Name: cochlear implantation in the guinea pig
Uploaded: 2018-06-15T13:00:00.000+00:00
Duration: 9 min 56 s
Description: Watch this Scientific Journal Video about Cochlear Implantation in the Guinea Pig at JoVE.com

The authors want to thank Sandra Peiritsch for the care of the animals and Noelani Peet for medical writing. The financial support by the Austrian Science Fund (FWF grant P 24260-B19) and MED-EL Austria is gratefully acknowledged.

All animal experiments were approved by the local animal welfare committee and the Austrian Federal Ministry for Science, Research and Economy.
1. Prepare the Equipment and Setup Required for Surgery
<ol>
	<li>Position the surgical microscope, drill, heating plate and pulse oximeter to allow easy and efficient handling during surgery. Check the function of the devices based on the manufacturer&#39;s manual. Make sure that the heating plate is set at 38 &#176;C in order to protect the animal from hypothermia during the surgery.</li>
	<li>Put on a hood and a mask.</li>
	<li>Perform surgical hand disinfection. Wash hands meticulously with soap. Dry hands and then use an alcoholic hand sanitizer to disinfect hands. Put on gloves after the hands are dry.</li>
	<li>Prepare the sterilized surgical instruments and the equipment required for cochlear implantation. See the Table of Materials for the surgical instruments and equipment used for guinea pig CI implantation in this protocol.</li>
	<li>When compound action potential (CAP) measurements are intended, prepare an approximately 3.5 cm-long piece of Teflon-insulated gold wire by carefully removing parts of the insulation from both ends using a micro forceps (approximately 3 mm from one end and 5 mm from the other end). Prepare a second piece of gold wire (approximately 2.5 cm) with uninsulated ends of approximately 5 mm. Place the prepared wires in alcohol or disinfectant.</li>
</ol>
2. Anesthesia, Medication and Animal Preparation
<ol>
	<li>Weigh the animal. 
	NOTE: Animals used are female Dunkin-Hartley albino guinea pigs. The weight of the animals used, which are approximately 4-6 weeks old, ranges from 300 to 400 g.</li>
	<li>Prepare the anesthetics and medication required for surgery based on the weight of the animal. Prepare 0.72 mL of the anesthetic mixture, consisting of 0.06 mL ketamine (100 mg/mL), 0.18 mL medetomidine (1 mg/mL), 0.12 mL of midazolam (5 mg/mL) and 0.36 mL of fentanyl (50 &#181;g/mL) for surgery on a 400 g guinea pig. See Table 1 for weight-based doses of the anesthetics.</li>
	<li>Inject the mixture of ketamine, medetomidine, midazolam and fentanyl subcutaneously to the fat pad in the neck of the animal using a 27 G needle. See Table 1 for weight based doses of the anesthetics. Cover the cage and leave the animal in a quiet place for 10 min before proceeding.</li>
	<li>Lubricate the eyes of the animal and keep them lubricated during the whole procedure. Shave the head of the animal, focusing on the retroauricular region to allow for a sufficient surgical access to the bulla.</li>
	<li>Position the animal on the heating plate in prone position and make sure that the animal is not in direct contact with the heating plate to prevent thermal burns. Place the pulse oximeter probe on a foot of the animal. Then carefully open the mouth of the animal with a small laryngoscope and clean the entire oral cavity from food rests by using a sucker.</li>
	<li>Keep the mouth of the animal opened with the laryngoscope. Carefully insert a stomach tube into the esophagus of the animal and slowly push it in the direction of the stomach until a resistance is felt.
	<ol>
		<li>Monitor the O2-saturation of the animal to make sure that the stomach tube is not in the trachea. Remove the stomach tube if there is a decrease in O2 saturation and try again after the animal is completely oxygenated.</li>
	</ol>
	</li>
	<li>Inject a mixture of physiological saline, 5% glucose and enrofloxacin into the fat pad in the neck of the animal using a 23 G needle.</li>
	<li>Use alcoholic hand sanitizer to disinfect hands again. Put on new clean gloves.</li>
	<li>Prepare the surgical field with&#160;alternating scrubs of povidone iodine and 70% ethanol and cover the animal. Use self-adhesive drapes or towel clamps to make sure that only the surgical field stays uncovered during the procedure.</li>
	<li>Subcutaneously inject 0.1 mL of 2% lidocaine solution into the area of the planned incision for sufficient local anesthesia and position the animal sideways.</li>
	<li>Re-dose the animal with &#188; of the initial dose of the anesthetic mixture every 30 min after the first injection to maintain sufficient anesthesia.</li>
</ol>
3. Cochlear Implantation
<ol>
	<li>Perform an approximately 2-3 cm skin incision 3-5 mm posterior to the pinna using a scalpel. Use bipolar cautery when necessary to minimize bleeding. 
	Note: Anesthetic depth should be confirmed by the lack of pedal withdrawal prior to the first incision. This test should be repeated every 15-20 minutes to monitor anesthetic depth throughout the procedure.&#160;</li>
	<li>Carefully cut the muscles in the retroauricular area after palpating the auditory bulla using a 15 scalpel or surgical scissors. 
	NOTE: Palpate the auditory bulla as a prominence under the muscles.</li>
	<li>Dissect the muscles from the bulla by pushing them gently aside using a raspatory or a cotton bud. Use a retractor to expose the full length of the incision and have unhindered access to the bulla.</li>
	<li>Use the tip of a 15 scalpel to drill a hole into the bulla. Carefully rotate the scalpel until the bone is perforated to allow for inspection of the middle ear structures.</li>
	<li>Enlarge the bullostomy as needed to make sure that the basal turn of the cochlea and the round window niche can be adequately visualized. Position the head of the animal in an inflected position in order to access these structures. Cover the area of the bullostomy with a small piece of a compress to prevent blood and extracellular fluid from running in to the middle ear. 
	NOTE: The inflected position of the head may obstruct the airway of the animal. Therefore, the oxygen saturation of the animal needs to be checked frequently.</li>
	<li>Position the animal in the prone position. Expose the vertex of the animal by performing a rectangular incision and removing the skin. Dissect the periosteum and clean the bone from any other kind of tissue or blood by using a scalpel.</li>
	<li>When CAP measurements are intended, finish the preparation of the 3.5 cm teflon insulated gold wire by forming a small hook with a micro forceps at the end of the wire, which has been uninsulated for 3 mm.</li>
	<li>When CAP measurements are intended, guide the end of the gold wire, which has been uninsulated for 5 mm, subcutaneously to the vertex through an 18 G peripheral venous catheter using a micro forceps. Use another micro forceps with the other hand to guide the hooked end of the wire carefully in to the middle ear.</li>
	<li>When CAP measurements are intended, inflect the head of the animal to visualize the area of the round window niche through the bullostomy. Use the other hand to hook the gold wire to the bony prominence of the round window niche by using a micro forceps.</li>
	<li>Maintain gentle tension at the gold wire and fix it to the cranial verge of the bullostomy with 10 - 15 &#181;L of tissue glue by using a 1 mL syringe with a 27 G needle. Avoid displacement of glue into the middle ear. See Figure 1 for an intraoperative picture of the round window area with the gold wire in situ.</li>
	<li>When CAP measurements are intended, connect the uninsulated end of the gold wire to the equipment used to measure auditory potentials and perform baseline CAP measurements. See Honeder et al., 2016, for a detailed description of the CAP measurements routinely performed in our laboratory.12</li>
	<li>Position the animal in prone position. Drill 2 holes 1 mm anterior to the lambda suture using a 1-mm burr without causing damage to the dura. Implant 2 stainless steel screws 2mm into the skull. 
	NOTE: The screws serve as fixation points for the connector of the electrode. Adapt the distance between the screws with respect to the size of the connector.</li>
	<li>Use an 18 G peripheral venous catheter to guide the electrode from the connector to the bulla in a tissue layer as close to the skull as possible.</li>
	<li>Mix the dental cement powder with the fluid for dental cement powder using a spatula according to the manufacturer&#39;s manual.</li>
	<li>Place 0.5 - 0.7 mL of semifluid dental cement between the screws by using a spatula. Position the connector of the electrode between the screws.</li>
	<li>Keep the connector in position until the dental cement is hardened. Make sure that the screws are encased by the cement to allow for stable fixation of the connector.</li>
	<li>Position the animal sideways. Carefully drill the cochleostomy 1 mm from the round window niche using a 0.5 mm diamond burr at a rotation rate of 5000 rounds per minute.</li>
	<li>Carefully insert the electrode in to the scala tympani to a depth of 4 mm.Withdraw the electrode and repeat the insertion. See Figure 2 for the electrode used for guinea pig cochlear implantation.</li>
	<li>Using a straight needle, seal the cochleostomy area with a small piece of muscle. Fix the electrode to the cranial verge of the bullostomy with 10 &#181;L of tissue glue by using a 1 mL syringe with a 27 G needle.</li>
	<li>Prepare the dental cement as mentioned in 3.14. Carefully close the bullostomy with approximately 0.3 mL of dental cement by using a spatula. Close the retroauricular incision using 5-0 absorbable sutures.</li>
	<li>Turn the animal to the prone position.</li>
	<li>When CAP measurements are intended, grab the posterior edge of the rectangular incision at the vertex of the animal by using a tissue forceps. Use a scissor with the other hand to make a subcutaneous tunnel of approximately 2 cm length in the neck of the animal.</li>
	<li>When CAP measurements are intended, implant the 2.5 cm gold wire subcutaneously into the neck of the animal using a forceps. Solder the shorter uninsulated end to the designated pin of the connector on the vertex of the animal.</li>
	<li>When CAP measurements are intended, solder the round window niche electrode (gold wire) to the appropriate pin of the connector on the vertex of the animal.</li>
	<li>Apply an additional amount of dental cement on the top of the connector to fully cover the insulated pins and the electrode.</li>
	<li>When CAP measurements are intended, perform postoperative measurements according to the research protocol.</li>
</ol>
4. Postoperative Care
<ol>
	<li>Apply atipamezole and flumazenil subcutaneously after the surgery and measurements in order to antagonize the anesthesia.</li>
	<li>Apply physiologic saline as fluid substitution to support the recovery of the animal from the surgery.</li>
	<li>Place the animal under a heating lamp until it has fully recovered from anesthesia and starts to move within the cage.
	<ol>
		<li>Prevent hyperthermia or burn by placing the heating lamp approximately 50 cm to the animal. Make sure that the body temperature of the animal is always between 37.5 &#176;C and 39 &#176;C.</li>
	</ol>
	</li>
	<li>Check the animal for symptoms of vestibular injury like nystagmus, circling or rolling over.13,14</li>
	<li>Apply buprenorphine for analgesia twice a day for two more days post-surgery. 
	NOTE: Before the first application of buprenorphine post-surgery make sure that the animal has completely recovered and has stable respiration. Application of this medication while the animal is still under anesthesia may lead to respiratory depression.</li>
	<li>Weigh the animal during the first 3 days post-surgery to detect possible weight loss as a surrogate marker of distress during this time. 
	NOTE: Weight loss of approximately 10% during the first 3 days post-surgery can be expected and should be considered common. This weight loss is temporary and will recover within a few days.</li>
</ol>

<ol>
	<li>Stevens, G., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Global+and+regional+hearing+impairment+prevalence:+an+analysis+of+42+studies+in+29+countries.">Global and regional hearing impairment prevalence: an analysis of 42 studies in 29 countries.</a> Eur J Public Health. 23 (1), 146-152 (2013).</li><li>Ciorba, A., Bianchini, C., Pelucchi, S., Pastore, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+impact+of+hearing+loss+on+the+quality+of+life+of+elderly+adults.">The impact of hearing loss on the quality of life of elderly adults.</a> Clin Interv Aging. 7, 159-163 (2012).</li><li>Arnoldner, C., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Electric+acoustic+stimulation+in+patients+with+postlingual+severe+high-frequency+hearing+loss:+clinical+experience.">Electric acoustic stimulation in patients with postlingual severe high-frequency hearing loss: clinical experience.</a> Adv Otorhinolaryngol. 67, 116-124 (2010).</li><li>Kral, A., O'Donoghue, G. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Profound+deafness+in+childhood.">Profound deafness in childhood.</a> N Engl J Med. 363 (15), 1438-1450 (2010).</li><li>DeMason, C., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Electrophysiological+properties+of+cochlear+implantation+in+the+gerbil+using+a+flexible+array.">Electrophysiological properties of cochlear implantation in the gerbil using a flexible array.</a> Ear Hear. 33 (4), 534-542 (2012).</li><li>Eshraghi, A. A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Pattern+of+hearing+loss+in+a+rat+model+of+cochlear+implantation+trauma.">Pattern of hearing loss in a rat model of cochlear implantation trauma.</a> Otol Neurotol. 26 (3), 442-447 (2005).</li><li>Mistry, N., Nolan, L. S., Saeed, S. R., Forge, A., Taylor, R. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cochlear+implantation+in+the+mouse+via+the+round+window:+effects+of+array+insertion.">Cochlear implantation in the mouse via the round window: effects of array insertion.</a> Hear Res. 312, 81-90 (2014).</li><li>Wysocki, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Topographical+anatomy+of+the+guinea+pig+temporal+bone.">Topographical anatomy of the guinea pig temporal bone.</a> Hear Res. 199 (1-2), 103-110 (2005).</li><li>Heffner, H. E., Heffner, R. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Hearing+ranges+of+laboratory+animals.">Hearing ranges of laboratory animals.</a> J Am Assoc Lab Anim Sci. 46 (1), 20-22 (2007).</li><li>Van Beek-King, J. M., Bhatti, P. T., Blake, D., Crawford, J., McKinnon, B. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Silicone-coated+thin+film+array+cochlear+implantation+in+a+feline+model.">Silicone-coated thin film array cochlear implantation in a feline model.</a> Otol Neurotol. 35 (1), 45-49 (2014).</li><li>Marx, M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cochlear+implantation+feasibility+in+rhesus+macaque+monkey:+anatomic+and+radiologic+results.">Cochlear implantation feasibility in rhesus macaque monkey: anatomic and radiologic results.</a> Otol Neurotol. 34 (7), 76-81 (2013).</li><li>Honeder, C., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Effects+of+sustained+release+dexamethasone+hydrogels+in+hearing+preservation+cochlear+implantation.">Effects of sustained release dexamethasone hydrogels in hearing preservation cochlear implantation.</a> Hear Res. , (2016).</li><li>Ris, L., Capron, B., de Waele, C., Vidal, P. P., Godaux, E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Dissociations+between+behavioural+recovery+and+restoration+of+vestibular+activity+in+the+unilabyrinthectomized+guinea-pig.">Dissociations between behavioural recovery and restoration of vestibular activity in the unilabyrinthectomized guinea-pig.</a> J Physiol. 500, 509-522 (1997).</li><li>Jin, Z., Mannstrom, P., Skjonsberg, A., Jarlebark, L., Ulfendahl, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Auditory+function+and+cochlear+morphology+in+the+German+waltzing+guinea+pig.">Auditory function and cochlear morphology in the German waltzing guinea pig.</a> Hear Res. 219 (1-2), 74-84 (2006).</li><li>Honeder, C., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Effects+of+intraoperatively+applied+glucocorticoid+hydrogels+on+residual+hearing+and+foreign+body+reaction+in+a+guinea+pig+model+of+cochlear+implantation.">Effects of intraoperatively applied glucocorticoid hydrogels on residual hearing and foreign body reaction in a guinea pig model of cochlear implantation.</a> Acta Otolaryngol. 135 (4), 313-319 (2015).</li><li>Moteki, H., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Long-term+results+of+hearing+preservation+cochlear+implant+surgery+in+patients+with+residual+low+frequency+hearing.">Long-term results of hearing preservation cochlear implant surgery in patients with residual low frequency hearing.</a> Acta Otolaryngol. 137 (5), 516-521 (2017).</li><li>Eshraghi, A. A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Clinical,+surgical,+and+electrical+factors+impacting+residual+hearing+in+cochlear+implant+surgery.">Clinical, surgical, and electrical factors impacting residual hearing in cochlear implant surgery.</a> Acta Otolaryngol. 137 (4), 384-388 (2017).</li><li>Reiss, L. A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Morphological+correlates+of+hearing+loss+after+cochlear+implantation+and+electro-acoustic+stimulation+in+a+hearing-impaired+Guinea+pig+model.">Morphological correlates of hearing loss after cochlear implantation and electro-acoustic stimulation in a hearing-impaired Guinea pig model.</a> Hear Res. 327, 163-174 (2015).</li><li>Chang, M. Y., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+Effect+of+Systemic+Steroid+on+Hearing+Preservation+After+Cochlear+Implantation+via+Round+Window+Approach:+A+Guinea+Pig+Model.">The Effect of Systemic Steroid on Hearing Preservation After Cochlear Implantation via Round Window Approach: A Guinea Pig Model.</a> Otol Neurotol. 38 (7), 962-969 (2017).</li><li>Eshraghi, A. A., Yang, N. W., Balkany, T. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Comparative+study+of+cochlear+damage+with+three+perimodiolar+electrode+designs.">Comparative study of cochlear damage with three perimodiolar electrode designs.</a> Laryngoscope. 113 (3), 415-419 (2003).</li></ol>

Christoph Arnoldner and Clemens Honeder are the holders of a research grant from MED-EL Austria. The electrodes used in this publication were provided by MED-EL Austria. The remaining authors have nothing to disclose.

The presented protocol describes how to perform cochlear implantation in the guinea pig model. This protocol can be used to evaluate different interventions for their effects on residual hearing and foreign body reaction to the CI electrode. Several precautions should be taken to achieve a high reproducibility and accuracy of the experiments.
Baseline hearing thresholds of all guinea pigs should be measured preoperatively using e.g. auditory brainstem responses. Some of the commercially available guinea pigs exhibit a relevant hearing loss and should therefore not be included in the experimental cohort. Depending on the length of the surgery and protocol this evaluation can be performed either immediately before surgery or a few days before cochlear implantation, giving the animal sufficient time to recover from anesthesia.
When performing surgery under general anesthesia in a spontaneously breathing animal, speed is important. Therefore, meticulous preparation before surgery is essential, as is the choice of the anesthetic protocol. The use of ketamine, medetomidine, midazolam and fentanyl in combination with local anesthesia results in a sufficient anesthesia and analgesia, while at the same time the animal continues to breathe spontaneously. Compared to the often-described use of ketamine and xylazine, this regimen results in better analgesia and reduced perioperative morbidity and mortality. It is important to have all instruments and medications (including a booster of the anesthetics) readily available before putting the animal to sleep.
Due to position changes of the animals during the surgery (changing from prone position to sideward position and back), there is a risk of aspiration of the stomach content in to the lungs. For this reason, the protocol also includes the application of a stomach tube, which is a fast and easy way to protect the animal from aspiration and reduce the perioperative mortality.
To maintain sterility during re-positioning, the areas where the animal is touched need to be covered by sterile drapes, gloves need to be changed thereafter or re-positioning needs to be done by another individual that is not sterile.
O2-saturation monitoring is also of utmost importance during the surgery. The positioning of the head required for visualization of the promontory and round window niche can cause an obstruction of the airway, which can easily be handled when identified early enough.
Usually the animals lose a great amount of body fluids (e.g. blood, extra cellular fluid, urine) during the surgery. Therefore, the fluid substitution protocol introduced in this manuscript represents a well-tolerated method to stabilize the hemodynamics of the animals and supports their fast recovery from anesthesia.
In order to avoid mistakes when performing audiometrical measurements, it is recommended to connect the same pin of the connector to a specific electrode during each surgery.
One limitation of this method is the relatively high variability in postoperative hearing threshold shifts, which often do not correlate well with the surgeon&#39;s perception. Even though this variability in outcomes resembles the situation in human CI recipients with residual hearing, it is not fully understood what the causes of the variable results are.16,17,18 In general, the variability decreases with time and the experience of the surgeon. It is important to avoid excessive forces when inserting the electrode, which can be achieved by a slow insertion speed. Because the careful insertion of a CI electrode can result in only very limited hearing loss, the presented protocol describes a repeated insertion of the electrode, which causes a higher and more predictable hearing loss. This hearing loss is most pronounced in the high frequency area, between 16 and 32 kHz. As the intracochlear trauma depends on the insertion depth, the morphology of the cochlea and the approach (cochleostomy versus round window insertion) need to be taken into account. Insertion of the CI electrode through the round window membrane, usually performed in human hearing preservation cochlear implantation, has also been used in the guinea pig model.19 Because the round window membrane is hidden in the guinea pig and electrode insertion through the round window membrane results in an unfavorable insertion angle, drilling a cochleostomy leads to more predictable hearing threshold shifts. This protocol proposes the use of a scalpel instead of a drill for the opening of the tympanic bulla, because this results in a reduced noise exposure of the ear to be implanted. A histological evaluation of the inner ears addressing the foreign body reaction to the electrode, the amount of hair cells and spiral ganglion cells as well as trauma to structures like the osseous spiral lamina and electrode translocation rates should be performed in all implanted ears, as these results facilitate better understanding of the functional results measured.12,20

More than 500 million people worldwide suffer from hearing loss.1 Impaired hearing has been linked to a higher rate of depression, lower self-esteem and lower feelings of self-worth, which all lead to reduced quality of life.2 While hearing aids are an adequate way to restore sensory function in cases of moderate hearing loss, the most effective treatment modality for patients suffering from profound hearing loss is the cochlear implant (CI). Due to the excellent outcomes with respect to speech perception, candidacy criteria for cochlear implantation now also include<s>s</s> patients who have substantial residual hearing in the low frequency region, but do not benefit from hearing aids.3 Since these patients can use combined electric and acoustic stimulation in the implanted ear, hearing preservation has become a major issue for CI surgeons. During cochlear implantation, an electrode array is inserted into the scala tympani of the cochlea, where it electrically stimulates the auditory nerve.4 The electrode insertion trauma poses a risk to residual hearing and induces fibrosis, which increases electrode impedances and battery consumption of the implant. Thus, models to study pharmaceutical interventions that can reduce the hearing loss and fibrosis caused by the insertion of the electrode are essential.
The guinea pig is a suitable and convenient animal model for CIs, because of the easier and more reproducible surgical access to the inner ear as compared to gerbils, rats or mice.5,6,7,8 Furthermore, the hearing range of this species is relatively comparable to human hearing.9 Larger species like cats or monkeys, which have been used to address specific research questions related to CIs, do not represent a reasonable choice for most CI studies due to both ethical and financial considerations.10,11
In summary, the guinea pig is a reliable and relatively cost-efficient model to evaluate the effects of pharmacological interventions in the setting of cochlear implantation.

<table><tbody><tr><td>Scale</td><td></td><td></td><td></td></tr><tr><td>Oxygen insufflator</td><td></td><td></td><td></td></tr><tr><td>Shaver</td><td></td><td></td><td></td></tr><tr><td>Sucker</td><td></td><td></td><td></td></tr><tr><td>Povidone Iodine Solution</td><td></td><td></td><td></td></tr><tr><td>Alcohol</td><td></td><td></td><td></td></tr><tr><td>Laryngoscope</td><td></td><td></td><td></td></tr><tr><td>Stomach tube&amp;nbsp;</td><td></td><td></td><td>Fr 06, Lg 80 cm</td></tr><tr><td>Surgical binocular microscope</td><td></td><td></td><td></td></tr><tr><td>Drill</td><td></td><td></td><td></td></tr><tr><td>0.5 mm diamond burr</td><td></td><td></td><td></td></tr><tr><td>1 mm diamond burr</td><td></td><td></td><td></td></tr><tr><td>Heating plate</td><td></td><td></td><td></td></tr><tr><td>Pulse oximeter</td><td></td><td></td><td></td></tr><tr><td>Tissue glue</td><td></td><td></td><td></td></tr><tr><td>Dental cement powder</td><td></td><td></td><td></td></tr><tr><td>Fluid for dental cement powder</td><td></td><td></td><td></td></tr><tr><td>Bipolar cautery</td><td></td><td></td><td></td></tr><tr><td>Gauze compress</td><td></td><td></td><td></td></tr><tr><td>Cotton bud</td><td></td><td></td><td></td></tr><tr><td>Cement mixing bowl</td><td></td><td></td><td></td></tr><tr><td>Teflon insulated gold wire</td><td></td><td></td><td>99.99 % gold, diameter: 0.125 mm, isolation: 0,016 mm, PTFE (Polytetrafluoroethylen)&amp;nbsp;</td></tr><tr><td>Scalpel with blade No. 10</td><td></td><td></td><td></td></tr><tr><td>Scalpel with blade No. 15</td><td></td><td></td><td></td></tr><tr><td>Scissors</td><td></td><td></td><td></td></tr><tr><td>Mosquito forceps</td><td></td><td></td><td></td></tr><tr><td>Dressing forceps</td><td></td><td></td><td></td></tr><tr><td>Tissue forceps</td><td></td><td></td><td></td></tr><tr><td>Delicate dressing forceps 2X</td><td></td><td></td><td></td></tr><tr><td>Micro forceps</td><td></td><td></td><td></td></tr><tr><td>Screw driver</td><td></td><td></td><td></td></tr><tr><td>Stainless steel screws</td><td></td><td></td><td>diameter: 1 mm</td></tr><tr><td>Retractor</td><td></td><td></td><td></td></tr><tr><td>Needle probe</td><td></td><td></td><td></td></tr><tr><td>Spatula</td><td></td><td></td><td></td></tr><tr><td>Needle holder</td><td></td><td></td><td></td></tr><tr><td>5-0 absorbable sutures&amp;nbsp;</td><td></td><td></td><td></td></tr><tr><td>Needle 23G</td><td></td><td></td><td></td></tr><tr><td>Needle 27G</td><td></td><td></td><td></td></tr><tr><td>Medetomidine 1 mg/mL</td><td></td><td></td><td>0.36 mg/kg</td></tr><tr><td>Midazolam g mg/mL</td><td></td><td></td><td>1.2 mg/kg</td></tr><tr><td>Fentanyl 50 &amp;micro;g/mL</td><td></td><td></td><td>0.036 mg/kg</td></tr><tr><td>Ketamine 100 mg/mL</td><td></td><td></td><td>12 mg/kg</td></tr><tr><td>Lidocaine (local anesthesia)</td><td></td><td></td><td>4 mg/kg</td></tr><tr><td>Atipamezole 5 mg/mL</td><td></td><td></td><td>1 mg/kg</td></tr><tr><td>Flumazenil 0.1 mg/mL</td><td></td><td></td><td>0.1 mg/kg</td></tr><tr><td>Enrofloxacin 100 mg/mL</td><td></td><td></td><td>7 mg/kg</td></tr><tr><td>Buprenorphin&amp;nbsp; 0.3 mg/mL</td><td></td><td></td><td>0.05 mg/kg&amp;nbsp;</td></tr><tr><td>Physiological Saline (at body temperature)</td><td></td><td></td><td>12.5 mL/Kg (pre-surgery)</td></tr><tr><td>Glucose 5 % (preoperative, at body temperature)</td><td></td><td></td><td>12.5 mL/Kg</td></tr><tr><td>Physiological Saline (at body temperature)</td><td></td><td></td><td>25 mL/kg (post-surgery)</td></tr></tbody></table>

cochlear implantation in the guinea pig

Cochlear implants are highly efficient devices that can restore hearing in subjects with profound hearing loss. Due to improved speech perception outcomes, candidacy criteria have been expanded over the last few decades. This includes patients with substantial residual hearing that benefit from electrical and acoustical stimulation of the same ear, which makes hearing preservation during cochlear implantation an important issue. Electrode impedances and the related issue of energy consumption is another major research field, as progress in this area could pave the way for fully implantable auditory prostheses. To address these issues in a systematic way, adequate animal models are essential. Therefore, the goal of this protocol is to provide an animal model of cochlear implantation, which can be used to address various research questions. Due to its large tympanic bulla, which allows easy surgical access to the inner ear, as well as its hearing range which is relatively similar to the hearing range of humans, the guinea pig is a commonly used species in auditory research. Cochlear implantation in the guinea pig is performed via a retroauricular approach. Through the bullostomy a cochleostomy is drilled and the cochlear implant electrode is inserted into the scala tympani. This electrode can then be used for electrical stimulation, determination of electrode impedances and the measurement of compound action potentials of the auditory nerve. In addition to these applications, cochlear implant electrodes can also be used as drug delivery devices, if a topical delivery of pharmaceutical agents to the cells or fluids of the inner ear is intended.

Usually surgical wounds heal fast and without complications in the guinea pig model and the contacts for postoperative electrophysiological measurements remain easily accessible at the vertex of the animal (Figure 3). Figure 4 shows the pre- and postoperative click-CAP measurement of a representative animal. Electrode insertion resulted in a threshold shift of 16 decibels (dB) (Figures 4A and 4B). Figure 4C illustrates the pre- and postoperative frequency-specific CAP thresholds of the same animal. CAP thresholds are almost unchanged in the low frequencies, whereas a threshold shift of approximately 25 to 30 dB is achieved in the high frequency area, starting at 8 kHz.
Electrode insertion can cause trauma to the inner ear. In addition to the acute surgical trauma, the foreign body reaction to the electrode negatively impacts cochlear implant performance. Figure 5 demonstrates the cochleae of guinea pigs after CI insertion and different histological procedures. In Figure 5A the electrode, which is positioned correctly in the scala tympani, was left in situ, whereas in Figure 5B the electrode was removed before histological workup. In Figure 5A almost no foreign body reaction is visible, whereas in Figure 5B a large area of the scala tympani is filled with fibrotic tissue. Figure 5C depicts the fracture of the osseous spiral lamina due to CI electrode insertion, which also caused a loss of spiral ganglion cells in this animal. Such fractures can explain higher than expected threshold shifts in some animals.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/56829/56829fig1.jpg" /> 
Figure 1: Round window area with gold wire in situ. An asterisk marks the round window, an x the basal turn of the cochlea. The gold wire is marked by an arrow. Scale bar 2 mm. <a href="/files/ftp_upload/56829/56829fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/56829/56829fig2.jpg" /> 
Figure 2: Guinea pig cochlear implant electrode. The electrode with two contacts is inserted for 4 mm. The diameter of the electrode is tapered from 0.3 mm at the tip to 0.5 mm. Lines indicate 0.5 mm. <a href="/files/ftp_upload/56829/56829fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/56829/56829fig3.jpg" /> 
Figure 3: Guinea pig approximately two weeks after cochlear implantation. The CI electrode is in situ and the contacts for electrophysiological measurements are easily accessible at the vertex of the animal. <a href="/files/ftp_upload/56829/56829fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/56829/56829fig4.jpg" /> 
Figure 4: Representative CAP thresholds (A) Preoperative click CAP threshold of a representative animal. (B) Postoperative click CAP threshold of the same animal, exhibiting a threshold shift of 16 dB. Lines indicate 10 dB. (C) Pre- and postoperative frequency-specific CAP thresholds. While low frequencies are almost unchanged, a threshold shift of 25-30 dB can be observed in the frequency range of 8 to 32 kHz. <a href="/files/ftp_upload/56829/56829fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/56829/56829fig5.jpg" /> 
Figure 5: Potential local consequences of CI electrode insertion. (A) Micrograph of a basal turn of a guinea pig cochlea with the CI electrode in situ (#) and only minimal foreign body reaction. Histological analysis was performed using a grinding and polishing technique after resin embedding and Giemsa staining.15 Scalebar 100 &#181;m (B) Micrograph of the tympanic duct of the upper basal turn of the cochlea with visible tissue response leaving a canal after removal of the CI electrode (#). Scalebar 100 &#181;m (C) Lower basal turn of cochlea with fractured osseous spiral lamina (bold arrow) and adjacent tissue response: (i) loss of spiral ganglion cells (arrow) in Rosenthal`s canal (ii) fibrosis and osteoneogenesis in vestibular duct (&#9679;), and (iii) loss of the organ of Corti (*). Drilling-hole for insertion of CI (&#9675;) with adjacent osteoneogenesis. Scalebar 500 &#181;m. Figures 5B and 5C were stained with hematoxylin (blue) &#38; eosin (red). <a href="/files/ftp_upload/56829/56829fig5large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Cochlear Implantation in the Guinea Pig at JoVE.com

Cochlear Implantation in the Guinea Pig

The goal of this protocol is to provide an animal model of cochlear implantation, which can be used to address a multitude of research questions. Potential applications include the evaluation of pharmaceutical interventions or electrical stimulation for beneficial effects on hearing thresholds or electrode impedances.

Cochlear implants are highly efficient devices that can restore hearing in subjects with profound hearing loss. Due to improved speech perception ...

Universidad San Sebastian

Research

JoVE Journal

Medicine

11.1K Views.  Medical University of Vienna. The goal of this protocol is to provide an animal model of cochlear implantation, which can be used to address a multitude of research questions. Potential applications include the evaluation of pharmaceutical interventions or electrical stimulation for beneficial effects on hearing thresholds or electrode impedances.

Video: Cochlear Implantation in the Guinea Pig

Surgical Induction of Endolymphatic Hydrops by Obliteration of the Endolymphatic Duct

Surgical induction of endolymphatic hydrops (ELH) in the guinea pig by obliteration and obstruction of the endolymphatic duct is a well-accepted animal model of the condition and an important correlate for human Meniere's disease. In 1965, Robert Kimura and Harold Schuknecht first described an intradural approach for obstruction of the endolymphatic duct (Kimura 1965). Although effective, this technique, which requires penetration of the brain's protective covering, incurred an undesirable level of morbidity and mortality in the animal subjects. Consequently, Andrews and Bohmer developed an extradural approach, which predictably produces fewer of the complications associated with central nervous system (CNS) penetration.(Andrews and Bohmer 1989) 
The extradural approach described here first requires a midline incision in the region of the occiput to expose the underlying muscular layer. We operate only on the right side. After appropriate retraction of the overlying tissue, a horizontal incision is made into the musculature of the right occiput to expose the right temporo-occipital suture line. The bone immediately inferio-lateral the suture line (Fig 1) is then drilled with an otologic drill until the sigmoid sinus becomes visible. Medial retraction of the sigmoid sinus reveals the operculum of the endolymphatic duct, which houses the endolymphatic sac. Drilling medial to the operculum into the area of the endolymphatic sac reveals the endolymphatic duct, which is then packed with bone wax to produce obstruction and ultimately ELH. 
In the following weeks, the animal will demonstrate the progressive, fluctuating hearing loss and histologic evidence of ELH.

This video shows how to surgically obstruct the guinea pig's endolymphatic duct to produce endolymphatic hydrops.

Surgical induction of endolymphatic hydrops (ELH) in the guinea pig by obliteration and obstruction of the endolymphatic duct is a well-accepted animal ...

Biology

Cochlear Implant Surgery and Electrically-evoked Auditory Brainstem Response Recordings in C57BL/6 Mice

Cochlear implants (CIs) are neuroprosthetic devices that can provide a sense of hearing to deaf people. However, a CI cannot restore all aspects of hearing. Improvement of the implant technology is needed if CI users are to perceive music and perform in more natural environments, such as hearing out a voice with competing talkers, reflections, and other sounds. Such improvement requires experimental animals to better understand the mechanisms of electric stimulation in the cochlea and its responses in the whole auditory system. The mouse is an increasingly attractive model due to the many genetic models available. However, the limited use of this species as a CI model is mainly due to the difficulty of implanting small electrode arrays. More details about the surgical procedure are therefore of great interest to expand the use of mice in CI research.
In this report, we describe in detail the protocol for acute deafening and cochlear implantation of an electrode array in the C57BL/6 mouse strain. We demonstrate the functional efficacy of this procedure with electrically-evoked auditory brainstem response (eABR) and show examples of facial nerve stimulation. Finally, we also discuss the importance of including a deafening procedure when using a normally hearing animal. This mouse model provides a powerful opportunity to study genetic and neurobiological mechanisms that would be of relevance for CI users.

Animal models of cochlear implants can advance knowledge of the technological bases of treating permanent sensorineural hearing loss with electrical stimulation. This study presents a surgical protocol for acute deafening and cochlear implantation of an electrode array in mice as well as the functional assessment with auditory brainstem response.

Cochlear implants (CIs) are neuroprosthetic devices that can provide a sense of hearing to deaf people. However, a CI cannot restore all aspects of ...

Neuroscience

Subretinal Implantation of RPE on a Carrier in Minipigs: Guidelines for Preoperative Preparations, Surgical Techniques, and Postoperative Care

Degenerative disorders of the retina (including age-related macular degeneration), which originate primarily at or within the retinal pigmented epithelial (RPE) layer, lead to a progressive disorganization of the retinal anatomy and the deterioration of visual function. The substitution of damaged RPE cells (RPEs) with in vitro cultured RPE cells using a subretinal cell carrier has shown potential for re-establishing the anatomical structure of the outer retinal layers and is, therefore, being further studied. Here, we present the principles of a surgical technique that allows for the effective subretinal transplantation of a cell carrier with cultivated RPEs into minipigs. The surgeries were performed under general anesthesia and included a standard lens-sparing three-port pars plana vitrectomy (PPV), subretinal application of a balanced salt solution (BSS), a 2.7 mm retinotomy, implantation of a nanofibrous cell carrier into the subretinal space through an additional 3.0 mm sclerotomy, fluid-air exchange (FAX), silicone oil tamponade, and closure of all the sclerotomies. This surgical approach was used in 29 surgeries (18 animals) over the past 8 years with a success rate of 93.1%. Anatomic verification of the surgical placement was carried out using in vivo fundus imaging (fundus photography and optical coherence tomography). The recommended surgical steps for the subretinal implantation of RPEs on a carrier in minipig eyes can be used in future preclinical studies using large-eye animal models.

The subretinal implantation of retinal pigmented epithelium (RPE) is one of the most promising approaches for the treatment of degenerative retinal diseases. However, the performance of preclinical studies on large-eye animal models remains challenging. This report presents guidelines for the subretinal transplantation of RPE on a cell carrier into minipigs.

Degenerative disorders of the retina (including age-related macular degeneration), which originate primarily at or within the retinal pigmented epithelial ...

Bioengineering

Subdural Soft Electrocorticography (ECoG) Array Implantation and Long-Term Cortical Recording in Minipigs

Neurological impairments and diseases can be diagnosed or treated using electrocorticography (ECoG) arrays. In drug-resistant epilepsy, these help delineate the epileptic region to resect. In long-term applications such as brain-computer interfaces, these epicortical electrodes are used to record the movement intention of the brain, to control the robotic limbs of paralyzed patients. However, current stiff electrode grids do not answer the need for high-resolution brain recordings and long-term biointegration. Recently, conformable electrode arrays have been proposed to achieve long-term implant stability with high performance. However, preclinical studies for these new implant technologies are needed to validate their long-term functionality and safety profile for their translation to human patients. In this context, porcine models are routinely employed in developing medical devices due to their large organ sizes and easy animal handling. However, only a few brain applications are described in the literature, mostly due to surgery limitations and integration of the implant system on a living animal.
Here, we report the method for long-term implantation (6 months) and evaluation of soft ECoG arrays in the minipig model. The study first presents the implant system, consisting of a soft microfabricated electrode array integrated with a magnetic resonance imaging (MRI)-compatible polymeric transdermal port that houses instrumentation connectors for electrophysiology recordings. Then, the study describes the surgical procedure, from subdural implantation to animal recovery. We focus on the auditory cortex as an example target area where evoked potentials are induced by acoustic stimulation. We finally describe a data acquisition sequence that includes MRI of the whole brain, implant electrochemical characterization, intraoperative and freely moving electrophysiology, and immunohistochemistry staining of the extracted brains.
This model can be used to investigate the safety and function of novel design of cortical prostheses; mandatory preclinical study to envision translation to human patients.

Subdural Soft Electrocorticography ECoG Array Implantation and Long-Term Cortical Recording in Minipigs

Here, we present a method for the long-term performance and safety assessment of soft subdural electrode arrays in a minipig model, describing surgical method and tools, postoperative magnetic resonance imaging, electrophysiology of the auditory cortex, electrochemical properties of the implant, and postmortem immunochemistry.

Neurological impairments and diseases can be diagnosed or treated using electrocorticography (ECoG) arrays. In drug-resistant epilepsy, these help ...

A Revised Surgical Approach to Induce Endolymphatic Hydrops in the Guinea Pig

Endolymphatic hydrops is an enlargement of scala media that is most often associated with Meniere&#39;s disease, though the pathophysiologic mechanism(s) remain unclear. In order to adequately study the attributes of endolymphatic hydrops, such as the origins of low-frequency hearing loss, a reliable model is needed. The guinea pig is a&#160;good model because it hears in the low-frequency regions that are putatively affected by endolymphatic hydrops. Previous research has demonstrated that endolymphatic hydrops can be induced surgically via intradural or extradural approaches that involve drilling on the endolymphatic duct and sac. However, whether it was possible to create an endolymphatic hydrops model using an extradural approach that avoided dangerous drilling on the endolymphatic duct and sac was unknown. The objective of this study was to demonstrate a revised extradural approach to induce experimental endolymphatic hydrops at 30 days post-operatively by obliterating the endolymphatic sac and injuring the endolymphatic duct with a fine pick. The sample size consisted of seven guinea pigs. Functional measurements of hearing were made and temporal bones were subsequently harvested for histologic analysis. The approach had a success rate of 86% in achieving endolymphatic hydrops. The risk of cerebrospinal fluid leak was minimal. No perioperative deaths or injuries to the posterior semicircular canal occurred in the sample. The presented method demonstrates a safe and reliable way to induce endolymphatic hydrops at a relatively quick time point of 30 days. The clinical implications are that the presented method provides a reliable model to further explore the origins of low-frequency hearing loss that can be associated endolymphatic hydrops.

This article demonstrates an extradural approach to obliterate the guinea pig endolymphatic sac and injure the endolymphatic duct with a fine pick in order to induce experimental endolymphatic hydrops.

Endolymphatic hydrops is an enlargement of scala media that is most often associated with Meniere&#39;s disease, though the pathophysiologic mechanism(s) ...

Implantation of hiPSC-derived Cardiac-muscle Patches after Myocardial Injury in a Guinea Pig Model

Due to the limited regeneration capacity of the heart in adult mammals, myocardial infarction results in an irreversible loss of cardiomyocytes. This loss of relevant amounts of heart muscle mass can lead to the heart failure. Besides heart transplantation, there is no curative treatment option for the end-stage heart failure. In times of organ donor shortage, organ independent treatment modalities are needed. Left-ventricular assist devices are a promising therapy option, however, especially as destination therapy, limited by its side-effects like stroke, infections and bleedings. In recent years, several cardiac repair strategies including stem cell injection, cardiac progenitors or myocardial tissue engineering have been investigated. Recent improvements in cell biology allow for the differentiation of large amounts of cardiomyocytes derived from human induced pluripotent stem cells (iPSC). One of the cardiac repair strategies currently under evaluation is to transplant artificial heart tissue. Engineered heart tissue (EHT) is a three-dimensional in vitro created cardiomyocyte network, with functional properties of native heart tissue. We have created EHT-patches from hiPSC derived cardiomyocytes. Here we present a protocol for the induction of left ventricular myocardial cryoinjury in a guinea pig, followed by implantation of hiPSC derived EHT on the left ventricular wall.

Here we present a protocol for the induction of left ventricular cryoinjury followed by the implantation of a cardiac muscle patch, derived from human iPS-cell cardiomyocytes in a guinea pig model.

Due to the limited regeneration capacity of the heart in adult mammals, myocardial infarction results in an irreversible loss of cardiomyocytes. This loss ...

Pupillometry to Assess Auditory Sensation in Guinea Pigs

Noise exposure is a leading cause of sensorineural hearing loss. Animal models of noise-induced hearing loss have generated mechanistic insight into the underlying anatomical and physiological pathologies of hearing loss. However, relating behavioral deficits observed in humans with hearing loss to behavioral deficits in animal models remains challenging. Here, pupillometry is proposed as a method that will enable the direct comparison of animal and human behavioral data. The method is based on a modified oddball paradigm - habituating the subject to the repeated presentation of a stimulus and intermittently presenting a deviant stimulus that varies in some parametric fashion from the repeated stimulus. The fundamental premise is that if the change between the repeated and deviant stimulus is detected by the subject, it will trigger a pupil dilation response that is larger than that elicited by the repeated stimulus. This approach is demonstrated using a vocalization categorization task in guinea pigs, an animal model widely used in auditory research, including in hearing loss studies. By presenting vocalizations from one vocalization category as standard stimuli and a second category as oddball stimuli embedded in noise at various signal-to-noise ratios, it is demonstrated that the magnitude of pupil dilation in response to the oddball category varies monotonically with the signal-to-noise ratio. Growth curve analyses can then be used to characterize the time course and statistical significance of these pupil dilation responses. In this protocol, detailed procedures for acclimating guinea pigs to the setup, conducting pupillometry, and evaluating/analyzing data are described. Although this technique is demonstrated in normal-hearing guinea pigs in this protocol, the method may be used to assess the sensory effects of various forms of hearing loss within each subject. These effects may then be correlated with concurrent electrophysiological measures and post-hoc anatomical observations.

Pupillometry, a simple and non-invasive technique, is proposed as a method to determine hearing-in-noise thresholds in normal hearing animals and animal models of various auditory pathologies.

Noise exposure is a leading cause of sensorineural hearing loss. Animal models of noise-induced hearing loss have generated mechanistic insight into the ...

The Miniature Pig: A Large Animal Model for Cochlear Implant Research

Cochlear implants (CI) are the most effective method to treat people with severe-to-profound sensorineural hearing loss. Although CIs are used worldwide, no standard model exists for investigating the electrophysiology and histopathology in patients or animal models with a CI or for evaluating new models of electrode arrays. A large animal model with cochlea characteristics similar to those of humans may provide a research and evaluation platform for advanced and modified arrays before their use in humans. 
To this end, we established standard CI methods with Bama mini-pigs, whose inner ear anatomy is highly similar to that of humans. Arrays designed for human use were implanted into the mini pig cochlea through a round window membrane, and a surgical approach followed that was similar to that used for human CI recipients. Array insertion was followed by evoked compound action potential (ECAP) measurements to evaluate the function of the auditory nerve. This study describes the preparation of the animal, surgical steps, array insertion, and intraoperative electrophysiological measurements. 
The results indicated that the same CI used for humans could be easily implanted in mini-pigs via a standardized surgical approach and yielded similar electrophysiological outcomes as measured in human CI recipients. Mini-pigs could be a valuable animal model to provide initial evidence of the safety and potential performance of novel electrode arrays and surgical approaches before applying them to human beings.

Miniature pigs (mini-pigs) are an ideal large animal model for research into cochlear implants. Cochlear implantation surgery in mini-pigs can be utilized to provide initial evidence of the safety and potential performance of novel electrode arrays and surgical approaches in a living system similar to human beings.

Cochlear implants (CI) are the most effective method to treat people with severe-to-profound sensorineural hearing loss. Although CIs are used worldwide, ...

Endoscopic Balloon Dilatation of the Eustachian Tube via the Soft Palate Approach in Miniature Pigs

The eustachian tube (ET) is one of the most complex organs in the human body, and its dysfunction may lead to a variety of diseases. In recent years, an increasing number of scholars have opted to conduct ET-related studies using large experimental animals such as miniature pigs or sheep, yielding promising results. Typically, conventional endoscopic procedures are performed through the nasal approach for large experimental animals. However, due to the elongated and narrow nasal cavity in these animals, transnasal surgeries are challenging. 
To address this issue, we explored an ET surgery approach via the soft palate. The animal was placed in a supine position. After endotracheal intubation under general anesthesia, a mouth opener was used to fully expose the upper palate. Local infiltration with diluted adrenal fluid was performed for anesthesia of the area. A sickle knife was then used to make a longitudinal soft palate incision at the junction of the soft and hard palates. After hemostasis, an endoscope was inserted into the nasopharynx cavity, allowing the visualization of the pharyngeal opening of the ET on the posterior lateral wall of the nasal cavity. 
Subsequently, a specialized pusher was used to insert a balloon into ET. The balloon was inflated, maintained at 10 bar for 2 min, and then removed. The incision in the soft palate was then sutured to ensure proper alignment. The soft palate healed well after the operation. This surgical approach is suitable for ET-related procedures in large experimental animals (e.g., miniature pigs, sheep, and dogs). The surgical procedure is simple, with a short surgical time, and wound healing is rapid. Under endoscopy, the pharyngeal opening of the ET is visible, and it is thus a good choice for procedures such as balloon dilation of the ET.

In this study, we present a protocol to explore an eustachian tube surgery approach via the soft palate in miniature pigs. The surgical procedure is simple, with short surgical time, and wound healing is rapid; it is thus a good choice for procedures such as balloon dilation of the eustachian tube.

The eustachian tube (ET) is one of the most complex organs in the human body, and its dysfunction may lead to a variety of diseases. In recent years, an ...

Guinea Pig Round Window Membrane Explantation

Guinea Pig Round Window Membrane Explantation for Ex Vivo Studies

Efficient and minimally invasive drug delivery to the inner ear is a significant challenge. The round window membrane (RWM), being one of the few entry points to the inner ear, has become a vital focus of investigation. However, due to the complexities of isolating the RWM, our understanding of its pharmacokinetics remains limited. The RWM comprises three distinct layers: the outer epithelium, the middle connective tissue layer, and the inner epithelial layer, each potentially possessing unique delivery properties.
Current models for investigating transport across the RWM utilize in vivo animal models or ex vivo RWM models which rely on cell cultures or membrane fragments. Guinea pigs serve as a validated preclinical model for the investigation of drug pharmacokinetics within the inner ear and are an important animal model for the translational development of delivery vehicles to the cochlea. In this study, we describe an approach for explantation of a guinea pig RWM with surrounding cochlear bone for benchtop drug delivery experiments. This method allows for preservation of native RWM architecture and may provide a more realistic representation of barriers to transport than current benchtop models.

Author Spotlight: Extraction of Guinea Pig Round Window Membrane to Facilitate Inner Ear Drug Delivery Research

This protocol outlines a method for the explantation of the round window membrane from guinea pig temporal bones, providing a valuable resource for ex vivo studies.

Efficient and minimally invasive drug delivery to the inner ear is a significant challenge. The round window membrane (RWM), being one of the few entry ...