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. Author manuscript; available in PMC: 2012 Dec 13.
Published in final edited form as: J Neurosci Res. 2010 Jun;88(8):1812–1821. doi: 10.1002/jnr.22333

Transcriptional changes in adhesion-related genes are site-specific during noise-induced cochlear pathogenesis

Qunfeng Cai 1, Minal Patel 1, Donald Coling 1, Bo Hua Hu 1,*
PMCID: PMC3521497  NIHMSID: NIHMS410934  PMID: 20091770

Abstract

Cell–cell junctions and junctions between cells and extracellular matrix are essential for maintenance of the structural and functional integrity of the cochlea, and are also a major target of acoustic trauma. While morphological assessments have revealed adhesion dysfunction in noise-traumatized cochleae, the molecular mechanisms responsible for adhesion disruption are not clear. Here, we screened the transcriptional expression of 49 adhesion-related genes in normal rat cochleae and measured the expression changes in the early phases of cochlear pathogenesis after acoustic trauma. We found that genes from four adhesion families, including the immunoglobulin superfamily and the integrin, cadherin, and selectin families, are expressed in the normal cochlea. Exposure to an intense noise at 120 dB sound pressure level (SPL) for 2 h caused site-specific changes in expression levels in the apical and the basal sections of the sensory epithelium. Expression changes that occurred in the cochlear sensory epithelium were biphasic, with early upregulation at 2 h post-noise exposure and subsequent downregulation at 1 day post-exposure. Importantly, the altered expression level of seven genes (Sgce, Sell, Itga5, Itgal, Selp, Cntn1 and Col5a1) is related to the level of threshold shift of the auditory brainstem response (ABR), an index reflecting functional change in the cochlea. Notably, the genes showing expression changes exhibited diverse constitutive expression levels and belong to multiple adhesion gene families. The finding of expression changes in multiple families of adhesion genes in a temporal fashion (2 h vs. 1 day) and a spatial fashion (the apical and the basal sensory epithelia as well as the lateral wall tissue) suggests that acoustic overstimulation provokes a complex response in adhesion genes, which likely involves multiple adhesion-related signaling pathways.

Keywords: Cochlea, Sensory epithelium, Hair cells, Acoustic trauma, Adhesion, Transcriptional expression, Noise

Introduction

Acoustic overstimulation is a common cause of acquired sensorineural hearing loss in the adult population. There is considerable evidence that functional loss of the cochlea is not simply a consequence of the initial mechanical destruction of the cochlea, but is also attributed to the development of complex secondary events that contributed to early as well as delayed cell damage. Gaining an understanding of the molecular mechanisms underlying cochlear damage is an important step in developing therapeutic strategies for the prevention of noise-induced hearing loss.

Cell adhesion is essential for maintenance of tissue integrity. Excessive movement of the basilar membrane of the cochlea due to acoustic overstimulation stretches the organ of Corti and compromises both cell–cell junctions and adhesion of cells to extracellular matrix. Morphological evidence for structural disruption of intercellular junctions has been observed in animal models of acoustic trauma (Hamernik et al., 1984; Henderson and Hamernik, 1986; Lim and Melnick, 1971; Saunders et al., 1985; Thorne et al., 1984), while alteration in the barrier function of the reticular lamina due to cell junction dysfunction has also been observed (Hu and Zheng, 2008).

Cell adhesion in the mammalian organ of Corti relies on the function of tight junctions, gap junctions, adherens junctions, and desmosomes (Gulley and Reese, 1976; Kikuchi et al., 2000; Nadol, 1978; Raphael and Altschuler, 1991), as well as focal adhesions that form junctions between cells and extracellular matrix (Jamesdaniel et al., in press; Littlewood Evans and Muller, 2000; Meyer zum Gottesberge et al., 2008; Tian et al., 2009). These junctions work coherently to maintain functional and structural integrity of tissues. Gap junctions are intercellular channels that facilitate direct communication of ions and small cytoplasmic molecules. In the cochlear sensory epithelium, gap junctions are distributed between supporting cells (Forge et al., 1999; Kikuchi et al., 1995), and mutation of gap junction genes has been linked to congenital hearing loss (Nickel and Forge, 2008). The tight junction is responsible for establishing compositionally distinct compartments by forming tight seals between cells (Anderson and Van Itallie, 1995; Schneeberger and Lynch, 1992). In the cochlea, the tight junction is important for maintenance of the barrier function of the reticular lamina (Gulley and Reese, 1976), and mutation of tight junction-associated genes causes hearing loss(Riazuddin et al., 2006; Wilcox et al., 2001). The adherens junction is present in both sensory cell and supporting cell junctions (Leonova and Raphael, 1997; Nunes et al., 2006). In the reticular lamina, adherens junction proteins are intermingled with the tight junction proteins, forming a hybrid tight junction that encircles the cells (Nunes et al., 2006). Adherens junction proteins also play an important role in maintaining the structural integrity of the cytoskeleton. Alteration of adherens proteins, such as cadherins, leads to disruption of the actin structure of cells (Tsukita et al., 1992).

Thus far, only a handful of adhesion-related molecules have been identified and their distributions in the cochlea documented (Davies and Holley, 2002; Simonneau et al., 2003; Suzuki and Harris, 1995; Toyama et al., 2005; Tsuprun and Santi, 1999; Whitlon and Rutishauser, 1990). These molecules belong to multiple adhesion gene families, including the immunoglobulin superfamily and the integrin and cadherin families. The involvement of adhesion molecules has been implicated in scar formation in the organ of Corti during the recovery phases of acoustic injury (Raphael and Altschuler, 1991). A recent study has shown that mice lacking the vezatin protein, a ubiquitous adherens junction protein, exhibit a high vulnerability to acoustic trauma (Bahloul et al., 2009), suggesting that this adherens junction gene regulates sensory cell responses to acoustic overstimulation. Although these studies have linked adhesion genes to noise-induced cochlear pathogenesis, our understanding of adhesion signaling pathways in cochlear pathogenesis remains limited.

In non-cochlear tissues, adhesion proteins have been implicated in mediating signaling pathways in various pathological conditions, including cell detachment (Frisch and Screaton, 2001; Grossmann, 2002; Haga et al., 2008; Malik, 1997). The detachment of cells from their anchorage has been shown to be a trigger for apoptosis (Frisch, 2000). For example, Frisch and Francis (Frisch and Francis, 1994) reported that disruption of normal epithelial cell–matrix interactions in culture resulted in apoptotic cell death. Because noise exposure stretches the basilar membrane and compromises cell–cell junctions, we suspected that acoustic trauma could alter adhesion molecule expression, which in turn could affect cell survival. One unresolved question is which adhesion-related genes participate in the early response of adhesion junctions to acoustic trauma, a possible early event in acute sensory cell death (Hu et al., 2006).

The mammalian cochlea consists of three major partitions: the sensory epithelium, the lateral wall, and the modiolus, each playing a distinct role in maintaining cochlear function. While the modiolus is protected by a bony shell, the sensory epithelium and the lateral wall tissue can sustain direct mechanical insult, and morphological observations have identified the sensory epithelium as the primary site of acoustic injury (Henderson and Hamernik, 1986; Saunders et al., 1985). In the sensory epithelium, the basal and the apical partitions differ in several ways. Anatomically, the basilar membrane is narrower, and sensory cells are smaller in the basal end. Physiologically, the cochlea is organized in a tonotopic manner, with lower frequencies encoded at the apical end and higher frequencies encoded at the basal end. The two partitions of the sensory epithelium differ also in certain biological properties. For example, the expression of certain genes exhibits a basal-to-apical gradient in the sensory epithelium (Weston et al., 2011). Functionally, the basal end exhibits less antioxidant capacity, and is prone to oxidative stress (Sha et al., 2001). Moreover, basal sensory cells exhibit a higher susceptibility to ototoxicity and age-related degeneration (Fechter et al., 1997, Forge and Schacht, 2000 and Sha et al., 2001). Importantly, previous investigations have demonstrated a greater vulnerability to acoustic injury in the basal sensory cells (Bohne et al., 1987; Hu et al., 2002b; Pouyatos et al., 2009; Thorne et al., 1984). These spatial differences in the properties of the sensory epithelium prompted us to investigate site-specific changes in the expression levels of adhesion-related genes.

In the current investigation, we examined the expression patterns of 49 adhesion-related genes in three cochlear partitions (the apical and the basal sections of the sensory epithelium and the lateral wall tissues) in normal and noise-traumatized rat cochleae. This study identified the expression of several adhesion-related genes, many of which have not been reported for cochlear tissues. This study also documents noise-induced differential changes in the expression pattern of the adhesion genes and reveals a significant correlation between the expression levels of certain adhesion genes and loss of the cochlear function. These results provide important clues for the understanding of adhesion signaling pathways in noise-induced cochlear pathogenesis.

Materials and Methods

Animals

Sasco Sprague Dawley rats (2–4 months old, 210–300 g, male and female, Charles River Laboratories, Wilmington, MA) were used in the current study. All animals received a baseline-hearing test. Only the animals with normal hearing sensitivity were included in the study. The procedures involving use and care of the animals were reviewed and approved by the State University of New York at Buffalo Institutional Animal Care and Use Committee.

Experimental Design

All animals received a baseline ABR test, and were then randomly assigned to either a normal control group (without receiving noise exposure) or a noise group. The noise group animals were exposed to intense noise for 2 h. The exposed animals were sacrificed after the final ABR test at either 2 h or 1 day post-exposure. Additional four animals were allowed to survive for 28 days for evaluation of permanent hearing loss. For the animals that were sacrificed at 2 h after noise exposure, one cochlea from each animal was used for assessment of mRNA expression levels of adhesion-related genes and the other cochlea was used for assessment of sensory cell damage. Five additional animals from the noise group were used for the analysis of Cdh1 protein expression. The separate experimental procedures are detailed below.

Noise exposure

A continuous noise (1–7 kHz) for 2 h at 120 dB sound pressure level (re 20 μPa) was used to induce acoustic trauma to the cochlea. The noise signal was generated with a real-time signal processor (RP2.1; TDT, Alachua, FL, USA). The signal was routed through an attenuator (PA5; TDT) and a power amplifier (Crown XLS 202; Harman International Company, China) to a loud speaker (NSD2005-8; Eminence). The noise level was calibrated using a sound level meter (Larson Davis 800 B; Depew, NY, USA), a preamplifier (Larson and Davis; model 825), and a condenser microphone (Larson and Davis; LDL 2559) that was placed at the position of the animals head in the sound field. The rats were individually exposed to the noise in a holding cage to ensure a consistent level of noise exposure.

Auditory Brainstem Response (ABR) test

ABR measurements were conducted pre- and three times post-exposure (2 h, 1 day and 28 days) to determine the hearing sensitivity of the animals. ABRs were recorded as previously described (Hu et al., 2009). Briefly, an animal was anesthetized with intraperitoneal injection of a mixture of ketamine (87 mg/kg) and xylazine (3 mg/kg). The body temperature was maintained at 37.5°C with a warming blanket (Harvard Apparatus). Stainless-steel needle electrodes were placed subdermally over the vertex (noninverting input) and posterior to the stimulated and nonstimulated ear (inverting input and ground) of the animal. ABRs were elicited with tone bursts at 5, 10, 20, 30, and 40 kHz (0.5 ms rise/fall Blackman ramp, 1 ms duration, alternating phase) at the rate of 21/s, which were generated digitally (SigGen; TDT) using a D/A converter (RP2.1; TDT; 100 kHz sampling rate) and fed to a programmable attenuator (PA5; TDT), an amplifier (SA1; TDT), and a closed-field loudspeaker (CF1; TDT). The electrode outputs were delivered to a pre-amplifier (RA4LI and RA4PA; TDT) and then to a medusa base station (RA16BA, TDT). Responses were filtered (100–3000 Hz), amplified (20×) and averaged using TDT hardware and software. These responses were then stored and displayed on a computer. The ABR threshold was defined as the lowest intensity that reliably elicited a detectable response.

Assessment of sensory cell damage

The organs of Corti were isolated for pathological examination 2 h post-exposure using a method that has been described in our previous publication (Hu and Cai, 2010). Propidium iodide (Invitrogen Inc.) was used as a nuclear marker to stain the cochleae for assessment of cell damage. Briefly, 2 h after noise exposure the animal was deeply anesthetized and decapitated. The cochlea (either right or left) was quickly removed from the skull, and perfused with a propidium iodide solution (5 μg/ml in 10 mM phosphate buffered saline, PBS) for 10min. The cochlea was then perfused with 10mM PBS to remove the propidium iodide solution and then fixed with 10% buffered formalin for at least 4 h. After dissection, the organ of Corti was collected and mounted on a slide with an antifade medium (Prolong® Gold antifade reagent, Invitrogen. Inc.).

To confirm that sensory cells showing condensed or fragmented nuclei were dying via the apoptotic pathway, caspase-3 activity, a biological marker of apoptosis, was detected using a fluorescent probe, FAM-DEVD-FMK (APT403; Millipore, Bedford, MA). The basic procedure for caspase-3 labeling has been described in detail in our previous publication (Hu and Cai, 2010). Briefly, the animals were scarified at 2 h post-noise exposure. The cochleae were quickly collected and perfused with approximately 20 μl of the freshly prepared staining solution diluted according to the manufacturer’s instructions. The staining solution remained in the cochlea for 50 min, and then the cochlea was fixed with 10% buffered formalin. After dissection, the organs of Corti were further stained with propidium iodide for 10 min.

The specimens were examined with a fluorescence microscope to identify hair cell lesions. The pathological criteria used for identifying damaged cells have been previously described (Hu et al., 2002b; Yang et al., 2004). Based on the nuclear morphology, the numbers of damaged and missing hair cells were identified and counted. The data were assembled into a cochleogram showing the frequency-place correlation for the rat (Muller, 1991).

mRNA expression levels of adhesion-related genes

We assessed the expression levels of 49 adhesion-related genes using PARN-013A SABiosciences adhesion arrays (Qiagen, Valencia, CA) (see Table 1 for the gene list). Methodology to isolate total RNA, synthesize and pre-amplify cDNA and run qRT-PCR reactions was performed as previously described (Hu and Cai, 2010; Hu et al., 2009). Briefly, the animal was decapitated and either the left or the right cochlea was quickly removed from the skull. The cochlea was perfused with an RNA stabilization reagent (RNAlater; Qiagen, Valencia, CA), and was dissected in the same reagent. Three samples (the apical and the basal sensory epithelium as well as the lateral wall tissue) were collected from the cochlea. The apical sample contained the apical 45% of the sensory epithelium and the basal sample contained the tissue from 45% to 90% from the apex of the sensory epithelium. The lateral wall sample consisted of the lateral wall tissue from the second cochlear turn. Each sample was run separately for qRT-PCR analysis. Total RNA was isolated from each sample using the RNeasy Micro Kit (Qiagen) as per manufacturer’s instruction. The concentration of isolated total RNA was measured with a NanoDrop instrument (NanoDrop 1000, Thermo Scientific). The isolated total RNA was used to generate cDNA and the cDNA was pre-amplified using the RT2 Nano PreAMP cDNA Synthesis Kit (Qiagen). Synthesized cDNA was mixed with RT2 Real-Time PCR SYBR Green/Fluorescein Master Mix (Qiagen) and transferred to a 96-well plate. qRT-PCR was performed using a Bio-Rad MyiQ Single-Color Real-Time PCR System. At least four biological repetitions were performed for each experimental condition.

Table 1.

The 49 adhesion-related genes examined.

GeneBank Symbol Name
NM_001007145 Catna1 Catenin (cadherin associated protein), alpha 1
NM_053357 Ctnnb1 Catenin (cadherin associated protein), beta 1
NM_031333 Cdh2 Cadherin 2
XM_001061943 Cdh4 Cadherin 4
XM_215375 Col6a1 Collagen, type VI, alpha 1
NM_012967 Icam1 Intercellular adhesion molecule 1
NM_031521 Ncam1 Neural cell adhesion molecule 1
NM_031591 Pecam1 Platelet/endothelial cell adhesion molecule 1
XM_340884 Itga3 Integrin alpha 3
XM_235707 Itga5 Integrin alpha 5 (fibronectin receptor alpha)
NM_031768 Itgae Integrin, alpha E, epithelial-associated
NM_012711 Itgam Integrin alpha M
NM_017022 Itgb1 Integrin beta 1 (fibronectin receptor beta)
NM_153720 Itgb3 Integrin beta 3
XM_237536 Lama1 Laminin, alpha 1
XM_226159 Lama3 Laminin, alpha 3
XM_223087 Lamb3 Laminin, beta 3
NM_022266 Ctgf Connective tissue growth factor
NM_057118 Cntn1 Contactin 1
XM_573983 Tgfbi Transforming growth factor, beta induced
NM_012924 Cd44 Cd44 molecule
NM_001002023 Sgce Sarcoglycan, epsilon
NM_138879 Sele Selectin, endothelial cell
NM_013114 Selp Selectin, platelet
XM_214778 Thbs2 Thrombospondin 2
XM_232077 Ctnna2 Catenin (cadherin associated protein), alpha 2
NM_031334 Cdh1 Cadherin 1
XM_226426 Cdh3 Cadherin 3, type 1, P-cadherin (placental)
NM_134452 Col5a1 Collagen, type V, alpha 1
XM_221536 Col8a1 Collagen, type VIII, alpha 1
NM_012889 Vcam1 Vascular cell adhesion molecule 1
NM_203409 Ncam2 Neural cell adhesion molecule 2
XM_345156 Itga2 Integrin, alpha 2
XM_230033 Itga4 Integrin alpha 4
NM_031691 Itgad Integrin, alpha D
NM_001033998 Itgal Integrin alpha L
XM_230950 Itgav Integrin alpha V
XM_001069791 Itgb2 Integrin beta 2
NM_013180 Itgb4 Integrin beta 4
XM_219866 Lama2 Laminin, alpha 2
NM_012974 Lamb2 Laminin, beta 2
XM_341133 Lamd Laminin, gamma 1
NM_012881 Spp1 Secreted phosphoprotein 1
XM_342245 Postn Periostin, osteoblast specific factor
NM_019156 Vtn Vitronectin
NM_001033680 Syt1 Synaptotagmin I
NM_019177 Sell Selectin, lymphocyte
NM_001013062 Thbs1 Thrombospondin 1
NM_022587 Entpd1 Ectonucleoside triphosphate diphosphohydrolase 1
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The quality control of mRNA quantification was performed using three integrated control assays in the PCR array: the reverse transcription control, positive PCR control, and genomic DNA control. All PCR runs passed the first two control tests. However, the DNA contamination analysis identified one array to be contaminated by genomic DNA. As a result, the data from this array were excluded from the final analysis.

Immunohistology

Immunohistochemistry was used to localize protein expression of Cdh1 in the organ of Corti. The control (without receiving noise exposure) and the noise-traumatized animals were sacrificed. The cochleae were fixed with 10% buffered formalin and organs of Corti were dissected. Dissected tissues were then permeabilized for 30 min with 0.2% Triton X-100 in 10 mM PBS, blocked using 10% goat serum in PBS and then incubated with purified mouse E-cadherin primary antibody (1:200, BD Biosciences) at 4°C overnight. The tissues were then rinsed with 10 mM PBS (3×), incubated with a secondary antibody (Alexa Fluor 488 goat anti-mouse, 1:500, Invitrogen) for 1 h and then counterstained with propidium iodide. Two additional organs of Corti, one from a control subject and one from a noise-traumatized subject, were stained with only the secondary antibody to assess nonspecific staining.

Data analyses

ABR measurements

A two-way ANOVA (post-exposure time × test frequency) was used to compare ABR threshold shifts measured at 2 h, 1 day and 28 days post-exposure and among the five tested frequencies. If a significant main effect occurred, post hoc testing with a Tukey test was performed to delineate the nature of the differences.

Sensory cell damage

All the specimens were observed with a fluorescence microscope to identify cochlear lesions, which were further observed with a confocal microscope (Zeiss LSM510 multichannel laser scanning confocal imaging system) using a method that has been reported previously (Hu et al., 2006). To determine the site of primary lesions, we divided the sensory epithelium into the apical section and the basal section. The apical section contained the apical 45% sensory epithelium and the basal section consisted of the sensory epithelium section 45% to 90% from the apex. A paired Student’s t test was performed to compare the numbers of damaged sensory cells between the apical and basal sections of the sensory epithelia. An α level of 0.05 was selected for significance.

Quantification of Cdh1 positive sensory cells

The numbers of Cdh1-positive sensory cells in the apical and basal sensory epithelium sections were quantified by counting the sensory cells exhibiting strong Cdh1 immunoreactivity in their top junctions with supporting cells. The numbers were compared using a paired Student’s t test. An α level of 0.05 was selected for significance.

Analyses of mRNA expression levels of adhesion-related genes

The raw Ct value of each gene was first normalized to reference genes. The reference genes were selected from the five housekeeping genes (Rplp1, Hprt1, Rpl13a, Ldha and Actb) that are integrated into the qRT-PCR array. We used geNorm software to calculate the internal control gene-stability measures (M value) (Vandesompele et al., 2002), a quantitative parameter for the assessment of expression level variation of reference genes. Based on the M value, we selected three most stable genes (Rplp1, Hprt1 and Actb) and used the arithmetic mean of their expression levels as the baseline to calculate the ΔCt of each targeted gene. The fold changes in the expression levels of adhesion-related genes were calculated across two groups as ΔΔCt = ΔCt (noise group) – ΔCt (control group). The significance of the changes was analyzed using SAM (Significance Analysis of Microarrays) (Tusher et al., 2001) with the number of permutation of 100. A significant change was defined as q-values less than or equal to 0.01 and fold differences equal to or greater than 2. Because a Ct value of 35 represents single molecule template detection (Guthrie et al., 2008), a gene expression level with the raw Ct value greater than 35 was considered to be below the detection level of the assay in the current investigation.

Results

Expression patterns of adhesion genes in the apical and basal sections of the normal sensory epithelium

To profile the expression pattern of adhesion-related genes under normal conditions, we examined the constitutive expression levels of 49 adhesion-related genes in the apical and the basal sections of the sensory epithelium and found diverse expression levels of these genes (Supplementary Table S1). The highest-expressed genes (Ctnnb1, Catna1, Thbs1, and Lamb2) had expression levels either greater or less than, but within 1 cycle difference of, the average expression level of the reference control genes. The lowest-expressed genes (Sele, Itgad, Itgae, and Sell) had raw Ct values of >35 or were undetectable.

Anatomically, the apical and basal sections of the sensory epithelium have the same cell populations, yet they differ in many ways in their physiological and biological characteristics. We therefore sought to determine whether these two sections have different expression patterns for adhesion genes. First, we calculated the Pearson’s correlation coefficient of the ΔCt values across the 49 genes and found that the ΔCt values of the genes are highly correlated between the two partitions (r = 0.95, p < 0.0001 and slope = 1.007, Fig. 1). However, certain genes appeared to be less coordinated in their expression. To define the difference, we examined the fold differences in gene expression level using the SAM algorithm. Among the 49 examined genes, four were more highly expressed in the apical sections, whereas 23 were more highly expressed in the basal sections, with fold differences ranging from 2.55 to 14.35 and a false discovery rate <0.015 (bar graph in Fig. 2). To further illustrate the difference, we ranked the 49 genes based on their expression levels and compared the rank positions of the genes between the two anatomic sites. We found differences in the rank positions for multiple genes (scatter plot in Fig. 2). Notably, the result of the rank analysis is consistent with the result of the fold-difference analysis. Together, these observations suggest that transcriptional expression patterns of adhesion-related genes are not completely parallel in the apical and basal sections of the cochlear sensory epithelium.

Fig. 1.

Fig. 1

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Scatter plot showing the relative expression level (ΔCt) of each adhesion-related gene in the apical samples versus the basal samples. Expression levels were normalized to the reference genes. Correlation coefficients (r) and p values were determined with a Pearson’s correlation test.

Fig. 2.

Fig. 2

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Comparisons of the expression levels of adhesion-related genes between apical and basal samples. The expression levels of the genes in the apical samples are plotted as the baseline value (the zero line). Bars represent the fold differences between the basal and apical expression levels, and dots represent the rank difference between the apical and basal expression levels. A positive value indicates higher expression of a gene in the basal samples, and a negative value represents higher expression of a gene in the apical samples. Asterisks mark the genes having a fold difference equal to or greater than 2 and q-values less than or equal to 0.01.

Exposure to intense noise causes both functional deterioration and sensory cell damage in the cochlea

Exposure to intense noise traumatizes cochlear structures. To provide the cochlear damage context for profiling the gene expression levels, we examined the functional and morphological changes of the cochleae following the acoustic trauma. ABRs were measured pre-exposure and at three time points post-exposure (2 h, 1 day, and 28 days). Threshold shifts obtained at the first two post-exposure time points represent temporary threshold shifts, while threshold shifts measured at the last time point represent the permanent threshold shifts. Figure 3A shows the pre- and post-exposure ABR thresholds at the five tested frequencies representing low-, middle-, and high-frequency hearing. The analysis of post-exposure threshold shifts with a two-way ANOVA (time × frequency factors) revealed a significant effect of time (F = 233.06; df 2,145; p < 0.001) and a significant effect of frequency (F = 31.63; df 4,145; p < 0.01). The elevation of the thresholds was evident at 2 h post-exposure, with an average threshold shift of 70.1 ± 16.0 dB (mean ± SD). The frequency dependence of the threshold shifts was relatively flat, with slightly greater loss in the low frequencies (5 and 10 kHz, both differ from other frequencies, Tukey test, p < 0.001) and a notch at 20 kHz (differing from all other frequencies, Tukey test, p < 0.001). The thresholds were partially recovered at 1 day post-exposure (Tukey test, p < 0.001) and further recovered at 28 days post-exposure (Tukey test, p < 0.001), leaving an average permanent threshold shift of 23.1 ± 14.6 dB (mean ± SD). This observation suggests that the noise level used in the current investigation was capable of inducing a permanent loss of hearing sensitivity.

Fig. 3.

Fig. 3

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Functional and pathological changes in the noise-traumatized cochleae. A, The threshold shifts of auditory brainstem responses (ABRs) at 2 h, 1 day, and 28 days post-exposure. B, An example of sensory cell damage illustrated by propidium iodide staining from a cochlea examined at 2 h post-noise exposure. Arrows indicate sensory cells with malformed nuclei. The double-arrow indicates the site of a missing cell. C, An example of caspase-3 staining from a cochlea examined at 2 h post-noise exposure. Arrows indicate the sensory cells having condensed or fragmented nuclei. These cells exhibit strong caspase-3 activity (green). Scale bar = 20 μm. D, An example of the nuclear morphology in a normal sensory epithelium from a control ear. IHC indicates inner hair cells, while OHC1, OHC2, and OHC3 indicate the first, second, and third rows of outer hair cells, respectively. E, Cochleogram showing the distribution of damaged sensory cells in the organ of Corti at 2 h post-noise exposure. The two horizontal arrows mark the regions of the sensory epithelium used for quantification of sensory cell damage in the apical and basal sections (see Fig. 3F). Noted that the counts from the basal region between 90 and 100% were not included in the calculation for Fig. 3F. F, Comparison of the numbers of degenerated sensory cells in the apical and basal sections of the sensory epithelium in normal and noise-damaged ears. The number of damaged sensory cells in the basal section is significantly higher than in the apical section (paired Student’s t test, *p < 0.05). Bars: one standard deviation. N = the number of the ears. (For interpretation of the references to color in this figure legend, the reader is referred to the web version of this article.)

To further evaluate the impact of noise, we examined the level of sensory cell damage at 2 h post-exposure. Nuclear morphology was used as the index for defining sensory cell damage, because its change is an early sign of damage and because its progression occurs in a step-by-step fashion (Hu et al., 2006; Yang et al., 2004). We found malformed sensory cell nuclei, as well as indications of missing cells, at this time point (Fig. 3B). Caspase-3 staining revealed that sensory cells having condensed or fragmented nuclei exhibited strong caspase-3 activity (Fig. 3C), suggesting that these cells were apoptotic cells. In contrast, the malformed or missing nuclei were rare in normal control ears (Fig. 3D). To quantify the level of damage, we counted the damaged cells (the cells with condensed or swollen nuclei, as well as missing cells) along the entire length of the organs of Corti and used the data to assemble a cochleogram. As shown in Figure 3E, there were two lesions in the organ of Corti, one in the apical partition and the other in the basal partition of the cochlea. We then compared the numbers of total damaged cells between the apical and the basal sections and found that the basal partition had more damaged cells (22.3 ± 27.6 vs. 62.1 ±64.8, mean ± SD; paired Student’s t test, p = 0.003, Fig. 3F), suggesting that the basal section is the major site of sensory cell pathogenesis. Notably, the noise-induced damage was significantly greater than the spontaneous sensory cell degeneration in normal cochleae (22.3 ± 27.6 vs.2.9 ± 3.4 for the apical partition; 62.1 ± 64.8 vs. 16.9 ± 13.4 for the basal partition; Student’s t test, p = 0.01 for both partitions). Together, these observations indicate that the noise condition used in the current study was capable of inducing not only hearing loss but also sensory cell death. As shown in our previous publications (Hu and Cai, 2010; Hu et al., 2009), this initial sensory cell damage will continue to develop at 1 day and 7 days post-noise exposure.

Acoustic overstimulation causes a site-specific change in mRNA expression levels of adhesion genes in the sensory epithelium

To define the transcriptional responses of the adhesion-related genes to acoustic overstimulation, we analyzed the fold change in expression level of the 49 genes using SAM software. In the apical samples, five genes (Sell, Thbs1, Itgae, Icam1, and Itga5) were significantly upregulated and none was downregulated at 2 h post-exposure (Table 2). These upregulated genes belong to different adhesion families, including the immunoglobulin superfamily (lcam1) and the selectin (Sell) and integrin (Itgae and Itga5) families. Among these upregulated genes, two (Sell and Itgae) were undetectable in the normal sensory epithelium, but became detectable after noise exposure.

Table 2.

Differential expression of adhesion-related genes 2 h post-exposure in the apical and basal sections. q Value <0.01 indicates a significant expression change.

Symbol Apical Section Basal Section
Fold change q value Fold change q value
Itgae 6.464 <0.01
Sell 6.233 <0.01
Itga5 2.063 <0.01
* Icam1 4.112 <0.01 8.206 <0.01
* Thbs1 4.765 <0.01 2.241 <0.01
Selp 376.5 <0.01
Sele 18.79 <0.01
Itgb2 13.35 <0.01
Itga3 3.180 <0.01
Cdh1 2.457 <0.01
Cdh2 3.191 <0.01
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Note: asterisks indicate the genes that are altered in both the apical and the basal sections

The basal samples also featured an upregulation of the gene expression level. However, the expression changes in the basal samples were not completely parallel to the apical samples. First, the total number of altered genes was more in the basal partition than in the apical partition (8 genes vs. 5 genes). Second, changes in expression at the two sites involved different gene sets. Three genes (Itgae, Itga5, and Sell) were upregulated in the apical partition, but not in the basal partition. Six genes (Itga3, Itgb2, Selp, Sele, Cdh1, and Cdh2) were upregulated in the basal partition, but not in the apical partition. At this time point, only two genes (Icam1 and Thbs1) co-varied in the two anatomic sites. Together, these data provide evidence that acoustic trauma provokes differential expression changes in adhesion-related genes between the apical and the basal sections of the sensory epithelium.

The site-specific transcriptional change in Cdh1 is consistent with its protein expression change

To provide further evidence for the site-specific response of adhesion genes to acoustic overstimulation in the sensory epithelium, we examined the protein expression of Cdh1 gene, because this gene had exhibited a significant transcriptional increase only in the basal samples (fold change = 2.46, q = 0 for the basal samples vs. fold change = 1.15, q = 0.47 for the apical samples). Employing immunohistology to identify the protein expression pattern, we found that in the normal cochlear sensory epithelium, strong immunoreactivity to the Cdh1 protein appeared in the intercellular junctions among supporting cells, including Deiters cells, pillar cells and Hensen cells (Fig. 4A), while staining in the junctions around sensory cells was weak or undetectable (data not shown). This finding is consistent with the previously reported distribution of the Cdh1 protein (Nunes et al., 2006; Whitlon, 1993).

Fig. 4.

Fig. 4

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Cdh1 immunoreactivity in the sensory epithelium of a normal and a noise-traumatized cochlea. A, A micrograph showing distribution of Cdh1 immunoreactivity in a normal organ of Corti. The Cdh1 immunoreactivity (green fluorescence) is located in the intercellular junctions between supporting cells, including pillar cells, Deiters cells, and Hensen cells. The nuclei of the cells, illustrated by propidium iodide staining (red fluorescence), appear normal. B and C, Cdh1 staining in a noise-traumatized organ of Corti. Some outer hair cells exhibit strong Cdh1 immunoreactivity in their top junctions with neighboring supporting cells (double-arrows) along with malformed nuclei (arrows). One OHC has strong Cdh1 immunoreactivity (the arrowhead in C), but its nucleus has a relatively normal shape (see the nucleus to the left of the asterisk). IHC represents inner hair cells, while OHC represents outer hair cells. Scale bar = 20 μm. D, Comparison of the numbers of Cdh1-positive sensory cells between the apical and the basal sections of the sensory epithelium. The basal section has more Cdh1-positive sensory cells than the apical section (paired Student’s t test, * p < 0.01). Bars: one standard deviation. N = the number of ears. (For interpretation of the references to color in this figure legend, the reader is referred to the web version of this article.)

After exposure to intense noise, a marked increase in Cdh1 immunoreactivity appeared in some of the outer hair cells in the damaged regions of the sensory epithelium. The fluorescence was located in the circumferential rings of the outer hair cells, the site where the outer hair cell forms junctions with the phalangeal processes of Deiters cells and pillar cells (Figs. 4B and 4C). To determine the viability of these Cdh1-positive cells, we counterstained the tissues with propidium iodide, a nuclear dye, to illustrate nuclear morphology. Most Cdh1-positive cells exhibited malformed nuclei with increased propidium iodide fluorescence, suggesting that these Cdh1-positive cells were damaged (Fig. 4B). However, some Cdh1-positive cells (less than 5% of total Cdh1-positive cells) showed a relatively normal nuclear morphology (Fig. 4C). This may suggest that the change in Cdh1 protein precedes nuclear degradation. Moreover, the result indicates that the site-specific upregulation in Cdh1 expression is spatially correlated with sensory cell degeneration.

To quantify the expression change, we counted the numbers of Cdh1-positive cells in the sensory epithelia, and compared their numbers between the apical and the basal sections. As shown in Figure 4D, the number of Cdh1-positive cells in the basal partition is significantly higher than that in the apical partition (75.5 ± 67.6 vs. 34.7 ± 34.6, mean ± SD, paired Student’s t test, p < 0.01). This finding is consistent with the observation of significant transcriptional upregulation of Cdh1 in the basal samples of the sensory epithelia, further suggesting the occurrence of site-specific alteration in expression of adhesion genes.

To verify the specificity of the immunolabeling, we checked the molecular weight of the protein targeted by the Cdh1 antibody using a western blotting assay and found two bands, 100 kDa and 120 kDa, consistent with the molecular weights of the two isoforms of the Cdh1 protein reported in non-cochlear tissues (Barshishat et al., 2000). Moreover, the primary antibody control observation, which omitted the primary antibody, exhibited no fluorescence at the cell–cell junctions in the organ of Corti (date not shown).

Individual variation in the functional loss of the cochleae is correlated with the expression levels of adhesion-related genes

Individual variation in cochlear responses to acoustic trauma has been reported in previous studies (Hunter-Duvar, 1977; Lipscomb et al., 1977; Saunders et al., 1985; Thorne et al., 1984). Given the finding of individual variation in transcriptional levels of adhesion-related genes, we sought to determine whether individual variation in transcription level is related to the level of hearing loss. To this end, we calculated the Pearson’s correlation between the average ABR threshold shift and the expression level for each gene. In the apical partition, two genes had expression levels that were correlated with the magnitude of hearing loss, one positively (Sgce) and the other negatively (Sell), with the positive correlation meaning that a higher expression level was associated with greater hearing loss (Fig. 5, r = 0.95 and −0.98, respectively, p < 0.05). In the basal partition, six genes showed expression levels that were correlated, four positively (Itga5, Itgal, Sell, and Selp) and two negatively (Cntn1 and Col5a1), with the level of hearing loss (Fig. 5, r = 0.97, 0.95, 0.97, 0.98, −0.99, and −0.96, respectively, p < 0.05). We suspect that these genes contribute to an individual’s susceptibility to acoustic trauma.

Fig. 5.

Fig. 5

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Correlations between the threshold shifts of auditory brainstem responses (ABRs) and the relative expression levels of seven adhesion-related genes from either apical or basal samples.

Expression changes occur in genes with diverse baseline levels of expression

Given the diversity of the constitutive expression levels of the 49 genes examined, we wondered whether the baseline transcriptional level of a gene contributes to its propensity to respond to mechanical stress. To address this question, we examined the rank distribution of the expression-altered genes among the 49 genes. The rank position of a gene was based on its constitutive expression level. As shown in Figure 6, the expression-altered genes are spread across the full range of the rank order, indicating that these genes have different constitutive expression levels.

Fig. 6.

Fig. 6

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The distribution of genes showing expression changes along the rank sequence of the 49 genes examined. Dots indicate the individual ranking position of the genes based on their constitutive expression levels.

The early transcriptional changes in adhesion-related genes are biphasic

Previous observations have revealed the dynamic nature of cochlear pathogenesis (Hu et al., 2002b; Pye, 1981; Thorne and Gavin, 1985; Yang et al., 2004). We therefore sought to determine whether the early expression change observed at 2 h post-exposure persists to 1 day post-exposure. In contrast to the up-regulation-dominated changes at 2 h post-exposure, the expression pattern at day 1 featured a trend of down-regulation. All the upregulated expression levels observed at 2 h post-exposure returned to their baseline levels. At 1 day post-exposure (Fig. 7), six genes (Cd44, Pecam1, Ctgf, Ctnna2, Lama2, and Cntn1) were significantly downregulated with fold-changes ranging from 2.47 to 8.29 (SAM analysis, q = 0). None of these downregulated genes was upregulated at 2 h post-exposure. The data obtained from the two time points demonstrate the presence of a biphasic change in the expression levels of adhesion-related genes in the first post-exposure day, indicating that the noise-induced transcriptional change is a time-dependent event.

Fig. 7.

Fig. 7

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Fold difference of the transcriptional expression levels of adhesion-related genes between normal tissues and noise-damaged tissues examined 1 day post-exposure. Negative values indicate the downregulation of genes. Asterisks indicate q-values < 0.01.

Noise-induced transcriptional changes in the lateral wall tissue of the cochlea

The lateral wall tissue, consisting of the stria vascularis and the spiral ligament, plays an important role in maintaining cochlear function. Although the sensory epithelium is the primary target of acoustic trauma, functional and morphological changes have been found in the lateral wall structures (Engstrom and Engstrom, 1979; Lim, 1976; Shi and Nuttall, 2007). To determine whether acoustic overstimulation causes a parallel change in adhesion gene expression in the lateral wall tissue, we profiled the lateral wall expression of adhesion-related genes and examined their changes at 2 h post-exposure. In contrast to the upregulation-dominated changes in the sensory epithelium, the lateral wall samples featured a downregulation-dominated change in gene expression (Table 3). Among the 25 genes with expression changes, 21 were downregulated, and only four were upregulated. Among the four upregulated genes, three (Sele, Selp, and Thbs1) co-varied in the sensory epithelium. In contrast, only one gene (Cdh2) among the 21 downregulated genes co-varied in the cochlear sensory epithelia. These observations suggest that these upregulated genes (Sele, Selp, and Thbs1) are more likely to co-vary in the lateral wall and the sensory epithelium, whereas the downregulated genes (see Table 3) tend to be site-specific. Together, our current investigation indicates that exposure to intense noise causes site-specific changes in the sensory epithelium and the lateral wall tissue.

Table 3.

Differential expression of adhesion-related genes 2 h post-exposure in the lateral wall tissues. q Value <0.01 indicates a significant expression change.

Genes Fold change q value
Thbs1 7.464 < 0.01
Selp 107.3 = 0.01
Upregulated Cdh3 2.799 = 0.01
Sele 6.600 = 0.01
Down regulated Ncam1 8.442 < 0.01
Lama2 5.260 < 0.01
Spp1 5.408 < 0.01
Itga4 3.850 < 0.01
Thbs2 2.694 < 0.01
Ncam2 8.770 < 0.01
Itgav 3.986 < 0.01
Lama1 5.306 < 0.01
Syt1 11.94 < 0.01
Cntn1 3.938 < 0.01
Lamb3 3.891 < 0.01
Col8a1 7.198 < 0.01
Cdh2 3.965 < 0.01
Entpd1 4.651 < 0.01
Lamc1 5.269 < 0.01
Itgb1 4.431 < 0.01
Vtn 7.438 < 0.01
Ctnna2 2.814 < 0.01
Vcam1 4.250 < 0.01
Col6a1 3.713 < 0.01
Itga2 2.374 = 0.01
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Discussion

The objectives of the current investigation were to profile the transcriptional pattern of adhesion-related genes and to define their expression changes induced by acoustic trauma in the rat cochlea. This study documents the constitutive expression of multiple adhesion-related genes in normal cochlear tissues, many of which have not been previously reported, and also documents noise-induced differential changes in the expression levels of these genes and a high correlation between the expression levels of certain adhesion genes and functional loss of the cochlea. These results provide important clues for further investigation of adhesion signaling pathways in noise-induced cochlear pathogenesis and the repair process.

The effort to understand transcriptional changes in different cochlear partitions, particularly in the different sections of the sensory epithelium, has been hampered by the scarceness of cochlear tissues available for RNA analyses and the difficulty of dissecting fresh cochlear tissues without contamination. Our method, using a combination of the techniques of fresh tissue dissection, cDNA pre-amplification, and RNA preservation, allowed us, for the first time, to investigate site-specific transcriptional changes in the cochlea and to correlate the transcriptional changes to functional change in the cochlea.

We demonstrate in the current study that the constitutive expression levels of certain adhesion-related genes differ in the apical and the basal sections of the sensory epithelium. We also present evidence of a site-specific change in adhesion-gene transcription after exposure to intense noise. Specifically, the apical and the basal samples exhibit different numbers of genes that are altered in expression level. Three genes show expression changes exclusively in the apical samples and six in the basal samples. Only two genes were co-regulated in both the apical and the basal samples. This observation suggests that the apical and basal sections of the sensory epithelium undergo different adhesion modulation with the basal section sustaining a greater adhesion change. This difference may also be attributed to the difference in the damage levels in the two sections of the sensory epithelium. Notably, analyses of gene families show that the site-specific change involves genes from the integrin, cadherin, and selectin gene families, whereas the change in the immunoglobulin family member (Icam1) is ubiquitous in the sensory epithelium.

Individual variation in the levels of functional and pathological changes is a hallmark of acoustic trauma (Hunter-Duvar, 1977; Lipscomb et al., 1977; Saunders et al., 1985; Thorne et al., 1984; Wagner et al., 2003), and is clearly evident in the current investigation. However, the molecular basis for the variation is not clear. Our correlation analysis identified seven genes showing a correlation between the expression level and the level of hearing loss. These genes may contribute to the individual variation in the levels of cochlear damage after acoustic overstimulation. However, due to the limited sample size and the limitation of the correlation analysis for defining causality, it is essential to confirm the current results using experimental approaches to manipulating gene expression, so that the roles of these genes in the development of cochlear damage can be more accurately defined.

Cochlear tissues consist of heterogeneous cell populations and the junctions connecting these cells involve multiple adhesion proteins. Previous studies have identified some of the adhesion-related proteins in cochlear tissues, such as proteins from the Cdh1, I-CAM, and integrin families (Davies and Holley, 2002; Suzuki and Harris, 1995; Toyama et al., 2005; Whitlon, 1993). The current study confirms the cochlear expression of the corresponding genes at the transcriptional level. Importantly, we have detected the expression of many genes that have not previously been linked to cochlear structures.

We found expression changes in all four adhesion gene families. Selectin family members are a family of transmembrane molecules that are expressed on the surface of leukocytes and activated endothelial cells (Chen and Geng, 2006; Stamenkovic, 1995). In the current study, we demonstrate that the constitutive expression of selectin family genes (Sell, Sele, and Selp) is either very low or undetectable, suggesting that these genes may not be major players in the adhesion functions of cochlear structures. However, the expression levels of this gene family were upregulated after noise exposure. Although the biological role of selectin gene activation in cochlear pathogenesis remains unclear, evidence suggests that these genes play an important role in cochlear inflammation. Selectin family members have been found to participate in the inflammatory response by mediating the interaction between leukocytes, platelets, and endothelia (Grailer et al., 2009). In the cochlea, recruitment of inflammatory cells to the injured cochlea has been found after acoustic trauma (Fredelius and Rask-Andersen, 1990; Hirose et al., 2005). We therefore suggest that selectin family genes are mediators of cochlear inflammation. It is likely that controlling expression of these inflammatory genes will serve as a new therapeutic target for suppression of inflammatory responses (Rossi and Constantin, 2008).

We found upregulation of three cadherin family members after acoustic overstimulation. Cadherin proteins are a group of calcium-dependent proteins playing an important role in maintaining cell-cell junction function and cellular architecture via interaction with actin molecules through catenin proteins (Leckband and Prakasam, 2006). Expression of cadherin family genes, including Cdh1 and Cdh2, in the cochlear tissues has been reported (Simonneau et al., 2003; Whitlon, 1993; Whitlon et al., 1999). Thus far, the precise role of the Cdh1 protein in sensory cell degeneration is not clear. Evidence from the current investigation and from previous publications suggests that this protein may be involved in both degenerative and repair processes. In noise-traumatized cochleae, the Cdh1 protein has been implicated in scar formation in the recovery phase of cochlear pathogenesis (Raphael and Altschuler, 1991). Our current investigation further links cadherin family genes to the initial phase of cochlear damage. We found that most sensory cells showing strong Cdh1 immunoreactivity have malformed nuclei. Notably, certain cells with strong Cdh1 immunoreactivity exhibit relatively normal nuclear morphology, suggesting that Cdh1 is involved in the early stages of the degenerative process in these cells. Cadherin proteins have been found to play an important role in maintaining cytoskeletal structures (Tsukita et al., 1992), and in sensory cells, the increased Cdh1 immunoreactivity is spatially close to the site of actin, the cuticular plates of the cells. We have previously demonstrated rapid degradation of F-actin protein in the early phase of acute sensory cell apoptosis (Hu et al., 2002a). Together, this evidence suggests the involvement of Cdh1 in sensory cell degeneration.

We have also demonstrated the involvement of immunoglobulin superfamily and integrin family genes in acoustic trauma. Icam1, an immunoglobulin gene, is upregulated in both the apical and the basal partitions of the sensory epithelium, and its expression has previously been linked to the inflammatory response (Fujimoto et al., 2008; Glushakova et al., 2008), including cochlear inflammation (Suzuki and Harris, 1995). In ischemia/reperfusion models, early Icam-1 expression has been observed (Olanders et al., 2002) and this expression has been linked to oxidative stress (Lange et al., 2008). Given the reduction in antioxidant capacity during noise trauma (Henderson et al., 2006), we argue that the increase in Icam1 expression may be linked to oxidative stress in the cochlea.

In summary, the current study was designed to investigate the involvement of adhesion-related genes in cochlear pathogenesis caused by acoustic trauma. This study, for the first time, examined site-specific gene expression patterns in the sensory epithelia in noise-injured cochleae. We describe multiple novel findings, including constitutive expression patterns of adhesion-related genes in normal rat cochleae and differential expression changes following acoustic trauma. Importantly, we uncover links between functional loss of the cochlea and gene expression changes. This study has implications for understanding of the molecular mechanisms of acute sensory cell damage and individual variation in cochlear susceptibility to acoustic overstimulation. Future investigations into the roles for these genes in the functional integrity of particular types of intercellular junctions in the cochlea will provide a better understanding of their contribution to noise-induced cochlear damage.

Supplementary Material

Supplementary Table 1s

Supplementary Table S1. Relative expression levels (ΔCt) of adhesion-related genes in the apical and the basal sections of the sensory epithelia.

Research Highlights.

  • Multiple families of adhesion-related genes are expressed in the rat cochlea.

  • Acoustic trauma alters expression patterns of adhesion-related genes.

  • Cochlear functional loss is related to the expression levels of adhesion genes.

  • Genes showing expression changes exhibit diverse constitutive expression levels

Acknowledgments

The research was supported by NIDCD1R01 DC010154-01 to BH Hu.

Abbreviation

ABR

Auditory brainstem response

SAM

Significance Analysis of Microarrays

PBS

phosphate buffered saline

Contributor Information

Qunfeng Cai, Email: qcai@buffalo.edu.

Minal Patel, Email: minalpat@buffalo.edu.

Donald Coling, Email: dcoling@buffalo.edu.

Bo Hua Hu, Email: bhu@buffalo.edu.

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Associated Data

This section collects any data citations, data availability statements, or supplementary materials included in this article.

Supplementary Materials

Supplementary Table 1s

Supplementary Table S1. Relative expression levels (ΔCt) of adhesion-related genes in the apical and the basal sections of the sensory epithelia.

NIHMS410934-supplement-Supplementary_Table_1s.tif (10.6MB, tif)

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