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.

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

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

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.

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 ...

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

Research

JoVE Journal

Medicine

476 Views.  National Defense Medical College. 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.

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

Controlled Cortical Impact Model for Traumatic Brain Injury

Every year over a million Americans suffer a traumatic brain injury (TBI). Combined with the incidence of TBIs worldwide, the physical, emotional, social, and economical effects are staggering. Therefore, further research into the effects of TBI and effective treatments is necessary. The controlled cortical impact (CCI) model induces traumatic brain injuries ranging from mild to severe. This method uses a rigid impactor to deliver mechanical energy to an intact dura exposed following a craniectomy. Impact is made under precise parameters at a set velocity to achieve a pre-determined deformation depth. Although other TBI models, such as weight drop and fluid percussion, exist, CCI is more accurate, easier to control, and most importantly, produces traumatic brain injuries similar to those seen in humans. However, no TBI model is currently able to reproduce pathological changes identical to those seen in human patients. The CCI model allows investigation into the short-term and long-term effects of TBI, such as neuronal death, memory deficits, and cerebral edema, as well as potential therapeutic treatments for TBI.

Traumatic brain injuries (TBIs) remain a serious health problem. Using the controlled cortical impact surgery model, research on the effects of TBI and possible treatment methods may be performed.

Every year over a million Americans suffer a traumatic brain injury (TBI).  Combined with the incidence of TBIs worldwide, the physical, emotional, ...

Renal Ischaemia Reperfusion Injury: A Mouse Model of Injury and Regeneration

Renal ischaemia reperfusion injury (IRI) is a common cause of acute kidney injury (AKI) in patients and occlusion of renal blood flow is unavoidable during renal transplantation. Experimental models that accurately and reproducibly recapitulate renal IRI are crucial in dissecting the pathophysiology of AKI and the development of novel therapeutic agents. Presented here is a mouse model of renal IRI that results in reproducible AKI. This is achieved by a midline laparotomy approach for the surgery with one incision allowing both a right nephrectomy that provides control tissue and clamping of the left renal pedicle to induce ischaemia of the left kidney. By careful monitoring of the clamp position and body temperature during the period of ischaemia this model achieves reproducible functional and structural injury. Mice sacrificed 24 hr&#160;following surgery demonstrate loss of renal function with elevation of the serum or plasma creatinine level as well as structural kidney damage with acute tubular necrosis evident. Renal function improves and the acute tissue injury resolves during the course of 7 days following renal IRI such that this model may be used to study renal regeneration. This model of renal IRI has been utilized to study the molecular and cellular pathophysiology of AKI as well as analysis of the subsequent renal regeneration.

The mouse model of renal ischaemia reperfusion injury described here comprises of a right nephrectomy that provides control tissue and clamping of the left renal pedicle to induce ischaemia that results in acute kidney injury. This model uses a midline laparotomy approach with all steps performed via one incision.

Renal ischaemia reperfusion injury (IRI) is a common cause of acute kidney injury (AKI) in patients and occlusion of renal blood flow is unavoidable ...

A Mouse Model for Laser-induced Choroidal Neovascularization

The mouse laser-induced choroidal neovascularization (CNV) model has been a crucial mainstay model for neovascular age-related macular degeneration (AMD) research. By administering targeted laser injury to the RPE and Bruch&#8217;s membrane, the procedure induces angiogenesis, modeling the hallmark pathology observed in neovascular AMD. 
First developed in non-human primates, the laser-induced CNV model has come to be implemented into many other species, the most recent of which being the mouse. Mouse experiments are advantageously more cost-effective, experiments can be executed on a much faster timeline, and they allow the use of various transgenic models. The miniature size of the mouse eye, however, poses a particular challenge when performing the procedure. Manipulation of the eye to visualize the retina requires practice of fine dexterity skills as well as simultaneous hand-eye-foot coordination to operate the laser. However, once mastered, the model can be applied to study many aspects of neovascular AMD such as molecular mechanisms, the effect of genetic manipulations, and drug treatment effects. 
The laser-induced CNV model, though useful, is not a perfect model of the disease. The wild-type mouse eye is otherwise healthy, and the chorio-retinal environment does not mimic the pathologic changes in human AMD. Furthermore, injury-induced angiogenesis does not reflect the same pathways as angiogenesis occurring in an age-related and chronic disease state as in AMD.
Despite its shortcomings, the laser-induced CNV model is one of the best methods currently available to study the debilitating pathology of neovascular AMD. Its implementation has led to a deeper understanding of the pathogenesis of AMD, as well as contributing to the development of many of the AMD therapies currently available.

Here, we present the mouse laser-induced choroidal neovascularization (CNV) protocol, an experimental model that re-creates the vascular hallmarks of neovascular age-related macular degeneration (AMD). Once mastered, it can reliably and effectively induce CNV as a model system to test various experimental measures.

The mouse laser-induced choroidal neovascularization (CNV) model has been a crucial mainstay model for neovascular age-related macular degeneration (AMD) ...

Neonatal Murine Cochlear Explant Technique as an In Vitro Screening Tool in Hearing Research

While there have been remarkable advances in hearing research over the past few decades, there is still no cure for Sensorineural Hearing Loss (SNHL), a condition that typically involves damage to or loss of the delicate mechanosensory structures of the inner ear. Sophisticated in vitro and ex vivo assays have emerged in recent years, enabling the screening of an increasing number of potentially therapeutic compounds while minimizing resources and accelerating efforts to develop cures for SNHL. Though homogenous cultures of certain cell types continue to play an important role in current research, many scientists now rely on more complex organotypic cultures of murine inner ears, also known as cochlear explants. The preservation of organized cellular structures within the inner ear facilitates the in situ evaluation of various components of the cochlear infrastructure, including inner and outer hair cells, spiral ganglion neurons, neurites, and supporting cells. Here we present the preparation, culture, treatment, and immunostaining of neonatal murine cochlear explants. The careful preparation of these explants facilitates the identification of mechanisms that contribute to SNHL and constitutes a valuable tool for the hearing research community.

Neonatal Murine Cochlear Explant Technique as an In Vitro Screening Tool in Hearing Research

The goal of this protocol is to demonstrate the preparation, culture, treatment, and immunostaining of neonatal murine cochlear explants. The technique can be utilized as an in vitro screening tool in hearing research.

While there have been remarkable advances in hearing research over the past few decades, there is still no cure for Sensorineural Hearing Loss (SNHL), a ...

Modified Heterotopic Hindlimb Osteomyocutaneous Flap Model in the Rat for Translational Vascularized Composite Allotransplantation Research

Vascularized composite allotransplantation (VCA) is a relatively new field in the reconstructive surgery. Clinical achievements in human VCA include hand and face transplants and, more recently, abdominal wall, uterus, and urogenital transplants. Functional outcomes have exceeded initial expectations, and most recipients enjoy an improved quality of life. However, as clinical experience accumulates, chronic rejection and complications from the immunosuppression must be addressed. In many cases where grafts have failed, the causative pathology has been ischemic vasculopathy. <span style="font-size: 11pt; line-height: 115%; font-family: Arial, sans-serif; color: rgb(41, 43, 49); background-image: initial; background-position: initial; background-size: initial; background-repeat: initial; background-attachment: initial; background-origin: initial; background-clip: initial;">The biological mechanisms of the acute and chronic rejection associated with VCA, especially ischemic vasculopathy, are important areas of research. However, due to the very small number of VCA patients, the evaluation of proposed mechanisms is better addressed in an experimental model. Multiple groups have used animal models to address some of the relevant unsolved questions in VCA rejection and vasculopathy. Several model designs involving a variety of species are described in the literature. Here we present a reproducible model of VCA heterotopic hindlimb osteomyocutaneous flap in the rat that can be utilized for translational VCA research. This model allows for the serial evaluation of the graft, including biopsies and different imaging modalities, while maintaining a low level of morbidity.

Vascularized composite allograft offers life-altering benefits to transplant recipients, but the biological causes of graft rejection and vasculopathy remain poorly understood. The rodent surgical model presented here offers a reproducible, clinically relevant model of transplantation, allowing researchers to evaluate rejection events and potential therapeutic strategies to prevent their occurrence.

Vascularized composite allotransplantation (VCA) is a relatively new field in the reconstructive surgery. Clinical achievements in human VCA include hand ...

Intrathecal Application of a Fluorescent Dye for the Identification of Cerebrospinal Fluid Leaks in Cochlear Malformation

In cases of cerebrospinal fluid (CSF) leaks, reliable detection of their origins is needed to seal the leak sufficiently and prevent complications, such as meningitis. A method is presented here using intrathecal administered fluorescein in a clinical case of bilateral congenital ear malformation. A fluorescent dye is administered intrathecally to achieve intraoperative visualization of CSF leaks. The dye is applied 20 min before surgery, and concentration of 5% is used. Per every 10 kg of body weight, 0.1 mL of the fluid is applied intrathecally. The fluorescein is visualized using a fully digital microscope. The origin of the fluid leak is identified in the stapes footplate. During primary surgery, it is sealed, and cochlea implantation is performed for hearing restoration. In this specific case, 6 weeks later, the implant was explanted due to acute meningitis, and the electrode array was left as a spacer. Postoperatively, in the aural smear, &#946;-transferrin was detected. During a revision mastoidectomy, dislocated coverage of the leak was found. The stapes was removed and oval window sealed. Five days after revision surgery, no &#946;-transferrin was detected in the aural smear. During the revision of cochlea implantation 6 months later, intact coverage of the oval niche was observed. Thus, intrathecal fluorescein application proves to be a reliable tool for the detection of CSF leaks. It facilitates the orientation in malformations and complicated or unknown surgical situs. In the literature, its use is described for CSF fistulas in endonasal surgery but is rarely described in skull base and mastoid surgeries. The method has been used successfully in several cases with CSF leaks, and the results confirm the feasibility of safely accessing the origin of the leak.

Intrathecally applied fluorescein is used to achieve intraoperative visualization of CSF leaks. This protocol describes a lumbar puncture, the application of 5% fluorescein, and intraoperative visualization using a fully digital microscope.

In cases of cerebrospinal fluid (CSF) leaks, reliable detection of their origins is needed to seal the leak sufficiently and prevent complications, such ...

Performing Intracochlear Electrocochleography During Cochlear Implantation

Electrocochleography (ECochG) measures inner ear potentials generated in response to acoustic stimulation of the ear. These potentials reflect the residual function of the cochlea. In cochlear implant candidates with residual hearing, the implant electrode can directly measure ECochG responses during the implantation process. Various authors have described the ability to monitor the inner ear function by continuous ECochG measurements during the surgery. The measurement of ECochG signals during surgery is not trivial. There are no interpretable signals in up to 20% of cases. For a successful recording, a standardized procedure is recommended to achieve the highest measurement reliability and avoid possible pitfalls. Therefore, seamless collaboration between the CI surgeon and CI technician is key. This video consists of an overview of the system setup and a stepwise procedure of performing intracochlear ECochG measurements during CI surgery. It shows the surgeon's and the CI technician's roles in the process, and how a smooth collaboration between the two is made possible.

Electrocochleography (ECochG) measures inner ear potentials generated in response to acoustic stimulation. In cochlear implant (CI) candidates, such inner ear potentials can be measured directly with the implant electrodes. In this video, we systematically explain how to perform ECochG recordings during CI surgery.

Electrocochleography (ECochG) measures inner ear potentials generated in response to acoustic stimulation of the ear. These potentials reflect the ...

Controlled Semi-Automated Laser-Induced Injuries for Studying Spinal Cord Regeneration in Zebrafish Larvae

Zebrafish larvae possess a fully functional central nervous system (CNS) with a high regenerative capacity only a few days after fertilization. This makes this animal model very useful for studying spinal cord injury and regeneration. The standard protocol for inducing such lesions is to transect the dorsal part of the trunk manually. However, this technique requires extensive training and damages additional tissues. A protocol was developed for laser-induced lesions to circumvent these limitations, allowing for high reproducibility and completeness of spinal cord transection over many animals and between different sessions, even for an untrained operator. Furthermore, tissue damage is mainly limited to the spinal cord itself, reducing confounding effects from injuring different tissues, e.g., skin, muscle, and CNS. Moreover, hemi-lesions of the spinal cord are possible. Improved preservation of tissue integrity after laser injury facilitates further dissections needed for additional analyses, such as electrophysiology. Hence, this method offers precise control of the injury extent that is unachievable manually. This allows for new experimental paradigms in this powerful model in the future.

The present protocol describes a method to induce tissue-specific and highly reproducible injuries in zebrafish larvae using a laser lesion system combined with an automated microfluidic platform for larvae handling.

Zebrafish larvae possess a fully functional central nervous system (CNS) with a high regenerative capacity only a few days after fertilization. This makes ...

Robotic Cochlear Implantation for Direct Cochlear Access

Robot-assisted systems offer great potential for gentler and more precise cochlear implantation. In this article, we provide a comprehensive overview of the clinical workflow for robotic cochlear implantation using a robotic system specifically developed for a minimally invasive, direct cochlear access. The clinical workflow involves experts from various disciplines and requires training to ensure a smooth and safe procedure. The protocol briefly summarizes the history of robotic cochlear implantation. The clinical sequence is explained in detail, beginning with the assessment of patient eligibility and covering surgical preparation, preoperative planning with the special planning software, drilling of the middle ear access, intraoperative imaging to confirm the trajectory, milling of the inner ear access, insertion of the electrode array, and implant management. The steps that require special attention are discussed. As an example, the postoperative outcome of robotic cochlear implantation in a patient with advanced otosclerosis is presented. Finally, the procedure is discussed in the context of the authors' experience.

Robotic cochlear implantation is a procedure for minimally invasive inner ear access. Compared to conventional surgery, robotic cochlear implantation involves additional steps that need to be carried out in the operating room. In this article we give a description of the procedure and highlight the important aspects of robotic cochlear implantation.

Robot-assisted systems offer great potential for gentler and more precise cochlear implantation. In this article, we provide a comprehensive overview of ...

The Miniature Pig: A Large Animal Model for Cochlear Implant Research

Cochlear implants (CI) are the most effective method to treat people with severe-to-profound sensorineural hearing loss. Although CIs are used worldwide, no standard model exists for investigating the electrophysiology and histopathology in patients or animal models with a CI or for evaluating new models of electrode arrays. A large animal model with cochlea characteristics similar to those of humans may provide a research and evaluation platform for advanced and modified arrays before their use in humans. 
To this end, we established standard CI methods with Bama mini-pigs, whose inner ear anatomy is highly similar to that of humans. Arrays designed for human use were implanted into the mini pig cochlea through a round window membrane, and a surgical approach followed that was similar to that used for human CI recipients. Array insertion was followed by evoked compound action potential (ECAP) measurements to evaluate the function of the auditory nerve. This study describes the preparation of the animal, surgical steps, array insertion, and intraoperative electrophysiological measurements. 
The results indicated that the same CI used for humans could be easily implanted in mini-pigs via a standardized surgical approach and yielded similar electrophysiological outcomes as measured in human CI recipients. Mini-pigs could be a valuable animal model to provide initial evidence of the safety and potential performance of novel electrode arrays and surgical approaches before applying them to human beings.

Miniature pigs (mini-pigs) are an ideal large animal model for research into cochlear implants. Cochlear implantation surgery in mini-pigs can be utilized to provide initial evidence of the safety and potential performance of novel electrode arrays and surgical approaches in a living system similar to human beings.

Cochlear implants (CI) are the most effective method to treat people with severe-to-profound sensorineural hearing loss. Although CIs are used worldwide, ...