The authors wish to thank the Genomics and Noninvasive Neurofunctional Evaluation facilities (IIBM, CSIC-UAM) for their technical support. This work was supported by grants of the Spanish &#34;Ministerio de Economia y Competitividad&#34; (FEDER-SAF2014-53979-R) and the European Union (FP7-AFHELO and FP7-PEOPLE-TARGEAR) to IVN.

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
<ol>
	<li>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.</li>
</ol>
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&#160;postmicrosurgery) with non-invasive procedures such as ABR9.
<ol>
	<li>For ABR testing, anesthetize mice with short duration effect protocols i.e.&#160;intraperitoneal injection of ketamine (100 mg/kg body weight (BW) and xylazine (10 mg/kg,&#160;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.</li>
	<li>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.</li>
	<li>Protect the eyes from desiccation and secondary keratoconjunctivitis sicca by topical administration of tear supplements, such as hydroxypropyl methylcellulose-based gels.</li>
	<li>Keep the mouse at physiological temperature (37.5-38 &#176;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. &#160;For anaesthesia induction and recovery, electric heating pads, incandescent or infrared lights can be used.</li>
	<li>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.&#160;Modern auditory workstations (i.e. Tucker Davis Techonologies) include all these components in a single compact system.&#160;
	<ol>
		<li>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).</li>
		<li>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.
		<ol>
			<li>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.</li>
		</ol>
		</li>
		<li>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).
		<ol>
			<li>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.</li>
		</ol>
		</li>
		<li>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 steps27,29,30.
		<ol>
			<li>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).</li>
		</ol>
		</li>
		<li>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 &#181;V (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.</li>
		<li>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.</li>
		<li>Perform data analysis either manually or automatically.
		<ol>
			<li>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).</li>
		</ol>
		</li>
	</ol>
	</li>
	<li>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 groups26,30. 
	NOTE: In longitudinal studies, many functional data are collected from the same animal at different temporal points (i.e.&#160;before and after microsurgery). In this case, a general linear model repeated measure test provides a detailed variance analysis.</li>
</ol>
3. Vehicle Preparation
<ol>
	<li>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.
	<ol>
		<li>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 &#176;C until use. 
		NOTE: The CGP-hydrogel is moderately viscous but still injectable at this temperature. Below 4 &#176;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 &#176;C.</li>
		<li>Aliquot (0.5 mL) commercial RL buffer and store at 4 &#176;C until use.</li>
	</ol>
	</li>
</ol>
4. Microsurgical Procedures
<ol>
	<li>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).
	<ol>
		<li>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.</li>
		<li>Maintain body temperature with heating pads during the entire procedure and protect the eyes from corneal keratitis with a hydroxypropyl methylcellulose-based gel.</li>
	</ol>
	</li>
	<li>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.).</li>
	<li>Microsurgery
	<ol>
		<li>Bullostomy 
		NOTE: Bullostomy is a unilateral procedure. Operate one ear of the mouse and use the contralateral ear as the control.
		<ol>
			<li>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.</li>
			<li>Using a scalpel, make a 2 cm longitudinal incision from the mandible to the clavicle.</li>
			<li>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.</li>
			<li>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.</li>
			<li>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.</li>
			<li>Using a 34 G catheter and a glass micro syringe, slowly inject 3-5 &#181;L&#160;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.</li>
			<li>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.&#160;NOTE: Absorbable and non-absorbable sutures could be used. Non-absorbable sutures must be removed &#160;in 2 weeks. Silk is not recommended for skin closure since its use is associated with incision infection and local tissue reactions.</li>
		</ol>
		</li>
		<li>Bilateral transtympanic injection
		<ol>
			<li>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.</li>
			<li>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).</li>
			<li>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).</li>
			<li>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 &#181;L&#160;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.</li>
			<li>Close the skin incisions with a 5-0 silk surgical suture and clean as described in 4.3.1.7.</li>
			<li>Place the mouse on its other side and operate the contralateral ear (steps 4.3.2.1-to 4.3.2.5).</li>
		</ol>
		</li>
	</ol>
	</li>
	<li>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.</li>
	<li>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.</li>
</ol>
5. Morphological Evaluation of Cochlear Cytoarchitecture
<ol>
	<li>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.</li>
	<li>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.</li>
	<li>Use a stereomicroscope to dissect out the inner ear from the temporal bone as described34,35 without separating the vestibular and cochlear components of the inner ear.</li>
	<li>Fix the isolated inner ear with 4% (wt/v) PFA in 0.1 M PBS, pH 7.5 at 4 &#176;C for 12 h with gentle shaking. Wash 3x for 5 min with 0.1 M PBS, pH 7.5.</li>
	<li>Decalcify the samples with 10% ethylenediaminetetraacetic acid (EDTA) prepared in 0.1 M PBS, pH 6.5 at 4 &#176;C for 10 d&#160;with constant shaking, changing the EDTA solution every 3 d.</li>
	<li>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.</li>
	<li>Embed the samples in paraffin wax as described34 and make 7 &#181;m thick cochlear sections parallel to the modiolus.</li>
	<li>To evaluate cochlear cytoarchitecture, stain sections with haematoxylin and eosin (H&#38;E)30 and use a light microscope connected to a digital camera to capture images with 4X and 20X lenses.</li>
</ol>
6. Cochlear Gene Expression
<ol>
	<li>Clean the working surface and surgical instruments with RNAse decontamination solution.</li>
	<li>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.</li>
	<li>Remove the remaining petrous bone with jeweller forceps and gently separate the cochlea from the vestibule using Vanna&#39;s eye scissors35.</li>
	<li>Immediately transfer the cochlea into a 2 mL&#160;microcentrifuge tube with 80 &#181;L&#160;RNA protector and stabilizer solution and freeze the tissue by placing the tube in dry ice. Preserve cochlear samples at -70 &#176;C until use.</li>
	<li>Isolate cochlear RNA as described35 and determine its quality and quantity spectrophotometrically.</li>
	<li>Generate cochlear cDNA from equal amounts of total mouse RNA using a reverse transcription commercial kit.</li>
	<li>Perform qRT-PCR to amplify complementary deoxyribonucleic acid (cDNA) in triplicate to measure gene transcripts35,36. 
	NOTE: In this work pro- and anti-inflammatory gene transcripts of Il1b, Il6, Tgfb1, Tnfa, Il10 and Dusp1 were measured.</li>
	<li>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.</li>
</ol>

The authors declare that they have no competing financial interests.

Local drug delivery to the inner ear can be done directly by intracochlear injection or indirectly by intratympanic administration, placing the drug in the middle ear4,19,39. Intracochlear administration provides controlled and precise drug delivery to the cochlea, avoiding diffusion through window membranes, basal-to-apical concentration gradients and clearance through the Eustachian tube. However, it is usually a highly invasive procedure that requires a complex and delicate microsurgery7,39. In this context, the industry is developing new, coated, implantable devices for sustained drug release40,41. On the other hand, intratympanic administration is a minimally invasive and easy to perform procedure that allows the injection of larger volumes of the drug into the middle ear, although the pharmacokinetics is not easy to control. The majority of the drug is cleared through Eustachian tube and the remaining fraction has to diffuse through the RW membrane to reach the cochlea18. RW is the site of maximum absorption of substances from the middle ear into the perilymph-filled tympanic duct of the cochlea7. It is a semipermeable three layer structure, although its permeability depends on the drug characteristics (size, concentration, solubility and electrical charge) and on transmembrane transport systems (diffusion, active transport or phagocytosis)42. The oval window and otic capsules are alternative but less effective entrances to cochlea43,44.
Here we demonstrate and compare two microsurgical methods for targeted drug delivery into the mouse middle ear: bullostomy and transtympanic injection procedures. Common critical steps to these procedures include: i) an evaluation of hearing before and after the microsurgery, ii) preparation of a homogeneous vehicle solution under sterile conditions, iii) careful supervision of the anaesthetic procedure and monitoring of animal body temperature and constants, iv) slow placement of the appropriate volume of vehicle targeting the RW, and iv) taking cochlear samples to complete molecular and morphological analysis.
Retroauricular and ventral approaches for bullostomy have been described7,45. We used the ventral approximation because in our experience it has resulted in&#160;less morbidity&#160;and provided better access to the RW46. Transtympanic injections are usually carried out through the pars tensa of the tympanic membrane, anterior or posterior to the malleus manubrium12. In this work we performed a modification of the technique, an injection through the pars flaccida beyond the malleus with a previous additional puncture of the pars tensa to allow air evacuation during injection.
The transtympanic injection was less invasive than the bullostomy, although both microsurgeries were rapid (20 and 5 min per ear for bullostomy and transtympanic approach respectively), with short postoperative recuperation times and no morbidity. Most importantly, both procedures maintained hearing and the ABR parameters were identical to those determined before the microsurgery. The transtympanic approach takes less time than the bullostomy and can be performed in both ears of the same animal during the same intervention. Advantages of the transtympanic injection are thus that it can be performed bilaterally and repeated, if required. On the other hand, bullostomy provides direct visual access to the RW membrane and allows the filling of the RW niche. In contrast, transtympanic injection does not allow for control of vehicle placement in the RW niche.
The procedures reported in this work describe how to perform a local drug vehicle delivery to the middle ear for pre-clinical applications such as evaluation of ototoxicity and evaluation of efficacy in hearing loss. Two microsurgery procedures are described that provide alternative methods with specific advantages and drawbacks. Both preserve hearing and do not cause morphological alterations. Local inflammation is described as a potential complication of bullostomy. A set of complementary techniques are also described for postsurgical procedures, including hearing, morphological and inflammatory marker expression evaluations. Future applications for these techniques include the preclinical evaluation of new therapies for hearing loss, including genetic, cellular and pharmacological approaches, in animal models. Intratympanic administrations ensure the delivery of the treatment in the middle ear, in contact with the round window membrane, facilitating the passage into the perilymph without evident cochlear damage.

Hearing impairment is the most frequent human sensory deficit and affects 5.3% of the worldwide population, and 30% of individuals over the age of 65 (<a href="http://www.who.int/topics/deafness/en">http://www.who.int/topics/deafness/en</a>, updated 2016). Hearing loss affects language acquisition in children and accelerates cognitive decline in older people. Therefore, it is a significant health-care problem with a tremendous socio-economic impact. It can be caused by genetic defects, environmental factors or a combination of both1, which in the end induce damage and death of hair cells and neurons in the cochlea. These cells do not regenerate in mammals, therefore cellular loss and concomitant hearing loss cannot be reversed. Clinical options are based on prosthetic devices, including hearing aids and cochlear, middle ear and bone conduction implants2. Unfortunately, there are no specific medical restorative treatments for hearing impairment and thus several research lines are focused on the development of preventive and reparative therapies. Novel therapeutic options include gene and cell therapies as well as development of small molecules for pharmacological therapy2.
One of the most important challenges in cochlear pharmacological therapy is drug delivery. Systemic treatments have limited efficacy in the cochlea due to the blood-labyrinth barrier3, continuous endothelium in contact with cochlear blood vessels, which acts as a physical and biochemical barrier to maintain inner ear fluid homeostasis, therefore limiting drug passage to the inner ear. It is permeable only to small liposoluble molecules, although permeability can be increased during cochlear inflammation, and also with the use of diuretics or osmotic agents. The volume of drug that eventually reaches the cochlea after systemic administration is reduced; therefore, high doses that could cause organic toxicity are required. In addition, hepatic metabolism of the drug can produce toxic or inactive metabolites4,5,6,7. In contrast, local interventions allow the placement of a known limited quantity of the drug into the middle or inner ear without undesirable side effects4,7,8,9. In current clinical practice, intratympanic administrations are limited to certain cochlear pathologies, such as gentamicin in Meniere&#39;s disease10, corticosteroids in sudden deafness, Meniere&#39;s disease, immune-mediated and noise induced hearing loss,11,12,13,14,15 and insulin-like growth factor 1 (IGF1) in sudden deafness4,16,17.
Formulations for local administration should preserve the delicate homeostasis (pH and osmolarity) of cochlear fluids. In addition, it is very important to maintain sterility throughout the process to avoid bacterial contamination of the cerebrospinal fluid. The excipient used for drug delivery should be biocompatible, nonototoxic and of the appropriate consistency. Liquid solutions are recommended for intracochlear injections, but are not suitable for the intratympanic route due to the clearance through the Eustachian tube. In this case, drugs are usually carried by semi-solid gels to increase their permanence in the middle ear4,18,19. Alternative delivery systems used as carriers to increase the passage of the drug to the inner ear are nanoparticles20 and adenoviruses21 Here we compared two vehicles: CGP and an RL solution. CGP is a hydrogel formed by chitosan, a linear polysaccharide composed of D-glucosamine and N-acetyl-D-glucosamine obtained from crustacean shells, and &#946;-glycerophosphate, a polyol that forms a shield of water around the chitosan chains and maintains it in liquid form. CGP is thermosensitive and can be degraded by lysozymes, allowing a sustained drug release in the middle ear22,23,24,25. Chitosan-base hydrogels are suitable vehicles for clinical applications such as drug delivery due to their lack of immunogenicity and lack of activation of local inflammatory reactions23,24. On the other hand, RL buffer is a non-pyrogenic isotonic solution (273 mOsm/L and pH 6.5) intended for intravenous administration in humans as a source of water and electrolytes, especially in blood loss, trauma or burn injuries because the byproducts of lactate metabolism in the liver counteract acidosis.
Here we describe and compare two surgical methods that have been refined for local drug delivery to the mouse inner ear. The safety profile of both techniques was evaluated by using functional, morphological and molecular tests. Hearing was evaluated using Auditory Brainstem Response (ABR)26,27 performed before and after microsurgery at different times. End-point procedures were used to dissect the cochlea and compare the anatomical, cellular and molecular impact of these two microsurgical procedures.

<table border="0" cellpadding="0" cellspacing="0" width="1375">
	<tbody>
		<tr height="43">
			<td height="43" style="height:43px;width:455px;">Ketamine (Imalgene)</td>
			<td style="width:251px;">Merial</td>
			<td style="width:207px;"># 2529</td>
			<td style="width:464px;">CAUTION: avoid contact of the drug with skin or eyes or accidental self-inflicted injections</td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:455px;">Xylacine (Xilagesic)&#160;</td>
			<td style="width:251px;">Calier</td>
			<td style="width:207px;"># 6200025225</td>
			<td style="width:464px;"></td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:455px;">Lubricant eye gel (Artific)</td>
			<td style="width:251px;">Angelini</td>
			<td style="width:207px;"># 784710</td>
			<td style="width:464px;"></td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:455px;">Water pump&#160;</td>
			<td style="width:251px;">Gaymar</td>
			<td style="width:207px;"># TP472</td>
			<td style="width:464px;"></td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:455px;">Subdermal needle electrodes&#160;</td>
			<td style="width:251px;">Spes Medica</td>
			<td style="width:207px;"># MN4013D10SM</td>
			<td style="width:464px;"></td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:455px;">Low Impedance Headstage&#160; (RA4LI)</td>
			<td style="width:251px;">Tucker-Davis Technologies</td>
			<td style="width:207px;"></td>
			<td style="width:464px;"></td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:455px;">Speakers (MF1 Multi-Field Magnetic Speaker)</td>
			<td style="width:251px;">Tucker-Davis Technologies</td>
			<td style="width:207px;"></td>
			<td style="width:464px;"></td>
		</tr>
		<tr height="64">
			<td height="64" style="height:64px;width:455px;">System 3 Evoked Potential Workstation</td>
			<td style="width:251px;">Tucker-Davis Technologies</td>
			<td style="width:207px;"></td>
			<td style="width:464px;">The System is composed of: RP2 processor, RA16 base station, PA5 attenuator, SA1 amplifier, MA3 microphone amplifier, RA4LI impedance headstage and RA4A medusa pre-amplifier&#160;</td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:455px;">SigGenRP software</td>
			<td style="width:251px;">Tucker-Davis Technologies</td>
			<td style="width:207px;"></td>
			<td style="width:464px;"></td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:455px;">Warming pads (TP pads)</td>
			<td style="width:251px;">Gaymar</td>
			<td style="width:207px;"># TP3E</td>
			<td style="width:464px;"></td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:455px;">Statistics software (SPSS)</td>
			<td style="width:251px;">IBM</td>
			<td style="width:207px;"></td>
			<td style="width:464px;"></td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:455px;">Chitosan (deacetylated)</td>
			<td style="width:251px;">Sigma-Aldrich</td>
			<td style="width:207px;"># C3646</td>
			<td style="width:464px;"></td>
		</tr>
		<tr height="43">
			<td height="43" style="height:43px;width:455px;">Acetic acid (glacial)</td>
			<td style="width:251px;">VWR</td>
			<td style="width:207px;"># 20103.295</td>
			<td style="width:464px;">CAUTION: flammable liquid, skin corrosion and respiratory and skin sensitizer</td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:455px;">Glycerophosphate</td>
			<td style="width:251px;">Sigma</td>
			<td style="width:207px;"># SLBG3671V</td>
			<td style="width:464px;"></td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:455px;">Ringer&#180;s lactate buffer</td>
			<td style="width:251px;">Braun</td>
			<td style="width:207px;"># 1520-ESP</td>
			<td style="width:464px;"></td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:455px;">Medetomidine (Domtor)</td>
			<td style="width:251px;">Esteve</td>
			<td style="width:207px;"># 02400190</td>
			<td style="width:464px;"></td>
		</tr>
		<tr height="43">
			<td height="43" style="height:43px;width:455px;">Phentanile (Fentanest)</td>
			<td style="width:251px;">Kern Pharma</td>
			<td style="width:207px;"># 756650.2</td>
			<td style="width:464px;">CAUTION: &#160;&#160;avoid contact of the drug with open wounds or accidental self-inflicted injections</td>
		</tr>
		<tr height="64">
			<td height="64" style="height:64px;width:455px;">Isoflurane (IsoVet)</td>
			<td style="width:251px;">Braun</td>
			<td style="width:207px;"># 469860</td>
			<td style="width:464px;">CAUTION: Avoid exposures at ceiling concentrations greater than 2ppm of any halogenated anesthetic agent over a sampling period not to exceed one hour.</td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:455px;">Surgical microscope (OPMI pico)</td>
			<td style="width:251px;">Zeiss</td>
			<td style="width:207px;"></td>
			<td style="width:464px;"></td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:455px;">Sterile drape (Foliodrape)</td>
			<td style="width:251px;">Hartmann</td>
			<td style="width:207px;"># 277546</td>
			<td style="width:464px;"></td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:455px;">Sterilizer&#160;</td>
			<td style="width:251px;">Fine Science Tools</td>
			<td style="width:207px;"># 18000-45</td>
			<td style="width:464px;"></td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:455px;">Scalpel blade</td>
			<td style="width:251px;">Swann Morton</td>
			<td style="width:207px;"># 0205</td>
			<td style="width:464px;">CAUTION</td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:455px;">Scalpel handle</td>
			<td style="width:251px;">Fine Science Tools</td>
			<td style="width:207px;"># 91003-12</td>
			<td style="width:464px;"></td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:455px;">Pividone iodine based antiseptic (Betadine)</td>
			<td style="width:251px;">Meda Pharma SAU</td>
			<td style="width:207px;"># M-12207</td>
			<td style="width:464px;"></td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:455px;">Adventitia scissors (SAS18-R8)</td>
			<td style="width:251px;">S&#38;T</td>
			<td style="width:207px;"># 12075-12</td>
			<td style="width:464px;"></td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:455px;">Curved scissors</td>
			<td style="width:251px;">CM Instrumente</td>
			<td style="width:207px;"># AJ023-18</td>
			<td style="width:464px;"></td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:455px;">Forceps</td>
			<td style="width:251px;">CM Instrumente</td>
			<td style="width:207px;"># BB019-18</td>
			<td style="width:464px;"></td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:455px;">Gelatine sponge (Spongostan)</td>
			<td style="width:251px;">ProNaMAc</td>
			<td style="width:207px;"># MS0001</td>
			<td style="width:464px;"></td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:455px;">Microlance 27G</td>
			<td style="width:251px;">Becton Dickinson</td>
			<td style="width:207px;"># 302200</td>
			<td style="width:464px;"></td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:455px;">Microliter syringe (701 RN SYR)</td>
			<td style="width:251px;">Hamilton</td>
			<td style="width:207px;"># 80330</td>
			<td style="width:464px;"></td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:455px;">Catheter (Microfil 34G)</td>
			<td style="width:251px;">World Precision Instruments&#160;</td>
			<td style="width:207px;"># MF34G-5</td>
			<td style="width:464px;"></td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:455px;">Tissue Adhesive (Vetbond)</td>
			<td style="width:251px;">3M</td>
			<td style="width:207px;"># 1469SB</td>
			<td style="width:464px;"></td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:455px;">Needle holder (Round handled needle holder)&#160;</td>
			<td style="width:251px;">Fine Science Tools</td>
			<td style="width:207px;"># 12075-12</td>
			<td style="width:464px;"></td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:455px;">Silk surgical suture (Braided Silk 5/0)</td>
			<td style="width:251px;">Arago</td>
			<td style="width:207px;"># 990011</td>
			<td style="width:464px;"></td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:455px;">Chlorhexidine (Cristalmina)</td>
			<td style="width:251px;">Salvat</td>
			<td style="width:207px;"># 787341</td>
			<td style="width:464px;"></td>
		</tr>
		<tr height="43">
			<td height="43" style="height:43px;width:455px;">Pentobarbital&#160; (Dolethal)</td>
			<td style="width:251px;">Ventoquinol</td>
			<td style="width:207px;"># VET00040</td>
			<td style="width:464px;">CAUTION: &#160;&#160;avoid contact of the drug with open wounds or accidental self-inflicted injections</td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:455px;">Stereomicroscope (Leica)</td>
			<td style="width:251px;">Meyer Instruments</td>
			<td style="width:207px;"># MZ75</td>
			<td style="width:464px;"></td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:455px;">Vannas Micro-dissecting (Eye) Scissors Spring Action</td>
			<td style="width:251px;">Harvard Apparatus</td>
			<td style="width:207px;"># 28483</td>
			<td style="width:464px;"></td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:455px;">Jeweller&#8217;s forceps (Dumont)</td>
			<td style="width:251px;">Fine Science Tools</td>
			<td style="width:207px;"># 11252-00</td>
			<td style="width:464px;"></td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:455px;">RNase Decontamination Solution&#160; (RNaseZap)</td>
			<td style="width:251px;">Sigma-Aldrich</td>
			<td style="width:207px;"># R2020</td>
			<td style="width:464px;"></td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:455px;">RNA Stabilization Solution&#160; (RNAlater)</td>
			<td style="width:251px;">Thermo Fisher Scientific</td>
			<td style="width:207px;"># R0901</td>
			<td style="width:464px;"></td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:455px;">Purification RNA kit (RNeasy)</td>
			<td style="width:251px;">Qiagen</td>
			<td style="width:207px;"># 74104</td>
			<td style="width:464px;"></td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:455px;">cDNA Reverse Transcription Kit</td>
			<td style="width:251px;">Thermo Fisher Scientific</td>
			<td style="width:207px;"># 4368814</td>
			<td style="width:464px;"></td>
		</tr>
		<tr height="22">
			<td height="154" rowspan="1" style="height:154px;width:455px;">Gene expression assay (TaqMan probes)</td>
			<td rowspan="1" style="width:251px;">Thermo Fisher Scientific</td>
			<td height="22" style="height:22px;width:207px;">Il1b: Mm00446190_m1 
			Il6: Mm00446190_m1 
			Tgfb1: Mm01178820_m1 
			Tnfa: Mm99999068_m1 
			Il10: Mm00439614_m1 
			Dusp1: Mm00457274_g1 
			Hprt1: Mm00446968_m1</td>
			<td rowspan="1" style="width:464px;"></td>
		</tr>
		<tr height="20">
			<td height="41" rowspan="2" style="height:41px;width:455px;">Real-time PCR System (7900HT)</td>
			<td rowspan="2" style="width:251px;">Applied Biosystems</td>
			<td rowspan="2" style="width:207px;"># 4329001</td>
			<td rowspan="2" style="width:464px;"></td>
		</tr>
		<tr height="21">
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:455px;">Paraformaldehyde (PFA)</td>
			<td style="width:251px;">Merck</td>
			<td style="width:207px;"># 1040051000</td>
			<td style="width:464px;">TOXIC: PFA is a potential&#160;carcinogen</td>
		</tr>
		<tr height="64">
			<td height="64" style="height:64px;width:455px;">Ethylenediaminetetraacetic acid (EDTA)</td>
			<td style="width:251px;">Merck</td>
			<td style="width:207px;"># 405491</td>
			<td style="width:464px;">CAUTION:&#160; harmful if inhaled, may cause damage to respiratory tract through prolonged or repeated exposure if 
			inhaled.</td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:455px;">Hematoxylin solution</td>
			<td style="width:251px;">Sigma-Aldrich</td>
			<td style="width:207px;"># HHS16</td>
			<td style="width:464px;"></td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:455px;">Eosin Y</td>
			<td style="width:251px;">Sigma-Aldrich</td>
			<td style="width:207px;"># E4382</td>
			<td style="width:464px;">Hazards: causes serious eye irritation</td>
		</tr>
	</tbody>
</table>

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.

Hearing was tested by ABR before and at several times after microsurgery to evaluate the impact on auditory function (Figure 1A). ABR registers were performed under anaesthesia to avoid animal movement and voltage artefacts and therefore improve its reproducibility27. Intraperitoneally administered ketamine based combinations or inhalatory isoflurane were usually employed to anesthetize animals during the ABR tests. The ketamine/xylazine combination provides a short-acting (2-3 min) induction and a stable, safe maintenance phase while performing ABR registers. It should be noted that isoflurane can affect ABR measurement sensitivity38. For ABR registers, subdermal electrodes are placed in specific locations (Figure 1B) and the electrical impedance is measured. If the impedance is 3 kOhm or higher, electrode positioning has to be checked to avoid alterations in ABR wave amplitude.
Intratympanic delivery is performed in mice by two microsurgical procedures (Figure 2). Exposure of the bulla during bullostomy involves retraction of the submandibular glands and digastric muscle. This procedure is carried out with extreme care because the carotid artery and vagal nerve are very close (Figure 2A). Next, the bulla is drilled to localize the stapedial artery and the RW membrane (Figure 2B). To avoid cracking bone, a small 0.5 mm aperture is made with a 27 G needle before drilling. The 34 G catheter is directed through the bullostomy towards the RW membrane and a small volume of vehicle is delivered onto the window niche (Figure 2C). The transtympanic injection is performed through an incision in the pars flaccida of the tympanic membrane with a 27 G needle; a larger one can provoke a tear in the membrane. Before the injection, we recommend making an additional incision in the pars tensa to allow the outflow of air during injection of the vehicle (Figure 2F). It is critical to avoid damage of the stapedial artery, a branch of internal carotid artery, which would lead to life-threatening bleeding.
Mice with bullostomy or transtympanic surgeries preserved hearing throughout the experiment, similar to non-operated controls (Figure 3). ABR thresholds in response to clicks and tone bursts did not change significantly after microsurgery compared to baseline values. No significant differences were observed between bullostomy and transtympanic approaches. Morphological studies were carried out to confirm correct vehicle delivery into the middle ear and to assess the potential changes caused by the procedures in cochlear cytoarchitecture. None of the main cochlear regions showed morphological alterations and animals from both procedures presented a similar morphology of all cochlear structures (Figure 4A). In addition, the cochlear profiles for gene expression of pro- and anti-inflammatory cytokines were also studied. Despite lack of functional differences in ABR data between the two procedures, bullostomy caused a stronger inflammatory response than the transtympanic approach (Figure 4B).
<img alt="Figure 1" src="/files/ftp_upload/54951/54951fig1.jpg" /> 
Figure 1. Experimental Design and Hearing Assessment. (A) Diagram of the experimental procedure. Hearing was evaluated with ABR before and after microsurgery. Cochlear samples were obtained 28 days after microsurgery. (B) Anesthetized mouse in prone position over the heating pad inside a sound-attenuating chamber, with subdermal electrodes placed in the scalp between the ears over the vertex of the skull (active, positive); in the parotid region below the pinna (reference, negative) and in the back (ground). The free-field speaker is placed at a fixed distance (5 cm) facing the right ear. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/54951/54951fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" src="/files/ftp_upload/54951/54951fig2.jpg" /> 
Figure 2. Microsurgery for Vehicle Application. (A) Ventral view of the tympanic bulla. The bullostomy is performed caudal to the facial nerve with a 27 G needle. (B) RWN and stapedial artery can be observed through the perforation. (C) The 34 G catheter is directed through the bullostomy toward the RW niche. (D) One month after the bullostomy, a small bony scar is present in the opening site (arrowhead). (E) Lateral view of the ear, showing the incision in the external ear canal and the tympanic membrane (square). (F) Detail of the tympanic membrane. A puncture was made at the caudal upper quadrant of the tympanic membrane using a 27 G needle (black asterisk, in the pars flaccida); the injection was made through this perforation using a 34 G catheter. An additional hole was made in the cranial inferior quadrant of the membrane (white asterisk, in the pars tensa) prior to the injection to balance the tympanic pressure. (G) View of the 34 G catheter through the puncture of the tympanic membrane. (H) View of the cochlear region 24 h after the microsurgery. RWN filled with vehicle solution (asterisk). Lat, lateral; Ro, rostral; Do, dorsal; Ma, malleus; Co, cochlea; OW, Oval window; RWN, round window niche. Scale bars = 200 &#181;m in A, D, F; Scale bars = 100 &#181;m in B, C, H; Scale bars = 1,000 &#181;m in E, G. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/54951/54951fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" src="/files/ftp_upload/54951/54951fig3.jpg" /> 
Figure 3. Hearing Assessment. Evolution of ABR thresholds (mean &#177;SEM, in dB SPL) in response to click (A) and tone burst (B) stimuli, before and 7, 14 and 28 days after micro-surgery in male eight-week-old C57BL/6J mice. Bullostomy (orange; n = 11); transtympanic injection (blue; n = 6); non-operated (grey; n = 11), <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/54951/54951fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" src="/files/ftp_upload/54951/54951fig4.jpg" /> 
Figure 4. Cochlear Morphology and Gene Expression Analysis. (A) Morphology of the main cochlear structures at the base of the cochlea. Haematoxilin-eosin staining of mid-modiolar paraffin sections (7 &#181;m), of ears from non-operated mice, and mice one month after microsurgery intervention by bullostomy or transtympanic injection. The scala media compartment (a,b,c) presents all the main components. Details of each of these structures (numbered boxes) are shown in the subsequent images: spiral ganglion (1), organ of Corti (2), spiral ligament (3) and stria vascularis (4). Inner hair cell (asterisk); outer hair cells (head of arrow). Scale bars = 100 &#181;m in a,b,c; Scale bars = 50 &#181;m in a-1,2,3,4. (B) Cochlear expression of inflammatory markers 28 d&#160;after the microsurgery. Comparison between bullostomy (orange) and transtympanic injection (blue) and to non-operated mice (white). *: non-operated vs. operated groups; ^: comparison between operated groups. Gene expression levels are represented as 2-&#916;&#916;Ct, or the n-fold difference relative to non-operated group.Values are presented as mean &#177; SEM of triplicates from pool samples of 3 mice per condition. Statistical significance: **p&#160;&#8804;0.01; ***p&#160;&#8804;0.001; ^^p&#160;&#8804;0.01; ^^^p&#160;&#8804;0.001. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/54951/54951fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about A Comparative Study of Drug Delivery Methods Targeted to the Mouse Inner Ear: Bullostomy Versus Transtympanic Injection at JoVE.com

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.

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. ...

a-comparative-study-drug-delivery-methods-targeted-to-mouse-inner-ear

Nanyang Technological University

Research

JoVE Journal

Biology

13.1K Views.  Instituto de Investigaciones Biomédicas (IIBm) Alberto Sols CSIC-UAM. 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.

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

Testing Visual Sensitivity to the Speed and Direction of Motion in Lizards

Testing visual sensitivity in any species provides basic information regarding behaviour, evolution, and ecology. However, testing specific features of the visual system provide more empirical evidence for functional applications. Investigation into the sensory system provides information about the sensory capacity, learning and memory ability, and establishes known baseline behaviour in which to gauge deviations (Burghardt, 1977). However, unlike mammalian or avian systems, testing for learning and memory in a reptile species is difficult. Furthermore, using an operant paradigm as a psychophysical measure of sensory ability is likewise as difficult. Historically, reptilian species have responded poorly to conditioning trials because of issues related to motivation, physiology, metabolism, and basic biological characteristics. Here, I demonstrate an operant paradigm used a novel model lizard species, the Jacky dragon (Amphibolurus muricatus) and describe how to test peripheral sensitivity to salient speed and motion characteristics. This method uses an innovative approach to assessing learning and sensory capacity in lizards. I employ the use of random-dot kinematograms (RDKs) to measure sensitivity to speed, and manipulate the level of signal strength by changing the proportion of dots moving in a coherent direction. RDKs do not represent a biologically meaningful stimulus, engages the visual system, and is a classic psychophysical tool used to measure sensitivity in humans and other animals. Here, RDKs are displayed to lizards using three video playback systems. Lizards are to select the direction (left or right) in which they perceive dots to be moving. Selection of the appropriate direction is reinforced by biologically important prey stimuli, simulated by computer-animated invertebrates.

Testing visual sensitivity in lizards using an operant conditioning paradigm that employs video playback of random-dot kinematograms and computer-generated invertebrates.

Testing visual sensitivity in any species provides basic information regarding behaviour, evolution, and ecology. However, testing specific features of ...

Gross and Fine Dissection of Inner Ear Sensory Epithelia in Adult Zebrafish (Danio rerio)

Neurosensory epithelia in the inner ear are the crucial structures for hearing and balance functions. Therefore, it is important to understand the cellular and molecular features of the epithelia, which are mainly composed of two types of cells: hair cells (HCs) and supporting cells (SCs). Here we choose to study the inner ear sensory epithelia in adult zebrafish not only because the epithelial structures are highly conserved in all vertebrates studied, but also because the adult zebrafish is able to regenerate HCs, an ability that mammals lose shortly after birth. We use the inner ear of adult zebrafish as a model system to study the mechanisms of inner ear HC regeneration in adult vertebrates that could be helpful for clinical therapy of hearing/balance deficits in human as a result of HC loss.
Here we demonstrate how to do gross and fine dissections of inner ear sensory epithelia in adult zebrafish. The gross dissection removes the tissues surrounding the inner ear and is helpful for preparing tissue sections, which allows us to examine the detailed structure of the sensory epithelia. The fine dissection cleans up the non-sensory-epithelial tissues of each individual epithelium and enables us to examine the heterogeneity of the whole epithelium easily in whole-mount epithelial samples.

Gross and Fine Dissection of Inner Ear Sensory Epithelia in Adult Zebrafish Danio rerio

The inner ear sensory epithelium of adult zebrafish is a good model system for understanding the mechanisms of hair cell regeneration in adult vertebrates. This protocol demonstrates the fine dissection of the epithelia, through which we can get tissue samples for studying the regenerative events at cellular and subcellular levels.

Neurosensory epithelia in the inner ear are the crucial structures for hearing and balance functions. Therefore, it is important to understand the ...

Methods for the Study of the Zebrafish Maxillary Barbel

Barbels are skin sensory appendages found in fishes, reptiles and amphibians. The zebrafish, Danio rerio, develops two pairs of barbels- a short nasal pair and a longer maxillary pair. Barbel tissue contains cells of ectodermal, mesodermal and neural crest origin, including skin cells, glands, taste buds, melanocytes, circulatory vessels and sensory nerves. Unlike most adult tissue, the maxillary barbel is optically clear, allowing us to visualize the development and maintenance of these tissue types throughout the life cycle. 
This video shows early development of the maxillary barbel (beginning approximately one month post-fertilization) and demonstrates a surgical protocol to induce regeneration in the adult appendage (&gt;3 months post-fertilization). Briefly, the left maxillary barbel of an anesthetized fish is elevated with sterile forceps just distal to the caudal edge of the maxilla. A fine, sterile spring scissors is positioned against the forceps to cut the barbel shaft at this level, establishing an anatomical landmark for the amputation plane. Regenerative growth can be measured with respect to this plane, and in comparison to the contralateral barbel. Barbel tissue regenerates rapidly, reaching maximal regrowth within 2 weeks of injury.
Techniques for analyzing the regenerated barbel include dissecting and embedding matched pairs of barbels (regenerate and control) in the wells of a standard DNA electrophoresis gel. Embedded specimens are conveniently photographed under a stereomicroscope for gross morphology and morphometry, and can be stored for weeks prior to downstream applications such as paraffin histology, cryosectioning, and/or whole mount immunohistochemistry. These methods establish the maxillary barbel as a novel in vivo tissue system for studying the regenerative capacity of multiple cell types within the genetic context of zebrafish.

The zebrafish maxillary barbel is an integumentary sense organ containing ectodermal, mesodermal and neural crest derivatives. Importantly, the adult barbel can regenerate after proximal amputation. This video introduces maxillary barbel development and demonstrates a surgical protocol to induce regeneration, followed by collection, embedding and downstream imaging of barbel specimens.

Barbels are skin sensory appendages found in fishes, reptiles and amphibians.  The zebrafish, Danio rerio, develops two pairs of barbels- a short nasal ...

Direct Delivery of MIF Morpholinos Into the Zebrafish Otocyst by Injection and Electroporation Affects Inner Ear Development

In recent years, electroporation has become a popular technique for in vivo transfection of DNA, RNA, and morpholinos into various tissues, including the eye, brain, and somites of zebrafish. The advantage of electroporation over other methods of genetic manipulation is that specific tissues can be targeted, both spatially and temporally, for the introduction of macromolecules by the application of electrical current. Here we describe the use of electroporation for transfecting mif and mif-like morpholinos into the tissues of the developing inner ear of the zebrafish. In past studies, mif morpholino injected into embryos at the 1- to 8-cell stage resulted in widespread morphological changes in the nervous system and eye, as well as the ear. By targeting the tissues of the inner ear at later stages in development, we can determine the primary effects of MIF in the developing inner ear, as opposed to secondary effects that may result from the influence of other tissues. By using phalloidin and acetylated tubulin staining to study the morphology of neurons, neuronal processes, and hair cells associated with the posterior macula, we were able to assess the efficacy of electroporation as a method for targeted transfection in the zebrafish inner ear. The otic vesicles of 24hpf embryos were injected with morpholinos and electroporated and were then compared to embryos that had received no treatment or had been only injected or electroporated. Embryos that were injected and electroporated showed a decrease in hair cell numbers, decreased innervation by the statoacoustic ganglion (SAG) and fewer SAG neurons compared with control groups. Our results showed that direct delivery of morpholinos into otocysts at later stages avoids the non-specific nervous system and neural crest effects of morpholinos delivered at the 1-8 cell stage. It also allows examination of effects that are directed to the inner ear and not secondary effects on the ear from primary effects on the brain, neural crest or periotic mesenchyme.

A method to deliver morpholinos directly into the zebrafish otocyst at 24hpf has been developed. Using microinjection of morpholinos into the lumen of otic vesicle and electroporation to effect penetration, we were able to bypass the effect of morpholinos on the brain and obtain effects specific to the inner ear.

In recent years, electroporation has become a popular technique for in vivo transfection of DNA, RNA, and morpholinos into various tissues, including the ...

A High Throughput in situ Hybridization Method to Characterize mRNA Expression Patterns in the Fetal Mouse Lower Urogenital Tract

Development of the lower urogenital tract (LUT) is an intricate process. This complexity is evidenced during formation of the prostate from the fetal male urethra, which relies on androgenic signals and epithelial-mesenchymal interactions1,2. Understanding the molecular mechanisms responsible for prostate development may reveal growth mechanisms that are inappropriately reawakened later in life to give rise to prostate diseases such as benign prostatic hyperplasia and prostate cancer.
The developing LUT is anatomically complex. By the time prostatic budding begins on 16.5 days post conception (dpc), numerous cell types are present. Vasculature, nerves and smooth muscle reside within the mesenchymal stroma3. This stroma surrounds a multilayered epithelium and gives rise to the fetal prostate through androgen receptor-dependent paracrine signals4. The identity of the stromal androgen receptor-responsive genes required for prostate development and the mechanism by which prostate ductal epithelium forms in response to these genes is not fully understood. The ability to precisely identify cell types and localize expression of specific factors within them is imperative to further understand prostate development. In situ hybridization (ISH) allows for localization of mRNAs within a tissue. Thus, this method can be used to identify pattern and timing of expression of signaling molecules and their receptors, thereby elucidating potential prostate developmental regulators.
Here, we describe a high throughput ISH technique to identify mRNA expression patterns in the fetal mouse LUT using vibrating microtome-cut sections. This method offers several advantages over other ISH protocols. Performing ISH on thin sections adhered to a slide is technically difficult; cryosections frequently have poor structural quality while both cryosections and paraffin sections often result in weak signal resolution. Performing ISH on whole mount tissues can result in probe trapping. In contrast, our high throughput technique utilizes thick-cut sections that reveal detailed tissue architecture. Modified microfuge tubes allow easy handling of sections during the ISH procedure. A maximum of 4 mRNA transcripts can be screened from a single 17.5dpc LUT with up to 24 mRNA transcripts detected in a single run, thereby reducing cost and maximizing efficiency. This method allows multiple treatment groups to be processed identically and as a single unit, thereby removing any bias for interpreting data. Most pertinently for prostate researchers, this method provides a spatial and temporal location of low and high abundance mRNA transcripts in the fetal mouse urethra that gives rise to the prostate ductal network.

A High Throughput in situ Hybridization Method to Characterize mRNA Expression Patterns in the Fetal Mouse Lower Urogenital Tract

Here, we describe an efficient high throughput in situ hybridization (ISH) method for visualizing patterns of mRNA expression in developing fetal mouse prostate tissue sections. The method can be easily adapted to visualize mRNA expression patterns in other mouse tissues or in tissues from other species.

Development of the lower urogenital tract (LUT) is an intricate process. This complexity is evidenced during formation of the prostate from the fetal male ...

Intraductal Injection for Localized Drug Delivery to the Mouse Mammary Gland

Herein we describe a protocol to deliver various reagents to the mouse mammary gland via intraductal injections. Localized drug delivery and knock-down of genes within the mammary epithelium has been difficult to achieve due to the lack of appropriate targeting molecules that are independent of developmental stages such as pregnancy and lactation. Herein, we describe a technique for localized delivery of reagents to the mammary gland at any stage in adulthood via intraductal injection into the nipples of mice. The injections can be performed on live mice, under anesthesia, and allow for a non-invasive and localized drug delivery to the mammary gland. Furthermore, the injections can be repeated over several months without damaging the nipple. Vital dyes such as Evans Blue are very helpful to learn the technique. Upon intraductal injection of the blue dye, the entire ductal tree becomes visible to the eye. Furthermore, fluorescently labeled reagents also allow for visualization and distribution within the mammary gland. This technique is adaptable for a variety of compounds including siRNA, chemotherapeutic agents, and small molecules.

A protocol for the non-invasive intraductal delivery of aqueous reagents to the mouse mammary gland is described. The method takes advantage of localized injection into the nipples of mammary glands targeting mammary ducts specifically. This technique is adaptable for a variety of compounds including siRNA, chemotherapeutic agents and small molecules.

Herein we describe a protocol to deliver various reagents to the mouse mammary gland via intraductal injections. Localized drug delivery and knock-down of ...

Phosphopeptide Analysis of Rodent Epididymal Spermatozoa

Spermatozoa are quite unique amongst cell types. Although produced in the testis, both nuclear gene transcription and translation are switched off once the pre-cursor round cell begins to elongate and differentiate into what is morphologically recognized as a spermatozoon. However, the spermatozoon is very immature, having no ability for motility or egg recognition. Both of these events occur once the spermatozoa transit a secondary organ known as the epididymis. During the ~12 day passage that it takes for a sperm cell to pass through the epididymis, post-translational modifications of existing proteins play a pivotal role in the maturation of the cell. One major facet of such is protein phosphorylation. In order to characterize phosphorylation events taking place during sperm maturation, both pure sperm cell populations and pre-fractionation of phosphopeptides must be established. Using back flushing techniques, a method for the isolation of pure spermatozoa of high quality and yield from the distal or caudal epididymides is outlined. The steps for solubilization, digestion, and pre-fractionation of sperm phosphopeptides through TiO2 affinity chromatography are explained. Once isolated, phosphopeptides can be injected into MS to identify both protein phosphorylation events on specific amino acid residues and quantify the levels of phosphorylation taking place during the sperm maturation processes.

Proteomic analysis of any cell type is highly dependent on both purity and pre-fractionation of the starting material in order to de-complexify the sample prior to liquid chromatography mass spectrometry (MS). By using back-flushing techniques, pure spermatozoa can be obtained from rodents. Following digestion, phosphopeptides can be enriched using TiO2.

Spermatozoa are quite unique amongst cell types. Although produced in the testis, both nuclear gene transcription and translation are switched off once ...

Trans-inner Cell Mass Injection of Embryonic Stem Cells Leads to Higher Chimerism Rates

In an effort to increase efficiency in the creation of genetically modified mice via ES Cell methodologies, we present an adaptation to the current blastocyst injection protocol. Here we report that a simple rotation of the embryo, and injection through Trans-Inner cell mass (TICM) increased the percentage of chimeric mice from 31% to 50%, with no additional equipment or further specialized training. 26 different inbred clones, and 35 total clones were injected over a period of 9 months. There was no significant difference in either pregnancy rate or recovery rate of embryos between traditional injection techniques and TICM. Therefore, without any major alteration in the injection process and a simple positioning of the blastocyst and injecting through the ICM, releasing the ES cells into the blastocoel cavity can potentially improve the quantity of chimeric production and subsequent germline transmission.

Here we present a protocol to increase chimera production without the use of new equipment. A simple orientation change of the embryo for injection can increase the number of embryos produced, and potentially reduce the timeline to germline transmission.

In an effort to increase efficiency in the creation of genetically modified mice via ES Cell methodologies, we present an adaptation to the current ...

Laser-assisted Lentiviral Gene Delivery to Mouse Fertilized Eggs

Lentiviruses are efficient vectors for gene delivery to mammalian cells. Following transduction, the lentiviral genome is stably incorporated into the host chromosome and is passed on to progeny. Thus, they are ideal vectors for creation of stable cell lines, in vivo delivery of indicators, and transduction of single cell fertilized eggs to create transgenic animals. However, mouse fertilized eggs and early stage embryos are protected by the zona pellucida, a glycoprotein matrix that forms a barrier against lentiviral gene delivery. Lentiviruses are too large to penetrate the zona and are typically delivered by microinjection of viral particles into the perivitelline cavity, the space between the zona and the embryonic cells. The requirement for highly skilled technologists and specialized equipment has minimized the use of lentiviruses for gene delivery to mouse embryos. This article describes a protocol for permeabilizing the mouse fertilized eggs by perforating the zona with a laser. Laser-perforation does not result in any damage to embryos and allows lentiviruses to gain access to embryonic cells for gene delivery. Transduced embryos can develop into blastocyst in vitro, and if implanted in pseudopregnant mice, develop into transgenic pups. The laser used in this protocol is effective and easy to use. Genes delivered by lentiviruses stably incorporate into mouse embryonic cells and are germline transmittable. This is an alternative method for creation of transgenic mice that requires no micromanipulation and microinjection of fertilized eggs.

Mouse fertilized eggs and early stage embryos are protected by the zona pellucida, a glycoprotein matrix that forms a barrier against gene delivery. This article describes a protocol for perforating the zona with a laser to transduce embryonic cells with lentiviral vectors and to create transgenic mice.

Lentiviruses are efficient vectors for gene delivery to mammalian cells. Following transduction, the lentiviral genome is stably incorporated into the ...

Efficient Dissection and Culture of Primary Mouse Retinal Pigment Epithelial Cells

Eye disorders affect millions of people worldwide, but the limited availability of human tissues hinders their study. Mouse models are powerful tools to understand the pathophysiology of ocular diseases because of their similarities with human anatomy and physiology. Alterations in the retinal pigment epithelium (RPE), including changes in morphology and function, are common features shared by many ocular disorders. However, successful isolation and culture of primary mouse RPE cells is very challenging. This paper is an updated audiovisual version of the protocol previously published by Fernandez-Godino et al. in 2016 to efficiently isolate and culture primary mouse RPE cells. This method is highly reproducible and results in robust cultures of highly polarized and pigmented RPE monolayers that can be maintained for several weeks on Transwells. This model opens new avenues for the study of the molecular and cellular mechanisms underlying eye diseases. Moreover, it provides a platform to test therapeutic approaches that can be used to treat important eye diseases with unmet medical needs, including inherited retinal disorders and macular degenerations.

This protocol, which was originally reported by Fernandez-Godino et al. in 20161, describes a method to efficiently isolate and culture mouse RPE cells, which form a functional and polarized RPE monolayer within one week on Transwell plates. The procedure takes approximately 3 hours.

Eye disorders affect millions of people worldwide, but the limited availability of human tissues hinders their study. Mouse models are powerful tools to ...