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EoE Sub Articles

All procedures involving animal models have been reviewed by the local institutional animal care committee and the JoVE veterinary review board.
1. Surgical implantation of soft electrocorticography (ECoG) arrays
<ol>
	<li>Anesthesia
	<ol>
		<li>Premedication: Isolate the Female G&#246;ttingen minipigs (n = 7) at 2-6 months of age (5-8 kg) and fast it overnight. Inject a mix of midazolam at 0.75 mg/kg, atropine at 0.25 &#181;g/kg, and haldol at 0.1 mg/kg intradermally, and wait until the animal is sedated. Weigh the animal before proceeding.</li>
		<li>Installation of intravenous (IV) lead:
		<ol>
			<li>Place the animal on the surgery table on a heating pad. Induce anesthesia by placing a face mask on the animal, using sevoflurane at 3%-3.5%.</li>
			<li>Place electrocardiogram leads on the abdomen, a blood saturation sensor on the tail, and a temperature sensor in the nostril.</li>
			<li>Place an IV lead on an ear vein and confirm blood access using a syringe filled with saline. Ensure the eyes are kept hydrated by using ointment.</li>
		</ol>
		</li>
		<li>Intubation: Inject a bolus of atracurium at 0.5 mg/kg, ketamine at 1 mg/kg, and fentanyl at 1-2 &#181;g/kg. Place the animal on its back for intubation. Insert a 4.5 mm tube.</li>
		<li>Medication: After intubation, stop the sevoflurane anesthesia and install a propofol infusion at 10 mg/kg/h, fentanyl at 2 &#181;g/kg/h, atracurium at 0.2-0.5 mg/kg/h, and saline at 4-7 mg/kg/h. Start an infusion of mannitol at 1 g/kg/h to reduce brain swelling during the surgery. 
		&#8203;NOTE: A multimodal analgesia regimen may be used if recommended by the local animal ethics committee.</li>
	</ol>
	</li>
	<li>Aseptic field and skin preparation: Shave the entire surface of the head beyond the surgical field. Using a sterile pad, thoroughly scrub the head with betadine. Next, place sterile drapes on the instrumentation table and on the animal to reveal only the surgical window. Finally, scrub the head again with a sterile pad using betadine.</li>
	<li>Craniotomy and durotomy
	<ol>
		<li>Skin cut: Incise the skin with a scalpel knife along the midline. Separate the muscle and periosteum (25 mm laterally from bregma on both sides and 40 mm anterior and posterior to bregma) from the bone using a raspatory and place spreaders to get optimum access for later drilling.</li>
		<li>Measurements and marking: Identify bregma and lambda and mark them with a sterile surgical pen (Figure 1A, B). Using a sterile ruler, define the bone flap outline centered around the implantation target on both hemispheres. In this specific case, the auditory cortex was chosen, with coordinates -5 mm to -15 mm from bregma and -4 mm to -20 mm laterally. Then, adjust the craniotomy to the size of the implant and anatomical landmarks, limiting the opening size.</li>
		<li>Craniotomy:
		<ol>
			<li>Using a bone drill with a round cutting bit, drill the outline of the craniotomy, taking into account the thickness of the skull. Irrigate the drilling location with saline solution to avoid overheating the bone.</li>
			<li>Carefully drill the outline homogeneously until reaching the dura mater. At the first breakthrough, finish drilling the outline until it has thinned enough to almost break through. Then, use a flat spatula (either on the midline or the lateral side) to break away the bone flap in one piece, using the craniotomy edge as leverage. If too much resistance is encountered, continue thinning the bone.</li>
			<li>Place the bone piece in sterile saline.</li>
			<li>Once the bone flap is removed, carefully chip the edge of the craniotomy away, using a Kerison to avoid the sharp bone edge from cutting into the dura mater.</li>
			<li>If excessive bleeding is encountered on the dura mater or the bone, use Gelfoam or bone wax, respectively. Place a wet compress (standard pad in sterile saline solution) in the craniotomy and repeat this step on the other hemisphere (Figure 1B).</li>
		</ol>
		</li>
		<li>Durotomy:
		<ol>
			<li>Using the needle from a 6-0 suture kit, carefully pierce and lift the dura mater at the anterior or posterior end of the craniotomy halfway between the medial and lateral side and create the beginning of an incision with the stab knife.</li>
			<li>Then, using a small flat spatula inserted in the subdural space acting as a cutting base to protect the cortex, create an anteroposterior slit in the dura mater by advancing simultaneously with both tools. Ensure that the slit is slightly larger than the width of the implant (Figure 1C). If any bleeding or damage occurs at this step, cover it with gel foam and wait until it stops. 
			&#8203;NOTE: The slit trajectory should be adapted if large blood vessels are present in the dura mater to avoid bleeding.</li>
		</ol>
		</li>
	</ol>
	</li>
	<li>Implantation
	<ol>
		<li>Device insertion:
		<ol>
			<li>Irrigate the implant (Figure 2A) with saline on both sides so it slides into the subdural space more easily. Place the implant above the dura mater slit and, with small forceps, subdurally insert the device by sliding it sequentially on each edge.</li>
			<li>Carefully hold the pedestal end of the device and advance with the implant in order not to create tension hindering the insertion. When the connector edge is located on top of the slit, stop the insertion.</li>
		</ol>
		</li>
		<li>Secure the implant: To secure the implant in place, place a titanium bridge over the cable after the edge of the craniotomy or in the anchoring wings and secure it with one or two titanium screws using the appropriate screwdriver (Figure 1D).</li>
		<li>Ground placement: Carefully remove 1 cm of the insulation of the ground wires and insert it epidurally at the posterior end of the craniotomy (or any epidural location far from the cortex of interest or large blood vessels) (wire in&#160;Figure 1E)</li>
		<li>Repeat the steps on the contra-lateral hemisphere.</li>
		<li>Dura mater closure: Suture the dura mater carefully around the implant cable using a 3-0 resorbable suture and a small needle holder. Bring the two dura mater edges together as much as possible without tearing through the thin membrane with the suture wire (Figure 1D, E).</li>
		<li>Bone flap placement: Fix a titanium bridge on the anterior and posterior part of each bone flap using a titanium screw. Be careful to plan the positioning of the Ti-bridges with respect to the placement of the footplate legs in the next steps. Screw the end of the titanium bridges to the skull (Figure 1F, G).</li>
	</ol>
	</li>
	<li>Pedestal and footplate placement
	<ol>
		<li>Positioning: In this configuration, the footplate has six legs with two screw holes each (Figure 2B). Plan the placement of the footplate on the skull to optimize the location of the screws (avoid placing them at the edge of the craniotomy or in the temporal muscle). Skip the holes in the legs if they cannot be screwed in.</li>
		<li>Footplate securing: Screw in the titanium screws of the footplate until it is firmly in place (see&#160;Figure 1H).</li>
		<li>Pedestal placement: Remove the titanium bridges over the connection cables and flip over the pedestal to land on the footplate. Screw the pedestal onto the footplate. Check that the pedestal is firmly in place (Figure 1I).</li>
	</ol>
	</li>
	<li>Suture and closure
	<ol>
		<li>Cleaning of the wound: Clean the subcutaneous space of any bone or other debris by flushing with saline. Cut away some skin around the pedestal edges to create a round edge following the cylinder.</li>
		<li>Subcutaneous sutures: Remove the spreaders and fold the skin flaps together. Create subcutaneous sutures with a 4-0 non-resorbable suture wire, 3 mm apart, using simple interrupted sutures or simple continuous sutures. Start away from the pedestal, moving toward it on both sides of the incision.</li>
		<li>Dermal sutures: Suture the skin using a 6-0 non-resorbable suture wire, with sutures 5 mm apart. Start away from the pedestal, moving toward it on both sides of the incision. Take care to achieve good tissue apposition between the two skin flaps and near the pedestal edge to avoid a void (Figure 1J).</li>
		<li>Wound dressing: Clean the wound area again with a sterile pad and betadine. Apply a self-adhesive sterile bandage over the wound.</li>
	</ol>
	</li>
	<li>Awakening: After all the measurements have been performed, take the animal off all anesthetics but keep under ventilation. For analgesia, apply a buprenorphine patch (25 mg/h) for 24 h. Place the animal on a heating pad covered with drapes to speed up the wake-up time. When spontaneous breathing is recovered, extubate the animal and put under an oxygen face mask until consciousness is recovered (which can take 1 to 4 h).</li>
	<li>Postoperative animal care: For 5 days, keep the animal under close surveillance.&#160;Give a dose of cephalexin at 75 mg twice daily with food, separated from other animals. Carry out disinfection of the wound daily by applying copious amounts of betadine with soaked sterile pads (best done during feeding). 
	NOTE: Long-term care and housing: The operated animal is kept isolated for 24 h. It is put back in its original social group if the animal is well enough to interact socially with its peers.&#160;Daily observation of the pedestal and skin opening needs to be conducted to follow the integration of the device on the head. When appropriate, clean the location around the pedestal with copious amounts of betadine.</li>
</ol>
2. Electrophysiological recording
<ol>
	<li>Spontaneous activity: Plug the wireless recording system through the pedestal and record the baseline activity for 2-3 min. These recordings will serve as a control to analyze the auditory evoked potentials.</li>
	<li>Auditory evoked potentials: In addition to the wireless system, insert speakers in a closed field in the animal ears. Play tone burst acoustic stimulation at different frequencies (ranging from 200-20,000 Hz) at around 70 dB sound pressure level (SPL) over 120 repetitions. Then, average the recordings and align them over the stimulus period for analysis.</li>
	<li>Sensory evoked potentials: Place the needles in the snout at three different positions. Evoke sensory potentials by stimulating the snout for ~30 s with the pulse generator at different amplitudes to obtain the recruitment curves.</li>
</ol>
3. Freely moving recording
<ol>
	<li>Follow the same procedure described in section 2 for recording awake signals from the brain. Plug the wireless headstage by either holding the animal in the experimenter&#39;s arms or feeding the animal with treats to distract it. Provide the acoustic stimulation using external speakers placed close to the animal.</li>
</ol>

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Bone drill</td>
			<td>BBraun</td>
			<td>Elan 4 with&#160; GA861 handpiece</td>
			<td></td>
		</tr>
		<tr>
			<td>Bone drill bit</td>
			<td>BBraun</td>
			<td>Neurocutter GP204R</td>
			<td></td>
		</tr>
		<tr>
			<td>Bonewax</td>
			<td>Ethicon</td>
			<td>W31G</td>
			<td></td>
		</tr>
		<tr>
			<td>Catheter</td>
			<td>Venisystems</td>
			<td>Abbocath 14G</td>
			<td></td>
		</tr>
		<tr>
			<td>Gelfoam</td>
			<td>Pfizer</td>
			<td>Gelfoam</td>
			<td></td>
		</tr>
		<tr>
			<td>Insert speakers</td>
			<td>Etymotic</td>
			<td>Etymotic ER2 insert Earphones</td>
			<td></td>
		</tr>
		<tr>
			<td>Potentiostat</td>
			<td>Gamry Instruments</td>
			<td>Reference 600</td>
			<td></td>
		</tr>
		<tr>
			<td>Pulse Generator</td>
			<td>AM Systems</td>
			<td>Model 2100 Isolated Pulse Stimulator</td>
			<td></td>
		</tr>
		<tr>
			<td>Recording headstage</td>
			<td>Multichannel systems</td>
			<td>W2100-HS32</td>
			<td></td>
		</tr>
		<tr>
			<td>Recording system</td>
			<td>Multichannel systems</td>
			<td>W2100</td>
			<td></td>
		</tr>
		<tr>
			<td>Screwdriver</td>
			<td>Medtronic</td>
			<td>Handle: 001201, Shaft: 8001205</td>
			<td></td>
		</tr>
		<tr>
			<td>Stab knife</td>
			<td>Fine Science Tools</td>
			<td>10316-14</td>
			<td></td>
		</tr>
		<tr>
			<td>Suture wire dermal</td>
			<td>Ethicon</td>
			<td>Vicryl 2-0</td>
			<td></td>
		</tr>
		<tr>
			<td>Suture wire dura mater</td>
			<td>Ethicon</td>
			<td>Mersilk 5-0&#160;</td>
			<td></td>
		</tr>
		<tr>
			<td>Suture wire for lifting dura</td>
			<td>Ethicon</td>
			<td>Prolene 6-0 with BV-1 needle</td>
			<td></td>
		</tr>
		<tr>
			<td>Suture wire subcutaneous</td>
			<td>Ethicon</td>
			<td>Vicryl 4-0</td>
			<td></td>
		</tr>
		<tr>
			<td>Titanium bridge</td>
			<td>Medtronic</td>
			<td>TiMesh 015-2001-4</td>
			<td>Cut out the required size</td>
		</tr>
		<tr>
			<td>Titanium screws</td>
			<td>Medtronic</td>
			<td>9001635, 9001640</td>
			<td></td>
		</tr>
		<tr>
			<td>X-ray system</td>
			<td>GE</td>
			<td>GE OEC 9800 Plus C-Arm</td>
			<td></td>
		</tr>
	</tbody>
</table>

subdural implantation of a soft electrocorticography array for cortical electrophysiology recording in minipigs

Source: Fallegger, F. et al., <a href="https://app.jove.com/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.

<img alt="Figure 1" src="/files/ftp_upload/22756/22756_Figure1.jpg" /> 
Figure 1: Minimally invasive implantation of soft ECoG onto the brain.&#160;(A) Surgical access to the skull, with indication of bregma. (B) Bilateral craniotomy with visible dura mater. (C) Slit durotomy on the first hemisphere. (D) Subdural implantation of soft ECoG and dura mater closure. (E) Slit durotomy on ...

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

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

Begin with an anesthetized minipig with a bilateral craniotomy over the auditory cortex.
Create slits in the dura mater, the brain&#39;s outer covering.
Take a soft electrocorticography array consisting of electrode strips, flexible cables, and a port with connectors.
Insert the strip until the cable edge rests atop the slit, ensuring proper electrode placement on the auditory cortex surface.
Repeat the procedure contralaterally.
Place a titanium bridge over the cables, securing the implant.
Insert the ground wires to establish a recording reference.
Suture the dura mater and replace and secure the bone flaps using titanium bridges.
Attach the footplate and pedestal to the skull. Suture the incision.
Allow the pig to recover and affix a wireless head stage to the pedestal.
The array captures electrical signals generated by cortical neurons, which are transmitted from the head stage to the recording system.
Apply sound stimulation and record the stimulus-induced activity of the auditory neurons.

subdural-implantation-soft-electrocorticography-array-for-cortical

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JoVE Encyclopedia of Experiments

Neuroscience

Neurophysiology

Electrophysiology Techniques

47 Views. Source: Fallegger, F. et al., Subdural Soft Electrocorticography (ECoG) Array Implantation and Long-Term Cortical Recording in Minipigs. 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's auditory cortex and recording both baseline and sound stimulus-evoked neuronal action potentials. 

Video: Subdural Implantation of a Soft Electrocorticography Array for Cortical Electrophysiology Recording in Minipigs - Concept

Microelectrode Guided Implantation of Electrodes into the Subthalamic Nucleus of Rats for Long-term Deep Brain Stimulation

Deep brain stimulation (DBS) is a widely used and effective therapy for several neurologic disorders, such as idiopathic Parkinson&#8217;s disease, dystonia or tremor. DBS is based on the delivery of electrical stimuli to specific deep anatomic structures of the central nervous system. However, the mechanisms underlying the effect of DBS remain enigmatic. This has led to an interest in investigating the impact of DBS in animal models, especially in rats. As DBS is a long-term therapy, research should be focused on molecular-genetic changes of neural circuits that occur several weeks after DBS. Long-term DBS in rats is challenging because the rats move around in their cage, which causes problems in keeping in place the wire leading from the head of the animal to the stimulator. Furthermore, target structures for stimulation in the rat brain are small and therefore electrodes cannot easily be placed at the required position. Thus, a set-up for long-lasting stimulation of rats using platinum/iridium electrodes with an impedance of about 1 M&#937; was developed for this study. An electrode with these specifications allows for not only adequate stimulation but also recording of deep brain structures to identify the target area for DBS. In our set-up, an electrode with a plug for the wire was embedded in dental cement with four anchoring screws secured onto the skull. The wire from the plug to the stimulator was protected by a stainless-steel spring. A swivel was connected to the circuit to prevent the wire from becoming tangled. Overall, this stimulation set-up offers a high degree of free mobility for the rat and enables the head plug, as well as the wire connection between the plug and the stimulator, to retain long-lasting strength.

A method for implanting electrodes into the subthalamic nucleus (STN) of rats is described. Better localization of the STN was achieved by using a microrecording system. Furthermore, a stimulation set-up is presented that is characterized by long-lasting connections between the head of the animal and the stimulator.

Deep brain stimulation (DBS) is a widely used and effective therapy for several neurologic disorders, such as idiopathic Parkinson&#8217;s disease, ...

JoVE Journal

Chronic Implantation of Multiple Flexible Polymer Electrode Arrays

Simultaneous recordings from large populations of individual neurons across distributed brain regions over months to years will enable new avenues of scientific and clinical development. The use of flexible polymer electrode arrays can support long-lasting recording, but the same mechanical properties that allow for longevity of recording make multiple insertions and integration into a chronic implant a challenge. Here is a methodology by which multiple polymer electrode arrays can be targeted to a relatively spatially unconstrained set of brain areas.
The method utilizes thin-film polymer devices, selected for their biocompatibility and capability to achieve long-term and stable&#160;electrophysiologic recording interfaces. The resultant implant allows accurate and flexible targeting of anatomically distant regions, physical stability for months, and robustness to electrical noise. The methodology supports up to sixteen serially inserted devices across eight different anatomic targets. As previously demonstrated, the methodology is capable of recording from 1024 channels. Of these, the 512 channels in this demonstration used for single neuron recording yielded 375 single units distributed across six recording sites. Importantly, this method also can record single units for at least 160 days.
This implantation strategy, including temporarily bracing each device with a retractable silicon insertion shuttle, involves tethering of devices at their target depths to a skull-adhered plastic base piece that is custom-designed for each set of recording targets, and stabilization/protection of the devices within a silicone-filled, custom-designed plastic case. Also covered is the preparation of devices for implantation, and design principles that should guide adaptation to different combinations of brain areas or array designs.

Described below is a method for implantation of multiple polymer electrode arrays across anatomically distant brain regions for chronic electrophysiological recording in freely moving rats. Preparation and surgical implantation are described in detail, with emphasis on design principles to guide adaptation of these methods for use in other species.

Simultaneous recordings from large populations of individual neurons across distributed brain regions over months to years will enable new avenues of ...

Stereotaxic Viral Injection and Gradient-Index Lens Implantation for Deep Brain In Vivo Calcium Imaging

A miniature fluorescence microscope (miniscope) is a potent tool for in vivo calcium imaging from freely behaving animals. It offers several advantages over conventional multi-photon calcium imaging systems: (1) compact; (2) light-weighted; (3) affordable; and (4) allows recording from freely behaving animals. This protocol describes brain surgeries for deep brain in vivo calcium imaging using a custom-developed miniscope recording system. The preparation procedure consists of three steps, including (1) stereotaxically injecting the virus at the desired brain region of a mouse brain to label a specific subgroup of neurons with genetically encoded calcium sensor; (2) implantation of gradient-index (GRIN) lens that can relay calcium image from deep brain region to the miniscope system; and (3) affixing the miniscope holder over the mouse skull where miniscope can be attached later. To perform in vivo calcium imaging, the miniscope is fastened onto the holder, and neuronal calcium images are collected along with simultaneous behavior recordings. The present surgery protocol is compatible with any commercial or custom-built single-photon and two-photon imaging systems for deep brain in vivo calcium imaging.

Stereotaxic Viral Injection and Gradient-Index Lens Implantation for Deep Brain In Vivo Calcium Imaging

Miniscope in vivo calcium imaging is a powerful technique to study neuronal dynamics and microcircuits in freely behaving mice. This protocol describes performing brain surgeries to achieve good in vivo calcium imaging using a miniscope.

A miniature fluorescence microscope (miniscope) is a potent tool for in vivo calcium imaging from freely behaving animals. It offers several advantages ...

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

Transcript

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

Microelectrode Guided Implantation of Electrodes into the Subthalamic Nucleus of Rats for Long-term Deep Brain Stimulation

Chronic Implantation of Multiple Flexible Polymer Electrode Arrays

Stereotaxic Viral Injection and Gradient-Index Lens Implantation for Deep Brain In Vivo Calcium Imaging