This work was supported by two grants from JSPS KAKENHI (Grant Numbers 21K09573 (K.M.) and 23K15901 (T.K.)).

All experimental procedures were approved by the Institutional Animal Care and Use Committee of the National Defense Medical College (approval #18050) and performed in accordance with the guidelines of the National Institutes of Health and the Ministry of Education, Culture, Sports, Science, and Technology of Japan. All efforts were made to minimize the number of animals and their suffering.
1. Animals
<ol>
	<li>Use 8-week-old male CBA/J mice to follow this protocol. Before the experiment, subject the mice to a hearing function test and endoscopic observation of the tympanic membrane to ensure normality.</li>
	<li>Divide 27 CBA/J mice into three groups: (1) 2.0 J/cm2 exposed group (n = 9 mice); (2) 2.25 J/cm2 exposed group (n = 9 mice); and (3) 2.5 J/cm2 exposed group (n = 9 mice). Remove all the ears for assessment 1 month after the LISW exposure.</li>
</ol>
2. Experimental settings of LISW exposure
<ol>
	<li>The laser target is a black, natural rubber disk, 10 mm in diameter and 0.5 mm thick. To increase the LISW impulse, use an acrylic resin welding adhesive to bond a 1.0 mm thick, transparent, polyethylene terephthalate sheet (PET) to the top of the target area. Irradiate a 532 nm Q-switched Nd: YAG laser to generate the LISW behind the target (Figure 1A).</li>
	<li>Focus the laser pulse with a plano-convex lens to a 3.0 mm diameter spot on the laser target.</li>
	<li>Use the LISW irradiation to generate plasma at the bonding surface of the two materials and vaporize the rubber (plasma-mediated ablation), leaving vaporized rubber in the cavity.</li>
	<li>Use a hydrophone to measure the pressure wave of LISW at 1.0 mm underwater, not in living tissue. Place a 0.25 mm diameter fiber optic hydrophone under the black rubber 1.0 mm below the water surface to record the LISW pressure waveforms and measure them using a digital oscilloscope. 
	NOTE: The pressure waveforms showed stable characteristics with similar maximum pressure and impulse as shown in Figure 1B.</li>
	<li>Perform all animal procedures under general anesthesia using intraperitoneal injections of 1 mg/kg medetomidine hydrochloride and 75 mg/kg ketamine. Apply an ophthalmic ointment to both eyes of the mouse to prevent drying and provide heat support.</li>
	<li>Carefully shave the postauricular regions to avoid retaining the trapped air in the fur. Fix the mice on a plate and position the postauricular regions in the focal area of the LISW in a vertically upward direction.</li>
	<li>Attach a black rubber target percutaneously to the postauricular region of the mouse ear. To ensure acoustic impedance matching, use an ultrasound conductive gel between the laser target and the skin surface.</li>
	<li>Apply a single LISW pulse to the cochlea via the temporal bone. Set the outputs of the laser pulses to three energy densities: 2.0 J/cm2 , 2.25 J/cm2, and 2.5 J/cm2.</li>
</ol>
3. Cochlear function test
NOTE: Auditory brainstem response (ABR) tests were performed as previously reported14,15.
<ol>
	<li>Perform the ABR measurement 1 day before and 1 day and 1 month after LISW exposure.</li>
	<li>ABR is an auditory evoked potential in response to auditory stimuli and is commonly used to assess hearing thresholds at four frequencies (12.0 kHz, 16.0 kHz, 20.0 kHz, and 24.0 kHz).</li>
	<li>Present the stimulation sound over a small earphone and measure the sound pressure level near the tympanic membrane of the mouse using a small microphone placed near the earphone. Output burst stimuli from a sound generator at 37 cycles/s and amplify the sound pressure from a 20 dB sound pressure level (SPL) to 80 dB SPL in 5 dB SPL steps.</li>
	<li>Insert a stainless steel needle electrode for electroencephalogram recording under the ear canal and frontal region of the ear and insert a ground electrode under the caudal region of the tail.</li>
	<li>Evaluate the cochlear functions by measuring the ABR peak I (P1) amplitude. Automatically analyze the ABR waveforms with respect to the hearing thresholds and ABR P1 amplitude using the ABR peak analysis software as previously reported18.</li>
	<li>Calculate the ABR threshold shifts by subtracting the thresholds obtained before exposure. Compare the ABR threshold shifts in the three exposed groups to those of the non-exposed contralateral ears (control). Measure ABR amplitudes using ABR waveform during 80 dB SPL stimulation.</li>
</ol>
4. Histological assessment
NOTE: Histological assessment was performed as previously described14,15.
<ol>
	<li>HCs and cochlear synapse
	<ol>
		<li>Perform the pathological examination of the cochlea 1 month after LISW exposure.</li>
		<li>Confirm the depth of anesthesia via toe pinch before perfusion. After hemoperfusion with lactated Ringer&#39;s solution, perform transcardiac perfusion with 1 mL/g of 4% paraformaldehyde (PFA). After decapitation, remove the cochlea and perfuse directly with 4% PFA, followed by fixation at 4 &#176;C overnight.</li>
		<li>After fixation, decalcify the cochlea by shaking in 0.5 mol/L ethylenediaminetetraacetic acid (EDTA) solution for 2 days.</li>
		<li>Divide the demineralized cochlea into four pieces. After freezing each piece of cochlea on dry ice for 10 min, perform blocking at room temperature for 1 h in 5% normal horse serum simply conjugated with 0.3% Triton X for permeabilization.</li>
		<li>Use anti-myosin 7a (Myo7A), anti-C-terminal binding protein (CtBP2), and anti-neurofilament (NF) antibodies as primary antibodies and incubated at 37 &#176;C overnight. Use Myo7A, CtBP2, and NF antibodies to evaluate HCs, presynaptic ribbons, and cochlear nerve fibers, respectively.</li>
		<li>Wash off the unbound primary antibody with phosphate-buffered saline (PBS) for 5 x 3 min. Incubate the specimens with the appropriate secondary antibodies at 37 &#176;C for 2 h. After staining, wash the specimens for 3 x 5 min with PBS, and encapsulate the specimens on the slide glass with a water-soluble encapsulant using cover glass.</li>
		<li>For evaluation, acquire the entire image of the cochlea (divided into four pieces) at 10x magnification, and compute the cochlear frequency map using the referenced ImageJ software plug-in to precisely localize the specific cochlear regions at 12.0 kHz, 16.0 kHz, 20.0 kHz, and 24.0 kHz frequency.</li>
		<li>Calculate the rates of HC survival and the number of synapses at each frequency.
		<ol>
			<li>To calculate the rates of HC survival, count the numbers of surviving and missing HCs per 200 &#181;m length at each frequency, and calculate the survival rate of the HCs using equation (1) shown below: 
			HC survival rate (%) = (Number of surviving HCs / Number of surviving and missing HCs) &#215; 100&#160; (1)</li>
			<li>To calculate the number of synapses, obtain high-resolution z-stack images of the inner HC area using an oil immersion objective lens (63&#215;) with a 3.1x digital zoom and a 0.25 &#181;m step size under confocal fluorescence microscopy. Import the image stacks to ImageJ, and automatically count the CtBP2 puncta per IHC within a 50 &#181;m range in each image stack. Calculate the synaptic ribbons survival rate using equation (2): 
			&#8203;Synaptic ribbons survival rate (%) = (Number of synaptic ribbons in LISW exposed ears / Number of synaptic ribbons in control ears) &#215; 100&#160; (2)</li>
		</ol>
		</li>
		<li>For scanning electron microscopy (SEM), remove the cochlea, as previously described, and then fix with 2% PFA and 2.5% glutaraldehyde together at 4 &#176;C overnight. After decalcification of the cochlea by shaking in 0.5 mol/L EDTA solution at 4 &#176;C for 7 days, dissect the cochleae into four pieces for whole-mount preparation.</li>
		<li>Fix the tissues with 1% osmium tetroxide at 4 &#176;C for 30 min, dehydrate in 50% ethanol at room temperature for 10 min, repeat with 70 %, 80 %, 95 %, and then 100 % ethanol, sputter coating with osmium, and examine under an electron microscope at 5.0 kV, as previously reported14.</li>
		<li>Conduct a quantitative analysis of stereociliary bundle disruption in outer hair cells (OHCs) by calculating the ratio of disrupted stereocilia (number of disrupted OHC stereocilia/total number of OHC stereocilia) in each energy group using the SEM images14. Count the number of stereocilia per 100 &#181;m at the center of the 16.0 and 24.0 kHz regions. Designate one or more rows of OHC bundles that are bent toward the lateral side, tangled, or lacking their base as disrupted.</li>
	</ol>
	</li>
	<li>SGNs
	<ol>
		<li>To quantitatively assess the number of SGNs at 1 month after LISW exposure, perform transcardiac perfusion with 4% PBS, decapitate, remove the cochlea, and perform posterior fixation under the same conditions as described above. Decalcify the cochlea in 0.5 M EDTA for 1 week.</li>
		<li>After decalcification, immerse the cochleae in 30% sucrose overnight, embed them in the cryosectioning compound, freeze in liquid nitrogen to prepare sections near Rosenthal&#39;s canal at a thickness of 15 &#181;m, stain with hematoxylin and eosin, and view them under a light microscope14.</li>
		<li>For SGN density measurement, count the number of SGNs in the middle turn of the Rosenthal&#39;s canal and calculate SGN survival per control.</li>
	</ol>
	</li>
</ol>
5. Statistical analysis
<ol>
	<li>Perform statistical analyses using the software of choice.</li>
	<li>Analyze statistical differences in ABR threshold shift, HC, SGN, and synaptic counts using two-way repeated-measures ANOVA [&#34;frequency or cochlear parts (apical/middle/base)&#34; &#215; &#34;animal groups&#34;], two-way analysis of variance (ANOVA), followed by post-hoc Tukey&#39;s multiple comparison test.</li>
	<li>Present all data as mean &#177; standard error and set the statistical significance level at p &#60; 0.05.</li>
</ol>

Characterization of LISW Effects on Cochlear Pathophysiology in Mice

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C., Epstein, M. J., Cleveland, S. S., Wang, H., Maison, S. F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Toward+a+differential+diagnosis+of+hidden+hearing+loss+in+humans.">Toward a differential diagnosis of hidden hearing loss in humans.</a> PLoS One. 11 (9), e0162726 (2016).</li><li>Koizumi, Y., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Y-27632,+a+rock+inhibitor,+improved+laser-induced+shock+wave+(lisw)-induced+cochlear+synaptopathy+in+mice.">Y-27632, a rock inhibitor, improved laser-induced shock wave (lisw)-induced cochlear synaptopathy in mice.</a> Mol Brain. 14 (1), 105 (2021).</li><li>Parthasarathy, A., Kujawa, S. 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The authors declare that they have no conflicts of interest.

This study aimed to validate a mouse model of blast-induced cochlear damage using LISW. Our findings demonstrated that following LISW application through the temporal bone, the exposed mice ear exhibited a consistent pathological and physiological decline in the cochlea, which was accompanied by an increase in LISW overpressure. These results indicate that this mouse model is appropriate for replicating various cochlear pathologies by adjusting the LISW output. Specifically, this LISW-induced cochlear dysfunction mouse m...

Hearing loss and tinnitus are among the most prevalent disabilities, reported in up to 62% of veterans1. Several blast-induced auditory complications, including sensorineural hearing loss (SNHL) and tympanic membrane perforation (TMP), have been reported in individuals exposed to blast overpressure2. Moreover, research on individuals exposed to blasts suggests that blast exposure frequently results in defects in auditory temporal resolution, even when the hearing thresholds are within normal range, which is known as &#34;hidden hearing loss (HHL)&#34;3. It is well established that there is a substantial loss of cochlear synapses between inner hair cells (IHCs) and auditory neurons (ANs) in blast-related cochlear pathology4. Synaptic degeneration results in impaired auditory processing and is a major contributing factor in the development of HHL5. Thus, auditory organs are fragile components containing complex and highly organized structures. However, the precise mechanism by which blast waves affect the inner ear at the cellular level remains unclear. This is because of the challenges in replicating the precise clinical and mechanical intricacies of blast injuries in laboratory settings and the complexity of blast-induced cochlear pathologies.
The primary component of a blast injury is the shock wave (SW), characterized by a rapid and high increase in peak pressure6. The complexity of blast injuries has been extensively investigated in numerous retrospective studies7,8,9. There are various devices for blast generation, such as compressed gas10, shock tubes11, and small-magnitude explosives12, at different levels of pressure. The pressure waveform of the SW generated by recently developed devices closely resembled that of an actual explosion. An important concept in establishing an animal model of blast-induced sensorineural hearing loss is to minimize concomitant injuries, other than auditory damage, to reduce animal death. Thus, blast injury studies have been developed in which shock tubes have been miniaturized and the output can be precisely controlled so that exposed animals rarely die. However, although these animal models usually develop complications, such as TMP, evaluation of cochlear function is difficult because of concurrent conductive hearing loss2. We previously performed an ear-protected animal study on blast injury using earplugs and found no incidence of TMP13. The earplugs could partially attenuate severe cochlear damage but not central auditory neurodegeneration or tinnitus development. Thus, earplugs protect the cochleae as well as the tympanic membrane. However, an animal model of blast-induced pure cochlear damage without TMP is required to study the cochlear pathophysiology caused by blast injuries.
We previously developed a topical blast injury model of the inner ear in rats and mice using a laser-induced shock wave (LISW)14,15. This method can be safely and easily performed at a standard laboratory level and has been used to generate models of lung and head blast injuries16,17. The energy of the LISW can be adjusted by changing the laser type and power, allowing control over the degree of cochlear damage. The LISW-induced cochlear injury model is valuable for studying the mechanisms of SNHL caused by blast injuries and investigating potential treatments. In this study, we describe detailed experimental protocols for creating a mouse model of blast-induced cochlear damage using LISW and demonstrate cochlear degeneration, including the loss of hair cells (HCs), cochlear synapses, and spiral ganglion neurons (SGNs), in an energy-dependent manner in mice following LISW exposure.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>532 nm Q-switched Nd:YAG laser&#160;</td>
			<td>Quantel</td>
			<td>Brilliant b</td>
			<td></td>
		</tr>
		<tr>
			<td>ABR peak analysis software</td>
			<td>Mass Eye and Ear</td>
			<td>N/A</td>
			<td>EPL Cochlear Function Test Suite</td>
		</tr>
		<tr>
			<td>Acrylic resin welding adhesive&#160;</td>
			<td>Acrysunday Co., Ltd</td>
			<td>N/A</td>
			<td></td>
		</tr>
		<tr>
			<td>confocal fluorescence microscopy</td>
			<td>Leica</td>
			<td>TCS SP8</td>
			<td></td>
		</tr>
		<tr>
			<td>cryosectioning compound</td>
			<td>Sakura</td>
			<td>Tissue-Tek O.C.T</td>
			<td></td>
		</tr>
		<tr>
			<td>CtBP2 antibody</td>
			<td>BD Transduction</td>
			<td>#612044</td>
			<td></td>
		</tr>
		<tr>
			<td>Dielectric multilayer mirrors</td>
			<td>SIGMAKOKI CO.,LTD</td>
			<td>TFMHP-50C08-532</td>
			<td>M1-M3</td>
		</tr>
		<tr>
			<td>Digital oscilloscope</td>
			<td>Tektronix</td>
			<td>DPO4104B</td>
			<td></td>
		</tr>
		<tr>
			<td>Earphone</td>
			<td>CUI</td>
			<td>CDMG15008-03A</td>
			<td></td>
		</tr>
		<tr>
			<td>Hydrophone</td>
			<td>RP acoustics e.K.</td>
			<td>FOPH2000</td>
			<td></td>
		</tr>
		<tr>
			<td>Image J software plug-in</td>
			<td>NIH</td>
			<td>measurement line</td>
			<td>https://myfiles.meei.harvard.edu/xythoswfs/webui/_xy-e693768_1-t_wC4oKeBD</td>
		</tr>
		<tr>
			<td>Light microscope</td>
			<td>Keyence Corporation</td>
			<td>BZ-X700</td>
			<td></td>
		</tr>
		<tr>
			<td>Myosin 7A antibody</td>
			<td>Proteus Biosciences</td>
			<td>#25&#8211;6790&#160;</td>
			<td></td>
		</tr>
		<tr>
			<td>Neurofilament antibody</td>
			<td>Sigma</td>
			<td>#AB5539</td>
			<td></td>
		</tr>
		<tr>
			<td>Plano-convex lens</td>
			<td>SIGMAKOKI CO.,LTD</td>
			<td>SLSQ-30-200PM</td>
			<td></td>
		</tr>
		<tr>
			<td>Prism software</td>
			<td>GraphPad</td>
			<td>N/A</td>
			<td>ver.8.2.1</td>
		</tr>
		<tr>
			<td>Scanning electron microscope</td>
			<td>JEOL Ltd</td>
			<td>JSM-6340F</td>
			<td></td>
		</tr>
		<tr>
			<td>Small digital endoscope</td>
			<td>AVS Co. Ltd</td>
			<td>AE-C1</td>
			<td></td>
		</tr>
		<tr>
			<td>Ultrasonic jelly</td>
			<td>Hitachi Aloka Medical</td>
			<td>N/A</td>
			<td></td>
		</tr>
		<tr>
			<td>Variable attenuator</td>
			<td>Showa Optronics Co.</td>
			<td>N/A</td>
			<td>Currenly avaiable successor: KYOCERA SOC Corporation, RWH-532HP II</td>
		</tr>
		<tr>
			<td>Water-soluble encapsulant&#160;</td>
			<td>Dako</td>
			<td>#S1964</td>
			<td></td>
		</tr>
	</tbody>
</table>

research application of laser-induced shock wave for studying blast-induced cochlear injury

The ear is the organ most susceptible to explosion overpressure, and cochlear injuries frequently occur after blast exposure. Blast exposure can lead to sensorineural hearing loss (SNHL), which is an irreversible hearing loss that negatively affects the quality of life. Detailed blast-induced cochlear pathologies, such as the loss of hair cells, spiral ganglion neurons, cochlear synapses, and disruption of stereocilia, have been previously documented. However, determining cochlear sensorineural deterioration after a blast injury is challenging because animals exposed to blast overpressure usually experience tympanic membrane perforation (TMP), which causes concurrent conductive hearing loss. To evaluate pure sensorineural cochlear dysfunction, we developed an experimental animal model of blast-induced cochlear injury using a laser-induced shock wave. This method avoids TMP and concomitant systemic injuries and reproduces the functional decline in the SNHL component in an energy-dependent manner after LISW exposure. This animal model could be a platform for elucidating the pathological mechanisms and exploring potential treatments for blast-induced cochlear dysfunction.

Author Spotlight: Development of a Laser-Induced Shock Wave Animal Model Without Tympanic Membrane Perforation

Research Application of Laser-Induced Shock Wave for Studying Blast-Induced Cochlear Injury

To evaluate pure sensory neuro cochlear dysfunction caused by blast exposure result tympanic membrane perforation. We develop an experimental animal model of blast induced cochlear injury using a razor induce shock wave. This method can avoid can tympanic membrane perforation and concomitant systemic injuries and reproduce the functional decline in the sensory neuro hearing loss component in an energy dependent manner after after LISW exposure.This animal model could be an ideal platform for stating the pathological mechanism and exploring potential treatment for the blast induced cochlear dysfunction.

LISW waveform 
The reproducibility of the LISW pressure waveform was measured 5x at 2.0 J/cm2 as follows. The waveforms were generally similar and stable and showed a sharp increase with time width, peak pressure, and impulse of 0.43&#177;0.4 &#181;s, 92.1 &#177; 6.8 MPa, and 14.1 &#177; 1.9 Pa&#8729;s (median &#177; SD), which corresponds to SW characteristics (Figure 1B). LISWs are characterized by a fast rise time, high peak pressure, short duration, and p...

Read this Scientific Article Protocol about Author Spotlight: Development of a Laser-Induced Shock Wave Animal Model Without Tympanic Membrane Perforation at JoVE.com

Here, we describe experimental protocols for creating an animal model of blast-induced cochlear injury using laser-induced shock wave (LISW). Exposure of the temporal bone to LISW allows the reproduction of blast-induced cochlear pathophysiology. This animal model could be a platform for elucidating cochlear pathology and exploring potential treatments for blast injuries.

The ear is the organ most susceptible to explosion overpressure, and cochlear injuries frequently occur after blast exposure. Blast exposure can lead to ...

research-application-laser-induced-shock-wave-for-studying-blast

Research

JoVE Journal

Medicine

471 vues.  National Defense Medical College. Évaluer le dysfonctionnement neurocochléaire sensoriel pur causé par l’exposition à l’explosion résultant de la perforation de la membrane tympanique. Nous développons un modèle animal expérimental de lésions cochléaires induites par l’explosion à l’aide d’une onde de choc induite par un rasoir. Cette méthode permet d’éviter la perforation de la membrane tympanique et les lésions systémiques concomitantes et de reproduire le déclin...

Vidéo: Pleins feux sur l’auteur : Développement d’un modèle animal d’ondes de choc induites par laser sans perforation de la membrane tympanique

Evaluating the Impact of Laser-Induced Shock Wave Exposure on Cochlear Function and Auditory Thresholds in Mice

To begin, prepare a laser-induced shockwave, or LISW exposure assembly. Irradiate the exposure assembly with a 532 nanometer Q-switched NDYAG laser to generate the LISW behind the target. Focus the laser pulse through a plano-convex lens to a three millimeter diameter spot on the laser target.Use the LISW irradiation to generate plasma at the bonding surface of the two materials, and vaporize the rubber, leaving vaporized rubber in the cavity. Carefully shave the postauricular regions of the anesthetized mouse to avoid retaining the trapped air in the fur. Fix the mouse on a plate with their postauricular regions positioned upwards towards the LISW focal area.To ensure acoustic impedance matching, use an ultrasound conductive gel between the laser target and the skin surface. Attach a black rubber target percutaneously to the postauricular region of the mouse ear. Apply a single LISW pulse to the cochlea via the temporal bone at three energy densities.Insert a ground electrode under the caudal region of the tail. Then, insert a stainless steel needle electrode for electroencephalogram recording under the ear canal and frontal region of the ear. Present the stimulation sound over a small earphone and measure the sound pressure level near the tympanic membrane of the mouse.Generate output burst stimuli from a sound generator at 37 cycles, and amplify the sound pressure from a 20 decibel sound pressure level to 80 decibel SPL in five decibel SPL steps. Measure the auditory brainstem response, or ABR peak one amplitude, to evaluate the cochlear functions. Then, ABR peak analysis software automatically analyzes the ABR waveforms with respect to the hearing thresholds and ABR P1 amplitude.An auditory function assessment using ABR post LISW exposure revealed significant ABR threshold shifts at one day, persisting for one month at higher energy levels. ABR thresholds in the two joules per square centimeters group showed recovery to pre-exposure levels, indicating energy dependent effects. After one day and one month of LISW exposure, significant decreases in ABR P1 amplitudes were observed in all LISW irradiated groups across frequencies.

Impact of Laser-Induced Shock Wave Exposure on Cochlear Histology and Cellular Survival in Mice

One month after LISW exposure, conduct a pathological examination of the mouse's cochlea. Start hemoperfusion with lactated Ringer's solution, followed by transcardiac perfusion with 4%paraformaldehyde. After decapitation of the mouse, remove the cochlea and perfuse directly with 4%paraformaldehyde at four degrees Celsius overnight.After acquiring the entire image of the cochlea, using ImageJ software, compute the cochlear frequency map to precisely localize the specific cochlear regions at different frequencies. Using the given formula, calculate the survival rate of the hair cells at each frequency. After acquiring high-resolution Z-stack images of the inner hair cells, calculate the number of synapses.Visualize the hematoxylin and eosin-stained cochlea sections and count the number of spiral ganglion neurons in the middle turn of the Rosenthal's canal to calculate the spiral ganglion neuron survival per control. Compared to the control, fluorescent immunostaining showed intact cochlear structures with no significant loss of hair cells or nerve fibers across all groups. However, quantitative assessments indicated that the survival of outer hair cells was significantly lower in the 2.5 joules per square centimeter group.Meanwhile, inner hair cell survival was consistent among groups. Synaptic ribbon counts were notably reduced in higher-frequency groups. This was paralleled by a decrease in spiral ganglion neuron density in the 2.5 joules per square centimeter group, indicating that cochlear degeneration is dependent on LISW energy exposure.