Name: cochlear implant surgery and electrically-evoked auditory brainstem response recordings in c57bl/6 mice
Uploaded: 2019-01-09T13:00:00.000+00:00
Duration: 9 min 6 s
Description: Watch this Scientific Journal Video about Cochlear Implant Surgery and Electrically-evoked Auditory Brainstem Response Recordings in C57BL/6 Mice at JoVE.com

The authors would like to thank Pierre Stahl, Oticon Medical, Nice, France, for providing the Animal Stimulation Platform and advice on stimulation paradigms, and James B. Fallon and Andrew K. Wise from Bionics Institute, Melbourne, Australia, for surgical advice. This work was supported by a grant from the Swiss National Science Foundation (ERC transfer grant to T.R.B.).

All procedures were carried out according to Basel University, Switzerland, animal care and guidelines. They were licensed by the Veterinary Office of the Canton of Basel, Switzerland.
NOTE: C57BL/6 adult mice, aged 8&#8211;12 weeks (weight 20&#8211;30 g), were used in this study. 
The left ear is used as the experimental ear. The right ear serves as an intra-animal control and is not surgically altered.
1. Preoperative Procedures
<ol>
	<li>Anaesthetize the animal 30 min prior to surgery via intra-peritoneal (i.p.) injection of ketamine/xylazine (80 mg/kg ketamine, 16 mg/kg xylazine, i.p., volume injected at 10 &#956;L/g body weight).
	<ol>
		<li>Supplement anesthesia as necessary, as judged by a positive pedal and palpebral (toe-pinch) reflex and movement of the whiskers, with a lower dose of ketamine (45 mg/kg, i.p., injected at 10 &#956;L/g body weight). Agents and dose regimes can be substituted per institutional guidelines. 
		Notes: In general, the animal will need an injection every 45&#8211;60 min with this agent and dose regime.&#160;The average time from initial incision to closure around the implanted electrode array is typically 1-1.5 hours.</li>
	</ol>
	</li>
	<li>Check for full sedation of the animal marked by a regular breathing rate and a lack of toe-pinch reflexes. Maintain this level of anesthesia.</li>
	<li>Maintain the animal&#39;s body temperature at 36.6 &#176;C with a closed-loop heating pad. Apply eye ointment to avoid dehydration of the cornea. This will also suppress the animal&#39;s blink reflex, which can add noise to the recoding.</li>
	<li>Administer local analgesic via subcutaneous injection (s.c.) of bupivacaine/Lidocaine (0.1 mg/mL bupivacaine and 0.4 mg/mL lidocaine, 0.1 mL administered s.c.) along the intended incision line to minimize any surgical discomfort. Agents and dose regimes can be substituted per institutional guidelines.</li>
	<li>Administer the muscarinic antagonist atropine (atropinesulfate amino, 0.1 mg/mL, 20 &#956;L administered s.c., dissolved in PBS) in the neck to reduce mucus secretion and to facilitate breathing. Agents and dose regimes can be substituted per institutional guidelines.</li>
</ol>
2. Pre-deafening Acoustic Auditory Brainstem Response (aABR)
NOTE: aABR is used to measure the status of hearing before and after deafening. Testing is performed on the left ear and in a soundproof electrically shielded booth. We recommend to test and later implant the left ear for a right-handed person. Further details on ABR in mice can be found in13,14. Tucker Davis Technologies (TDT) hardware and software (BioSig) are used to record ABR but other systems can be used.
<ol>
	<li>Block the contralateral (right) ear with acoustic foam to isolate the ABR response from the ipsilateral (left) ear. Put the foam in a 1 mL syringe and inject it into the right ear canal of the mouse to cover the whole ear canal with foam (0.1&#8211;0.2 mL of foam). Make sure the syringe seals closely to the ear so that the foam gets all the way into the ear canal.</li>
	<li>Place the speaker 10 cm from the left ear. 
	NOTE: The speaker for this setup was calibrated using a PCB microphone as described in refefence15.</li>
	<li>Clean the ABR electrodes with 70% ethanol solution. Place the electrodes under the skin: active (Ch1) on the vertex, reference (-) below the pinna of the ipsilateral ear, and ground in the hind leg (Figure 1).</li>
	<li>Connect the head-stage and pre-amplifier to the auditory processor via the optic fiber port.</li>
	<li>Check the impedance of the active and reference electrode.
	<ol>
		<li>If the impedance is over 3 Ohm, re-arrange them and re-take the measurement. The best recordings are obtained when the electrodes have the same impedance. Close the sound-proof booth.</li>
	</ol>
	</li>
	<li>Present click stimulation and record ABR in a free-field condition with a complex auditory processor and software. Standardize the click stimulus in the software: 0.1 ms single-channel monophasic clicks are presented at 21 Hz; Click level decreases from 90 dB SPL to 10 dB SPL in 10 dB steps; 10 ms recording window. Average a total of 512 responses at each dB level.</li>
	<li>Apply a 2,000 Hz lowpass filter and a 300 Hz highpass filter offline to reduce noise in the recording using a custom-made Matlab script.</li>
	<li>Determine the ABR threshold as the lowest dB level with a recognizable ABR wave response (Figure 2, Figure 3).</li>
</ol>
3. Surgery
NOTE: Typical instruments used include a scissor, a scalpel, a pair of metallic forceps with straight or curved tips, a tissue retractor tool, several suction wedges and absorbable paper points. The surgery is performed on the left ear.
<ol>
	<li>Put the mouse on its right side. Avoid undue torsional stress on the cervical vertebrae. Make sure to keep the body straight to keep the airways open.</li>
	<li>Cut the fur behind the left ear with a scissor (or shave it with a shaver) to expose the skin. Sterilize the skin with 70% ethanol solution and betadine (povidone/iodine).</li>
	<li>Under microscopic magnification (16x), make a 1&#8211;1.5 cm post-auricular incision with the scalpel.</li>
	<li>Switch to higher microscopic magnification (25x).</li>
	<li>Perform blunt dissection through the subcutaneous fat layer, which can be of variable thickness, with forceps. 
	NOTE: Be careful when dissecting as the external jugular vein traverses this area. Damage to this structure can cause excessive bleeding.</li>
	<li>Retract the sternocleidomastoid muscle to reveal the tympanic bulla periosteum. Use the facial nerve as a key anatomical landmark to aid identification of the auditory bulla. The facial nerve wraps around the posterior/dorsal edge of the sternocleidomastoid muscle and runs rostrally along the ear canal towards the pinna. Gently place the self-retaining retractor tool in the incision to ease access to the bulla (Figure 4).</li>
	<li>Remove the tissue overlying the medio-dorsal area of the bulla to allow clear visualization of the ridge between the bulla and the mastoid process.</li>
	<li>Gently rotate a 30 G needle to pierce the bulla and make a hole (bullostomy) on the posterior-superior side of the ridge (the bone is thinner on this side). Alternatively, use a dental surgical drill. 
	NOTE: This and the following steps can be done with even higher microscopic magnification (40x) if preferred. Also, change the position of the microscope if needed. It is important to maximize the surgical view of the middle ear space.</li>
	<li>Widen the bullostomy by pinching small bone pieces using fine tipped forceps to expose the middle ear cavity. Extend the bullostomy dorsally towards the mastoid process until the round window niche is clear of overlying bone. The stapedial artery, a branch of the internal carotid artery, runs ventral to the round window niche.
	<ol>
		<li>Be careful not to damage the vessel as excessive bleeding may be fatal. Small bleeds can be stopped by pressing a small piece of spongostan in the inner ear cavity.</li>
		<li>Extend the bullostomy toward the anterior-superior direction to visualize the stapes, the middle ear bone connected to the oval window.</li>
	</ol>
	</li>
	<li>Remove the stapes with forceps to expose the oval window.</li>
</ol>
4. Round Window Application of Ototoxic Agent
<ol>
	<li>Gently perforate the round window and oval window membranes using a blunted 30 G needle. Check that perilymph runs out.</li>
	<li>Slowly perfuse 0.05% weight/volume neomycin dissolved in PBS (adjusted to pH 7.4) through the oval window. Liquid should flush out of the round window. Repeat the same procedure on the round window. Be careful not to damage the window bone structures with the needle used to perfuse.</li>
	<li>Place a small piece (1 mm2) of spongostan soaked in neomycin within the round window and oval window niche.</li>
	<li>Remove the retractor tool, close the incision and wait 30 min.</li>
</ol>
5. Post-deafening Acoustic ABR
<ol>
	<li>Record aABR in a similar way as before deafening (Steps 2.2 to 2.8) (Figure 2b, Figure 3).</li>
</ol>
6. Insertion of CI Electrode Array
NOTE: The intracochlear electrode array consists of four platinum bands (&#216;0.2 mm) with platinum/iridum parylene insulated wire shielded in a silicone tube (Figure 5).
<ol>
	<li>Place the retractor tool in the incision to re-access the bulla.</li>
	<li>Insert the electrode array into the round window (scala tympani) at a depth where the 4th platinum ring is located just inside the round window. This gives an insertion depth of ~2 mm, corresponding to an intracochlear position at ~30 kHz16.</li>
	<li>Coil the lead wire inside the bulla and glue the wire to the tissue above the bulla. Coiling the wire helps to keep the array in place throughout the experiment.</li>
	<li>Carefully remove the retractor and close the insertion with tissue glue.</li>
	<li>Make a small incision (0.5 mm) in the neck perpendicular to the line between where the active and reference ABR electrodes will be using a tissue scissor. Place the platinum ground ball in the subcutaneous pocket and close the small incision with tissue glue (Figure 6).</li>
	<li>Connect the electrode array board to the Animal Stimulator Platform.</li>
</ol>
7. Electric Auditory Brainstem Response (eABR)
NOTE: An Animal Stimulator Platform (ASP) is used to electrically stimulate the electrode array. Other current sources and software systems can be used.
<ol>
	<li>Place the ABR electrodes as before (Steps 2.3 to 2.5) (Figure 6).</li>
	<li>Open the ASP software and define the electric pulse stimulation paradigm. We use a charge-balanced biphasic pulses with 50 &#956;s/phase and 10 &#956;s interphase gap presented at 23.3 pulses per second (pps). The electric stimulation is delivered in monopolar electrode configuration with increasing current levels. A total of 400 responses are averaged at each current level.</li>
	<li>Present the electric pulse trains and record the evoked eABR response continuously via the TDT headstage, pre-amplifier and auditory processor.</li>
	<li>Plot and analyze the eABR data via a custom-made matlab script (Figure 7). The script and an example of a recording are provided in the Supplementary.</li>
</ol>
8. End of Experiment
<ol>
	<li>At the end of the experiment, euthanize the animal according to institutional guidelines.</li>
	<li>Carefully open the incision and remove the implant.</li>
	<li>Ultra-sonicate the electrode array in distilled water for 10 min to remove tissue debris. 
	NOTE: The implant can be reused several times if the electrodes are intact and properly conducting. To check this, measure the impedance of the electrodes with a multimeter when the array is dry.</li>
	<li>Store the electrode array in a dry place.</li>
</ol>

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No competing financial interests. The authors have nothing to disclose.

This manuscript describes the surgical approach for acute deafening and cochlear implantation in the mouse, as well as the functional assessment of CI stimulation with auditory brainstem response. Although the mouse cochlea is small and the surgery challenging, the CI mouse model is feasible and serves as a valuable tool in auditory research.
The stapedial artery is present in the middle ear of the mouse. The artery enters the bulla posterior-medially, and runs inferiorly to the round window niche and then superiorly to the oval window niche. In the initial development of the mouse model, we experienced fatal intra-operative bleeding following trauma to the stapedial artery, mainly while accessing the bulla. As a consequence, we adapted a more limited approach and accessed the bulla in smaller, refined dissection steps. No further complications due to bleeding were thereafter observed. Despite the fact that stapedial artery cauterization has no significant effect on hearing threshold or number of spiral ganglion neurons in mice5, in our opinion, it is unnecessary so long as great care and attention are taken during the surgery. We suggest taking the time needed to develop fine psychomotor skills and reach technical proficiency. The average time from initial incision to closure around the implanted electrode array is typically 1&#8211;1.5 h.
The described acute CI surgery in mice is similar to the &#34;ventral&#34; procedure and round window insertion used in other rodents, including rats and gerbils20,21,22. Other rodent studies have used the &#34;dorsal approach&#34; with a basal turn cochleostomy instead of a round window insertion, avoiding the SA entirely and inserting the array more deeply6,23,24. The implantation of a chronic stimulation assembly in mice follows the same steps as described in this protocol with the addition of a Dacron mesh to fix the implant and postoperative care5.
The main technical challenges when performing CI surgeries in mice are the small size of the cochlea compared to the cochlea of rats and gerbils, and the presence of a large SA. The SA is also present in rats but not in gerbils. In addition, since mice are smaller than rats and gerbils, they are more vulnerable to surgical procedures.
To eliminate electrophonic responses in eABR recordings and to mimic hair cell loss found in most CI users, we deafened the animals before CI insertion. Mice are difficult to deafen ototoxically in vivo25 because the systemic concentrations of aminoglycosides required to cause ototoxicity has a narrow dose window: lower doses given over several days results in no hair cell loss whereas a single injection of a higher dose can be lethal26. Also, susceptibility to aminoglycosides is strain dependent26. However, it has been shown that a single dose of aminoglycosides in combination with a loop diuretic can produce excessive outer hair cell loss in CBA/CaJ mice without fatal consequences27. Delayed inner hair cell death was reported in half of all cochleae examined27.
In this manuscript, we used topical application of the aminoglycosides neomycin inspired by the protocol recently established for C57BL/6 mice5. Acute application of neomycin significantly increased the click-evoked hearing threshold by 46 dB &#177; 6.1. Although this increase is larger than the 35 dB increase reported by Irving et al. (pre- vs post-surgery: 41.6 dB &#177; 3.3 vs 76.6 dB &#177; 4.4, p = 0.02, n = 3)5, we achieved the same post-deafening threshold (75.7 dB &#177; 3.7 vs 76.6 dB &#177; 4.4). 0.05% neomycin is thought to cause a partial loss of hearing, mainly by rapid outer hair cell death, as inner hair cell loss takes longer to occur27. It is therefore possible that electrophonic response, which is generated both by inner and outer hair cells4,8,9,10,11,12, is only partially eliminated in deafened animals with residual hearing. Even though 0.05% (weight/volume) neomycin does not decrease the number of spiral ganglion neurons 4 weeks post-deafening5, it is yet unknown if neomycin in our acute setup affects the auditory nerve fibers or promotes synaptopathy (loss of synapses between inner hair cells and type I auditory nerve fibers). Another uncertainty is that the topical neomycin treatment may not produce a uniform distribution of hair cell loss along the length of the cochlea. Future studies are required to answer these questions.
In summary, the growing number of genetic models for human deafness and the biochemical tools available make the mouse an attractive animal model for auditory research, including the field of CIs.

Cochlear implants (CIs) are electronic devices that can provide a sense of hearing to people with severe and profound hearing loss. It uses electrodes surgically implanted in the cochlea of the inner ear to directly stimulate the auditory nerve. To date, the CI is the most successful sensory prosthesis and has helped more than 600,000 people worldwide1. However, the device has shortcomings. First, the benefits provided by the device vary greatly among recipients. Second, speech in noisy environments and music are still poorly perceived by most CI users.
For many years, animal models have been used to better understand these issues in CI research and to continuously improve safety and efficacy of the devices. The models have given valuable insight into several phenomena, such as plastic changes in the brain taking place following CI implantation2, the effect of applying gene therapy to preserve residual hearing3, and biophysical properties of the electrically stimulated auditory nerve4, among many other examples.
Mice are a powerful model organism due to the large availability of genetic models of deafness. Other advantages include the ability to manipulate the mouse genome (e.g., via the CRISPR-Cas system), the opportunity to use advanced imaging techniques to study mechanisms, particularly in the brain, the high reproduction rate, rapid development and easy breeding and handling. The main technical challenges in performing CI surgeries in mice are the small size of the cochlea and the presence of a large stapedial artery (SA). The SA usually disappears during embryonic development in humans but persists throughout life in a number of rodents, including mice, rats, and gerbils. The SA runs below the round window niche, which complicates access to the cochlea and increases surgical risk.
Previous studies have shown the feasibility of CI implantation in mice5,6,7. Irving et&#160;al. demonstrated that chronic intracochlear electrical stimulation can be achieved for up to one month. Acute stimulation was also performed but the recordings were not presented. They showed that cauterizing the stapedial artery had no significant effect on the hearing threshold or the number of spiral ganglion neurons and that topical application of the aminoglycoside neomycin, an ototoxic drug, was an effective deafening procedure in mice5. Soken et al. described a modified dorsal approach to the mouse cochlea through the round window to better preserve hearing status6. Following insertion of a platinum-iridium wire, substantial residual hearing was observed with an increased auditory brainstem response (ABR) threshold of 28 dB. Otoacoustic emissions (OAE) were lost in animals with large ABR threshold shifts6. Mistry et al. tested the functional and histopathological effects of implantation in the absence of electric stimulation7. Even though hearing was preserved in both 3 and 6 months-old implanted mice at low frequencies, implantation resulted in fibrosis-like tissue around the implant and osteoneogenesis around the bullostomy7.
In short, out of the three studies on CIs in mice, only one demonstrates functional recording of CI stimulation. Irving and colleagues performed both acute and chronic eABR recordings but only showed data from chronic CI stimulation5. However, the chronic model with a fully implantable device developed by Irving et al. is technically challenging. It is not yet known if acute CI stimulation, both less challenging and faster, can achieve similar results.
CIs are used by people with severe and profound hearing loss who no longer benefit from hearing aids. Animal models for CI users should therefore include a deafening procedure when normally hearing animals are used. Another reason to deafen hearing animals is that the electrical stimulation of a deaf or hearing cochlea produces different neural responses4,8,9,10,11,12. Electrical stimulation of a deaf cochlea directly activates the auditory nerve fibers and generates an electroneural response (&#945;). It is characterized by short latency and a small dynamic range in the periphery8,10. On the other hand, electrical stimulation of a hearing cochlea also excites the hair cells in an electrophonic response (&#946;) that is characterized by longer latencies and larger dynamic range4,11. The electrophonic response is attributed to normal excitation of nerve fibers by inner hair cells, electrically induced contraction of outer hair cells, and generation of a travelling wave4. Electroneural and electrophonic responses also result in two different activity patterns in the central nervous system9. Sato et al. recorded midbrain neurons of a CI implanted guinea pig before and after deafening with neomycin, which eliminates the electrophonic contribution. They showed that the slope of the rate-level function was steeper and firing rates higher in the deafened condition compared to the hearing condition9. Therefore, depending on the research question stated, it is important to consider including deafening to separate electrophonic and electroneural responses upon electric stimulation of the auditory nerve.
Here, we describe the procedure for acute deafening and the cochlear implantation of an electrode array in a mouse as well as the functional recording of intracochlear electric stimulation with electrically-evoked auditory brainstem response (eABR).

<table><tbody><tr><td>&lt;strong&gt;Hardware&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>Sound-proof booth</td><td>IAC Acoustics, Winchester, UK</td><td>Mac-2 Enclosure RF Shielded Box 2A</td><td></td></tr><tr><td>MF1 Speaker</td><td>Tucker Davis Technologies (TDT), FL, USA</td><td></td><td></td></tr><tr><td>PCB microphone</td><td>PCB Piezotronics, Inc, NY, USA</td><td>Model 378C01</td><td></td></tr><tr><td>Low impedance headstage</td><td>TDT, FL, USA</td><td>RA4LI</td><td></td></tr><tr><td>Medusa pre-amplifier</td><td>TDT, FL, USA</td><td>RA4PA</td><td></td></tr><tr><td>RZ6 auditory processor</td><td>TDT, FL, USA</td><td></td><td></td></tr><tr><td>Animal Stimulator Platform</td><td>ASP, Oticon Medical, Nice, France</td><td></td><td></td></tr><tr><td>Multimeter</td><td>Fluks</td><td>#115</td><td></td></tr><tr><td>&lt;strong&gt;Surgical equipment&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>Closed-loop heating pad</td><td>FHC, Inc. ME, USA</td><td></td><td></td></tr><tr><td>Eye ointment</td><td>Alcon, CH</td><td>Lacrinorm Augengel</td><td></td></tr><tr><td>Acoustic foam</td><td>Otoform Ak, Dreve Otoplastik GmbH</td><td>#464</td><td></td></tr><tr><td>Disposable subdermal needle electrodes</td><td>Horizon, Rochester Electro-Medical Inc.</td><td>S83018-R9, 27G</td><td></td></tr><tr><td></td><td></td><td></td><td></td></tr><tr><td>Self-retaining retractor tool (Mini Collibri Retractor)</td><td>Fine Science Tools</td><td>#17000-01</td><td></td></tr><tr><td>Suction wedges</td><td>Agnthos, SE</td><td>#42-886-460</td><td></td></tr><tr><td>Absorbable paper point (Medium)</td><td>WPI, FL, USA</td><td>#504182</td><td></td></tr><tr><td>Intracochlear electrode array</td><td>Bionics Institute, Melbourne, Australia</td><td>4 channel</td><td></td></tr><tr><td>Spongostan Standard</td><td>Ferrosan Medical Devices</td><td>#MS0002</td><td></td></tr><tr><td>Tissue glue. Loctite 4161 Superbond</td><td>Henkel</td><td>Part No 19743</td><td></td></tr><tr><td>Animal Stimulator Platform (ASP)</td><td>Oticon Medical, Nice, France</td><td></td><td></td></tr><tr><td>&lt;strong&gt;Drugs/chemicals&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>Ketamine (Narketan)</td><td>Provet AG, CH</td><td>100mg/mL, #VQ_320265</td><td></td></tr><tr><td>Xylazine (Rompun)</td><td>Provet AG, CH</td><td>Inj Diss 2%, # 1315</td><td></td></tr><tr><td>Bupivacaine</td><td>Compendium, CH</td><td>Bupivacain Sintetica inj Diss 0.5%</td><td></td></tr><tr><td>Atropine (Atropinesulfat Amino)</td><td>Amino AG, CH</td><td>1 mg/mL</td><td></td></tr><tr><td>Betadine (Povidone/iodine)</td><td>Provedic, CH</td><td></td><td></td></tr><tr><td>Neomycin (Neomycin trisulfate salt)</td><td>Sigma</td><td>N1876-25G, Lot#WXBB7516V</td><td></td></tr><tr><td>&lt;strong&gt;Software&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>BioSigRZ</td><td>TDT, FL, USA</td><td></td><td></td></tr><tr><td>Matlab</td><td>MathWorks, MA, USA</td><td></td><td></td></tr><tr><td>ASP software</td><td>Oticon Medical, Nice, France</td><td></td><td></td></tr></tbody></table>

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.

The purpose of this study was to describe a reliable model for acute CI stimulation in the deafened mouse. Pre- and post-surgical hearing thresholds served as a functional readout of the deafening procedure. Topical application of 0.05% neomycin in the oval and round window significantly increased click-evoked hearing thresholds by 46 dB &#177; 6 (pre- vs post-neomycin: 30.0 dB &#177; 3.8 vs 75.7 dB &#177; 3.7, p = 0.0003, paired t-test, n= 7) (Figure 3). The mouse-sized electrode array was hereafter inserted into the round window (Figure 4, Figure 5). Electric simulation of an intracochlear electrode could reliably generate eABR activity. (Figure 7). In some cases, CI stimulation activated the facial nerve and produced a high amplitude wave with either short or long latency (Figure 8A and Figure 8B, respectively). The short latency response was characterized by a rapid amplification of wave IV around 3 ms and is likely to be a direct response of the facial nerve. The long latency response appeared around 5&#8211;6 ms and is likely to be a non-auditory muscle (myogenic) response evoked indirectly by the facial nerve. Facial nerve responses are rarely reported in animal studies in literature but is a well-known complication in human CI users17,18,19. In Figure 8, facial nerve stimulation appeared at relatively medium current levels (150&#8211;200 &#956;A) and in two different animals. In other cases, both responses could appear in the same animal at very high current levels (not shown). We recommend limiting the current level to levels below the appearance of facial nerve stimulation.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/58073/58073fig1.jpg" /> 
Figure 1: Auditory Brainstem Response (ABR) setup. Subdermal electrodes are placed at the vertex (active/channel 1 [Ch1]), behind the ipsilateral ear (reference [Ref]) and at the hind leg (ground [Gnd]) of the anesthetized mouse. Electrode signals are amplified and then recorded by a TDT system. Acoustic and electric stimulation are presented via a microphone and an Animal Stimulator Platform, respectively. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/58073/58073fig1large.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/58073/58073fig2.jpg" /> 
Figure 2: Representative aABR waves to click stimulation from a wild-type mouse before and after deafening with 0.05% neomycin. (A) The normal-hearing aABR pattern is characterized by waves labelled I-V and a low hearing threshold, here 30 dB SPL (arrow). (B) The deafened aABR pattern shows an increased hearing threshold, here 70 dB SPL (arrow). The waves have a longer latency and more temporal jitter. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/58073/58073fig2large.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/58073/58073fig3.jpg" /> 
Figure 3: aABR threshold before and after deafening. Application of neomycin significantly increased aABR thresholds by 46 dB &#177; 6. Pre- vs post-neomycin: 30.0 dB &#177; 3.8 vs 75.7 dB &#177; 3.7, p = 0.0003, paired t-test, n = 7. Errors are standard error of the means. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/58073/58073fig3large.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/58073/58073fig4.jpg" /> 
Figure 4: The surgery. (A) Exposure to the auditory bulla. The bullostomy is performed (white dotted circle) along the ridge on the tympanic bulla (black dotted line). (B) The bullostomy allows visualization of the round window, stapedial artery, and oval window. Neomycin is gently flushed through first the oval window, then the round window. (C) The electrode array is inserted until the 4th electrode is located just inside the round window niche. The electrode wire is coiled inside the bulla to keep the array in place before the incision is closed. CN VII = cranial nerve VII (facial nerve), OW = oval window, RW = round window, SA = stapedial artery, SCM = sternocleidomastoid muscle, TB = tympanic bulla. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/58073/58073fig4large.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/58073/58073fig5.jpg" /> 
Figure 5: The mouse cochlear implant. (A) The intracochlear electrode array consists of four platinum bands spaced at a 0.4 mm interval with a diameter d: 0 [tip] (d=0.21), 1 (d=0.23), 2 (d=0.25), 3 (d=0.27). The width of each electrode is 0.2 mm. The four platinum/iridium (90/10) parylene insulated wires are shielded in a silicone tube. (B) Magnification of the electrode array tip (red dotted square). The electrode array and a platinum reference ball are connected to a print board. Scale bar = 1 mm. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/58073/58073fig5large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 6" class="xfigimg" src="/files/ftp_upload/58073/58073fig6.jpg" /> 
Figure 6: Electrically-evoked ABR (eABR) setup. The CI platinum ground ball (Gnd, red) is placed in a subcutaneous pocket in the neck of the mouse. The line between active (Ch1(+) at the vertex) and reference (Ref (-) at the ipsilateral ear) ABR electrodes is perpendicular to the line between the electrode array and the ground in order to obtain the best eABR response. The eABR ground electrode (Gnd, black) is placed in the hind leg. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/58073/58073fig6large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 7" class="xfigimg" src="/files/ftp_upload/58073/58073fig7.jpg" /> 
Figure 7: Representative eABR waves to CI stimulation in a deafened mouse. A biphasic pulse train is presented to electrode #1 in monopolar configuration at 23.3 pulses per second (pps) with 400 repetitions. Stimuli level 0-175 &#956;A is shown in 25 &#956;A steps (see stimulation details in Step 7.2). Roman numerals denote eABR wave number. The wave amplitudes and latency increase and decrease, respectively, with increasing current level. In this example, wave II appeared around 1 ms, wave III around 2 ms, wave IV around 3 ms, wave V around 4 ms. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/58073/58073fig7large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 8" class="xfigimg" src="/files/ftp_upload/58073/58073fig8.jpg" /> 
Figure 8: Example of facial nerve stimulation. In some cases, CI stimulation can activate the facial nerve and evoke a direct response with short latency (A) (arrow) or indirect response with longer latency (B) (arrow). The examples shown are from two CI-implanted animals stimulated with a biphasic pulse train using 0&#8211;300 &#956;A in 50 &#956;A steps (see stimulation details in Step 7.2). Roman numerals denote eABR wave numbers. * denotes clipping of the eABR wave due to saturation of the amplifier. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/58073/58073fig8large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Cochlear Implant Surgery and Electrically-evoked Auditory Brainstem Response Recordings in C57BL/6 Mice at JoVE.com

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

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 ...

cochlear-implant-surgery-electrically-evoked-auditory-brainstem

Nanyang Technological University

Research

JoVE Journal

Neuroscience

13.8K Views.  Technical University of Denmark. Diermodellen van cochleaire implantaten kunnen onze kennis van de technologische basis van de behandeling van permanent sensorisch zenuwverlies met elektrische stimulatie bevorderen. Dit protocol toont acute oorverdovende en cochleaire implantatie in de muis aan. En de functionele beoordeling van cochleaire implantaatstimulatie met auditieve hersenstamrespons.Enkele belangrijke uitdagingen van het uitvoeren van cochleaire implantaatchirurgie bij muizen zijn de kleine omvang van het sla...

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

Direct Visualization of the Murine Dorsal Cochlear Nucleus for Optogenetic Stimulation of the Auditory Pathway

Investigation into the use of virus-mediated gene transfer to arrest or reverse hearing loss has largely been relegated to the peripheral auditory system. Few studies have examined gene transfer to the central auditory system. The dorsal cochlear nucleus (DCN) of the brainstem, which contains second order neurons of the auditory pathway, is a potential site for gene transfer. In this protocol, a technique for direct and maximal exposure of the murine DCN via a posterior fossa approach is demonstrated. This approach allows for either acute or survival surgery. Following direct visualization of the DCN, a host of experiments are possible, including injection of opsins into the cochlear nucleus and subsequent stimulation by an optical fiber coupled to a blue light laser.&#160;Other neurophysiology experiments, such as electrical stimulation and neural injector tracings are also feasible.&#160;The level of visualization and the duration of stimulation achievable make this approach applicable to a wide range of experiments.

The goal of this protocol is to outline a surgical approach to provide direct access to the dorsal cochlear nucleus in a murine model.

Investigation into the use of virus-mediated gene transfer to arrest or reverse hearing loss has largely been relegated to the peripheral auditory system. ...

Morphological and Functional Evaluation of Ribbon Synapses at Specific Frequency Regions of the Mouse Cochlea

Cochlear inner hair cells (IHCs) transmit acoustic signals to spiral ganglion neurons (SGNs) through ribbon synapses. Several experimental studies have indicated that hair cell synapses may be the initial targets in sensorineural hearing loss (SNHL). Such studies have proposed the concept of cochlear &#34;synaptopathy&#34;, which refers to alterations in ribbon synapse number, structure, or function that result in abnormal synaptic transmission between IHCs and SGNs. While cochlear synaptopathy is irreversible, it does not affect the hearing threshold. In noise-induced experimental models, restricted damage to IHC synapses in select frequency regions is employed to identify the environmental factors that specifically cause synaptopathy, as well as the physiological consequences of disturbing this inner ear circuit. Here, we present a protocol for analyzing cochlear synaptic morphology and function at a specific frequency region in adult mice. In this protocol, cochlear localization of specific frequency regions is performed using place-frequency maps in conjunction with cochleogram data, following which the morphological characteristics of ribbon synapses are evaluated via synaptic immunostaining. The functional status of ribbon synapses is then determined based on the amplitudes of auditory brainstem response (ABR) wave I. The present report demonstrates that this approach can be used to deepen our understanding of the pathogenesis and mechanisms of synaptic dysfunction in the cochlea, which may aid in the development of novel therapeutic interventions.

This manuscript describes an experimental protocol for evaluating the morphological characteristics and functional status of ribbon synapses in normal mice. The present model is also suitable for noise-induced and age-related cochlear synaptopathy-restricted models. The correlative results of previous mouse studies are also discussed.

Cochlear inner hair cells (IHCs) transmit acoustic signals to spiral ganglion neurons (SGNs) through ribbon synapses. Several experimental studies have ...

Data Acquisition and Analysis In Brainstem Evoked Response Audiometry In Mice

Brainstem evoked response audiometry (BERA) is of central relevance in the clinical neurophysiology. As other evoked potential (EP) techniques, such as visually evoked potentials (VEPs) or somatosensory evoked potentials (SEPs), the auditory evoked potentials (AEPs) are triggered by the repetitive presentation of identical stimuli, the electroencephalographic (EEG) response of which is subsequently averaged resulting in distinct positive (p) and negative (n) deflections. In humans, both the amplitude and the latency of individual peaks can be used to characterize alterations in synchronization and conduction velocity in the underlying neuronal circuitries. Importantly, AEPs are also applied in basic and preclinical science to identify and characterize the auditory function in pharmacological and genetic animal models. Even more, animal models in combination with pharmacological testing are utilized to investigate for potential benefits in the treatment of sensorineural hearing loss (e.g., age- or noise-induced hearing deficits). Here we provide a detailed and integrative description of how to record auditory brainstem-evoked responses (ABRs) in mice using click and tone-burst application. A specific focus of this protocol is on pre-experimental animal housing, anesthesia, ABR recording, ABR filtering processes, automated wavelet-based amplitude growth function analysis, and latency detection.

Brainstem evoked response audiometry is an important tool in clinical neurophysiology. Nowadays, brainstem evoked response audiometry is also applied in the basic science and preclinical studies involving both pharmacological and genetic animal models. Here we provide a detailed description of how auditory brainstem responses can be successfully recorded and analyzed in mice.

Brainstem evoked response audiometry (BERA) is of central relevance in the clinical neurophysiology. As other evoked potential (EP) techniques, such as ...

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

We present two minimally invasive microsurgical techniques in rodents for specific drug delivery into the middle ear so that it may reach the inner ear. The first procedure consists of perforation of the tympanic bulla, termed bullostomy; the second one is a transtympanic injection. Both emulate human clinical intratympanic procedures.
Chitosan-glycerophosphate (CGP) and Ringer&#180;s Lactate buffer (RL) were used as biocompatible vehicles for local drug delivery. CGP is a nontoxic biodegradable polymer widely used in pharmaceutical applications. It is a viscous liquid at RT&#160;but it congeals to a semi solid phase at body temperature. RL is an isotonic solution used for intravenous administrations in humans. A small volume of this vehicle is precisely placed on the Round Window (RW) niche by means of a bullostomy. A transtympanic injection fills the middle ear and allows less control but broader access to the inner ear.
The safety profiles of both techniques were studied and compared by using functional and morphological tests. Hearing was evaluated by registering the Auditory Brainstem Response (ABR) before and several times after microsurgery. The cytoarchitecture and preservation level of cochlear structures were studied by conventional histological techniques in paraformaldehyde-fixed and decalcified cochlear samples. In parallel, unfixed cochlear samples were taken and immediately frozen to analyze gene expression profiles of inflammatory markers by quantitative Reverse Transcriptase Polymerase Chain Reaction (qRT-PCR).
Both procedures are suitable as drug delivery methods into the mouse middle ear, although transtympanic injection proved to be less invasive compared to bullostomy.

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

Two microsurgery approaches for local drug delivery to the inner ear are described here and compared in terms of impact on hearing parameters, cochlear cytoarchitecture and expression of inflammatory markers.

We present two minimally invasive microsurgical techniques in rodents for specific drug delivery into the middle ear so that it may reach the inner ear. ...

Biology

Flexible Multi-Region Neural Recording Device for Epilepsy Research

An Integrated Method for Crafting Flexible and Convenient Electrophysiological Optrodes for Multi-Region In Vivo Recording

Epilepsy is a neurological disorder characterized by synchronized abnormal discharges involving multiple brain regions. Focal lesions facilitate the propagation of epileptic signals through associated neural circuits. Therefore, in vivo recording of local field potential (LFP) from the critical brain regions is&#160;essential for deciphering the circuits involved in seizure propagation. However, current methods for electrode fabrication and implantation lack flexibility. Here, we present a handy device designed for electrophysiological recordings (LFPs and electroencephalography [EEG]) across multiple regions. Additionally, we seamlessly integrated optogenetic manipulation and calcium signaling recording with LFP recording. Robust after-discharges were observed in several separate regions during epileptic seizures, accompanied by increasing&#160;calcium signaling. The approach used in this study offers a convenient and flexible strategy for synchronous neural recordings across diverse regions of the brain. It holds the potential for advancing research on neurological disorders by providing insights into the neural profiles of multiple regions involved in these disorders.

Author Spotlight: Unraveling Neural Communication and Circuit Interactions in Health and Disease

We have developed a simplified and cost-effective approach for electrode fabrication and conducted recordings of signals across multiple regions in freely moving mice. Utilizing optogenetics, alongside multi-region electrophysiology and calcium signal recording, enabled the revelation of neuronal activities across regions in the seizure kindling model.

Epilepsy is a neurological disorder characterized by synchronized abnormal discharges involving multiple brain regions. Focal lesions facilitate the ...

Investigating Auditory Neuronal Responses in an Awake Mouse in the Presence of an Inhibitory Neurotransmitter Blocker

Source: Ayala, Y. A., et al. <a href="https://app-jove-com.remotexs.ntu.edu.sg/v/53914">Extracellular Recording of Neuronal Activity Combined with Microiontophoretic Application of Neuroactive Substances in Awake Mice.</a> J. Vis. Exp. (2016).
This video demonstrates the study of neuronal activity in a specific region of the mouse brain that processes auditory stimuli. The technique involves applying an inhibitory neurotransmission blocker and comparing neuronal responses to auditory stimuli before and after the drug is applied to the localized brain region.

All procedures involving animal models have been reviewed by the local institutional animal care committee and the JoVE veterinary review board.
1. Drugs ...

JoVE Encyclopedia of Experiments

Neurophysiology

Stimulation and Modulation Techniques

Subdural Implantation of a Soft Electrocorticography Array for Cortical Electrophysiology Recording in Minipigs

Source: Fallegger, F. et al., <a href="https://app-jove-com.remotexs.ntu.edu.sg/v/64997">Subdural Soft Electrocorticography (ECoG) Array Implantation and Long-Term Cortical Recording in Minipigs.</a>&#160;J. Vis. Exp. (2023).
This video demonstrates the implantation of a soft electrocorticography array and the long-term cortical recordings in a minipig. It outlines the steps involved in surgically implanting the array over the minipig&#39;s auditory cortex and recording both baseline and sound stimulus-evoked neuronal action potentials.

All procedures involving animal models have been reviewed by the local institutional animal care committee and the JoVE veterinary review board.
1. ...

Electrophysiology Techniques

Evaluation of Ribbon Synapses at Specific Frequency Regions of Mouse Cochlea

Source: Yu, S., et al. <a href="https://app-jove-com.remotexs.ntu.edu.sg/v/59189">Morphological and Functional Evaluation of Ribbon Synapses at Specific Frequency Regions of the Mouse Cochlea.</a> J. Vis. Exp. (2019).
This video demonstrates a method for evaluating ribbon synapses in specific regions of the mouse cochlea. It utilizes immunostaining and confocal imaging techniques to identify functional synapses across different frequency regions in each cochlear turn.

All procedures involving animal handling have been performed in accordance with the institute&#39;s IRB guidelines.
1. Animal Selection

	For all ...

Recording Electrically-Evoked Auditory Brainstem Responses in a Deafened Mouse

Source: Navntoft, C. A., et al. <a href="https://app-jove-com.remotexs.ntu.edu.sg/v/58073">Cochlear Implant Surgery and Electrically-evoked Auditory Brainstem Response Recordings in C57BL/6 Mice.</a> J. Vis. Exp. (2019)
This video demonstrates the procedure to implant a cochlear device in a deafened mouse and apply electrical stimuli to record auditory brainstem responses. This procedure enables to understand the basic mechanisms of electrical hearing and can contribute to advancements in cochlear implant technology and auditory neuroscience.

Implantation of an Electrode Assembly in a Mouse Brain for Electrophysiological Recording

Before proceeding with the surgery, record the weight of the 8-week-old VGAT-Cre mice. Then, place the animal in an induction chamber to induce anesthesia. Place the anesthetized mouse on a heating pad at 37 degrees Celsius. If using a feedback-controlled temperature system, insert the temperature probe into the rectum of the animal for temperature monitoring during the surgery. To start the surgery, redirect the inhalant anesthesia flow to the nose cone and mount the animal in the stereotaxic frame by gently placing the front upper teeth into the incisor bar and move the cone onto its nose, then place the ear bars into the ears and affix to the stereotaxic frame.
Ensure that the animal&#39;s head is leveled and centered and cannot be moved when slightly probed. Once the animal is mounted, inject 0.5 milliliters of normosol onto the mice for hydration and apply ocular lubricant to prevent corneal drying. Monitor the depth of anesthesia by pinching a hind-limb toe. If the withdrawal reflex is absent in the animal, proceed to remove the scalp hair of the mouse around the surgical area. When done, disinfect the clean skin area thrice with an alternating application of ethanol and iodine finishing with iodine, and inject 0.05 milliliters of 0.25% bupivacaine subcutaneously.
Make an incision on the skull using a scalpel, and then, cut out a part of the skin with surgical scissors exposing the skull. Push the skin aside before cleaning the skull from all underlying tissues with a cotton swab. Clean the skull with hydrogen peroxide using sterile cotton swabs, and make the skull sutures, bregma, and lambda visible. With the stereotaxically mounted drill, Move the drill tip onto bregma and zero x, y, and z coordinates, then, raise and move the drill to lambda to determine its coordinates to determine if the head is level.
After drying the skull thoroughly, apply one drop of self-etching dental adhesive with the applicator. Wait 60 seconds before curing the dental adhesive with a dental UV light for 40 seconds. A glossy surface indicates that the adhesive has been effectively cross-linked with the skull. With the help of a 0.031-inch drill bit, carefully drill two boreholes bilaterally into the skull at 5,000 rotations per minute, or RPM, avoiding drilling into the brain. Apply a drop of saline to enhance the drilling of the burr holes. For implantation of hippocampal depth electrodes, drill at 3 millimeters posterior and 3 millimeters lateral from bregma.
Drill one burr hole for the reference electrode above the cerebellum, behind the lambda at 6 millimeters posterior and 0 millimeters lateral from bregma. Next, mount the six-pin pedestal with assembled electrodes into the electrode holder on a stereotactic frame. By aligning the electrodes above the corresponding burr holes, slowly lower and navigate the headset to locate the two bipolar electrode tips right on top of the left and the right hippocampal burr holes, and if needed, adjust the reference electrode to hover above hole, over the cerebellum.
When the hippocampal twist electrodes are right above the holes, zero the z-axis, and slowly lower the electrode to minus three millimeters into the right and left hippocampus, and guide the reference electrode into the hole above the cerebellum. Prepare the dental cement by mixing the powder and liquid in a mixing bowl, then cover the skull surface, the electrodes, and the space between the skull surface and the bottom of the pedestal with dental cement.
Once the cement dries and hardens, detach the electrode holder from the stereotactic arm, raise arm, and remove the holder from the pedestal. After administering the mouse subcutaneously with ketoprofen and normosol, remove the animal from the stereotactic frame to move the animal onto the heated pad in the vivarium cage. Once fully awake, return the single animal to the vivarium cage, feed the animal with soft food, and monitor the weight loss daily up to 72 hours post-surgery.

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

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