S.S. is partially funded by a Four-Year Doctoral Fellowship from the University of British Columbia. A.M. is partially supported by a Canada Graduate Scholarship - Master&#39;s from the Canadian Institute of Health Research (CIHR). D.S. acknowledges funding from the Michael Smith Health Research British Columbia Scholar Award. This work was partially funded by the Government of Canada&#39;s New Frontiers in Research Fund - Transformation (NFRFT-2020-00238). The schematic in Figure 1 was generated using Biorender.com, and the 3D model was obtained with permission from sketchfab.com.

All procedures were conducted according to the guidelines of the Canadian Council for Animal Care and overseen by the University of British Columbia Animal Care Committee. Female Long-Evans rats, weighing 350-450 g and aged 6-8 months, were group-housed (21 &#176;C; 12 h:12 h light&#160;cycle) and given ad libitum access to a standard rodent diet prior to and after the surgery. The details of the reagents and equipment used for this study are listed in the Table of Materials.
1. Pre-operative preparation
<ol>
	<li>Sterilize all surgical tools using an autoclave.</li>
	<li>Anesthetize the animal with isoflurane (5% for induction and 2% for maintenance) delivered in oxygen at a flow rate of 1 L/min.</li>
	<li>Transfer the animal from the induction chamber to a heating pad and promptly connect the isoflurane nose cone. Verify that the animal is under a surgical plane of anesthesia by ensuring a complete loss of toe-pinch reflex on both legs.</li>
	<li>Shave the back of the rat starting at the base of the ears, as shown in Figure 2A.</li>
	<li>Apply a generous amount of sterile eye ointment over the eyes and inject buprenorphine (diluted to 0.03 mg/kg) and 10 mL lactated ringers subcutaneously (warmed to body temperature).</li>
	<li>Wipe the shaved area with antiseptic surgical scrub (chlorhexidine) followed by isopropyl alcohol in a circular motion starting from the center of the shaved area and widening the circle diameter. Repeat the soap/alcohol process two more times.</li>
	<li>Secure the animal in a stereotaxic frame, positioning the head using lubricated ear bars for stability (Figure 2A). 
	NOTE: Throughout the entire procedure, consistently provide thermal support, verify anesthesia depth via toe-pinch reflex, and monitor vital signs.</li>
	<li>Place a sterile surgical drape on top of the animal.</li>
</ol>
2. Cervical spinal cord exposure
<ol>
	<li>Using sterile forceps, begin by palpating the base of the skull. Feel for a prominent spinous process that extends rostrocaudally near the base of the skull; this is C214 (Figures 1B and Figure 2B).</li>
	<li>Proceed with palpation caudal to C2 to find a notably sharp and pointed spinous process, identifiable as T215 (Figures 1B and Figure 2C).</li>
	<li>Using a scalpel, create an incision in the skin, starting from C2 and extending caudally for about 1.5 cm (Figure 2D). 
	NOTE: The incision size may vary among animals of different sizes. Ensure identification of the anatomical landmarks in advance and proceed accordingly.</li>
	<li>Carefully cut through the subcutaneous adipose layer with the scalpel to expose the intact underlying dorsal musculature.</li>
	<li>Once the dorsal muscles are exposed, perform blunt dissection by pulling them apart from the midline using two Adson forceps (Figure 2E). 
	NOTE: It is important to perform blunt dissection (pulling the muscle fibers apart) rather than cutting the muscles to minimize bleeding. Adequate exposure of the dorsal musculature should reveal a ball-shaped muscle (Figure 2G). This muscle completely covers C2 and partially covers C3.</li>
	<li>Using rongeurs and sterile forceps, lift a flap of skin immediately caudal to the incised area. Use rongeurs to create a small subcutaneous pocket-this will be the location for the device (Figure 2F). 
	NOTE: The subcutaneous pocket should be larger than the device itself. It is recommended that the surgeon(s) reposition themselves to face the animal anteriorly to improve control and visibility when opening the subcutaneous pocket.</li>
	<li>Place a retractor to expose the vertebral column (Figure 2G).</li>
	<li>Using rongeurs, remove any remaining muscles or tissues covering the vertebrae and begin to identify the spinal cord segments. Immediately caudal to the ball-shaped muscle is C4, followed by C5 and C6 (Figure 2G-I). Once complete, rinse the surgical area with sterile saline and dry with sterile gauze. 
	NOTE: The ball-shaped muscle fully envelops C2 and partially extends over C3. The spinal cord segment directly caudal to it, with minimal contact, is designated as C4.</li>
	<li>Perform laminectomies depending on the intended purpose of the probe.
	<ol>
		<li>For a probe under the lamina, perform a lateral laminectomy at C5 and C6, creating a lateralized opening in the lamina for future probe placement (Figure 2H, Supplementary Figure 1A).</li>
		<li>For a probe over the lamina, perform a medial laminectomy of C5, ensuring not to remove the lateral aspects of the spinous process-merely exposing a medial pathway for the probe placement (Figure 2I, Supplementary Figure 1B).</li>
		<li>Following the laminectomy, rinse the region with sterile saline and dry with sterile gauze to remove any bone debris. 
		NOTE: When performing the laminectomy, it is critical to push the rongeurs up against the bone and avoid any downward movements to prevent damage to the cord.</li>
	</ol>
	</li>
</ol>
3. Epidural placement of the device
<ol>
	<li>Hold the sterile device with plastic-tip sterile forceps (Figure 3A) and drive it inside the subcutaneous opening made earlier in step 2.9. 
	NOTE: It is critical to avoid touching the device with non-sterile gloves/tools to maintain device sterility. Use designated sterile forceps for device placement and positioning under the skin.</li>
	<li>Suture or glue the device to the neighboring muscle layer to keep it secure, ensuring all sutures and procedures remain sterile. Use 6-0 or 7-0 non-absorbable sutures for optimal securement (Figure 3B). 
	NOTE: Use non-absorbable sutures if suturing the device to the muscles. Otherwise, the device is prone to movement after suture absorption in the body. If using glue, ensure the use of a commercial sterile surgical adhesive which ensures its long-term stability and biocompatibility. Avoid securing the device to the subcutaneous fat/adipose layer to ensure reliable anchoring points.
	<ol>
		<li>Plan the device body position before proceeding with this step. Since the device will be permanently fastened, issues with probe placement (step 3.4) can arise if the device is located too close or too far from the desired level at the spinal cord.</li>
	</ol>
	</li>
	<li>Place the retractor around the spinal cord and open a suitable window to place the probe on the spinal cord.</li>
	<li>Probe placements:
	<ol>
		<li>Probe placement under the lamina: Carefully insert the probe using plastic-tip forceps under the (C5 and C6) laminae by sliding it through the lateralized channels made in step 2.9.1 (Figure 3D).</li>
		<li>Probe placement over the spinal cord:
		<ol>
			<li>Adjust the probe using plastic-tip forceps to align and place the probe tip on top of the medial window created at C5 in step 2.9.2 (Figure 3C).</li>
		</ol>
		</li>
	</ol>
	</li>
	<li>In a sterile, small, and preferably ceramic container, prepare cement by mixing one scoop of dental cement powder, 3 drops of high tech grease, and one drop of catalyst. Mix with sterile toothpicks until a viscous consistency is achieved. 
	NOTE: It is highly recommended that one person who is not performing the surgery prepares the cement so that the surgeon can hold the probe in place and dry the desired area immediately before applying the cement. This step is time-sensitive. Ensure that the surgeon positions the probe prior to preparing the cement. Once the catalyst is applied, the cement will thicken too much to effectively bond with the bone if there is a delay. Practice with the cement prior to surgery to ensure achieving the proper consistency before placing it on the probe/bone.</li>
	<li>Dry the intended cementing area on the vertebrae completely to make a reliable anchor point. Apply 1-2 drops of dental cement at the tip of the probe, which should be placed on top of the intended vertebral level, in this case, C4 (Figure 3C,D). 
	NOTE: It is critical for the vertebrae to be as dry as possible prior to cementing. Otherwise, the cement will not adhere, and the probe will not be secured.</li>
	<li>Pause for 30 s and gently touch the cement to verify it is cured.</li>
	<li>If the cement is not completely cured, wait an additional 20 s and re-apply freshly made cement until the probe is robustly secured to the bone. 
	NOTE: The cement has bonded properly when it becomes hard and stiff to the touch. 
	CAUTION: Applying the cement while it is too liquid may cause it to seep into the spinal cord tissue between the vertebrae. If this occurs, allow approximately 10 s for the cement to thicken to a more gum-like consistency before gently using forceps to remove cement that has come into contact with the spinal cord.</li>
</ol>
4. Post-surgical procedures
<ol>
	<li>Once the surgery is complete, suture the incision site using 5-0 vicryl sutures. Gently remove the animal from the stereotaxic apparatus and transfer it to a heated recovery chamber. For the initial 3-5 days post-surgery, provide the animal with soft, moistened chow, treats, and hydrogel water.</li>
	<li>Thoroughly monitor the animal twice daily during the first-week post-surgery. Administer buprenorphine and warmed lactated Ringer&#39;s injections twice daily for the subsequent 2 days, or longer if signs of pain persist.&#160;Weigh the animal daily for one week post-implantation, then weekly. Consult a veterinarian if weight loss is observed. Continue daily checks until the animal shows no clinical health concerns. Thereafter, check on the animal at least once per week. 
	NOTE: Monitor animals for weight loss over 20%, severe pain unresponsive to treatment, exposed implants or large seromas (3x implant size), autophagy, dehydration, or combined symptoms raising the clinical health score to require euthanasia. Consult a veterinarian for treatment options as needed.</li>
	<li>(Optional) Outline the device perimeter on the skin after surgery and incision closure for a visual indication of the device location in the body (Figure 4A). 
	NOTE: This step is particularly useful when the device requires external access, such as devices needing to be powered wirelessly through an external transmitter where antenna coupling is critical, or drug delivery pumps that must be refilled through the skin16,17,18.</li>
</ol>

Spinal Cord Surgery and Optical Shank Implantation in Rodents

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The authors have no conflicts of interest.

Neuromodulation and therapeutic interventions of the spinal cord often require the placement of probes in precise, targeted segments3,4,7,13. Given the inherent mobility of the spinal cord, the probe must be reliably secured to enable chronic studies. Based on the specific application, it may be important to control whether the probe is in physical contact with the spinal cord, or if the contact can be reduced to lessen the inflammatory tissue response when possible. Therefore, surgical steps for each of the two methods are described. The protocol specifically details how to place a probe in the cervical segment of the spinal cord at C5. Nonetheless, using the described landmark for T2 or T10 of the spinal cord, the probe can be similarly placed in a precise location over the thoracic or the lumbar region by counting down the vertebrae from T2 or T10, respectively, once they are exposed. Furthermore, to minimize spinal cord tissue damage, we secured the device body, which is often larger and more rigid compared to the connected probe, in a subcutaneous space away from the spinal cord.
There are some critical points to implanting the device that is coupled with the probe. First, it is critical to decide on the location of the device body prior to cementing the probe. This ensures the distance between the tip of the probe and the device body is optimized to reduce tension on the probe as well as avoid having extra probe length, which can, for example cause probe twisting or displacement. Essentially, the goal is to ensure that the length of the probe is similar to the distance from the subcutaneous space where the device body is placed to the targeted spinal cord region where the probe is cemented. By conducting terminal surgery procedures in which various probe lengths are tested, the optimum size can be determined for a targeted segment.
To maintain sterility, the device should be handled carefully to prevent contact with the outer layer of the skin during insertion into the subcutaneous pocket. Such contact can compromise the device&#39;s sterility, potentially leading to post-operative infection. In addition, it is important to minimize the amount of force applied to the device when holding it with forceps to prevent damaging its coating, which is typically a thin protective, insulative, and sterile layer20,21. Removing the coating may drastically reduce the lifespan of the device by, for example, shortening the circuit, causing electrical shock to the animal, and/or provoking an inflammatory response in the body. Handling the device with plastic tip forceps may help reduce such complications.
When suturing the device to soft tissue, it is important to avoid suturing to subcutaneous adipose tissue. As observed in preliminary trials, fat layers are not a reliable anchoring point for sutures since they are prone to rupture. Instead, the device body was sutured to an adjacent muscle layer in the subcutaneous space using non-absorbable sutures for the permanent placement of the device in the body. On the other hand, when securing the probe to the spinous processes, it is important to ensure the site to which the probe is being secured is dry before applying the cement. Wet bone/probe prolongs the curing time and may result in the complete failure of the process.
There are some critical considerations associated with an implantable device that need to be carefully addressed prior to implantation surgery. (1) Electrically active parts of the device must be encapsulated by an insulative passivation layer. Any deprivation in the passivating layer might cause device functional failure. (2) The implantable must be thoroughly sterilized according to the facility animal protocol. (3) The junction between the device and the neural probes or the stimulatory shanks must be securely formed. The connection will go through repeatable mechanical stress due to constant animal movements. (4) The neural probes or stimulatory shanks attached to the device must be flexible and stretchable enough to avoid snapping at various points.
The described protocol may be extended to implant devices in animal models of different sizes. After identifying the anatomical landmarks, the described surgical methods can be methodically customized to secure any neural probes or stimulatory shanks at targeted segments of the spinal cord and implant their associated control modules. However, depending on the application, different devices can have varying sizes, materials, and thicknesses from the one implanted in this paper; for example, devices connected to an external control module require additional considerations. Additionally, it must be noted that while this protocol is tailored for optogenetic stimulation, other neuromodulatory applications, such as drug delivery or electrical stimulation/recording, require slightly different surgical procedures. Specifically, these applications need subdural implantation to ensure direct contact with the spinal cord beneath the dura mater7. However, for optogenetics, intimate contact with the tissue is typically unnecessary because the rodent dura mater does not significantly impede light penetration, which enables light sources to be placed epidurally10.

The spinal cord facilitates a range of essential central nervous system functions, from coordinating motor behaviors to regulating homeostatic processes such as respiration1,2. Elucidating the role of the sophisticated network of circuitry across the spinal cord requires interfaces, whether for electrical stimulation, recording, drug delivery, or optical stimulation to targeted areas3,4,5,6. Although devices have been developed to enable such interrogations7,8,9,10,11, specialized surgical techniques are required for their chronic implantation in the spinal cord4. In particular, the spinal cord and associated vertebrae have increased susceptibility to mechanical deformations caused by natural movements such as extension and bending8,12,13. These unique characteristics of the spinal cord make it intrinsically challenging to ensure that the implanted probes remain stable, functional, and secured at a specific segment over extended periods of time.
Herein, a surgical protocol is described for inserting and securing an optical shank in a targeted segment of the spinal cord (Figure 1A). Since interfacing with the cervical region in particular has been shown to introduce unique challenges9, the implantation steps are specifically demonstrated over the C5 cervical region. It is posited that the complexity of the cervical spine arises from its deeper positioning and the abundance of musculature, a characteristic not as prominent along the rest of the spinal cord. Regardless, the procedures outlined in this protocol are designed to be adaptable for surgeries across various spinal cord regions. Stepwise instructions are provided to locate and identify spinal cord segments using pronounced anatomical &#34;landmarks&#34; identifiable from over the skin (Figure 1B). The protocol then elucidates two techniques for surgical implantation: one tailored for probes that require direct contact with the spinal cord, and another for probes that may not require direct contact. The described steps are designed to be reproduced by any researcher with training in rodent survival surgery.
This protocol encompasses step-by-step instructions for the implantation of an optoelectronic device (18 mm x 13 mm) with an attached flexible optical shank over the C5 cervical level. The implantable device is secured subcutaneously caudal to C5 and consists of a microscale light-emitting diode (&#181;LED) indicator, which illuminates when spinal cord optical stimulation occurs, providing live feedback of device functionality. The effect of the implanted optical shank on the natural motor function was assessed on rodents who had received implants and was compared to rodents with sham surgeries. Results indicate that the probes do not adversely affect the natural hindlimb and forelimb function of the animal seven days post-implantation.

<table><tbody><tr><td>Adson Forceps&amp;nbsp;</td><td>Fine Science Tools</td><td>11027-12</td><td></td></tr><tr><td>Alm 3 Point Retractor</td><td>Fine Science Tools</td><td>17010-10</td><td></td></tr><tr><td>Buprenorphine / Vetergesic&amp;nbsp;</td><td>CDMV</td><td>124918</td><td>Manufacturer provides at 0.3 mg/mL but must be diluted to 0.03 mg/kg for use in rats</td></tr><tr><td>Chlorhexidine 2% Solution</td><td>Partnar</td><td>PCH-020&amp;nbsp;</td><td></td></tr><tr><td>Curved Long Hemostat Forceps</td><td>KaamKaaj Tools</td><td>14.5</td><td>Curved Long Hemostat Forceps with A Stainless Steel Ratchet Locking Tweezer</td></tr><tr><td>CVD Parylene Machine: SCS Labcoter 2</td><td>Specialty Coating Systems</td><td>PDS 2010</td><td></td></tr><tr><td>Dental Cement - Catalyst&amp;nbsp;</td><td>Parkell, Inc</td><td>S371</td><td></td></tr><tr><td>Dental Cement - Metabond</td><td>Parkell, Inc</td><td>S398</td><td></td></tr><tr><td>Dental Cement - Powder&amp;nbsp;</td><td>Parkell, Inc</td><td>S396</td><td></td></tr><tr><td>Forceps with Replaceable Plastic Tips</td><td>Fine Science Tools</td><td>11980-13</td><td></td></tr><tr><td>Friedman-Pearson Rongeurs&amp;nbsp;</td><td>Fine Science Tools</td><td>16121-14</td><td></td></tr><tr><td>Isoflurane USP</td><td>Fresenius Kabi&amp;nbsp;</td><td>CP0406V2</td><td>Provided at 5% for induction and 2% for mainentance through precision vaporizer&amp;nbsp;</td></tr><tr><td>Isopropyl Alcohol 70%</td><td>McKesson</td><td>350600</td><td></td></tr><tr><td>Lacri-Lube Sterile Eye Ointment&amp;nbsp;</td><td>Refresh&amp;nbsp;</td><td></td><td></td></tr><tr><td>Long Evans Rats</td><td>Charles River Laboratories</td><td>6</td><td></td></tr><tr><td>Low temperature solder paste</td><td>Chip Quik Inc.</td><td>11.38</td><td></td></tr><tr><td>Magnets</td><td>Radial Magnets, Inc.</td><td>0.53</td><td>Magnet Neodymium Iron Boron (NdFeB) N35 (3.00 mm x 1.00 mm)</td></tr><tr><td>Olsen-Hegar Needle Holders with Suture Cutters&amp;nbsp;</td><td>Fine Science Tools</td><td>12002-12</td><td></td></tr><tr><td>PDMS: SYLGARD&amp;nbsp;184</td><td>Sigma Aldrich</td><td>761036</td><td></td></tr><tr><td>Scalpel Blades - #15</td><td>Fine Science Tools</td><td>10015-00</td><td></td></tr><tr><td>Scalpel Handle - #3</td><td>Fine Science Tools</td><td>10003-12</td><td></td></tr><tr><td>Solder flux</td><td>Chip Quik Inc.</td><td>14.25</td><td></td></tr><tr><td>Stereotaxic Frame&amp;nbsp;</td><td>David Kopf Instruments</td><td>Model 900</td><td></td></tr><tr><td>Sterile Kwik-Sil Adhesive</td><td>World Precision Instruments</td><td>KWIK-SIL-S</td><td></td></tr><tr><td>UV Flashlight</td><td>Vansky</td><td>19.99</td><td></td></tr><tr><td>Wireless Charger</td><td>Nilkin</td><td>NKT06</td><td></td></tr><tr><td>Wireless Charging coil</td><td>TDK Corporation</td><td>WT202012-15F2-ID</td><td></td></tr></tbody></table>

implantation of optoelectronic devices in the rodent spinal cord

Neuromodulation can provide diagnostic, modulatory, and therapeutic applications. While extensive work has been conducted in the brain, modulation of the spinal cord remains relatively unexplored. The inherently delicate and mobile spinal cord tissue imposes constraints that make the precise implantation of neural probes challenging. Despite recent advances in neuromodulation devices, particularly flexible bioelectronics, opportunities to expand their use in the spinal cord have been limited by the surgical complexities of device implantation. Here, we provide a series of surgical protocols tailored specifically for the implantation of a custom-made optoelectronic device that interfaces with the spinal cord in rodents. The steps to place and anchor an optical shank on a specific segment of the spinal cord via two different surgical implantation methods are detailed here. These methods are optimized for a diverse range of devices and applications, which may or may not require direct contact with the spinal cord for optical stimulation. To elucidate the methodology, the vertebral anatomy is referenced first to identify prominent landmarks before making a skin incision. The surgical steps to secure an optical shank over the cervical spine in rodents are demonstrated. Procedures are then outlined for securing the optoelectronic device connected to the optical shank in a subcutaneous space away from the spinal cord, minimizing unnecessary direct contact. Behavioral studies comparing animals receiving the implants to those undergoing sham surgeries indicate that the optical shanks did not adversely affect hindlimb or forelimb function seven days post-implantation. The present work broadens the neuromodulation toolkit for use in future studies aimed at investigating various spinal cord interventions.

An optoelectronic device with its detailed functional diagram shown in Supplementary Figure 2 was implanted in four Long Evans rats. Supplementary Figure 3 shows the final optoelectronic device ready to implant. Three other animals received sham surgeries, which involved a medial laminectomy at C5 without device implantation. The optoelectronic device consisted of a flexible probe with an embedded &#181;LED at the tip which was activated by an integrated LED driver. The LED driver is controlled by a microcontroller with programmable firmware. It also consisted of a device body that was sutured to the muscle layer immediately under the skin. A layer of Parylene-C (~10 &#181;m) is deposited on the whole device using chemical vapor deposition (CVD). A second layer of Polydimethylsiloxane (PDMS) (~800 &#181;m) covered the optoelectronic device body (Supplementary Figure 3) to form a soft interface with the tissue. The probe tip was secured at C4 with the &#181;LED hovering over C5. A &#181;LED indicator was utilized on the device (with its light visible from under the skin) that simultaneously turned on with the &#181;LED of the optical shank for live verification of device functionality. The animals were monitored for a period of 7 days following surgery to confirm the sustained reliability of their performance over time (Figure 4B).
The motor functions of the animals were evaluated using the Martinez open-field locomotor rating scale19. To assess open-field behavior, two trained observers who were unaware of the treatment groups conducted the tests before the operations as well as on days three, five, and seven post-surgery. Following data collection, the Mann-Whitney U test was conducted to determine differences at each timepoint for both the forelimb and hindlimb scores between the implant and sham groups. Our analysis indicates a similar forelimb function score in implant and sham groups by day seven (Figure 5A). Similarly, there were no statistically significant differences between the groups for the hindlimb scores across all timepoints (Figure 5B).
Post-mortem verification was performed 7 days post-implantation to confirm if the probe and the device body had remained in place. No visible detachment of the suture or the device was found. Furthermore, pulling on the device body did not cause its detachment from the tissue (Supplementary Figure 4A). The previously dissected and sutured muscles were then exposed over the spinal cord, and it was confirmed that the probes remained securely cemented over the spinal cord (Supplementary Figure 4B). Similar to the device body, the probe head was pulled back successively against the cementing point to assess its attachment to the probe-lamina mechanical joint.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/66992/66992fig01.jpg" /> 
Figure 1: Schematic overview of device implantation and anatomical landmarks. (A) Demonstrating the placement of the probe over the spinal cord and subcutaneous placement of the device. (B) 3D model indicating the landmarks used to determine spinal cord levels. The C2, T2, and T10 spinous processes are shown for reference. Darker colors indicate the corresponding level. <a href="https://www.jove.com/files/ftp_upload/66992/66992fig01large.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/66992/66992fig02.jpg" /> 
Figure 2: Exposing the spinal cord and preparing a subcutaneous pocket. (A) The stereotaxic is positioned on the animal. (B) C2 spinous process and (C) T2 spinous process are identified via palpation. (D) An incision through the skin and subcutaneous adipose layer is made to expose dorsal musculature at the point of interest at the cervical level. (E) Through blunt dissection of the dorsal musculature, the cervical vertebrae are exposed. (F) A subcutaneous pocket is created to secure the implantable device caudal to the incision site. (G) A retractor is placed following adequate dissection to expose cervical vertebrae and the ball-shaped muscle, completely covering C2 and partially masking C3. The dashed line indicates the ball-shaped muscle. Once the cervical spinal cord has been exposed, either (H) two lateral laminectomies at C5 and C6 are done for probe placement under the vertebrae, or (I) medial laminectomy is made at C5 for probe placement over the vertebrae. Asterisks indicate the site of lateral laminectomy. Scale bars = 3 mm. <a href="https://www.jove.com/files/ftp_upload/66992/66992fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/66992/66992fig03.jpg" /> 
Figure 3: Device implantation and probe placement. (A) The device is placed in the subcutaneous pocket. (B) The device is sutured to the musculature. (C) The probe is secured on top of the C5 lamina, which had received medial laminectomy. (D) The device is placed under the C5 and C6 lamina which both received lateral laminectomy. In both (C) and (D), the tip of the probe is cemented at an intact C4. Scale bars = 3 mm. <a href="https://www.jove.com/files/ftp_upload/66992/66992fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/66992/66992fig04.jpg" /> 
Figure 4: Marking device and verifying functionality post-surgery. (A) The location of the device may be optionally marked over the skin after suturing for the ease of its identification post-surgery. (B) The figure depicts an animal post-implantation. The functionality of the device was validated by observing the indicator &#181;LED visible beneath the skin, confirming the successful operation of the device (the bump on the right side of the animal is where the device body is implanted). <a href="https://www.jove.com/files/ftp_upload/66992/66992fig04large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/66992/66992fig05.jpg" /> 
Figure 5: Martinez open field behavioral scores in sham and implant groups for forelimb and hindlimb performance over time. The plots illustrate the mean behavioral scores for (A) forelimb and (B) hindlimb open field assessments across four timepoints: 0 (baseline), 3 days, 5 days, and 7 days post-implantation (DPI). Error bars represent the standard error of the mean (SEM). Significant differences (p &#60; 0.05) between the sham and implant groups are indicated with asterisks (*) at specific timepoints. The figure legend indicates the sham groups displayed by the dotted line, while the implant group is shown by the solid line. The sham sample size was n = 3, and the implant was n = 4. The non-parametric Mann-Whitney U test was used to evaluate the significance of differences between groups at each timepoint. <a href="https://www.jove.com/files/ftp_upload/66992/66992fig05large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Supplementary Figure 1: Illustration of laminectomy. Dashed lines indicate the regions to resect for (A) two lateral laminectomies for probe placement under the vertebrae, and (B) medial laminectomy for probe placement over the vertebrae. <a href="https://www.jove.com/files/ftp_upload/66992/Supplementary Figure 1.zip" target="_blank">Please click here to download this File.</a>
Supplementary Figure 2: Schematic of the optoelectronic device. The detailed block diagram of the device is shown. The top left block depicts a wireless power receiver antenna resonant LC tank. The received power is rectified and fed into a low-dropout voltage regulator (LDO). A microcontroller unit automatically activates the device based on the programmed parameters, and an LED driver powers any &#181;LEDs embedded in the probe. <a href="https://www.jove.com/files/ftp_upload/66992/Supplementary Figure 2.zip" target="_blank">Please click here to download this File.</a>
Supplementary Figure 3: Optoelectronic device. The final optoelectronic device with biocompatible encapsulation connected to an optical shank comprising 1 &#181;LED at the tip. The dashed rectangle depicts the location of the &#181;LED. <a href="https://www.jove.com/files/ftp_upload/66992/Supplementary Figure 3.zip" target="_blank">Please click here to download this File.</a>
Supplementary Figure 4: Post-mortem verification of device stability. Seven days after implantation, (A) the device body had remained sutured to the musculature in the same position it was implanted, and (B) the cemented probe remained secured on top of the C4 lamina. <a href="https://www.jove.com/files/ftp_upload/66992/Supplementary Figure 4.zip" target="_blank">Please click here to download this File.</a>

Author Spotlight: Innovative Methodology for Implanting and Securing Neural Probes in the Rodent Spinal Cord

This protocol details a surgical procedure for performing spinal cord surgery and for implanting and securing an optical shank over the spinal cord in rodents.

Implantation of Optoelectronic Devices in the Rodent Spinal Cord

Neuromodulation can provide diagnostic, modulatory, and therapeutic applications. While extensive work has been conducted in the brain, modulation of the ...

implantation-of-optoelectronic-devices-in-the-rodent-spinal-cord

Research

JoVE Journal

Neuroscience

1.1K Views.  University of British Columbia. This protocol details a surgical procedure for performing spinal cord surgery and for implanting and securing an optical shank over the spinal cord in rodents.

Video: Author Spotlight: Innovative Methodology for Implanting and Securing Neural Probes in the Rodent Spinal Cord

Implantation and Recording of Wireless Electroretinogram and Visual Evoked Potential in Conscious Rats

The full-field electroretinogram (ERG) and visual evoked potential (VEP) are useful tools to assess retinal and visual pathway integrity in both laboratory and clinical settings. Currently, preclinical ERG and VEP measurements are performed with anesthesia to ensure stable electrode placements. However, the very presence of anesthesia has been shown to contaminate normal physiological responses. To overcome these anesthesia confounds, we develop a novel platform to assay ERG and VEP in conscious rats. Electrodes are surgically implanted sub-conjunctivally on the eye to assay the ERG and epidurally over the visual cortex to measure the VEP. A range of amplitude and sensitivity/timing parameters are assayed for both the ERG and VEP at increasing luminous energies. The ERG and VEP signals are shown to be stable and repeatable for at least 4 weeks post surgical implantation. This ability to record ERG and VEP signals without anesthesia confounds in the preclinical setting should provide superior translation to clinical data.

We show surgical implantation and Recording procedures to measure visual electrophysiological signals from the eye (electroretinogram) and brain (visual evoked potential) in conscious rats, which is more analogous to the human condition where recordings are conducted without anesthesia confounds.

The full-field electroretinogram (ERG) and visual evoked potential (VEP) are useful tools to assess retinal and visual pathway integrity in both ...

Behavior

Simultaneous Electroencephalography, Real-time Measurement of Lactate Concentration and Optogenetic Manipulation of Neuronal Activity in the Rodent Cerebral Cortex

Although the brain represents less than 5% of the body by mass, it utilizes approximately one quarter of the glucose used by the body at rest1. The function of non rapid eye movement sleep (NREMS), the largest portion of sleep by time, is uncertain. However, one salient feature of NREMS is a significant reduction in the rate of cerebral glucose utilization relative to wakefulness2-4. This and other findings have led to the widely held belief that sleep serves a function related to cerebral metabolism. Yet, the mechanisms underlying the reduction in cerebral glucose metabolism during NREMS remain to be elucidated.
	One phenomenon associated with NREMS that might impact cerebral metabolic rate is the occurrence of slow waves, oscillations at frequencies less than 4 Hz, in the electroencephalogram5,6. These slow waves detected at the level of the skull or cerebral cortical surface reflect the oscillations of underlying neurons between a depolarized/up state and a hyperpolarized/down state7. During the down state, cells do not undergo action potentials for intervals of up to several hundred milliseconds. Restoration of ionic concentration gradients subsequent to action potentials represents a significant metabolic load on the cell8; absence of action potentials during down states associated with NREMS may contribute to reduced metabolism relative to wake.
	Two technical challenges had to be addressed in order for this hypothetical relationship to be tested. First, it was necessary to measure cerebral glycolytic metabolism with a temporal resolution reflective of the dynamics of the cerebral EEG (that is, over seconds rather than minutes). To do so, we measured the concentration of lactate, the product of aerobic glycolysis, and therefore a readout of the rate of glucose metabolism in the brains of mice. Lactate was measured using a lactate oxidase based real time sensor embedded in the frontal cortex. The sensing mechanism consists of a platinum-iridium electrode surrounded by a layer of lactate oxidase molecules. Metabolism of lactate by lactate oxidase produces hydrogen peroxide, which produces a current in the platinum-iridium electrode. So a ramping up of cerebral glycolysis provides an increase in the concentration of substrate for lactate oxidase, which then is reflected in increased current at the sensing electrode. It was additionally necessary to measure these variables while manipulating the excitability of the cerebral cortex, in order to isolate this variable from other facets of NREMS.
	We devised an experimental system for simultaneous measurement of neuronal activity via the elecetroencephalogram, measurement of glycolytic flux via a lactate biosensor, and manipulation of cerebral cortical neuronal activity via optogenetic activation of pyramidal neurons. We have utilized this system to document the relationship between sleep-related electroencephalographic waveforms and the moment-to-moment dynamics of lactate concentration in the cerebral cortex. The protocol may be useful for any individual interested in studying, in freely behaving rodents, the relationship between neuronal activity measured at the electroencephalographic level and cellular energetics within the brain.

A procedure is described for manipulating the activity of cerebral cortical pyramidal neurons optogenetically while the electroencephalogram, electromyogram, and cerebral lactate concentration are monitored. Experimental recordings are performed on cable-tethered mice while they undergo spontaneous sleep/wake cycles. Optogenetic equipment is assembled in our laboratory; recording equipment is commercially available.

Although the brain represents less than 5% of the body  by mass, it utilizes approximately one quarter of the glucose used by the body  at rest1. The ...

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 ...

Implanting Glass Spinal Cord Windows in Adult Mice with Experimental Autoimmune Encephalomyelitis

Experimental autoimmune encephalomyelitis (EAE) in adult rodents is the standard experimental model for studying autonomic demyelinating diseases such as multiple sclerosis. Here we present a low-cost and reproducible glass window implantation protocol that is suitable for intravital microscopy and studying the dynamics of spinal cord cytoarchitecture with subcellular resolution in live adult mice with EAE. Briefly, we surgically expose the vertebrae T12-L2 and construct a chamber around the exposed vertebrae using a combination of cyanoacrylate and dental cement. A laminectomy is performed from T13 to L1, and a thin layer of transparent silicone elastomer is applied to the dorsal surface of the exposed spinal cord. A modified glass cover slip is implanted over the exposed spinal cord taking care that the glass does not directly contact the spinal cord. To reduce the infiltration of inflammatory cells between the window and spinal cord, anti-inflammatory treatment is administered every 2 days (as recommended by ethics committee) for the first 10 days after implantation. EAE is induced only 2-3 weeks after the cessation of anti-inflammatory treatment.
Using this approach we successfully induced EAE in 87% of animals with implanted windows and, using Thy1-CFP-23 mice (blue axons in dorsal spinal cord), quantified axonal loss throughout EAE progression. Taken together, this protocol may be useful for studying the recruitment of various cell populations as well as their interaction dynamics, with subcellular resolution and for extended periods of time. This intravital imaging modality represents a valuable tool for developing therapeutic strategies to treat autoimmune demyelinating diseases such as EAE.

We describe a method for implanting and maintaining glass windows over the exposed spinal cords of adult mice for studying experimental autoimmune encephalomyelitis. These windows provide chronic optical access to the spinal cord for monitoring cell dynamics at the subcellular level in live animals using two-photon microscopy.

Experimental autoimmune encephalomyelitis (EAE) in adult rodents is the standard experimental model for studying autonomic demyelinating diseases such as ...

Medicine

Optogenetic Manipulation of Neuronal Activity to Modulate Behavior in Freely Moving Mice

Optogenetic modulation of neuronal circuits in freely moving mice affects acute and long-term behavior. This method is able to perform manipulations of single neurons and region-specific transmitter release, up to whole neuronal circuitries in the central nervous system, and allows the direct measurement of behavioral outcomes. Neurons express optogenetic tools via an injection of viral vectors carrying the DNA of choice, such as Channelrhodopsin2 (ChR2). Light is brought into specific brain regions via chronic optical implants that terminate directly above the target region. After two weeks of recovery and proper tool-expression, mice can be repeatedly used for behavioral tests with optogenetic stimulation of the neurons of interest.
Optogenetic modulation has a high temporal and spatial resolution that can be accomplished with high cell specificity, compared to the commonly used methods such as chemical or electrical stimulation. The light does not harm neuronal tissue and can therefore be used for long-term experiments as well as for multiple behavioral experiments in one mouse. The possibilities of optogenetic tools are nearly unlimited and enable the activation or silencing of whole neurons, or even the manipulation of a specific receptor type by light.
The results of such behavioral experiments with integrated optogenetic stimulation directly visualizes changes in behavior caused by the manipulation. The behavior of the same animal without light stimulation as a baseline is a good control for induced changes. This allows a detailed overview of neuronal types or neurotransmitter systems involved in specific behaviors, such as anxiety. The plasticity of neuronal networks can also be investigated in great detail through long-term stimulation or behavioral observations after optical stimulation. Optogenetics will help to enlighten neuronal signaling in several kinds of neurological diseases.

With optogenetic manipulation of specific neuronal populations or brain regions, behavior can be modified with high temporal and spatial resolution in freely moving animals. By using different optogenetic tools in combination with chronically implanted optical fibers, a variety of neuronal modulations and behavioral testing can be performed.

Optogenetic modulation of neuronal circuits in freely moving mice affects acute and long-term behavior. This method is able to perform manipulations of ...

In vivo Optogenetic Stimulation of the Rodent Central Nervous System

The ability to probe defined neural circuits in awake, freely-moving animals with cell-type specificity, spatial precision, and high temporal resolution has been a long sought tool for neuroscientists in the systems-level search for the neural circuitry governing complex behavioral states. Optogenetics is a cutting-edge tool that is revolutionizing the field of neuroscience and represents one of the first systematic approaches to enable causal testing regarding the relation between neural signaling events and behavior. By combining optical and genetic approaches, neural signaling can be bi-directionally controlled through expression of light-sensitive ion channels (opsins) in mammalian cells. The current protocol describes delivery of specific wavelengths of light to opsin-expressing cells in deep brain structures of awake, freely-moving rodents for neural circuit modulation. Theoretical principles of light transmission as an experimental consideration are discussed in the context of performing in vivo optogenetic stimulation. The protocol details the design and construction of both simple and complex laser configurations and describes tethering strategies to permit simultaneous stimulation of multiple animals for high-throughput behavioral testing.

In vivo Optogenetic Stimulation of the Rodent Central Nervous System

Optogenetics has become a powerful tool for use in behavioral neuroscience experiments. This protocol offers a step-by-step guide to the design and set-up of laser systems, and provides a full protocol for carrying out multiple and simultaneous in vivo optogenetic stimulations compatible with most rodent behavioral testing paradigms.

The ability to probe defined neural circuits in awake, freely-moving animals with cell-type specificity, spatial precision, and high temporal resolution ...

A Procedure for Implanting a Spinal Chamber for Longitudinal In Vivo Imaging of the Mouse Spinal Cord

Studies in the mammalian neocortex have enabled unprecedented resolution of cortical structure, activity, and response to neurodegenerative insults by repeated, time-lapse in vivo imaging in live rodents. These studies were made possible by straightforward surgical procedures, which enabled optical access for a prolonged period of time without repeat surgical procedures. In contrast, analogous studies of the spinal cord have been previously limited to only a few imaging sessions, each of which required an invasive surgery. As previously described, we have developed a spinal chamber that enables continuous optical access for upwards of 8 weeks, preserves mechanical stability of the spinal column, is easily stabilized externally during imaging, and requires only a single surgery. Here, the design of the spinal chamber with its associated surgical implements is reviewed and the surgical procedure is demonstrated in detail. Briefly, this video will demonstrate the preparation of the surgical area and mouse for surgery, exposure of the spinal vertebra and appropriate tissue debridement, the delivery of the implant and vertebral clamping, the completion of the chamber, the removal of the delivery system, sealing of the skin, and finally, post-operative care. The procedure for chronic in vivo imaging using nonlinear microscopy will also be demonstrated. Finally, outcomes, limitations, typical variability, and a guide for troubleshooting are discussed.

A Procedure for Implanting a Spinal Chamber for Longitudinal In Vivo Imaging of the Mouse Spinal Cord

In this video, we describe a procedure for implanting a chronic optical imaging chamber over the dorsal spinal cord of a live mouse. The chamber, surgical procedure, and chronic imaging are reviewed and demonstrated.

Studies in the mammalian neocortex have enabled unprecedented resolution of cortical structure, activity, and response to neurodegenerative insults by ...

Laminectomy and Spinal Cord Window Implantation in the Mouse

This protocol describes a method for spinal cord laminectomy and glass window implantation for in vivo imaging of the mouse spinal cord. An integrated digital vaporizer is utilized to achieve a stable plane of anesthesia at a low-flow rate of isoflurane. A single vertebral spine is removed, and a commercially available cover-glass is overlaid on a thin agarose bed. A 3D-printed plastic backplate is then affixed to the adjacent vertebral spines using tissue adhesive and dental cement. A stabilization platform is used to reduce motion artifact from respiration and heartbeat. This rapid and clamp-free method is well-suited for acute multi-photon fluorescence microscopy. Representative data are included for an application of this technique to two-photon microscopy of the spinal cord vasculature in transgenic mice expressing eGFP:Claudin-5 &#8212; a tight junction protein.

This protocol describes implantation of a glass window onto the spinal cord of a mouse to facilitate visualization by intravital microscopy.

This protocol describes a method for spinal cord laminectomy and glass window implantation for in vivo imaging of the mouse spinal cord. An integrated ...

Optrode Array for Simultaneous Optogenetic Modulation and Electrical Neural Recording

During the last decade, optogenetics has become an essential tool for the investigation of neural signaling due to its unique capability of selective neural modulation or monitoring. As specific types of neuronal cells can be genetically modified to express opsin proteins, optogenetics enables optical stimulation or inhibition of the selected neurons. There have been several technological advances in the optical system for optogenetics. Recently, it was proposed to combine the optical waveguide for light delivery with electrophysiological recording to simultaneously monitor the neural responses to optogenetic stimulation or inhibition. In this study, an implantable optrode array (2x2 optical fibers) was developed with embedded multichannel electrodes. 
A light-emitting diode (LED) was employed as a light source, and a microfabricated microlens array was integrated to provide sufficient light power at the tip of the optical fibers. The optrode array system comprises the disposable part and the reusable part. The disposable part has optical fibers and electrodes, while the reusable part has the LED and electronic circuitry for light control and neural signal processing. The novel design of the implantable optrode array system is introduced in the accompanying video in addition to the procedure of the optrode implantation surgery, optogenetic light stimulation, and the electrophysiological neural recording. The results of in vivo experiments successfully showed time-locked neural spikes evoked by the light stimuli from hippocampal excitatory neurons of mice.

Here, we present the fabrication method of an optrode system with optical fibers for light delivery and an electrode array for neural recording. In vivo experiments with transgenic mice expressing channelrhodopsin-2 show the feasibility of the system for simultaneous optogenetic stimulation and electrophysiological recording.

During the last decade, optogenetics has become an essential tool for the investigation of neural signaling due to its unique capability of selective ...

Development of a Surgical Technique for Subretinal Implants in Rats

Retinal degeneration, such as age-related macular degeneration (AMD), is a leading cause of blindness worldwide. A myriad of approaches have been undertaken to develop regenerative medicine-based therapies for AMD, including stem cell-based therapies. Rodents as animal models for retinal degeneration are a foundation for translational research, due to the broad spectrum of strains that develop retinal degeneration diseases at different stages. However, mimicking human therapeutic delivery of subretinal implants in rodents is challenging, due to anatomical differences such as lens size and vitreous volume. This surgical protocol aims to provide a guided method for transplanting implants into the subretinal space in rats. A user-friendly comprehensive description of the critical steps has been included. This protocol has been developed as a cost-efficient surgical procedure for reproducibility across different preclinical studies in rats. Proper miniaturization of a human-sized implant is required prior to conducting the surgical experiment, which includes adjustments to the dimensions of the implant. An external approach is used instead of an intravitreal procedure to deliver the implant to the subretinal space. Using a small sharp needle, a scleral incision is performed in the temporal superior quadrant, followed by paracentesis to reduce intraocular pressure, thereby minimizing resistance during the surgical implantation. Next, a balanced salt solution (BSS) injection through the incision is carried out to achieve focal retinal detachment (RD). Lastly, insertion and visualization of the implant into the subretinal space are conducted. Post-operative assessment of the subretinal placement of the implant includes imaging by spectral domain optical coherence tomography (SD-OCT). Imaging follow-ups ascertain the subretinal stability of the implant, before the eyes are harvested and fixated for histological analysis.

The present protocol describes the scleral approach for subretinal device implantation, a feasible surgical technique for implementation in animal models of retinal diseases in research.

Implantation of Optoelectronic Devices in the Rodent Spinal Cord

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Development of a Surgical Technique for Subretinal Implants in Rats