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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J., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Light-induced+rescue+of+breathing+after+spinal+cord+injury.">Light-induced rescue of breathing after spinal cord injury.</a> J Neurosci. 28 (46), 11862-11870 (2008).</li><li>Geng, Y., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Advances+in+optogenetics+applications+for+central+nervous+system+injuries.">Advances in optogenetics applications for central nervous system injuries.</a> J Neurotrauma. 40 (13-14), 1297-1316 (2023).</li><li>Montgomery, K. L., Iyer, S. M., Christensen, A. J., Deisseroth, K., Delp, S. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Beyond+the+brain:+Optogenetic+control+in+the+spinal+cord+and+peripheral+nervous+system.">Beyond the brain: Optogenetic control in the spinal cord and peripheral nervous system.</a> Sci Transl Med. 8 (337), 5 (2016).</li><li>Tan, T., Watts, S. W., Davis, R. 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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 border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Adson Forceps&#160;</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&#160;</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&#160;</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&#160;</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&#160;</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&#160;</td>
			<td>Fine Science Tools</td>
			<td>16121-14</td>
			<td></td>
		</tr>
		<tr>
			<td>Isoflurane USP</td>
			<td>Fresenius Kabi&#160;</td>
			<td>CP0406V2</td>
			<td>Provided at 5% for induction and 2% for mainentance through precision vaporizer&#160;</td>
		</tr>
		<tr>
			<td>Isopropyl Alcohol 70%</td>
			<td>McKesson</td>
			<td>350600</td>
			<td></td>
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			<td>Lacri-Lube Sterile Eye Ointment&#160;</td>
			<td>Refresh&#160;</td>
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			<td>Long Evans Rats</td>
			<td>Charles River Laboratories</td>
			<td>6</td>
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			<td>Low temperature solder paste</td>
			<td>Chip Quik Inc.</td>
			<td>11.38</td>
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		<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&#160;</td>
			<td>Fine Science Tools</td>
			<td>12002-12</td>
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			<td>PDMS: SYLGARD&#160;184</td>
			<td>Sigma Aldrich</td>
			<td>761036</td>
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			<td>Scalpel Blades - #15</td>
			<td>Fine Science Tools</td>
			<td>10015-00</td>
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			<td>Scalpel Handle - #3</td>
			<td>Fine Science Tools</td>
			<td>10003-12</td>
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			<td>Solder flux</td>
			<td>Chip Quik Inc.</td>
			<td>14.25</td>
			<td></td>
		</tr>
		<tr>
			<td>Stereotaxic Frame&#160;</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>
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		<tr>
			<td>UV Flashlight</td>
			<td>Vansky</td>
			<td>19.99</td>
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		<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>
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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

Dissection and Culture of Commissural Neurons from Embryonic Spinal Cord

Commissural neurons have been widely used to investigate the mechanisms underlying axon guidance during embryonic spinal cord development. The cell bodies of these neurons are located in the dorsal spinal cord and their axons follow stereotyped trajectories during embryonic development. Commissural axons initially project ventrally towards the floorplate. After crossing the midline, these axons turn anteriorly and project towards the brain. Each of these steps is regulated by the action of several guidance cues. Cultures highly enriched in commissural neurons are ideally suited for many experiments addressing the mechanisms of axon pathfinding, including turning assays, immunochemistry and biochemistry. Here, we describe a method to dissect and culture commissural neurons from E13 rat dorsal spinal cord. First, the spinal cord is isolated and dorsal strips are dissected out. The dorsal tissue is then dissociated into a cell suspension by trypsinization and mechanical disruption. Neurons are plated onto poly-L-lysine-coated glass coverslips or tissue-culture dishes. After 30 hours in vitro, most neurons have extended an axon. The purity of the culture (Yam et al. 2009), typically over 90%, can be assessed by immunolabeling with the commissural neuron markers DCC, LH2 and TAG1 (Helms and Johnson, 1998). This neuronal preparation is a useful tool for in vitro studies of the cellular and molecular mechanisms of commissural axon growth and guidance during spinal cord development.

This video demonstrates a method to dissect and culture commissural neurons from E13 rat dorsal spinal cord. Dissociated commissural neurons are useful to study the cellular and molecular mechanisms of axon growth and guidance.

Commissural neurons have been widely used to investigate the mechanisms underlying axon guidance during embryonic spinal cord development. The cell bodies ...

Spinal Cord Electrophysiology II: Extracellular Suction Electrode Fabrication

Development of neural circuitries and locomotion can be studied using neonatal rodent spinal cord central pattern generator (CPG) behavior. We demonstrate a method to fabricate suction electrodes that are used to examine CPG activity, or fictive locomotion, in dissected rodent spinal cords. The rodent spinal cords are placed in artificial cerebrospinal fluid and the ventral roots are drawn into the suction electrode. The electrode is constructed by modifying a commercially available suction electrode. A heavier silver wire is used instead of the standard wire given by the commercially available electrode. The glass tip on the commercial electrode is replaced with a plastic tip for increased durability. We prepare hand drawn electrodes and electrodes made from specific sizes of tubing, allowing consistency and reproducibility. Data is collected using an amplifier and neurogram acquisition software. Recordings are performed on an air table within a Faraday cage to prevent mechanical and electrical interference, respectively.

A demonstration of the fabrication and use of an extracellular suction electrode used to measure electrophysiological recordings of neonatal rodent spinal cords in vitro

Development of neural circuitries and locomotion can be studied using neonatal rodent spinal cord central pattern generator (CPG) behavior. We demonstrate ...

Spinal Cord Transection in the Larval Zebrafish

Mammals fail in sensory and motor recovery following spinal cord injury due to lack of axonal regrowth below the level of injury as well as an inability to reinitiate spinal neurogenesis. However, some anamniotes including the zebrafish Danio rerio exhibit both sensory and functional recovery even after complete transection of the spinal cord. The adult zebrafish is an established model organism for studying regeneration following spinal cord injury, with sensory and motor recovery by 6&#160;weeks post-injury. To take advantage of in vivo analysis of the regenerative process available in the transparent larval zebrafish as well as genetic tools not accessible in the adult, we use the larval zebrafish to study regeneration after spinal cord transection. Here we demonstrate a method for reproducibly and verifiably transecting the larval spinal cord. After transection, our data shows sensory recovery beginning at 2&#160;days post-injury (dpi), with the C-bend movement detectable by 3 dpi and resumption of free swimming by 5 dpi. Thus we propose the larval zebrafish as a companion tool to the adult zebrafish for the study of recovery after spinal cord injury.

After spinal transection, adult zebrafish have functional recovery by six weeks post-injury. To take advantage of larval transparency and faster recovery, we present a method for transecting the larval spinal cord. After transection, we observe sensory recovery beginning at 2&#160;days post-injury, and C-bend movement by 3 days post-injury.

Mammals fail in sensory and motor recovery following spinal cord injury due to lack of axonal regrowth below the level of injury as well as an inability ...

An Ex Vivo Laser-induced Spinal Cord Injury Model to Assess Mechanisms of Axonal Degeneration in Real-time

Injured CNS axons fail to regenerate and often retract away from the injury site. Axons spared from the initial injury may later undergo secondary axonal degeneration. Lack of growth cone formation, regeneration, and loss of additional myelinated axonal projections within the spinal cord greatly limits neurological recovery following injury. To assess how central myelinated axons of the spinal cord respond to injury, we developed an ex vivo living spinal cord model utilizing transgenic mice that express yellow fluorescent protein in axons and a focal and highly reproducible laser-induced spinal cord injury to document the fate of axons and myelin (lipophilic fluorescent dye Nile Red) over time using two-photon excitation time-lapse microscopy. Dynamic processes such as acute axonal injury, axonal retraction, and myelin degeneration are best studied in real-time. However, the non-focal nature of contusion-based injuries and movement artifacts encountered during in vivo spinal cord imaging make differentiating primary and secondary axonal injury responses using high resolution microscopy challenging. The ex vivo spinal cord model described here mimics several aspects of clinically relevant contusion/compression-induced axonal pathologies including axonal swelling, spheroid formation, axonal transection, and peri-axonal swelling providing a useful model to study these dynamic processes in real-time. Major advantages of this model are excellent spatiotemporal resolution that allows differentiation between the primary insult that directly injures axons and secondary injury mechanisms; controlled infusion of reagents directly to the perfusate bathing the cord; precise alterations of the environmental milieu (e.g., calcium, sodium ions, known contributors to axonal injury, but near impossible to manipulate in vivo); and murine models also offer an advantage as they provide an opportunity to visualize and manipulate genetically identified cell populations and subcellular structures. Here, we describe how to isolate and image the living spinal cord from mice to capture dynamics of acute axonal injury.

An Ex Vivo Laser-induced Spinal Cord Injury Model to Assess Mechanisms of Axonal Degeneration in Real-time

We present a protocol utilizing two-photon excitation time-lapse microscopy to simultaneously visualize the dynamics of axon and myelin injuries in real time. This proposed protocol permits studies of both intrinsic and extrinsic factors which can influence central myelinated axon fate after injury and contribute to permanent clinical disability.

Injured CNS axons fail to regenerate and often retract away from the injury site. Axons spared from the initial injury may later undergo secondary axonal ...

Diffusion Imaging in the Rat Cervical Spinal Cord

Magnetic resonance imaging (MRI) is the state of the art approach for assessing the status of the spinal cord noninvasively, and can be used as a diagnostic and prognostic tool in cases of disease or injury. Diffusion weighted imaging (DWI), is sensitive to the thermal motion of water molecules and allows for inferences of tissue microstructure. This report describes a protocol to acquire and analyze DWI of the rat cervical spinal cord on a small-bore animal system. It demonstrates an imaging setup for the live anesthetized animal and recommends a DWI acquisition protocol for high-quality imaging, which includes stabilization of the cord and control of respiratory motion. Measurements with diffusion weighting along different directions and magnitudes (b-values) are used. Finally, several mathematical models of the resulting signal are used to derive maps of the diffusion processes within the spinal cord tissue that provide insight into the normal cord and can be used to monitor injury or disease processes noninvasively.

The goal of this protocol is to obtain high-quality diffusion weighted magnetic resonance imaging (DWI) of the rat spinal cord for noninvasive characterization of tissue microstructure. This protocol describes optimizations of the MRI sequence, radiofrequency coil, and analysis methods to enable DWI images free from artifacts.

Magnetic resonance imaging (MRI) is the state of the art approach for assessing the status of the spinal cord noninvasively, and can be used as a ...

Two-photon Imaging of Cellular Dynamics in the Mouse Spinal Cord

Two-photon (2P) microscopy is utilized to reveal cellular dynamics and interactions deep within living, intact tissues. Here, we present a method for live-cell imaging in the murine spinal cord. This technique is uniquely suited to analyze neural precursor cell (NPC) dynamics following transplantation into spinal cords undergoing neuroinflammatory demyelinating disorders. NPCs migrate to sites of axonal damage, proliferate, differentiate into oligodendrocytes, and participate in direct remyelination. NPCs are thereby a promising therapeutic treatment to ameliorate chronic demyelinating diseases. Because transplanted NPCs migrate to the damaged areas on the ventral side of the spinal cord, traditional intravital 2P imaging is impossible, and only information on static interactions was previously available using histochemical staining approaches. Although this method was generated to image transplanted NPCs in the ventral spinal cord, it can be applied to numerous studies of transplanted and endogenous cells throughout the entire spinal cord. In this article, we demonstrate the preparation and imaging of a spinal cord with enhanced yellow fluorescent protein-expressing axons and enhanced green fluorescent protein-expressing transplanted NPCs.

A new ex vivo preparation for imaging the mouse spinal cord. This protocol allows for two-photon imaging of live cellular interactions throughout the spinal cord.

Two-photon (2P) microscopy is utilized to reveal cellular dynamics and interactions deep within living, intact tissues. Here, we present a method for ...

Imaging Neural Activity in the Primary Somatosensory Cortex Using Thy1-GCaMP6s Transgenic Mice

The mammalian brain exhibits marked symmetry across the sagittal plane. However, detailed description of neural dynamics in symmetric brain regions in adult mammalian animals remains elusive. In this study, we describe an experimental procedure for measuring calcium dynamics through dual optical windows above bilateral primary somatosensory corticies (S1) in Thy1-GCaMP6s transgenic mice using 2-photon (2P) microscopy. This method enables recordings and quantifications of neural activity in bilateral mouse brain regions one at a time in the same experiment for a prolonged period in vivo. Key aspects of this method, which can be completed within an hour, include minimally invasive surgery procedures for creating dual optical windows, and the use of 2P imaging. Although we only demonstrate the technique in the S1 area, the method can be applied to other regions of the living brain facilitating the elucidation of structural and functional complexities of brain neural networks.

Imaging Neural Activity in the Primary Somatosensory Cortex Using Thy1-GCaMP6s Transgenic Mice

We describe an experimental procedure for measuring neuronal activity through dual optical windows above bilateral primary somatosensory corticies (S1) in Thy1-GCaMP6s transgenic mice using 2-photon (2P) microscopy in vivo.

The mammalian brain exhibits marked symmetry across the sagittal plane. However, detailed description of neural dynamics in symmetric brain regions in ...

Novel Passive Clearing Methods for the Rapid Production of Optical Transparency in Whole CNS Tissue

Since the development of CLARITY, a bioelectrochemical clearing technique that allows for three-dimensional phenotype mapping within transparent tissues, a multitude of novel clearing methodologies including CUBIC (clear, unobstructed brain imaging cocktails and computational analysis), SWITCH (system-wide control of interaction time and kinetics of chemicals), MAP (magnified analysis of the proteome), and PACT (passive clarity technique), have been established to further expand the existing toolkit for the microscopic analysis of biological tissues. The present study aims to improve upon and optimize the original PACT procedure for an array of intact rodent tissues, including the whole central nervous system (CNS), kidneys, spleen, and whole mouse embryos. Termed psPACT (process-separate PACT) and mPACT (modified PACT), these novel techniques provide highly efficacious means of mapping cell circuitry and visualizing subcellular structures in intact normal and pathological tissues. In the following protocol, we provide a detailed, step-by-step outline on how to achieve maximal tissue clearance with minimal invasion of their structural integrity via psPACT and mPACT.

Here, we present two novel methodologies, psPACT and mPACT, for achieving maximal optical transparency and subsequent microscopic analysis of tissue vasculature in the intact rodent whole CNS.

Since the development of CLARITY, a bioelectrochemical clearing technique that allows for three-dimensional phenotype mapping within transparent tissues, ...

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 ...

Preparation of Rhythmically-active In Vitro Neonatal Rodent Brainstem-spinal Cord and Thin Slice

Mammalian inspiratory rhythm is generated from a neuronal network in a region of the medulla called the preB&#246;tzinger complex (pBC), which produces a signal driving the rhythmic contraction of inspiratory muscles. Rhythmic neural activity generated in the pBC and carried to other neuronal pools to drive the musculature of breathing may be studied using various approaches, including en bloc nerve recordings and transverse slice recordings. However, previously published methods have not extensively described the brainstem-spinal cord dissection process in a transparent and reproducible manner for future studies. Here, we present a comprehensive overview of a method used to reproducibly cut rhythmically-active brainstem slices containing the necessary and sufficient neuronal circuitry for generating and transmitting inspiratory drive. This work builds upon previous brainstem-spinal cord electrophysiology protocols to enhance the likelihood of reliably obtaining viable and rhythmically-active slices for recording neuronal output from the pBC, hypoglossal premotor neurons (XII pMN), and hypoglossal motor neurons (XII MN). The work presented expands upon previous published methods by providing detailed, step-by-step illustrations of the dissection, from whole rat pup, to in vitro slice containing the XII rootlets.

This protocol both visually communicates the brainstem-spinal cord preparation and clarifies the preparation of brainstem transverse slices in a comprehensive step-by-step manner. It was designed to increase reproducibility and enhance the likelihood of obtaining viable, long lasting, rhythmically-active slices for recording neural output from the respiratory regions of the brainstem.

Mammalian inspiratory rhythm is generated from a neuronal network in a region of the medulla called the preB&#246;tzinger complex (pBC), which produces a ...