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

Ensure that the animal handling procedures are in accordance with international and national regulations. The protocol follows the European Community 2010/63/EU and Spanish RD 53/2013 guidelines, respectively.

1. General Animal Handling

1. Feed mice ad libitum with a standard diet and drinking water. Control health and well-being by following Federation for Laboratory Animal Science Associations (FELASA) recommendations.

2. Hearing Assessment

NOTE: Track functional impact of microsurgical procedures by testing hearing before and many times after surgery (in this work 2, 7, 14 and 28 d postmicrosurgery) with non-invasive procedures such as ABR⁹.

1. For ABR testing, anesthetize mice with short duration effect protocols i.e. intraperitoneal injection of ketamine (100 mg/kg body weight (BW) and xylazine (10 mg/kg, BW). Alternatively, perform hearing tests under inhalant anaesthesia.
   NOTE: Since ABR parameters can be influenced by the anaesthetic protocol²⁸, use the same one throughout the experiment.
2. Check the depth of anaesthesia by testing the toe-pinch reflex.
   NOTE: When the withdrawal reflex disappears, the animal has reached an appropriate depth of anaesthesia to perform auditory testing.
3. Protect the eyes from desiccation and secondary keratoconjunctivitis sicca by topical administration of tear supplements, such as hydroxypropyl methylcellulose-based gels.
4. Keep the mouse at physiological temperature (37.5-38 °C) during the entire procedure. To avoid electrical interference, use a warm water pump and heating pads. Monitor body temperature with rectal probes. Always take care not to overheat the animal.
   NOTE: We recommend cleaning the heating pad with a surface disinfectant between mice.  For anaesthesia induction and recovery, electric heating pads, incandescent or infrared lights can be used.
5. ABR procedure
   NOTE: For ABR registration, use a computer workstation with an inboard sound card to create waveforms (Digital to analog, D-A output, conversion) and to digitize electric response waveforms (analog to digital, A-D input), an attenuator, an oscilloscope and a low-impedance amplifier. Modern auditory workstations (i.e. Tucker Davis Techonologies) include all these components in a single compact system. 
   1. Place the anesthetized mouse in prone position on the heating pads within a sound-attenuating chamber to avoid ambient noise interference and reverberation (Figure 1).
   2. Deliver acoustic stimuli into the external auditory canal. Use preset stimuli or new signals designed with the appropriate software. Connect the D-A workstation output to the selected speaker.
      NOTE: Free-field or closed-field speakers inserted in the external auditory canal could be used. The former are preferred when working with mice because of the difficulty in probe insertion and sound calibration in closed systems. Free field speakers stimulate both ears and elicit a binaural response. To obtain predominantly monaural responses with free-field speakers, contralateral activity has to be eliminated by occlusion (i.e. with ear plugs) or by masking the noise.
      1. Place free-field speaker at a fixed distance (usually 5-20 cm) facing the head or selected ear with the center of the speaker aligned with the external auditory canal. Ensure that no obstacles are between the speaker and the ear and that the pinna is completely opened.
   3. Place stainless steel subdermal needle electrodes as follows: i) the active (positive) electrode in the scalp between the ears (over the vertex of the skull), ii) the reference (negative) electrode, in the parotid region below the pinna, and iii) the ground electrode in the back, tail or hind limb region (Figure 1).
      1. Check the electrical impedance in the positive and negative electrodes. Ensure that the impedance is less than 3 kOhm (ideally 1 kOhm). If it is higher, reposition, clean with alcohol or replace the electrodes.
   4. For ABR recording, generate broadband clicks and pure tone frequencies and present at decreasing intensities from 90 to 10 dB relative to the sound pressure level (SPL) in 5-10 dB SPL steps^27,29,30.
      1. Present brief click or tone burst stimuli (1-5 ms) beginning with high level (i.e. 80 or 90 dB SPL) and reducing the intensity in 5-10 dB SPL steps. Register the electrical response in the first 10 ms after stimulation (evoked ABR responses appear at 6-8 ms).
         NOTE: For this reason, stimulation rates should not be higher than 50 stimuli/s (normal rate 20-50).
   5. Amplify, record and average the evoked electrical response to each stimulus and intensity. Use an amplifier with low noise and a good signal-to-noise ratio, and connect it to the A-D input.
      NOTE: ABRs have very low amplitudes, typically below 1 µV (peak-to-peak) and must be recorded using an amplifier with very low noise. In normal hearing mice, clear ABR waves emerge after averaging 100 – 200 responses, however to obtain high quality recordings, or in the case of hearing impairment, more repetitions are recommended (750-1,000)²⁷.
   6. Visually determine the ABR threshold during the test.
      NOTE: The ABR threshold is the lowest sound stimuli intensity that elicits a reliable ABR recording with waves I to IV clearly visible and medium peak-to-peak voltage 2 SD above the mean background activity³¹. This data has to be confirmed during off-line analysis, along with other parameters including peak and interpeak latency, and wave amplitudes.
   7. Perform data analysis either manually or automatically.
      1. For manual analysis, identify the 4-5 ABR waves (I, II, III…etc.) and mark out the peaks (P1, P2, P3…) and valleys (N1, N2, N3…) for each wave. Once the analysis is completed, export data to spreadsheet or text file.
         NOTE: Specific software for electrical response recording usually performs the analysis automatically. Additional measurements can be determined in the ABR recording in response to a fixed intensity (i.e. 70 or 80 dB SPL) or at intensities relative to individual click thresholds (i.e. 15 dB SPL over threshold).
6. Perform statistical analysis of ABR data using the appropriate software. Depending on the experimental design, use standard paired T-test or analysis of variance (ANOVA) to compare main ABR parameters in the different groups^26,30.
   NOTE: In longitudinal studies, many functional data are collected from the same animal at different temporal points (i.e. before and after microsurgery). In this case, a general linear model repeated measure test provides a detailed variance analysis.

3. Vehicle Preparation

1. Prepare and use vehicle solutions under sterile conditions.
   NOTE: Liquid solutions are usually cleared quickly through the Eustachian tube. Different injectable delivery systems could be used to increase the residence time of the drug in the middle ear, including hydrogels and nanoparticles³².
   1. To prepare CGP-hydrogel, dissolve 75% deacetylated chitosan in 0.2 M acetic acid yielding a 1.5-2% (wt/wt) chitosan solution. Add 9% glycerophosphate (wt/wt) to this solution⁷. Prepare the solution just before administration and store the hydrogel at 4 °C until use.
      NOTE: The CGP-hydrogel is moderately viscous but still injectable at this temperature. Below 4 °C it changes to a solid phase, blocking its application. After application, CGP undergoes a phase transition to a semi-solid gel in about 15 min at 37 °C.
   2. Aliquot (0.5 mL) commercial RL buffer and store at 4 °C until use.

4. Microsurgical Procedures

1. Induce general anaesthesia with ketamine based combinations of sedatives and analgesics (i.e. ketamine 100 mg/Kg, medetomidine 0.05 mg/Kg and phentanile 0.025 mg/Kg) by intraperitoneal injection, followed by inhalant agents (i.e. isoflurane).
   1. After administering injectable agents, adjust the anaesthetic facemask to the mouse snout and connect the O₂ supply to isoflurane vapor. Maintain the inhalation anaesthesia during the microsurgery and monitor the anesthetic plane with the toe-pinch reflex and breathing pattern. Begin surgical preparation when the reflex is totally abolished and the mouse presents regular breathing.
   2. Maintain body temperature with heating pads during the entire procedure and protect the eyes from corneal keratitis with a hydroxypropyl methylcellulose-based gel.
2. Prepare a clean surgical area by using sterile drapes. Sterilize the microsurgical instruments with a glass-bead sterilizer prior to surgery. Maintain sterile conditions during the whole surgical procedure (sterile gloves, drapes, surgical instruments, etc.).
3. Microsurgery
   1. Bullostomy
      NOTE: Bullostomy is a unilateral procedure. Operate one ear of the mouse and use the contralateral ear as the control.
      1. Place the mouse in a decubitus supine position. Prepare the surgical area at the ventral surface of the neck using clippers to remove the fur. Clean the skin with povidone iodine based antiseptic solution, and cover it with sterile drapes.
      2. Using a scalpel, make a 2 cm longitudinal incision from the mandible to the clavicle.
      3. Under magnification with a surgical microscope, identify the submandibular glands and separate both with forceps. Retract the submandibular glands and localize the origin of the digastric muscle and the facial nerve.
      4. Make an incision in the origin of the digastric muscle with a scissors, and retract it ventrally, exposing the underlying inferior-medial aspect of the tympanic bulla.
      5. Make an opening in the bulla by drilling into it with a 27 G needle (Figure 2A). Localize the stapedial artery and the RW membrane caudal to it (Figure 2B). Clean the blood from the drilled area with an absorbable gelatin sponge.
      6. Using a 34 G catheter and a glass micro syringe, slowly inject 3-5 µL of vehicle solution (CGP-hydrogel or RL) through the bullostomy directly onto the RW niche, filling it (Figure 2C). Seal the bullostomy with 1-2 drops of tissue adhesive.
      7. Return the submandibular glands to their initial position and close the skin incisions with 5-0 silk surgical suture. Apply a chlorhexidine-based antiseptic around the incision to avoid wound infection. NOTE: Absorbable and non-absorbable sutures could be used. Non-absorbable sutures must be removed  in 2 weeks. Silk is not recommended for skin closure since its use is associated with incision infection and local tissue reactions.
   2. Bilateral transtympanic injection
      1. Place the mouse in the lateral decubitus position and prepare an aseptic surgical area below the external auditory meatus as described in 4.3.1.1.
      2. Make a 0.5 cm longitudinal incision in the vertical part of the outer ear canal close to the tragus and section the internal cutaneous fold of the pinna (optional).
      3. Locate the tympanic membrane at the end of the outer ear canal using a surgical microscope (Figure 2E) and identify the upper pars flaccida and the inferior pars tensa, which is divided into anterior and posterior sections by the handle of the malleus (Figure 2F).
      4. Make a small myringostomy in the caudal portion of the pars flaccida. Make an additional incision in the pars tensa of the tympanic membrane, to allow air evacuation during injection³³. Gently inject 10-15 µL of vehicle (CGP-hydrogel or RL) solution with a glass micro syringe connected to a 34 G catheter through the pars flaccida, close to the RW niche until the middle ear is clearly full.
      5. Close the skin incisions with a 5-0 silk surgical suture and clean as described in 4.3.1.7.
      6. Place the mouse on its other side and operate the contralateral ear (steps 4.3.2.1-to 4.3.2.5).
4. Keep the mouse on a heating pad until it has regained sufficient consciousness to maintain sternal recumbency. Do not return an animal that has undergone surgery to the company of other animals until fully recovered.
5. Monitor the body condition, activity and the presence of signs of pain or stress. Provide analgesics if needed (i.e. Buprenorphine 0.05 mg/kg, Carprofen 5-10 mg/kg). Review the surgical wound daily and remove skin closures 7-14 days postoperatively after verifying that the wound has healed.

5. Morphological Evaluation of Cochlear Cytoarchitecture

1. Euthanize the mouse at the end of the experiment (in this work 28 days postmicrosurgery), with an intraperitoneal pentobarbital overdose (100 mg/kg) to study the long term effects of the surgery.
2. Perform a transcardial perfusion with cold 0.1 M Phosphate-buffered Saline (PBS), pH 7.5, followed by 4% (wt/v) paraformaldehyde (PFA) in 0.1 M PBS, pH 7.5 as described²⁶.
   CAUTION: Paraformaldehyde is highly toxic; avoid contact with skin, eyes or mucous membranes. Avoid breathing the powder during measuring and preparation.
3. Use a stereomicroscope to dissect out the inner ear from the temporal bone as described^34,35 without separating the vestibular and cochlear components of the inner ear.
4. Fix the isolated inner ear with 4% (wt/v) PFA in 0.1 M PBS, pH 7.5 at 4 °C for 12 h with gentle shaking. Wash 3x for 5 min with 0.1 M PBS, pH 7.5.
5. Decalcify the samples with 10% ethylenediaminetetraacetic acid (EDTA) prepared in 0.1 M PBS, pH 6.5 at 4 °C for 10 d with constant shaking, changing the EDTA solution every 3 d.
6. When cochleae acquire a soft consistency, remove EDTA and wash 3x for 5 min with 0.1 M PBS, pH 7.5, with shaking at RT.
7. Embed the samples in paraffin wax as described³⁴ and make 7 µm thick cochlear sections parallel to the modiolus.
8. To evaluate cochlear cytoarchitecture, stain sections with haematoxylin and eosin (H&E)³⁰ and use a light microscope connected to a digital camera to capture images with 4X and 20X lenses.

6. Cochlear Gene Expression

1. Clean the working surface and surgical instruments with RNAse decontamination solution.
2. Euthanize the mouse as described in 5.1 and quickly dissect out the inner ear from the temporal bone using a microscope. Immerse the inner ear in a glass dish containing Ribonucleic acid (RNA) protector and stabilizer reagent.
3. Remove the remaining petrous bone with jeweller forceps and gently separate the cochlea from the vestibule using Vanna's eye scissors³⁵.
4. Immediately transfer the cochlea into a 2 mL microcentrifuge tube with 80 µL RNA protector and stabilizer solution and freeze the tissue by placing the tube in dry ice. Preserve cochlear samples at -70 °C until use.
5. Isolate cochlear RNA as described³⁵ and determine its quality and quantity spectrophotometrically.
6. Generate cochlear cDNA from equal amounts of total mouse RNA using a reverse transcription commercial kit.
7. Perform qRT-PCR to amplify complementary deoxyribonucleic acid (cDNA) in triplicate to measure gene transcripts^35,36.
   NOTE: In this work pro- and anti-inflammatory gene transcripts of Il1b, Il6, Tgfb1, Tnfa, Il10 and Dusp1 were measured.
8. Calculate relative expression ratios by normalizing target transcript cycle threshold (Ct) levels to the arithmetic mean of the reference gene level and the relative quantification by normalizing the problem group transcript levels to the arithmetic mean of the calibrator group³⁷.