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

All aspects of animal research were conducted in accordance with the guidelines of the appropriate Institutional Animal Care and Use Committee. All vertebrate animal experimental procedures described here were approved under the guidelines of the Medical University of South Carolina’s (MUSC) Institutional Animal Care and Use Committee (IACUC).

1. Model Selection

1. Maintain the animal model in a low noise vivarium with routine caretaking per institutional protocols until ready for use. In animal research facilities (ARFs), maintain vibration stability, noise damping, diurnal lighting, prep space, surface finishes, sealing and caulking, and ventilation that meet NIH standards.
   Note: National Institutes of Health (NIH) guidelines for ARFs and maintenance of a low noise vivarium may be reviewed at: http://www.orf.od.nih.gov/PoliciesAndGuidelines/BiomedicalandAnimalResearchFacilitiesDesignPoliciesandGuidelines/
2. For all experiments, use the right ear as the experimental ear. The left ear serves as an intra-animal control and is not surgically altered.
3. Inspect the model animal pre-operatively for signs of middle or outer ear infection. Potential signs may include drainage of fluid or pus from the ear, inflamed pinna tissue, and/or lethargy of the animal. This is unusual, but if noted, avoid further surgery and treat the animal appropriately.

2. Preoperative Procedures

1. Anesthetize the animal 30 min prior to surgery and any perioperative procedures via intra-peritoneal (IP) injection of ketamine (100 mg/kg IP) and xylazine (20 mg/kg IP). Supplement anesthesia as necessary, as judged by a positive toe-pinch reflex, with a lower dosage of ketamine (25 mg/kg IP) and xylazine (5 mg/kg IP). Determine dosing in compliance with institutionally allowed protocols for mice with age appropriate adjustments in dose.
2. Check for full sedation of the model. Check for a stage 3 plane of anesthesia for the entirety of the described protocol marked by a regular breathing rate, a lack of righting reflex (in mice), and lack of pedal and palpebral (toe-pinch) reflexes. Maintain the animal at this level of anesthesia. This is paramount for the minimization of pain and intraoperative movement, bleeding, and leakage of interstitial fluids during surgery.
3. Maintain the animal’s body temperature at 37.5 °C with a closed-loop heating pad.
4. Administer analgesia via subcutaneous injection of buprenorphine. Deliver buprenorphine (0.1 mg/kg SQ once 30 min before surgery) as analgesia to minimize any surgical discomfort. Base dosing and alternatives suitable to the selected model on institutionally approved protocols. Antibiotics use is not necessary if good aseptic technique has been practiced. The animals receive post-operative analgesia if there are signs of pain and distress.
5. Perform physiologic tests preoperatively. Performing auditory brainstem response (ABR) testing and/or distortion product optoacoustic emissions testing (DPOAE) both pre-operatively and just prior to sacrifice of the animal serves as an objective measurement for the effect of the chosen ototoxic agent on the mouse’s hearing.

3. Surgical Preparation and Positioning

1. Sterilize all instruments pre-operatively per institutional standards. Prepare surgical instruments and field in a consistent, sterile, and organized manner to avoid unneeded effort and movement during the procedure. Typical instruments required includes sharp dissecting scissors, several pairs of metallic forceps with jeweler’s tips, an otologic pick, an otologic curette, and a bipolar electrocautery unit (Figure 1).
2. Prepare and sterilize paper wicks made from labwipes in advance by cutting a small triangular piece of the wipe (~ 0.7 in x 1.25 in x 1.75 in) and twisting it tightly between the thumb and forefinger of one hand (~ 1.25 in). Creation of a tightly twisted, extremely thin wick is paramount to success of the procedure. Prepare a total of 15-20 wicks prior to surgery (Figure 2).
3. Use a dental drill to quickly perforate the bone of the tympanic bulla in a controlled fashion. Use of a belt-driven dental hand drill with a 1 or 2 mm tapered tip is preferable. An operative microscope is necessary to complete the protocol. (see Step 4.7)
4. Pre-draw 0.2 ml of an aqueous solution containing the selected ototoxic agent into a 1 ml tuberculin syringe with a 28 G, 1/2' needle. Typically 1 drop (~10 µl) of the agent is enough to completely fill the mouse RWN. A metallic, blunt tip syringe needle eases the delivery of the agent while preventing damage to underlying structures by a sharp bevel tip. Expel any air within the syringe as bubbles can inadvertently fill the RWN and/or prevent proper application of the agent to the round window itself.
5. Remove fur from the post-auricular skin using electrical grooming clippers. Remove fur in a zone extending from the auriculo-cephalic crease rostrally to the shoulder girdle caudally. Extend hair removal from the dorsal, sagittal midline to the mandibular angle laterally. Proper fur removal is paramount to maintaining a clean and clear surgical field. Gently brush clippings from the intended surgical site.
6. Sterilize skin of prepared area per institutional protocols. Apply betadine (povidone/ iodine) alternated with ethanol topically in a circular manner for 2 min. Other agents can be substituted per institutional guidelines.
7. Place the animal on a flat surface to maintain constant body positioning during the procedure. Use a small heating pad placed under the body onto the platform to control body temperature during the procedure to maintain the animal’s body temperature at 36-38 °C. Do not overheat the animal as this may lead to mild to severe skin damage and/or early euthanasia.
   1. Secure the head of the animal via a bite block/head holder apparatus. Utilize a 0.5 cm by 2 cm bite block machined from brass with 2-4 mm diameter holes drilled through it at 5 mm intervals along the long axis. Open the animal’s mouth around this fixture and fit the upper central incisors into one of the holes depending on the animal’s size. Gently tighten a small clamp over the dorsum of the animal’s snout to hold it in place (Figure 3).
   2. Confirm that the bite block/head holder is rigidly connected to the center of a U-shaped articulating arm (Figure 3). Secure a 1cm wide rod to the left arm of the “U” and use rod as a left-side head rest (the right side is always the operative side).
   3. Once firmly secured in the head holder, rotate the mouse to a left lateral decubitus position. Position the body carefully on the flat operating surface to ensure it will be stable throughout the procedure and avoid undue torsional stress on the cervical vertebrae.
8. Position an operating microscope capable of 4x, 10x, and 20x magnification over the surgical field. Confirm that the microscope can maintain its position in a hands-free manner as this is ideal for the two-handed surgical protocol discussed below.
9. Place a bipolar-enabled cautery unit with fine tipped jeweler’s hand piece in a position that is immediately available for the assistance in cauterization of small vessels and dissection of tissue. This may also be necessary should heavier bleeding be encountered.

4. Surgical Approach

1. Under microscopic magnification, use sharp scissors or a scalpel blade to create a 1-1.5 cm postauricular incision, approximately 6-8 mm caudal to the auriculocephalic crease. In the adult mouse, an incision from the dorsal midline laterally to a point near the angle of mandible is adequate. Carefully avoid cutting deeply to preserve underlying vascular structures.
2. Conduct careful blunt dissection through the subcutaneous fat layer which can be of variable thickness. Fat may be safely removed if needed to improved exposure. Take caution when dissecting in a ventral-medial direction as the external jugular vein traverses this area and damage to this structure may cause large amounts of blood loss and flooding of the surgical field. Control any excessive bleeding with absorbable gelatin sponges and/or cotton pellets. Use bipolar cautery for heavier bleeding.
3. Once the fat layer is properly divided, expose the cervical musculature. Note important structures including the large muscle body of the cleidomastoideus centrally within the exposed surgical field, the external jugular vein ventrally, and parotid tissue rostrally overlying the angle of mandible. An important landmark is a small nerve branch (of cranial nerve XI) that wraps around the posterior/dorsal edge of the cleidomastoideus to extend rostrally toward the pinna (Figure 4).
4. Gently retract the cleidomastoideus muscle body in a posterior/dorsal direction (Figures 4, 5). Gently divide the transparent fascia enveloping the muscle body. In a similar manner, gently retract the parotid and external jugular vein in the opposite (anterior/ventral) direction (Figure 5).
5. With good retraction of the cleidomastoideus muscle body, the shiny dome of the tympanic bulla periosteum will come into view (Figure 6). At the caudal aspect of the bulla, the insertion of a deeper cervical muscle, the sternomastoideus, will come into view (Figure 6). Preserve the facial nerve, which becomes visible at the dorsal and rostral aspect of the bulla dome. Place a self-retaining retractor (Sterile titanium shaft– embedded in disposable silicone paste) prior to drilling.
6. With two-handed technique, gently divide the bulla periosteum with bipolar diathermy to expose the underlying bone. Using forceps or an otologic curette, gently elevate and push the periosteum in a peripheral direction to widely expose the bulla dome.
   Note: Step 4.6 is critical to maximize surgical view of the middle ear space. Utilize meticulous and gentle soft tissue handling to avoid bleeding and/or leakage of interstitial fluid into the bulla cavity after drilling.
7. Upon a properly exposed bulla dome, drill a 2 mm pilot hole through the bulla bone with a dental surgical drill between the caudal margin of the dome and the visibly opaque line (representing the tympanic membrane) extending across the rostral aspect of the bulla (Figure 7). Take care to drill only through bone to preserve underlying structures, such as the stapedial artery. Drill a second pilot hole nearby to facilitate un-roofing of the bulla bone (Figure 7).
8. Using a pair of jeweler’s tip forceps, uncap the bulla bone in a dorsal and caudal direction (Figure 8). Remove the bone in a piecemeal fashion under high magnification. Do not puncture the stapedial artery, which lies directly beneath the bulla cap, as bleeding from this artery may compromise the procedure. Minimize the amount of bone removed to prevent excessive fluid entry to the middle ear while still allowing excellent visualization and access to the round window niche (Figure 8).

5. Round Window Application of Ototoxic Agent

1. Make subtle rotational adjustments to the head holder to bring the RWN squarely into view. The RWN is typically located at the dorsal and caudal aspect of the middle ear space and appears as a cup-like indention of the otic capsule bone. In most cases, the stapedial artery runs 1-2 mm ventral/rostral to this. The RWN can sometimes be tucked peripherally under an acute angulation of the bulla dome. In such cases, careful positioning of the animal’s head is paramount.
   1. Using the paper wicks prepared preoperatively, remove all visible fluid in the middle ear and RWN until dry bone is visualized.
      Note: This is the most critical step of the entire protocol, as the ~10 µl (1 drop) of ototoxic agent applied to the RWN can easily be diluted by this fluid.
2. Under maximum magnification, use a fine caliber needle on a 1 ml tuberculin syringe to apply one drop (~10 µl) of ototoxic agent directly to the RWN, filling it completely. Take care to not disrupt the stapedial artery and observe closely for the replacement of a small light reflection at the base of a dry niche with a dull and hazy fluid meniscus as an indicator that the niche is filling properly.
   1. Allow the agent to rest within the RWN for approximately 10 min exposure time. After this, completely wick out the agent and replace it with a new application of the same agent. Determine repetitions of application according to agent specifications. Total exposure time in this procedure typically ranges between 30 to 60 min.
   2. At the end of the procedure, completely wick away the agent one final time and apply fresh agent to the RWN. Leave the bulla uncapped and use forceps to close the soft tissue over the surgical site.
3. Seal surgical site at the level of the skin with a 4-0, non-absorbable monofilament suture. Place the animal in a recovery cage/station. Monitor the animal’s clinical condition regularly following the operation. Maintain the animal in appropriate environment conditions, including housing with soft bedding and supplementation with soft food, to minimize stress. Determine future procedures and post-operative conditions according to the institutional IACUC protocols.
4. Use institutional IACUC protocols to sterilize instruments before use on the next animal. Allow for proper cooling time between instrument use.

6. Postoperative Procedures and Cochlear Tissue Harvesting

1. As described in step 2.5, perform physiologic testing of hearing postoperatively at desired points and prior to sacrifice. Perform Postoperative procedures according to experimental aims. Sacrifice the animal on any desired postoperative day.
2. Upon full anesthesia via IP injection, as required per institutional IACUC protocols, sacrifice the animal. Using sharp scissors, decapitate animal just caudal to the occiput. Sharply open the cranial vault with scissors along the dorsal and ventral midline and spread widely. Gently scoop out brain tissue to expose the temporal bones.
   Note: A number of procedures for investigation of post exposure changes in the cochlea can branch from this point and will be mentioned in the discussion section. Determine the investigative method most relevant to the experimental aims. If electron microscopy is planned after the cochlea are sectioned, use cardiac catheterization to pre-fix the tissue. This is beyond the scope of this discussion and has been covered in depth elsewhere.¹³
3. Cut the temporal bones out of the skull base with the scissors and immediately placed in fixative solution. Immerse bones directly in 4% paraformaldehyde for 1.5 hr at RT. Monitor fixation time, as longer duration can limit the outcome of histological analysis at later steps. Decalcify dissected bones for a variable period of time via submersion in 1 mM Ethylenediaminetetraacetic acid (EDTA).