implantation of an electrode assembly in a mouse brain for electrophysiological recording

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.

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Universidad de Cartagena UDEC

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JoVE Encyclopedia of Experiments

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Electrophysiology Techniques

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All procedures involving animal models have been reviewed by the local institutional animal care committee and the JoVE veterinary review board.
1. Making headsets with two bipolar electrodes (Figure 1)
<ol>
	<li>Cut 3.5 cm of stainless polytetrafluoroethylene-coated stainless-steel wire.</li>
	<li>Strip about 1 mm of the insulation coat off the wire on both ends. Do not have too much of the wire stripped.</li>
	<li>Put two pins on a vise holder with the bottom part of the pin that has a longer slit facing down.</li>
	<li>Apply flux onto the stripped ends of the wire and onto the tops of the pins.</li>
	<li>Tin the stripped part of the wire with just enough solder to coat it.</li>
	<li>Add a minimal amount of solder to the top of the pin without overflowing onto the sides.</li>
	<li>Place one end of the stripped ends of the wire into the pin as deep as it will allow while the solder is melted. 
	NOTE: There is a side hole where the wire can come out - do not let the wire come out of the pin. All of the stripped wire must stay within the pin.</li>
	<li>Repeat steps 1.6-1.7 for the second pin, now with the other stripped ends of the wire.</li>
	<li>Let the pins sit for 30 s to set, remove them from the vise holder, and then pull on them to make sure that the connection between the wire and the pins is strong and holds.</li>
	<li>Rinse the pins in cold water, and then dry.</li>
	<li>Verify conductance between pin 1 and pin 2 using an ohmmeter.</li>
	<li>Bring the pins at the ends of the wire together; hold them parallel and close. Clamp a hemostat to the center of the wire. Then, rotate the hemostat, so the wire twists around itself fairly tight. Remove the hemostat.</li>
	<li>Clamp a forceps onto the twisted wire 2 mm below the pins and bend the wire at 90&#730;.</li>
	<li>Push the same wire again 90&#730; back over the forceps creating another bend 1 mm from the first.</li>
	<li>Cut the twisted wire at a 45&#176; angle below the bend at 3.5 mm with small sharp scissors.</li>
	<li>Prepare two of these bipolar (double-pin twisted) electrodes for each headset (optional, second is back-up in case of electrical problems with the first).</li>
	<li>Prepare one single reference electrode by cutting a wire with pins soldered on both ends into two (steps 1.1-1.17, Figure 1A).</li>
	<li>Cut the wire at 7 mm.</li>
	<li>Bend the end 1 mm below the tip.</li>
	<li>Then, cover the bent 1 mm tip of the wire with forceps and rotate the wire tight around the forceps to create a small loop (1 mm in diameter).</li>
	<li>Bend the loop perpendicular to the straight part of the wire to make the wire tip point outward again.</li>
	<li>Assemble the two bipolar electrodes and the single reference electrode into the six-pin pedestal in a way that the bipolar electrodes are side by side with 6 mm distance between them, and that the reference electrode is placed in the middle outer hole (Figure 1B). 
	NOTE: An alternative method is to implant the electrodes, cement them in place, and then insert their pins into the pedestal.</li>
</ol>
2. Stereotaxic electrode implantation
<ol>
	<li>Sterilize all surgical tools and the six-pin electrode assembly by autoclaving before the surgery. A sterile surgical field must be maintained during the surgery, and sterile surgical gloves must be used. Sterile drapes (e.g., Press n&#39; Seal) are recommended to cover the animal except for the surgical area.</li>
	<li>Use 8-week-old VGAT-Cre mice (expressing Cre recombinase under the control of the vesicular GABA transporter [VGAT] gene, age-matched, both male and female) for surgeries 4 weeks after weaning. Record the weight of the animal before surgery to allow the measurement of post-surgical weight loss.</li>
	<li>Use a certified isoflurane vaporizer or a low-flow anesthesia system equipped with a precision syringe pump, integrated digital vaporizer, and feedback heat pad. 
	NOTE: Low-flow systems are capable of delivering anesthesia at low flow rates proportionate to the animal&#39;s size into either an induction chamber or through a nose cone on the stereotaxic frame (70 mL/min, isoflurane concentration in air is 4% for induction and 2% for surgery). Using less anesthesia not only benefits the animal during surgeries but also reduces the risk of lab personnel&#39;s exposure to isoflurane.</li>
	<li>Place the anesthetized animal on a heated pad warmed to 37 &#176;C to keep it warm during surgery. If using a feedback-controlled temperature system, insert the lightly lubricated temperature probe into the rectum of the animal for temperature monitoring during surgery.</li>
	<li>Mount the animal onto the stereotaxic frame by gently placing ear bars into the ears and the front upper teeth into the incisor bar. Place the nose cone over the nose for proper anesthesia delivery. Make sure that the head is leveled and centered and cannot be moved when slightly probed.</li>
	<li>Subcutaneously inject 0.5 mL of normosol for hydration.</li>
	<li>Apply ocular lubricant to prevent corneal drying.</li>
	<li>Monitor the depth of anesthesia by the absence of the withdrawal reflex after pinching a hindlimb toe, and then decrease isoflurane to 1.5%-2.0% during surgery.</li>
	<li>Remove hair at and around the surgical area by plucking or using clippers (shaving) or depilatory cream and disinfect the skin with three cycles of alternating application of iodine and ethanol, finishing with iodine. Removing hair away from the surgical site is recommended only if there are means to maintain anesthesia at that location. If necessary, use alcohol-dipped cotton tip applicators to remove hair from the immediate area surrounding the head. Inject 0.05 mL of the local analgesic bupivacaine (0.25%) subcutaneously.</li>
	<li>Make an incision on the skull using a scalpel, and then cut out a part of the skin with sharp surgical scissors, exposing the skull. Pushing the skin aside, using a cotton swab, clean the skull from all the muscles and underlying tissues obstructing the view. 
	NOTE: To stop accidental bleeding, apply pressure over the bleeding site with a sterile cotton swab until it stops.</li>
	<li>Clean the skull with hydrogen peroxide using sterile cotton swabs to make the skull sutures and both bregma and lambda visible.</li>
	<li>Dry the skull thoroughly, and then apply one drop of self-etching dental adhesive using its applicator. Brush it into the skull, wait 60 s, and cure with a dental UV light for 40 s. A glossy surface indicates that the adhesive has been effectively cross-linked with the skull. 
	NOTE: This step is critical for secure attachment of the headset.</li>
	<li>Use a 0.031&#34; drill bit (0.79 mm) to drill two burr holes bilaterally for implantation of hippocampal depth electrodes (approximately 5,000 rpm). Drill one extra burr hole for the reference electrode above the cerebellum behind the lambda. 
	&#8203;NOTE: When drilling, take care to lower the drill slowly and avoid drilling into the brain.</li>
	<li>The coordinates for the electrodes are as follows (from bregma in mm): hippocampal electrodes at 3 mm posterior, 3 mm lateral, and 3 mm depth; and cerebellar reference electrode at 6 mm posterior, 0 mm lateral, and 0 mm depth (subdural).</li>
	<li>To increase the accuracy, use a stereotaxically mounted drill, zero the stereotaxic manipulator X/Y axis when touching bregma - this is the point of reference for the coordinates.</li>
	<li>Assemble a headset by inserting all the electrodes in the six-pin pedestal, making sure the pins are pushed all the way into the pedestal. Mount the pedestal into the electrode holder on a stereotaxic frame (Figure 1B).</li>
	<li>Align electrodes above the corresponding burr holes. Stereotaxically implant twisted bipolar stainless-steel wire electrodes in the right and left hippocampus and reference electrode into the cerebellum by slowly lowering the headset and guiding the electrodes into the burr holes.</li>
	<li>When the hippocampal twist electrode is right above the hole, zero the Z axis and slowly lower to -3.0 mm.</li>
	<li>Cover the skull surface and electrodes with dental cement and fill in the space between the skull surface and the bottom of the pedestal. The skin edges will be adjacent to the dental cement such that no underlying tissue will remain exposed. Wait for the cement to dry and harden. Then detach the electrode holder from the stereotaxic arm and remove the holder from the pedestal.</li>
	<li>Inject 0.1 mL of ketoprofen (1 mg/mL, subcutaneously [SC]) for analgesia and a second dose of 0.5 mL of normosol (SC) for hydration and remove the animal from the stereotaxic frame.</li>
	<li>Place an isothermal pad preheated to 37 &#176;C inside an empty vivarium cage covered with a paper towel. Once it is fully awake, place the animal in a clean cage with bedding and soft food, and return it to the vivarium. Animals are housed singly from this point on to prevent chewing on each other&#39;s headsets. Wire bar hoppers are not used to avoid headsets getting stuck, and instead, water bottles are tied to the underside of the cage tops, and chow is present in the bedding.</li>
	<li>Feed the animal some soft food for 72 h after surgery and monitor for weight loss and body condition score. Dehydrated animals may be administered 0.5 mL of normosol subcutaneously if dehydrated (weight loss, increased skin turgor, sunken eyes). Ketoprofen can be given subcutaneously once daily for 2 more days following surgery (follow the analgesia regimen according to the local institutional animal care and use committee [IACUC] guidelines). Allow animals to fully recover in their cages for 4-7 days before being transferred to the electroencephalography (EEG) recording system.</li>
</ol>

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>16 Channel Extracellular Differential AC Amplifier (115V/60Hz)</td>
			<td>AD Instruments</td>
			<td>AM3500-115-60</td>
			<td>Alternate EEG amplifier</td>
		</tr>
		<tr>
			<td>363/CP PLUG COLLAR, PINS SLEEVE</td>
			<td>P1 Technologies</td>
			<td>363SLEEVPIN0NL</td>
			<td>For electrode holder</td>
		</tr>
		<tr>
			<td>Cable, 363-363 5CM - 100CM W/MESH 6TCM</td>
			<td>P1 Technologies</td>
			<td>363363XXXXCM004</td>
			<td>mouse-to-commutator cable</td>
		</tr>
		<tr>
			<td>CCTV cameras Qcwox HD Sony IR LED</td>
			<td>Sony</td>
			<td>QC-SP316</td>
			<td></td>
		</tr>
		<tr>
			<td>Commutator SL6C/SB (single brush)</td>
			<td>P1 Technologies</td>
			<td>8BSL6CSBC0MT</td>
			<td>formerly Plastics One, Inc.</td>
		</tr>
		<tr>
			<td>Current amplifier</td>
			<td>A-M Systems</td>
			<td>Model 2100</td>
			<td></td>
		</tr>
		<tr>
			<td>Dental cement</td>
			<td>Stoelting</td>
			<td>51459</td>
			<td></td>
		</tr>
		<tr>
			<td>Drill bits, #75, OD 0.310&#34; LOC 130 PT</td>
			<td>Kyocera</td>
			<td>105-0210.310</td>
			<td></td>
		</tr>
		<tr>
			<td>E363/0 SOCKET CONTACT SKEWED</td>
			<td>P1 Technologies</td>
			<td>8IE3630XXXXE</td>
			<td>pins for connector</td>
		</tr>
		<tr>
			<td>iBond Self Etch glue</td>
			<td>Kulzer</td>
			<td>CE0197</td>
			<td></td>
		</tr>
		<tr>
			<td>MS363 PEDESTAL 2298 6 PIN WHITE</td>
			<td>P1 Technologies</td>
			<td>8K000229801F</td>
			<td>EEG headset connector</td>
		</tr>
		<tr>
			<td>Ohmeter</td>
			<td>Simpson</td>
			<td>260</td>
			<td>High sensitivity</td>
		</tr>
		<tr>
			<td>PowerLab 16/35 and LabChart Pro</td>
			<td>AD Instruments</td>
			<td>PL3516/P</td>
			<td>Alternate EEG software</td>
		</tr>
		<tr>
			<td>SomnoSuite</td>
			<td>Kent Scientific Corp.</td>
			<td>SS-01</td>
			<td>anesthesia unit &#38; RightTemp monitoring</td>
		</tr>
		<tr>
			<td>Stereotactic drill and micromotor kit</td>
			<td>Foredom Electric Co.</td>
			<td>K.1070</td>
			<td></td>
		</tr>
		<tr>
			<td>Stereotactic frame</td>
			<td>David Kopf Instruments</td>
			<td>Model 940</td>
			<td></td>
		</tr>
		<tr>
			<td>Teflon-coated wire for depth electrode, OD 0.008&#39;</td>
			<td>A-M Systems</td>
			<td>791400</td>
			<td></td>
		</tr>
		<tr>
			<td>VGAT-Cre mice on congenic C57BL/6J background</td>
			<td>The Jackson Laboratory</td>
			<td>664</td>
			<td></td>
		</tr>
	</tbody>
</table>

Source: Straub, J., et al. <a href="https://appjove.unicartagenaproxy.elogim.com/v/62929">Preparation and Implantation of Electrodes for Electrically Kindling VGAT-Cre Mice to Generate a Model for Temporal Lobe Epilepsy.</a> J. Vis. Exp. (2021).
This video demonstrates a technique for implanting an electrode assembly into a mouse brain. The assembly contains bipolar electrodes to record activity from the hippocampus and a reference electrode to obtain baseline activity from the cerebellum. An anesthetized mouse is placed on a stereotaxic frame, and the electrode assembly is implanted in the target brain region. The animal is then allowed to recover before performing electrophysiological recording.

<img alt="Figure 1" src="/files/ftp_upload/22847/22847_Figure1.jpg" /> 
Figure 1: Key steps in EEG headset fabrication. (A) Appearance of the electrodes at the various steps in the protocol (numbers in the left match step). (B) A picture of the final product mounted in a homemade holder that fits the stereotaxic frame. Note, the holder ends with a collar pin assembly that fits into the headset pedestal.

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

Secure an anesthetized mouse on a stereotaxic frame containing a heating pad to maintain body temperature.
Apply lubricant to prevent the eyes from drying and inject an electrolyte solution subcutaneously to maintain hydration.
Remove the fur from the scalp, disinfect the area, and inject an analgesic subcutaneously to reduce pain.
Make an incision to expose the skull, then clean it to make anatomical landmarks visible for identifying brain regions.
Apply dental adhesive and harden it using a curing light to aid in electrode implantation.
Drill holes above the hippocampus and cerebellum, and mount an electrode assembly using a holder on the stereotaxic frame.
The bipolar electrodes record hippocampal activity, while a reference electrode obtains baseline activity from the cerebellum.
Insert the electrodes, secure the assembly with dental cement, and detach from the holder.
Administer the electrolyte solution and an analgesic again.
Allow the mouse to completely recover before performing electrophysiological recording.

70 ビュー. 電気生理学的記録のためのマウス脳への電極アセンブリの埋め込み

ビデオ: 電気生理学的記録のためのマウス脳への電極アセンブリの埋め込み - 実験

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

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

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

人工内耳手術と c57bl/6 マウスに電気的誘発脳幹の応答録音

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

JoVE Journal

神経科学

Using a Bipolar Electrode to Create a Temporal Lobe Epilepsy Mouse Model by Electrical Kindling of the Amygdala

The amygdala is one of the most common origins of seizures, and the amygdala mouse model is essential for the illustration of epilepsy. However, few studies have described the experimental protocol in detail. This paper illustrates the whole process of amygdala electrical kindling epilepsy model making, with the introduction of a method of bipolar electrode fabrication. This electrode can both stimulate and record, reducing brain injury caused by implanting separate electrodes for stimulation and recording. For long-term electroencephalogram (EEG) recording purposes, slip rings were used to eliminate the record interruption caused by cable tangles and falling off.
After periodic stimulation (60 Hz, 1 s every 15 min) of the basolateral amygdala (AP: 1.67 mm, L: 2.7 mm, V: 4.9 mm) for 19.83 &plusmn; 5.742 times, full kindling was observed in six mice (defined as induction of three continuous grade V episodes classified by Racine's scale). An intracranial EEG was recorded throughout the entire kindling process, and an epileptic discharge in the amygdala lasting 20-70 s was observed after kindling. Therefore, this is a robust protocol for modeling epilepsy originating from the amygdala, and the method is suitable for revealing the role of the amygdala in temporal lobe epilepsy. This research contributes to future studies on the mechanisms of mesial temporal lobe epilepsy and novel antiepileptogenic drugs.

The amygdala plays a key role in temporal lobe epilepsy, which originates in and propagates from this structure. This article provides a detailed description of the fabrication of deep brain electrodes with both recording and stimulating functions. It introduces a model of medial temporal lobe epilepsy originating from the amygdala.

バイポーラ電極を用いて扁桃体の電気キンドリングによる側頭葉てんかんマウスモデルの作成

The amygdala is one of the most common origins of seizures, and the amygdala mouse model is essential for the illustration of epilepsy. However, few ...

網膜神経節細胞層におけるカルシウム活性の評価

Calcium Imaging In Electrically Stimulated Flat-Mounted Retinas

Retinal dystrophies are a leading cause of blindness worldwide. Extensive efforts are underway to develop advanced retinal prostheses that can bypass the impaired light-sensing photoreceptor cells in the degenerated retina, aiming to partially restore vision by inducing visual percepts. One common avenue of investigation involves the design and production of implantable devices with a flexible physical structure, housing a high number of electrodes. This enables the efficient and precise generation of visual percepts. However, with each technological advancement, there arises a need for a reliable and manageable ex vivo method to verify the functionality of the device before progressing to in vivo experiments, where factors beyond the device's performance come into play. This article presents a comprehensive protocol for studying calcium activity in the retinal ganglion cell layer (GCL) following electrical stimulation. Specifically, the following steps are outlined: (1) fluorescently labeling the rat retina using genetically encoded calcium indicators, (2) capturing the fluorescence signal using an inverted fluorescence microscope while applying distinct patterns of electrical stimulation, and (3) extracting and analyzing the calcium traces from individual cells within the GCL. By following this procedure, researchers can efficiently test new stimulation protocols prior to conducting in vivo experiments.

著者スポットライト:カルシウムイメージングを使用した神経電気刺激への革新的なアプローチ 

Retinal prostheses have the ability to generate visual perceptions. To advance the development of new prostheses, ex vivo methods are needed to test devices before implantation. This article provides a comprehensive protocol for studying calcium activity in the retinal ganglion cell layer when subjected to electrical stimulation.

電気刺激フラットマウント網膜におけるカルシウムイメージング

Retinal dystrophies are a leading cause of blindness worldwide. Extensive efforts are underway to develop advanced retinal prostheses that can bypass the ...