Thyroid hormone increases fibroblast growth factor receptor expression and disrupts cell mechanics in the developing organ of corti
© Szarama et al; licensee BioMed Central Ltd. 2013
Received: 22 August 2012
Accepted: 29 January 2013
Published: 9 February 2013
Thyroid hormones regulate growth and development. However, the molecular mechanisms by which thyroid hormone regulates cell structural development are not fully understood. The mammalian cochlea is an intriguing system to examine these mechanisms, as cellular structure plays a key role in tissue development, and thyroid hormone is required for the maturation of the cochlea in the first postnatal week.
In hypothyroid conditions, we found disruptions in sensory outer hair cell morphology and fewer microtubules in non-sensory supporting pillar cells. To test the functional consequences of these cytoskeletal defects on cell mechanics, we combined atomic force microscopy with live cell imaging. Hypothyroidism stiffened outer hair cells and supporting pillar cells, but pillar cells ultimately showed reduced cell stiffness, in part from a lack of microtubules. Analyses of changes in transcription and protein phosphorylation suggest that hypothyroidism prolonged expression of fibroblast growth factor receptors, and decreased phosphorylated Cofilin.
These findings demonstrate that thyroid hormones may be involved in coordinating the processes that regulate cytoskeletal dynamics and suggest that manipulating thyroid hormone sensitivity might provide insight into the relationship between cytoskeletal formation and developing cell mechanical properties.
KeywordsYoung’s modulus Hair cell Pillar cell Hypothyroid Cell mechanics
The cytoskeleton plays a key role in modulating the morphological changes of cells and tissues, impacting both the resistance to deformation and the exertion of force by the cell [1, 2]. In the central nervous system, thyroid hormones are thought to act as master regulators of tissue growth and cytoskeletal development [3, 4]. Thyroid hormone action can be mediated through traditional genomic mechanisms such as the regulation of transcription [5, 6] in association with co-activators and co-repressors . However, a direct transcriptional regulation of cytoskeletal elements by thyroid hormone receptors has not been described. This suggests that thyroid hormones might act indirectly by controlling intermediary signaling cascades, which could coordinate the regulatory components of cytoskeletal formation [8, 9].
The sensory epithelium of the mammalian inner ear is an excellent model system to understand basic questions of cytoskeletal organization, as structural elements, such as actin and microtubules, facilitate the organization and architecture of this tissue. The auditory epithelium of the inner ear contains one row of inner hair cells and three rows of outer hair cells (OHCs), which detect and amplify sound, and several types of non-sensory supporting cells, which withstand the mechanical stress of sound vibrations. In particular, supporting pillar cells (PCs), which form the fluid-filled tunnel of Corti , have a highly organized cytoskeletal network composed primarily of microtubules [11, 12]. Disruptions in the formation of this tunnel have been shown to negatively impact hearing function [13–15]. Therefore, it is imperative that we understand the regulatory cues that facilitate normal structural development.
There is reason to suggest that thyroid hormone signaling has an important role in development of the cochlea. Both the sensory and supporting cells in this epithelium have unique expression patterns of thyroid hormone receptors [16, 17], and deiodinase enzymes [18, 19], which control the availability of thyroid hormone. More recently, thyroid hormone transporters have also been localized to this sensory epithelium , and the developmental expression profile contributes some control over the accessibility of ligand to the cell-specific targets. Consistent with this hypothesis, deletion of all known thyroid hormone receptors leads to hearing impairment and inner ear cell structural defects, including malformation of cells in the organ of Corti, and a collapsed tunnel of Corti . However, the factors that mediate development of the inner ear structures are largely unknown. In the cochlea, cytoskeletal formation, including the actin-based cuticular plate in OHCs  and the dense 15-protofilament microtubule network in PCs  that is mostly acetylated , follows cell differentiation [10–12]. However, the signaling pathways mediating cell structural development are not well understood.
Interestingly, mutations in some components of the Fibroblast growth factor (Fgf) signaling pathway cause inner ear structural defects that appear similar to those of the hypothyroid phenotype. In particular, mutations in Fgf receptor 3 (Fgfr3) lead to disruptions in tissue structure, including a collapsed tunnel of Corti, and cause auditory defects in both mice and humans [13, 22–24]. These disruptions have been attributed to both a lack of differentiation of PCs and malformation in PC microtubule development . However, there must also be some regulation of Fgf-receptor expression. One possibility that has been observed in other organ systems is the impact of thyroid hormone regulation of the Fgf-signaling pathway. For example, thyroid hormone stimulates Fgfr expression in undifferentiated cartilage [25, 26]. Furthermore, a thyroid hormone response element is present in the promoter region of Fgfr1 . Together, these data suggest that Fgf-signaling could act as an intermediary between thyroid hormone signaling and cytoskeletal development, and motivate further examination of thyroid hormone action specifically in the cochlea.
In this study, we examined the mechanism of thyroid hormone action on inner ear structural development. We found that hypothyroidism led to higher mRNA expression of Fgf-receptors relative to controls, leading to a delay in the down-regulation of Fgfr3. Hypothyroidism also led to delayed OHC and PC differentiation, which may be mediated in part by the aberrant expression of Fgf-receptors. Finally, hypothyroidism disrupted OHC and supporting PC structure, and aberrantly stiffened embryonic and early postnatal epithelial cells. Here, we show that the hypothyroid-induced cell stiffening may be mediated in part by the disrupted phosphorylation of Cofilin, which has the potential to alter actin dynamics.
Thyroid hormone levels regulate Fgfr expression in the cochlea
In order to localize the increase in expression of these receptors in the cochlea, we examined Fgfr mRNA expression by in situ hybridization in control and hypothyroid cochleae. At P0, Fgfr1 is localized to cell populations in both the greater epithelial ridge, a collection of cells medial to the sensory epithelium that gives rise to the inner sulcus , and the lesser epithelial ridge, which is located lateral to the sensory epithelium and gives rise to the spiral ligament  (Figure 1C). In hypothyroid conditions, expression of Fgfr1 appeared more intense at P0 relative to controls at basal and apical regions of the cochlear duct (Figure 1C). Expansion of the expression domain of Fgfr1 was not observed. In contrast with Fgfr1, Fgfr3 is normally initially expressed broadly within the sensory epithelium in progenitors that will give rise to both hair cells and supporting cells , but by P0 is down-regulated in sensory OHCs of the more mature basal region of the cochlea and maintained in non-sensory supporting pillar and Deiter’s cells (Figure 1D). At later developmental time points, Fgfr3 expression in the apex resembles expression in the base . In hypothyroid conditions, Fgfr3 expression in the base persisted in OHCs at P0, indicating a delay in down-regulation (Figure 1D). Taken together, these results show that hypothyroidism leads to a delay in development of the inner ear, and suggest that there may also be a delay in differentiation of both OHCs and PCs at postnatal stages.
To further assess postnatal PC development, expression of S100-A1 protein was labeled and compared between cochleae in hypothyroid and control conditions. S100 proteins are initially highly enriched throughout the cochlear duct at embryonic stages , and are later down-regulated in differentiated PCs after the first postnatal week. While S100-A1 immunofluorescence intensity appeared to be lower in hypothyroid than in control conditions at P0, by P3, S100-A1 immunofluorescence appeared higher in hypothyroid than in control conditions (Figure 2B), which is consistent with the developmental delay observed in hypothyroid cochleae . To quantify this difference, measurements of relative fluorescence intensity (mean ± s.e.m. A.U.) were calculated and compared between hypothyroid and control conditions. In PCs, S100-A1 intensity decreased from 987 ± 119 A.U. in control conditions to 319 ± 116 A.U. in hypothyroid conditions at P0, as was the case for the OHCs (Figure 2C; p-value < 0.05). Later, at P6, expression of S100-A1 was significantly increased from 609 ± 48 A.U. in controls to 1257 ± 18 A.U. in hypothyroid conditions (Figure 2C; p-value < 0.01) showing that disruptions in these cells persist at later postnatal stages. Additionally, the relative fluorescence intensity of phalloidin in hypothyroid OHCs and PCs showed about a three-fold increase relative to control conditions (Figure 2C; p-value < 0.05).
Finally, to assess late postnatal differentiation of PCs, we examined the relative immunofluorescence of CD44, a protein that is restricted to supporting PCs at P0, increases in intensity through P7  and is reported to be responsive to changes in thyroid hormone levels in the cerebellum . Confocal micrographs of cochlear cross-sections at P6 showed that CD44 fluorescence appeared lower in hypothyroid conditions relative to controls (Figure 2D), suggesting that PCs maintain a sensitivity to thyroid hormone levels during postnatal development that may contribute to cell maturation beyond the effects of Fgfr3. Taken together, these data suggest that hypothyroidism leads to a delay in the timing of PC maturation, as early markers of development that are normally down-regulated persist and late markers of development are delayed under experimental conditions. Additionally, the corresponding delay in down-regulation of Fgfr expression also suggests that some of these early effects are mediated through prolonged Fgfr expression.
Hypothyroidism disrupts microtubule formation in the cochlea
Hypothyroidism stiffens outer hair cells and supporting pillar cells
To examine the effects of hypothyroidism on the mechanical properties of developing PCs, we also calculated average Young’s modulus for PCs in control and hypothyroid conditions. We found that in controls, PC Young’s modulus was approximately five-fold higher between E16 and P5 (Figure 5C). This corresponds to increased microtubule acetylation  and suggests that increased cytoskeletal formation leads to cell stiffening. In contrast with controls, hypothyroid PCs were significantly stiffer at E16 and P0 (Figure 5C; p-value < 0.05). However by P5, hypothyroid PCs showed significantly reduced Young’s modulus relative to controls (Figure 5C; p-value < 0.05). In summary, these results indicate that hypothyroidism leads to a stiffening of OHCs and PCs early in development, and that at later time points hypothyroid PCs fail to develop mature mechanical properties.
Actin is responsible for the aberrant increase in pillar cell and outer hair cell stiffness
Cofilin activity mediates changes in actin dynamics under hypothyroid conditions
Effects of thyroid hormone signaling in the inner ear may be regulated through prolonged Fgf-signaling
Our data support previous findings that thyroid hormone levels coordinate the timing of development in many target tissues, including the skeletal  and central nervous systems [61, 62]. However, the specific molecular pathways that mediate the effects of thyroid hormone are largely still unknown. Thyroid hormones primarily regulate transcriptional activity through thyroid hormone receptors (reviewed in ). Based on similarity in phenotypes and existing data from other systems, we examined Fibroblast growth factor receptors after altering thyroid hormone levels. In the organ of Corti, decreased levels of thyroid hormone were shown to result in prolonged expression of Fgfr1 and Fgfr3, respectively (Figure 1), suggesting that thyroid hormone receptors negatively regulate Fgf-receptor expression in the developing cochlea. Interestingly, Fgfr1 has been shown to accelerate differentiation when inactivated in differentiating osteoblasts  and a thyroid hormone response element has been identified in the promoter region of Fgfr1. In the organ of Corti, it seems that Fgfr3 regulates the timing of hair cell and supporting cell development as shown by experiments in which ectopic activation of Fgfr3 delays differentiation of supporting cells [23, 32]. The observation of similar delays in the down-regulation of p75ntr and S100 proteins (Figure 2) in differentiating cells in hypothyroid conditions is consistent with the idea that the maintenance of Fgfr3 signaling mediates at least some of the effects observed in hypothyroidism of the cochlea. However, these data do not rule out an additional layer of regulation by thyroid hormones on Fgf-signaling. Indeed, thyroid hormone has been shown to play a role in heparin sulfate expression in the developing growth plate , which suggests an additional pathway, through which thyroid hormone could enhance Fgf-signaling. Overall, these findings suggest that the timing of down regulation of Fgf-signaling is important for the developing inner ear. However, the persistent expression of both Fgfr3 and of Fgf8, the most likely cognate ligand for Fgfr3 in the organ of Corti, into adult stages, suggests that Fgf-signaling may also have a continuous role in inner ear tissue morphology. While it seems reasonable to suggest that the mechanism to decrease Fgfr3-signaling would be to decrease the presence of ligand [66, 67], we propose that thyroid hormones might also mediate Fgf-signaling at this stage of inner ear development by regulating the level of Fgf-receptor expression in the organ of Corti.
Using the drug methimazole, hypothyroidism was induced in utero to examine the cellular response to a lack of thyroid hormone in the organ of Corti. Thyroid hormone levels in utero are thought to be primarily from a maternal source and transmitted through the placenta [68, 69]. Methimazole transmission has also been shown to cross the placenta  and to reduce the serum levels of thyroid hormone . While the effectiveness of this drug has led to widespread use as a treatment in clinical cases of hyperthyroidism for adults and, in particular, during pregnancy , methimazole is not without side effects. In less than 1% of human subjects, agranulocytosis, a lowered white blood cell count, and liver toxicity resulting in jaundice were reported [73, 74]. However, considering the relatively limited time course of methimazole treatment in these studies, it seems unlikely that the treatment led to these side effects.
Thyroid hormone may disrupt cytoskeletal formation through prolonged Fgf-signaling
A key observation of the experiments described here was the alteration in cytoskeletal formation in response to hypothyroidism. While disruption of the cytoskeleton in cases of hypothyroidism has long been known [3, 7, 42], the molecular basis for these effects is not yet fully understood. The results presented here suggest that Fgf signaling may act as an intermediary, with prolonged Fgf-signaling leading to disruptions in microtubule formation. In the case of microtubule structure, we extend existing research by providing two complementary mechanisms for this hypothyroid-induced structural defect. First, deletion of Fgfr3 leads to decreased β-tubulin , further supporting the relationship between Fgf-signaling and microtubule number. Second, the increase in S100-A1 in PCs is representative of elevated intracellular calcium levels [75, 76], which could directly impair microtubule formation and is observed in Fgfr3-deficient mice .
In addition to the microtubule disruptions in PCs (Figures 3 and 4), there also appeared to be a higher fluorescence intensity of phalloidin-labeled actin in hypothyroid conditions relative to controls. Phalloidin, a member of the phallotoxin group of F-actin binding peptides, has been used to quantify the amount of F-actin in cells [78, 79]. This probe has a diameter of 12–15 Å , which has the advantages of low steric hindrance to bind close to a 1:1 ratio  and therefore does not compete with actin binding proteins . This probe also has a similar affinity for both large and small filaments , but does not bind monomeric G-actin. This is a limitation of the technique as it is not possible with only one probe to examine whether or not the ratio of F-actin-to-G-actin has been altered in hypothyroid relative to control conditions. However, the advantages of this probe support our conclusion that stronger phalloidin fluorescence intensity, combined with electron micrographs showing no apparent difference in cuticular plate thickness, suggests that there is a higher density of actin, which may cause in part measured stiffening at the lumenal surface of the cochlea.
There is increasing evidence that Fgf-signaling mediates actin dynamics through signaling , which may be the key link between hypothyroidism and the observed cell structural defects. Indeed, Fgf-signaling is necessary for the dynamic changes in actin remodeling that lead to invagination of the developing otocyst . In the hypothyroid cochlea, Fgf-receptors might mediate actin dynamics through activation of p21-activated kinases (PAKs) or ROCK [85, 86] that can then phosphorylate LIMK to inhibit Cofilin activity via phosphorylation [87, 88]. The prolonged Fgfr-expression shown in hypothyroid conditions coincides with a decrease in phosphorylation of Cofilin (Figure 7), which is consistent with the role for this protein in the rate of actin filament turnover. Since both increased actin filaments and decreased Cofilin phosphorylation were observed in hypothyroid cochleae, these results are consistent with this proposed mode of regulation. This might in part explain the disruption to OHC morphology (Figure 3), which is interesting given that recently it has been proposed in other developing tissues that the rearrangement of actin filaments generally would produce a disruption in cell morphology . Additionally, disrupted p75ntr expression has also been shown to mediate actin dynamics through down-regulation of Rho-GTP signaling . Since p75ntr expression appeared to persist in hypothyroid conditions at P3 (Figure 2), the contribution of Fgf-signaling may also contribute indirectly through a p75ntr-dependent mechanism.
Alterations in cytoskeletal dynamics may affect cell mechanical properties
Ultimately, impaired functional ability of supporting PCs to resist deformation at P5 (Figure 5) is not exactly concomitant with the decrease in microtubule number observed in electron micrographs of hypothyroid pillar cells relative to controls. However, it is possible that the lack of acetylation observed at P5 (Figure 4), and not absolute microtubule number, contributes to the formation of the tunnel of Corti by the pillar cells. It is also possible that a lack of acetylation has other effects on microtubule length and polymerization [91, 92]. Future studies directly disrupting acetylation, while also controlling for microtubule length and protofilament number, will add important in vitro data that can be brought back into microtubule-based cell structural assays to further support what has been shown here.
In the case of actin structure, the findings that hypothyroid OHCs and PCs are aberrantly stiffer than controls (Figure 5) and are increasingly sensitive to Latrunculin A (Figure 6) implicate the actin cytoskeleton, rather than microtubule development, in hypothyroid-induced cell stiffening observed from E16 to P3. In this study, atomic force microscopy (AFM) has been used to probe cytoskeletal structures in living cells without membrane disruption or fixation. While the cell membrane is an integral part of stabilizing the cytoskeleton, previous results have shown that membrane stiffness is negligible in eukaryotic cells relative to cytoskeletal stiffness [93, 94]. Furthermore, with increasing indentation depth—in this study indentation was 1.5 μm—the bulk of Young’s modulus has been shown to be the result of cytoskeletal stiffness , and in the case of the developing cochlea, reflects the stiffness of the actin mesh at the cell apex. Indeed, by combining AFM with live-cell fluorescence imaging techniques, it is possible to get single cell specificity. For example, after treatment with the cytoskeletal disrupting agent Latrunculin A, the OHC but not the neighboring PC Young’s modulus was decreased in cochlear explant cultures. Overall, these AFM data suggest that the calculated Young’s modulus reflects cell surface mechanical properties, which are dominated by cytoskeletal components.
Cell-specific responses to thyroid hormone may lend insight into cell structural development
While data presented here were only able to measure the consequences of disrupted actin dynamics in hair cells and supporting PCs, disruptions to actin dynamics could have broad consequences for many supporting cell types in the developing organ of Corti. For example, the greater epithelial ridge is composed of columnar epithelial cells that undergo structural changes leading to the formation of the fluid filled space known as the inner sulcus . While we know that these cells support synapse formation of sensory inner hair cells in the organ of Corti [96, 97], the aberrant cell cytoplasm in this region may explain part of the pathology of hypothyroidism. By localizing phosphorylated Cofilin expression in control and hypothyroid cochlear cross-sections (Figure 8), we observed not only the aberrant defect in the sensory epithelium, but also the cell-specific differences in phosphorylated cofilin expression throughout the cochlear duct. Cofilin phosphorylation has also been observed to mediate actin dynamics in a number of developing systems [98, 99]. The relationship between hypothyroidism and cell-structural defects in the developing cochlea might prove to be the basis for additional studies of morphogenesis in developing systems that rely on coordination by thyroid hormones [60, 62]. Future studies also examining actin dynamics under reduced thyroid hormone levels could lend insight into a mechanism by which developmental malformations to the structure and function of the organ of Corti may be prevented.
We find that hypothyroidism leads to a delay in the down-regulation of Fgf-receptors and to a decrease in microtubule formation and acetylation at early postnatal stages. While the lack of microtubules ultimately reduce supporting pillar cell stiffness, we find that hypothyroidism actually stiffens outer hair cells at P3 and pillar cells at E16 and P0 as a result of increased F-actin. Our data also show that the increased dynamics of actin in these cells might be the result of hypothyroid-induced Fgf signaling and a decreased phosphorylation of Cofilin. Together, these data implicate the sensitivity of cell structural development to thyroid hormones, and suggest that thyroid hormone signaling might coordinate the time-course of tissue morphogenesis.
Cochlear explant cultures and pharmacological treatments
Cochleae of Institute for Cancer Research (ICR) mice (Charles River Laboratories, Frederick, Maryland, USA) were cultured at specific stages between E16 and P5 as previously described  and plated on No.1 glass coverslips (Corning, New York, USA). To increase thyroid hormone signaling, explants were treated with either 5 μM triiodothyronine or 5 μM reverse triiodothyronine (Sigma, St. Louis, Missouri, USA), an inactive form of thyroid hormone, as a control, in DMSO. To decrease thyroid hormone signaling, animals were treated with 0.02% methimazole (Sigma) and 10% sucrose in drinking water and low iodine feed administered ad libitum. All animal care and procedures were approved by the Animal Care and Use Committee at NIH and complied with the NIH guidelines for the care and use of animals.
Transmission electron microscopy
Cochleae from 3 control and 3 hypothyroid animals at P3 were isolated and placed immediately in 0.1 M phosphate buffer (pH 7.4) containing 4% paraformaldehyde and 2% glutaraldehyde for 30 minutes at room temperature followed by 2 hours at 4°C. The inner ears were then washed in phosphate buffer and cacodylate buffer, post-fixed with 1% osmium tetroxide, and dehydrated through a graded alcohol series before being embedded in Poly/BED 812 resin (Polysciences Inc., Warrington, Pennsylvania, USA) as previously described . Thin sections of about 75 nm were cut using a Leica Reichert ultramicrotome, stained with lead citrate, mounted on 200-Cu mesh grids, and examined at room temperature using a JEOL transmission electron microscope (Akishima, Tokyo, Japan) at 80 kV with 15,000× magnification. Images were acquired using a Hamamatsu Camera (Hamamatsu Photonics K.K., Japan) and Advanced Microscopy Techniques Camera System software version 534.4 (Woburn, Massachusetts, USA). Images were cropped in Adobe Photoshop CS4 (Adobe). To quantify morphological changes in OHCs and PCs, transverse sections from three animals for each condition were analyzed with ImageJ  analysis software for the presence of microtubules at distances 2, 4, and 6 μm from the lumenal surface of PCs. Measurements of OHC length and width were also made. Analysis of statistical significance was determined using student’s T-test.
Immunohistochemistry and confocal image analysis
Samples of the same time point were harvested on the same day under the same conditions, fixed in 4% paraformaldehyde for 4 hours, rinsed, passed through an increasing sucrose gradient, and embedded in OCT (Sakura, New York, USA) in parallel. All sections to be examined from the same time point for comparison were cryosectioned at 12 μm thickness on the same day. Sections were processed in parallel with the same stock solutions, which included permeabilization with 0.2% Tween-20 in PBS, blocking with 10% normal horse serum and incubation overnight in primary antibody (p75ntr, Covance, Princeton, New Jersey, USA, 1:1000; ZO-1, Millipore, 1:1000, S100-A1, Neomarkers, Kalamazoo, Michigan, USA, 1:500; CD44, BD Pharmigen, Franklin Lakes, New Jersey, USA, 1:200; p-Cofilin, Santa Cruz Biotechnology, Santa Cruz, California, USA, 1:500 ) at 4°C. Primary antibodies were detected using either Alexa Fluor 488 or 546 (Invitrogen, 1:1000) conjugated secondary antibodies. Directly conjugated Phalloidin 633 (Invitrogen, 1:5000) was applied to all samples. Cochleae prepared for whole mount immunohistochemistry were fixed for 2 hours and were labeled with a primary antibody against acetylated tubulin (Sigma, 1:750). Samples were mounted in Fluoromount-G (Southern Biotech, Birmingham, Alabama, USA), using No. 1Â½ glass coverslips (Corning, New York, USA) adhered with nail polish. All fluorescence images were acquired at room temperature with LSM 510 acquisition software as 12 μm Z-stacks with 1 μm optical sectioning using a Zeiss 510 LSM Confocal Microscope with 40X oil objective [1.3 numerical aperture (NA); Plan-Neofluar]. Using line scan analysis, fluorescence intensity was measured from projected Z-stacks with ImageJ  analysis software in 2 μm areas at the lumenal surfaces of both PCs and OHCs, as this region was previously observed with Transmission Electron Microscopy to maintain a homogeneous cell cytoplasm from E16 through P5 . Average fluorescence intensity (mean ± s.e.m. Arbitrary Units) was calculated from 6 samples, and compared using student’s T-test.
In situ hybridization
Cochleae from 6 animals per condition—hypothyroid or control—were harvested on the same day at the same time point and fixed in a stock solution of 4% paraformaldehyde in 1X PBS overnight. Samples were then rinsed, passed through an increasing sucrose gradient, and embedded in OCT (Sakura, New York, USA) in parallel using the same solutions under the same conditions. Samples were cryosectioned in parallel at 12 μm thickness. All steps of the in situ hybridization were carried out in parallel on 6 sections from 6 animals per condition as previously described  with probes specific to Fgfr1 and Fgfr3. All samples had the same exposure to the same reaction mixture, as previously described  for the same length of time.
Quantitative real-time polymerase chain reaction (qPCR)
Cochleae were removed from the temporal bone, and the surrounding mesenchyme, scala vestibuli and scala tympani were removed. 6–8 cochleae were pooled and total RNA was isolated using RNAqueous (Ambion, Grand Island, New York, USA) reagents. cDNA was synthesized from 500 ng total RNA for each condition using a Superscript III first strand synthesis kit (Invitrogen). Amplification was performed with SYBR Green (Applied Biosystems, Foster City, California, USA). Amplification of all mRNA was performed on an ABI Prism 7000 (Applied Biosystems) with the following cycling conditions: 40 cycles of 95°C for 15 seconds, and 60°C for 1 minute. To calculate fold change, gene expression was normalized to glyceraldehyde 3-phosphate dehydrogenase (GAPDH) and statistical significance was confirmed with student’s T-test. Primer sequences used are as follows: Fgfr1,F 3′-ATGGTTGACCGTTCTGGAAG-5′; Fgfr1,R 3′-AGAAAAGGGTACGCAGCAGA-5′; Fgfr3,F 3′-GAGACTTGGCTGCCAGAAAC-5′; Fgfr3, R 3′-GGGCTCACATTTGTGGTCTT-5′, GAPDH, F 3′-ATCCTGTAGGCCAGGTCATG-5′; GAPDH, R 3′-TATGCCCGAGGACAATAAGG-5′.
Western blot analysis
Cochlear explants were freshly isolated before total protein extraction. Proteins were extracted from 8–10 cochleae in RIPA buffer containing complete mini protease inhibitor cocktail (Roche), complete phosphatase inhibitor cocktail (Roche), 1 mM Na3VO4, and 500 mM NaF. Protein was quantified using the Dc protein assay kit (Bio Rad) and Lowry method using a ND-1000 Spectrophotometer (Nanodrop, Wilmington, Deleware, USA). 25 μg total protein per condition was loaded onto 4-12% SDS-PAGE gels (Invitrogen) run for 2 hours at 120 V, transferred to nitrocellulose membrane (Invitrogen), and run for 3 hours at 80 V at 4°C. Membranes were blocked in 0.05% TBS-T containing BLOTTO (Rockland, Gilbertsville, Pennsylvania, USA) and 1% BSA (Sigma). Primary antibodies were incubated in blocking solution at the following concentrations: p-Cofilin (1:500, Abcam); Cofilin (1:5000, Abcam). Primary antibodies were conjugated to horseradish peroxidase anti-rabbit secondary antibody (1:5000, Amersham) and detected using ECL Detection Reagents (Amersham). Membranes were visualized using Image Station 4000R (KODAK) and Carestream Molecular Imaging Software (New Haven, Connecticut, USA). Image analysis was performed with ImageJ  using the Gel Analysis plug-in method to calculate relative density. Relative density of phosphorylated protein signal was normalized to β-actin (Sigma, 1:5000) signal for the same tissue and under the same experimental conditions. Statistical significance was determined with Welch’s T-test .
Atomic force microscopy and live cell imaging
Experiments were performed using a Bioscope II and Bioscope Catalyst AFM (Bruker, Santa Barbara, California, USA) head mounted onto a Zeiss Axiovert 200 inverted microscope and controlled via a Nanoscope V controller. Pyramidal shaped, gold-coated, silicon nitride cantilever probes with 0.03 N/m spring constant (Bruker) were used for all measurements. Cells were identified after being loaded with 500 nM Calcein AM vital dye (Invitrogen) in Leibovitz’s media (Invitrogen) for 30 minutes and rinsed with fresh Leibovitz’s media. Contact mode AFM was applied to all samples in Leibovitz’s media at 1.5 μm maximum indentation using 3 μm ramps at 1 Hz continuous force ramping to collect 3 force-distance curves for the center of each identified hair cell or supporting cell. All force-distance curves were analyzed with custom analysis software in MATLAB (Mathworks, Natick, Massachusetts, USA) and fit to the Sneddon model  to measure Young’s modulus, which is a material property of the cellular resistance to deformation and was calculated with the formula F = (2/πtanα)(E/(1-ν2))δ2; where F is applied force, ν is Poisson’s ratio and assumed to be 0.5, δ is cantilever indentation, and α is cantilever tip angle. Average Young’s modulus (mean ± s.e.m. kPa) was calculated from the average of sample measurements of 10 cells within a given region of interest such as the base or the apex of the cochlea. Statistical analyses were performed using Welch’s T-test .
Outer hair cells
Fibroblast growth factor
Fibroblast growth factor receptor 3
Neurotrophin receptor p75
Embryonic day 16
Postnatal day 0
Rho-associated protein kinase.
We thank Ya-Xian Wang for help with tissue preparation for TEM, and S. Raft, D.S. Sharlin, and A. Fridberger for comments on earlier versions of this manuscript. This research was supported by the National Institute on Deafness and Other Communication Disorders (NIDCD) Intramural Research Program [DC000059 to M.W.K., DC00003333 to R.S.C.] and in part with an Intramural Fellowship to Promote Diversity, from the NIH Office of the Director to K.B.S.
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