We thank Dr. A. Jacobo, Dr. J. Salvi, and A. Petelski for their contributions to the original research on which this protocol is based. We also thank J. Llamas and W. Makmura for technical assistance and animal husbandry. We acknowledge NIDCD Training grant T32 DC009975, NIDCD grant R01DC015530, Robertson Therapeutic Development Fund, and the Caruso Family Foundation for funding. Finally, we acknowledge support from Howard Hughes Medical Institute, of which Dr. Hudspeth is an Investigator.

All methods described here have been approved by the Animal Care and Use Committees of Rockefeller University and of the University of Southern California.
1. (Optional) Preparation of Collagen I Solution from Mouse-tail Tendons
Note: Collagen I solutions are available commercially. Follow the manufacturer&#39;s instructions for gel preparation.
<ol>
	<li>Euthanize 5 - 10 young adult (3 - 5 weeks old) mice of any wild type strain with carbon dioxide in accordance with the protocol approved by the relevant Institutional Animal Care and Use Committee16. Collect the tails and disinfect them by submerging in 70% ethanol for a minimum of 4 h at room temperature. 
	Note: Incubation for over 48 h should be avoided, as it results in excessive tissue dehydration and impedes the tendon extraction process.</li>
	<li>Remove the skin from each tail by introducing a longitudinal cut with a scalpel blade and retracting the whole skin with forceps. Transfer the skinned tails to a 100-mm Petri dish filled with clean 70% ethanol and cut them into 10-mm segments. 
	Note: Discard the thinnest parts of the tails, which are difficult to manipulate.</li>
	<li>Using forceps, secure each segment of the tail to the bottom of the Petri dish and use a second pair of fine forceps to extract the tendons from the tail one at a time. Individual tendon fibers should emerge with minimal resistance. 
	Note: Old, dull #5 forceps work well for this step.</li>
	<li>Prepare 100 mL of a 0.1% solution (by volume) of acetic acid in sterile, molecular-grade water and add 10 mL of that solution to a sterile 100-mm Petri dish. Transfer the tendons to the Petri dish and leave for 1 h at room temperature to denature the collagen I.</li>
	<li>Use a sterile scalpel or iridectomy scissors with blunt, curved tips to mince the tendon into 1 - 2 mm fragments. Transfer the minced tendons to a sterile 50-mL tube and add 0.1% acetic acid to bring the volume to 50 mL. 
	Note: Use four or five tails for each 50 mL of acid to achieve a collagen I concentration of 2.0 - 2.5 mg/mL.</li>
	<li>Refrigerate the collagen I solution at 4 &#176;C for a minimum of 48 h to facilitate complete protein denaturation. Vortex with a tabletop vortex set to the maximal speed for 1 min twice a day.</li>
	<li>Measure the protein concentration of the solution using the bicinchoninic acid assay, adjust the collagen I concentration to 2.0 - 2.5 mg/mL by adding 0.1% solution of acetic acid, and store at 4 &#176;C. 
	Note: Collagen I solution can be stored for one to two years.</li>
	<li>(Optional) Centrifuge the collagen I solution at 2,000 x g for 1 h at 4 &#176;C and use the translucent top fraction (approximately half of the volume), to achieve optically clear collagen gels in Section 4.</li>
</ol>
2. Dissection of Vestibular and Auditory Organs
<ol>
	<li>Euthanize pregnant mice of any strain with carbon dioxide in accordance with the protocol approved by the relevant Institutional Animal Care and Use Committee16. Extract and decapitate the embryos. 
	Note: Lfng-CreERT2/tdTomato mice can be used to allow permanent labeling of supporting cells upon exposure to 4-hydroxytamoxifen17.</li>
	<li>Sterilize all the working surfaces and dissection instruments, including two pairs of #5 forceps and a hair knife18, by cleaning them with 70% ethanol.</li>
	<li>Split each head in two halves by introducing a longitudinal cut with a sterile scalpel blade or by using two pairs of #5 forceps. Extract the temporal bones, which contain the inner ears19, and place them into 15 mL of ice-cold Hank&#39;s balanced salt solution (HBSS) in a 60-mm Petri dish. Keep the dish on ice. 
	Note: Embryos at a range of stages from E13.5 to E18.5 have been used successfully.</li>
	<li>(Optional) Wash the inner ears by gently shaking them in a 50 mL conical tube containing 30 mL ice-cold HBSS and replacing the HBSS three or four times. Keep the tube on ice. 
	Note: This step limits possible contamination of the tissue culture and quickly brings the temperature to 4 &#176;C, which prevents tissue degradation and cell death.</li>
	<li>Using two pairs of fine #5 forceps, separate the inner ears from the temporal bone and transfer the ears, three or four at a time, into a 60-mm Petri dish filled with ice-cold HBSS.</li>
	<li>Dissect the vestibular sensory organs.
	<ol>
		<li>Orient the ears medial side up and locate the utricle. With two pairs of #5 forceps, remove the cartilage surrounding the vestibular organs. Gently sever the vestibular nerve, the connection between the utricle and saccule, and the semicircular canals, as shown in Figure 1. Gently pull the utricle and the attached ampullae of the superior and horizontal semicircular canals from the ear. 
		Note: This method can be used on inner ears of stages E16.5 - E18.5. Preserve the cartilage fragments for Section 3 below.</li>
	</ol>
	</li>
	<li>Dissect the cochlea.
	<ol>
		<li>Remove the cartilaginous tissue surrounding the hearing organ with two pairs of #5 forceps. Gently sever the connection between the cochlear base and the saccule as shown in Figure 1. 
		Note: For stages E13.5 - E14.5, soften the cartilage prior to dissection by treating the inner ears with 0.25% collagenase I in phosphate-buffered saline (PBS) solution for 5 min at room temperature. Preserve the cartilage fragments for Section 3 below.</li>
	</ol>
	</li>
	<li>Using a 200-&#181;L pipet fitted with a wide nonstick tip, transfer the utricles and cochleae to a 30-mm Petri dish filled with growth medium comprising DMEM/F12 supplemented with 33 mM Dglucose, 19 mM sodium bicarbonate, 15 mM 4(2hydroxyethyl)-1piperazineethanesulfonic acid (HEPES), 1 mM glutamine, 1 mM nicotinamide, 20 mg/L epidermal growth factor, 20 mg/L fibroblast growth factor, 10 mg/L insulin, 5.5 mg/L transferrin, and 5 &#181;g/L sodium selenite.</li>
	<li>Maintain the utricle preparations for up to 3 h at 37 &#176;C in a tissue-culture incubator gassed with 5% carbon dioxide to allow healing of the cuts in the epithelium introduced during the dissection. Cochlea preparations should be transferred to the collagen gel 10 min after the dissection. 
	Note: Because the dissection is performed in ice-cold HBSS, there is no need to pre-warm the growth medium to 37 &#176;C.</li>
</ol>
3. (Optional) Adjust Collagen I Gel Stiffness by Adding Varying Concentrations of Chondrocytes
Note: The method for chondrocyte isolation was modified from Gosset et al.20
<ol>
	<li>Prepare a 1% solution of collagenase I in sterile 1x PBS and store 100 - 200 &#181;L aliquots at -80 &#176;C. Defrost on ice when needed, avoiding multiple freeze-thaw cycles.</li>
	<li>Use fine forceps to collect the pieces of cartilage left over from the dissection of the vestibular and auditory organs from 10 - 12 ears. Separate connective and inner ear tissues from the cartilage. Transfer the cartilage to a 30-mm Petri dish and add 300 &#181;L of sterile growth medium.</li>
	<li>Use a sterile scalpel blade or iridectomy scissors with blunt curved tips to mince the tissue to achieve cartilage pieces approximately 0.5 mm in length.</li>
	<li>Add collagenase I to the medium with cartilage to achieve a final enzyme concentration of 0.25%. Transfer the dish to a tissue-culture incubator at 37 &#176;C gassed with 5% carbon dioxide. Pipet vigorously using a 1,000 &#181;L pipet every 20 min until pieces of cartilage dissociate and are no longer distinguishable in the solution, then incubate for an additional 20 min. 
	Note: In case of the utricle preparations, chondrocyte isolation can be performed during step 2.9; it takes approximately 2 h (3 - 4 pipetting rounds).</li>
	<li>Collect the cell suspension in a 15 mL conical tube and adjust the volume to 10 mL with sterile 1x PBS. Centrifuge at 800 x g for 5 min at 4 &#176;C. Remove the supernatant and resuspend the cells in 10 mL of sterile 1x PBS. Centrifuge at 800 x g for 5 min at 4 &#176;C and remove the supernatant. 
	Note: It is critical to wash the cells twice; any remaining collagenase I will digest the collagen I gel.</li>
	<li>Using a 200-&#181;L pipet, add 100 &#181;L of culture medium to the cell pellet and pipet gently to resuspend. Count the cells using a hemocytometer and maintain them on ice prior to use.</li>
	<li>To increase gel stiffness, add chondrocytes to the neutralized collagen I gel solution (Section 1) and mix rapidly to distribute the cells throughout the gel prior to solidification. 
	Note: The elastic modulus of the collagen I gel (a measure of stiffness), increases linearly with the number of chondrocytes added11. The relationship is described by the experimentally determined linear function E = 206&#183;Nc + 15, in which E is the elastic modulus in Pascals and Nc is the number of chondrocytes in millions. For a detailed protocol for gel stiffness measurements please refer to the original article11.</li>
</ol>
4. Place the Vestibular or Auditory Sensory Organ in a Collagen I Gel
<ol>
	<li>In a sterile 1.5-mL tube, prepare collagen I polymerization solution by mixing 160 &#181;L of 10x PBS with phenol red pH indicator, 133 &#181;L of 0.34 M sodium hydroxide, 70 &#181;L of 0.9 M sodium bicarbonate, and 40 &#181;L of 1 M HEPES. Keep the tube on ice. 
	Note: This recipe provides the polymerization solution needed to prepare 2 mL of collagen I gel, but can be scaled as needed.</li>
	<li>Mix 100 &#181;L of polymerization solution and 400 &#181;L of collagen I solution on ice by gently pipetting up and down in a chilled 1.5-mL tube. To increase gel stiffness, add 50 &#181;L of growth medium containing chondrocytes and mix gently as described in Section 3.</li>
	<li>Transfer 500 &#181;L of the neutralized collagen I solution to a chilled 30-mm Petri dish with a 10-mm glass-bottom insert or to a well of a four-well plate. 
	Note: It is critical to keep all the reagents on ice to prevent rapid and uneven polymerization of the collagen I.</li>
	<li>Quickly transfer the cochleae or utricles to the neutralized collagen I solution and adjust the organs to their desired positions with a sterile hair knife18 or pair of fine forceps. 
	Note: Collagen I polymerization becomes noticeable after 1 - 2 min as the solution becomes turbid.</li>
	<li>After the solution around the tissue has polymerized, place the Petri dish or four-well plate for 20 min at 37 &#176;C in a tissue-culture incubator gassed with 5% carbon dioxide to ensure complete polymerization.</li>
	<li>Add 3 mL of growth medium supplemented with 0.5% fetal bovine serum (FBS) per Petri dish or 500 &#181;L of the same medium per well of a four-well plate. Maintain the culture at 37 &#176;C in a tissue-culture incubator gassed with 5% carbon dioxide. If desired, supplement the growth medium with 10 &#181;M 5-ethynyl-2&#180;-deoxyuridine (EdU) to label proliferating cells. 
	Note: Higher FBS concentrations can be used if desired.</li>
</ol>
5. Viral Injections in Three-dimensional Cultures of Vestibular and Auditory Sensory Organs
<ol>
	<li>Defrost the desired virus on ice and mix with trypan blue solution in a 0.5 mL conical tube to achieve a final dye concentration of 0.05%. Use 10 - 20x dye to avoid substantial dilution of the virus. Keep on ice. 
	Note: Adenovirus serotype 5 works best for infecting supporting cells in the utricle19, whereas adeno-associated virus Anc80 can infect both hair cells and supporting cells in the utricle and the cochlea21.</li>
	<li>Break the tip of a glass needle prepared on a micropipette puller with clean fine forceps while observing it at the highest magnification of a binocular dissecting microscope. 
	Note: It is important to optimize the settings on the needle puller. Needles with 9 - 12-mm shanks and 20 - 30-&#181;m openings work best.</li>
	<li>Remove a sensory-organ culture from the incubator. Attach the needle to the microinjector and fill it with 2 - 3 &#181;L of dye and virus mixture. Advance the needle into the sensory organ while observing it under the binocular dissecting microscope.</li>
	<li>Gently drive the needle tip through mesenchymal and epithelial layers of the roof of a three-dimensional utricular culture. Inject the viral mixture until the cavities of the utricle and ampullae fill with the blue dye. 
	Note: Because it allows easy access to sensory organs, a 10-mm glass-bottom Petri dish is optimal for the injections.</li>
	<li>Incubate at 37 &#176;C in a tissue-culture incubator gassed with 5% carbon dioxide. 
	Note: When green or red fluorescent protein is used in the viral construct, the fluorescence is apparent 24 h after viral transduction.</li>
</ol>

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The authors have nothing to disclose.

The molecular signals that mediate growth and differentiation in the inner ear during development have been studied extensively5,6,7,8,9,10. However, evidence obtained from the utricular model system suggests that mechanical cues, sensed through cell junctions and the activation of Hippo signaling, also play an important role...

The study of most aspects of mammalian organ development has been facilitated by in vitro systems. Two principal methods are now used for the culture of vestibular sensory organs: free-floating1 and adherent2 preparations. Both methods permit the investigation of hair cell vulnerabilities3 and regeneration1,4&#160;in vitro. In addition, the developmental roles of the Notch5,6, Wnt7,8, and epidermal growth factor receptor (EGFR)9,10 signaling cascades in the inner ear have been established, in part, through the use of in vitro cultures of sensory epithelia. However, cell growth and differentiation are controlled, not only through signaling by morphogens, but also through physical and mechanical cues such as intercellular contacts, the stiffness of extracellular matrix, and mechanical stretching or constriction. The role of such mechanical stimuli is challenging to investigate in the developing inner ear in vivo. Moreover, existing free-floating and adherent culture methods are not suitable for such studies in vitro. Here we describe a method for three-dimensional organotypic culture in collagen I gels of varying stiffness. This method largely preserves the in vivo architecture of the vestibular and cochlear sensory organs and allows investigation of the effects of mechanical force on growth and differentiation11.
Because mechanical stimuli are known to activate downstream molecular events, such as the Hippo signaling pathway12,13,14,15, it is important to be able to combine mechanical stimulation with biochemical and genetic manipulations. The culture method described here permits virus-mediated gene delivery and can therefore be used to study both mechanical and molecular signaling during inner ear development11.

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three-dimensional organotypic cultures of vestibular and auditory sensory organs

The sensory organs of the inner ear are challenging to study in mammals due to their inaccessibility to experimental manipulation and optical observation. Moreover, although existing culture techniques allow biochemical perturbations, these methods do not provide a means to study the effects of mechanical force and tissue stiffness during development of the inner ear sensory organs. Here we describe a method for three-dimensional organotypic culture of the intact murine utricle and cochlea that overcomes these limitations. The technique for adjustment of a three-dimensional matrix stiffness described here permits manipulation of the elastic force opposing tissue growth. This method can therefore be used to study the role of mechanical forces during inner ear development. Additionally, the cultures permit virus-mediated gene delivery, which can be used for gain- and loss-of-function experiments. This culture method preserves innate hair cells and supporting cells and serves as a potentially superior alternative to the traditional two-dimensional culture of vestibular and auditory sensory organs.

Vestibular and auditory sensory organs from embryonic ears, cultured in 40-Pa collagen I gels mimicking low stiffness embryonic conditions11, retain relatively normal three-dimensional structures (Figure 1) and maintain hair cells and supporting cells (Figure 2 and Figure 3). Although supporting cell density decreases by over 30% (Student&#39;s t-test: n = 4, p</e...

Watch this Scientific Journal Video about Three-dimensional Organotypic Cultures of Vestibular and Auditory Sensory Organs at JoVE.com

Three-dimensional Organotypic Cultures of Vestibular and Auditory Sensory Organs

Three-dimensional organotypic cultures of the murine utricle and cochlea in optically clear collagen I gels preserve innate tissue morphology, allow for mechanical stimulation through adjustment of matrix stiffness, and permit virus-mediated gene delivery.

The sensory organs of the inner ear are challenging to study in mammals due to their inaccessibility to experimental manipulation and optical observation. ...

three-dimensional-organotypic-cultures-vestibular-auditory-sensory

Research

JoVE Journal

Developmental Biology

8.2K Views.  Keck School of Medicine of the University of Southern California. Driedimensionale organotypic culturen van de lymfkliertest utricle en het slakkenhuis in optisch duidelijk collageen ik gels behouden aangeboren weefsel morfologie, toestaan voor mechanische stimulatie via aanpassing van matrix stijfheid en virus-gemedieerde gene levering toestaan.

Video: Three-dimensional Organotypic Cultures of Vestibular and Auditory Sensory Organs

Vibratome Sectioning for Enhanced Preservation of the Cytoarchitecture of the Mammalian Organ of Corti

The mammalian organ of Corti is a highly ordered cellular mosaic of mechanosensory hair and nonsensory supporting cells (reviewed in 1,2).Visualization of this cellular mosaic often requires that the organ of Corti is cross-sectioned. In particular, the nonsensory pillar and Deiters' cells, whose nuclei are located basally with respect to the hair cells, cannot be visualized without cross-sectioning the organ of Corti. However, the delicate cytoarchitecture of the mammalian organ of Corti, including the fine cytoplasmic processes of the pillar and Deiters' cells, is difficult to preserve by routine histological procedures such as paraffin and cryo-sectioning, which are compatible with standard immunohistochemical staining techniques.
Here I describe a simple and robust procedure consisting of vibratome sectioning of the cochlea, immunohistochemical staining of these vibratome sections in whole mount, followed by confocal microscopy. This procedure has been used widely for immunhistochemical analysis of multiple organs, including the mouse limb bud, zebrafish gut, liver, pancreas, and heart (see 3-6 for selected examples). In addition, this procedure was sucessful for both imaging and quantitificaton of pillar cell number in mutant and control organs of Corti in both embryos and adult mice 7. This method, however, is currently not widely used to examine the mammalian organ of Corti. The potential for this procedure to both provide enhanced preservation of the fine cytoarchitecture of the adult organ of Corti and allow for quantification of various cell types is described.

A simple procedure of vibratome sectioning the organ of Corti, followed by immunohistochemistry and confocal microscopy is described. This procedure allows for improved preservation of the fine cytoarchitecture of the mammalian organ of Corti, and consequently allows for accurate quantification of cell types.

The mammalian organ of Corti is a highly ordered cellular mosaic of mechanosensory hair and nonsensory supporting cells (reviewed in 1,2).Visualization of ...

Neuroscience

Selective Tracing of Auditory Fibers in the Avian Embryonic Vestibulocochlear Nerve

The embryonic chick is a widely used model for the study of peripheral and central ganglion cell projections. In the auditory system, selective labeling of auditory axons within the VIIIth cranial nerve would enhance the study of central auditory circuit development. This approach is challenging because multiple sensory organs of the inner ear contribute to the VIIIth nerve 1. Moreover, markers that reliably distinguish auditory versus vestibular groups of axons within the avian VIIIth nerve have yet to be identified. Auditory and vestibular pathways cannot be distinguished functionally in early embryos, as sensory-evoked responses are not present before the circuits are formed. Centrally projecting VIIIth nerve axons have been traced in some studies, but auditory axon labeling was accompanied by labeling from other VIIIth nerve components 2,3. Here, we describe a method for anterograde tracing from the acoustic ganglion to selectively label auditory axons within the developing VIIIth nerve. First, after partial dissection of the anterior cephalic region of an 8-day chick embryo immersed in oxygenated artificial cerebrospinal fluid, the cochlear duct is identified by anatomical landmarks. Next, a fine pulled glass micropipette is positioned to inject a small amount of rhodamine dextran amine into the duct and adjacent deep region where the acoustic ganglion cells are located. Within thirty minutes following the injection, auditory axons are traced centrally into the hindbrain and can later be visualized following histologic preparation. This method provides a useful tool for developmental studies of peripheral to central auditory circuit formation.

Here we describe a microdissection technique followed by fluorescent dye injection into the acoustic ganglion of early chick embryos for selective tracing of auditory axon fibers in the nerve and hindbrain.

The embryonic chick is a widely used model for the study of peripheral and central ganglion cell projections. In the auditory system, selective labeling ...

An Isolated Semi-intact Preparation of the Mouse Vestibular Sensory Epithelium for Electrophysiology and High-resolution Two-photon Microscopy

Understanding vestibular hair cells function under normal conditions, or how trauma, disease, and aging disrupt this function is a vital step in the development of preventative approaches and/or novel therapeutic strategies. However, the majority of studies looking at abnormal vestibular function have not been at the cellular level but focused primarily on behavioral assays of vestibular dysfunction such as gait analyses and vestibulo-ocular reflex performance. While this work has yielded valuable data about what happens when things go wrong, little information is gleaned regarding the underlying causes of dysfunction. Of the studies that focus on the cellular and subcellular processes that underlie vestibular function, most have relied on acutely isolated hair cells, devoid of their synaptic connections and supporting cell environment. Therefore, a major technical challenge has been access to the exquisitely sensitive vestibular hair cells in a preparation that is least disrupted, physiologically. Here we demonstrate a semi-intact preparation of the mouse vestibular sensory epithelium that retains the local micro-environment including hair cell/primary afferent complexes.

Analysis of vestibular hair cell function is complicated by their location deep within the hardest part of the skull, the petrous temporal bone. Most functional hair cell studies have used acutely isolated hair cells. Here we describe a semi-intact preparation of mouse vestibular epithelium for electrophysiological and two-photon microscopy studies.

Understanding vestibular hair cells function  under normal conditions, or how trauma, disease, and aging disrupt this  function is a vital step in the ...

Primary Culture and Plasmid Electroporation of the Murine Organ of Corti. 

In all mammals, the sensory epithelium for audition is located along the spiraling organ of Corti that resides within the conch shaped cochlea of the inner ear (fig 1). Hair cells in the developing cochlea, which are the mechanosensory cells of the auditory system, are aligned in one row of inner hair cells and three (in the base and mid-turns) to four (in the apical turn) rows of outer hair cells that span the length of the organ of Corti. Hair cells transduce sound-induced mechanical vibrations of the basilar membrane into neural impulses that the brain can interpret. Most cases of sensorineural hearing loss are caused by death or dysfunction of cochlear hair cells.
An increasingly essential tool in auditory research is the isolation and in vitro culture of the organ explant 1,2,9. Once isolated, the explants may be utilized in several ways to provide information regarding normative, anomalous, or therapeutic physiology. Gene expression, stereocilia motility, cell and molecular biology, as well as biological approaches for hair cell regeneration are examples of experimental applications of organ of Corti explants.
This protocol describes a method for the isolation and culture of the organ of Corti from neonatal mice. The accompanying video includes stepwise directions for the isolation of the temporal bone from mouse pups, and subsequent isolation of the cochlea, spiral ligament, and organ of Corti. Once isolated, the sensory epithelium can be plated and cultured in vitro in its entirety, or as a further dissected micro-isolate that lacks the spiral limbus and spiral ganglion neurons. Using this method, primary explants can be maintained for 7-10 days. As an example of the utility of this procedure, organ of Corti explants will be electroporated with an exogenous DsRed reporter gene. This method provides an improvement over other published methods because it provides reproducible, unambiguous, and stepwise directions for the isolation, microdissection, and primary culture of the organ of Corti.

Primary Culture and Plasmid Electroporation of the Murine Organ of Corti.

This procedure describes a method for the isolation and culture of the murine organ of Corti with or without the spiral limbus and spiral ganglion neurons. We also demonstrate a method for the expression of an exogenous reporter gene in the organ of Corti explant by electroporation.

In all mammals, the sensory epithelium for audition is located along the spiraling organ of Corti that resides within the conch shaped cochlea of the ...

Biology

Long-term Time Lapse Imaging of Mouse Cochlear Explants

Here we present a method for long-term time-lapse imaging of live embryonic mouse cochlear explants. The developmental program responsible for building the highly ordered, complex structure of the mammalian cochlea proceeds for around ten days. In order to study changes in gene expression over this period and their response to pharmaceutical or genetic manipulation, long-term imaging is necessary. Previously, live imaging has typically been limited by the viability of explanted tissue in a humidified chamber atop a standard microscope. Difficulty in maintaining optimal conditions for culture growth with regard to humidity and temperature has placed limits on the length of imaging experiments. A microscope integrated into a modified tissue culture incubator provides an excellent environment for long term-live imaging. In this method we demonstrate how to establish embryonic mouse cochlear explants and how to use an incubator microscope to conduct time lapse imaging using both bright field and fluorescent microscopy to examine the behavior of a typical embryonic day (E) 13 cochlear explant and Sox2, a marker of the prosensory cells of the cochlea, over 5 days.

Live imaging of the embryonic mammalian cochlea is challenging because the developmental processes at hand operate on a temporal gradient over ten days. Here we present a method for culturing and then imaging embryonic cochlear explant tissue taken from a fluorescent reporter mouse over five days.

Here we present a method for long-term time-lapse imaging of live embryonic mouse cochlear explants. The developmental program responsible for building ...

In Ovo and Ex Ovo Methods to Study Avian Inner Ear Development

The inner ear perceives sound and maintains balance using the cochlea and vestibule. It does this by using a dedicated mechanosensory cell type known as the hair cell. Basic research in the inner ear has led to a deep understanding of how the hair cell functions, and how dysregulation can lead to hearing loss and vertigo. For this research, the mouse has been the pre-eminent model system. However, mice, like all mammals, have lost the ability to replace hair cells. Thus, when trying to understand cellular therapies for restoring inner ear function, complementary studies in other vertebrate species could provide further insights. The auditory epithelium of birds, the basilar papilla (BP), is a sheet of epithelium composed of mechanosensory hair cells (HCs) intercalated by supporting cells (SCs). Although the anatomical architecture of the basilar papilla and the mammalian cochlea differ, the molecular mechanisms of inner ear development and hearing are similar. This makes the basilar papilla a useful system for not only comparative studies but also to understand regeneration. Here, we describe dissection and manipulation techniques for the chicken inner ear. The technique shows genetic and small molecule inhibition methods, which offer a potent tool for studying the molecular mechanisms of inner ear development. In this paper, we discuss in ovo electroporation techniques to genetically perturb the basilar papilla using CRIPSR-Cas9 deletions, followed by dissection of the basilar papilla. We also demonstrate the BP organ culture and optimal use of culture matrices, to observe the development of the epithelium and the hair cells.

In Ovo and Ex Ovo Methods to Study Avian Inner Ear Development

The chick is a cost-effective, accessible, and widely available model organism for a variety of studies. Here, a series of protocols is detailed to understand the molecular mechanisms underlying avian inner ear development and regeneration.

The inner ear perceives sound and maintains balance using the cochlea and vestibule. It does this by using a dedicated mechanosensory cell type known as ...

In vitro Time-lapse Live-Cell Imaging to Explore Cell Migration toward the Organ of Corti

To study the effects of mesenchymal stem cells (MSCs) on cell regeneration and treatment, this method tracks MSC migration and morphological changes after co-culture with cochlear epithelium. The organ of Corti was immobilized on a plastic coverslip by pressing a portion of the Reissner's membrane generated during the dissection. MSCs confined by a glass cylinder migrated toward cochlear epithelium when the cylinder was removed. Their predominant localization was observed in the modiolus of the organ of Corti, aligned in a direction similarly to that of the nerve fibers. However, some MSCs were localized in the limbus area and showed a horizontally elongated shape. In addition, migration into the hair cell area was increased, and the morphology of the MSCs changed to various forms after kanamycin treatment. In conclusion, the results of this study indicate that the coculture of MSCs with cochlear epithelium will be useful for the development of therapeutics via cell transplantation and for studies of cell regeneration that can examine various conditions and factors.

In this study, we present a real-time imaging method using confocal microscopy to observe cells moving toward damaged tissue by ex vivo incubation with the cochlear epithelium containing the organ of Corti.

To study the effects of mesenchymal stem cells (MSCs) on cell regeneration and treatment, this method tracks MSC migration and morphological changes after ...

Medicine

Isolating and Culturing Vestibular and Spiral Ganglion Somata from Neonatal Rodents for Patch-Clamp Recordings

The compact morphology of isolated and cultured inner ear ganglion neurons allows for detailed characterizations of the ion channels and neurotransmitter receptors that contribute to cell diversity across this population. This protocol outlines the steps necessary for successful dissecting, dissociating, and short-term culturing of the somata of inner ear bipolar neurons for the purpose of patch-clamp recordings. Detailed instructions for preparing vestibular ganglion neurons are provided with the necessary modifications needed for plating spiral ganglion neurons. The protocol includes instructions for performing whole-cell patch-clamp recordings in the perforated-patch configuration. Example results characterizing the voltage-clamp recordings of hyperpolarization-activated cyclic nucleotide-gated (HCN)-mediated currents highlight the stability of perforated-patch recording configuration in comparison to the more standard ruptured-patch configuration. The combination of these methods, isolated somata plus perforated-patch-clamp recordings, can be used to study cellular processes that require long, stable recordings and the preservation of intracellular milieu, such as signaling through G-protein coupled receptors.

Presented here are methods providing detailed instructions for dissecting, dissociating, culturing, and patch-clamp recording from vestibular ganglion and spiral ganglion neurons of the inner ear.

The compact morphology of isolated and cultured inner ear ganglion neurons allows for detailed characterizations of the ion channels and neurotransmitter ...

Whole Neonatal Cochlear Explants as an In vitro Model

Untreated hearing loss imposes significant costs on the global healthcare system and impairs individuals' quality of life. Sensorineural hearing loss is characterized by the cumulative and irreversible loss of sensory hair cells and auditory nerves in the cochlea. Entire and vital cochlear explants are one of the fundamental tools in hearing research to detect hair cell loss and to characterize the molecular mechanisms of the inner ear cells. Many years ago, a protocol for neonatal cochlear isolation was developed, and although it has been modified over time, it still holds potential for improvement. 
This paper presents an optimized protocol for isolating and culturing whole neonatal cochlear explants in multi-well culture chambers that enables the study of hair cells and spiral ganglion neuron cells along the entire length of the cochlea. The protocol was tested using cochlear explants from mice and rats. Healthy cochlear explants were obtained to study the interaction between hair cells, spiral ganglion neuron cells, and the surrounding supporting cells. 
One of the main advantages of this method is that it simplifies the organ culture steps without compromising the quality of the explants. All three turns of the organ of Corti are attached to the bottom of the chamber, which facilitates in vitro experiments and the comprehensive analysis of the explants. We provide some examples of cochlear images from different experiments with live and fixed explants, demonstrating that the explants retain their structure despite exposure to ototoxic drugs. This optimized protocol can be widely used for the integrative analysis of the mammalian cochlea.

Whole Neonatal Cochlear Explants as an In vitro Model

The current protocol updates previous protocols and incorporates relatively straightforward approaches for culturing high-quality cochlear explants. This provides reliable data acquisition and high-resolution imaging in live and fixed cells. This protocol supports the ongoing trend of studying inner ear cells.

Untreated hearing loss imposes significant costs on the global healthcare system and impairs individuals' quality of life. Sensorineural hearing loss is ...

Harvest of Vestibular End-Organs under Physiologic Conditions during Labyrinthectomy

The living human inner ear is challenging to study because it is encased within dense otic capsule bone that limits access to biological tissue. Traditional temporal bone histopathology methods rely on lengthy, expensive decalcification protocols that take 9-10 months and reduce the types of tissue analysis possible due to RNA degradation. There is a critical need to develop methods to access fresh human inner ear tissue to better understand otologic diseases, such as M&#233;ni&#232;re's disease, at the cellular and molecular level. This paper describes a technique for the harvest of human vestibular end organs from a living donor under physiologic conditions. An individual with M&#233;ni&#232;re's disease and 'drops attacks' that were refractory to intratympanic gentamicin injection underwent labyrinthectomy. A traditional mastoidectomy was first performed, and the horizontal and superior semicircular canals (SCC) were identified. The mastoid cavity was filled with a balanced salt solution so that the labyrinth could be opened under more physiologic conditions to preserve cellular integrity. A zero-degree endoscope fit with a lens-cleaning sheath irrigation system was used to visualize the submerged mastoid cavity, and a 2 mm diamond burr was used to skeletonize and open the horizontal and superior SCCs, followed by the vestibule. The ampullae and portion of the canal ducts for the superior and lateral SCCs were harvested. The utricle was similarly harvested. Harvested tissue was immediately placed in an ice-cold buffer and then fixed for one hour in 4% paraformaldehyde in phosphate-buffered saline (PBS). The tissue was rinsed several times in 1x PBS and stored for 48 h at 4 &#176;C. The tissue samples underwent immunostaining with a combination of primary antibodies against tenascin-C (Calyx), oncomodulin (streolar hair cells), calretinin (Calyx and Type II hair cells), synaptic vesicle protein 2 (efferent fibers and boutons), &#946;-tubulin 1 (Calyx and afferent boutons), followed by incubation with fluorophore-conjugated secondary antibodies. The tissue samples were then rinsed and mounted for confocal microscopy examination. Images revealed the presence of ampullar and macular hair cells and neural structures. This protocol demonstrates that it is possible to harvest intact, high-quality human inner ear tissue from living donors and may provide an important tool for the study of otologic disease.

Here, we describe a technique for harvesting human vestibular end-organs under physiologic conditions during labyrinthectomy and their analysis using immunostaining.

Three-dimensional Organotypic Cultures of Vestibular and Auditory Sensory Organs

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