Name: preparation of peripheral nerve stimulation electrodes for chronic implantation in rats
Uploaded: 2020-07-14T13:00:00.000+00:00
Duration: 9 min 39 s
Description: Watch this Scientific Journal Video about Preparation of Peripheral Nerve Stimulation Electrodes for Chronic Implantation in Rats at JoVE.com

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 approaches16,26,27,28. Care is required throughout assembly to avoid excessive application of UV cure adhesive while still ensuring adequate mechanical stability of the sutures and Pt:Ir leads for long-term cuff function. Excessive adhesive complicates device implantation and may irritate surrounding tissue post-surgery, while insufficient adhesive increases the probability that over time the leads may not maintain good contact with the nerve, resulting in decreased device performance or failure. Consistent placement of the de-insulated Pt:Ir wires inside the cuff lumen is also critical for achieving low impedances and good device performance. Care should be taken to properly align the de-insulated wire such that the maximum possible surface of exposed wire sits inside the cuff, while no exposed wire exists externally.
We have validated that our approach produces cuffs of similar size and reliability as those currently in common use by several labs for chronic VNS delivery in rats4,5,6,7,8,9,10,11,12,13,14,15,19,20,21,22. Recent studies indicate that peripheral nerve fibers are similarly recruited using a wide variety of electrode contact sizes and orientations16,29, suggesting that this protocol can be adapted for many experiments requiring peripheral nerve stimulation, and that small variations in lead spacing or surface area that arise from assembly of cuffs by hand will not critically impact most experimental results. During each stimulation session, we monitored the voltage across the cuff leads using an oscilloscope to ensure that the leads were not shorted or broken, but we did not track changes in impedance for specific implanted devices for the duration of the 5&#8211;10 weeks post-implant. One study of a similar implanted device reported that impedance does significantly increase during the first 4 weeks after surgical implantation, presumably as the acute injury stabilizes30. In this study, however, changes in device impedance were not correlated with device performance over 8 weeks of chronic implantation: the authors reported no significant change in the relationship between VNS intensity and compound action potential amplitude over several weeks post-implant. Here, we were similarly able to functionally validate cuff performance after 5&#8211;10 weeks of implantation by (i) verifying that VNS could still evoke a cessation of breathing and drop in SpO2 consistent with the Hering-Breuer reflex, and (ii) replicating prior work demonstrating VNS-induced motor map reorganization. In our own work, we have found induction of the Hering-Breuer reflex to be the most reliable way to validate long-term functionality of implanted VNS cuffs, which may exhibit reduced device performance or failure due to a number of factors unrelated to cuff assembly; these include surgical complications, nerve damage, and/or mechanical damage to the cuff or headcap. Excellent surgical technique and application-specific validation of device functionality is crucial for stable and successful usage of chronically implanted stimulating cuff electrodes.
We have described a simple, inexpensive approach for assembly of peripheral nerve cuff electrodes for chronic implantation in small animals and demonstrated its usefulness for VNS delivery during rat behavioral experiments. VNS is increasingly under investigation for a wide range of clinical indications, including inflammatory diseases such as rheumatoid arthritis1,2 and Crohn&#8217;s disease,31 as well as neurological disorders such as stroke5,6,7,8 and PTSD10,11. This accessible method for making stimulating cuff electrodes should facilitate the use of preclinical rodent models in a variety of translational research studies into the mechanisms and efficacy of VNS. The protocol is easily adaptable, further increasing the versatility of the approach. For example, the diameter and/or length of the polyurethane tubing can be modified to accommodate chronic stimulation experiments in other species or at other peripheral nerve sites (e.g., sciatic, phrenic, or sacral nerves). Alternatively, configurations with additional leads could enable stimulation at multiple sites along the nerve, or could accommodate simultaneous recording of a stimulation-evoked compound action potential.

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.

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

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 than 2 k&#937; in saline were implanted; all cuffs met this criterion (0/9 cuffs excluded). During implantation surgeries, functional validation of all cuffs was performed by testing for a stimulation-induced brief cessation of breathing and subsequent drop in blood oxygen saturation attributed to the Hering-Breuer reflex. To evoke this response, a 10 s train of 30 Hz, 0.8 mA current stimulation (100 &#181;s biphasic pulses) was delivered across the cuff leads. For 9/9 implanted cuffs, we observed a VNS-induced cessation of breathing for the duration of the 10 sec stimulation, which was accompanied by a drop in SpO2 of at least 5% (% change in SpO2 = -10.3 &#177; 3.2%, mean &#177; std; range = -5.7 to -14.5%), confirming cuff function and proper implantation. During initial implantation, we found a significant correlation between initial SpO2 readings and the percent change in SpO2 evoked by VNS (Figure 2B; R2 = 0.60, p = 0.0083, Pearson&#39;s linear correlation), consistent with published literature demonstrating that anesthesia depth impacts the magnitude of the Hering-Breuer reflex23,24. To test the long-term functionality of the chronically implanted cuffs, rats were anesthetized again 6 weeks after device implantation and VNS was applied to evoke the Hering-Breuer reflex response. For 7 of 9 devices, we observed a greater than 5% drop in SpO2 using 10 s trains of 0.8 mA, 30 Hz stimulation (Figure 2C). In these devices, the magnitude of stimulation-evoked change in SpO2 did not differ from that observed at initial implantation, suggesting excellent continued performance of the chronically implanted devices (initial % change in SpO2 = -9.7 &#177; 3.4%, final % change in SpO2 = -15.8 &#177; 6.5%, mean &#177; std; p = 0.08, paired t-test). In the remaining 2 devices, increasing the stimulation amplitude to 1.6 mA was sufficient to evoke a reliable reduction in SpO2 of at least 5%, suggesting that these devices continued to function, but that changes in impedance, nerve damage, or cuff orientation over time may have resulted in reduced performance.
To further test the long-term functionality of our chronically implanted stimulating electrodes, a second group of rats was trained on a simplified version of a skilled reaching lever-press task developed by Hays et al. to quantitatively assess forelimb motor performance25. Multiple studies have demonstrated that pairing VNS with correct motor performance on this task results in the expansion of the proximal forelimb representation in primary motor cortex19,20,21,22. In our simplified version of the task, rats were required to reach 2 cm outside the training booth to fully depress a lever, and then to release it within 2 s in order to receive a food reward (Figure 2D). Animals received two 30 min training sessions per day until they achieved stable proficiency on the task (&#62;65% correct, &#62;100 trials/session, for at least 8/10 consecutive sessions). Rats then underwent surgery to implant a stimulating cuff electrode around their left vagus nerve. After recovery from surgery, acclimation to stimulating cables, and return to proficient behavioral performance, rats received an additional 10 training sessions in which VNS (0.5 s train of 0.8 mA, 30 Hz pulses; 100 &#181;s biphasic pulse width), or sham stimulation (no stimulation), was delivered at the time of correct lever release. Within 24 h after the last VNS-paired training session, rats were anesthetized with ketamine/xylazine (80/10 mg/kg, i.p.), cuff electrode function was tested, and cortical motor mapping was performed according to published procedures22. Consistent with prior studies demonstrating that VNS drives expansion of task-relevant motor map representations, VNS treated rats (N = 3) exhibited significantly larger proximal forelimb (PFL) representations than sham treated rats (N = 4) in our study (Figure 2D; PFL % of total map area, mean + SEM: sham = 15.6 &#177; 6.7%, VNS = 38.3 &#177; 1.0%; p = 0.035, 2-sample t-test, test power = 0.8). In all VNS-treated animals, cuff function was validated after induction of anesthesia at the time of mapping, 5&#8211;10 weeks post-implant, by confirming a greater than 5% change in SpO2 occurred in response to VNS (10 s train of 0.8 mA, 30 Hz pulses; 100 &#181;s biphasic pulse width).
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/61128/61128fig1.jpg" /> 
Figure 1: Assembly of the stimulating cuff electrodes. (A) After securing the sutures on either side of the cuff, Pt:Ir wire can be threaded through the cuff wall at hole #1 (white arrowhead) using a sewing needle. (B) Pt:Ir wire is properly threaded and ready for de-insulation after creating a wire loop around the cuff edge and threading the wire again through hole #1 (white arrowhead) and across the cuff through hole #2 (yellow arrowhead). (C) Once both leads are in place, secure the first lead by threading suture through the midline hole and around the cuff edge (yellow arrowhead). (D) Close the loop around the second lead with a half-knot and glue in place to complete cuff assembly. <a href="https://www.jove.com/files/ftp_upload/61128/61128fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/61128/61128fig2.jpg" /> 
Figure 2: Device usage for chronic vagus nerve stimulation in awake behaving rats.&#160;&#160;(A) Headcap assembly. (B) During device implantation, VNS-evoked reductions in SpO2 were correlated with initial SpO2 readings (R2 = 0.602, p = 0.008, Pearson&#8217;s linear correlation). (C) Comparison of VNS-evoked SpO2 drops obtained at device implantation versus at the termination of stimulation experiments 6 weeks later. Lines indicate pairs of measurements for individual rats. Stimulation in panels B and C consisted of a single 10 s train of 100 &#181;s biphasic pulses delivered at 0.8 mA and 30 Hz. (D) Rat with chronically implanted VNS cuff electrodes performing the lever-press task. (E) VNS (0.5 s train of 0.8 mA, 30 Hz, 100 &#181;s biphasic pulses) paired with correct lever-press performance expanded the map representation of task-relevant musculature in motor cortex. Rats that received VNS paired with correct lever press performance (N = 3) exhibited a significantly larger percentage of motor map area devoted to proximal forelimb (PFL) representation compared to rats that received Sham stimulation (N = 4). Dots show PFL representations for individual subjects; error bars indicate SEM. VNS treatment followed by motor cortical mapping was performed 5&#8211;10 weeks post-implantation. <a href="https://www.jove.com/files/ftp_upload/61128/61128fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

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

Nanyang Technological University

Research

JoVE Journal

Neuroscience

7.6K 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

A Wireless, Bidirectional Interface for In Vivo Recording and Stimulation of Neural Activity in Freely Behaving Rats

In vivo electrophysiology is a powerful technique to investigate the relationship between brain activity and behavior at a millisecond and micrometer scale. However, current methods mostly rely on tethered cable recordings or only use unidirectional systems, allowing either recording or stimulation of neural activity, but not at the same time or same target. Here, a new wireless, bidirectional device for simultaneous multichannel recording and stimulation of neural activity in freely behaving rats is described. The system operates through a single portable head stage that both transmits recorded activity and can be targeted in real-time for brain stimulation using a telemetry-based multichannel software. The head stage is equipped with a preamplifier and a rechargeable battery, allowing stable long-term recordings or stimulation for up to 1 h. Importantly, the head stage is compact, weighs 12 g (including battery) and thus has minimal impact on the animal&#180;s behavioral repertoire, making the method applicable to a broad set of behavioral tasks. Moreover, the method has the major advantage that the effect of brain stimulation on neural activity and behavior can be measured simultaneously, providing a tool to assess the causal relationships between specific brain activation patterns and behavior. This feature makes the method particularly valuable for the field of deep brain stimulation, allowing precise assessment, monitoring, and adjustment of stimulation parameters during long-term behavioral experiments. The applicability of the system has been validated using the inferior colliculus as a model structure.

A Wireless, Bidirectional Interface for In Vivo Recording and Stimulation of Neural Activity in Freely Behaving Rats

A wireless, bidirectional system for multi-channel neural recordings and stimulation in freely behaving rats is introduced. The system is light and compact, thus having minimal impact on the animal&#180;s behavioral repertoire. Moreover, this bidirectional system provides a sophisticated tool to assess causal relationships between brain activation patterns and behavior.

In vivo electrophysiology is a powerful technique to investigate the relationship between brain activity and behavior at a millisecond and micrometer ...

Behavior

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 ...

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, ...

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 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 ...

Implantation and Control of Wireless, Battery-free Systems for Peripheral Nerve Interfacing

Peripheral nerve interfaces are frequently used in experimental neuroscience and regenerative medicine for a wide variety of applications. Such interfaces can be sensors, actuators, or both. Traditional methods of peripheral nerve interfacing must either tether to an external system or rely on battery power that limits the time frame for operation. With recent developments of wireless, battery-free, and fully implantable peripheral nerve interfaces, a new class of devices can offer capabilities that match or exceed those of their wired or battery-powered precursors. This paper describes methods to (i) surgically implant and (ii) wirelessly power and control this system in adult rats. The sciatic and phrenic nerve models were selected as examples to highlight the versatility of this approach. The paper shows how the peripheral nerve interface can evoke compound muscle action potentials (CMAPs), deliver a therapeutic electrical stimulation protocol, and incorporate a conduit for the repair of peripheral nerve injury. Such devices offer expanded treatment options for single-dose or repeated dose therapeutic stimulation and can be adapted to a variety of nerve locations.

This is a protocol for the surgical implantation and operation of a wirelessly powered interface for peripheral nerves. We demonstrate the utility of this approach with examples from nerve stimulators placed on either the rat sciatic or phrenic nerve.

Peripheral nerve interfaces are frequently used in experimental neuroscience and regenerative medicine for a wide variety of applications. Such interfaces ...

Preparation of Rat Sciatic Nerve for Ex Vivo Neurophysiology

Ex vivo preparations enable the study of many neurophysiological processes in isolation from the rest of the body while preserving local tissue structure. This work describes the preparation of rat sciatic nerves for ex vivo neurophysiology, including buffer preparation, animal procedures, equipment setup and neurophysiological recording. This work provides an overview of the different types of experiments possible with this method. The outlined method aims to provide 6 h of stimulation and recording on extracted peripheral nerve tissue in tightly controlled conditions for optimal consistency in results. Results obtained using this method are A-fibre compound action potentials (CAP) with peak-to-peak amplitudes in the millivolt range over the entire duration of the experiment. CAP amplitudes and shapes are consistent and reliable, making them useful to test and compare new electrodes to existing models, or the effects of interventions on the tissue, such as the use of chemicals, surgical alterations, or neuromodulatory stimulation techniques. Both conventional commercially available cuff electrodes with platinum-iridium contacts and custom-made conductive elastomer electrodes were tested and gave similar results in terms of nerve stimulus strength-duration response.

Preparation of Rat Sciatic Nerve for Ex Vivo Neurophysiology

This protocol describes the preparation of rat whole sciatic nerve tissue for ex vivo electrophysiological stimulation and recording in an environmentally-regulated, two-compartment, perfused saline bath.

Ex vivo preparations enable the study of many neurophysiological processes in isolation from the rest of the body while preserving local tissue structure. ...

 Fabricating Epimysial EMG Electrodes for Preclinical Rodent Models 

Development of a Low-cost Epimysial Electromyography Electrode: A Simplified Workflow for Fabrication and Testing

Electromyography (EMG) is a valuable diagnostic tool for detecting neuromuscular abnormalities. Implantable epimysial electrodes are commonly used to measure EMG signals in preclinical models. Although classical resources exist describing the principles of epimysial electrode fabrication, there is a sparsity of illustrative information translating electrode theory to practice. To remedy this, we provide an updated, easy-to-follow guide on fabricating and testing a low-cost epimysial electrode. 
Electrodes were made by folding and inserting two platinum-iridium foils into a precut silicone base to form the contact surfaces. Next, coated stainless steel wires were welded to each contact surface to form the electrode leads. Lastly, a silicone mixture was used to seal the electrode. Ex vivo testing was conducted to compare our custom-fabricated electrode to an industry standard electrode in a saline bath, where high levels of signal agreement (sine [intraclass correlation - ICC= 0.993], square [ICC = 0.995], triangle [ICC = 0.958]), and temporal-synchrony (sine [r = 0.987], square [r = 0.990], triangle [r= 0.931]) were found across all waveforms. Low levels of electrode impedance were also quantified via electrochemical impedance spectroscopy. 
An in vivo performance assessment was also conducted where the vastus lateralis muscle of a rat was surgically instrumented with the custom-fabricated electrode and signaling was acquired during uphill and downhill walking. As expected, peak EMG activity was significantly lower during downhill walking (0.008 &plusmn; 0.005 mV) than uphill (0.031 &plusmn; 0.180 mV, p = 0.005), supporting the validity of the device. The reliability and biocompatibility of the device were also supported by consistent signaling during level walking at 14 days and 56 days post implantation (0.01 &plusmn; 0.007 mV, 0.012 &plusmn; 0.007 mV respectively; p &gt; 0.05) and the absence of histological inflammation. Collectively, we provide an updated workflow for the fabrication and testing of low-cost epimysial electrodes.

Author Spotlight: Epimysial Electrode Fabrication and Testing in ACL Injury Studies

Our purpose was to provide an updated, easy-to-follow guide on the fabrication and testing of epimysial electromyography electrodes. To that end, we provide instructions for material sourcing and a detailed walkthrough of the fabrication and testing process.

Electromyography (EMG) is a valuable diagnostic tool for detecting neuromuscular abnormalities. Implantable epimysial electrodes are commonly used to ...

Flexible Multi-Region Neural Recording Device for Epilepsy Research

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

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

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

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

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

Implanting a Wireless Bidirectional Microelectrode System in a Rat Brain

Source: Melo-Thomas, L., et al., <a href="https://app.jove.com/v/56299">A Wireless, Bidirectional Interface for In Vivo Recording and Stimulation of Neural Activity in Freely Behaving Rats.</a> J. Vis. Exp. (2017)
The video demonstrates a procedure for implanting a wireless bidirectional multichannel electrode unit into a rat brain to record and stimulate multichannel neural networks.

All procedures involving animal models have been reviewed by the local institutional animal care committee and the JoVE veterinary review board.
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Preparation of Peripheral Nerve Stimulation Electrodes for Chronic Implantation in Rats

Summary

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Chapters in this video

A Wireless, Bidirectional Interface for In Vivo Recording and Stimulation of Neural Activity in Freely Behaving Rats

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

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

Chronic Transcranial Electrical Stimulation and Intracortical Recording in Rats

Chronic Implantation of Multiple Flexible Polymer Electrode Arrays

Implantation and Control of Wireless, Battery-free Systems for Peripheral Nerve Interfacing

Preparation of Rat Sciatic Nerve for Ex Vivo Neurophysiology

Development of a Low-cost Epimysial Electromyography Electrode: A Simplified Workflow for Fabrication and Testing

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

Implanting a Wireless Bidirectional Microelectrode System in a Rat Brain