JARO

JARO 17: 289–302 (2016) DOI: 10.1007/s10162-016-0567-7 D 2016 Association for Research in Otolaryngology

Research Article

Journal of the Association for Research in Otolaryngology

Inhibitors of Histone Deacetylases Attenuate Noise-Induced Hearing Loss JUN CHEN,1,2 KAYLA HILL,1

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SU-HUA SHA1

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Department of Pathology and Laboratory Medicine, Medical University of South Carolina, Walton Research Building, Room 403-E, 39 Sabin Street, Charleston, SC 29425, USA

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Department of Otolaryngology-Head and Neck Surgery, Xijing Hospital, Fourth Military Medical University, Xi’an, 710032, China Received: 13 November 2015; Accepted: 30 March 2016; Online publication: 19 April 2016

ABSTRACT Loss of auditory sensory hair cells is the major pathological feature of noise-induced hearing loss (NIHL). Currently, no established clinical therapies for prevention or amelioration of NIHL are available. The absence of treatments is due to our lack of a comprehensive understanding of the molecular mechanisms underlying noise-induced damage. Our previous study indicates that epigenetic modification of histones alters hair cell survival. In this study, we investigated the effect of noise exposure on histone H3 lysine 9 acetylation (H3K9ac) in the inner ear of adult CBA/J mice and determined if inhibition of histone deacetylases by systemic administration of suberoylanilide hydroxamic acid (SAHA) could attenuate NIHL. Our results showed that H3K9ac was decreased in the nuclei of outer hair cells (OHCs) and marginal cells of the stria vascularis in the basal region after exposure to a traumatic noise paradigm known to induce permanent threshold shifts (PTS). Consistent with these results, levels of histone deacetylases 1, 2, and 3 (HDAC1, HDAC2 and HDAC3) were increased predominately in the nuclei of cochlear cells. Silencing of HDAC1, HDAC2, or HDAC3 with siRNA reduced the expression of the target HDAC in OHCs, but did not attenuate noiseinduced PTS, whereas treatment with the pan-HDAC inhibitor SAHA, also named vorinostat, reduced OHC

Correspondence to: Su-Hua Sha & Department of Pathology and Laboratory Medicine & Medical University of South Carolina & Walton Research Building, Room 403-E, 39 Sabin Street, Charleston, SC 29425, USA. Telephone: 843-792-8324; email: [email protected]

loss, and attenuated PTS. These findings suggest that histone acetylation is involved in the pathogenesis of noise-induced OHC death and hearing loss. Pharmacological targeting of histone deacetylases may afford a strategy for protection against NIHL. Keywords: prevention of noise-induced hearing loss, histone acetylation, histone deacetylase inhibitor SAHA

INTRODUCTION Noise-induced hearing loss (NIHL) is increasing in industrialized countries stemming both from workplace exposures and leisure activities. The major pathological feature of NIHL is loss of cochlear hair cells, which adult mammals lack the ability to regenerate, resulting in permanent hearing loss. Although NIHL is a detriment to the quality of life of affected individuals and confers a great economic cost, there currently are no established clinical therapies for prevention or treatment of NIHL (Oishi and Schacht 2011). Post-translational modification of histones is an important form of chromatin regulation. By altering their interactions with DNA and nuclear proteins without changing the DNA sequence, local histone modifications impact the activation of gene transcription, including the transcription of genes related to stress response and cell fate. Histones, the primary components of chromatin organization, wrap DNA into packaged units called nucleosomes. Each nucle289

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osome core is an octamer comprised of the histone proteins H2A, H2B, H3, and H4. All histone core proteins share long amino-terminal tails that are accessible to post-translational enzymatic modifications including methylation, acetylation, and phosphorylation (Kouzarides 2007). In general, acetylation of histone H3 lysine 9 (H3K9ac) increases gene expression by altering DNA and protein interaction and thus accessibility to transcriptional machinery, whereas methylation of histone H3 lysine 9 is associated with gene silencing. Histone acetyltransferases (HATs) and histone deacetylases (HDACs) are enzymes that regulate acetylation of histone proteins at lysine residues. One of the most common transcriptional coactivators of HATs is p300, which contains a HAT domain and promotes histone acetylation (Chakravarti et al. 1999), while HDACs remove those acetyl groups, often condensing chromatin and repressing gene transcription. Four classes of mammalian HDACs have been identified based on homology to yeast HDACs (Dietz and Casaccia 2010). Among the HDACs, HDAC2 has been recognized as having a pathogenic role in neurodegenerative diseases such as Alzheimer’s disease (Falkenberg and Johnstone 2014) and HIV-infected neuronal cells (Atluri et al. 2014). Epigenetic aberrations are involved in several human pathologies, including cancers, neurological diseases, and immune disorders, as well as wide range of brain disorders. Histone deacetylase inhibitors have emerged as a promising new strategy for therapeutic intervention, including in neurodegenerative diseases (Saha and Pahan 2006; Dompierre et al. 2007). Among several histone deacetylase inhibitors, suberoylanilide hydroxamic acid (SAHA), also known in medicine as vorinostat, crosses the blood-brain barrier and is the first of the HDAC inhibitors to be approved by the Food and Drug Administration for clinical use in cancer treatment (Xu et al. 2007). SAHA binds to the catalytic domains of HDACs and efficiently blocks class I, II, and IV HDACs, leading to an accumulation of acetylated histones. Interestingly, decreasing histone acetylation or treatment with the HDAC inhibitor trichostatin A attenuated gentamicinand cisplatin-induced ototoxicity (Drottar et al. 2006; Jiang et al. 2006; Chen et al. 2009; Wang et al. 2013). Recently, SAHA treatment was shown to attenuate hearing loss caused by treatment with the combination of kanamycin with furosemide as well as noiseinduced hearing loss in mice (Layman et al. 2015; Wen et al. 2015). In this study, we investigated changes of H3K9ac, HDAC1, HDAC2, and HDAC3 in the inner ear using adult male CBA/J mice under noise conditions that induced permanent threshold shifts (PTS noise) with loss of outer hair cells (OHCs), as we have previously

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described (Yuan et al. 2015). We also silenced HDAC1, HDAC2, and HDAC3 individually (using siRNA) to determine if blockade of one isoform of HDACs could attenuate noise-induced permanent hearing loss. Finally, we employed SAHA treatment to assess if blocking HDACs could attenuate noiseinduced OHC loss and NIHL. Our study supports the hypothesis that histone modifications are involved in the sequelae of noise exposure and that treatment with the HDAC inhibitor SAHA attenuates noiseinduced OHC loss and subsequent NIHL.

MATERIALS AND METHODS Materials The antibodies used in this study were monoclonal rabbit anti-H3K9ac (Cell Signaling Technology, #9649), polyclonal rabbit anti-histone H3 (Cell Signaling Technology, #9715), monoclonal mouse antiHDAC1, 2, and 3 (Cell Signaling Technology, #9928), polyclonal mouse anti-GAPDH (Millipore, #ABS16), and polyclonal rabbit anti-myosin VII (Proteus Biosciences, #25-6790). Secondary antibodies for Western blotting were obtained from Jackson ImmunoResearch Laboratories and secondary Alexa 488 fluorescent antibodies from Invitrogen. Suberoylanilide hydroxamic acid (SAHA) was from Cayman Chemical Inc. (#10009929). ECL and Supersignal Dura-Enhanced Chemiluminescence Plus (ECL-Plus) for Western blotting detection and PageRuler Plus protein ladder were purchased from Fisher Scientific. All other reagents were purchased from Sigma-Aldrich.

Animals Male CBA/J mice at 11 weeks of age (Harlan Sprague Dawley) were allowed free access to water and a regular mouse diet (Purina 5025), and were kept at 22 ± 1 °C under a standard 12-h light/dark cycle to acclimate for 1 week before the experiments. All research protocols were approved by the Institutional Animal Care and Use Committee at the Medical University of South Carolina (MUSC). Animal care was under the supervision of the Division of Laboratory Animal Resources at MUSC.

Noise Exposure Unrestrained CBA/J male mice at 12 weeks of age (one mouse per stainless steel wire cage, approximately 9 cm3) were exposed to a broadband noise (BBN) with a frequency spectrum from 2 to 20 kHz for 2 h at 98 dB sound pressure level (SPL) to induce PTS with loss of OHCs (Yuan et al. 2015). Four

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individual stainless steel wire cages were located in the center of the sound chamber and the level of sound was consistent throughout the field during the noise exposure. The sound exposure chamber was fitted with a loudspeaker (model 2450H; JBL) driven by a power amplifier (model XLS 202D; Crown Audio) fed from a CD player (model CD-200; Tascam TEAC American). Audio CD sound files are created and equalized with audio editing software (Audition 3; Adobe System, Inc.). The background sound intensity of the environment surrounding the cages was 65 dB as measured with a sound level meter (model 1200, Quest Technologies). Sound levels for noise exposure are calibrated with a sound level meter at multiple locations within the sound chamber to ensure uniformity of the sound field and are measured before and after exposure to ensure stability. Control mice were kept in silence (without use of the loudspeaker) within the same chamber for 2 h.

Treatment of Mice with SAHA SAHA was dissolved in dimethyl sulfoxide (DMSO) as stock solutions (50 mg/mL SAHA) as previously described (Faraco et al. 2006; Guan et al. 2009; Li et al. 2010). Just prior to injection, the stock solution of SAHA was diluted with saline. The final concentration of SAHA for injection was 50-mg/kg body weight. Mice received intraperitoneal (IP) injections of SAHA 6 times: once per day at 1 day before and three consecutive days after noise exposure, as well as 30 min before and 8 h after the noise exposure on the day of noise exposure. Control mice received the same amounts of DMSO at 1 μL/g body weight.

Intra-tympanic Delivery of siRNA The methods for small-interfering RNA (siRNA) delivery into the adult mouse inner ear in vivo were modified from a recent study (Oishi et al. 2013). Briefly, after anesthesia, a retroauricular incision was made to approach the temporal bone. The otic bulla was identified ventral to the facial nerve, and a shallow hole was made in the thin part of the otic bulla with a 30-G needle and enlarged with a dental drill to a diameter of 2 mm in order to visualize the round window. A customized sterile micro medical tube was inserted into the hole just above the round window niche to slowly deliver 0.6 μg (10 μL) of pre-designed HDAC1, HDAC2, or HDAC3 siRNA (Life Technologies siRNA ID# s67416) to cover the round window niche. After the siRNA was delivered, the hole was covered with surrounding muscle and glued with tissue adhesive. Finally, the skin incision was closed with tissue adhesive and the mouse was kept in the surgical position for about 1 h. Based on our previous

experiments (Oishi et al. 2013; Zheng et al. 2014; Yuan et al. 2015), local intra-tympanic siRNA delivery resulted in temporary elevation of thresholds that completely recovered by 72 h. Therefore, noise exposure was performed 72 h after siRNA delivery. The animals were exposed to BBN or kept in silence in the noise chamber for 2 h. Although the volume of the mouse middle ear cavity is 5–8 μL, we slowly flushed 10 μL of the siRNA solution into the cavity, as some of the solution flows out through the 2-mm made for visualization of the round window niche.

Auditory Brainstem Response (ABR) Mice were anesthetized with an intraperitoneal injection of xylazine (10 mg/kg) and ketamine (100 mg/ kg), and then placed in a sound-isolated and electrically shielded booth (Acoustic Systems). Body temperature was monitored and maintained near 37 °C with a heating pad. Acoustic stimuli were delivered monaurally to a Beyer earphone attached to a customized plastic speculum inserted into the ear canal. Subdermal electrodes were inserted at the vertex of the skull, under the left ear, and under the right ear (ground). ABRs were measured at 8, 16, and 32 kHz. Tucker Davis Technology (TDT) System III hardware and SigGen/Biosig software were used to present the stimuli (15 ms duration tone bursts with 1 ms risefall time) and record the response. Up to 1024 responses were averaged for each stimulus level. Thresholds were determined for each frequency by reducing the intensity in 10-dB increments and then in 5-dB steps near threshold until no organized responses were detected. Thresholds were estimated between the lowest stimulus level where a response was observed and the highest level without response. ABR waves I and II were monitored to assess thresholds. All ABR measurements were conducted by the same experimenter. The ABR scores were assigned by an expert who was blinded to the treatment conditions.

Immunocytochemistry for Cochlear Surface Preparations and Quantification of the Immunofluorescence Signals We have followed a procedure as previously described (Chen et al. 2012; Zheng et al. 2014; Yuan et al. 2015). The temporal bones were removed 1 h after noise exposure and perfused through the scala with a solution of 4 % paraformaldehyde in PBS, pH 7.4, and kept in this fixative overnight at 4 °C. The cochleae were then rinsed in PBS. Before decalcification of each cochlea in a 4 % solution of sodium EDTA (adjusted with HCl to pH 7.4), the apical otic capsule was removed. The EDTA solution was

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changed daily for 3 days at 4 °C. Following decalcification with 4 % EDTA, cochleae were used for immunocytochemistry on surface preparations. The cochleae were dissected under a microscope by removing the softened otic capsule, stria vascularis, Reissner’s membrane, and tectorial membrane. The remaining tissue, including the modiolus and cochlear sensory epithelium, was permeabilized in 3 % Triton X-100 solution for 30 min at room temperature. The specimens were washed three times with PBS and blocked with 10 % normal goat serum for 30 min at room temperature, followed by incubation with primary antibodies anti-H3K9ac or antiHDAC2 at 4 °C for 48 h. After washing three times (10 min each wash), the tissues were incubated with the Alexa-488-conjugated secondary antibody at a concentration of 1:200 at 4 °C overnight in darkness. The specimens were washed three times with PBS and incubated with propidium iodide (PI) at 1 μg/mL in PBS for 1 h at room temperature. After the final wash with PBS, the tissue was dissected by removing the modiolus. The epithelia were divided equally into three segments (apex, middle, and base). Specimens were mounted on slides with anti-fade mounting media. Control incubations were routinely processed without primary antibody treatments. Immunofluorescence signals for H3K9ac and HDAC2 in OHCs on surface preparations were quantified from original confocal images, each taken with a 63× −magnification lens under identical conditions and equal parameter settings for laser gains and photomultiplier tube gains, using Image J software (National Institutes of Health). The cochleae were fixed and stained simultaneously with identical solutions and processed in parallel. All the surface preparations were counterstained with PI for labeling OHC nuclei to identify the comparable parts for capture of confocal images. The immunofluorescence was measured in the cochlear surface preparations in 0.12-mm segments, each containing about 60 OHCs. The intensity of the background fluorescence was subtracted, and the average fluorescence per cell was then calculated. The relative fluorescence was quantified by normalizing the ratio of the average fluorescence of noise-exposed hair cells to the average fluorescence of the unexposed hair cells. Each condition was replicated in at least three different animals. For analysis of changes in H3K9ac distribution from the center of the nucleus to its periphery after SAHA treatment, signal quantification was performed using Image J software by calculating the signal intensity along a line crossing the nuclei’s axes. An experimenter who was blinded to the treatment conditions conducted the fluorescence measurements.

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Immunohistochemistry for Cochlear Cryosections Following decalcification with 4 % EDTA for 72 h at 4 °C, cochleae were processed for immunocytochemistry on frozen sections. Sections of 8-μm thickness were incubated in 0.3 % Triton X-100 in PBS for 30 min at room temperature. The specimens were washed three times with PBS and blocked with 10 % normal goat serum for 30 min at room temperature, followed by incubation with the primary antibody antiH3K9ac (1:50) at 4 °C for 48 h. After washing three times (10 min each), the tissues were incubated with the Alexa-488-conjugated secondary antibody at a concentration of 1:200 at 4 °C overnight in darkness. The specimens were washed three times with PBS and incubated with propidium iodide (PI) at 1 μg/mL in PBS for 1 h at room temperature. After the final wash with PBS, specimens were mounted on slides with anti-fade mounting media. Secondary antibody controls were routinely processed without primary antibody treatments. Immunolabeled cryosections were imaged with a Zeiss laser confocal microscope (Zeiss LSM 510).

Immunocytochemistry for Paraffin-Embedded Cochlear Sections Following decalcification with 4 % EDTA, each cochlea was transferred to 70 % ethanol and embedded in paraffin for sections. Five-micrometer sections were routinely deparaffinized in xylene, rehydrated in alcohol, and processed as follows: The sections were incubated with target retrieval solution (Dako #S2367) in a steamer (Oster #CKSTSTMD5-W) for 8 min and then with a 3 % hydrogen peroxide solution for 10 min and protein block (Dako #x0909) for 20 min at room temperature. Sections were first titrated with concentrations of each primary antibody (HDAC1, HDAC2, or HDAC3) to identify the appropriate dilution. Sections were then incubated with primary antibodies (HDAC1 1:12,000, HDAC2 1:1000, HDAC3 1:500) in a humidified chamber at 4 °C overnight, followed by biotinylated secondary antibodies (Vector) for 30 min and ABC reagent (Vector) for 30 min. Immunocomplexes of horseradish peroxidase were visualized by DAB (Dako) reaction, and sections were counterstained with hematoxylin before mounting. Images were taken with Zeiss microscope.

Surface Preparations and Diaminobenzidine (DAB) Staining of Myosin-VII-Labeled Hair Cells for Hair Cell Counts This procedure was described in our previous reports (Chen et al. 2012; Zheng et al. 2014; Yuan et al. 2015). Briefly, following decalcification, the cochleae were

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placed into 3 % hydrogen peroxide for 2.5 h to quench endogenous peroxidases. After incubation in a solution for blocking nonspecific antibody binding overnight at 4 °C, the tissues were incubated with a primary antibody (rabbit polyclonal anti-myosin VII at a 1:100 dilution) for 4 days at 4 °C on a nutator mixer, washed in PBS, and then incubated overnight at 4 °C with biotinylated goat anti-rabbit antibody at a 1:100 dilution. The specimens were rinsed again and then incubated in ABC solution (Vector Laboratories #PK4001) overnight. Following another washing, the cochleae were incubated in DAB for 3 h, as necessary for sufficient staining intensity, followed by washing to stop the DAB reaction. Finally, the cochleae were microdissected under a microscope into apical, middle, and basal segments and mounted on slides with Fluoromount-G mounting media. Images were taken with a Zeiss AxioCam MRc5 camera with Axioplan 2 imaging software under a Zeiss microscope. Images from the apex through the base of the DAB-stained preparations were captured using a 40× lens on the Zeiss microscope. Lengths of the cochlear epithelia were measured and recorded in millimeters. OHCs were counted from the apex to the base along the entire length of the mouse cochlear epithelium. The percentage of hair cell loss in each 0.5-mm length of epithelium was plotted as a function of the cochlear length as a cytocochleogram.

Extraction of Total Cochlear Protein Cochleae were rapidly removed and dissected in icecold PBS containing complete™ mini EDTA-free protease inhibitor cocktail tablets (Roche Diagnostic) at pH 7.4. To extract total protein, tissues from the cochleae of one mouse were homogenized in ice-cold radioimmunoprecipitation assay (RIPA) lysis buffer containing RIPA lysis buffer base (50 mM Tris-HCl, 1 % IGEPAL, 0.25 % Na-deoxycholate, 150 mM NaCl, 1 mM EDTA, 1 mM PMSF, 1 mM NaF) plus Phosphatase Inhibitor Cocktails II and III, and Roche Protease Inhibitor by using a glass/glass micro Tissue Grind pestle and vessel for 30 s. After 30 min on ice, tissue debris was removed by centrifugation at 15,000×g at 4 °C for 10 min, and the supernatants were retained as the total protein fractions. Protein concentrations were determined using the Bio-Rad Protein Assay dye reagent with bovine serum albumin (BSA) as a protein standard. Two cochleae from the same mouse were pooled for each sample.

Western Blot Analysis Protein samples (30 μg) were separated by SDS-PAGE. After electrophoresis, the proteins were transferred onto a nitrocellulose membrane (Pierce) and blocked

with 5 % solution of nonfat dry milk in PBS-0.1 % Tween 20 (PBS-T). The membranes were incubated with anti-H3K9ac (1:1000), anti-histone H3 (1:1000), or anti-HDAC2 (1:1000) antibodies at 4 °C overnight, and then washed three times (10 min each) with PBST buffer. Membranes were then incubated with an appropriate secondary antibody at a concentration of 1:2500 for 1 h at room temperature. Following extensive washing of the membrane, the immunoreactive bands were visualized by SuperSignal® West Dura Extended Duration Substrate or Pierce® ECL Western Blotting Substrate (Thermo Scientific). The membranes were then stripped and re-probed for GAPDH at a concentration of 1:20,000 as a control for sample loading. X-ray films of Western blots were scanned and analyzed using Image J software. First, the background staining density for each band was subtracted from the band density. Next, the probing protein/GAPDH ratio was calculated from the band densities run on the same gel to normalize for differences in protein loading. Finally, the difference in the ratio of the control and experimental bands was tested for statistical significance.

Statistical Analysis Data were analyzed using IBM SPSS Statistics Premium V21 and GraphPad software for Windows. The group size (n) in vivo was determined by the variability of measurements and the magnitude of the differences between groups. Based on our previous and current preliminary studies, we determined that five to eight animals per group provide sufficient statistical power. Data were statistically evaluated by one-way analysis of variance (ANOVA) or repeated measures ANOVA with Tukey’s multiple comparisons or posthoc testing using IBM SPSS Statistics or by two-tailed unpaired t-tests or one sample t tests using GraphPad Software (GraphPad Software Inc., San Diego, CA, USA). A p value G 0.05 was considered statistically significant. Data are presented as means ± SD or SEM.

RESULTS Noise Exposure Reduces H3K9ac Levels in the Nuclei of OHCs and Strial Marginal Cells in the Basal Turn Using the noise condition that leads to PTS in adult CBA/J mice (Chen et al. 2012; Zheng et al. 2014; Yuan et al. 2015), we first assessed the expression of H3K9ac using immunohistochemistry on cryosections 1 h after the exposure. Immunolabeling for H3K9ac decreased in the nuclei of OHCs (Fig. 1A, OC, white rectangle) and strial marginal cells (Fig. 1B, stria,

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Fig. 1.

Noise exposure reduces the levels of H3K9ac immunolabeling in the nuclei of OHC and strial marginal cells in the basal turn. A Cryosections of the basal turn of the organ of Corti (OC) revealed localization of H3K9ac in nuclei of OHCs (white rectangle) and supporting cells. The immunolabeling for H3K9ac in OHCs was decreased 1 h after 98-dB noise exposure. B The exposure also reduced immunolabeling for H3K9ac in strial marginal cells (Stria, arrow). C There was no alteration of immunolabeling for H3K9ac in spiral ganglion neurons between control and noise-

exposed mice. OC organ of Corti, Stria stria vascularis, SGN spiral ganglion neurons, red propidium iodide (PI)-labeled nuclei, green H3K9ac, n = 3. Scale bar = 10 μm. D and E Western blots detected a single protein band for H3K9ac and total histone H3 in cochlear homogenates with no difference between the controls and the noiseexposed mice examined 1 h after noise exposure. Data are presented as means + SD, n = 4.

arrow), without obvious alteration in the nuclei of spiral ganglion neurons (Fig. 1C, SGN). Additionally, we evaluated the levels of H3K9ac and total histone H3 on Western blots of total cochlear homogenates. Immunoblots showed a single 17-kDa band for each with no alteration in the band density between control and PTS noise-exposed cochleae (Fig. 1D, E). In order to quantify the immunolabeling for H3K9ac in OHCs, cochlear surface preparations were immunolabeled with an anti-H3K9ac antibody, revealing a reduction in H3K9ac in basal OHC nuclei (Fig. 2C), but not in the apical or middle turn OHC nuclei 1 h after PTS noise exposure (Fig. 2A, B). Quantification of H3K9acimmunolabeling in basal OHC nuclei confirmed a significant reduction by about 60 % 1 h after the

noise exposure (t4 = 10.883, p G 0.001). The amount of H3K9ac in basal OHCs was recovered slightly at 5 h, but did not recover further at 24 h after the noise exposure and remained significantly lower than those of control un-exposed mice at 5 h (t4 = 3.03, p = 0.0354) and 24 h (t4 = 4.812, p = 0.0084) (Fig. 2D). These results demonstrate that PTS-noise exposure results in modifications of H3K9ac in the nuclei of OHCs and strial marginal cells in the basal turn.

Noise Exposure Increases HDAC1, HDAC2, and HDAC3 in the Cochlear Tissues In order to probe for a potential cause for the decreased H3K9ac, we first evaluated the levels of

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Fig. 2. PTS noise exposure reduces immunolabeling for H3K9ac in nuclei of basal OHCs, but not in apical and middle OHCs on surface preparations. Representative images of OHC nuclei (red) from the apical turn (A) and middle turn (B) showed no alteration of H3K9ac (green) between control and the noise-exposed mice examined 1 h after the noise exposure. C Representative images of OHC nuclei from the basal turn showed a reduction of H3K9ac labeling 1 h after

the noise exposure compared to controls. D Quantification of H3K9ac immunolabeling in basal OHC nuclei displays significant reductions that are not recovered 24 h after PTS noise exposure. Control: n = 5, 1 h: n = 5, 5 h: n = 3, 24 h: n = 3. Data are presented as means + SD; n = 5, ***p G 0.001. All images are representative of five mice for each condition. Scale bar = 10 μm.

HDAC1, 2, and 3 using paraffin-embedded cochlear sections from the mid-modiolar region. Noise exposure caused an increase in HDAC1, 2, and 3 in the basal turn of the organ of Corti, including in the

nuclei of in OHCs, IHCs, and supporting cells such as Deiters cells (Fig. 3A, B, C; OC), in the spiral ganglion cells (Fig. 3A, B, C; SGN), and in endothelial cells of stria vascularis vessels (Fig. 3A, B and C, Stria), which

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Fig. 3. Noise exposure increases levels of HDAC1, 2, and 3 in cochlear tissues. A Paraffin-embedded cochlear sections showed an increase in immunostaining for HDAC1 in organ of Corti (OC), spiral ganglion cells (SGN) and the stria vascularis (Stria) 1 h after the noise exposure compared to control mice. B Immunostaining for HDAC2 increased in the organ of Corti (OC), spiral ganglion cells (SGN), and the stria vascularis (Stria) 1 h after the noise exposure compared to

control mice. C Immunostaining for HDAC3 increased in the organ of Corti (OC), spiral ganglion cells (SGN), and the stria vascularis (Stria) 1 h after the noise exposure compared to control mice. Blue hematoxylin counterstaining for nuclei, brown DAB staining of immunolabeled HDAC1, 2, and 3. Images were taken from basal turn, n = 3. Scale bar = 10 μm.

appeared to be related to affects in the microvasculature. In order to quantify the changes of HDACs in the cochlear tissue, we selected HDAC2 to analyze using Western blots of total cochlear homogenates because of its known role in disease pathology. Immunoblots showed two bands, a 60-

kDa band specific for HDAC2 and a 25-kDa band for mouse IgG light chain. Quantification of the 60-kDa band densities revealed a significant twofold increase at 1 h after the exposure (Fig. 4, t5 = 4.244, p = 0.0132). These results demonstrate that PTS-noise exposure does provoke elevation of

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Fig. 4.

Noise exposure increases HDAC2 levels in basal OHCs and the cochlear tissue homogenates. A OHC nuclei (red) from the base show increased immunolabeling for HDAC2 (green) 1 h after noise exposure compared to control mice. Red propidium iodide (PI)labeled nuclei. Images are representative of five mice for each condition. Scale bar = 10 μm. A’ Quantification of HDAC2associated immunofluorescence in basal OHCs confirmed a significant increase 1 h after the noise exposure. Data are

presented as means + SD; n = 5, *p G 0.05. B Western blots of total cochlear homogenates showed an increased 60-kDa HDAC2 band after the noise exposure and no changes of the 25-kDa IgG band. B’ Quantification of the 60-kDa band density confirmed a significant increase. GAPDH was used as sample loading control. Data are presented as means + SD; n = 6, *p G 0.05.

HDACs in the nuclei of all cell types of cochlear tissues.

SAHA Treatment Attenuates PTS-Noise-Induced Loss of OHCs and Subsequent Hearing Loss

Silencing HDAC1, 2, or 3 Alone Does Not Attenuate Noise-Induced Permanent Threshold Shifts As noise exposure increased HDAC1, 2 and 3, we tested if silencing HDAC1, 2, or 3 alone in CBA/J mice could attenuate NIHL as assessed by auditory brainstem response measurements. First, we evaluated HDAC2 siRNA (siHDAC2). Local delivery of 0.6-μg siHDAC2 resulted in diminished expression of HDAC2 protein in OHCs 72 h after silencing treatment (Fig. 5A). Quantitative analysis of the silencing efficiency showed that the expression of HDAC2 was reduced by 30 % in OHCs compared to control cochleae that were treated with scrambled control siRNA (Fig. 5A’, t2 = −10.645, p = 0.009). However, Pretreatment with siHDAC2 did not attenuate PTSnoise-induced threshold shifts at any of the measured frequencies (8, 16, or 32 kHz, Fig. 5B). Furthermore, pretreatment with either siHDAC1 or siHDAC3 also did not alter PTS-noise-induced threshold shifts at any of the test frequencies. These results indicate that blockade of HDAC1, 2, or 3 alone in cochlear tissues is not sufficient to reduce PTS.

Two weeks after the PTS noise exposure, OHC loss followed a base-to-apex gradient beginning at 3.5 mm from the apex and increasing towards the base, and reached 100 % at 5–5.5 mm from the apex (Fig. 6B, C). Treatment with SAHA significantly decreased noise-induced OHC loss (F 1, 12 = 10.875, p = 0.006). OHC loss was reduced by 40 % at 4 mm (p = 0.032), 50 % at 4.5 mm (p = 0.002), and 20 % at 5 mm (p = 0.059) from the apex. Consequently, auditory threshold shifts were significantly different between the three groups (PTS noise only, DMSO vehicle treatment plus PTS noise, and SAHA treatment plus PTS noise) at 16 kHz (Fig. 6A, F2, 36 = 8.707, p = 0.001) 14 days after the noise exposure, but not at 32 kHz. Treatment with SAHA attenuated PTS by about 20 dB at 16 kHz compared to vehicle-treated mice (p = 0.014), whereas treatment with SAHA alone did not affect auditory function. These results imply that treatment with the HDAC inhibitor SAHA attenuates PTS. SAHA, however, was ineffective at preventing hearing loss after a noise paradigm known in our lab to induce more severe permanent hearing loss.

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Treatment with SAHA Temporarily Alters the Distribution of Histone Acetylation in the Nuclei of OHCs Since SAHA treatment attenuated hair cell loss and NIHL, we also assessed if treatment with SAHA in vivo alters histone acetylation in OHCs. The levels of H3K9ac were examined 1 and 3 h after a single dose of SAHA (50-mg/kg IP injection) without noise exposure. SAHA treatment changed the distribution of H3K9ac in the nuclei of OHCs 1 h after the treatment, but not at the 3-h time point. (Fig. 7A). One hour after SAHA treatment, there was clear enrichment of H3K9ac at the nuclear periphery that was not observed in the vehicle-only group. We quantified the immunolabeling of H3K9ac across the OHC nuclei (Fig. 7B, enlarged example of a single OHC). The pixel profiles across the center of OHC nuclei show redistribution of H3K9ac from the center to the periphery after SAHA treatment (Fig. 7B’). This observation suggests that SAHA treatment changes the distribution of H3K9ac in the OHC nuclei.

DISCUSSION Post-translational modification of histones alters their interaction with DNA and nuclear proteins, influencing gene expression and cell fate. Our results show

Fig. 5. Pretreatment with siHDAC2 reduces HDAC2 expression in OHCs. (A) Local delivery of 0.6 μg HDAC2 siRNA (siHDAC2) decreased HDAC2 expression in OHCs 72 h after the delivery compared to control siRNA (siControl)-treated mice. Images were taken from the upper basal turn. Scale bar = 10 μm. A’ Quantification of HDAC2-associated immunofluorescence confirmed a

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that noise exposure decreases H3K9ac in the nuclei of OHCs and marginal cells of the stria vascularis, suggesting that epigenetic modification of the histone H3 tail is involved in noise-induced OHC loss and NIHL. OHCs play a specific role in refining sensitivity and frequency selectivity of the mechanical vibrations of the cochlea and are referred to as cochlear amplifiers. The marginal cells are involved primarily in ion transport, generating a positive transepithelial potential by secreting potassium ions (Wangemann et al. 1995). They are important in maintaining the endocochlear potential. However, the detailed alterations in signaling and other molecular changes resulting from the decrease in H3K9ac in the nuclei of OHCs and strial marginal cells after the noise exposure remain unknown. There may be modulation of gene expression associated with noise-induced loss of OHCs and hearing loss. The two opposing pathways of histone acetylation and deacetylation are regulated by HATs and HDACs. The immunoreactivity of the three HDACs was localized entirely within the nuclei of cochlear cells of control mice without noise exposure, but had weak staining in the cytosol of some of the cochlear cells after the noise exposure regardless of the dilutions of the antibodies. This rules out non-specific staining and might be due to increased transcription of HDACs in the cytosol to meet the cellular needs under stress. Elevation of HDAC1, 2, and 3 levels in

significant decrease within OHCs. Data are presented as means + SD; n = 3, **p G 0.01. B Noise-induced auditory threshold shifts were not different between siControl and siHDAC2 treatment 14 days after the exposure. Data are presented as means + SD; n = 6.

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the nuclei of OHCs indicates that the decreased H3K9ac levels in OHCs are likely caused by activation of HDACs. However, HDACs are also augmented in other cochlear tissues, such as the stria vascularis and spiral ganglion neuron cells, and are significantly increased in total cochlear tissue homogenates, whereas H3K9ac levels remain steady in total cochlear tissue homogenates. The other tissues in cochlear homogenates dilute the population of cells that respond to H3K9ac, implying distinct responses to PTS noise in different cell types. Additionally, the effect on HAT levels in OHCs after the noise exposure is not clear. Although we attempted to immunolabel HAT/p300 with surface preparations and cochlear paraffin sections, the large amount of non-specific immunolabeling with the antibody made it difficult to draw any conclusions. Our silencing results suggest that blocking only one isoform of HDACs is not sufficient to attenuate noiseinduced permanent hearing loss. However, systemic administration of HDAC inhibitor SAHA attenuates PTS (98 dB SPL) at 16 kHz, which is in line with a recent publication (Wen et al. 2015). Additionally, the activity of HDACs is not restricted to the core histones, it also targets non-histone substrates, including transcription factors, chaperones, and structural proteins (Spange et al. 2009). The attenuation of PTSnoise-induced OHC loss and subsequent NIHL by treatment with SAHA indeed may not rely solely on the blockade of HDAC activity on histones, although HDAC inhibitors act primarily through modulation of chromatin condensation by regulation of histone acetylation. SAHA has a broad spectrum of functions (Iyer et al. 2010; Layman and Zuo 2014), including reducing apoptotic-signaling pathways, which are involved in NIHL (Pirvola et al. 2000; Hu et al. 2002; Wang et al. 2003; Zheng et al. 2014). Thus, it is highly likely that attenuation of NIHL by SAHA treatment is due, at least to some extent, to targeting of nonhistone core proteins. Although this study does not have such explicit results, SAHA treatment reduces ischemic injury in the mouse brain by increasing the expression of protective genes such as Hsp70 and Bcl2 (Faraco et al. 2006). The HDAC inhibitor valproic acid enhances NF-κB and protects neurons from hypoxia-induced apoptosis in vitro (Li et al. 2008). Furthermore, SAHA treatment attenuates acute ototoxicity by activating the NF-κB pathway (Layman et al. 2015), implying that HDAC inhibitors act on multiple targets (Choudhary et al. 2009). Treatment with SAHA alone changes the distribution of H3K9ac from the center of nuclei to the periphery in basal OHCs, in line with the function of other HDAC inhibitors such as trichostatin A and sodium butyrate, which are found to increase the levels of acetylated histones preferentially at the

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nuclear periphery in vitro (Gilchrist et al. 2004; Bartova et al. 2005; Rada-Iglesias et al. 2007). The movement of H3K9ac to the periphery of OHC nuclei by SAHA treatment supports the notion of alterations in gene expression, as trichostatin A alters the

Fig. 6. Treatment with SAHA reduces noise-induced OHC loss and attenuates auditory threshold shifts. A SAHA treatment attenuated auditory threshold shifts at 16 kHz 14 days after the exposure compared to vehicle control mice. Data are presented as means ± SEM; n = 13 in each group of PTS noise only, PTS noise + vehicle, and PTS noise + SAHA, *p G 0.05. B SAHA treatment appeared to reduce PTS noise-induced OHC loss. The representative images were taken in the basal turn 4 mm from the apex of the cochlear epithelium. OHC 1, 2, and 3 label the three rows of OHCs. Scale bar = 10 μm. C Quantification revealed that SAHA treatment reduced OHC loss. OHCs were counted along the entire length of the cochlear epithelium from the apex to base. Data are presented as means ± SEM; n = 7, *p G 0.05; **p G 0.01.

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Fig. 7.

SAHA treatment temporarily alters the distribution of H3K9ac in the nuclei of OHCs. A Representaive images revealed changes in H3K9ac (green) distribution in nuclei of OHCs (red) 1 h and 3 h after SAHA treatment. Images were taken from the basal turn; red: propidium iodide (PI)-labled nuclei of OHCs, green: H3K9ac, n = 5. Scale bar = 10 μm. B Enlarged images of OHC nuclei

revealed redistribution of H3K9ac from the center to the periphery 1 h after SAHA treatment. Scale bar = 5 μm. B’ The graph represents quantification of H3K9ac immunolabeling (green) along the yellow line drawn across nuclei of OHCs from vehicle- and SAHA-treated mice, n = 5.

expression of genes that are responsible for regulating intracellular calcium homeostasis, synaptic plasticity, and apoptosis after cisplatin treatment (Wang et al. 2013). It is a common notion that SAHA-treatmentincreased histone acetylation is dose dependent. Our results are in agreement with the recent publication showing that SAHA treatment increased histone acetylation in OHCs and cochlear tissues at SAHA doses of 100 mg/kg body weight, but not at 50 mg/kg body weight (Layman et al. 2015). In summary, our results reveal epigenetic mechanisms involved in the pathogenesis of noise-induced loss of OHCs and subsequent NIHL. This study points out a possible strategy for development of preventive compounds against NIHL by blockade of HDAC activity. Although SAHA treatment attenuates PTSnoise-induced OHC loss and NIHL, we should emphasize that a more complete picture of the molecular mechanisms underlying the effects of

SAHA treatment should be elucidated in the future to further explore its effectiveness as an oto-protective agent. In addition, we also need to consider further examination of the toxicity of the compound in order to determine if it is suitable for prevention or treatment of NIHL.

ACKNOWLEDGMENTS The research project described was supported by grant R01 DC009222 from the National Institute on Deafness and Other Communication Disorders, National Institutes of Health. This work was conducted in the WR Building at MUSC in renovated space supported by grant C06 RR014516. Animals were housed in MUSC CRI animal facilities supported by grant C06 RR015455 from the Extramural Research Facilities Program of the National Center for Research Resources. We thank MUSC Biorepository and Tissue Analysis Shared Resource for

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technical assistance for cochlear paraffin sections. We also thank Andra Talaska for proofreading of the manuscript.

COMPLIANCE WITH ETHICAL STANDARDS All research protocols were approved by the Institutional Animal Care and Use Committee at the Medical University of South Carolina (MUSC). Animal care was under the supervision of Division of Laboratory Animal Resources at MUSC.

Conflict of Interest The authors declare that they have no conflict of interest. The data contained in the manuscript have not been previously published and have not been submitted elsewhere, and will not be submitted elsewhere while under review. All authors have reviewed the contents of the manuscript, approve of its contents, and validate the accuracy of the data.

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