This work was funded by the University of Texas at Dallas and the UT Board of Regents. We thank Solomon Golding, Bilaal Hassan, Marghi Jani, and Ching-Tzu Tseng for technical assistance.

All procedures described in this protocol are carried out in accordance with the NIH Guide for the Care and Use of Laboratory Animals and were approved by the Institutional Animal Care and Use Committee of The University of Texas at Dallas.
1. Stimulating cuff electrode fabrication
<ol>
	<li>Prepare the cuff tubing.
	<ol>
		<li>Using a razor blade, cut a piece of polymer tubing 2.5 mm in length. Insert forceps tips or a paper clip through the tubing and use the blade to make a slit lengthwise through the wall of the tubing on one side the cuff.</li>
		<li>Remove the forceps from the tubing and insert a large sewing needle through the midline of the cuff, perpendicular to the long axis. Insert the needle through the slit (top) and into the center of the tubing opposite (bottom). Place the needle into the foam board to pin the cuff in place during the remaining assembly steps.</li>
	</ol>
	</li>
	<li>Place suture for securing cuff closure during implantation.
	<ol>
		<li>Insert the small sewing needle through the wall of the cuff, on the midline, approximately 0.5 mm from the top slit on one side. Insert the needle from interior to exterior to avoid damaging the cuff tubing. Insert a 2 cm length of 6/0 suture through the eye of the needle and pull the needle through the wall of the tubing to thread the suture into the cuff.</li>
		<li>Leaving the thread in place, remove the needle and puncture a second hole through the tubing wall approximately 0.5 mm below the first hole, along the midline of the cuff. Insert the suture through the eye of the needle and pull the needle through the tubing wall to again thread the suture through the cuff.</li>
		<li>Both ends of the suture thread should now be on the exterior side of the cuff. Adjust the suture so that ~1.5 cm extends from the top hole, and ~0.5 mm extends from the bottom hole.</li>
		<li>Apply a small amount of UV cure adhesive to the short end of the suture extending from the lower hole and pull the longer suture end until the lower tail is nearly flush with the exterior wall of the tubing. Use the UV wand to cure the adhesive and hold the suture firmly in place.</li>
		<li>Repeat steps 1.2.1 through 1.2.3 on the opposite side of the cuff.</li>
	</ol>
	</li>
	<li>Place the Platinum:Iridium (Pt:Ir) wire leads.
	<ol>
		<li>Use the small sewing needle to make 4 holes in the cuff wall. Each pair of holes should be placed approximately 0.5&#8211;0.8 mm from the perpendicular midline, with a hole approximately 0.5&#8211;0.8 mm from the top slit on either side of the cuff. 
		CAUTION: For the most consistent and accurate placement of the leads, insert the needle from interior to exterior to make all holes, using the suture placement as a guide.</li>
		<li>Insert the sewing needle again, this time working from exterior to interior, through lead hole 1. Insert approximately 0.5 cm of a 7.5 cm length of Pt:Ir wire through the eye of the needle and pull the needle through the tubing to thread the wire lead through the cuff wall. Adjust the wire so that ~4.5 cm extends on the exterior side of the cuff (Figure 1A).</li>
		<li>Insert the needle through lead hole 1 again, again working exterior-to-interior, and additionally insert the needle through lead hole 2 directly across from lead hole 1. Insert ~0.5 cm of the shorter (interior) end of the Pt:Ir wire through the eye of the needle and pull the needle through the tubing to thread the wire lead through the cuff walls. 
		NOTE: Both ends of the Pt:Ir wire should now be on the exterior side of the cuff, and a wire loop is formed around the slit edge and through lead hole 1 (Figure 1B).</li>
		<li>Repeat steps 1.3.1 through 1.3.3 to place Pt:Ir wire through lead holes 3 and 4.</li>
		<li>Using a butane lighter, carefully remove the insulation from a 5&#8211;6 mm length at the end of Pt:Ir wires extending from lead hole 2 and lead hole 4. 
		CAUTION: Isolate the ends of the leads from the rest of the cuff assembly carefully to avoid damaging to the cuff. Use tools to hold the wires to avoid injury.</li>
		<li>Align the bare wire inside the cuff to place the leads in their final locations. To do this, gently pull on the end of the Pt:Ir wire extending from hole 1 until the uninsulated portion of wire is flush with hole 1. Repeat with the other lead to align the uninsulated end of the wire threaded through lead holes 3 and 4.</li>
		<li>Apply a small amount of UV cure adhesive to the wire loops on the exterior side of the cuff at lead holes 1 and 3. Use the UV wand to cure the adhesive and secure the leads in place.</li>
		<li>Use a small pipette tip to push the uninsulated Pt:Ir wire leads against the interior wall of the cuff. Once the leads are in place, cut the ends of the wires extending from lead holes 2 and 4 so that approximately 1 mm of wire extends beyond the exterior of the cuff wall.</li>
		<li>Fold the 1 mm tails of the wire flat against the exterior surface of the cuff, taking care not to short them together. Apply a small amount of UV cure adhesive to just cover the two tails and cure the adhesive to secure lead placement and provide electrical insulation. 
		CAUTION: It is important to fully cover the externally exposed Pt:Ir surfaces with adhesive to insulate the wires and avoid off-target stimulation.</li>
	</ol>
	</li>
	<li>Secure the Pt:Ir wire leads in place with suture securement.
	<ol>
		<li>Remove the large needle with the cuff assembly from the foam board. Insert a 3 cm length of 6/0 suture through the eye of the needle and pull the needle through the tubing to thread the suture through the bottom of the cuff at the midpoint.</li>
		<li>Switch to the small sewing needle to complete suture threading for Pt:Ir lead securement. Insert the needle through the same midline hole, working again from interior to exterior to avoid deformation of the tubing and the wire leads. Insert the exterior tail of the suture through the eye of the needle and pull the needle through the cuff wall to create a loop of suture around the edge of the cuff (Figure 1C). 
		NOTE: Use forceps and&#160;work under the microscope to ensure the suture is oriented along the long axis of the cuff and lies flat against the tubing. This step ensures the leads remain separated on the interior side of the cuff and are held in place lateral to the cuff midline.</li>
		<li>Create a second loop around the opposite end of the cuff by tying the ends of the suture in a half knot, on the exterior side of the cuff. Ensure the suture runs along the long axis of the cuff and lies flat against the tubing. While holding the knot tight so it lays flat against the tubing, apply a small amount of UV cure adhesive to the half-knot and cure to hold in place.</li>
		<li>Carefully cut the ends of the suture thread as close to the knot as possible. If necessary, use a small amount of additional UV cure adhesive to glue the short ends of suture so they lay flat against the tubing (Figure 1D).</li>
	</ol>
	</li>
	<li>Solder connector pins to the Pt:Ir wire leads.
	<ol>
		<li>Using a butane lighter, remove the insulation from ~3 mm at the end of each of the Pt:Ir wire leads. Solder the cup side of a gold pin (see Table of Materials) to the uninsulated end of each lead.</li>
	</ol>
	</li>
	<li>Test the impedance of the assembled device.
	<ol>
		<li>Connect the gold pins to the inputs of an LCR meter or electrode impedance check module and set the test frequency to 1 kHz. Submerge the cuff tubing (and Pt:Ir stimulation contacts interior to the cuff) in a small beaker filled with saline, taking care to keep the gold lead pins and probe connectors dry. Verify that the assembled cuff has an impedance at 1 kHz of less than 2 k&#937; before proceeding with implantation. 
		NOTE: High impedance often indicates inadequate Pt:Ir surface area exposed, which can arise due to factors such as insufficient removal of insulation, accidental application of adhesive in the cuff interior, broken wire strands, etc. Cuffs should also be inspected for broken or poorly placed wire strands that could result in shorted contacts with long-term use.</li>
	</ol>
	</li>
</ol>
2. Head-cap construction
NOTE: Headcap assembly procedures are similar to those published previously (Childs et al.17), and are summarized here for convenience.
<ol>
	<li>Assemble the headcap17
	<ol>
		<li>Cut two small pieces of 30 AWG wire wrap, one ~13 mm in length and one ~10 mm in length. Strip the ~1.5 mm of insulation off each end of both wires. Solder the pin side of a gold pin to one end of each wire, as close to the cup as possible. Use wire cutters to cut off excess length of pin beyond the solder joint.</li>
		<li>Solder the other ends of the AWG wires to the two central solder cups of a 4-pin microstrip connector.</li>
		<li>Bend the wire headcap leads up toward the connector and place the gold pins flat against the connector, parallel to each other, as shown in Figure 2A. The pin connected to the shorter wire should be placed below the pin connected to the longer wire. Use nail acrylic, dental cement, or UV cure adhesive to secure the headcap leads in place.</li>
	</ol>
	</li>
</ol>
3. Device usage
<ol>
	<li>Implant the cuff electrodes for chronic vagus nerve stimulation. 
	NOTE: All surgical procedures should be performed using sterile or aseptic technique under appropriate anesthesia, in accordance with NIH Guidelines for the Care and Use of Laboratory animals and with local IACUC approval. The following procedures are meant to illustrate a representative usage of the device and are not intended to be comprehensive.
	<ol>
		<li>Place the rat in a stereotaxic frame and make a sagittal incision over the parietal and occipital bones to reveal the skull surface for implantation of the headcap/connector. Carefully drill 4 holes in the skull and place jeweler&#8217;s screws.&#160;Use dental acrylic to secure the headcap to the skull and screws.</li>
		<li>Remove the rat from the stereotaxic frame and lay on its right side. Make a vertical incision in the skin on the left side of the neck, and carefully dissect the left vagus nerve from the carotid artery, located between the sternomastoid and sternohyoid muscles and underneath the omohyoid muscle.</li>
		<li>Tunnel the cuff leads subcutaneously toward the skull. Connect the leads to the headcap using the gold pins.</li>
		<li>Place the vagus nerve inside the cuff and secure the device closed by tying a double knot in the cuff sutures. Be careful to avoid damaging the nerve during implantation by manipulating the nerve with blunt, nonconductive hooks or by grasping connective tissue surrounding the nerve.</li>
		<li>Test the implant by applying stimulation to the device (10 s train of 0.8 mA, 30 Hz, 100 &#181;s biphasic pulses). Proper implantation will result in cessation of breathing and a drop in SpO2 of 5% or more.</li>
		<li>Cover the gold pins and exposed leads with dental acrylic, close wounds with sutures, and clean the incision sites with saline, alcohol, and povidone iodine solution.</li>
		<li>Provide replacement fluids, analgesics and postoperative care in line with NIH guidelines and IACUC approval.</li>
	</ol>
	</li>
	<li>Stimulate the vagus nerve during awake behavior. 
	NOTE: Delivery of VNS as animals perform specific motor tasks has previously been shown to expand the motor map representation of task-relevant musculature. We use this validated paradigm to provide a representative example of device usage, but many other behavioral paradigms and/or stimulation parameters may be relevant to alternative applications. Rats were trained to proficiency on the lever press task used here prior to device implantation. Post-surgery, good performance was again verified prior to VNS delivery: rats performed at least 100 successful trials in two 30 min training sessions per day. VNS was paired with correct lever presses during 10 subsequent training sessions over 5 days.
	<ol>
		<li>Connect the rat to a stimulus generator via implanted head-cap and adjust to appropriate stimulation settings. For VNS-induced reorganization of the motor cortical map, pair each correct lever press with a&#160;single train of 15 biphasic pulses, each with a width of 100 &#181;s and amplitude of 800 &#181;A, delivered at a frequency of 30 Hz.</li>
		<li>A stimulation train is delivered immediately after detection of each successful lever press throughout ten 30 min training sessions. During VNS-delivery, use an oscilloscope to monitor successful delivery of current stimulation.</li>
	</ol>
	</li>
	<li>Validate chronically implanted cuff function.
	<ol>
		<li>Within 24 h of the last VNS-paired training session, use intracranial microstimulation (ICMS) to quantify the functional somatotopic map in the motor cortex19,20,21,22.</li>
		<li>After induction of anesthesia for ICMS mapping of the motor cortex, validate cuff function again by applying a 10 s train of 30 Hz, 0.8 mA current stimulation (100 &#181;s biphasic pulses), which should result in a cessation of breathing and reduction in SpO2 levels of at least 5%, consistent with the Hering-Breuer reflex. 
		NOTE: Depending on the application, cuff function may be considered acceptable if a reliable SpO2 drop of less than 5% is observed, or if higher current amplitudes (up to 1.6 mA) reliably produce at least a 5% reduction in SpO2. Failure to observe a cessation of breathing and/or a reliable decrease in SpO2 is indicative of implant failure.</li>
	</ol>
	</li>
</ol>

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The authors have nothing to disclose.

Here we describe a simple, low-cost approach for assembly of chronically implantable stimulating cuff electrodes for use in rodents, facilitating preclinical investigations of this emerging therapy. This simplified method requires no specialized training or equipment, and uses a small number of tools and supplies that are easily accessible to most research labs, reducing both the monetary and labor costs of device manufacture compared to other approaches16,26<sup...

Recently, the demand for chronically implantable cuff electrodes for stimulation of peripheral nerves has grown, as studies increasingly demonstrate the preclinical usefulness of this technique for the treatment of numerous inflammatory diseases1,2,3 and neurological disorders4,5,6,7,8,9,10,11,12,13,14,15. Chronic VNS, for example, has been shown to enhance neocortical plasticity in a variety of learning contexts, improving motor rehabilitation4,5,6,7,8, extinction learning10,11,12,13,14, and sensory discrimination15. Commercially available peripheral nerve cuff electrodes are often associated with extended times for order fulfillment and relatively high costs, which can limit their accessibility. Alternatively, protocols for &#8220;in-house&#8221; fabrication of chronically implantable cuff electrodes remain limited, and rodent anatomy presents particular challenges due to their small size. Current protocols for constructing cuff electrodes for chronic rodent experiments often require the use of complex equipment and techniques, as well as extensively trained personnel. In this protocol, we demonstrate a simplified approach to cuff electrode fabrication based on previously published and widely used methods16,17. We validate the functionality of our chronically implanted electrodes in rats by demonstrating that, at the time of cuff implantation around the left cervical vagus nerve, stimulation applied to the cuff electrodes successfully produced a cessation of breathing and drop in SpO2. Stimulation of afferent pulmonary receptor vagal fibers is known to engage the Hering-Breuer reflex, in which the inhibition of several respiratory nuclei in the brainstem results in the suppression inspiration18. Thus, cessation of breathing consistent with the Hering-Breuer reflex, and the resulting drop in SpO2, provide a straightforward test for proper electrode implantation and cuff function in anesthetized rats. To validate the long-term functionality of chronically implanted cuff electrodes, reflex responses were measured at the time of implantation and compared to the responses obtained in the same animals six weeks after implantation. A second group of rats was implanted with VNS cuff electrodes after behavioral training on a lever pressing task. In these rats, VNS paired with correct task performance produced reorganization of the cortical motor map, consistent with previously published studies19,20,21,22. At the time of motor cortical mapping under anesthesia, which occurred 5&#8211;10 weeks after device implantation, we further validated cuff function in VNS-treated animals by confirming that VNS successfully induced a cessation of breathing and a greater than 5% drop in SpO2.
The recently published protocols from Childs et al.17 and Rios et al.16 provide a well-validated starting point for a simplified cuff electrode fabrication approach, as this popular method has been utilized by multiple labs conducting chronic VNS studies in rodents1,2,3,4,5,6,7,8,9,10,11. The original method involves several high-precision steps for manipulating the fine microwires such that cuff electrode fabrication takes over an hour to complete, and extensive training to perform reliably. The simplified approach described here requires significantly fewer materials and tools and can be completed in under one hour by minimally trained personnel.

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			<td height="21" style="height:21px;width:441px;">Biocompatible polyurethane-based polymer tubing, 0.080&#34; OD x 0.040&#34; ID</td>
			<td style="width:178px;">Braintree Scientific</td>
			<td style="width:174px;">MRE080 36 FT</td>
			<td style="width:542px;"></td>
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		<tr height="21">
			<td height="21" style="height:21px;">Dissecting microscope</td>
			<td style="width:178px;">AM Scopes</td>
			<td style="width:174px;">#SM-6T-FRL</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Fine Serrated Scissors, straight, 22mm cutting edge</td>
			<td style="width:178px;">Fine Science Tools</td>
			<td style="width:174px;">#14058-09</td>
			<td>for cutting Pt/Ir wire and suture thread</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Forceps, #5 Dumont forceps, straight, 11 cm, 0.1 x 0.06 mm tip</td>
			<td style="width:178px;">Fine Science Tools</td>
			<td style="width:174px;">#11626-11</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Forceps, ceramic tipped forceps, 0.3 mm x 30 mm tips</td>
			<td style="width:178px;">Electron Microscopy Sciences</td>
			<td style="width:174px;">#78127-71</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Gold Pins, PCB Press Fit Socket</td>
			<td style="width:178px;">Mill-Max</td>
			<td style="width:174px;">#1001-0-15-15-30-27-04-0</td>
			<td>or similar small pins for connecting cuff leads to headcap</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Isobutane lighter</td>
			<td style="width:178px;">BIC</td>
			<td style="width:174px;">#LCP21-AST</td>
			<td>for de-insulating Pt/Ir wire</td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;">Micro strip connector with latch, 4-pin</td>
			<td style="width:178px;">Omnetics</td>
			<td style="width:174px;">A24002-004 / PS1-04-SS-LT</td>
			<td></td>
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			<td height="21" style="height:21px;">Pipette tip, 10 uL</td>
			<td style="width:178px;">VWR</td>
			<td>89079-464</td>
			<td></td>
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		<tr height="21">
			<td height="21" style="height:21px;">Platinum-Iridium (90/10%) Wire, 0.001&#34; (diameter) x 9 strands, PTFE insulated</td>
			<td style="width:178px;">Sigmund Cohn</td>
			<td style="width:174px;">10IR9/49T</td>
			<td></td>
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		<tr height="21">
			<td height="21" style="height:21px;">Razor Blade, Single Edge, Surgical Carbon Steel No.9</td>
			<td style="width:178px;">VWR</td>
			<td style="width:174px;">#55411-050</td>
			<td>for cutting MicroRenathane tubing</td>
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		<tr height="21">
			<td height="21" style="height:21px;">Sewing needle, ca. 4.0 cm length x 0.7 mm diameter (size 6-7)</td>
			<td style="width:178px;">Singer</td>
			<td style="width:174px;">00276</td>
			<td>Smaller needle for threading Pt/Ir wire</td>
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		<tr height="21">
			<td height="21" style="height:21px;">Sewing needle, ca. 4.5 cm length x 0.8 mm diameter (size 2-3)</td>
			<td style="width:178px;">Singer</td>
			<td style="width:174px;">00276</td>
			<td>Larger needle for pinning cuff during assembly and for threading suture</td>
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		<tr height="21">
			<td height="21" style="height:21px;">Small foam board</td>
			<td style="width:178px;">Juvo+/Amazon</td>
			<td>B07C9637SJ</td>
			<td>for fabrication platform; our dimensions are ca. 2.5&#34; x 3.5&#34; x 1&#34; (L x W x H)</td>
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		<tr height="21">
			<td height="21" style="height:21px;">Solder, multicore lead-free, 0.38mm diameter</td>
			<td style="width:178px;">Loctite/Multicore</td>
			<td style="width:174px;">#796037</td>
			<td></td>
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		<tr height="21">
			<td height="21" style="height:21px;">Soldering station</td>
			<td style="width:178px;">Weller</td>
			<td style="width:174px;">WES51</td>
			<td>or similar soldering iron compatible with long conical tips (this part has been discontinued)</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Soldering tip, long conical, 0.01&#34; / 0.4 mm</td>
			<td style="width:178px;">Weller</td>
			<td style="width:174px;">1UNF8</td>
			<td></td>
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		<tr height="21">
			<td height="21" style="height:21px;">Suture, nonabsorbable braided silk ,size 6/0</td>
			<td style="width:178px;">Fine Science tools</td>
			<td style="width:174px;">#18020-60</td>
			<td></td>
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		<tr height="21">
			<td height="21" style="height:21px;">UV (405 nm) spot light</td>
			<td style="width:178px;">Henkel/Loctite</td>
			<td style="width:174px;">#2182207</td>
			<td></td>
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		<tr height="21">
			<td height="21" style="height:21px;">UV Light Cure Adhesive 25 ml</td>
			<td style="width:178px;">Henkel/Loctite</td>
			<td style="width:174px;">AA 3106</td>
			<td>or similar biocompatible UV cure adhesive</td>
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		<tr height="21">
			<td height="21" style="height:21px;">Wire wrapping wire, 30 AWG</td>
			<td style="width:178px;">Digikey</td>
			<td style="width:174px;">K396-ND</td>
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preparation of peripheral nerve stimulation electrodes for chronic implantation in rats

Peripheral nerve cuff electrodes have long been used in the neurosciences and related fields for stimulation of, for example, vagus or sciatic nerves. Several recent studies have demonstrated the effectiveness of chronic VNS in enhancing central nervous system plasticity to improve motor rehabilitation, extinction learning, and sensory discrimination. Construction of chronically implantable devices for use in such studies is challenging due to rats&#8217; small size, and typical protocols require extensive training of personnel and time-consuming microfabrication methods. Alternatively, commercially available implantable cuff electrodes can be purchased at a significantly higher cost. In this protocol, we present a simple, low-cost method for construction of small, chronically implantable peripheral nerve cuff electrodes for use in rats. We validate the short and long-term reliability of our cuff electrodes by demonstrating that VNS in ketamine/xylazine anesthetized rats produces decreases in breathing rate consistent with activation of the Hering-Breuer reflex, both at the time of implantation and up to 10 weeks after device implantation. We further demonstrate the suitability of the cuff electrodes for use in chronic stimulation studies by pairing VNS with skilled lever press performance to induce motor cortical map plasticity.

Vagus nerve cuff electrodes and headcaps were chronically implanted in rats according to previously published surgical procedures17,19,20,21,22. Prior to implantation, impedance at 1 kHz was measured across the cuff leads with the cuff tubing submerged in saline (impedance = 1.2 &#177; 0.17 k&#937; [mean &#177; std]; N = 9). Only cuffs with impedances less tha...

Watch this Scientific Journal Video about Preparation of Peripheral Nerve Stimulation Electrodes for Chronic Implantation in Rats at JoVE.com

Preparation of Peripheral Nerve Stimulation Electrodes for Chronic Implantation in Rats

Existing approaches for constructing chronically implantable peripheral nerve cuff electrodes for use in small rodents often require specialized equipment and/or highly trained personnel. In this protocol we demonstrate a simple, low-cost approach for fabricating chronically implantable cuff electrodes, and demonstrate their effectiveness for vagus nerve stimulation (VNS) in rats.

Peripheral nerve cuff electrodes have long been used in the neurosciences and related fields for stimulation of, for example, vagus or sciatic nerves. ...

preparation-peripheral-nerve-stimulation-electrodes-for-chronic

Research

JoVE Journal

Neuroscience

7.5K Views.  University of Texas at Dallas. Existing approaches for constructing chronically implantable peripheral nerve cuff electrodes for use in small rodents often require specialized equipment and/or highly trained personnel. In this protocol we demonstrate a simple, low-cost approach for fabricating chronically implantable cuff electrodes, and demonstrate their effectiveness for vagus nerve stimulation (VNS) in rats.

Video: Preparation of Peripheral Nerve Stimulation Electrodes for Chronic Implantation in Rats

Surgical Implantation of Chronic Neural Electrodes for Recording Single Unit Activity and Electrocorticographic Signals

The success of long-term electrophysiological recordings often depends on the quality of the implantation surgery. Here we provide useful information for surgeons who are learning the process of implanting electrode systems. We demonstrate the implantation procedure of both a penetrating and a surface electrode. The surgical process is described from start to finish, including detailed descriptions of each step throughout the procedure. It should also be noted that this video guide is focused towards procedures conducted in rodent models and other small animal models. Modifications of the described procedures are feasible for other animal models.

We provide useful information for surgeons who are learning the process of implanting chronic neural recording electrodes. Techniques for both penetrating and surface electrode systems are described in a rodent animal model.

The success of long-term electrophysiological recordings often depends on the quality of the implantation surgery.  Here we provide useful information for ...

Fiber-optic Implantation for Chronic Optogenetic Stimulation of Brain Tissue

Elucidating patterns of neuronal connectivity has been a challenge for both clinical and basic neuroscience. Electrophysiology has been the gold standard for analyzing patterns of synaptic connectivity, but paired electrophysiological recordings can be both cumbersome and experimentally limiting. The development of optogenetics has introduced an elegant method to stimulate neurons and circuits, both in vitro1 and in vivo2,3. By exploiting cell-type specific promoter activity to drive opsin expression in discrete neuronal populations, one can precisely stimulate genetically defined neuronal subtypes in distinct circuits4-6. Well described methods to stimulate neurons, including electrical stimulation and/or pharmacological manipulations, are often cell-type indiscriminate, invasive, and can damage surrounding tissues. These limitations could alter normal synaptic function and/or circuit behavior. In addition, due to the nature of the manipulation, the current methods are often acute and terminal. Optogenetics affords the ability to stimulate neurons in a relatively innocuous manner, and in genetically targeted neurons. The majority of studies involving in vivo optogenetics currently use a optical fiber guided through an implanted cannula6,7; however, limitations of this method include damaged brain tissue with repeated insertion of an optical fiber, and potential breakage of the fiber inside the cannula. Given the burgeoning field of optogenetics, a more reliable method of chronic stimulation is necessary to facilitate long-term studies with minimal collateral tissue damage. Here we provide our modified protocol as a video article to complement the method effectively and elegantly described in Sparta et al.8 for the fabrication of a fiber optic implant and its permanent fixation onto the cranium of anesthetized mice, as well as the assembly of the fiber optic coupler connecting the implant to a light source. The implant, connected with optical fibers to a solid-state laser, allows for an efficient method to chronically photostimulate functional neuronal circuitry with less tissue damage9 using small, detachable, tethers. Permanent fixation of the fiber optic implants provides consistent, long-term in vivo optogenetic studies of neuronal circuits in awake, behaving mice10 with minimal tissue damage.

The development of optogenetics now provides the means to precisely stimulate genetically defined neurons and circuits, both in vitro and in vivo. Here we describe the assembly and implantation of a fiber optic for chronic photostimulation of brain tissue.

Elucidating patterns of neuronal connectivity has been a challenge for both clinical and basic neuroscience. Electrophysiology has been the gold standard ...

Transplantation of Olfactory Ensheathing Cells to Evaluate Functional Recovery after Peripheral Nerve Injury

Olfactory ensheathing cells (OECs) are neural crest cells which allow growth and regrowth of the primary olfactory neurons. Indeed, the primary olfactory system is characterized by its ability to give rise to new neurons even in adult animals. This particular ability is partly due to the presence of OECs which create a favorable microenvironment for neurogenesis. This property of OECs has been used for cellular transplantation such as in spinal cord injury models. Although the peripheral nervous system has a greater capacity to regenerate after nerve injury than the central nervous system, complete sections induce misrouting during axonal regrowth in particular after facial of laryngeal nerve transection. Specifically, full sectioning of the recurrent laryngeal nerve (RLN) induces aberrant axonal regrowth resulting in synkinesis of the vocal cords. In this specific model, we showed that OECs transplantation efficiently increases axonal regrowth.
OECs are constituted of several subpopulations present in both the olfactory mucosa (OM-OECs) and the olfactory bulbs (OB-OECs). We present here a model of cellular transplantation based on the use of these different subpopulations of OECs in a RLN injury model. Using this paradigm, primary cultures of OB-OECs and OM-OECs were transplanted in Matrigel after section and anastomosis of the RLN. Two months after surgery, we evaluated transplanted animals by complementary analyses based on videolaryngoscopy, electromyography (EMG), and histological studies. First, videolaryngoscopy allowed us to evaluate laryngeal functions, in particular muscular cocontractions phenomena. Then, EMG analyses demonstrated richness and synchronization of muscular activities. Finally, histological studies based on toluidine blue staining allowed the quantification of the number and profile of myelinated fibers.
All together, we describe here how to isolate, culture, identify and transplant OECs from OM and OB after RLN section-anastomosis and how to evaluate and analyze the efficiency of these transplanted cells on axonal regrowth and laryngeal functions.

Olfactory ensheathing cells (OECs) are neural crest cells which allow growth of the primary olfactory neurons. This specific property can be used for cellular transplantation. We present here a model of cellular transplantation based on the use of OECs in a laryngeal nerve injury model.

Olfactory ensheathing cells (OECs) are neural crest cells which allow growth and regrowth of the primary olfactory neurons. Indeed, the primary olfactory ...

Optogenetic Stimulation of the Auditory Nerve

Direct electrical stimulation of spiral ganglion neurons (SGNs) by cochlear implants (CIs) enables open speech comprehension in the majority of implanted deaf subjects1-6. Nonetheless, sound coding with current CIs has poor frequency and intensity resolution due to broad current spread from each electrode contact activating a large number of SGNs along the tonotopic axis of the cochlea7-9. Optical stimulation is proposed as an alternative to electrical stimulation that promises spatially more confined activation of SGNs and, hence, higher frequency resolution of coding. In recent years, direct infrared illumination of&#160;the cochlea has been used to evoke responses in the auditory nerve10. Nevertheless it requires higher energies than electrical stimulation10,11 and uncertainty remains as to the underlying mechanism12. Here we describe a method based on optogenetics to stimulate SGNs with low intensity blue light, using transgenic mice with neuronal expression of channelrhodopsin 2 (ChR2)13 or virus-mediated expression of the ChR2-variant CatCh14. We used micro-light emitting diodes (&#181;LEDs) and fiber-coupled lasers to stimulate ChR2-expressing SGNs through a small artificial opening (cochleostomy) or the round window. We assayed the responses by scalp recordings of light-evoked potentials (optogenetic auditory brainstem response: oABR) or by microelectrode recordings from the auditory pathway and compared them with acoustic and electrical stimulation.

Cochlear implants (CIs) enable hearing by direct electrical stimulation of the auditory nerve. However, poor frequency and intensity resolution limits the quality of hearing with CIs. Here we describe optogenetic stimulation of the auditory nerve in mice as an alternative strategy for auditory research and developing future CIs.

Direct electrical stimulation of spiral ganglion neurons (SGNs) by cochlear implants (CIs) enables open speech comprehension in the majority of implanted ...

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, ...

Fabrication of High Contact-Density, Flat-Interface Nerve Electrodes for Recording and Stimulation Applications

Many attempts have been made to manufacture multi-contact nerve cuff electrodes that are safe, robust and reliable for long term neuroprosthetic applications. This protocol describes a fabrication technique of a modified cylindrical nerve cuff electrode to meet these criteria. Minimum computer-aided design and manufacturing (CAD and CAM) skills are necessary to consistently produce cuffs with high precision (contact placement 0.51 &#177; 0.04 mm) and various cuff sizes. The precision in spatially distributing the contacts and the ability to retain a predefined geometry accomplished with this design are two criteria essential to optimize the cuff&#39;s interface for selective recording and stimulation. The presented design also maximizes the flexibility in the longitudinal direction while maintaining sufficient rigidity in the transverse direction to reshape the nerve by using materials with different elasticities. The expansion of the cuff&#39;s cross sectional area as a result of increasing the pressure inside the cuff was observed to be 25% at 67 mm Hg. This test demonstrates the flexibility of the cuff and its response to nerve swelling post-implant. The stability of the contacts&#39; interface and recording quality were also examined with contacts&#39; impedance and signal-to-noise ratio metrics from a chronically implanted cuff (7.5 months), and observed to be 2.55 &#177; 0.25 k&#8486; and 5.10 &#177; 0.81 dB respectively.

This article provides a detailed description on the fabrication process of a high contact-density flat interface nerve electrode (FINE). This electrode is optimized for recording and stimulating neural activity selectively within peripheral nerves.

Many attempts have been made to manufacture multi-contact nerve cuff electrodes that are safe, robust and reliable for long term neuroprosthetic ...

Chronic Transcranial Electrical Stimulation and Intracortical Recording in Rats

Transcranial electrical stimulation (TES) is a powerful and relatively simple approach to diffusely influence brain activity either randomly or in a closed-loop event-triggered manner. Although many studies are focusing on the possible benefits and side-effects of TES in healthy and pathologic brains, there are still many fundamental open questions regarding the mechanism of action of the stimulation. Therefore, there is a clear need for a robust and reproducible method to test the acute and the chronic effects of TES in rodents. TES can be combined with regular behavioral, electrophysiological, and imaging techniques to investigate neuronal networks in vivo. The implantation of transcranial stimulation electrodes does not impose extra constraints on the experimental design while it offers a versatile, flexible tool to manipulate brain activity. Here we provide a detailed, step-by-step protocol to fabricate and implant transcranial stimulation electrodes to influence brain activity in a temporally constrained manner for months.

This detailed protocol describes transcranial stimulation electrode placement on the temporal bone in order to investigate the short- and long-term effects of transcranial electrical stimulation in freely moving rats.

Transcranial electrical stimulation (TES) is a powerful and relatively simple approach to diffusely influence brain activity either randomly or in a ...

Chronic Implantation of Whole-cortical Electrocorticographic Array in the Common Marmoset

Electrocorticography (ECoG) allows the monitoring of electrical field potentials from the cerebral cortex with high spatiotemporal resolution. Recent development of thin, flexible ECoG electrodes has enabled conduction of stable recordings of large-scale cortical activity. We have developed a whole-cortical ECoG array for the common marmoset. The array continuously covers almost the entire lateral surface of cortical hemisphere, from the occipital pole to the temporal and frontal poles, and it captures whole-cortical neural activity in one shot. This protocol describes a chronic implantation procedure of the array in the epidural space of the marmoset brain. Marmosets have two advantages regarding ECoG recordings, one being the homologous organization of anatomical structures in humans and macaques, including frontal, parietal, and temporal complexes. The other advantage is that the marmoset brain is lissencephalic and contains a large number of complexes, which are more difficult to access in macaques with ECoG, that are exposed to the brain surface.These features allow direct access to most cortical areas beneath the surface of the brain. This system provides an opportunity to investigate global cortical information processing with high resolutions at a sub-millisecond order in time and millimeter order in space.

We have developed a whole-cortical electrocorticographic array for the common marmoset that continuously covers almost the entire lateral surface of cortex, from the occipital pole to the temporal and frontal poles. This protocol describes a chronic implantation procedure of the array in the epidural space of the marmoset brain.

Electrocorticography (ECoG) allows the monitoring of electrical field potentials from the cerebral cortex with high spatiotemporal resolution. Recent ...

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 ...

Preparation and Implantation of Electrodes for Electrically Kindling VGAT-Cre Mice to Generate a Model for Temporal Lobe Epilepsy

It was discovered that electrical kindling of VGAT-Cre mice led to the spontaneous motor and electrographic seizures. A recent paper focused on how unique VGAT-Cre mice were used in developing spontaneous recurring seizures (SRS) after kindling and a likely mechanism - insertion of Cre into the VGAT gene - disrupted its expression and reduced GABAergic tone. The present study extends these observations to a larger cohort of mice, focusing on key issues such as how long the SRS continues after kindling and the effect of the animal's sex and age. This report describes the protocols for the following key steps: making headsets with hippocampal depth electrodes for electrical stimulation and for reading the electroencephalogram; surgery to affix the headset securely on the mouse's skull so that it does not fall off; and key details of the electrical kindling protocol such as duration of the pulse, frequency of train, duration of train, and amount of current injected. The kindling protocol is robust in that it reliably leads to epilepsy in most VGAT-Cre mice, providing a new model to test for novel antiepileptogenic drugs.

This report describes the methods to generate a model of temporal lobe epilepsy based on the electrical kindling of transgenic VGAT-Cre mice. Kindled VGAT-Cre mice may be useful in determining what causes epilepsy and for screening novel therapies.

It was discovered that electrical kindling of VGAT-Cre mice led to the spontaneous motor and electrographic seizures. A recent paper focused on how unique ...