Development of the rat abdominal VNS implant was funded by the Defense Advanced Research Projects Agency (DARPA) BTO, under the auspices of Dr. Doug Weber and Dr. Eric Van Gieson through the Space and Naval Warfare Systems Center (Contract No. N66001-15-2-4060). Research reported in this publication was supported by the Bionics Institute Incubation Fund. The Bionics Institute acknowledges the support they receive from the Victorian Government through its Operational Infrastructural Support Program. We would like to thank Mr. Owen Burns for mechanical design, Prof. John B Furness for anatomical expertise, Prof. Robert K Shepherd for peripheral interface, neuromodulation and recording expertise, Ms. Philippa Kammerer and Ms. Amy Morley for animal husbandry and testing, Ms. Fenella Muntz and Dr. Peta Grigsby for their advice on post-operative animal care, and Ms. Jenny Zhou and the electrode fabrication team from NeoBionica for production of the VNS arrays.

All procedures involving animals were approved by the Animal Ethics Committee of St. Vincent&#39;s Hospital (Melbourne) and complied with the Australian Code for the Care and Use of Animals for Scientific Purposes (National Health and Medical Research Council of Australia) and the Prevention of Cruelty to Animals (1986) Act. In total, 24 female Dark Agouti rats (8-9 weeks old) were used for this study. The experimental groups consisted of: a normal cohort (n = 8) that received no collagen injection or VNS implant; an unstimulated disease cohort (n = 8) that received an implant and a collagen injection (no electrophysiological tests conducted); and a stimulated disease cohort (n = 8) that received an implant, a collagen injection, electrophysiological testing and VNS therapy. Implantation surgery occurred 5 days prior to collagen injection, and habituation for VNS therapy started 4 days after collagen injection and occurred over 7 days. VNS therapy was applied from day 11 to 17 (inclusive) following collagen injection15. For the stimulated disease cohort, electrophysiological testing was performed immediately after the implantation surgery under anesthesia, on the day of collagen injection, 10 days after collagen injection, and the day of termination (17 days after collagen injection).
1. Sonication and sterilization of electrode array
<ol>
	<li>Set the ultrasonic cleaner to a frequency of 80 kHz and fill the ultrasonic tank with tap water. Submerge the electrode array in the cleaning solution in a clean plastic container and place it in the ultrasonic tank. 
	NOTE: The cleaning solution and sonication time to be used for each step are summarized in Table 1. Use a clean container for each step.</li>
	<li>Place the sonicated electrode array into a sterilization bag using clean forceps sonicated with 0.5% liquid cleaning solution in distilled water and rinsed in distilled water. Autoclave the electrode array for 45 min with maximum temperature of 130 &#176;C, and let it dry on a clean bench.</li>
</ol>
2. Implantation of electrode array on the abdominal vagus nerve
NOTE: In this study we have used female dark agouti rats (8-9 weeks of age)15. We have also successfully used this protocol to chronically implant adult male Sprague-Dawley rats (10-14 weeks of age)10,16. Surgery is conducted under aseptic conditions, and all instruments, electrode array and consumables such as gauze and cotton tips are sterilized by autoclaving.
<ol>
	<li>Anesthetize the rat in an induction chamber using 3% isoflurane and 1 L/min oxygen. Once there is no pedal reflex to toe pinching, move the rat to the heat mat with a thermostat on the surgical table and place an isoflurane mask over the nose.</li>
	<li>Monitor the respiratory rate and rectal temperature throughout the surgery and adjust the isoflurane level between 1.5% and 2.5% to maintain the respiratory rate between 40 to 62 breaths per min. Adjust the heat mat setting if required, to maintain the rectal temperature range between 35.9 - 37.5 &#176;C.</li>
	<li>Administer analgesia pre-medication subcutaneously using 1 mL syringes with 25G needles (carprofen 5 mg/kg and buprenorphine 0.03 mg/kg subcutaneously) prior to the start of surgery.</li>
	<li>Shave generously around the incision site, including the area along the ventral midline from the xyphoid process to the end of the rib cage, the lumbar aspect of the back along the dorsal midline, and the left-hand side of the body between the forelimb and the hindlimb to allow subcutaneous tunneling of the array.</li>
	<li>Clean the surgical sites in a circular motion three times with alternating rounds of betadine and alcohol and place a surgical drape over the animal. Administer bupivacaine (1-2 mg/kg) subcutaneously using a 1 mL syringe with a 25G needle at the dorsal and ventral incision sites.</li>
	<li>Place the animal in ventral recumbency and make a 2 cm long incision on the back where the percutaneous pedestal will be anchored using a scalpel blade.</li>
	<li>Turn the rat to dorsal recumbency and make a 3 cm incision on the skin along the midline just below the xyphoid process using a scalpel blade. Hold up the skin near the incision site, and using dissecting scissors, blunt-dissect the skin layer from the muscle layer around the incision.</li>
	<li>To allow subcutaneous tunneling of the array from the pedestal to site of implantation, place the animal on its right side, insert a hemostat from the ventral incision and blunt dissect towards the dorsal incision site. Cut off the rim of a needle cap and insert the electrode array to protect it while in transit (Figure 1B). Using hands (wear sterile gloves), tunnel the electrode array under the skin towards the ventral incision.</li>
	<li>To access the esophagus and the vagus nerve, lay the animal in dorsal recumbency again. Make a 3 cm incision on the muscle layer along the midline below the xyphoid process, large enough to expose the whole length of the liver. Avoid damaging the liver during this step.</li>
	<li>Make a smaller incision (less than 1 cm) on the muscle layer laterally (the left side of the animal) to the main ventral incision. Tunnel the electrode array through this small incision using the needle cap used in step 2.8 to insert the array into the abdominal cavity. 
	NOTE: This step reduces tension applied to the main incision site and reduces the risk of sutures bursting.</li>
	<li>Retract the skin and muscle layers to hold the abdominal cavity open. Ensure to keep tissues moist by using cotton tips and gauze soaked in sterile saline to handle the tissue.</li>
	<li>Gently retract the liver by cutting the connective tissue around it using Vannas scissors and placing a retractor over a small piece of gauze soaked in saline for protection. Gently retract the stomach, to allow straightening of the esophagus and overlying vagus nerve, by placing a retractor between the esophagus and the stomach. 
	NOTE: Retractors are made by rounding the pointy end of fishhooks.</li>
	<li>Following exposure of the ventral surface of the esophagus, identify the abdominal vagus nerve and its sub-branches, including the hepatic nerve, the celiac nerve and two gastric branches (Figure 1D).</li>
	<li>Cut the connective tissue securing the abdominal vagus nerve to the esophagus using fine forceps and Vannas scissors and dissect the length of the nerve from just above the hepatic and celiac branches towards the diaphragm. Ensure to not tear, stretch, or pinch the nerve. Place an electrode array next to the nerve to confirm that enough length of the nerve is separated from the connective tissue to fit the array.</li>
	<li>Once connective tissue had been cleared around the nerve, pass the silk sutures (7-0) on the electrode-side of the array cuff under the nerve. Open the cuff of the array and place the nerve carefully into the array channel.</li>
	<li>Ensure that the entire length of the nerve sits inside the array channel. Tie the sutures around the cuff together to securely shut the cuff to ensure the nerve does not slip out of the channel. Trim the sutures.</li>
	<li>Using 7-0 silk suture, suture the tab of the array onto the esophagus to secure the array in place and prevent it from twisting. Avoid damaging the other branches of the vagus nerve or inserting needle too deep into the esophagus smooth muscle.</li>
	<li>Gently remove retractors and ensure all gauze has been removed from the abdominal cavity. Administer 1-2 mL of warm sterile saline using a 1 mL syringe in the abdominal cavity and reposition the liver to the correct position.</li>
	<li>Close the muscle layer with 3-0 silk suture using the simple running suture technique, making secure square knots with at least 3 throws at both ends. Space stitches closely together (approximately 3 mm apart) to prevent complications such as hernia/protrusion of the xyphoid process.</li>
	<li>Use suture to close the incision of the peritoneum together with the incision of the muscle layer, to reduce the chance of tissue adhesion.</li>
	<li>Using an absorbable suture material (Vicryl 4-0), close the skin incision. Use a buried suturing technique such as the running buried vertical mattress suture, or the running buried dermal suture to prevent the animal from removing the suture.</li>
	<li>Turn the animal to ventral recumbency, and using scissors, extend the dorsal incision to 4-5 cm, and blunt dissect between the muscle and the skin layer further so the connector base of the percutaneous connector can sit flat on the muscle layer.</li>
	<li>Using silk 3-0 suture, make 6 to 8 simple interrupted sutures around the connector base to secure it to the muscle layer underneath. Close the skin incision with silk 3-0 suture, using the horizontal mattress suture technique, ensuring secure square knots with at least 3 throws. 
	NOTE: In this step, braided silk sutures are preferred for their ease of handling and their capacity to create more secure knots&#160;compared to monofilament sutures.</li>
	<li>At the completion of surgery, administer Hartmann&#39;s solution subcutaneously (1 mL/100 g/h). Turn off the isoflurane and let the animal recover on a heat mat while running oxygen (1.5 L/min). Once the rat is conscious and fully mobile, return the rat to its home cage, placed on a heat pad, until fully recovered from anesthetic.</li>
	<li>Closely observe the animal&#39;s recovery from isoflurane and ensure that the animal is able to access food and drink. In the next two days, provide subcutaneous administration of post-surgical analgesia (carprofen 5 mg/kg, daily) to alleviate pain. Monitor the animal at least 2x per day and check for evidence of defecation, quality of the coat, activity level and presence of any swelling or discharge from the surgical wounds.</li>
	<li>Record weight of the animal, and in the rare event the animal loses 10% or more, start intensive treatment. Intensive treatment includes subcutaneous administration of&#160;fluids (Hartmann&#39;s solution, 2x 10 mL) each day, providing additional food such as fresh vegetables and dietary gel supplement, and placing half the cage on a heat pad with a thermostat for extra warmth. Increase the frequency of monitoring until the animal recovers. Continue with administration of analgesia (carprofen 5 mg/kg, SQ, daily) if required based on a Grimace scale.</li>
</ol>
3. Electrophysiological testing
NOTE: Recording evoked compound action potentials (ECAPs) confirms appropriate placement of the electrode array on the vagus nerve. Additionally, recording of ECAPs using the electrode array described above provides likely confirmation of electrical activation of vagal C-fibers and suprathreshold VNS10,15.
<ol>
	<li>Measure the common ground impedance of electrodes to assess their integrity and detect any open or short circuits of wires prior to recording ECAPs. Functioning abdominal vagus nerve electrodes in vivo should have impedance values between 4 - 20 k&#8486;.</li>
	<li>Test animals while anaesthetized i.e., immediately after surgery, or awake and freely moving. Perform awake testing at least 2-3 days after the surgery to allow for the surgical skin wounds to heal and stabilize. Gather the equipment required for impedance and electrophysiological testing which include a custom-made stimulator, a data acquisition device, an isolated differential amplifier and a data acquisition and analysis software as listed in the Table of Materials.</li>
	<li>Wrap the animal in a towel if required, connect a cable to the back percutaneous connector, and connect the other end of the cable to a stimulator. To test the common ground impedance of electrodes, apply biphasic current pulses (100 &#181;s per phase and current of 107 &#181;A) between the electrode of interest and all other electrodes on the array.</li>
	<li>Measure the peak voltage at the end of the first phase of the voltage waveform (Vtotal) and calculate total impedance (Ztotal) using Ohm&#39;s law (Z = voltage/current).</li>
	<li>Connect a pair of electrodes to the stimulator and a pair of electrodes to the recording equipment and apply bipolar stimulation to generate ECAPs using the reference electrode of the VNS implant placed under the skin as a reference for the differential recording of ECAPs. Make two sets of recordings averaged from a total of 50 repetitions using the data acquisition and analysis software.</li>
	<li>Use the following settings for measurements. 
	Currents: 0 to 2 mA in 0.1 mA increments; 
	Pulse width: 25 - 200 &#181;s; 
	Interphase gap: 8 - 50 &#181;s; 
	Stimulation rate: 10 - 30 pulses/s; 
	Sampling rate: 100 kHz; 
	Filter: High pass 200 Hz, low pass 2000 Hz, voltage gain 1 x 102.</li>
	<li>Using the data analysis software, analyze the ECAP response by measuring the peak-to-peak voltage of the waveforms within the analysis window (4 - 10 ms post-stimulus, indicated by shading in Figure 3A,B). ECAP threshold is defined as the minimum stimulus current intensity producing a response amplitude of at least 0.1 &#181;Vpeak-peak in both sets of average electrophysiology recordings. A valid response will be repeated for at least two current levels above the threshold, and not present for at least two current levels below the threshold10,15.</li>
</ol>
4. Chronic abdominal VNS in awake rats
NOTE: Abdominal VNS can be applied to awake animals once the surgical wound around the percutaneous connector has healed and stabilized. To reduce any stress response and allow for better data collection, animals are habituated to the testers&#39; handling and stimulation environment, one hours a day over seven days prior to the implantation surgery and commencement of VNS therapy.
<ol>
	<li>Measure the impedance of each electrode as described in step 3.4, prior to applying any VNS. Ensure that the impedance of stimulating electrodes is below 20 k&#8486;.</li>
	<li>Connect a cable to the back percutaneous connector and connect the other end of the cable to a stimulator programmed to apply appropriate stimulation (e.g., 27 Hz, 1.6 mA, 200 &#181;s pulse-width with 50 &#181;s interphase gap, 30 s ON, 2.5 min off15), and turn on the stimulator. 
	NOTE: Although animals are often observed falling asleep during stimulation if appropriately habituated, use a cable with protective outer material such as steel coils where possible to prevent it from being chewed.</li>
	<li>Observe the animal at the start of each VNS therapy session to ensure there is no adverse reaction such as excessive grooming or sudden increase/decrease in activity level in sync with the timing of stimulation.</li>
	<li>Monitor every 30 min to check for twisting or disconnection of the cable. To apply VNS chronically (e.g., 3 h a day over 7 days15), repeat steps 4.1-4.3 at the start of each session. 
	NOTE: The use of a commutator may reduce the chance of cables becoming twisted and could require less frequent monitoring.</li>
</ol>

Abdominal VNS for Treating Inflammatory Conditions in Rat Models

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This research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.

This method of abdominal VNS implant surgery and chronic stimulation of the vagus nerve and recording of ECAPs have been successfully used and well-tolerated for 5 weeks in rats following implantation10,15,16. Retraction of the stomach, liver, and gut to gain a good view of the esophagus and the vagus nerve is one of the key steps in the surgery. Once these organs are retracted, the vagus nerve becomes accessible. Retraction of ...

Vagus nerve stimulation (VNS) delivered at the cervical site in the neck is The United States Food and Drug Administration (FDA)-approved treatment for refractory epilepsy, refractory depression, and post-ischemic stroke rehabilitation1, and European Commission-approved for heart failure in Europe2. Non-invasive cervical VNS is FDA-approved for migraine and headache1. Its application is expected to expand, with recent clinical trials showing efficacy of VNS in other indications such as Crohn&#39;s disease3, rheumatoid arthritis4,5 and impaired glucose tolerance and type 2 diabetes6,7. Although promising, cervical VNS can cause bradycardia and apnea due to off-target activation of the nerve fibers that innervate the lungs and the heart8,9,10. Side effects such as cough, pain, voice alteration, headache and increase in apnea-hypopnea index are commonly reported in patients receiving cervical VNS11,12. Reduction in stimulation strength is a common strategy for reducing these side effects, however reduced charge may limit efficacy of VNS therapy by failing to activate therapeutic fibers11. In support of this hypothesis, the responder rate of patients receiving high intensity stimulation for the treatment of epilepsy was higher than that of patients receiving low intensity stimulation13.
Abdominal VNS is applied on the subdiaphragmatic vagus nerve, above the hepatic and celiac branches14 (Figure 1). Our previous study demonstrated that in rats abdominal VNS does not cause cardiac or respiratory side effects associated with cervical VNS10. Earlier studies also demonstrate anti-inflammatory effects of abdominal VNS in a rat model of inflammatory bowel disease and rheumatoid arthritis10,15 as well as reduction in glycemia in a rat model of type 2 diabetes16. Recently, the abdominal VNS technology has been translated for a first-in-human clinical trial for the treatment of inflammatory bowel disease (NCT05469607).
The peripheral nerve electrode array used to deliver stimulation to the abdominal vagus nerve (WO201909502017) has been custom developed for use in rats, and comprises of two to three platinum electrode pairs placed 4.7 mm apart, supported by a medical-grade silicone elastomer cuff, a suturing tab to anchor the array to the esophagus, a lead wire and a percutaneous connector to be mounted on the lumbar region (Figure 2). The lead wire is tunneled under the skin on the left side of the animal. Multiple electrode pair design allows for electrical stimulation of the nerve as well as recording electrically evoked compound action potentials (ECAPs), which confirms correct placement of the implant onto the nerve and suprathreshold stimulation intensities. Abdominal VNS is well tolerated in freely moving rats for months10,15,16. This allows for assessment of its efficacy on disease models.
This manuscript describes the methods for the electrode array sterilization, abdominal vagus nerve implantation surgery, and chronic stimulation and recording of ECAPs in awake rats for studying the efficacy of abdominal VNS in a variety of disease models. These methods were originally developed for studying the efficacy of abdominal VNS in the rat model of inflammatory bowel disease10 and have also been successfully used for a rat model of rheumatoid arthritis15 and diabetes16.

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			<td>Dumont tweezer, style 5</td>
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			<td>Dumont tweezer, style N7, self-closing</td>
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			<td>EMS72864-D</td>
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			<td>Elmasonic P sonicator</td>
			<td>Elma</td>
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			<td>Hartmann&#39;s solution</td>
			<td>Baxter</td>
			<td>AHB2323</td>
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			<td>Hemostat</td>
			<td>ProSciTech</td>
			<td>TS1322-140</td>
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			<td>HPMC/PAA Moisturising Eye Gel</td>
			<td>Alcon</td>
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			<td>Igor Pro-8 software</td>
			<td>Wavemetrics, Inc</td>
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			<td>Isoflo (Isoflurane)</td>
			<td>Zoetis</td>
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			<td>Isolated differential amplifier</td>
			<td>World Precision Instruments</td>
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			<td>Liquid pyroneg</td>
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			<td>HH12291</td>
			<td>cleaning solution</td>
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			<td>Plastic drape</td>
			<td>Multigate</td>
			<td>22-203</td>
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			<td>Neo-Bionica</td>
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			<td>Ultrapure water</td>
			<td>Merck Millipre</td>
			<td>Milli-Q Direct</td>
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			<td>Zebravet</td>
			<td>UP10SM</td>
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			<td>Vannas scissors</td>
			<td>ProSciTech</td>
			<td>EMS72933-01</td>
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			<td>Vicryl suture 4-0</td>
			<td>Ethicon</td>
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implantation surgery for abdominal vagus nerve stimulation and recording studies in awake rats

Abdominal vagus nerve stimulation (VNS) can be applied to the subdiaphragmatic branch of the vagus nerve of rats. Due to its anatomical location, it does not have any respiratory and cardiac off-target effects commonly associated with cervical VNS. The lack of respiratory and cardiac off-target effects means that the intensity of stimulation does not need to be lowered to reduce side effects commonly experienced during cervical VNS. Few recent studies demonstrate the anti-inflammatory effects of abdominal VNS in rat models of inflammatory bowel disease, rheumatoid arthritis, and glycemia reduction in a rat model of type 2 diabetes. Rat is a great model to explore the potential of this technology because of the well-established anatomy of the vagus nerve, the large size of the nerve that allows easy handling, and the availability of many disease models. Here, we describe the methods for cleaning and sterilizing the abdominal VNS electrode array and surgical protocol in rats. We also describe the technology required for confirmation of suprathreshold stimulation by recording evoked compound action potentials. Abdominal VNS has the potential to offer selective, effective treatment for a variety of conditions, including inflammatory diseases, and the application is expected to expand similarly to cervical VNS.

Recording evoked compound action potentials (ECAPs, Figure 3A,B) immediately after surgery is a technique that can be used to help confirm correct placement of the nerve within the array channel, and that stimulation is efficacious in activating the vagus nerve.
In Figure 3, female dark agouti rats (8-9 weeks of age) were implanted with the VNS electrode array. In rats randomly selected to receive therapeutic stimulat...

Watch this Scientific Journal Video about Author Spotlight: Exploring Abdominal VNS for Inflammatory Conditions at JoVE.com

Author Spotlight: Exploring Abdominal VNS for Inflammatory Conditions

The present protocol describes the surgical technique for implanting an electrode array onto the abdominal vagus nerve in rats, along with methods for chronic electrophysiology testing and stimulation using the implanted device.

Implantation Surgery for Abdominal Vagus Nerve Stimulation and Recording Studies in Awake Rats

Abdominal vagus nerve stimulation (VNS) can be applied to the subdiaphragmatic branch of the vagus nerve of rats. Due to its anatomical location, it does ...

implantation-surgery-for-abdominal-vagus-nerve-stimulation-recording

Research

JoVE Journal

Neuroscience

1.4K Views.  Bionics Institute. The present protocol describes the surgical technique for implanting an electrode array onto the abdominal vagus nerve in rats, along with methods for chronic electrophysiology testing and stimulation using the implanted device.

Video: Author Spotlight: Exploring Abdominal VNS for Inflammatory Conditions

Preparation of an Awake Mouse for Recording Neural Responses and Injecting Tracers

It is well known that anesthesia alters neural response properties in various regions of the brain.13. In the auditory system, fundamental response properties of brainstem neurons including threshold, frequency specificity, and inhibitory sidebands are altered in significant ways under anesthesia1-2. These observations prompted physiologists to seek ways to record from single neurons without the contaminating effects of anesthesia. One result was a decerebrate preparation, where the brainstem was completely transected at the level of the midbrain4. The drawbacks of this preparation are a formidable surgery, the elimination of descending projections from the forebrain, and an inability to use sensory stimulation to examine structures above the midbrain. A different strategy has been to implant electrode arrays chronically to record from single neurons and multiunit clusters while the animal is awake and/or behaving5,6. These techniques however are not compatible with injecting tracer dyes after first electrophysiologically characterizing a brain structure. To avoid altering neural response properties with anesthetics while recording electrophysiological response properties from single neurons, we have adapted a head restraint technique long used in bats7-9 to mouse10-12. Using this method, we are able to conduct electrophysiological recordings over several days in the unanesthetized mouse. At the end of the recording sessions, we can then inject a dye to reconstruct electrode positions and recording sites or inject a tracer so that pathways to and from the recording loci can be determined. This method allows for well isolated single neuron recordings over multiple days without the use anesthetics.

Electrophysiological characterization of neuronal responses is important for understanding brain function and for guiding the placement of dyes for pathway tracing. However, many studies are performed in anesthetized animals. To understand brain function without anesthetics, we developed a method to record neuronal response properties and inject dyes in awake mouse.

It is well known that anesthesia alters neural response properties in various regions of the brain.13.  In the auditory system, fundamental response ...

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 ...

Recording and Analysis of Circadian Rhythms in Running-wheel Activity in Rodents

When rodents have free access to a running wheel in their home cage, voluntary use of this wheel will depend on the time of day1-5. Nocturnal rodents, including rats, hamsters, and mice, are active during the night and relatively inactive during the day. Many other behavioral and physiological measures also exhibit daily rhythms, but in rodents, running-wheel activity serves as a particularly reliable and convenient measure of the output of the master circadian clock, the suprachiasmatic nucleus (SCN) of the hypothalamus. In general, through a process called entrainment, the daily pattern of running-wheel activity will naturally align with the environmental light-dark cycle (LD cycle; e.g. 12 hr-light:12 hr-dark). However circadian rhythms are endogenously generated patterns in behavior that exhibit a ~24 hr period, and persist in constant darkness. Thus, in the absence of an LD cycle, the recording and analysis of running-wheel activity can be used to determine the subjective time-of-day. Because these rhythms are directed by the circadian clock the subjective time-of-day is referred to as the circadian time (CT). In contrast, when an LD cycle is present, the time-of-day that is determined by the environmental LD cycle is called the zeitgeber time (ZT).
Although circadian rhythms in running-wheel activity are typically linked to the SCN clock6-8, circadian oscillators in many other regions of the brain and body9-14 could also be involved in the regulation of daily activity rhythms. For instance, daily rhythms in food-anticipatory activity do not require the SCN15,16 and instead, are correlated with changes in the activity of extra-SCN oscillators17-20. Thus, running-wheel activity recordings can provide important behavioral information not only about the output of the master SCN clock, but also on the activity of extra-SCN oscillators. Below we describe the equipment and methods used to record, analyze and display circadian locomotor activity rhythms in laboratory rodents.

Circadian rhythms in voluntary wheel-running activity in mammals are tightly coupled to the molecular oscillations of a master clock in the brain. As such, these daily rhythms in behavior can be used to study the influence of genetic, pharmacological, and environmental factors on the functioning of this circadian clock.

When rodents have free access to a running wheel in their home cage, voluntary use of this wheel will depend on the time of day1-5. Nocturnal rodents, ...

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 ...

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 ...

Laboratory Administration of Transcutaneous Auricular Vagus Nerve Stimulation (taVNS): Technique, Targeting, and Considerations

Non-invasive vagus nerve stimulation (VNS) may be administered via a novel, emerging neuromodulatory technique known as transcutaneous auricular vagus nerve stimulation (taVNS). Unlike cervically-implanted VNS, taVNS is an inexpensive and non-surgical method used to modulate the vagus system. taVNS is appealing as it allows for rapid translation of basic VNS research and serves as a safe, inexpensive, and portable neurostimulation system for the future treatment of central and peripheral disease. The background and rationale for taVNS is described, along with electrical and parametric considerations, proper ear targeting and attachment of stimulation electrodes, individual dosing via determination of perception threshold (PT), and safe administration of taVNS.

Laboratory Administration of Transcutaneous Auricular Vagus Nerve Stimulation taVNS: Technique, Targeting, and Considerations

A methodological description of the technique, potential targets, and proper administration of transcutaneous auricular vagus nerve stimulation (taVNS) on the human ear is described.

Non-invasive vagus nerve stimulation (VNS) may be administered via a novel, emerging neuromodulatory technique known as transcutaneous auricular vagus ...

Stereotaxic Surgery for Implantation of Microelectrode Arrays in the Common Marmoset (Callithrix jacchus)

Marmosets (Callithrix jacchus) are small non-human primates that are gaining popularity in biomedical and preclinical research, including the neurosciences. Phylogenetically, these animals are much closer to humans than rodents. They also display complex behaviors, including a wide range of vocalizations and social interactions. Here, an effective stereotaxic neurosurgical procedure for implantation of recording electrode arrays in the common marmoset is described. This protocol also details the pre- and postoperative steps of animal care that are required to successfully perform such a surgery. Finally, this protocol shows an example of local field potential and spike activity recordings in a freely behaving marmoset 1 week after the surgical procedure. Overall, this method provides an opportunity to study the brain function in awake and freely behaving marmosets. The same protocol can be readily used by researchers working with other small primates. In addition, it can be easily modified to allow other studies requiring implants, such as stimulating electrodes, microinjections, implantation of optrodes or guide cannulas, or ablation of discrete tissue regions.

Stereotaxic Surgery for Implantation of Microelectrode Arrays in the Common Marmoset Callithrix jacchus

This work presents a protocol to perform a stereotaxic, neurosurgical implantation of&#160;microelectrode arrays in the common marmoset. This method specifically enables electrophysiological recordings in freely behaving animals but can be easily adapted to any other similar neurosurgical intervention in this species (e.g., cannula for drug administration or electrodes for brain stimulation).

Marmosets (Callithrix jacchus) are small non-human primates that are gaining popularity in biomedical and preclinical research, including the ...

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.

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. ...

Implantation of a Cranial Window for Repeated In Vivo Imaging in Awake Mice

To fully understand the cellular physiology of neurons and glia in behaving animals, it is necessary to visualize their morphology and record their activity in vivo in behaving mice. This paper describes a method for the implantation of a chronic cranial window to allow for the longitudinal imaging of brain cells in awake, head-restrained mice. In combination with genetic strategies and viral injections, it is possible to label specific cells and regions of interest with structural or physiological markers. This protocol demonstrates how to combine viral injections to label neurons in the vicinity of GCaMP6-expressing astrocytes in the cortex for simultaneous imaging of both cells through a cranial window. Multiphoton imaging of the same cells can be performed for days, weeks, or months in awake, behaving animals. This approach provides researchers with a method for viewing cellular dynamics in real time and can be applied to answer a number of questions in neuroscience.

Implantation of a Cranial Window for Repeated In Vivo Imaging in Awake Mice

Presented here is a protocol for the implantation of a chronic cranial window for the longitudinal imaging of brain cells in awake, head-restrained mice.

To fully understand the cellular physiology of neurons and glia in behaving animals, it is necessary to visualize their morphology and record their ...