E,G,K,M,Q). In contrast, experimental ears showed exposure time-dependent abnormalities. A 4-hour DOX exposure starting at 9 AM on E8.5 (termed E8.25 for convenience) caused mild PSCC reductions in a few ears (n=3/14; Fig. 5B) and had no discernable effect on the remaining ears, whereas 6-hour or 24-hour exposures had increasingly severe consequences. Most of the 6-hour group (n=14/19) and some of the 24-hour group (n=2/16) lacked an EDS (Figs. 5C’,D). The majority of 24-hour exposures blocked most development of the otocyst, leaving a small vesicle (n=6/16; Fig. 5D’) or no ear tissue (n=8/16; data not shown). By delaying DOX administration to 9 PM on E8.5 (called E8.75) all experimental samples that had any ear tissue (n=17/32) showed at least an EDS and most (n=12/32) had a central (vestibular) segment and a linear CD (Figs. 5F,F’). A 2-hour DOX pulse starting at 9 AM on E9.5 (called E9.25) had no effect on morphogenesis (data not shown), but 4- or 6-hour exposures consistently caused only PSCC defects (Figs. 5H,I,I’). The 24-hour exposures allowed EDS outgrowth, but consistently blocked most vestibular and cochlear outgrowth (n=40/40; Figs. 5J,J’). 12-hour DOX exposures starting at 9 PM on E9.5 permitted EDS outgrowth and formation of at least a vertical canal pouch, but no SCC formation. In addition, the CD was short and narrow (n=14/16) or not present (n=2/16; Fig. 5L,L’). DOX exposures starting at 9 AM on E10.5 (termed E10.25) and extending for 4 or 6 hours consistently blocked normal PSCC formation (n=19/20; both groups considered together) and the 6-hour exposure sometimes affected the ASCC as well (n=3/10), but had virtually no effect on development of the utricle, saccule or CD (Figs. 5N,O,O’) until the exposure reached 24 hours (Figs. 5P,P’). By starting DOX at 9 PM on E10.5 (termed E10.75), the most severe defects were avoided, nevertheless, the SCCs appeared thin, the utricle and saccule were reduced and the CD was short (n=5/5; Fig. 5R). In summary, we found that the earlier DOX was started and the longer it was present, the more severe were the morphogenesis defects. Furthermore, some of the DOX pulses gave such consistent outcomes that it seemed possible to identify acute epithelial transcriptional targets of FGFR2b ligands mediating particular morphogenetic events.
RNA-Seq reveals transcriptional targets of FGFR2b ligands during early otic morphogenesis
To identify transcriptional targets of FGFR2b ligands during early otocyst morphogenesis, when they are required for both vestibular and cochlear outgrowth, we chose three DOX exposures (Fig. 6A) that gave similar morphogenesis outcomes: E9.75+12 hours (Seq1, Figs. 5L,L’), E10.25+24 hours (Seq2, Figs. 5P,P’) and E9.25+24 hours (Seq3, Figs. 5J,J’). Immediately following DOX exposure, otocysts were microdissected, cleaned of mesenchyme (Fig. 6B), and pooled into separate control and experimental groups from each female. RNA was isolated, processed for RNA-Seq and analyzed for differential expression under both unpaired (genotype only) and paired (genotype and litter) statistical models.
Differentially expressed genes with an adjusted (adj) P<0.05 in each paired dataset were visualized with volcano plots (Fig. 6C; Fig. S4). In all 3 datasets the maximum fold-changes were relatively modest, perhaps reflecting the short periods of inhibition, but for many genes the differences were highly significant. Fgfr2 and Ighg1 were among the most highly differentially expressed Seq1 genes (5.3-fold and 633-fold induced, respectively, but Ighg1 was omitted from the plot for legibility; Fig 6C). Inspection of Fgfr2 reads showed upregulation was due to expression of the Fgfr2b splice isoform specifically in transgene-containing samples. The Ighg1 reads were also transgene-specific, thus validating the efficacy of the inductions. Excluding Fgfr2 and Ighg1 as vector-specific, there were 968 genes >1.5-fold upregulated and 631 genes >1.5-fold downregulated (adjP<0.05) in experimental otocyst RNA. Significantly downregulated genes included well-known transcriptional targets of FGF signaling including Etv1, Etv4, Etv5, Dusp6, Spry2 and Spry1 (indicated in Fig. 6C and listed, Excel S1). Similar analyses of Seq2 and Seq3 also showed significant upregulation of vector-specific sequences and significant downregulation of known FGF target genes (Fig. S4, listed in Excel S1). To validate an FGF target gene significantly downregulated in all 3 datasets, we detected Etv5 by ISH of otocyst sections. Seq1 control otocysts showed lateral and ventromedial Etv5 expression, whereas experimental embryos did not express otocyst Etv5 (Fig. 6D,E). Similar results were obtained with Seq3 samples (Fig. S5).
Fgfr1 sequence reads in each dataset showed no changes in level between control and experimental samples, which were similar to control levels of Fgfr2. Interestingly, Fgfr1c, thought to be mesenchymal, was the predominant splice isoform, but Fgfr1b was also detected. Fgfr3 sequences were present at levels at least 20-fold below those of Fgfr1 or Fgfr2 in control samples and were unchanged by dnFGFR2 induction (Excel S1). The only FGFR2b ligand genes expressed at significant levels in control or experimental otocysts were Fgf3 and Fgf10 (Excel S1), consistent with ISH surveys (Wright et al., 2003 and data not shown). Interestingly, Fgf3 was slightly, but significantly upregulated in all 3 datasets. However, as dnFGFR2b inhibition acts at the level of protein, this is unlikely to impact the phenotypes.
FGFR2b ligands are required to promote otocyst cell proliferation
To explore functional relationships between significantly differentially expressed genes in each dataset (either up or downregulated; adjP<0.05), we used Ingenuity Pathway Analysis. In each case, the top 5 affected pathways included cell cycle and DNA damage/repair pathways (Excel S2; -log(P-value)=8-13). In most cases these genes were downregulated in our datasets. In addition, we used GOrilla software to identify gene ontology terms for processes enriched in the downregulated Seq1 dataset (adjP<0.05), and the results were similar (top 5 shown in Excel S2; FDR q-values 1.55x10-21-7.84x10-13). To assess proliferation more directly, we quantified pHH3-positive cells per otic epithelial area in Seq3 otocyst sections and found a significant ~1.8-fold decrease in pHH3 labeling of experimental samples relative to controls (Fig. 6F-H), suggesting that one role for FGFR2b ligands during E9.25-E10.25 is to control the rate of otic epithelial proliferation.
Signaling by FGFR2b ligands in the early otocyst represses genes that function later in otic epithelial development or hearing
Among the significantly upregulated genes in dnFgfr2b samples from the Seq1 paired analysis, we noticed a gene that, when mutated, causes human hearing loss (Gjb6; Fig. 6C). To determine whether other such genes, or those responsible for mouse hearing loss and/or cochlear development, were enriched in either the up- or downregulated genesets, we conducted a gene set enrichment analysis (Subramanian et al., 2005) on all 16,232 genes detected in the Seq1 paired analysis using two partially overlapping gene lists: 95 human hereditary hearing loss genes identified by Nishio et al. (2015; rank listed in Excel S3) and 258 mouse genes involved in inner ear development or function collated by Ohlemiller et al. (2016; rank listed in Excel S4). Both genesets were highly enriched in the upregulated Seq1 dataset (normalized enrichment score=2.09 for the human genes and 2.06 for the mouse genes; both nominal P-values, <1x10-3), but not in the downregulated set. Similar results were obtained with the Seq2 dataset (data not shown). Together, these analyses suggest that one role of FGFR2b ligands at this early stage of morphogenesis is to prevent premature expression of epithelial genes that have later roles in development or function of the inner ear.
Validation of new genes regulated by FGFR2b ligands during the early stages of otocyst morphogenesis
To validate new FGFR2b target genes by ISH we focused first on downregulated genes and assayed selected genes based on overlap in multiple RNA-Seq datasets (Excel S5), relatively high degree of differential expression, and normalized read count above that of Fgf3 (Excel S1), which is relatively difficult to detect by ISH, and novelty with respect to inner ear development and/or FGF/MAPK signaling. Seven genes validated as downregulated in Seq1 otocysts are illustrated in Figure 7. Spred3 was expressed in E10.25 control otocysts in a ventromedial domain (Fig. 7A), and was greatly reduced in experimental otocysts (Fig. 7A’). Six2 was detected in the ventral-most region of control otocyts as well as laterally (Fig. 7B), but was absent from dnFGFR2b otocysts (Fig. 7B’). Prdm1(Blimp1) was expressed similarly to Six2 in control otocysts (Fig. 7C) and was strongly downregulated in the corresponding experimental otocysts (Fig. 7C’). Crlf1 was also expressed similarly to Six2 (Fig. 7D) in controls, but ventral expression was absent and lateral expression was downregulated in dnFGFR2b otocysts (Fig. 7D’). Tspan15 was expressed in a broad ventrolateral domain in controls (Fig. 7E) and was largely absent from dnFGFR2b otocysts (Fig. 7E’). Pou3f3 was expressed in the ventral-most region of controls (Fig. 7F) and was absent from dnFGFR2b otocysts (Fig. 7F’). Finally, Gchfr was expressed similarly to Pou3f3 (Fig. 7G) and was absent from dnFGFR2b otocysts (Fig. 7G’). We observed similar downregulation of Spred3, Prdm1, Crlf1, Tspan15, and Gchfr expression in dnFGFR2b otocysts subjected to the Seq2 and Seq3 DOX exposures (Figs. S6,S7). Six2 and Pou3f3 downregulation was confirmed by ISH in Seq3 otocysts (Fig. S7), but not tested in Seq2 otocysts, as these genes were not significantly affected in the Seq2 dataset.
We also tested several genes common to the upregulated lists, but most were widely expressed in controls and any changes in expression levels were not revealed by ISH (data not shown). However, Bmper transcripts were confined to the dorsomedial region of control otocysts (Fig. 7H), but the expression domain expanded to encompass most of the otic epithelium in experimental otocysts (Fig. 7H’). Similar results obtained with Seq2 and Seq3 otocysts (Figs. S6,S7). Therefore, these datasets are a rich source of novel FGFR2b signaling targets in the otocyst.
DISCUSSION
Fgfr2b and two genes encoding FGFR2b ligands, Fgf3 and Fgf10, are required individually for otic morphogenesis (Hatch et al., 2007; Mansour et al., 1993; Pauley et al., 2003; Pirvola et al., 2000; Urness et al., 2015). We found that Fgf3 and Fgf10 are expressed continuously during otic morphogenesis, however, their requirement for otic placode induction (Alvarez et al., 2003; Urness et al., 2010; Wright and Mansour, 2003a) obscured potential combinatorial roles during otocyst morphogenesis. Here, we blocked FGF3 and FGF10 signaling after otic placode induction by using conditional gene inactivation and temporally controlled inhibition via a drug-inducible ligand trap. Analysis of conditional mutants revealed that both genes are required in the otic lineage after E8.5 for both vestibular and cochlear development, but that the role of Fgf3 is only revealed in the absence of Fgf10, and that these signal-encoding genes control expression of genes that function in otic epithelial morphogenesis. Furthermore, we found that FGF3 and FGF10 signal redundantly to maintain otic neuroblasts. Temporally regulated inhibition showed that FGFR2b signaling acts continuously throughout otic morphogenesis and that pulses of inhibition can be used to identify the timing of particular morphogenetic events. We used brief windows of inhibition at early stages of morphogenesis to conduct the first genomewide identification of proximal targets of FGFR2b signaling in the otocyst, and validated several genes that are novel candidates for involvement in the initial stages of otocyst morphogenesis.
Fgf3 and Fgf10 are expressed and function continuously to control otocyst formation and morphogenesis
Fgf3 and Fgf10 are expressed continuously throughout otocyst development and cochlear morphogenesis. These are likely the only relevant FGFR2b (or FGFR1b) ligand-encoding genes for early otic morphogenesis as, based on RNA-Seq data, the others are either not expressed (Fgf22) or are detected at negligible levels (Fgf1 and Fgf7). The use of stage-matched tissue throughout morphogenesis helped to define the relative temporal and spatial dynamics of the expression patterns and aided in interpreting functional perturbations. We found that Fgf10 has an earlier and broader distribution than Fgf3 in the otic epithelium,and both transcripts are present in the otic ganglion, but Fgf3 is seen only transiently, whereas once Fgf10 starts expressing, it is present continuously at high levels in the cochlear ganglion. Although Fgf10 is expressed in mesenchyme underlying the preplacodal ectoderm, neither gene appears in periotic mesenchyme during otic cup formation or later (Schimmang, 2007; Urness et al., 2015; Wright and Mansour, 2003b). ISH data for Fgfr2b and Fgfr1b, which encode the receptors for FGF3 and FGF10, are limited because of the small size of probes that distinguish them from “c” isoforms, but extant data are consistent with the idea that they are also primarily epithelial (Orr-Urtreger et al., 1993; Pirvola et al., 2000; Wright et al., 2003; Wright and Mansour, 2003a) and, indeed, the “b” isoforms are evident in our otocyst RNA-Seq datasets. Therefore, taken together, the ligand and receptor expression data are consistent with our findings of continuous and combinatorial roles for Fgf3 and Fgf10 in otocyst epithelial morphogenesis and otic ganglion development. They also raise the interesting possibility that signaling involves epithelial and/or ganglion ligands activating epithelial, rather than mesenchymal or ganglionic receptors, except perhaps as neuroblasts begin delamination from the epithelium. This contrasts with epithelial ligands FGF9 and FGF20, which signal initially to “c” type receptors in the mesenchyme, inducing signals that control epithelial proliferation (Huh et al., 2015).
Fgf3 and Fgf10 are both required for vestibular and cochlear morphogenesis and for maintenance of otic neuroblasts
Conditional mutant analyses showed that both genes are required after otic placode induction, but deletion of Fgf3 alone from the Pax2-Cre lineage was inconsequential. This contrasts with the variably penetrant otic dysmorphology of F3KO mutants, the most severe of which initiate with alterations of dorsal otocyst patterning, loss of the EDS, and subsequent cystic development of the epithelium, ultimately resulting in hearing loss and circling behavior (Hatch et al., 2007; Mansour et al., 1993). The normal phenotype of F3cKO ears points to a critical role for Fgf3 expression in the hindbrain. Indeed, we found that F10KO embryos in which only hindbrain sources of Fgf3 were deleted (using Sox1Cre) had very small otocysts. In contrast, F10cKO ears had abnormalities very similar to those of F10KO ears. This demonstrates that the unique functions of Fgf10 in otic morphogenesis arise from its expression in the placodal lineage rather than earlier in the mesenchyme. Analysis of conditional mutants that separate epithelial from ganglion sites of Fgf10 expression will be needed to further dissect the spatial requirements for Fgf10 function.
Although Pax2-Cre is active in the placode at E8.5, we found no obvious effects at E9.5 on otocyst morphology in F3cKO;F10cKO embryos, and only two tested genes, Etv5 and Tbx1, were lost or altered in these otocysts. The first major losses in expression of multiple genes required for morphogenesis occurred at E10.5. This shows that both Fgf3 and Fgf10 are required in the placode lineage for normal otocyst morphogenesis and suggests that overlapping expression of Fgf3 and Fgf10 starting at E9.5 may be critical for both cochlear and vestibular outgrowth and morphogenesis. However, the phenotypes of triple allelic conditional mutants point to functional differences between Fgf3 and Fgf10. Both the cochlear and vestibular morphology of F3cKO/F10cHet ears were less severely affected than in F3cHet/F10cKO ears. This may reflect the relatively larger domain and higher level of epithelial Fgf10 than of Fgf3, and may be presaged by the differential effects on Etv5 expression in the two types of E9.5 otocysts. The loss of dorsolateral Etv5 when Fgf10 is the only remaining allele, and of ventromedial Etv5 when Fgf3 is the only remaining allele, suggest that Fgf3 is particularly important dorsally and Fgf10 ventrally, at least initially. The loss of dorsolateral Tbx1 in the two most severely affected genotypes likely reflects effects of FGF3/FGF10 signaling on development of the vertical canal pouch, the derivatives of which (PSCC and ASCC) are strongly affected in these and in Tbx1 mutants (Freyer et al., 2013; Macchiarulo and Morrow, 2017). Whether this is a direct or indirect effect on Tbx1 expression is not yet clear, but it is interesting to note that Tbx1 is slightly, but significantly downregulated in the Seq2 and Seq3 datasets (Excel S1).
The presence of an EDS in both triple allelic conditional mutants and normal Gbx2 expression in these mutants and in F3cKO/F10cKO mutants contrasts with findings from F3KO mutants (Hatch et al., 2007), which usually lack an EDS and lose Gbx2 expression by E10.5. This is consistent with the idea that hindbrain, rather than epithelial Fgf3, induces the EDS. It is possible that the further shortening of F3cHet/F10cKO CDs results from reduced FGF3-stimulated proliferation rather than alterations in molecular patterning. This is supported by preliminary analyses of E18.5 ears that did not reveal any exacerbation of changes to CD marker genes analyzed previously in the F10KO mutant (data not shown, see Urness et al., 2015). However, the timing of such proliferative effects in F3cKO;F10cKO mutants must be later than E10.5, when differences in pHH3 labeling between control and F3cKO/F10cKO otocysts were not significant.
We suggested previously that Fgf3 plays a role in otic ganglion development, as the F3KO ganglion, like that of Fgfr2b null mutants (Pirvola et al., 2000), is smaller than normal (Mansour et al., 1993). In contrast, F10KO early otic ganglia and later cochlear ganglia appear normal (Urness et al., 2015) despite defects of vestibular innervation consequent to midgestation loss of vestibular sensory epithelia (Pauley et al., 2003). In contrast to zebrafish (Vemaraju et al., 2012), our present data from F3cKO;F10cKO mutants suggest that Fgf3 and Fgf10 are required together for maintenance of Neurog1 and Neurod1 expression, and development of an otic ganglion, rather than specification of otic neuroblasts. Our data do not address whether this requirement involves ligand expression in the epithelium or ganglion or both. However, by restricting dnFGFR2b expression to the placodal lineage by using Pax2-Cre in combination with the unrecombined Rosa26lslrtTA allele, it may be possible to avoid disrupting otic induction and determine whether FGFR2b ligands play any role in mouse otic neuroblast specification. In addition, this paradigm of tissue restricted and timed induction of dnFGFR2b could also be used to identify candidate genes responsible for neuroblast maintenance. Determining the role of the transient burst of Fgf3 in delaminating neuroblasts will be more challenging.
Temporally controlled inhibition of FGFR2b signaling during otocyst morphogenesis reveals requirements at multiple stages
Observations of embryos expressing dnFGFR2b between E5.5 and E10.5 showed that this inhibition strategy effectively phenocopies F3KO;F10KO mutants without causing abnormalities characteristic of disrupting FGFs that signal through “c”-type receptors. Thus, dnFGFR2b has the expected specificity and the only relevant FGFR2b ligands for otocyst morphogenesis are likely to be FGF3 and FGF10. Paintfilling of embryonic ears subjected to ubiquitous and chronic dnFGFR2b expression starting on different days of development revealed that FGFR2b ligands are required continuously for otic development at least through E13.5. Pulses of dnFGFR2b caused highly specific and penetrant otic malformations, supporting the idea that unlike irreversible CRE-mediated deletion of coding exons, the signaling inhibition effected by dnFGFR2b is reversible.
Together, the chronic and pulsed inhibition paradigms suggest a distinct progression of roles for FGFR2b ligands: first in inducing the placode, then in stimulating EDS, vestibular pouch and CD outgrowth, and finally in sculpting the SCCs, outgrowth of utricle and saccule, and specification of CD nonsensory tissue. As otic induction is complete by ~E8.5, it was surprising that starting chronic dnFGFR2b expression on E9.5 blocked ear development in 6/10 cases. Since dnFGFR2b embryos from Seq3 inductions (E9.25-E10.25) always had otocysts, and these had significantly reduced proliferation, we suggest that the loss of otic tissue in embryos induced chronically after E8.5 reflects degeneration of mitotically blocked cells rather than failure of otic induction. Some phenotypes revealed the timing of roles for FGFR2b ligands. For example, the 6–24 hours starting at E8.25 is particularly important for EDS formation, potentially reflecting FGF3 inhibition (Mansour et al., 1993; Hatch et al., 2007) and the 6-hours starting at E9.25 is important for PSCC formation, potentially reflecting FGF10 inhibition (Pauley et al., 2003; Urness et al., 2015). Other vestibular phenotypes are particularly interesting as they reveal three potential and previously unsuspected functions for FGFR2b signaling. Chronic dnFGFR2b induction starting at E10.5, or 4–24 hour pulses starting at E10.25 blocked fusion of vestibular pouches, suggesting requirements for FGFR2b ligands in fusion plate formation and chronic induction starting between E11.5-E13.5 or 12-hour pulses starting E9.75-E10.75 caused very thin semicircular canals, suggesting roles for FGFR2b ligands in limiting resorption of fusion plates. Finally, several conditions reduced utricle and saccule development. Thus, it will be interesting to explore regulatory relationships between FGFR2b signaling and genes already known to regulate vestibular mophogenesis (Alsina and Whitfield, 2017), as well as to induce dnFGFR2b in particular temporal/spatial windows and pursue unbiased identification of effector genes involved in the development of particular structures of interest.
FGFR2b ligands promote otic epithelial proliferation and prevent premature expression of genes required for hearing
Our RNA-Seq datasets revealed significant downregulation of genes involved in the cell cycle and DNA repair, and indeed, immunostaining of Seq3 samples showed that mitotic cell numbers were significantly reduced in E10.25 dnFGFR2b-containing otocysts. This result differed from that obtained with E10.5 F3cKO/F10cKO otocysts, which did not show a mitotic defect. Given that hindbrain Fgf3 is unaffected in Pax2-Cre;F3cKO/F10cKO mutants, and is extinguished by E10.5, it is likely that otocyst proliferation defects in these mutants would be detected at later stages of morphogenesis.
The RNA-Seq datasets also revealed significantly upregulated genes. We found that in Seq1 and Seq2, these genes are highly enriched for human hereditary hearing loss genes and mouse genes that are expressed and/or function later in the inner ear. These include Pax2, which was expanded in E10.5 F3cKO/F10cKO otocysts. This suggests that at early stages, FGFR2b signaling normally represses many genes important for later development and function of the cochlea, or alternatively, that the proliferative block imposed by dnFGFR2b expression promotes early differentiation of the epithelium. The latter possibility however, does not apply to the earliest genes required for sensory cell differentiation (Atoh1, Pou4f3 and Gfi1), which were detected at very low levels and were unaffected in any of the RNA-Seq datasets. We suggest that the upregulated genesets are worth mining for new candidates for hearing loss genes, of which many remain to be identified (Bowl and Brown, 2018).
Although we show that FGFR2b ligands are required to activate a Bmp (Bmp4) and repress a BMP regulatory gene (Bmper), we found no evidence for FGFR2b ligand regulation of key downstream components of the BMP or SHH pathways, suggesting that while FGFR2b ligands may regulate individual components of these pathways, at least at the stages investigated, they are not exclusively upstream of these key programs directing dorsal and ventral otic morphogenesis, respectively.
Novel targets of FGFR2b signaling in early otocyst development
We validated by ISH seven novel genes downregulated and one gene upregulated by FGFR2b ligands in the RNA-seq datasets. Some genes may be regulated directly by the intracellular signaling pathway activated by FGFR2b, as one of our analysis points was only 12 hours after induction. As it was not possible to study more than a few differentially expressed genes, it is difficult to speculate about their combinatorial functions in inner ear development. Nevertheless, it is interesting to note that the downregulated genes in our most robust dataset (Seq1) are enriched for transcription factor-coding genes (Excel S2), including those validated here, Six2, Prdm1 and Pou3f3. The first two have otocyst expression patterns similar to Spred3, Crlf1 and Tspan15, whereas Pou3f3 expression appears to overlap with Gchfr. This suggests that further mining of the existing differential expression data and generation of additional targets by employing different windows of FGFR2b inhibition, combined with promoter analysis and genomewide studies of otocyst chromatin modification could suggest important new gene regulatory networks acting to shape the epithelium.
The only upregulated gene validated by ISH was Bmper, which encodes BMP-binding endothelial regulator, an ortholog of Drosophila crossveinless-2 (cv-2) (Coffinier et al., 2002). cv-2 modulates BMP signaling biphasically in the fly wing with the direction of action dependent upon the concentration of Cv-2 and the concentration and types of local BMP ligands (Conley et al., 2000; Serpe et al., 2008). Similarly, in mouse, Bmper modulates the availability of BMPs, enhancing signaling when ligands are low and limiting signaling when ligands are high (Dyer et al., 2014; Kelley et al., 2009). Thus, Bmper null mutants have some phenotypes suggestive of a classic BMP signaling antagonist (Moser et al., 2003) and others suggestive of a BMP signaling agonist (Ikeya et al., 2006). Multiple Bmps and their receptors are expressed in and required for otocyst morphogenesis (Chang et al., 2008; Hwang et al., 2010; Ohyama et al., 2010) and misexpression of BMP ventral to the otic placode blocks outgrowth of the chick cochlea (Ohta et al., 2016). Thus, it will be interesting to determine whether the otic phenotype of a Bmper null mutant reflects a loss or gain of BMP signaling, and whether this differs in different regions of the developing otocyst. Our results also showed that at early stages of otocyst morphogenesis, Fgf3 and Fgf10 are required for expression of Bmp4, which is itself required for both vestibular and cochlear development (Chang et al., 2008). Determining whether FGFR2b ligand-dependent upregulation of Bmper functions in this context to further antagonize BMP signaling, or alternatively, to mitigate the reduction in Bmp4 by increasing signaling by other BMP ligands will require additional studies of otic Bmp expression and manipulation of Bmper allele levels in combination with dnFGFR2b induction at different stages.
The identification of several FGFR2b target genes not implicated previously in ear development or hearing loss syndromes provides a tantalizing glimpse into a new set of potential otocyst morphogenetic factors. Given the novelty of these targets, it is tempting to speculate that previously unappreciated regulatory pathways may be at play during otic morphogenesis, as has been postulated for otic placode induction (Anwar et al., 2017). Functional studies will be required to address the roles of each of these new genes.
MATERIALS AND METHODS
Mouse models and genotyping
Mice were maintained and euthanized in accordance with protocols approved by the University of Utah Institutional Animal Care and Use Committee. All Fgf mutant alleles were kept on a mixed genetic background comprised of C57Bl/6 and various 129 substrains. CD-1 outbred mice (Charles River Laboratory) were used to generate embryos for studies of normal expression patterns and for generating embryos for induction of dnFGFR2b. Noon of the day a mating plug was observed was considered E0.5.
Generation and PCR genotyping of the Fgf3 and Fgf10 null alleles (Fgf3-, formally designated Fgf3tm1.1Sms = MGI:3767558 and Fgf10-, formally designated Fgf10tm1.1Sms = MGI:3526181) and Fgf3 and Fgf10 and conditional alleles (Fgf3c; Fgf3tm1.2Sms = MGI:4456396] and Fgf10c; Fgf10tm1.2Sms = MGI:4456398) were described previously (Hatch et al., 2007; Urness et al., 2010). Tg(Pax2-Cre) mice (Tg(Pax2-cre)1Akg = MGI:3046196) were obtained from Dr. Andrew Groves (Ohyama and Groves, 2004). Tg(Pax2-Cre) was detected by PCR using primers specific to the transgene (5’ GGGGATCCCGACTACAAGG 3’; 5’ TAGTGAGTCGTATTAATTTCGATAAGC 3’). The Sox1Cre allele (Takashima et al., 2007) was transferred from Dr. Mario Capecchi with permission from Dr. Shin-Ichi Nishikawa (RIKEN) and genotyped using generic Cre primers. Rosa26lslLacZ reporter mice (Gt(ROSA)26Sortm1Sor = MGI:1861932) (Soriano, 1999) were maintained as homogyzotes.
Single conditional mutants were generated by crossing Fgf3c/c females to Fgf3-/+;Tg(Pax2-Cre)/+ males or Fgf10c/c females to Fgf10-/+;Tg(Pax2-Cre)/+ males. Combinations of Fgf3 and Fgf10 conditional mutants were obtained by crossing Fgf3c/c;Fgf10c/c females to Fgf3-/+;Fgf10-/+;Tg(Pax2-Cre)/+ or Fgf3-/+;Fgf10-/+; Sox1Cre/+ males. CRE activity was confirmed by mating males to Rosa26LacZR/LacZR females harvesting embryos at the indicated stages and staining with X-gal as described (Yang and Mansour, 1999).
The germline recombined Rosa26rtTA allele (derived from Gt(ROSA)26Sortm1(rtTA,EGFP)Nagy; MGI:3583817) (Belteki et al., 2005; Parsa et al., 2008) and Tg(tetO-dnFgfr2b) (Tg(tetO-Fgfr2b/Igh1.3Jaw; MGI 5582625) (Hokuto et al., 2003) alleles were transferred from the laboratory of Dr. Saverio Bellusci with permission from Dr. Jeffery Whitsett (Cincinnati Children’s Medical Center). Genotyping primers to detect Tg(tetO-dnFgfr2b) were 5’ CAGGCCAACCAGTCTGCCTGGC 3’ and 5’ CGTCTGAGCTGTGTGCACCTCC 3’. ROSA26rtTA genotyping primers were ROSA5 (5’ GAGTTCTCTGCTGCCTCCTG 3’) and ROSA3 (5’ CGAGGCGGATCACAAGCAATA 3’), which generate a wild type band of 322 bp and ROSA5 and RTTA3 (5’ AAGACCGCGAAGAGTTTGTC 3’), which generate a 215 bp rtTA-specific product. Double heterozygotes were obtained initially by crossing single heterozygotes. For most of the studies described here, we crossed wild type CD-1 females to Rosa26rtTA/rtTA;Tg(tetO-dnFgfr2b)/+ males, generating 50% each of control and experimental genotypes.
RNA in situ hybridization
Embryos were harvested and fixed in 4% PFA and stored in methanol at -20°C. RNA ISH to whole mount embryos or paraffin-embedded sections were performed as described (Urness et al., 2008; Urness et al., 2010). Probes for Sox9, Fgf3, Fgf10, Bmp4, Etv5, Gbx2, Sox2, Foxg1, Pax2, NeuroD1, Ngn1 and Crlf1 were generated by transcription of cDNA-containing plasmids. Template plasmids and acknowledgements are shown in Table S1. All other RNA probes were generated by transcription of a PCR-amplified, gene-specific 3’ UTR fragment containing a T7 promoter. The primer sequences are shown in Table S2. Whole embryos were photographed using a stereomicroscope (Zeiss Discovery.V12) fitted with a digital camera (QImaging Micropublisher 5.0). Hybridized tissue sections were photographed under DIC illumination (Zeiss Axioskop) using a digital camera (Zeiss Axiovision or Lumenera Infinity3).
Immunostaining of frozen tissue sections for quantification of mitotic cells in the otocyst
Embryos were fixed in 4% paraformaldehyde solution and cryosectioned in the transverse plane for immunostaining as described (Urness et al., 2015). Rabbit anti-phosphohistone H3 (Millipore 06-570) was applied at a dilution of 1:400 and mouse monoclonal anti-E-cadherin (BD Biosciences 610181) was diluted 1:60. Secondary antibodies were from Invitrogen and diluted 1:400 into PBST/5% normal serum (Alexa Fluor® 488 goat anti-rabbit (A11034) and Alexa Fluor® 594 goat anti-mouse (A11032)). DAPI was included in the mounting medium (Vectashield, Vector Labs). Fluorescent signals were observed under epi-illumination on a Zeiss Axioskop and captured using an Infinity3 camera (Lumenera) driven by InfinityAnalyze software. Channels were overlaid using Photoshop CS5. All pHH3-positive cells in the otocysts (defined by E-Cadherin staining) were counted from 6 μm (Pax2-Cre cross) or 8 μm (dnFgfr2b cross) sections extending from anterior to posterior. N=8 control (either Fgf3-/c;Fgf10-/c or Fgf3c/+;Fgf10c/+;Pax2-Cre/+) and n=6 experimental (Fgf3-/c;Fgf10-/c;Pax2-Cre/+) samples were counted for the Fgf3/Fgf10/Pax2-Cre conditional cross. N=3 control (Rosa26rtTA/+) and n=3 experimental (Rosa26rtTA/+;tetO-dnFgfR2b/+) samples for the dnFGFR2b cross. pHH3-positive cells per ear were normalized to the cross-sectional area counted. Statistical significance was determined using an unpaired Student’s t-test (Prism software 7.0).
Paint filling of embryonic inner ears
Filling of embryonic inner ears with latex paint and photography was as described previously (Urness et al., 2015).
Induction of dnFGFR2b expression
Initial inductions of dnFGFR2b designed to phenocopy Fgf3/Fgf10 double mutants were achieved by feeding pregnant females DOX chow (200 mg/kg, Custom Animal Diets, LLC) ad libitum for the indicated time periods (E5.5-E10.5, E5.5-E11.5 or E6.5-E11.5). All subsequent inductions to generate samples for paintfilling, RNA-seq, ISH or immunostaining were initiated by a single intraperitoneal injection of the pregnant dam with 0.1 ml/10 g body weight of 0.15 mg/ml (1.5mg/kg body weight) doxycycline hyclate (Sigma-Aldrich) prepared in PBS followed by provision of DOX chow ad libitum for the indicated time periods. We avoided using female Rosa26rtTA parents, as these seemed to require larger and variable amounts of DOX to see phenotypes than when rtTA was contributed by the male parent, presumably because the widespread, ubiquitous expression of rtTA in females served to sequester DOX. We did not measure the time needed to reactivate signaling after DOX withdrawal, but based on studies of the limb (Danopoulos et al., 2013), we expect that signaling resumes after 12-24 hours.
Otic vesicle preparation and RNA isolation
Embyos from timed matings of CD-1 females and Rosa26rtT/ArtTA;Tg(tetO-(s)dnFgfr2b)/+ males, with DOX exposures as specified, were dissected and the yolk sacs saved for genotyping. The otic vesicles, including surrounding mesenchyme, were crudely dissected from the head. Isolation of the vesicles free of mesenchyme was accomplished similarly to methods previously described (Urness et al., 2010) with the following modifications. Otocysts with adherent mesenchyme were incubated in 50 μl ice-cold PT solution (25 mg/ml pancreatin (Sigma), 5 mg/ml trypsin (Sigma), and 5 mg/ml polyvinylpyrrolidone MW360 (Sigma) in Tyrode’s solution) for ~7 min. (E10.25), or ~8 min. (E11.25) to promote separation of the mesenchyme. Otocysts were aspirated to Hepes-DMEM-10% FBS, where the digested mesenchyme could be gently teased from the underlying epithelium using fine forceps or tungsten needles, and by “rolling” the vesicle over the bottom of the dish to detach the mesenchyme as it adhered to the plastic. The two otocysts from each embryo were aspirated into 100 μl RNALater (Ambion) and stored at -20°C prior to genotyping. For each of four pregnant females per DOX induction regime, all otocysts of the same genotype were combined into paired control (Rosa26rtTA/+) and experimental (Rosa26rtTA/+;Tg-(tetO-(s)dnFgfr2b)/+) pools (n = 6-12 otocysts/pool).
Total RNA from each control and experimental otocyst pool was prepared using a Micro RNAeasy kit (Qiagen #74004) and analyzed for quantity and quality on a BioAnalyzer RNA TapeStation. All 24 samples (2 genotypes x 4 females x 3 DOX exposures) exceeded a RIN quality control number of 8.
RNA-Seq and bioinformatics
RNA library preparation, sequencing and analyses were conducted by the University of Utah/Huntsman Cancer Institute High-Throughput Genomics and Bioinformatic Analysis Shared Resource. Each RNA library was prepared using a TruSeq Stranded mRNA Sample Prep kit (Illumina) with oligo(dT) selection. 50-cycle single read sequencing of each library was conducted on an Illumina Hi-Seq 2500. Sequencing reads were aligned to mm10 + splice junctions (Ensembl build 74) using Novoalign (v2.08.03). Spliced alignments were converted back to genomic space, sorted and indexed using USeq (v8.8.8) SamTranscriptomeParser. Normalized coverage tracks (coverage per million mapped reads) were generated using USeq Sam2USeq and USeq2UCSCExe. Read counts for each gene were generated using USeq DefinedRegionDifferentialSeq (Nix et al., 2008) and differential expression analysis was performed using DESeq2 (Love et al., 2014).
To inspect Fgfr splicing we merged each set of control and dnFgfr2b alignments to separate .bam files, uploaded them to IGV 2.4.10 (Robinson et al., 2011; Thorvaldsdottir et al., 2013) and generated Sashimi plots. To identify significantly regulated pathways (P<0.05, Fisher’s Exact Test), all differentially expressed genes were loaded into Ingenuity Pathway Analysis (QIAGEN Inc., https://www.qiagenbioinformatics.com/products/ingenuitypathway-analysis). For GSEA analysis, two custom gene sets based on human hearing loss genes from Nishio et al. (2015) and mouse inner ear genes from Ohlemiller et al. (2016) were loaded into the Broad Institute GSEA website (Subramanian et al., 2005) and compared to ranked lists of otocyst genes sorted by fold change from DESeq2.
COMPETING INTERESTS
No competing interests declared.
FUNDING
Funded by grants from the National Institute of Health to SLM (R01 DC011819 and R01 DC002043). EG-M was supported by an SDB CHOOSE Development! fellowship.
DATA AVAILABILITY
The three RNA-seq datasets are deposited in GEO under accession #GSE116404.
ACKNOLWLEGMENTS
We thank Katia Hatch for auditory testing of F3cKO mice, Leslie Slota for unpublished work on marker gene expression in F3cHet/F10cKO cochleae, Saverio Bellusci and Denise Al-Alam for transferring Rosa26rtTA and Tg(tetO-dnFgfr2b) mice, Mario Capecchi for transferring Sox1Cre mice, Tim Mosbrugger and Chris Stubben for bioinformatics support, Shannon Odelberg for help with Prism and Gary Schoenwolf for experimental and editorial advice.