The study was performed with support by the Schulze Family Foundation (to A.S.B.).

All methods described here have been approved by the Institutional Animal Care and Use Committee (IACUC) of Mayo Clinic.
1. Prerequisites of Rigor and Reproducibility
<ol>
	<li>To ensure rigor of design, follow national standards of good laboratory practices are at all times and obtain internal approval by IACUC (or similar committee) prior to any animal involvement in experiments. 
	NOTE: This protocol was designed to maintain a clinically faithful approach. Thus, the materials, instruments, and techniques involved are described in a fashion identical to the highest clinical standards in humans. For instance, strict sterile technique is followed and expired materials should never be used.</li>
	<li>To support reproducibility of this methodology in an experimental setting, develop internal standard operating procedures and control for variability in swine breed, weight, gender, and age between separate cohorts. 
	NOTE: The design of this protocol was based on the use of swine weighing 38 - 53 kg.</li>
</ol>
2. Pre-operative Animal Care
<ol>
	<li>Administer prophylactic intramuscular (IM) ceftiofur, given at 5 mg/kg, 1 day prior to the procedure.</li>
	<li>Fast animals of solid food and restrict animals from cosmetic treatments, i.e. oil baths, 12 h prior to the procedure.</li>
	<li>Induce general anesthesia within 1 h of the procedure, using IM tiletamine and zolazepam, given as Telazol at 5 mg/kg, and IM xylazine, given at 2 mg/kg.</li>
	<li>Once induced, administer subcutaneous (SC) buprenorphine sustained release (SR), given at 0.18 mg/kg.</li>
	<li>Place an ear-vein catheter and perform rapid sequence intubation to place an endotracheal tube.</li>
	<li>Attach a pulse oximeter with heart rate monitoring function to the tongue to monitor oxygenation and heart rate.</li>
	<li>Place the animal in the prone position and clip the skin over the dorsum using an electric clipper. Clip hair over a large, bilateral area extending from the midline sagittal plane to the midline coronal plane and longitudinally from the sacral apex to the scapular spines. Use adhesive tape to remove hair and free-floating skin.</li>
	<li>Scrub the clipped area up to 3 times with warm water and soap and dry the skin using a lint-free towel.</li>
	<li>Mark bilateral anatomical landmarks using a surgical marking pen. Mark the last ribs, iliac crests, spinous processes, and transverse processes. 
	NOTE: By marking the last ribs, iliac crests, and transverse processes, the lumbar spine is demarcated along its superior, inferior, and lateral boundaries, respectively. This protocol has been designed to guide access and injection of any lumbar DRG of interest. For reference, the superior iliac crests align with the L3 or L4 vertebral level.</li>
	<li>Cover the animal with a warm blanket for transport to the operative suite.</li>
</ol>
3. Positioning in the Operative Suite
<ol>
	<li>Gently lift and position the animal prone into a modified large humane animal sling with padded abdominal aperture. 
	NOTE: The abdominal aperture allows for reduction in abdominal pressure similar to the Wilson frame used during human surgery of the spine. In turn, this decreases intraoperative bleeding from the spinal blood vessels. The sling is advantageous because the legs are allowed to hang freely through padded openings which protect the animal from peripheral nerve impingement. However, because the sling frame is made of metal, it should be padded with insulation to avoid electrical short-circuiting and inadvertent animal burn. A blanket roll can be placed to position the head and neck in a comfortable position, depending on the size of the animal.</li>
	<li>Maintain general anesthesia using 1 - 3% inhaled (IH) isoflurane, titrated to effect. Moisten eyes with ophthalmic ointment and gently tape them shut.</li>
	<li>Place lines for vital sign monitoring to document temperature, blood pressure, heart rate, and oxygenation. Monitor ventilation by capnography.</li>
	<li>Place an adhesive, disposable electrosurgical dispersive electrode over the left or right scapula.</li>
	<li>Administer warmed Lactated Ringer&#39;s as maintenance fluids through the ear-vein catheter. Give fluids at a rate of 5 - 10 mL/kg/h.</li>
	<li>Place a forced-air warming device over the thoracic and cervical region and avoid covering the last ribs.</li>
</ol>
4. Sterile Preparation of Operative Field for a Left-sided Injection
NOTE: From this point forward, proceed in strict sterile fashion.
<ol>
	<li>Prepare the skin overlying the lumbar spine beginning with wide application of 0.7% iodine povacrylex and 74% isopropyl alcohol according to the manufacturer&#39;s instructions. To ensure that the guide needle can later be placed in a sterile fashion, lateralize application to the side of planned injection by extending antisepsis towards the midline coronal plane past the marked transverse processes.</li>
	<li>Place disposable surgical towels in rectangular fashion to outline the planned incision site, which is over the midline along the marked lumbar spinous processes.</li>
	<li>Apply an adhesive antimicrobial incise drape over the operative towels and exposed skin. Clamp drapes in place and extend the drapery edge off the operative field.</li>
	<li>Secure a vertical drape to poles at the head of the sling between the operative field and the monitoring technician.</li>
	<li>Secure lines for suction and electrosurgery within the operative field by clamping to the sterile drapes. Pass the free ends of tubes and wires out of the sterile field.</li>
</ol>
5. Skin Incision and Subperiosteal Dissection
<ol>
	<li>Palpate the lumbar spinous processes along the midline and identify 3 consecutive vertebral levels.</li>
	<li>Use a #15 scalpel to open an 8 - 12 cm midline sagittal incision through the incise drape directly posterior to the spinous processes. Maintain hemostasis using gauze tamponade and monopolar electrosurgery. 
	NOTE: Care should be taken not to deviate from the midline as the incision is advanced in the anterior direction because this limits bleeding from the paraspinal muscles. Periodic palpation for the spinous processes facilitates advancement. Self-retaining Weitlaner, Meyerding, or Gelpi retractors may be placed and repositioned as needed to facilitate the dissection. Suction is used to maintain visibility.</li>
	<li>Dissect the subcutaneous tissue and fat using monopolar electrosurgery until the thoracolumbar fascia is reached. Palpate the spinous processes deep to the thoracolumbar fascia and cut the fascia along the midline to expose the supraspinous ligament spanning between spinous processes. 
	NOTE: The thoracolumbar fascia is identified as an organized, aponeurotic sheath with a connective tissue grain that interweaves in an oblique, lateral to medial direction. At this point, the incision may be lengthened in either the superior or inferior direction to ensure that 3 spinous processes are fully visible with the centermost spinous process aligned within the center of the dissection field.</li>
	<li>Use a #15 blade to place a 2-mm deep parasagittal incision through the supraspinous ligament posterior to each spinous process. Place each incision along the left third of the posterior surface of the spinous process.</li>
	<li>Gently release the supraspinous ligament at each level along each incision using a 5-mm Freer elevator.</li>
	<li>Identify the subperiosteal plane and dissect within that plane along the lateral surface of each spinous process.</li>
	<li>Perform subperiosteal dissection at each spinous process in a parallel fashion to ensure that a gentle, even dissection is achieved.</li>
	<li>Incise the paraspinal muscle attachment along the interspinous spaces using monopolar electrosurgery in concert with subperiosteal dissection.</li>
	<li>Identify the lamina at each level and continue subperiosteal dissection laterally to reach the lateral border of the 2 zygapophyseal joints that connect the 3 exposed vertebrae and to reach the lateral edge of the lamina between the joints, called the pars interarticularis. 
	NOTE: The pars interarticularis is the posterior border of the intervertebral foramen wherein the DRG resides. Occasionally, a small vein arises from a foramen located on the posterior surface of the lamina. These veins have a tendency to rupture during the subperiosteal dissection. Hemostasis is easily achieved by using a combination of bipolar electrosurgery and bone wax applied to the foramen.</li>
</ol>
6. Single-level Laminotomy
<ol>
	<li>Identify the target of laminotomy as the centermost lamina located between and medial to the 2 zygapophyseal joints.</li>
	<li>Trace the lamina to its inferior edge, to a point just medial to the contiguous inferior articular process of the caudal zygapophyseal joint.</li>
	<li>Use a 5-mm Freer elevator or curette to palpate the transition between the caudalmost edge of the lamina and central canal. 
	NOTE: Care is taken to not force the palpating instrument anterior as this will contact the dural sac and spinal cord. Note that the spinal cord in swine extends past the lumbar spine 18. An intervertebral disc rongeur can be used to remove additional soft tissue overlying this area to facilitate palpation.</li>
	<li>Use a 2-mm up-biting, 45-degree Kerrison rongeur to extract bone in a piece-wise fashion. Remove bone along the base of the spinous process superiorly to a level just caudal to the caudal surface of the pedicle and out laterally to its full extent.</li>
	<li>Use angled bone rongeurs to assist with bone removal. Leave the inferior articular process that was connected to the lamina in place until the laminotomy is largely completed.</li>
	<li>Confirm that the inferior articular process is freely mobile and attached only by the zygapophyseal joint capsule. Incise the capsule using a #15 or #11 blade.</li>
	<li>Remove the inferior articular process in a piece-wise fashion but leave the adjoining superior articular process intact. 
	NOTE: As the laminotomy is completed, hemostasis is performed with bipolar electrosurgery. Monopolar electrosurgery is not used because of the proximity of neural structures. Bone wax can be placed along sites of bleeding from exposed bone and absorbable gelatin sponges can be used to obtain hemostasis near soft tissue. Cottonoid is a helpful tool to wick serous fluid and blood away from the dissection.</li>
</ol>
7. Dissection of DRG
<ol>
	<li>Evacuate the epidural fat in a piece-wise fashion from superficial to deep beginning medially and proceeding laterally. Remove fat by gentle dissection using bipolar forceps and suction with 6 - 10 French Frazier suction tips. 
	NOTE: Loupe magnification or the use of a dissecting microscope is helpful in providing the level of detail needed to safely evacuate the epidural fat and achieve meticulous hemostasis of the epidural venous plexus using bipolar electrosurgery.</li>
	<li>Identify the dural sac along the midline running in a superoinferior direction, parallel to the axis of the skin incision. Remove epidural fat along the dural sac until the dural sac can be seen to give rise to the dural nerve root sleeve.</li>
	<li>Trace the dural sleeve laterally and inferiorly by epidural fat evacuation until it is seen to enlarge around the DRG. 
	NOTE: Identify the DRG for its oval shape and yellow to orange color. At the mid-lumbar spine, the DRG is typically 4 - 6 mm in size, longest in the medial to lateral direction, and located directly inferior or 2 - 3 mm medial to its respective pedicle. A blunt, right-angled nerve hook can be used to gently palpate for the pedicle.</li>
	<li>Evacuate epidural fat laterally, past the DRG, until the adjoining spinal nerve is seen. 
	NOTE: If durotomy occurs, repair it by watertight closure using 6-0 nylon suture and smooth micro needle driver in a simple running stich.</li>
</ol>
8. Injection of DRG
<ol>
	<li>Use a 22-gauge spinal needle to guide the trajectory of a 32-gauge convection enhanced delivery (CED) needle. Puncture the 22-gauge guide needle through the skin and paraspinal muscles. 
	NOTE: The CED needle is designed to achieve fluid convection in tissue, also known as bulk flow, due to pressure gradients 19,20.</li>
	<li>Aim the guide needle along a trajectory that approximates the longitudinal axis of the DRG and results in the needle tip emerging from the lateral paraspinal wall of the dissection field.</li>
	<li>Fine tune the needle path until the needle lumen aligns with the center of the DRG. 
	NOTE: The guide needle should never be allowed to contact the DRG.</li>
	<li>Draw up sterile injectate into a sterile syringe and connect the syringe to the sterile infusion tubing.</li>
	<li>Secure the tubing to the CED needle and hand the syringe out of the sterile field. Connect the syringe to a programmable syringe pump. 
	NOTE: Tubing is prepared to a length of 5 feet to ensure that sterility and mobility are maintained. Also, it is of paramount importance that no air bubbles be introduced into solution.</li>
	<li>Advance the injectate until expression is seen from the CED needle tip.</li>
	<li>Place the CED needle within the guide needle lumen and slowly advance the CED needle until it emerges from the guide needle tip. Ensure that the DRG is not punctured during needle alignment.</li>
	<li>Fine tune the guide needle position along the long axis of its trajectory to determine the final location of the CED tip.</li>
	<li>Secure the guide needle and CED needle together using interlocking needle hubs once depth and alignment of the guide and CED needles is achieved.</li>
	<li>Confirm that all injection apparatus connections are fully secured, including the guide needle, CED needle, and connected tubing loaded with injectate.</li>
	<li>Advance the guide needle along its long axis to approximate the CED needle tip and DRG.</li>
	<li>Puncture the DRG with the CED needle tip.</li>
	<li>Submerse the CED needle tip into the three-dimensional center of the DRG. 
	NOTE: Because the DRG is a three-dimensional structure of variable size and shape, DRG exposure must be complete in order to accurately place the CED needle tip at the true center of the DRG. The true DRG center is located at the intersection of its three anatomical axes, namely, the anterior to posterior, lateral to medial, and superior to inferior axes.</li>
	<li>Deliver 100 &#956;L of injectate by CED using a graduated rate and volume of 3 steps.</li>
	<li>Deliver 4 &#956;l at 2 &#956;L/min for the first step. Deliver 8 &#956;L at 4 &#956;L/min for the second step. Deliver 88 &#956;L at 8 &#956;L/min for the third and final step. 
	NOTE: Allow a 3-min pause between steps and after the final step to allow for pressure equilibration.</li>
	<li>Withdraw the injection apparatus after the final injection step and 3-min pause along its long axis in a smooth, gentle motion. 
	NOTE: For injected solutions that are colorless, colored dye is included in the solution at a concentration of 0.1% weight/volume to assist in visual assessment of injectate distribution 12. Also, the vital dye 4&#39;,6-diamidino-2-phenylindole (DAPI) is included in solution at a concentration of 0.25 &#956;g/&#956;L when the study design requires histologic assessment of injectate distribution 5.</li>
</ol>
9. Closure
<ol>
	<li>Apply 3 rounds of warm saline irrigation to the surgical site prior to closure to mobilize and flush the site of debris, i.e. bone fragments. Use suction to recover the saline and debris. 
	NOTE: Meticulous hemostasis is ensured when irrigation remains clear. Hemostatic agents (gelatin sponge) and cottonoid are removed at this time. Ensure that all materials and instruments have been cleared from the incision site prior to closure.</li>
	<li>Use a 3-layered technique for closure.</li>
	<li>Suture the thoracolumbar fascia using 0 suture in a simple, interrupted, noninverted fashion. Place a stich every 5 - 8 mm to achieve watertight closure.</li>
	<li>Suture the subcutaneous tissue using 2-0 suture in a simple, interrupted, inverted fashion with a stitch placed every 5- to 8-mm to achieve adequate strength.</li>
	<li>Close the skin using 0 suture in a simple, running or interrupted fashion.</li>
	<li>Use a needle counter to ensure that no sharps are unaccounted for.</li>
	<li>Irrigate the skin with saline, dry the skin, and place adhesive bandage strips perpendicular to the incision.</li>
	<li>Place gauze on top of the bandage strips and attach a final adhesive antimicrobial incise drape.</li>
</ol>
10. Post-operative Animal Care
<ol>
	<li>Extubate, cover with warm blankets, and transport the animal to recovery.</li>
	<li>Follow standard institutional operating procedures for post-operative monitoring and recovery from survival surgery. At a minimum, observe the animal every 15 min until return of consciousness, hourly until full anesthetic recovery is achieved, and twice-daily thereafter.</li>
	<li>Provide post-operative pain management by administering IM or oral carprofen, given at 4 mg/kg, once daily for 5 days beginning on post-operative day 0. Administer SC buprenorphine SR, given at 0.18 mg/kg, once on post-operative day 2.</li>
	<li>Administer post-operative antisepsis by giving IM ceftiofur, given at 5 mg/kg, once on post-operative day 4.</li>
	<li>Remove the bandage on post-operative day 5 - 7. Remove the sutures when wound healing is complete, typically on post-operative day 10 - 14.</li>
	<li>Humanely euthanize the animal according to standard institutional operating procedures once the study endpoint is reached.</li>
</ol>

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None; the authors have no conflicts of interest related to this study.

We sought to describe a method for surgical exposure of DRG via laminotomy and intraganglionic injection in a healthy large animal species, specifically, swine. In rodents, a similar method has been detailed 12 and used to deliver conventional pharmacologic agents 8,10 and viral vectors 6,7,9,12 to DRG. ...

Dorsal root ganglia (DRG) are anatomically discrete, neuronal collections located along the vertebral column. Each DRG contains the primary sensory neurons that encode and relay peripheral stimuli to the central nervous system (CNS) from specific body regions. For instance, the pain of osteoarthritis begins when pain receptors located about a joint perceive noxious stimuli. This process is termed nociception. Long-term awareness of noxious stimuli leads to chronic pain 1.
Chronic pain is a frequent subject of preclinical study 2 where one goal is to develop useful methods for targeted delivery of analgesics to DRG, such as intraganglionic injection 3. However, DRG are difficult to access because they reside within the boney confines of intervertebral foramina 4. Several groups have successfully overcome this obstacle through the use of spine surgery in rodents 5,6,7,8,9,10.
In the clinic, laminectomy is a common spine operation and refers to surgical removal of the vertebral lamina, thereby unroofing the vertebral canal 11. Incorporation of surgical techniques to afford direct DRG access has been successful in rodents 5,12, however, translation may be unrealistic considering differences in size of relevant structures and how that influences pharmacokinetics or technical feasibility 13,14. For example, one study determined the transverse spinal cord diameter at T10 to be 3.0, 7.0, and 8.2 mm for rat, pig, and human, respectively 15. Thus, large animal models better approximate human dimensions of nervous structures.
In swine, Raore et al. used multi-level laminectomy to gain access to the cervical spinal cord for multiple intraspinal injections 16. The procedure was well tolerated and led to a phase I clinical trial where comparable surgical outcomes were documented 17. These results encourage continued use of preclinical large animal models as predictors of technical feasibility and safety in humans.
To date, no detailed methodology exists for surgical access and injection of DRG in a large animal species. To narrow this translational gap, we report a protocol for DRG exposure and injection via laminotomy in swine. Standard neurosurgical techniques, instruments, and materials were used and the method was designed to mimic modern surgical practice. We demonstrate intraganglionic injection using an aqueous solution for lumbar DRG and confirm successful delivery via histology after post-operative day 21.

<table><tbody><tr><td>Large humane animal sling</td><td>Britz &amp;amp; Company</td><td>002539</td><td>Modified to include abdominal aperture</td></tr><tr><td>Adhesive patient return electrode - 9 inch</td><td>Medtronic</td><td>E7506</td><td>-</td></tr><tr><td>Ranger blood &amp;amp; fluid warming system</td><td>3M</td><td>24500</td><td>-</td></tr><tr><td>Lactated Ringer&#039;s fluid</td><td>Hospira</td><td>0409-7953-09</td><td>-</td></tr><tr><td>Force air warming device</td><td>3M</td><td>77500</td><td>-</td></tr><tr><td>Duraprep solution with applicator, 26 mL (0.7% iodine povacrylex, 74% isopropyl alcohol)</td><td>3M</td><td>8630</td><td>-</td></tr><tr><td>Sterile disposable surgical towels</td><td>Medline</td><td>MDT2168286</td><td>-</td></tr><tr><td>Ioban 2 incise drape</td><td>3M</td><td>6651EZSB</td><td>-</td></tr><tr><td>Disposable suction canister and tubing</td><td>Medline</td><td>DYND44703H</td><td>-</td></tr><tr><td>Button switch electrosurgical monopolar pencil</td><td>Medtronic</td><td>E2450H</td><td>-</td></tr><tr><td>Fine smooth straight bipolar electrosurgical forceps, 4 1/2 inch</td><td>Bovie</td><td>A826</td><td>-</td></tr><tr><td>#15 blade</td><td>Miltex</td><td>4-315</td><td>-</td></tr><tr><td>#11 blade</td><td>Miltex</td><td>4-311</td><td>-</td></tr><tr><td>4 x 4 surgical gauze</td><td>Dynarex</td><td>3262</td><td>-</td></tr><tr><td>Weitlaner self-retaining retractor, 8 inch</td><td>Miltex</td><td>11-618</td><td>-</td></tr><tr><td>Meyerding self-retaining retractor, 1 x 2 3/8 inch</td><td>Sklar</td><td>42-2078</td><td>-</td></tr><tr><td>Gelpi self-retaining retractor, 7 inch</td><td>Sklar</td><td>60-6570</td><td>-</td></tr><tr><td>Freer elevator, 5 mm</td><td>Medline</td><td>MDS4641518F</td><td>-</td></tr><tr><td>Bone wax</td><td>Ethicon</td><td>W31G</td><td>-</td></tr><tr><td>Spurling intervertebral disc rongeur, 3 mm</td><td>Sklar</td><td>42-2852</td><td>-</td></tr><tr><td>Spurling 45-degree, up-biting Kerrison rongeur, 2 mm</td><td>Medline</td><td>MDS4052802</td><td>-</td></tr><tr><td>Leksell angled rongeur, 2 mm</td><td>Sklar</td><td>40-4097</td><td>-</td></tr><tr><td>Gelfoam, size 50</td><td>Pfizer</td><td>AZL32301</td><td>-</td></tr><tr><td>Cottonoid patty</td><td>Medtronic</td><td>8004007</td><td>-</td></tr><tr><td>Frazier suction tip, 6 Fr</td><td>Sklar</td><td>50-2006</td><td>-</td></tr><tr><td>Frazier suction tip, 10 Fr</td><td>Sklar</td><td>50-2010</td><td>-</td></tr><tr><td>Dandy blunt right angle nerve hook</td><td>Medline</td><td>MDS4005220</td><td>-</td></tr><tr><td>Nylon suture, 6-0</td><td>Ethicon</td><td>697G</td><td>-</td></tr><tr><td>Castroviejo smooth micro needle holder</td><td>Medline</td><td>MDG2428614</td><td>-</td></tr><tr><td>22 gauge Quinke point spinal needle</td><td>Halyard Health</td><td>18397</td><td>-</td></tr><tr><td>32 gauge CED needle with locking Luer hub</td><td>See comments</td><td>n/a</td><td>As in: Pleticha, J., Maus, T.P., Christner, J.A., Marsh, M.P., Lee, K.H., Hooten, W.M., Beutler, A.S. Minimally invasive convection-enhanced delivery of biologics into dorsal root ganglia: validation in the pig model and prospective modeling in humans. Technical note. J Neurosurg. 121(4), 851-8 (2014).</td></tr><tr><td>Polyethylene tubing, 5 feet</td><td>Scientific Commodities</td><td>BB31695-PE/05</td><td>-</td></tr><tr><td>Monoject syringe, 3 mL</td><td>Kendall</td><td>SY15352</td><td>-</td></tr><tr><td>NanoJet syringe pump</td><td>Chemyx</td><td>10050</td><td>-</td></tr><tr><td>DAPI</td><td>Sigma-Aldrich</td><td>D9542</td><td>-</td></tr><tr><td>Fast Green FCF</td><td>Sigma-Aldrich</td><td>F7252</td><td>-</td></tr><tr><td>Bulb irrigation syringe</td><td>Medline</td><td>DYND20125</td><td>-</td></tr><tr><td>Fine-toothed Adson forceps</td><td>Medline</td><td>MDS1000212</td><td>-</td></tr><tr><td>Vicryl suture, 0</td><td>Ethicon</td><td>J603H</td><td>-</td></tr><tr><td>Vicryl suture, 2-0</td><td>Ethicon</td><td>J317H</td><td>-</td></tr><tr><td>Needle counter</td><td>Medline</td><td>NC20FBRGS</td><td>-</td></tr><tr><td>Steri-strip skin closure, 1/2x4 inch</td><td>3M</td><td>R1547</td><td>-</td></tr></tbody></table>

laminotomy for lumbar dorsal root ganglion access and injection in swine

Dorsal root ganglia (DRG) are anatomically well defined structures that contain all primary sensory neurons below the head. This fact makes DRG attractive targets for injection of novel therapeutics aimed at treating chronic pain. In small animal models, laminectomy has been used to facilitate DRG injection because it involves surgical removal of the vertebral bone surrounding each DRG. We demonstrate a technique for intraganglionic injection of lumbar DRG in a large animal species, namely, swine. Laminotomy is performed to allow direct access to DRG using standard neurosurgical techniques, instruments, and materials. Compared with more extensive bone removal via laminectomy, we implement laminotomy to conserve spinal anatomy while achieving sufficient DRG access. Intraoperative progress of DRG injection is monitored using a non-toxic dye. Following euthanasia on post-operative day 21, the success of injection is determined by histology for intraganglionic distribution of 4&#39;,6-diamidino-2-phenylindole (DAPI). We inject a biologically inactive solution to demonstrate the protocol. This method could be applied in future preclinical studies to target therapeutic solutions to DRG. Our methodology should facilitate testing the translatability of intraganglionic small animal paradigms in a large animal species. Additionally, this protocol may serve as a key resource for those planning preclinical studies of DRG injection in swine.

Histologic assessment of injectate spread 
Successful delivery of injectate to DRG is determined by histologic assessment of DAPI spread. The technique involves positioning the needle tip in the three-dimensional center of the DRG. Therefore, successful delivery is determined by evaluating the extent of DAPI staining from histologic sections both near (central DRG sections) and distant (peripheral DRG sections) to the needle tip. Figure 1A and Figur...

Watch this Scientific Journal Video about Laminotomy for Lumbar Dorsal Root Ganglion Access and Injection in Swine at JoVE.com

Laminotomy for Lumbar Dorsal Root Ganglion Access and Injection in Swine

We describe a method for laminotomy in swine that provides access to lumbar dorsal root ganglia (DRG) for intraganglionic injection. Injection progress is monitored intraoperatively and histologically confirmed up to 21 days after surgery. This protocol could be used for future preclinical studies involving DRG injection.

Dorsal root ganglia (DRG) are anatomically well defined structures that contain all primary sensory neurons below the head. This fact makes DRG attractive ...

laminotomy-for-lumbar-dorsal-root-ganglion-access-injection

Research

JoVE Journal

Neuroscience

12.7K Views.  Mayo Clinic, Translational Science Track, Mayo Graduate School. Beschrijven we een methode voor laminotomy in varkens die toegang tot lumbale dorsal root ganglia (DRG) voor intraganglionic injectie geeft. Injectie is toezicht op de vorderingen en en histologisch bevestigde tot 21 dagen na de operatie. Dit protocol kan worden gebruikt voor toekomstige Preklinische studies waarbij DRG injectie.

Video: Laminotomy for Lumbar Dorsal Root Ganglion Access and Injection in Swine

Intraspinal Cell Transplantation for Targeting Cervical Ventral Horn in Amyotrophic Lateral Sclerosis and Traumatic Spinal Cord Injury

Respiratory compromise due to phrenic motor neuron loss is a debilitating consequence of a large proportion of human traumatic spinal cord injury (SCI) cases 1 and is the ultimate cause of death in patients with the motor neuron disorder, amyotrophic laterals sclerosis (ALS) 2.
ALS is a devastating neurological disorder that is characterized by relatively rapid degeneration of upper and lower motor neurons. Patients ultimately succumb to the disease on average 2-5 years following diagnosis because of respiratory paralysis due to loss of phrenic motor neuron innnervation of the diaphragm 3. The vast majority of cases are sporadic, while 10% are of the familial form. Approximately twenty percent of familial cases are linked to various point mutations in the Cu/Zn superoxide dismutase 1 (SOD1) gene on chromosome 21 4. Transgenic mice 4,5 and rats 6 carrying mutant human SOD1 genes (G93A, G37R, G86R, G85R) have been generated, and, despite the existence of other animal models of motor neuron loss, are currently the most highly used models of the disease.
Spinal cord injury (SCI) is a heterogeneous set of conditions resulting from physical trauma to the spinal cord, with functional outcome varying according to the type, location and severity of the injury 7. Nevertheless, approximately half of human SCI cases affect cervical regions, resulting in debilitating respiratory dysfunction due to phrenic motor neuron loss and injury to descending bulbospinal respiratory axons 1. A number of animal models of SCI have been developed, with the most commonly used and clinically-relevant being the contusion 8.
Transplantation of various classes of neural precursor cells (NPCs) is a promising therapeutic strategy for treatment of traumatic CNS injuries and neurodegeneration, including ALS and SCI, because of the ability to replace lost or dysfunctional CNS cell types, provide neuroprotection, and deliver gene factors of interest 9.
Animal models of both ALS and SCI can model many clinically-relevant aspects of these diseases, including phrenic motor neuron loss and consequent respiratory compromise 10,11. In order to evaluate the efficacy of NPC-based strategies on respiratory function in these animal models of ALS and SCI, cellular interventions must be specifically directed to regions containing therapeutically relevant targets such as phrenic motor neurons. We provide a detailed protocol for multi-segmental, intraspinal transplantation of NPCs into the cervical spinal cord ventral gray matter of neurodegenerative models such as SOD1G93A mice and rats, as well as spinal cord injured rats and mice 11.

Neural precursor transplantation is a promising strategy for protecting and/or replacing lost/dysfunctional cervical phrenic motor neurons in spinal cord injury (SCI) and the motor neuron disorder, amyotrophic laterals sclerosis (ALS). We provide a protocol for cell delivery to cervical spinal cord ventral horn in rodent models of ALS and SCI.

Respiratory compromise due to phrenic motor neuron loss is a debilitating consequence of a large proportion of human traumatic spinal cord injury (SCI) ...

Medicine

Live Imaging of Dorsal Root Axons after Rhizotomy

The primary sensory axons injured by spinal root injuries fail to regenerate into the spinal cord, leading to chronic pain and permanent sensory loss. Regeneration of dorsal root (DR) axons into spinal cord is prevented at the dorsal root entry zone (DREZ), the interface between the CNS and PNS. Our understanding of the molecular and cellular events that prevent regeneration at DREZ is incomplete, in part because complex changes associated with nerve injury have been deduced from postmortem analyses. Dynamic cellular processes, such as axon regeneration, are best studied with techniques that capture real-time events with multiple observations of each living animal. Our ability to monitor neurons serially in vivo has increased dramatically owing to revolutionary innovations in optics and mouse transgenics. Several lines of thy1-GFP transgenic mice, in which subsets of neurons are genetically labeled in distinct fluorescent colors, permit individual neurons to be imaged in vivo1. These mice have been used extensively for in vivo imaging of muscle2-4 and brain5-7, and have provided novel insights into physiological mechanisms that static analyses could not have resolved. Imaging studies of neurons in living spinal cord have only recently begun. Lichtman and his colleagues first demonstrated their feasibility by tracking injured dorsal column (DC) axons with wide-field microscopy8,9. Multi-photon in vivo imaging of deeply positioned DC axons, microglia and blood vessels has also been accomplished10. Over the last few years, we have pioneered in applying in vivo imaging to monitor regeneration of DR axons using wide-field microscopy and H line of thy1-YFP mice. These studies have led us to a novel hypothesis about why DR axons are prevented from regenerating within the spinal cord11.
In H line of thy1-YFP mice, distinct YFP+ axons are superficially positioned, which allows several axons to be monitored simultaneously. We have learned that DR axons arriving at DREZ are better imaged in lumbar than in cervical spinal cord. In the present report we describe several strategies that we have found useful to assure successful long-term and repeated imaging of regenerating DR axons. These include methods that eliminate repeated intubation and respiratory interruption, minimize surgery-associated stress and scar formation, and acquire stable images at high resolution without phototoxicity.

An in vivo imaging protocol to monitor primary sensory axons following dorsal root crush is described. The procedures utilize wide-field fluorescence microscopy and thy1-YFP transgenic mice, and permit repeated imaging of axon regeneration over 4 cm in the PNS and axon interactions with the interface of the CNS.

The primary sensory axons injured by spinal root injuries fail to regenerate into the spinal cord, leading to chronic pain and permanent sensory loss. ...

Surgical Technique for Spinal Cord Delivery of Therapies: Demonstration of Procedure in Gottingen Minipigs

This is a compact visual description of a combination of surgical technique and device for the delivery of (gene and cell) therapies into the spinal cord. While the technique is demonstrated in the animal, the procedure is FDA-approved and currently being used for stem cell transplantation into the spinal cords of patients with ALS. While the FDA has recognized proof-of-principle data on therapeutic efficacy in highly characterized rodent models, the use of large animals is considered critical for validating the combination of a surgical procedure, a device, and the safety of a final therapy for human use. The size, anatomy, and general vulnerability of the spine and spinal cord of the swine are recognized to better model the human. Moreover, the surgical process of exposing and manipulating the spinal cord as well as closing the wound in the pig is virtually indistinguishable from the human. We believe that the healthy pig model represents a critical first step in the study of procedural safety.

Short visual description of the surgical technique and device used for the delivery of (gene and cell) therapies into the spinal cord. The technique is demonstrated in the animal but is entirely translatable and currently being used for human application.

This is a  compact visual description of a combination of surgical technique and device  for the  delivery of (gene and cell) therapies into the spinal ...

A Method of Nodose Ganglia Injection in Sprague-Dawley Rat

Afferent signaling via the vagus nerve transmits important general visceral information to the central nervous system from many diverse receptors located in the organs of the abdomen and thorax. The vagus nerve communicates information from stimuli such as heart rate, blood pressure, bronchopulmonary irritation, and gastrointestinal distension to the nucleus of solitary tract of the medulla. The cell bodies of the vagus nerve are located in the nodose and petrosal ganglia, of which the majority are located in the former. The nodose ganglia contain a wealth of receptors for amino acids, monoamines, neuropeptides, and other neurochemicals that can modify afferent vagus nerve activity. Modifying vagal afferents through systemic peripheral drug treatments targeted at the receptors on nodose ganglia has the potential of treating diseases such as sleep apnea, gastroesophageal reflux disease, or chronic cough. The protocol here describes a method of injection neurochemicals directly into the nodose ganglion. Injecting neurochemicals directly into the nodose ganglia allows study of effects solely on cell bodies that modulate afferent nerve activity, and prevents the complication of involving the central nervous system as seen in systemic neurochemical treatment. Using readily available and inexpensive equipment, intranodose ganglia injections are easily done in anesthetized Sprague-Dawley rats.

Afferent vagal signaling transmits important information to central nervous system from receptors located in organs of the abdomen and thorax. The nodose ganglia of vagus nerves contain many types of receptors that modulate vagal activity. This protocol describes a method of local injections of neurochemicals into the nodose ganglia.

Afferent signaling via the vagus nerve transmits important general visceral information to the central nervous system from many diverse receptors located ...

Dorsal Root Ganglion Injection and Dorsal Root Crush Injury as a Model for Sensory Axon Regeneration

Achieving axon regeneration after nervous system injury is a challenging task. As different parts of the central nervous system (CNS) differ from each other anatomically, it is important to identify an appropriate model to use for the study of axon regeneration. By using a suitable model, we can formulate a specific treatment based on the severity of injury, the neuronal cell type of interest, and the desired spinal tract for assessing regeneration. Within the sensory pathway, DRG neurons are responsible for relaying sensory information from the periphery to the CNS. We present here a protocol that uses a DRG injection with a viral vector and a concurrent dorsal root crush injury in the lower cervical spinal cord of an adult rat as a model to study sensory axon regeneration. As demonstrated using a control virus, AAV5-GFP, we show the effectiveness of a direct DRG injection in transducing DRG neurons and tracing sensory axons into the spinal cord. We also show the effectiveness of the dorsal root crush injury in denervating the forepaw as an injury model for evaluating axon regeneration. Despite the requirement for specialized training to perform this invasive surgical procedure, the protocol is flexible, and potential users can modify many parts to accommodate their experimental requirements. Importantly, it can serve as a foundation for those in search of a suitable animal model for their studies. We believe that this article will help new users to learn the procedure in a very efficient and effective manner.

This protocol presents the use of a dorsal root ganglion (DRG) injection with a viral vector and a concurrent dorsal root crush injury in an adult rat as a model to study sensory axon regeneration. This model is suitable for investigating the use of gene therapy to promote sensory axon regeneration.

Achieving axon regeneration after nervous system injury is a challenging task. As different parts of the central nervous system (CNS) differ from each ...

Real-Time Assessment of Spinal Cord Microperfusion in a Porcine Model of Ischemia/Reperfusion

Spinal cord injury is a devastating complication of aortic repair. Despite developments for the prevention and treatment of spinal cord injury, its incidence is still considerably high and therefore, influences patient outcome. Microcirculation plays a key role in tissue perfusion and oxygen supply and is often dissociated from macrohemodynamics. Thus, direct evaluation of spinal cord microcirculation is essential for the development of microcirculation-targeted therapies and the evaluation of existing approaches in regard to spinal cord microcirculation. However, most of the methods do not provide real-time assessment of spinal cord microcirculation. The aim of this study is to describe a standardized protocol for real-time spinal cord microcirculatory evaluation using laser-Doppler needle probes directly inserted in the spinal cord. We used a porcine model of ischemia/reperfusion to induce deterioration of the spinal cord microcirculation. In addition, a fluorescent microsphere injection technique was used. Initially, animals were anesthetized and mechanically ventilated. Thereafter, laser-Doppler needle probe insertion was performed, followed by the placement of cerebrospinal fluid drainage. A median sternotomy was performed for exposure of the descending aorta to perform aortic cross-clamping. Ischemia/reperfusion was induced by supra-celiac aortic cross-clamping for a total of 48 min, followed by reperfusion and hemodynamic stabilization. Laser-Doppler Flux was performed in parallel with macrohemodynamic evaluation. In addition, automated cerebrospinal fluid drainage was used to maintain a stable cerebrospinal pressure. After completion of the protocol, animals were sacrificed, and the spinal cord was harvested for histopathological and microsphere analysis. The protocol reveals the feasibility of spinal cord microperfusion measurements using laser-Doppler probes and shows a marked decrease during ischemia as well as recovery after reperfusion. Results showed comparable behavior to fluorescent microsphere evaluation. In conclusion, this new protocol might provide a useful large animal model for future studies using real-time spinal cord microperfusion assessment in ischemia/reperfusion conditions.

Spinal cord microcirculation plays a pivotal role in spinal cord injury. Most methods do not allow real-time assessment of spinal cord microcirculation, which is essential for the development of microcirculation-targeted therapies. Here, we propose a protocol using Laser-Doppler-Flow Needle probes in a large animal model of ischemia/reperfusion.

Spinal cord injury is a devastating complication of aortic repair. Despite developments for the prevention and treatment of spinal cord injury, its ...

Advanced Imaging of Dorsal Root Ganglion Neurons for Sensory Modality Research

In Vivo Calcium Imaging of Dorsal Root Ganglia Neurons' Response to Somatic and Visceral Stimuli

A technique is described for surgically exposing the dorsal root ganglion (DRG) of the lumbar-6 in a live, anesthetized laboratory mouse, along with the protocol for in vivo calcium imaging of the exposed DRG in response to various visceral and somatic stimuli. Pirt-GCaMP6s mice or C57BL6 mice intrathecally injected with AAV viruses packaged with GCaMP6s were utilized to capture Ca2+ transients. The amplitude of these transients indicates sensitivity to specific sensory modalities. Afferent fibers originate from internal organs, with primary neuronal cell bodies in spinal or vagal ganglia. Studies on visceral nociception and acupuncture analgesia can potentially be conducted on primary sensory neurons using advanced imaging technologies like in vivo calcium imaging, allowing for the recording of neuronal activity ensembles in the intact animal during stimulation or intervention. The responses of DRG neuron ensembles to somatic and visceral stimuli applied to their corresponding receptive fields were recorded. This technique illustrates how neuronal populations react to various types of somatic and visceral stimuli. It is possible to comprehensively compare neuronal ensemble responses to different stimuli, which is a particularly valuable approach in research on visceral pain and segmental mechanisms of somatic stimulation, such as acupuncture.

The present protocol outlines in vivo calcium imaging for measuring the responses of ensembles of lumbar-6 DRG neurons to somatic and visceral stimuli. Thorough comparisons can be made among neurons responding to different stimuli. This protocol is valuable for investigating mechanisms of visceral pain and somatic stimulation, such as acupuncture.

A technique is described for surgically exposing the dorsal root ganglion (DRG) of the lumbar-6 in a live, anesthetized laboratory mouse, along with the ...

Lumbar Puncture Delivery of AAV9 in Juvenile Rats for CNS Gene Therapy

Intrathecal Vector Delivery in Juvenile Rats via Lumbar Cistern Injection

Gene therapy is a powerful technology to deliver new genes to a patient for the treatment of disease, be it to introduce a functional gene, inactivate a toxic gene, or provide a gene whose product can modulate the biology of the disease. The delivery method for the therapeutic vector can take many forms, ranging from intravenous infusion for systemic delivery to direct injection into the target tissue. For neurodegenerative disorders, it is often desirable to skew transduction towards the brain and/or spinal cord. The least invasive approach to target the entire central nervous system involves injection into the cerebrospinal fluid (CSF), allowing the therapeutic to reach a large fraction of the central nervous system. The safest approach to deliver a vector into the CSF is the lumbar intrathecal injection, where a needle is introduced into the lumbar cistern of the spinal cord. This technique, also known as a lumbar puncture, has been widely used in neonatal and adult rodents and in large animal models. While the technique is similar across species and developmental stages, subtle differences in size, structure, and elasticity of tissues surrounding the intrathecal space require accommodations in the approach. This article describes a method for performing lumbar puncture in juvenile rats to deliver an adeno-associated serotype 9 vector. Here, 25-35 &#181;L of vector were injected into the lumbar cistern, and a green fluorescent protein (GFP) reporter was used to evaluate the transduction profile resulting from each injection. The benefits and challenges of this approach are discussed.

Author Spotlight: Assessing Intrathecal Gene Therapy Efficacy in Juvenile Rats

A surgical procedure is described to perform injections into the lumbar cistern of the juvenile rat. This approach has been used for the intrathecal delivery of gene therapy vectors, but it is anticipated that this approach can be used for a variety of therapeutics, including cells and drugs.

Gene therapy is a powerful technology to deliver new genes to a patient for the treatment of disease, be it to introduce a functional gene, inactivate a ...

Surgical Technique for Lumbar Spinal Catheter Insertion in Pigs Enabling Continuous Access to the Thecal Sac in a Terminal Setup

Pigs are increasingly used as a large animal model for pharmacologic CNS research due to the anatomical and physiological similarities between the porcine and human central nervous systems (CNS). However, accessing the cerebrospinal fluid (CSF) in larger pig breeds by conventional lumbar puncture techniques can be challenging due to an oblique orientation of the spinal spinous processes and a limited interlaminar space. Accordingly, an open surgical procedure for inserting a lumbar spinal catheter for continuous CSF sampling at the L4/L5 level in pigs is thoroughly described in this work. After positioning the pig and identifying the anatomical landmarks, a dorsal midline surgical incision is made to expose the spinous processes. By advancing the introducer needle, the spinal catheter is inserted inside the thecal sac of the spinal canal while leaving the bone structures of the spine intact. This method allows continuous infusion into or sampling from the porcine thecal sac with minimal bleeding or CSF leakage. The procedure is simple, time-efficient, and reproducible across different experimental setups, offering significant potential for various pre-clinical studies, including pharmacokinetic research, surgical training, and spinal cord injury models.

We present a technique for inserting a lumbar spinal catheter at the L4-L5 level in a 3-month-old Danish Landrace pig as part of a terminal research protocol, enabling continuous infusion or CSF sampling from the thecal sac.

Pigs are increasingly used as a large animal model for pharmacologic CNS research due to the anatomical and physiological similarities between the porcine ...

Lumbar Intrathecal Injection of Gene Therapy Vectors for Central Nervous System Targeting in Mice and Rats

One method to target the central nervous system for treating neurological diseases involves utilizing the lumbar intrathecal route of administration. This approach bypasses the blood-brain barrier to directly access the cerebrospinal fluid and preferentially target cells within the central nervous system. Multiple published preclinical studies employing the lumbar intrathecal injection route have contributed to the development of gene therapy clinical trials; however, the described protocols are variable and dispersed across multiple resources. Here, a comprehensive set of protocols for lumbar intrathecal injections in neonatal, juvenile, and adult mice and rats for preclinical gene therapy studies is presented. With proper training, this injection technique can be performed quickly and reliably. In addition to detailing the injection protocol at each developmental stage, associated parameters, such as injection volume, that can influence study outcomes are discussed. To demonstrate the application of lumbar intrathecal injections for targeting the central nervous system, the expression of adeno-associated virus serotype 9 in the brain, spinal cord, and peripheral tissues is presented following a successful or unsuccessful injection.

A lumbar intrathecal injection represents a translationally relevant route of administration for delivering gene therapy to the central nervous system. This comprehensive standardized protocol for lumbar intrathecal injections in neonatal, juvenile, and adult mice and rats aims to guide researchers in adopting this technique for preclinical gene therapy studies.

One method to target the central nervous system for treating neurological diseases involves utilizing the lumbar intrathecal route of administration. This ...

Laminotomy for Lumbar Dorsal Root Ganglion Access and Injection in Swine

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