Introduction: Noise-induced hearing loss is one of the most frequent recognized occupational diseases. The time course of the involved pathologies is still under investigation. Several studies have demonstrated an acute damage of the sensory tissue, but only few experiments investigated the degeneration of (type I) spiral ganglion neurons (SGNs), representing the primary neurons in the auditory system. The aim of the present study was to investigate the time course of SGN degeneration within a 7-day period after traumatic noise exposure starting immediately after trauma. Methods: Young adult normal hearing mice were noise exposed for 3 h with a broadband noise (5–20 kHz) at 115 dB SPL. Auditory threshold shift was measured by auditory brainstem recordings, and SGN densities were analyzed at different time points during the first week after acoustic trauma. Results: Significant reduction of SGN densities was detected and is accompanied by a significant hearing loss. Degeneration starts within hours after the applied trauma, further progressing within days post-exposure. Discussion: Early neurodegeneration in the auditory periphery seems to be induced by direct overstimulation of the auditory nerve fibers. SGN loss is supposed to be a result of inflammatory responses and neural deprivation, leading to permanent hearing loss and auditory processing deficits.

Hearing loss affects more than 5% of the worldwide population and noise exposure is a major cause of auditory dysfunction [1]. Particularly in industrial countries, noise-induced hearing loss (NIHL) is one of the most frequent recognized occupational diseases, is suspected to be responsible for audiological symptoms like tinnitus or hyperacusis, and is frequently followed by emotional and cognitive psychic disorders [2‒4]. Although a lot of research has focused on the physiological and anatomical effects of NIHL, the time course of the involved pathologies and its underlying mechanisms are still under investigation.

Several studies in the peripheral auditory system, particularly the cochlea, were able to show an acute damage of the sensory tissue including the inner and outer hair cells, the supporting cells, and the stria vascularis. Immediate pathologies are induced by mechanical forces and electrochemical hyperexcitation and are followed by early necrotic and apoptotic cell death pathways [5‒11]. A detailed investigation of the temporal structure of sensory degeneration has been performed by Bohne and colleagues, showing acute noise-induced loss of hair cells and supporting cells, further progressing over days, weeks, and months [12].

A few experiments investigated the degeneration of (type I) spiral ganglion neurons (SGN) over time, representing the primary neurons in the auditory system. The strength and time course of neurodegeneration are correlated with the intensity and duration of acoustic trauma and could be even present upon moderate sound exposure. It has been particularly demonstrated that apoptosis-related structural changes are present within a few days after noise exposure. However, it is still unclear if the observations are due to an early, direct impact of overexcitation or related to deprivation-induced neuroplasticity in response to destruction of cochlear tissue or excitation-induced degeneration of primary auditory afferences, i.e., the loss of synaptic connections and dendritic processes between inner hair cells and SGNs [13‒17].

Therefore, the aim of the present study was to investigate the time course of SGN degeneration within a 7-day period after traumatic noise exposure starting immediately after trauma. The results of the experiments gain insight into the time dependency of structural changes in primary auditory neurons and could help develop possibilities for clinical intervention for the preservation of auditory structures and hearing function.

Animals

Twenty-six young adult (30–40 days of age) female normal hearing mice (Mus musculus, NMRI strain) with a fully developed auditory system were used in the present study. Age, sex, and mouse strain were chosen according to earlier investigations [18, 19]. The experimental protocol was approved by the governmental commission for animal studies (LaGeSo, Berlin, Germany; Approval No. G0416/10). Experiments were carried out in accordance with the EU Directive 2010/63/EU on the protection of animals used for scientific purposes.

Noise Exposure

The noise exposure paradigm was similar as described in earlier publications of our group [19, 20]. Animals were exposed to a broadband flat spectrum noise (3 h, 5–20 kHz, 115 dB SPL peak-to-peak) under anesthesia (6 mg/kg xylazine and 60 mg/kg ketamine) in a soundproof chamber (80 cm × 80 cm × 80 cm, minimal attenuation 60 dB). An amplifier (Tangent AMP-50; Aulum, Denmark) and a DVD player were connected to loudspeakers (HTC 11.19; Visaton, Germany) placed above the animal´s heads. A sound level meter (Voltcraft 329; Conrad Electronic, Germany) was placed next to the animal’s ear to calibrate SPL. A heating pad (Thermolux CM 15W; Acculux, Germany) was placed below the animals to keep body temperature constant at 37°C during video camera-controlled anesthesia.

Auditory Brainstem Recordings

Hearing thresholds were measured in 6 animals in a separate subset of experiments under anesthesia at different time points after noise exposure by auditory brainstem recording (ABR; pre-exposure, 1 day post-exposure, 7 days post-exposure, respectively). Subdermal needle electrodes were placed at the vertex (reference), mastoid (active), and leg (ground). Tone burst stimuli of 3-ms duration and using a Blackman envelope (1.5 ms raise/fall time) were delivered through a free-field high-frequency transducer placed 10 cm above the animal’s head within a soundproof chamber. Recording and data acquisition parameters used a 31.1 bursts per second stimulus rate having an alternating phase. There were 1,024 sweeps made per frequency and level. The recording parameters included an amplifier gain of 100 k, recording via a bandpass filter of 100–3,000 Hz. The sampling rate used to collect electrophysiological data was 32 kHz. Stimulation and recording were controlled using the SmartEP software version 5.33 (Intelligent Hearing Systems, FL, USA). The hearing threshold was estimated by a stepwise decrease of the stimulus intensity until no electrophysiological response was visually detectable. Threshold shift was calculated for each subject and group data are given as mean hearing loss ± standard deviation.

Experimental Groups

Different groups of animals were investigated at various points in time after noise exposure. Experimental groups were chosen in accordance with earlier studies after single noise exposure by our group [18‒20]. At day 0, 19 animals of the experimental groups were noise exposed and randomly assigned to one of the following groups: acute group (investigated immediately after noise exposure, n = 3), 6 h group (investigated 6 h after the end of the noise exposure, n = 3), 24 h group (investigated 24 h after the end of the noise exposure, n = 5), and 7-day group (investigated 7 days after the end of the noise exposure, n = 4). At each experimental day, noise trauma was always started in the morning to avoid different noise impact due to diurnal variation in noise susceptibility [21]. Five mice were used as unexposed control animals.

Tissue Preparation and Histology

At the day of experiments, animals were perfused via the left heart chamber with a fixation solution (4% paraformaldehyde). The temporal bones including the cochlea were removed carefully from the skull and transferred into 0.2 m EDTA for 12 days to decalcify the tissue. Afterward, cochleae were extracted, embedded in paraffin and micro-sliced (6 μm) in the transversal plane using a rotation microtome (Leica RM2235, Leica Biosystems, Germany). Slices were stained on object slides using a modified hemalum-eosin staining. For removal of the paraffin from the tissue, slices were washed twice for 5 min in Rotihistol (Carl Roth, Karlsruhe, Germany) and rehydrated in a descending ethanol series (90%, 70% ethanol) and distilled water (5 min each). Nuclei were stained for 5 min in a hemalum solution and differentiated under flowing water for 10 min. Afterward, the slices were stained for 1 min in 0.1% eosin, washed shortly in distilled water and differentiated in 70% Ethanol. Tissue was dehydrated in an ascending ethanol series (90% ethanol, isopropanol) and kept in Rotihistol (Carl Roth, Karlsruhe, Germany) for storage. Object slides were mounted with Roti Histokitt (Carl Roth, Karlsruhe, Germany).

Image Analysis

The stained tissue was microscopically magnified (Zeiss AxioLab A1, Carl Zeiss, Germany) and pictures from each Rosenthal canal including the type I SGN were digitized by a digital camera (Zeiss ICc1, Carl Zeiss, Germany). For image analysis, each Rosenthal canal in the modiolus was surrounded to calculate the specific area. SGN were counted manually and the SGN density was calculated. SGN densities were normalized to the mean Rosenthal canal area observed before statistical comparison (mean area: 0.0383 mm2). Approximately 10 Rosenthal canals were analyzed per animal. Values are given as mean ± standard deviation.

Despite total SGN densities, basal (high-frequency area) and apical (low-frequency area) Rosenthal canals were additionally analyzed separately to give insight into frequency-specific differences in SGN densities over time. FIJI (ImageJ version 1.53c, National Institutes of Health, USA) software was used for image analysis.

Statistics

Data of the experimental groups and control group were analyzed statistically with SPSS software (SPSS version 25, IBM, USA). Data distribution was tested using the Shapiro-Wilk test. For normally distributed data, t test was used to compare the data between each experimental group and the control group. U test was applied if data were not normally distributed. Level of significance for all tests was set to p < 0.05, and the Bonferroni correction was applied to account for multiple comparisons.

Auditory Threshold Shift

Shifts in hearing thresholds in the animals in response to noise treatment was indicated by a significant ABR threshold shift in the investigated frequency range between 4 and 32 kHz. Frequency-dependent mean hearing loss ranged between 53 and 61 dB 1 day after noise exposure. Seven days following acoustic trauma, threshold shift was between 42 and 50 dB compared to the hearing level before noise exposure. Threshold shift showed a highly significant difference for all frequencies and for both time points compared to the pre-exposure levels (p < 0.001). However, hearing thresholds showed a slight recovery within 1 week post-exposure, which was even significant at lower frequencies (4 kHz: p = 0.007; 8 kHz: p = 0.012; 12 kHz: p = 0.002; 16 kHz: p = 0.031; 20 kHz: p = 0.007; 24 kHz: p = 0.02; 28 kHz: p = 0.123; 32 kHz: p = 0.038) (Fig. 1).

Fig. 1.

Mean (±SD) auditory threshold shift across the measured frequency range in noise-exposed animals at different time point post-exposure. Asterisks represent statistically significant differences after noise trauma compared to pre-exposure hearing thresholds. Black asterisks indicate significant differences between pre- and post-exposure hearing levels. Gray asterisks indicate significant recovery in auditory threshold shift from day 1 to day 7 after trauma (***p < 0.001; **p < 0.01; *p < 0.05).

Fig. 1.

Mean (±SD) auditory threshold shift across the measured frequency range in noise-exposed animals at different time point post-exposure. Asterisks represent statistically significant differences after noise trauma compared to pre-exposure hearing thresholds. Black asterisks indicate significant differences between pre- and post-exposure hearing levels. Gray asterisks indicate significant recovery in auditory threshold shift from day 1 to day 7 after trauma (***p < 0.001; **p < 0.01; *p < 0.05).

Close modal

SGN Densities

Mean density of SGNs was analyzed in the different experimental groups (noise-exposed and investigated at different time points within 2 weeks post-exposure). The resulting data were normalized on the mean Rosenthal canal area to ensure comparability of the results and statistically compared to the unexposed, normal hearing control group.

Total SGN density was shown to be reduced in the noise-exposed animals in relation to the controls, being significant already immediately after trauma. Mean (±SD) SGN number (normalized on mean RC area) was 82.2 ± 17.8 in the control group, whereas it has been 74.0 ± 11.6 in the acute group (p = 0.005). Significant reduction of SGN densities in relation to the controls were also present in the 6 h group (72.7 ± 11.4; p = 0.001), as well as in the 24 h group (67.7 ± 12.3; p < 0.001) and the 7 days group (67.0 ± 21.6; p < 0.001) (Fig. 2).

Fig. 2.

Mean total spiral ganglion neuron (SGN) density (basal and apical turn) of the control and noise-exposed groups (±SD). SGN numbers have been normalized to the mean analyzed Rosenthal canal area across all subjects. Asterisks indicate statistically significant difference of the experimental groups to the control group (***p < 0.001; **p < 0.01).

Fig. 2.

Mean total spiral ganglion neuron (SGN) density (basal and apical turn) of the control and noise-exposed groups (±SD). SGN numbers have been normalized to the mean analyzed Rosenthal canal area across all subjects. Asterisks indicate statistically significant difference of the experimental groups to the control group (***p < 0.001; **p < 0.01).

Close modal

Mean SGN densities in the basal part cochlear (i.e., the high-frequency region) between the experimental and control groups, the effect of noise exposure on mean SGN densities is comparable to the total SGN data. The difference of the noise-exposed groups was statistically significant at 24 h (68.5 ± 12.5; p < 0.001) and 7 days (67.5 ± 21.9; p = 0.005) after exposure compared to unexposed controls (81.0 ± 18.3). There was also a reduction of SGN densities in the acute group (74.2 ± 11.9; p = 0.035) and in the 6 h group (75.3 ± 10.5; p = 0.059) in relation to the control group. However, the effect was less pronounced in these data subset of these groups and statistical significance was not reached (Fig. 3).

Fig. 3.

Mean spiral ganglion neuron (SGN) density (basal turn only) of the control and noise-exposed groups (±SD). SGN numbers have been normalized to the mean analyzed Rosenthal canal area across all subjects. Asterisks indicate statistically significant difference of the experimental groups to the control group (***p < 0.001; **p < 0.01).

Fig. 3.

Mean spiral ganglion neuron (SGN) density (basal turn only) of the control and noise-exposed groups (±SD). SGN numbers have been normalized to the mean analyzed Rosenthal canal area across all subjects. Asterisks indicate statistically significant difference of the experimental groups to the control group (***p < 0.001; **p < 0.01).

Close modal

When looking at the SGN densities in the apical cochlear region (i.e., the low-frequency area) between the experimental and control groups, it became evident that the effect is more pronounced compared to the basal cochlear turn, meaning that the relative reduction of SGN densities is higher. The difference between the noise-exposed groups was statistically significant at 6 h (61.4 ± 7.9; p < 0.001), 24 h (61.8 ± 8.3; p < 0.001), and 7 days (63.0 ± 19.3; p = 0.0014) after exposure compared to unexposed controls (89.9 ± 13.3). There was also a strong reduction of SGN densities in the acute group in relation to the control level, although it did not reach statistical significance (71.5 ± 9.0; p = 0.042) (Fig. 4).

Fig. 4.

Mean spiral ganglion neuron (SGN) density (apical turn only) of the control and noise-exposed groups (±SD). SGN numbers have been normalized to the mean analyzed Rosenthal canal area across all subjects. Asterisks indicate statistically significant difference of the experimental groups to the control group (***p < 0.001; **p < 0.01).

Fig. 4.

Mean spiral ganglion neuron (SGN) density (apical turn only) of the control and noise-exposed groups (±SD). SGN numbers have been normalized to the mean analyzed Rosenthal canal area across all subjects. Asterisks indicate statistically significant difference of the experimental groups to the control group (***p < 0.001; **p < 0.01).

Close modal

In the present study, we investigated the development of SGN densities in mice after PTS-inducing noise exposure. Our results show a significant reduction of SGN densities after trauma, starting immediately after the applied trauma and further progressing within a few hours post-exposure. The effect is present in the high-frequency basal cochlear turn as well as in the low-frequency apical part, whereby it is more pronounced in the apical region.

Several studies have demonstrated the damaging effect of acoustic overexposure on cochlear structures, including hair cells, supporting cells, the stria vascularis and SGNs [6, 7, 9‒11, 22]. Concerning acute destruction (i.e., direct, immediate damage in response to noise exposure), outer and inner hair cells have been localized to serve as the main targets for noise impact and being highly responsible for a permanent shift in auditory thresholds [12]. Neurodegeneration in the auditory nerve as well as in the central auditory system has been characterized as subsequent pathologies due to deprived input from the sensory cells [16, 23].

However, it remains unclear in how far the observed changes in auditory thresholds in the present study are directly correlated with the significant SGN degeneration. Although several studies support the idea of a connection between hearing loss and SGN densities, auditory thresholds recover slightly within the first day after acoustic trauma. Therefore, different effects are supposed to contribute to the hearing loss, whereby some of them are transient only. Recent studies have identified reactive oxygen species and inflammatory responses in the inner ear as key factors at the acute stage of NIHL, whereby the upregulation of cholesterol metabolic pathways in the hair cells seem to play an important role in auditory threshold recovery within 1 week post-exposure. In permanent auditory threshold shift, cell loss in the organ of Corti and the spiral ganglion seem to represent correlates of NIHL [24‒26]. Recent work by our group was able to demonstrate an early effect of acoustic trauma on neuronal structures, showing acute apoptosis-related degeneration in several structures of the ascending auditory system, particularly located in the auditory brainstem [18‒20].

With the results of the current experiments, we can extend this assumption toward the peripheral auditory neurons. Loud sound exposure is leading to overexcitation in the auditory nerve, which is thought to induce apoptosis in the affected neuronal structures. Recently, we pointed out that intrinsic apoptotic mechanisms participate in the initial period of degeneration in the cochlear nucleus. This led us to the conclusion that acute overstimulation activates this pathway by glutamate-induced excitotoxicity [27]. However, an additional contribution of extrinsic apoptosis (e.g., by activating the TNF-alpha cascade) is supposed to play a role during this process as well.

Inflammatory responses in the inner ear are also participating during pathological effects after noise-induced injury. Upregulation of several pro-inflammatory cytokines and chemokines as well as macrophages support this idea [28‒30].

The long-lasting findings are in line with earlier investigations, showing degeneration on the level of the auditory nerve within days and weeks after traumatic injury of the inner ear [13‒17]. However, the present study is to our knowledge the first one closely monitoring the early effects of acoustic overexposure.

It has already been shown that glutamate, the main excitatory neurotransmitter in the inner ear, plays a crucial role in cochlear histopathologies. Excessive glutamate release at the synaptic contacts between the inner hair cells and the neurites of the SGNs lead to swelling and disruption of the synaptic connectivity due to calcium overload in the postsynaptic cells and leading oxidative stress in the impaired cellular structures by formation of reactive oxygen species, supporting our hypothesis of acute excitotoxic neurodegeneration on the SGN level [17, 31].

In contrast to temporary impact on auditory processing, neuronal degeneration is irreversible, and the observed pathologies are not able to recover. Therefore, our findings are of great significance for therapeutic treatment of acoustic overexposure. This is of particular importance regarding the timing of clinical intervention after traumatic noise injury since the results of the present study indicate only a short time window to reduce the observed neuropathologies in the auditory periphery. Treatments (e.g., pharmacological applications using antiapoptotic and anti-inflammatory drugs, physical intervention by near-infrared-light treatment) should therefore be applied as earliest possible after trauma [32, 33].

The larger effect in the apical (low-frequency) region of the cochlea in our study is probably due to the frequency band of the applied trauma, which is located in the low-frequency area of the mouse hearing range (i.e., in the apical turn) [34]. One reason for the missing statistical significance of the acute effect in the subdivision analysis compared to total SGN density comparison could be related to the lower sample size after separating the data between the different frequency regions.

The results are also of clinical relevance from a prosthetic point of view. The loss of SGNs and their peripheral neurites would largely influence the outcome of cochlear implant supply. The degeneration of the target structures responsible for electrical stimulation could result in reduced temporal and spatial resolution at the stimulation sites and has a negative influence on perception of environmental sound sources [35]. Early interventions would lead to a better electrode-neuron interface and an improved electrical transmission with a strong therapeutic benefit for the patients.

The present study was supported by the Deutsche Forschungsgemeinschaft DFG (GR 3519/3-1).

The experimental protocol was approved by the governmental commission for animal studies (LaGeSo, Berlin, Germany; Approval No. G0416/10). Experiments were carried out in accordance with the EU Directive 2010/63/EU on the protection of animals used for scientific purposes.

The authors have no conflicts of interest to declare.

This study was not supported by any sponsor or funder.

Conceptualization: M.G., A.E., and D.B.; data analysis: M.G., T.M., S.V., and D.B.; investigation: M.G., T.M., S.V., and F.F.; methodology: M.G., T.M. F.F., and D.B.; project administration and writing: M.G. and D.B.

The analyzed data of the present study, which underpin the results, are all included in the manuscript. The raw data are provided by the authors without reservation. Inquiries can be directed to the corresponding author [M.G.].

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