The current project was funded by the University of Kansas Proof of Concept Fund. We would like to thank the nursing staff at KUMC (Kansas City, KS) for assisting us in obtaining human umbilical cords, and David Jorgensen for assisting with cochleae cultures.

All procedures, including animal euthanasia, were conducted according to the approved Institutional Animal Care and Use Committee (IACUC) protocol (ACUP #2014-2234) at the University of Kansas Medical Center (KUMC).
NOTE: HWJCs were isolated from human umbilical cords that were donated by patients that provided informed consent and specimens were used in accordance with the protocols approved by the University of Kansas Human Subjects Committee (KU-IRB #15402).
1. Temporal Bone Harvest and Cochlea Isolation
<ol>
	<li>Decapitate an appropriately euthanized, 15-week-old mouse by cutting between the skull and first cervical vertebra with a sharp pair of surgical scissors, then spray down the head with 70% ethanol to disinfect (Figure 1A).</li>
	<li>Bisect the skull in a mid-sagittal plane using the same sharp pair of surgical scissors (Figure 1B-C, dashed lines).</li>
	<li>Remove the brain tissue from the skull using a pair of forceps (Figure 1D, arrowhead).</li>
	<li>Identify the temporal bone through the presence of the auditory and vestibular nerve roots (Figure 1E-F, arrowheads) on the interior of the skull and the ear canal from the exterior (Figure 1F, arrow). Carefully cut through the skull to isolate the temporal bone from the remainder of the skull (Figure 1G). 
	NOTE: In some instances, it may be possible to delicately pry the surrounding bones of the skull away from the temporal bone without requiring cutting.</li>
	<li>Insert the fine forceps into the opening of the ear canal approximately 5 mm (Figure 1H), so the tips do not risk puncturing the cochlea. Gently break open the bone of the bulla (Figure 1H, top red shaded area) so it fractures (Figure 1I, red dotted line). 
	NOTE: If necessary, a dissection microscope may aid in this process to ensure accurate placement and manipulation of instruments.</li>
	<li>Using fine forceps, pry away the remainder of the bulla from the temporal bone, exposing the bony labyrinth of the cochlea (Figure 1J, red brackets).</li>
	<li>Place a Petri dish large enough to hold the temporal bone (e.g., 30 mm or greater) underneath the dissecting scope field of view and fill it with enough Phosphate Buffered Saline (PBS) to cover the temporal bone (e.g., 0.5 to 1 mL).</li>
	<li>Submerge the temporal bone with the bulla removed in the PBS with the oval and round windows of the cochlea facing up. 
	NOTE: Removal of the vestibular system from the cochlea is not necessary, as the vestibular system serves well as a handle for manipulating and orienting features of the cochlea (Figure 1I and Figure 2B).</li>
	<li>Using an ultra-fine pair of forceps, remove the stapedial artery, which passes through the stapes.</li>
	<li>Insert one tip of the ultra-fine forceps through the arch of the stapes where the artery passed through, and delicately lift the stapes upward. The disarticulated stapes should lift off of the oval window.</li>
	<li>Using one tip of the ultra-fine forceps, puncture the oval and round windows.</li>
	<li>Using a 1x PBS filled 28.5-gauge syringe with tubing (inner diameter 0.28 mm, outer diameter 0.61 mm) attached, position the tubing over the oval window (Figure 2A - B). 
	NOTE: The opening of the tube can be manipulated using ultra-fine forceps helping to ensure accurate placement.
	<ol>
		<li>Use a fresh syringe and tubing for each cochlea.</li>
	</ol>
	</li>
	<li>Perfuse 2 mL of 10% antibiotic-antimycotic (anti-anti) and 10% penicillin-streptomycin (pen-strep) in PBS through the cochlea over 5 min to remove the perilymph. Perfusing too fast with too much pressure will damage the fine structures within the cochlea. 
	NOTE: Manually driving the syringe by hand to perfuse the fluid through the cochlea is effective, although a perfusion pump could be used as an alternative.
	<ol>
		<li>If manually perfusing, use a pair of fine, self-closing forceps to stabilize the cochlea (Figure 2A) during perfusion by grasping the vestibular portion of the temporal bone. 
		NOTE: Although not required, this leaves both hands free to handle instruments; one hand can drive the syringe (Figure 2A) while the other can position the tubing appropriately (Figure 2B).</li>
		<li>From this step forward, take care to avoid exposing the cochlea unnecessarily to air. Introducing air bubbles into the fluid spaces will block fluid circulation, and further drying out will damage the fine structures within the cochlea.</li>
	</ol>
	</li>
	<li>Remove and discard any remaining muscle tissue and bone fragments using fine forceps.</li>
	<li>If necessary, store isolated cochleae at 4 &#176;C in 10% anti-anti and 10% pen-strep solution in PBS for up to seven days with regular changes of antibiotic solution every 48 h.</li>
</ol>
2. Cochlear Processing
<ol>
	<li>Decellularization
	<ol>
		<li>For each cochlea, fill a 20-mL glass scintillation vial with a 1% sodium dodecyl sulfate (SDS) solution in de-ionized (DI) water, which is used to decellularize the cochlea.</li>
		<li>Cut the tip off of a plastic 7-mL transfer pipette using a razor blade so that the opening is large enough for a cochlea to pass through.</li>
		<li>Using the transfer pipette, gently draw the cochlea up into the pipette so it just passes the cut-off edge by a few millimeters.</li>
		<li>Expel the cochlea into the 1% SDS solution-filled scintillation vial.</li>
		<li>Circulate 2 mL of 1% SDS in de-ionized (DI) water through the cochlea in the scintillation vial&#160;using a transfer pipette.</li>
		<li>Place the scintillation vial into a rotator, and allow the scintillation vial to rotate at 10 rpm for 72 h at room temperature.
		<ol>
			<li>Change 1% SDS solution every 24 h over a 72 h period. Take care when changing SDS to never expose the cochlea to air.</li>
		</ol>
		</li>
		<li>Wash the cochlea three times for 30 min each with DI water. Perform the washes in the same scintillation vial used for the SDS charges; at this stage, the cochlea is decellularized.</li>
	</ol>
	</li>
	<li>Decalcification
	<ol>
		<li>Remove the remaining DI water from the scintillation vial, leaving just enough to keep the cochlea submerged.</li>
		<li>To begin decalcification, add 5 mL of 10% ethylenediaminetetraacetic acid (EDTA) (0.02 M) in DI water to the scintillation vial containing the cochlea.</li>
		<li>Circulate 2 mL of 10% EDTA in DI water through the cochlea in the scintillation vial&#160;using a transfer pipette.</li>
		<li>Place the scintillation vial back in the rotator, and allow the scintillation vial to rotate at 10 rpm for 72 h at room temperature. Change the 10% EDTA solution every 24 h over a 72 h period. Take care when changing solutions so that the cochlea is not exposed to air.</li>
		<li>Rinse the cochlea 3 times for 2 h each with 1x PBS; at this stage, the cochlea is decalcified.</li>
	</ol>
	</li>
	<li>Storage
	<ol>
		<li>Store the cochleae for up to 72 h in a 10% anti-anti, 10% pen-strep solution in PBS at 4 &#176;C. 
		NOTE: Longer storage periods may be possible with regular changes of antibiotic solution; however, extended storage of processed cochleae has not been validated in this protocol.</li>
	</ol>
	</li>
</ol>
3. Procurement and Expansion of hWJCs
<ol>
	<li>Isolate hWJCs according to previously published protocols22. Briefly, segment umbilical cords into 3-cm sections.</li>
	<li>Carefully make a shallow incision with a scalpel lengthwise on each umbilical cord segment to unroll and expose the Wharton&#39;s jelly of the umbilical cord. Use forceps to remove the blood vessels.</li>
	<li>Wash umbilical cord segments twice with enough PBS (e.g., 40 mL in a 60 mm Petri dish) to submerge the tissue. Finely mince tissue segments with a scalpel. Digest tissue in a 100 mm Petri dish with 50 mL of digestion medium (0.2% type 2 collagenase, 1% penicillin-streptomycin, in low-glucose Dulbecco&#39;s Modified Eagle&#39;s Medium (DMEM)).</li>
	<li>Place tissue in digesting medium on an orbital shaker at 50 rpm overnight in a 5% CO2, 37 &#176;C incubator. The following day, dilute digestion medium at a ratio of 1:16 in 2% antibiotic-antimycotic in PBS, and pellet cells by centrifugation at 500 x g at room temperature (~ 27 &#176;C) for 10 min. Discard supernatant.</li>
	<li>Resuspend pellets in Mesenchymal Stem Cell Growth Medium (MSCGM) and recombine cells. Plate cells at a density of 7 x 103 cells/cm2 in tissue culture treated T-75 flasks. Culture HWJCs in MSCGM, and expand to passage 5 for experiments. 
	NOTE: HWJCs may be sub-cultured into larger T-flasks (e.g., T-150, T-300) when passaging to increase yield of hWJCs for experiments.</li>
</ol>
4. Infusion of hWJCs into Decellularized Cochleae
<ol>
	<li>Prepare decellularized cochleae
	<ol>
		<li>Using aseptic technique, wash stored cochleae in 1x PBS three times for 30 min each.</li>
		<li>Transfer cochleae into separate wells of a 24-well plate, and add 1 mL of 37 &#176;C pre-warmed MSCGM.</li>
		<li>Incubate cochleae in a 5% CO2, 37 &#176;C cell culture incubator for a minimum of 1 h before infusion of cells.</li>
	</ol>
	</li>
	<li>Dissociate hWJCs and resuspend
	<ol>
		<li>Wash hWJCs with 37 &#176;C pre-warmed 1x PBS twice.</li>
		<li>Add enough 37 &#176;C pre-warmed Trypsin with 0.5% EDTA to cover the cell vessel surface.</li>
		<li>Incubate hWJCs for up to 5 min in a 5% CO2, 37 &#176;C cell culture incubator.
		<ol>
			<li>Verify that 90% of cells have detached from the cell culture surface by viewing cells under an inverted microscope. 
			NOTE: Cells that move when the culture vessel is gently tapped, have detached. Cells that remain stationary, have not detached.</li>
			<li>Gently tap the sides of the flask to further dislodge attached cells.</li>
		</ol>
		</li>
		<li>Using aseptic technique, transfer hWJCs from the culture vessel to a 50-mL conical tube containing an equivalent volume of MSCGM to the volume of Trypsin used to dissociate the cells.</li>
		<li>Pellet the hWJCs by centrifuging at 500 x g at room temperature (~ 27 &#176;C) for 5 min.</li>
		<li>Aspirate supernatant, and resuspend hWJCs in MSCGM at a concentration of 500,000 cells/mL.</li>
	</ol>
	</li>
	<li>Perfuse cochleae
	<ol>
		<li>Transfer cochleae to a new 24-well plate using a transfer pipette.</li>
		<li>Add a drop of MSCGM to each cochlea to prevent any cochlea from drying out.</li>
		<li>Delicately orient the cochlea using ultra-fine forceps so that the oval and round windows are facing up. 
		NOTE: Take extreme care when directly handling each cochlea as the cochlea is easily damaged.</li>
		<li>Using a sterile 28.5-gauge insulin syringe with connected tubing, draw up 0.2 mL resuspended hWJCs (100,000 cells).</li>
		<li>Using sterilized fine forceps, position tubing over the oval window.</li>
		<li>Delicately and slowly perfuse the cochlea with 0.2 mL of resuspended hWJCs using a 28.5-gauge insulin syringe connected to polyethylene tubing (Outer Diameter: 0.61 mm, Inner Diameter: 0.28 mm) over approximately 5 min.</li>
		<li>After perfusion, add 0.8 mL of 37 &#176;C pre-warmed MSCGM to the well containing the cochlea to bring the total volume in the well to 1 mL.</li>
		<li>Repeat steps 4.3.3-4.3.7 for each cochlea, using a fresh syringe and tubing each time.</li>
		<li>Place perfused cochleae in a 5% CO2, 37 &#176;C cell culture incubator, and change media three times per week.</li>
	</ol>
	</li>
</ol>
5. Cochlea Harvest and Preservation
NOTE: Cochleae may be cultured and harvested at any time point up to 30 days post-perfusion.
<ol>
	<li>To preserve cochleae, fix in 4% paraformaldehyde in 1x PBS overnight at 4 &#176;C on a rocking platform.</li>
	<li>After fixation, wash cochleae with 1x PBS three times for 5 min each.</li>
	<li>Gradually dehydrate cochlea with ethanol and clear with an aliphatic hydrocarbon solvent before embedding in paraffin.</li>
	<li>Section samples to a thickness of 10 &#181;m using a microtome and mount on glass microscope slides. 
	NOTE: Samples are now ready for histological or immunohistochemical processing using standard protocols.</li>
</ol>

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</ol>

The authors have nothing to disclose.

We have successfully demonstrated that native cochlear cells can be removed from the cochlea via a decellularization process, which allows for the use of the cochlea as an intricate, three-dimensional tissue scaffold. Santi et al.15 developed the initial method for decellularizing cochleae, and have accurately estimated the volumes of many cochlear structures through with the aid of light sheet microscopy23. Such early work served as a strong basis for the tissue engineering and cell culture techniques presented here. Decellularized cochlea can successfully be infused with cells, and then cultured for a prolonged period of time. Cochleae from a wide array of animals could theoretically be utilized in combination with a similarly wide array of cells types. These combinations allow the techniques presented here to be utilized in numerous possible experimental designs, such as tissue engineering applications that examine sensory epithelium development14, drug testing of new pharmaceutical agents24, and development of cochlear repair models25. However, despite the broad applicability of these techniques they are not without limitations.
One of the limitations of the current technique is that the perfusion is variable based on individual. Developing a holder, for the cochlea, and using a syringe pump to conduct perfusions may increase the accuracy and success of the process. Additionally, definitively proving successful decellularization could potentially be accomplished through several methods. We employed DAPI staining that was used to quantify remaining cell nuclei, and we observed no residual nuclei post-decellularization. Additionally, standard DNA quantification assays utilize the reagent, green dye detecting picogram quantities of DNA, to identify nucleic acid remnants, which may visually be more sensitive than DAPI for identifying the presence of small fragments of nucleic acids in decellularized tissues26. Regardless of which techniques are used to validate successful decellularization, it may not be possible to show complete decellularization of an individual cochlea prior to utilizing it as a scaffold. Nevertheless, we have found the decellularization process to be successful and robust.
The two most critical steps in the current protocol are the isolation and handling of the cochleae, and the perfusion of the cochleae. Great care must be taken when the cochlea is initially isolated, as the cochlea is delicate and brittle. If too much force is applied while handling the cochlea with forceps, the cochlea could fragment. Thus, it is imperative to handle each cochlea gently before decellularization and decalcification. The vestibular system, if left intact, serves as a great handle for grabbing the inner ear complex with forceps, and orienting the cochlea. Similarly, when perfusing the cochlea with cells, the pressure behind the perfusion must not be too great, otherwise cells may not survive the perfusion. As noted in the current protocol, attaching tubing to the end of a 1-mL insulin syringe allows for easier handling of the cochlea with one hand, and greater precision in perfusing cells through the cochlea with the other hand.
As a next step, it would be natural to assess developmental and functional markers to determine if the cochlear ECM is inducing differentiation of stem cells, and if so, which specific ECM components in the cochlea are inducing changes in cells. Perfusing different types of cells (e.g., stem cells, neuroprogenitors, fibroblasts, etc.) through the cochlea to monitor cell differentiation and behavior would be another fascinating follow-up investigation. In addition, cells that have been genetically modified or are exposed to a growth factor differentiation protocol may provide new insights into how the cochlear ECM supports neurosensory development, and possibly even function in hearing. Thus, the current protocol is a significant first step in using the cochlear ECM to study inner ear neurosensory development and maintenance, which may one day lead to the development of new therapies to restore hearing loss.

The cochlea is an intricate spiral structure found in the temporal bone. It is composed of an outer bony labyrinth and a concentric, inner membranous labyrinth1. The membranous labyrinth consists of three fluid spaces: Scala vestibuli, Scala media, and Scala tympani1. The scala media houses the sensory epithelium, which is composed of a multitude of cell types, but the sensory hair cells (HC), which transduce mechanical energy in sound waves to nerve impulses2, are of particular interest. Exposure to acoustic trauma3,4,5, medication6, disease7,8, and aging9 can all result in impaired auditory function via HC death. Hair cell loss in mammals is permanent, unlike avian HCs, which can regenerate after injury10.
A variety of contemporary research efforts have sought to restore lost HCs, although the specific experimental approaches vary. Manipulation of gene expression in the sensory epithelium and implantation of stem cells differentiated outside the body are dominant approaches in the field, although induction methods that seek to differentiate stem cells into cochlear organoids have been attempted11,12,13. Each of these approaches is either reliant directly on stem cells, or the developmental cues used by stem cells; however, a second shared, and potentially critical, element is the ECM of the cochlea itself14,15.
The ECM not only provides physical support for cells and tissue, which includes a surface for cell adhesion, proliferation, survival, and migration, but also plays critical roles in the development of HCs and the spiral ganglion15,16,17. Naturally occurring ECM provides inductive signals that may guide cell phenotype determination and/or cell adhesion, proliferation, and survival18. Consequently, the use of decellularized cochlea in combination with cultured hWJCs offer a unique opportunity to explore the role of the ECM and HC regeneration. HWJCs are a readily available, non-controversial cell type isolated from human umbilical cords that behave like mesenchymal stem cells19. HWJCs have shown the ability to differentiate down neurosensory cell lineages20,21. Thus, the current protocol details the isolation, decellularization, and perfusion of cochleae from C57BL mouse carcasses with hWJCs for inner ear tissue engineering.

<table><tbody><tr><td>Allegra X-14R Centrifuge</td><td>Beckman-Coulter</td><td>B08861</td><td></td></tr><tr><td>Intramedic Semi-Rigid Tubing</td><td>Becton Dickinson</td><td>427401</td><td></td></tr><tr><td>New Brunswick Innova 2000 Orbital Shaler</td><td>Eppendorf</td><td>M1190-0002</td><td></td></tr><tr><td>Surgical Scissors</td><td>Fine Science Tools</td><td>14060-10</td><td></td></tr><tr><td>Fine Forceps</td><td>Fine Science Tools</td><td>11370-40</td><td></td></tr><tr><td>Ultra-Fine Forceps</td><td>Fine Science Tools</td><td>18155-13</td><td></td></tr><tr><td>50-mL Conical Tubes</td><td>Fisher Scientific</td><td>12565271</td><td></td></tr><tr><td>Petri Dish</td><td>Fisher Scientific</td><td>FB087579B</td><td></td></tr><tr><td>U-100 Insulin Syringe</td><td>Fisher Scientific</td><td>14-829-1B</td><td></td></tr><tr><td>Scintillation Vial</td><td>Fisher Scientific</td><td>03-341-73</td><td></td></tr><tr><td>Rotator</td><td>Fisher Scientific</td><td>88-861-049</td><td></td></tr><tr><td>Transfer Pipette</td><td>Fisher Scientific</td><td>22-170-404</td><td></td></tr><tr><td>Razor Blade</td><td>Fisher Scientific</td><td>12-640</td><td></td></tr><tr><td>Antibiotic-Antimycotic</td><td>Fisher Scientific</td><td>15-240-062</td><td></td></tr><tr><td>Penicillin-Streptomycin</td><td>Fisher Scientific</td><td>15-140-122</td><td></td></tr><tr><td>24-Well Plate</td><td>Fisher Scientific</td><td>07-200-84</td><td></td></tr><tr><td>SuperFrost PLUS Glass Microscope Slides</td><td>Fisher Scientific</td><td>12-550-15</td><td></td></tr><tr><td>Transfer Pipette</td><td>Fisher Scientific</td><td>22-170-404</td><td></td></tr><tr><td>ProLong Gold Antifade Mountant with DAPI</td><td>Fisher Scientific</td><td>P36935</td><td></td></tr><tr><td>Clear-Rite 3</td><td>Fisher Scientific</td><td>22-046341</td><td></td></tr><tr><td>Thermo Scientific Forma Series II 3110 Water-Jacekted CO&lt;sub&gt;2&lt;/sub&gt; Incubator</td><td>Fisher Scientific</td><td>13-998-078</td><td></td></tr><tr><td>Mesenchymal Stem Cell Growth Medium</td><td>Lonza</td><td>PT-3001</td><td></td></tr><tr><td>Trypsin-EDTA</td><td>Lonza</td><td>CC-3232</td><td></td></tr><tr><td>TPP T-75 Culture Flask</td><td>MidSci</td><td>TP90076</td><td></td></tr><tr><td>TPP T-150 Culture Flask</td><td>MidSci</td><td>TP90151</td><td></td></tr><tr><td>TPP T-300 Culture Flask</td><td>MidSci</td><td>TP90301</td><td></td></tr><tr><td>Dissection Microscope</td><td>Nikon Instruments</td><td>SMZ800</td><td></td></tr><tr><td>Nikon Eclipse Ts2R-FL Inverted Microscope</td><td>Nikon Instruments</td><td>MFA51010</td><td></td></tr><tr><td>NuAire Class II, Type A2 Biosafety Cabinet</td><td>NuAire</td><td>NU-425-600</td><td></td></tr><tr><td>1X PBS</td><td>Sigma-Aldrich</td><td>P5368-10PAK</td><td></td></tr><tr><td>1% SDS Solution</td><td>Sigma-Aldrich</td><td>436143-100G</td><td></td></tr><tr><td>10% EDTA</td><td>Sigma-Aldrich</td><td>E9884-100G</td><td></td></tr></tbody></table>

a protocol for decellularizing mouse cochleae for inner ear tissue engineering

In mammals, mechanosensory hair cells that facilitate hearing lack the ability to regenerate, which has limited treatments for hearing loss. Current regenerative medicine strategies have focused on transplanting stem cells or genetic manipulation of surrounding support cells in the inner ear to encourage replacement of damaged stem cells to correct hearing loss. Yet, the extracellular matrix (ECM) may play a vital role in inducing and maintaining function of hair cells, and has not been well investigated. Using the cochlear ECM as a scaffold to grow adult stem cells may provide unique insights into how the composition and architecture of the extracellular environment aids cells in sustaining hearing function. Here we present a method for isolating and decellularizing cochleae from mice to use as scaffolds accepting perfused adult stem cells. In the current protocol, cochleae are isolated from euthanized mice, decellularized, and decalcified. Afterward, human Wharton&#39;s jelly cells (hWJCs) that were isolated from the umbilical cord were carefully perfused into each cochlea. The cochleae were used as bioreactors, and cells were cultured for 30 days before undergoing processing for analysis. Decellularized cochleae retained identifiable extracellular structures, but did not reveal the presence of cells or noticeable fragments of DNA. Cells perfused into the cochlea invaded most of the interior and exterior of the cochlea and grew without incident over a duration of 30 days. Thus, the current method can be used to study how cochlear ECM affects cell development and behavior.

Using the methods presented here, successful decellularization of cochleae was assessed by examining the presence or absence of DNA through 4&#39;,6-diamidino-2-phenylindole (DAPI) staining. Cochleae were considered fully decellularized if DNA was not identified within the decellularized cochlea. A native cochlea from a previous experiment that did not undergo decellularization or decalcification was used as a positive control to illustrate the structures and cells traditionally found in a C57BL mouse cochlea. The decellularized-decalcified cochlea, which did not have any hWJCs infused, was used as a negative control. No quantifiable DAPI stained cell nuclei were observed in any region of the tissue section (Figure 3). Two cochleae were decellularized and decalcified, and then infused with hWJCs for the current project. The decellularized-decalcified cochleae that were perfused with hWJCs were observed to have numerous DAPI stained nuclei within the cochlea and growing on the bony shell outside of the cochlea (Table 1). Total estimated cell densities for the cell infused first cochlea were 113,759 cells/cochlea after 30 days of culture (Figure 4 and Figure 5) and for the second cochlea were 118,732 cells/cochlea after 30 days of culture (Figure 6 and Figure 7). These cell density estimates include the Scala tympani, Scala media, and Scala vestibule, and volumes of the structures found within the three fluid spaces, but these estimates excluded any of the remaining bony labyrinth. In addition, samples from each cochlea were stained with Hematoxylin and Eosin (H&#38;E) to show the presence of cells and anatomical features within each cochlea (Figure 8). The gross anatomy of the cochlear microstructures remained predominantly unchanged throughout all stained sections; however, no cells were identifiable in the decellularized-decalcified section (Figure 8B). The identification of cell nuclei was dramatically different between the native cochlea (Figure 8A) and the decellularized-decalcified cochleae infused with cells (Figure 8C-D).
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/56523/56523fig1.jpg" /> 
Figure 1: Mouse temporal bone isolation. Once the mouse has been appropriately euthanized, decapitate the animal by cutting between the base of the skull and the first cervical vertebra, and spray the head with ethanol to disinfect (A). Bisect the skull in a sagittal plane of section (B-C) using a sharp pair of surgical scissors, cutting through both the brain tissue and bone. The bisected mouse head (D, lower mandible removed for unrelated experimental work) must then have the brain tissue removed to reveal the temporal bone. The two foramina through which the auditory and vestibular nerves pass (E, red arrowheads) are useful cues for successfully identifying the temporal bone. Remove the flesh from the skull (F). The external ear canal (F, arrow, top panel) provides a useful external cue as well. Carefully trim away the excess bone, leaving just the isolated temporal bone (G). The bulla (H, top panel, red highlighted area) must then be delicately removed using fine forceps (H, lower panel). The bulla is thin bone and can be readily chipped away (I) until the bony labyrinth of the cochlea is revealed (J, red brackets). Scale bars on all images are 1 mm. <a href="https://cloudflare.jove.com/files/ftp_upload/56523/56523fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/56523/56523fig2.jpg" /> 
Figure 2: Cochlear Perfusion. Once the temporal bone has been isolated and the bulla has been removed, the cochlea can then be set up for perfusion at a microscope. When perfusing using a manually driven syringe with tubing attached (A, dotted line) it may be helpful to stabilize the cochlea using a pair of self-closing forceps. Gently attach the forceps to the vestibular part of the temporal bone (B) and rest the forceps against the lip of the Petri dish (A). This leaves both hands free to manipulate instruments during perfusion. One hand can then manipulate the syringe, controlling the flow rate of the fluid, while the other hand can position the tubing over the oval window using a second pair of forceps (B). Cutting the free end of the tubing at an angle allows for easier positioning, allowing the tubing to be placed correctly without blocking the field of view. Scale bar is 1 mm. <a href="https://cloudflare.jove.com/files/ftp_upload/56523/56523fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/56523/56523fig3.jpg" /> 
Figure 3: Tissue section of a decellularized mouse cochlea. An upright epi-fluorescent microscope was used to image samples at a magnification of 200X (10X eyepiece and 20X objective). Images were stitched together to create approximately a 4 x 5 montage. DAPI nuclear staining showing absence of cells is shown in panel A using a DAPI common filter set (350 nm excitation, 470 nm emission), and an overexposed, grayscale image of the autofluorescence using a fluorescein isothiocyanate (FITC) common filter set (490 nm excitation, 525 nm emission), which provides anatomical landmarks, is shown in panel B. Automated cell counts were performed in three regions of interest, which are delineated on both panels A and B. Total cell counts were calculated for the entire tissue section (1, solid white line) and the cochlea (2, dashed line). These two numbers were used to calculate the number of cells in the bony structures outside of the cochlea but within the exterior, bony edges of the tissue section (3, space between the dotted line and dashed line). Scale bars on all images are 500 &#181;m. <a href="https://cloudflare.jove.com/files/ftp_upload/56523/56523fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/56523/56523fig4.jpg" /> 
Figure 4: A near-modiolar section of decellularized mouse cochlea 1 infused with hWJCs. DAPI nuclear staining is shown in panel A overlaid onto an overexposed, grayscale autofluorescence from the green channel, which provides anatomical landmarks. Automated cell counts were performed in three regions of interest, which are delineated in panel A. Total cell counts were calculated for the entire tissue section (1, solid white line) and the cochlea (2, dashed line). These two numbers were used to calculate the number of cells in the bony structures outside of the cochlea but within the exterior, bony edges of the tissue section (3, space between the dotted line and dashed line). Isolated DAPI staining is shown in panel B, and the threshold of the DAPI staining used during automated quantification is shown in panel C. Scale bars on all images are 500 &#181;m. <a href="https://cloudflare.jove.com/files/ftp_upload/56523/56523fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/56523/56523fig5.jpg" /> 
Figure 5: High magnification view of a near-modiolar section of decellularized mouse cochlea 1 infused with hWJCs. DAPI nuclear staining is shown in panel A overlaid onto an overexposed, grayscale image of the autofluorescence from the green channel, which provides anatomical landmarks. Isolated DAPI staining is shown in panel B, and the threshold image used during automated cell counting is shown in panel C. The fluid spaces and finer microstructures housed within the cochlea are nicely preserved during the decellularization and cell culture phases of the experiment. Scala Vestibuli (SV), Scala Tympani (SM), Basilar Membrane (BM), Tectorial Membrane (TM), Spiral Limbus (L), Rosenthal&#39;s Canal (RC), Modiolus (M), Spiral Ligament (SL), and Stria Vascularis (*). Reissner&#39;s membrane was not clearly defined in tissue sections, and as a result the Scala Media was not clearly defined. Scale bars on all images are 250 &#181;m. <a href="https://cloudflare.jove.com/files/ftp_upload/56523/56523fig5large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 6" class="xfigimg" src="/files/ftp_upload/56523/56523fig6.jpg" /> 
Figure 6: A near-modiolar section of decellularized mouse cochlea 2 that was infused with hWJCs. DAPI nuclear staining is shown in panel A, overlaid onto grayscale autofluorescence from the green channel, which provides anatomical landmarks. Automated cell counts were performed in three regions of interest, which are delineated in panel A. Total cell counts were calculated for the entire tissue section (1, solid white line) and the cochlea (2, dashed line). These two numbers were used to calculate the number of cells in the bony structures outside of the cochlea but within the exterior, bony edges of the tissue section (3, space between the dotted line and dashed lines). Isolated DAPI staining is shown in panel B, and the threshold of the DAPI staining used during automated quantification is shown in panel C. Scale bars on all images are 500 &#181;m. <a href="https://cloudflare.jove.com/files/ftp_upload/56523/56523fig6large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 7" class="xfigimg" src="/files/ftp_upload/56523/56523fig7.jpg" /> 
Figure 7: High magnification view of a near-modiolar section of decellularized mouse cochlea 2 that was infused with hWJCs. DAPI nuclear staining is shown in panel A, overlaid onto grayscale autofluorescence from the green channel, which provides anatomical landmarks. Isolated DAPI staining is shown in panel B, and the threshold image used during automated cell counting is shown in panel C. The fluid spaces and finer microstructures housed within the cochlear are preserved during the decellularization and cell culture phases of the experiment. Scala Vestibuli (SV), Scala Tympani (SM), Basilar Membrane (BM), Tectorial Membrane (TM), Spiral Limbus (L), Rosenthal&#39;s Canal (RC), Modiolus (M), Spiral Ligament (SL), and Stria Vascularis (*). Reissner&#39;s Membrane was not clearly defined in tissue sections, and as a result the Scala Media was not clearly defined. Scale bars on all images are 250 &#181;m. <a href="https://cloudflare.jove.com/files/ftp_upload/56523/56523fig7large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 8" class="xfigimg" src="/files/ftp_upload/56523/56523fig8.jpg" /> 
Figure 8: Hematoxylin and Eosin staining of control and treated cochleae. Hematoxylin and Eosin stained sections displaying a typical cochlea with native cells present is shown in panel A. Panel B contains the same structures as panel A, but is from a fully decellularized cochlea, which leaves behind the extracellular matrix. The two previously shown cochleae infused with hWJCs are seen in panels C and D. The gross cochlear anatomy remains predominantly unchanged through the decellularization and cell culture processes, although the specific cell populations and locations are dramatically altered. Scala Vestibuli (SV), Scala Tympani (SM), Basilar Membrane (BM), Tectorial Membrane (TM), Spiral Limbus (L), Rosenthal&#39;s Canal (RC), Modiolus (M), Spiral Ligament (SL), and Stria Vascularis (*). Scale bars on all images are 250 &#181;m. <a href="https://cloudflare.jove.com/files/ftp_upload/56523/56523fig8large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td></td>
			<td>A1 Cochlea + Cells</td>
			<td>A2 Cochlea + Cells</td>
			<td>B2 Neg. Control Cochlea</td>
		</tr>
		<tr>
			<td>Outside Bony Labyrinth</td>
			<td>3,758</td>
			<td>549</td>
			<td>0</td>
		</tr>
		<tr>
			<td>Bone Outside Cochlea Spaces</td>
			<td>248</td>
			<td>586</td>
			<td>0</td>
		</tr>
		<tr>
			<td>Within Cochlea</td>
			<td>518</td>
			<td>701</td>
			<td>0</td>
		</tr>
		<tr>
			<td>Total Cells</td>
			<td>4,524</td>
			<td>1,836</td>
			<td>0</td>
		</tr>
		<tr>
			<td>Section Volume (&#956;l)</td>
			<td>0.0077</td>
			<td>0.0100</td>
			<td>0.0147</td>
		</tr>
		<tr>
			<td>Percent of Cochlea Volume</td>
			<td>0.46%</td>
			<td>0.59%</td>
			<td>0.87%</td>
		</tr>
		<tr>
			<td>Estimated Cell Density within Cochlea</td>
			<td>113,759</td>
			<td>118,732</td>
			<td>0</td>
		</tr>
	</tbody>
</table>
Table 1: Cell Counts. DAPI stained cochlear sections were thresholded and quantified using automatic counting tools in ImageJ in 3 non-overlapping regions of interest (outside the bony labyrinth, temporal bone outside of the cochlear fluid spaces, and within the cochlea). Cross-sectional cochlear area was also measured in ImageJ, and this value was used to calculate section volume in combination with section thickness (10 &#181;m). Using an estimate of total cochlear volume published by Santi et al.23, the relative cochlear volume of a given section was calculated. The cell density of the entire cochlea was estimated using the relative cochlear volume of a given section in combination with the paired cell count.

Watch this Scientific Journal Video about A Protocol for Decellularizing Mouse Cochleae for Inner Ear Tissue Engineering at JoVE.com

A Protocol for Decellularizing Mouse Cochleae for Inner Ear Tissue Engineering

The goal of this protocol is to demonstrate an effective method to decellularize and decalcify mouse cochleae for utilization as scaffolds for tissue engineering applications.

In mammals, mechanosensory hair cells that facilitate hearing lack the ability to regenerate, which has limited treatments for hearing loss. Current ...

a-protocol-for-decellularizing-mouse-cochleae-for-inner-ear-tissue

Universidad Científica del Sur

Research

JoVE Journal

Bioengineering

10.7K Views.  University of Kansas Medical Center. The goal of this protocol is to demonstrate an effective method to decellularize and decalcify mouse cochleae for utilization as scaffolds for tissue engineering applications.

Video: A Protocol for Decellularizing Mouse Cochleae for Inner Ear Tissue Engineering

Culture of Embryonic Mouse Cochlear Explants and Gene Transfer by Electroporation

Auditory hair cells located within the mouse organ of Corti detect and transmit sound information to the central nervous system. The mechanosensory hair cells are aligned in one row of inner hair cells and three rows of outer hair cells that extend along the basal to apical axis of the cochlea. The explant culture technique described here provides an efficient method to isolate and maintain cochlear explants from the embryonic mouse inner ear. Also, the morphology and molecular characteristics of sensory hair cells and nonsensory supporting cells within the cochlear explant cultures resemble those observed in vivo and can be studied within its intrinsic cellular environment. The cochlear explants can serve as important experimental tools for the identification and characterization of molecular and genetic pathways that are involved in cellular specification and patterning. Although transgenic mouse models provide an effective approach for gene expression studies, a considerable number of mouse mutants die during embryonic development thereby hindering the analysis and interpretation of developmental phenotypes. The organ of Corti from mutant mice that die before birth can be cultured so that their in vitro development and responses to different factors can be analyzed. Additionally, we describe a technique for electroporating embryonic cochlear explants ex vivo which can be used to downregulate or overexpress specific gene(s) and analyze their potential endogenous function and test whether specific gene product is necessary or sufficient in a given context to influence mammalian cochlear development1-8.

We present a method that describes isolation and culture of cochlear explants from embryonic mouse inner ear. We also demonstrate a method for gene transfer into cochlear explants via square-wave electroporation. The in vitro explant culture coupled with gene transfer technique enables researchers to study the effects of altering gene expression during development.

Auditory hair cells located within the mouse organ of Corti detect and transmit sound information to the central nervous system. The mechanosensory hair ...

Developmental 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 ...

Neuroscience

The Mouse Round-window Approach for Ototoxic Agent Delivery: A Rapid and Reliable Technique for Inducing Cochlear Cell Degeneration

Investigators have utilized a wide array of animal models and investigative techniques to study the mammalian auditory system. Much of the basic research involving the cochlea and its associated neural pathways entails exposure of model cochleae to a variety of ototoxic agents. This allows investigators to study the effects of targeted damage to cochlear structures, and in some cases, the self-repair or regeneration of those structures. Various techniques exist for delivery of ototoxic agents to the cochlea. When selecting a particular technique, investigators must consider a number of factors, including the induction of inadvertent systemic toxicity, the amount of cochlear damage produced by the surgical procedure itself, the type of lesion desired, animal survivability, and reproducibility/reliability of results. Currently established techniques include parenteral injection, intra-peritoneal injection, trans-tympanic injection, endolymphatic sac injection, and cochleostomy with perilymphatic perfusion. Each of these methods has been successfully utilized and is well described in the literature; yet, each has various shortcomings. Here, we present a technique for topical application of ototoxic agents directly to the round window niche. This technique is non-invasive to inner ear structures, produces rapid onset of reliably targeted lesions, avoids systemic toxicity, and allows for an intra-animal control (the contra-lateral ear). Results stemming from this approach have helped deeper understanding of auditory pathophysiology, cochlear cell degeneration, and regenerative capacity in response to an acute injury. Future investigations may use this method to conduct interventional studies involving gene therapy and stem cell transplantation to combat hearing loss.

Various methods exist for introducing ototoxic agents to the cochleae of animal models. Presented is a surgical protocol for delivery of ototoxic agents to the round window niche. The procedure is reliable, creates targeted intra-cochlear lesions, and avoids mechanical damage to the microarchitecture. Examination of cochlear self-repair/regeneration is possible.

Investigators have utilized a wide array of animal models and investigative techniques to study the mammalian auditory system. Much of the basic research ...

Medicine

Evaluation of Planar-Cell-Polarity Phenotypes in Ciliopathy Mouse Mutant Cochlea

In recent years, primary cilia have emerged as key regulators in development and disease by influencing numerous signaling pathways. One of the earliest signaling pathways shown to be associated with ciliary function was the non-canonical Wnt signaling pathway, also referred to as planar cell polarity (PCP) signaling. One of the best places in which to study the effects of planar cell polarity (PCP) signaling during vertebrate development is the mammalian cochlea. PCP signaling disruption in the mouse cochlea disrupts cochlear outgrowth, cellular patterning and hair cell orientation, all of which are affected by cilia dysfunction. The goal of this protocol is to describe the analysis of PCP signaling in the developing mammalian cochlea via phenotypic analysis, immunohistochemistry and scanning electron microscopy. Defects in convergence and extension are manifested as a shortening of the cochlear duct and/or changes in cellular patterning, which can be quantified following dissection from developing mouse mutants. Changes in stereociliary bundle orientation and kinocilia length or positioning can be observed and quantitated using either immunofluorescence or scanning electron microscopy (SEM). A deeper insight into the role of ciliary proteins in cellular signaling pathways and other biological phenomena is crucial for our understanding of cellular and developmental biology, as well as for the development of targeted treatment strategies.

Primary cilia influence various signaling pathways. The mammalian cochlea is ideal for examining planar cell polarity (PCP) signaling. Cilia dysfunction affects cochlear outgrowth, cellular patterning and hair cell orientation, readouts of PCP. Our goal is to analyze PCP signaling in mouse cochlea via phenotypic analysis, immunohistochemistry and scanning electron microscopy.

In recent years, primary cilia have emerged as key regulators in development and disease by influencing numerous signaling pathways. One of the earliest ...

Cryosectioning and Immunostaining Mouse Inner Ear Tissue: From Embryonic to Adult Stages

This protocol details general histology methods for preparing inner ear samples from embryonic, neonatal, and adult mice. The purpose of this protocol is to provide a straightforward and standardized method for inner ear tissue processing for researchers, possibly new to the field, interested in working with the mouse cochlea. Included here are protocols for dissection, fixation, embedding, cryosectioning, and immunostaining. Section immunostaining is one of the best methods for examining individual cells within the context of the entire cochlea.
Key steps include dissecting the inner ear away from the skull, fixing the tissue using 4% paraformaldehyde, embedding with Optimal Cutting Temperature compound, cryosectioning, and immunostaining with antibodies targeting specific proteins expressed by the cochlea.
Included in our methods are special considerations made for the cochlea given its 
shape and structure.&#160;Reasonably good focus and dissection skills are needed, as tissue damage can affect the quality of immunostaining and consistency among samples. However, with the procedures we outline, most users can be trained in a few weeks to make beautiful preparations. Overall, this protocol offers a valuable way to promote research to understand auditory development, function, and disease in mouse models.

This protocol, intended for novice users of mouse inner ear tissue, comprehensively details steps for processing mouse inner ears at different developmental stages for section immunostaining.

This protocol details general histology methods for preparing inner ear samples from embryonic, neonatal, and adult mice. The purpose of this protocol is ...

Dextran Labeling and Uptake in Live and Functional Murine Cochlear Hair Cells

The hair cell mechanotransduction (MET) channel plays an important role in hearing. However, the molecular identity and structural information of MET remain unknown. Electrophysiological studies of hair cells revealed that the MET channel has a large conductance and is permeable to relatively large fluorescent cationic molecules, including some styryl dyes and Texas Red-labeled aminoglycoside antibiotics. In this protocol, we describe a method to visualize and evaluate the uptake of fluorescent dextrans in hair cells of the organ of Corti explants that can be used to assay for functional MET channels. We found that 3 kDa Texas Red-labeled dextran specifically labels functional auditory hair cells after 1&#8211;2 h incubation. In particular, 3 kDa dextran labels the two shorter stereocilia rows and accumulates in the cell body in a diffuse pattern when functional MET channels are present. An additional vesicle-like pattern of labeling was observed in the cell body of hair cells and surrounding supporting cells. Our data suggest that 3 kDa Texas-Red dextran can be used to visualize and study two pathways for cellular dye uptake; a hair cell-specific entry route through functional MET channels and endocytosis, a pattern also available to larger dextran.

Here, we present a method for visualizing the uptake of 3 kDa Texas Red-labeled dextran in auditory hair cells with functional mechanotransduction channels. In addition, dextrans of 3&#8211;10 kDa can be used to study endocytosis in hair and supporting cells of the organ of Corti.

The hair cell mechanotransduction (MET) channel plays an important role in hearing. However, the molecular identity and structural information of MET ...

Cochlear Surface Preparation in the Adult Mouse

Auditory processing in the cochlea depends on the integrity of the mechanosensory hair cells. Over a lifetime, hearing loss can be acquired from numerous etiologies such as exposure to excessive noise, the use of ototoxic medications, bacterial or viral ear infections, head injuries, and the aging process. Loss of sensory hair cells is a common pathological feature of the varieties of acquired hearing loss. Additionally, the inner hair cell synapse can be damaged by mild insults. Therefore, surface preparations of cochlear epithelia, in combination with immunolabeling techniques and confocal imagery, are a very useful tool for the investigation of cochlear pathologies, including losses of ribbon synapses and sensory hair cells, changes in protein levels in hair cells and supporting cells, hair cell regeneration, and determination of report gene expression (i.e., GFP) for verification of successful transduction and identification of transduced cell types. The cochlea, a bony spiral-shaped structure in the inner ear, holds the auditory sensory end organ, the organ of Corti (OC). Sensory hair cells and surrounding supporting cells in the OC are contained in the cochlear duct and rest on the basilar membrane, organized in a tonotopic fashion with high-frequency detection occurring in the base and low-frequency in the apex. With the availability of molecular and genetic information and the ability to manipulate genes by knockout and knock-in techniques, mice have been widely used in biological research, including in hearing science. However, the adult mouse cochlea is miniscule, and the cochlear epithelium is encapsulated in a bony labyrinth, making microdissection difficult. Although dissection techniques have been developed and used in many laboratories, this modified microdissection method using cell and tissue adhesive is easier and more convenient. It can be used in all types of adult mouse cochleae following decalcification.

This article presents a modified cochlear surface preparation method that requires decalcification and use of a cell and tissue adhesive to adhere the pieces of cochlear epithelia to 10 mm round cover slips for immunohistochemistry in adult mouse cochleae.

Auditory processing in the cochlea depends on the integrity of the mechanosensory hair cells. Over a lifetime, hearing loss can be acquired from numerous ...

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 ...

A Comparative Study of Drug Delivery Methods Targeted to the Mouse Inner Ear: Bullostomy Versus Transtympanic Injection

We present two minimally invasive microsurgical techniques in rodents for specific drug delivery into the middle ear so that it may reach the inner ear. The first procedure consists of perforation of the tympanic bulla, termed bullostomy; the second one is a transtympanic injection. Both emulate human clinical intratympanic procedures.
Chitosan-glycerophosphate (CGP) and Ringer&#180;s Lactate buffer (RL) were used as biocompatible vehicles for local drug delivery. CGP is a nontoxic biodegradable polymer widely used in pharmaceutical applications. It is a viscous liquid at RT&#160;but it congeals to a semi solid phase at body temperature. RL is an isotonic solution used for intravenous administrations in humans. A small volume of this vehicle is precisely placed on the Round Window (RW) niche by means of a bullostomy. A transtympanic injection fills the middle ear and allows less control but broader access to the inner ear.
The safety profiles of both techniques were studied and compared by using functional and morphological tests. Hearing was evaluated by registering the Auditory Brainstem Response (ABR) before and several times after microsurgery. The cytoarchitecture and preservation level of cochlear structures were studied by conventional histological techniques in paraformaldehyde-fixed and decalcified cochlear samples. In parallel, unfixed cochlear samples were taken and immediately frozen to analyze gene expression profiles of inflammatory markers by quantitative Reverse Transcriptase Polymerase Chain Reaction (qRT-PCR).
Both procedures are suitable as drug delivery methods into the mouse middle ear, although transtympanic injection proved to be less invasive compared to bullostomy.

A Comparative Study of Drug Delivery Methods Targeted to the Mouse Inner Ear: Bullostomy Versus Transtympanic Injection

Two microsurgery approaches for local drug delivery to the inner ear are described here and compared in terms of impact on hearing parameters, cochlear cytoarchitecture and expression of inflammatory markers.

We present two minimally invasive microsurgical techniques in rodents for specific drug delivery into the middle ear so that it may reach the inner ear. ...

Biology

Extracting the Cochlea from a Human Temporal Bone: A Cadaveric Protocol

The extraction of the cochlea from a cadaver human temporal bone may be required for different studies of the inner ear. For histological evaluations, the inner ear must be extracted from the temporal bone to facilitate histologic processing; likewise, some micro-computed tomography devices are too small to accommodate the complete temporal bones; additionally, the image quality can be enhanced when the cochlea is isolated.
The inner ear is located within the petrous part of the temporal bone. The inner ear can be divided into the osseous labyrinth or otic capsule and the membranous labyrinth inside the otic capsule. Furthermore, the inner ear can be divided into the vestibular system (the semicircular canals and the vestibule) and the cochlea. The appreciation of the location and orientation of the cochlea within the temporal bone is difficult, as it is embedded within bony structures and thus cannot be directly visualized. Nevertheless, there are distinct anatomical structures that can help guide the process to allow a reliable drill-out of the cochlea. The landmarks in the posterior parts of the cochlea are the facial nerve, semicircular canals, and the vestibule. In the middle, the inferior borders of the cochlea are identified by the round window and the basal turn of the cochlea. In the anterior border, one encounters the carotid artery; the landmark for the superior border is the genicular ganglion (GG) of the facial nerve. The medial structures are determined by the locations of the internal auditory canal, the superior semicircular canal, and the canal of the internal carotid artery.
In this article, we present a method for extracting the cochlea reliably out of the temporal bone by drill-out while following several anatomical landmarks.

This article presents a reliable method for extracting the human cochlea from the cadaver temporal bone by drill-out while following distinct anatomical landmarks.

The extraction of the cochlea from a cadaver human temporal bone may be required for different studies of the inner ear. For histological evaluations, the ...

A Protocol for Decellularizing Mouse Cochleae for Inner Ear Tissue Engineering

Summary

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Cochlear Surface Preparation in the Adult Mouse

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A Comparative Study of Drug Delivery Methods Targeted to the Mouse Inner Ear: Bullostomy Versus Transtympanic Injection

Extracting the Cochlea from a Human Temporal Bone: A Cadaveric Protocol