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. 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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 border="0" cellpadding="0" cellspacing="0" style="width:1104px;" width="1103">
	<colgroup>
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		<col />
		<col />
		<col />
	</colgroup>
	<tbody>
		<tr height="36">
			<td height="36" style="height:36px;width:308px;">Scale</td>
			<td style="width:155px;"></td>
			<td style="width:123px;"></td>
			<td style="width:519px;"></td>
		</tr>
		<tr height="36">
			<td height="36" style="height:36px;">Oxygen insufflator</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr height="36">
			<td height="36" style="height:36px;">Shaver</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr height="36">
			<td height="36" style="height:36px;">Sucker</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr height="36">
			<td height="36" style="height:36px;">Povidone Iodine Solution</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr height="36">
			<td height="36" style="height:36px;">Alcohol</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr height="36">
			<td height="36" style="height:36px;">Laryngoscope</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr height="36">
			<td height="36" style="height:36px;">Stomach tube&#160;</td>
			<td></td>
			<td></td>
			<td>Fr 06, Lg 80 cm</td>
		</tr>
		<tr height="36">
			<td height="36" style="height:36px;">Surgical binocular microscope</td>
			<td></td>
			<td></td>
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		</tr>
		<tr height="36">
			<td height="36" style="height:36px;">Drill</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr height="36">
			<td height="36" style="height:36px;">0.5 mm diamond burr</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr height="36">
			<td height="36" style="height:36px;">1 mm diamond burr</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr height="36">
			<td height="36" style="height:36px;">Heating plate</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr height="36">
			<td height="36" style="height:36px;">Pulse oximeter</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr height="36">
			<td height="36" style="height:36px;">Tissue glue</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr height="36">
			<td height="36" style="height:36px;">Dental cement powder</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr height="36">
			<td height="36" style="height:36px;">Fluid for dental cement powder</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr height="36">
			<td height="36" style="height:36px;">Bipolar cautery</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr height="36">
			<td height="36" style="height:36px;">Gauze compress</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr height="36">
			<td height="36" style="height:36px;">Cotton bud</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr height="36">
			<td height="36" style="height:36px;">Cement mixing bowl</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr height="36">
			<td height="36" style="height:36px;">Teflon insulated gold wire</td>
			<td></td>
			<td></td>
			<td>99.99 % gold, diameter: 0.125 mm, isolation: 0,016 mm, PTFE (Polytetrafluoroethylen)&#160;</td>
		</tr>
		<tr height="36">
			<td height="36" style="height:36px;">Scalpel with blade No. 10</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr height="36">
			<td height="36" style="height:36px;">Scalpel with blade No. 15</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr height="36">
			<td height="36" style="height:36px;">Scissors</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr height="36">
			<td height="36" style="height:36px;">Mosquito forceps</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr height="36">
			<td height="36" style="height:36px;">Dressing forceps</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr height="36">
			<td height="36" style="height:36px;">Tissue forceps</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr height="36">
			<td height="36" style="height:36px;">Delicate dressing forceps 2X</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr height="36">
			<td height="36" style="height:36px;">Micro forceps</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr height="36">
			<td height="36" style="height:36px;">Screw driver</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr height="36">
			<td height="36" style="height:36px;">Stainless steel screws</td>
			<td></td>
			<td></td>
			<td>diameter: 1 mm</td>
		</tr>
		<tr height="36">
			<td height="36" style="height:36px;">Retractor</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr height="36">
			<td height="36" style="height:36px;">Needle probe</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr height="36">
			<td height="36" style="height:36px;">Spatula</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr height="36">
			<td height="36" style="height:36px;">Needle holder</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr height="36">
			<td height="36" style="height:36px;">5-0 absorbable sutures&#160;</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr height="36">
			<td height="36" style="height:36px;">Needle 23G</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr height="36">
			<td height="36" style="height:36px;">Needle 27G</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr height="36">
			<td height="36" style="height:36px;">Medetomidine 1 mg/mL</td>
			<td></td>
			<td></td>
			<td>0.36 mg/kg</td>
		</tr>
		<tr height="36">
			<td height="36" style="height:36px;">Midazolam g mg/mL</td>
			<td></td>
			<td></td>
			<td>1.2 mg/kg</td>
		</tr>
		<tr height="36">
			<td height="36" style="height:36px;">Fentanyl 50 &#181;g/mL</td>
			<td></td>
			<td></td>
			<td>0.036 mg/kg</td>
		</tr>
		<tr height="36">
			<td height="36" style="height:36px;">Ketamine 100 mg/mL</td>
			<td></td>
			<td></td>
			<td>12 mg/kg</td>
		</tr>
		<tr height="36">
			<td height="36" style="height:36px;">Lidocaine (local anesthesia)</td>
			<td></td>
			<td></td>
			<td>4 mg/kg</td>
		</tr>
		<tr height="36">
			<td height="36" style="height:36px;">Atipamezole 5 mg/mL</td>
			<td></td>
			<td></td>
			<td>1 mg/kg</td>
		</tr>
		<tr height="36">
			<td height="36" style="height:36px;">Flumazenil 0.1 mg/mL</td>
			<td></td>
			<td></td>
			<td>0.1 mg/kg</td>
		</tr>
		<tr height="36">
			<td height="36" style="height:36px;">Enrofloxacin 100 mg/mL</td>
			<td></td>
			<td></td>
			<td>7 mg/kg</td>
		</tr>
		<tr height="36">
			<td height="36" style="height:36px;">Buprenorphin&#160; 0.3 mg/mL</td>
			<td></td>
			<td></td>
			<td>0.05 mg/kg&#160;</td>
		</tr>
		<tr height="36">
			<td height="36" style="height:36px;">Physiological Saline (at body temperature)</td>
			<td></td>
			<td></td>
			<td>12.5 mL/Kg (pre-surgery)</td>
		</tr>
		<tr height="36">
			<td height="36" style="height:36px;">Glucose 5 % (preoperative, at body temperature)</td>
			<td></td>
			<td></td>
			<td>12.5 mL/Kg</td>
		</tr>
		<tr height="36">
			<td height="36" style="height:36px;">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 ...

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

The Polyvinyl Alcohol Sponge Model Implantation

Wound healing is a complicated, multistep process involving many cell types, growth factors and compounds1-3. Because of this complexity, wound healing studies are most comprehensive when carried out in vivo. There are many in vivo models available to study acute wound healing, including incisional, excisional, dead space, and burns. Dead space models are artificial, porous implants which are used to study tissue formation and the effects of substances on the wound. Some of the commonly used dead space models include polyvinyl alcohol (PVA) sponges, steel wire mesh cylinders, expanded polytetrafluoroethylene (ePTFE) material, and the Cellstick1,2.
Each dead space model has its own limitations based on its material's composition and implantation methods. The steel wire mesh cylinder model has a lag phase of infiltration after implantation and requires a long amount of time before granulation tissue formation begins1. Later stages of wound healing are best analyzed using the ePTFE model1,4. The Cellstick is a cellulose sponge inside a silicon tube model which is typically used for studying human surgery wounds and wound fluid2. The PVA sponge is limited to acute studies because with time it begins to provoke a foreign body response which causes a giant cell reaction in the animal5. Unlike other materials, PVA sponges are easy to insert and remove, made of inert and non-biodegradable materials and yet are soft enough to be sectioned for histological analysis2,5.
In wound healing the PVA sponge is very useful for analyzing granulation tissue formation, collagen deposition, wound fluid composition, and the effects of substances on the healing process1,2,5. In addition to its use in studying a wide array of attributes of wound healing, the PVA sponge has also been used in many other types of studies. It has been utilized to investigate tumor angiogenesis, drug delivery and stem cell survival and engraftment1,2,6,7. With its great alterability, prior extensive use, and reproducible results, the PVA sponge is an ideal model for many studies1,2.
Here, we will describe the preparation, implantation and retrieval of PVA sponge disks (Figure 1) in a mouse model of wound healing.

A useful tool to analyze the effects of drugs, growth factors, and/or manipulated cells in an animal model of wound repair is described. This technique utilizes the properties of a polyvinyl alcohol (PVA) sponge to deliver and contain the desired treatment and also provide a platform to be excised and analyzed.

Wound healing is a complicated, multistep process involving many cell types, growth factors and compounds1-3. Because of this complexity, wound healing ...

Implantation of the Syncardia Total Artificial Heart

With advances in technology, the use of mechanical circulatory support devices for end stage heart failure has rapidly increased. The vast majority of such patients are generally well served by left ventricular assist devices (LVADs). However, a subset of patients with late stage biventricular failure or other significant anatomic lesions are not adequately treated by isolated left ventricular mechanical support. Examples of concomitant cardiac pathology that may be better treated by resection and TAH replacement includes: post infarction ventricular septal defect,&#160;aortic root aneurysm / dissection, cardiac allograft failure, massive ventricular thrombus, refractory malignant arrhythmias (independent of filling pressures), hypertrophic / restrictive cardiomyopathy, and complex congenital heart disease. Patients often present with cardiogenic shock and multi system organ dysfunction. Excision of both ventricles and orthotopic replacement with a total artificial heart (TAH) is an effective, albeit extreme, therapy for rapid restoration of blood flow and resuscitation. Perioperative management is focused on end organ resuscitation and physical rehabilitation. In addition to the usual concerns of infection, bleeding, and thromboembolism common to all mechanically supported patients, TAH patients face unique risks with regard to renal failure and anemia. Supplementation of the abrupt decrease in brain natriuretic peptide following ventriculectomy appears to have protective renal effects. Anemia following TAH implantation can be profound and persistent. Nonetheless, the anemia is generally well tolerated and transfusion are limited to avoid HLA sensitization. Until recently, TAH patients were confined as inpatients tethered to a 500 lb pneumatic console driver. Recent introduction of a backpack sized portable driver (currently under clinical trial) has enabled patients to be discharged home and even return to work. Despite the profound presentation of these sick patients, there is a 79-87% success in bridge to transplantation.

The purpose of this manuscript is to briefly review indications, management, and outcomes for the total artificial heart. Video operative techniques for device implantation are presented.

With advances in technology, the use of mechanical circulatory support devices for end stage heart failure has rapidly increased. The vast majority of ...

Implantation of Total Artificial Heart in Congenital Heart Disease

In patients with end-stage heart failure (HF), a total artificial heart (TAH) may be implanted as a bridge to cardiac transplant. However, in congenital heart disease (CHD), the malformed heart presents a challenge to TAH implantation. 
In the case presented here, a 17 year-old patient with congenital transposition of the great arteries (CCTGA) experienced progressively worsening HF due to his congenital condition. He was hospitalized multiple times and received an implantable cardioverter defibrillator (ICD). However, his condition soon deteriorated to end-stage HF with multisystem organ failure. 
Due to the patient's grave clinical condition and the presence of complex cardiac lesions, the decision was made to proceed with a TAH. The abnormal arrangement of the patient's ventricles and great arteries required modifications to the TAH during implantation.
With the TAH in place, the patient was able to return home and regain strength and physical well-being while awaiting a donor heart. He was successfully bridged to heart transplantation 5 months after receiving the device. This report highlights the TAH is feasible even in patients with structurally abnormal hearts, with technical modification.

This is a case report of a patient with congenitally corrected transposition of the great arteries (CCTGA) who received a total artificial heart (TAH) as a bridge to heart transplant. The TAH was successfully implanted with modifications to accommodate the patient's congenitally malformed heart.

In patients with end-stage heart failure (HF), a total artificial heart (TAH) may be implanted as a bridge to cardiac transplant. However, in congenital ...

Techniques for Processing Eyes Implanted with a Retinal Prosthesis for Localized Histopathological Analysis: Part 2 Epiretinal Implants with Retinal Tacks

Retinal prostheses for the treatment of certain forms of blindness are gaining traction in clinical trials around the world with commercial devices currently entering the market. In order to evaluate the safety of these devices, in preclinical studies, reliable techniques are needed. However, the hard metal components utilised in some retinal implants are not compatible with traditional histological processes, particularly in consideration for the delicate nature of the surrounding tissue. Here we describe techniques for assessing the health of the eye directly adjacent to a retinal implant secured epiretinally with a metal tack.
Retinal prostheses feature electrode arrays in contact with eye tissue. The most commonly used location for implantation is the epiretinal location (posterior chamber of the eye), where the implant is secured to the retina with a metal tack that penetrates all the layers of the eye. Previous methods have not been able to assess the proximal ocular tissue with the tack in situ, due to the inability of traditional histological techniques to cut metal objects. Consequently, it has been difficult to assess localized damage, if present, caused by tack insertion.
Therefore, we developed a technique for visualizing the tissue around a retinal tack and implant. We have modified an established technique, used for processing and visualizing hard bony tissue around a cochlear implant, for the soft delicate tissues of the eye. We orientated and embedded the fixed eye tissue, including the implant and retinal tack, in epoxy resin, to stabilise and protect the structure of the sample. Embedded samples were then ground, polished, stained, and imaged under various magnifications at incremental depths through the sample. This technique allowed the reliable assessment of eye tissue integrity and cytoarchitecture adjacent to the metal tack.

Here we describe histological techniques for visualising ocular tissue directly adjacent to a metal epiretinal tack and retinal prosthesis.

Retinal prostheses for the treatment of certain forms of blindness are gaining traction in clinical trials around the world with commercial devices ...

Primary Outcome Assessment in a Pig Model of Acute Myocardial Infarction

Mortality after acute myocardial infarction remains substantial and is associated with significant morbidity, like heart failure. Novel therapeutics are therefore required to confine cardiac damage, promote survival and reduce the disease burden of heart failure. Large animal experiments are an essential part in the translational process from experimental to clinical therapies. To optimize clinical translation, robust and representative outcome measures are mandatory. The present manuscript aims to address this need by describing the assessment of three clinically relevant outcome modalities in a pig acute myocardial infarction (AMI) model: infarct size in relation to area at risk (IS/AAR) staining, 3-dimensional transesophageal echocardiography (TEE) and admittance-based pressure-volume (PV) loops. Infarct size is the main determinant driving the transition from AMI to heart failure and can be quantified by IS/AAR staining. Echocardiography is a reliable and robust tool in the assessment of global and regional cardiac function in clinical cardiology. Here, a method for three-dimensional transesophageal echocardiography (3D-TEE) in pigs is provided. Extensive insight into cardiac performance can be obtained by admittance-based pressure-volume (PV) loops, including intrinsic parameters of myocardial function that are pre- and afterload independent. Combined with a clinically feasible experimental study protocol, these outcome measures provide researchers with essential information to determine whether novel therapeutic strategies could yield promising targets for future testing in clinical studies.

Reliable and accurate outcome assessment is the key for translation of preclinical therapies into clinical treatment. The current paper describes how to assess three clinically relevant primary outcome parameters of cardiac performance and damage in a pig acute myocardial infarction model.

Mortality after acute myocardial infarction remains substantial and is associated with significant morbidity, like heart failure. Novel therapeutics are ...

Neonatal Murine Cochlear Explant Technique as an In Vitro Screening Tool in Hearing Research

While there have been remarkable advances in hearing research over the past few decades, there is still no cure for Sensorineural Hearing Loss (SNHL), a condition that typically involves damage to or loss of the delicate mechanosensory structures of the inner ear. Sophisticated in vitro and ex vivo assays have emerged in recent years, enabling the screening of an increasing number of potentially therapeutic compounds while minimizing resources and accelerating efforts to develop cures for SNHL. Though homogenous cultures of certain cell types continue to play an important role in current research, many scientists now rely on more complex organotypic cultures of murine inner ears, also known as cochlear explants. The preservation of organized cellular structures within the inner ear facilitates the in situ evaluation of various components of the cochlear infrastructure, including inner and outer hair cells, spiral ganglion neurons, neurites, and supporting cells. Here we present the preparation, culture, treatment, and immunostaining of neonatal murine cochlear explants. The careful preparation of these explants facilitates the identification of mechanisms that contribute to SNHL and constitutes a valuable tool for the hearing research community.

Neonatal Murine Cochlear Explant Technique as an In Vitro Screening Tool in Hearing Research

The goal of this protocol is to demonstrate the preparation, culture, treatment, and immunostaining of neonatal murine cochlear explants. The technique can be utilized as an in vitro screening tool in hearing research.

While there have been remarkable advances in hearing research over the past few decades, there is still no cure for Sensorineural Hearing Loss (SNHL), a ...

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 ...

Performing Intracochlear Electrocochleography During Cochlear Implantation

Electrocochleography (ECochG) measures inner ear potentials generated in response to acoustic stimulation of the ear. These potentials reflect the residual function of the cochlea. In cochlear implant candidates with residual hearing, the implant electrode can directly measure ECochG responses during the implantation process. Various authors have described the ability to monitor the inner ear function by continuous ECochG measurements during the surgery. The measurement of ECochG signals during surgery is not trivial. There are no interpretable signals in up to 20% of cases. For a successful recording, a standardized procedure is recommended to achieve the highest measurement reliability and avoid possible pitfalls. Therefore, seamless collaboration between the CI surgeon and CI technician is key. This video consists of an overview of the system setup and a stepwise procedure of performing intracochlear ECochG measurements during CI surgery. It shows the surgeon's and the CI technician's roles in the process, and how a smooth collaboration between the two is made possible.

Electrocochleography (ECochG) measures inner ear potentials generated in response to acoustic stimulation. In cochlear implant (CI) candidates, such inner ear potentials can be measured directly with the implant electrodes. In this video, we systematically explain how to perform ECochG recordings during CI surgery.

Electrocochleography (ECochG) measures inner ear potentials generated in response to acoustic stimulation of the ear. These potentials reflect the ...

Robotic Cochlear Implantation for Direct Cochlear Access

Robot-assisted systems offer great potential for gentler and more precise cochlear implantation. In this article, we provide a comprehensive overview of the clinical workflow for robotic cochlear implantation using a robotic system specifically developed for a minimally invasive, direct cochlear access. The clinical workflow involves experts from various disciplines and requires training to ensure a smooth and safe procedure. The protocol briefly summarizes the history of robotic cochlear implantation. The clinical sequence is explained in detail, beginning with the assessment of patient eligibility and covering surgical preparation, preoperative planning with the special planning software, drilling of the middle ear access, intraoperative imaging to confirm the trajectory, milling of the inner ear access, insertion of the electrode array, and implant management. The steps that require special attention are discussed. As an example, the postoperative outcome of robotic cochlear implantation in a patient with advanced otosclerosis is presented. Finally, the procedure is discussed in the context of the authors' experience.

Robotic cochlear implantation is a procedure for minimally invasive inner ear access. Compared to conventional surgery, robotic cochlear implantation involves additional steps that need to be carried out in the operating room. In this article we give a description of the procedure and highlight the important aspects of robotic cochlear implantation.

Robot-assisted systems offer great potential for gentler and more precise cochlear implantation. In this article, we provide a comprehensive overview of ...

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, ...