This work was funded by a grant from the National Institute of Neurological Disorders and Stroke R01 R01NS103481 and the Shriners Hospital for Pediatric Research grants SHC 84051 and SHC 86000 and the Department of Defense (SC140089).

All of the following surgical and animal care procedures have been approved by the Animal Care and Use Committee of Temple University.
1. Pre-surgical preparations
<ol>
	<li>Prepare pulled glass needles for viral injection a few days before surgery using 3.5 nanoliter glass capillary pipettes designed for nanoliter injectors. Pull each pipette on a two-step needle puller according to the manufacturer&#8217;s instructions to create two needle templates.</li>
	<li>Refine the tip of the needle templates by cutting off approximately 1-2 mm of excess glass with microscissors. Measure approximate aperture size under a microscope with a microscope calibration slide to isolate needles with 30-40 &#181;m apertures.</li>
	<li>With the needle positioned at 30&#176;, use a micropipette beveller to create a tip with a 30-40 &#181;m aperture and a 45&#176; beveled angle. Verify aperture width with the Vernier scale on the calibration slide. Pass water and ethanol through the glass needle using a syringe with a flexible needle attachment to wash away debris and mark the needle at regular intervals with a black marker.</li>
	<li>Place the needles in a covered Petri dish previously cleaned with 70% ethanol and sterilize for 30 min in a Biosafety hood under UV light.</li>
	<li>Prepare HiRet lentivirus by removing a suitable volume from the freezer immediately prior to the procedure. 
	NOTE: A suitable volume includes the amount needed for injection (1 &#181;L per injection x number of injections) plus a small amount of extra volume to account for pipetting and loading losses. Transport and store the virus on ice when not in use.</li>
	<li>Prepare the injector by plugging it into the micropump and placing it into a micromanipulator with a Vernier scale.</li>
	<li>To prepare the glass needle, carefully load a colored dye such as red oil with a syringe outfitted with a flexible needle. Ensure that no bubbles remain in the needle.&#160;Use aseptic technique when handling the needle, and refrain from touching the tip.</li>
	<li>Insert the glass needle into the injector, ensuring that the needle is seated correctly into the washers, the injector cap is screwed on tight, and the steel injector needle is extended approximately &#190; the length of the glass needle. Virus can be loaded into the needle in a later step.</li>
</ol>
2. Anesthesia and surgical site preparation
<ol>
	<li>Weigh the animal on a digital scale. Record the pre-operative weight to determine the volume of anesthetic required and to allow for monitoring of weight post-surgery. Female Sprague-Dawley rats approximately 200&#8211;250 g were used in this protocol.</li>
	<li>Anesthetize the rat using either isoflurane inhalation or an injected ketamine/xylazine solution (k/x). Here, ketamine is injected intraperitoneally at a 67 mg/kg and xylazine at a 6.7 mg/kg dosage.</li>
	<li>Confirm an appropriate anesthetic plane by pinching the foot firmly. If reflexive withdrawal occurs, wait several additional minutes before proceeding. 
	NOTE: Also observe the whiskers, eyes and breathing rate for signs of consciousness. If the whiskers are twitching, the eye blinks when touched gently, or breathing is rapid and shallow, wait until the anesthetic plane is deeper to proceed with the protocol. Also monitor these signs throughout the laminectomy and injection surgery. If the animal displays a shallow anesthetic plane, administer a booster shot of ketamine-only equal to &#189; the original k/x dosage.</li>
	<li>Shave the rat along the dorsal midline from the hips to the inferior angle of the scapulae. Pull the skin of the animal taut for an easier and more precise shave.</li>
	<li>Apply ophthalmic ointment to both eyes.</li>
	<li>Apply antiseptic to the shaved area to sterilize the site. For the first scrub, soak sterile gauze with a 5% iodine solution and wipe away all hair and debris. Follow this with a unidirectional swipe with sterile gauze soaked in 70% ethanol, so that no area is contacted twice. Use this same technique with alternating iodine and ethanol-soaked gauze twice more.</li>
</ol>
3. Surgical field and instrument preparation
<ol>
	<li>Prepare a set of autoclaved surgical tools that include a scalpel, rongeurs, rat tooth forceps, spring scissors, hemostats, medium point curved forceps and retractors or weighted hooks by unwrapping the sterile wrap to create a sterile field.</li>
	<li>Open a package of sterile surgical gloves and place the sterile glove wrap on the table. Use this as an additional sterile field for used tools to prevent contamination of the sterile wrap.</li>
	<li>Drop a #10 scalpel blade onto the sterile field. Secure the blade to a handle with hemostats. Position sterile saline, 4.0 chromic catgut suture, and materials to control bleeding such as a cauterizer, sterile gauze, sterile cotton-tipped applicators (for muscle bleeds), or gelfoam or bonewax (for bone bleeds) in an accessible place.</li>
	<li>Retrieve the animal and set it on a sterile cloth. Place gauze underneath the bladder to collect urine. Prop up the target area with a rolled towel under the abdomen. If available, place a surgical heating pad underneath the sterile cloth, especially for longer procedures. 
	NOTE: Sterility is important during survival surgery. Keep a spray bottle of 70% ethanol on hand to maintain sterility of gloved hands, and a use bead sterilizer if instrument sterility is compromised, or between individual surgeries.</li>
</ol>
4. Exposing the vertebral column and identifying the laminectomy site
<ol>
	<li>Identify the area where a skin incision will be made by pressing the fingers gently at the last rib to locate the L1 vertebra. Using this as a landmark, make a 3-4 cm skin incision with a #10 surgical scalpel ending just inferior to L1 to expose the muscle. Hold the skin taut by gentle spreading and press firmly with the scalpel blade to ensure a clean incision.</li>
	<li>Cut and spread superficial fat&#160;with forceps and scissors&#160;if necessary. (Depending on target vertebra, there may or not be a large fat pad superficial to the muscle).&#160;</li>
	<li>Feel for the spinous processes with the flat of the scalpel blade or a finger. Often the midline area will be outlined by a &#8220;V&#8221; of white fascia on either side. Make a small rostral cut to allow room to grab securely onto an upper process with rat tooth forceps, then make 2 long, deep cuts as close to the processes as possible. At the deepest point of the cut, the dorsal surface of the vertebrae can be felt with the scalpel blade.</li>
	<li>Hold the lateral muscles aside with retractors or weighted hooks to improve visibility. Clear muscle around the processes with a scalpel, spring scissors or rongeurs to determine the shape of their heads. 
	NOTE: Remember that the spinal cord does not extend the full length of the vertebral column, as spinal cord tissue stops growing earlier in development than bone. This means that the target spinal level may be underneath a differently named vertebra.</li>
	<li>Locate the T11 and the adjoining T12 and T13 processes. 
	NOTE: Assistance in targeting correct vertebral levels can be found in a rat spinal cord atlas and previous studies outlining landmarks in the mouse, which has a very similar vertebral structure6,33. Leave a rostral spinous process such as T9 undisturbed to give a midline landmark.</li>
</ol>
5. Performing a laminectomy
<ol>
	<li>Once the target area has been correctly identified, perform laminectomies of the dorsal aspects of T11-T13. Gently spread the vertebrae to reveal intervertebral ligaments, which are good sites to insert rongeurs for the initial bite of bone. Hold the rongeurs in a half-closed position to increase fine control.</li>
	<li>Remove the spinous processes and the dorsal aspect of the vertebrae by taking small bites with the rongeurs. Be careful not to damage the spinal cord or disturb the dura. Lift slightly with the rat tooth forceps to help pull the spinal cord away from the vertebrae and decrease the tendency to hit spinal cord tissue.</li>
	<li>Clear bone away from the midline so that the midline blood vessel can be observed. Leave a window that clearly shows the spinal cord tissue and is free of debris.</li>
	<li>Gently touch the spinal cord with forceps. Some animals may reflexively jump even if their anesthetic plane is deep. Apply a few drops of a numbing agent such as lidocaine directly to the spinal cord to prevent jumping during the injection procedure.</li>
	<li>Secure the animal in a spinal holder by fastening stabilizing forceps to spinous processes rostral and caudal to the laminectomy window. Raise the abdomen of the animal using the spinal holder to negate the effect of breathing movements. This will increase needle stability and ensure appropriate depth of injection.</li>
</ol>
6. Loading virus and positioning the injector
<ol>
	<li>Load virus into the injector by pipetting approximately 5 &#181;L onto a piece of parafilm and positioning the needle so that the tip is inside the drop.</li>
	<li>Use the micropump to withdraw up to 4 &#181;L of virus at a rate of 20&#8211;100 nL/s.</li>
	<li>Set the controller to inject and release a small amount of virus from the needle to ensure the tip of the needle is not blocked. Wipe off excess virus with a laboratory wipe. 
	NOTE: A Hamilton syringe with a steel needle may be used as an alternative to pulled glass pipettes.</li>
	<li>Position the micromanipulator so that the Vernier scale is visible and position the needle at the midline of the spinal cord. 
	NOTE: The midline can sometimes be located by a large blood vessel running on the anterior surface of the spinal cord. However, this can vary in individual rats, and midline targeting should be confirmed by comparison with an intact spinous process.</li>
	<li>Direct the needle laterally by 0.8 mm using the Vernier scale on the micromanipulator.</li>
	<li>Lower the needle to the spinal cord until it is indenting, but not puncturing, the dura. Using a quick twisting motion, puncture the dura with the needle until it has sunk to a depth of 1.5 mm.</li>
</ol>
7. Injecting virus into the spinal cord
<ol>
	<li>Once the needle is in place, program the injector to inject at a rate of 400 nL/min. Confirm that virus is entering the spinal cord by observing the progress of the dye front. There should be no obvious leakage or bulging of spinal cord tissue. If leakage is observed, this can sometimes be alleviated by reducing the injection speed to 200 nL/min.</li>
	<li>Once the injection is finished, allow the needle to rest in the spinal cord for 2&#8211;5 min (depending on volume injected) to facilitate diffusion of the virus.</li>
	<li>Slowly withdraw the needle and move to the next injection site. Inject 1 &#181;L of virus into each of 6 evenly spaced sites approximately 1 mm apart along the length of the L1-L4 spinal tissue. The same needle may be used for each injection as long as it continues to function properly.</li>
</ol>
8. Wound closure and post-operative care
<ol>
	<li>Remove the animal from the spinal holder and take out retractors or hooks used to spread lateral muscle. Ensure that the wound is clear of all debris before closing.</li>
	<li>Suture the muscle using a 4.0 chromic catgut suture. Cut suture threads close to the knot to reduce likelihood of internal skin irritation.</li>
	<li>Staple the skin closed using 9 mm wound clips. To allow for optimal healing, line up the edges of the skin before stapling.</li>
	<li>Place the animal on a water convection warming pad and monitor until wakeful.</li>
	<li>Inject 5&#8211;10 mL of sterile saline subcutaneously to replenish fluids and an antibiotic such as cefazolin to prevent infection.&#160;When the animal is ambulatory, place it back in its home cage and provide initial analgesics.&#160; Monitor the rats for any sign of pain and distress and treat according to your IACUC approved procedure for alleviation of pain.</li>
</ol>

The authors have nothing to disclose.

Genetic manipulation of neurons in the brain and spinal cord has served to highlight sensory, motor and autonomic pathways via fluorescent tracing and to explore regrowth potential of neuronal tracts after injury27,28,29,30,31,32,33. Direct injection of a retrogradely transportable viral vec...

Viral vectors are important biological tools that can introduce genetic material into cells in order to compensate for defective genes, upregulate important growth proteins or manufacture marker proteins that highlight the structure and synaptic connections of their targets. This article focuses on direct injection of a highly efficient retrogradely transportable lentiviral vector into the rat spinal cord in order to highlight major motor pathways with fluorescent tracing.&#160; This method is also highly appropriate for axonal regeneration and regrowth studies to introduce proteins of interest into diverse populations of neurons and has been used to silence neurons for functional mapping studies1,2.
Many of the anatomical details of spinal motor pathways were elucidated through direct injection studies with classical tracers such as BDA and fluoro-gold3,4,5,6,7,8. These tracers are considered gold standard but may have certain disadvantages such as uptake by damaged axons, or axons in passage in the white matter surrounding an injection site9,10,11. This could lead to incorrect interpretations of pathway connectivity and may be a drawback in regeneration studies where dye absorption by damaged or severed axons could be mistaken for regenerating fibers during later analysis12.
Lentiviral vectors are popular in gene therapy studies, as they provide stable, long-term expression in neuronal populations13,14,15,16,17,18,19. However, traditionally packaged lentiviral vectors can have limited retrograde transport and may trigger immune system response when used in vivo4,20,21. A highly-efficient retrograde transport vector termed HiRet has been produced by Kato et al. by modifying the viral envelope with a rabies virus glycoprotein to create a hybrid vector that improves retrograde transport22,23.
Retrograde tracing introduces a vector into the synaptic space of a target neuron, allowing it to be taken up by that cell&#8217;s axon and transported to the cell body. Successful transport of HiRet has been demonstrated from neuronal synapses into the brains of mice and primates23,24 and from the muscle into motor neurons22. This protocol demonstrates injection into the lumbar spinal cord, specifically targeting the synaptic terminals of propriospinal interneurons and brainstem neurons. PNs receive connections from many different spinal pathways and can thus be utilized to target a diverse population of neurons in the spinal cord and brainstem. Labeled neurons in this study represent circuits innervating motor neuron pools relating to hindlimb motor function. Robust labeling is seen in the spinal cord and brainstem, including high fidelity details of dendritic arbors and axon terminals. We have also used this method in previous studies within the cervical spinal cord to label propriospinal and brainstem reticulospinal pathways25.
This protocol demonstrates injection of a viral vector into the lumbar spinal cord of a rat. As seen in Movie 1, the incision is targeted by identifying the L1 vertebra located at the last rib. This is used as a caudal landmark for a 3-4 cm incision that exposes musculature over the L1-L4 spinal cord. Laminectomies of the dorsal aspects of the T11-T13 vertebrae are performed and a beveled glass needle is directed 0.8 mm lateral from the midline and lowered 1.5 mm deep into the gray matter to inject virus.

<table><tbody><tr><td>#10 Scalpel Blades</td><td>Roboz</td><td>RS-9801-10</td><td>For use with the scalpel.</td></tr><tr><td>1 mL Syringes</td><td>Becton, Dickinson and Company</td><td>309659</td><td>For anesthetic IP injection, potential anesthetic booster shots, and antibiotic injections.</td></tr><tr><td>10mL Syringes</td><td>Becton, Dickinson and Company</td><td>309604</td><td>For injecting saline into the animal, post-surgery.</td></tr><tr><td>4.0 Chromic Catgut Suture</td><td>DemeTECH</td><td>NN374-16</td><td>To re-bind muscle during closing.</td></tr><tr><td>48000 Micropipette Beveler</td><td>World Precision Instruments</td><td>32416</td><td>Used to bevel the tips of the pulled glass capillary tubes to form functional glass needles.</td></tr><tr><td>5% Iodine Solution</td><td>Purdue Products L.P.</td><td>L01020-08</td><td>For use in sterilzation of the surgical site.</td></tr><tr><td>70% Ethanol</td><td>N/A</td><td>N/A</td><td>For sterilization of newly prepared glass needles, animal models during surgical preparation, and surgeon&#039;s hands during surgery, as well as all other minor maintainances of sterility.</td></tr><tr><td>Anesthetic (Ketamine/Xylazine Solution)</td><td>Zoetis</td><td>240048</td><td>For keeping the animal in the correct plane of consciousness during surgery.</td></tr><tr><td>Antibiotic (Cefazolin)</td><td>West-Ward Pharmaceuticals</td><td>NPC 0143-9924-90</td><td>To be injected subcutaneously to prevent infection post-surgery.</td></tr><tr><td>Bead Sterilizer</td><td>CellPoint</td><td>5-1450</td><td>To heat sterilize surgical instruments.</td></tr><tr><td>Bonewax</td><td>Fine Science Tools</td><td>19009-00</td><td>To seal up bone in the case of bone bleeding.</td></tr><tr><td>Cauterizer</td><td>Fine Science Tools</td><td>18010-00</td><td>To seal any arteries or veins severed during surgery to prevent excessive blood loss.</td></tr><tr><td>Digital Scale</td><td>Okaus</td><td>REV.005</td><td>For weighing the animal during surgical preparation.</td></tr><tr><td>Flexible Needle Attachment</td><td>World Precision Instruments</td><td>MF34G-5</td><td>For cleaning glass needles and loading red oil into glass needles.</td></tr><tr><td>Gelfoam</td><td>Pfizer</td><td>H68079</td><td>To seal up bone in the case of bone bleeding.</td></tr><tr><td>Glass Capillary Tubes</td><td>World Precision Instruments</td><td>4878</td><td>For pulled glass needles - should be designed for nanoliter injectors.</td></tr><tr><td>Hair Clippers</td><td>Oster</td><td>111038-060-000</td><td>For clearing the surgical site of hair.</td></tr><tr><td>Hemostats</td><td>Roboz</td><td>RS-7231</td><td>For general use in surgery.</td></tr><tr><td>Kimwipes</td><td>Kimtech</td><td>34155</td><td>For general use in surgery.</td></tr><tr><td>Medium Point Curved Forceps</td><td>Roboz</td><td>RS-5136</td><td>For general use in surgery.</td></tr><tr><td>Micromanipulator with a Vernier Scale</td><td>Kanetec</td><td>N/A</td><td>For precise targeting during surgery.</td></tr><tr><td>Microscissors</td><td>Roboz</td><td>RS-5621</td><td>For cutting glass whisps off of freshly pulled glass capillary tubes.</td></tr><tr><td>Microscope with Light and Vernier Scale Ocular</td><td>Leitz Wetzlar</td><td>N/A</td><td>Used to visualize and measure beveling of pulled glass capillary tubes into functional glass needles.</td></tr><tr><td>MicroSyringe Pump Controller</td><td>World Precision Instruments</td><td>62403</td><td>To control the rate of injection.</td></tr><tr><td>Nanoliter 2000 Pump Head Injector</td><td>World Precision Instruments</td><td>500150</td><td>To load and inject virus in a controlled fashion.</td></tr><tr><td>Needle Puller</td><td>Narishige</td><td>PC-100</td><td>To heat and pull apart glass capillary tubes to form glass needles.</td></tr><tr><td>Ophthalamic Ointment</td><td>Dechra Veterinary Products</td><td>RAC 0119</td><td>To protect the animal&#039;s eyes during surgery.</td></tr><tr><td>Parafilm</td><td>Bemis</td><td>PM-996</td><td>To assist with loading virus into the nanoinjector.</td></tr><tr><td>PrecisionGlide Needles (25G x 5/8)</td><td>Becton, Dickinson and Company</td><td>305122</td><td>For use with the 1mL and 10 mL syringes to allow injection of the animal model.</td></tr><tr><td>Rat Tooth Forceps</td><td>Roboz</td><td>RS-5152</td><td>For griping spinous processes.</td></tr><tr><td>Red Oil</td><td>N/A</td><td>N/A</td><td>To provide a front for visualization of virus entering tissue during injection.</td></tr><tr><td>Retractors</td><td>Roboz</td><td>RS-6510</td><td>To hold open the surgical wound.</td></tr><tr><td>Rimadyl Tablets</td><td>Bio Serv</td><td>MP275-050</td><td>For pain management post-surgery.</td></tr><tr><td>Rongeurs</td><td>Roboz</td><td>RS-8300</td><td>To remove muscle from the spinal column during surgery.</td></tr><tr><td>Scalpel Blade Handle</td><td>Roboz</td><td>RS-9843</td><td>To slice open skin and fat pad of animal model during surgery.</td></tr><tr><td>Scissors</td><td>Roboz</td><td>RS-5980</td><td>For general use in surgery.</td></tr><tr><td>Stainless Steal Wound Clips</td><td>CellPoint</td><td>201-1000</td><td>To bind the skin of the surgical wound during closing.</td></tr><tr><td>Staple Removing Forceps</td><td>Kent Scientific</td><td>INS750347</td><td>To remove the staples, should they be applied incorrectly.</td></tr><tr><td>Sterile Cloth</td><td>Phenix Research Products</td><td>BP-989</td><td>To provide a sterile surface for the operation.</td></tr><tr><td>Sterile Cotton-Tipped Applicators</td><td>Puritan</td><td>806-WC</td><td>To soak up blood in the surgical wound while maintaining sterility.</td></tr><tr><td>Sterile Gauze</td><td>Covidien</td><td>2146</td><td>To clean the surgical area and surgical tools while maintaining sterility.</td></tr><tr><td>Sterile Saline</td><td>Baxter Healthcare Corporation</td><td>281324</td><td>For use in blood clearing, and for replacing fluids post-surgery.</td></tr><tr><td>Surgical Gloves</td><td>N/A</td><td>N/A</td><td>For use by the surgeon to maintain sterile field during surgery.</td></tr><tr><td>Surgical Heating Pad</td><td>N/A</td><td>N/A</td><td>For maintaining the body temperature of the animal model during surgery.</td></tr><tr><td>Surgical Microscope</td><td>N/A</td><td>N/A</td><td>For enhanced visualization of the surgical wound.</td></tr><tr><td>Surgical Stapler</td><td>Kent Scientific</td><td>INS750546</td><td>To apply the staples.</td></tr><tr><td>T/Pump Heat Therapy Water Pump</td><td>Gaymar</td><td>TP500C</td><td>To pump warm water into the water convection warming pad.</td></tr><tr><td>Water Convection Warming Pad</td><td>Baxter Healthcare Corporation</td><td>L1K018</td><td>For use in the post-operational recovery area to maintain the body temperature of the unconscious animal.</td></tr><tr><td>Weighted Hooks</td><td>N/A</td><td>N/A</td><td>To hold open the surgical wound.</td></tr></tbody></table>

direct injection of a lentiviral vector highlights multiple motor pathways in the rat spinal cord

Introducing proteins of interest into cells in the nervous system is challenging due to innate biological barriers that limit access to most molecules. Injection directly into spinal cord tissue bypasses these barriers, providing access to cell bodies or synapses where molecules can be incorporated. Combining viral vector technology with this method allows for introduction of target genes into nervous tissue for the purpose of gene therapy or tract tracing. Here a virus engineered for highly efficient retrograde transport (HiRet) is introduced at the synapses of propriospinal interneurons (PNs) to encourage specific transport to neurons in the spinal cord and brainstem nuclei. Targeting PNs takes advantage of the numerous connections they receive from motor pathways such as the rubrospinal and reticulospinal tracts, as well as their interconnection with each other throughout spinal cord segments. Representative tracing using the HiRet vector with constitutively active green fluorescent protein (GFP) shows high fidelity details of cell bodies, axons and dendritic arbors in thoracic PNs and in reticulospinal neurons in the pontine reticular formation. HiRet incorporates well into brainstem pathways and PNs but shows age dependent integration into corticospinal tract neurons. In summary, spinal cord injection using viral vectors is a suitable method for introduction of proteins of interest into neurons of targeted tracts.

Successful injection and transport of the viral vector should result in transduction of a robust population of unilateral neurons in the spinal cord and in certain brainstem nuclei. Figure 1 demonstrates stereotypical labeling of neurons and axons in the thoracic spinal cord and in the pontine reticular formation of the brainstem at four weeks post-injection. Significant GFP expression is seen in neurons in the gray matter of the thoracic spinal cord on the side ipsilater...

Watch this Scientific Journal Video about Direct Injection of a Lentiviral Vector Highlights Multiple Motor Pathways in the Rat Spinal Cord at JoVE.com

Direct Injection of a Lentiviral Vector Highlights Multiple Motor Pathways in the Rat Spinal Cord

This protocol demonstrates injection of a retrogradely transportable viral vector into rat spinal cord tissue. The vector is taken up at the synapse and transported to the cell body of target neurons. This model is suitable for retrograde tracing of important spinal pathways or targeting cells for gene therapy applications.

Introducing proteins of interest into cells in the nervous system is challenging due to innate biological barriers that limit access to most molecules. ...

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8.6K Views.  Temple University. This protocol demonstrates injection of a retrogradely transportable viral vector into rat spinal cord tissue. The vector is taken up at the synapse and transported to the cell body of target neurons. This model is suitable for retrograde tracing of important spinal pathways or targeting cells for gene therapy applications.

Video: Direct Injection of a Lentiviral Vector Highlights Multiple Motor Pathways in the Rat Spinal Cord

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

Stereotaxic Injection of a Viral Vector for Conditional Gene Manipulation in the Mouse Spinal Cord

Intraparenchymal injection of a viral vector enables conditional gene manipulation in distinct populations of neurons or particular regions of the central nervous system. We demonstrate a stereotaxic injection technique that allows targeted gene expression or silencing in the dorsal horn of the mouse spinal cord. The surgical procedure is brief. It requires laminectomy of a single vertebra, providing for quick recovery of the animal and unimpaired motility of the spine. Controlled injection of a small vector suspension volume at low speed and use of a microsyringe with beveled glass cannula minimize the tissue lesion. The local immune response to the vector depends on the intrinsic properties of the virus employed; in our experience, it is minor and short-lived when a recombinant adeno-associated virus is used. A reporter gene such as enhanced green fluorescent protein facilitates monitoring spatial distribution of the vector, and the efficacy and cellular specificity of the transfection.

Viral vectors allow for targeted gene manipulation. We demonstrate a method for conditional gene expression or ablation in the mouse spinal cord, using stereotaxic injection of a viral vector into the dorsal horn, a prominent site of synaptic contact between primary somatosensory afferents and neurons of the central nervous system.

Intraparenchymal injection of a viral vector enables conditional gene manipulation in distinct populations of neurons or particular regions of the central ...

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 ...

Methods and Tips for Intravenous Administration of Adeno-associated Virus to Rats and Evaluation of Central Nervous System Transduction

Adeno-associated virus (AAV) vectors are a key reagent in the neurosciences for clustered regularly interspaced short palindromic repeats (CRISPR), optogenetics, cre-lox targeting, etc. The purpose of this manuscript is to aid the investigator attempting expansive central nervous system (CNS) gene transfer in the rat via tail vein injection of AAV. Wide-scale expression is relevant for conditions with widespread pathology, and a rat model is significant due to its greater size and physiologic similarities to humans compared to mice. In this example application, a wide-scale neuronal transduction is used to mimic a neurodegenerative disease that affects the entire spinal cord, amyotrophic lateral sclerosis (ALS). The efficient wide-scale CNS transduction can also be used to deliver therapeutic protein factors in pre-clinical studies. After a post-injection expression interval of several weeks, the effects of the transduction are evaluated. For a green fluorescent protein (GFP) control vector, the amount of GFP in the cerebellum is estimated quickly and reliably by a basic imaging program. For motor disease phenotypes that are induced by the ALS related protein transactive response DNA-binding protein of 43 kDa (TDP-43), the deficits are scored by escape reflex and rotarod. Beyond disease modeling and gene therapy, there are diverse potential applications for the wide-scale gene targeting described here. The expanded use of this method will aid in expediting hypothesis testing in the neurosciences and neurogenetics.

Methods for a wide-scale central nervous system gene delivery in the rat are covered. In this example, the purpose is to mimic a disease that affects the entire spinal cord. The widespread transduction can be used to deliver a therapeutic protein to the CNS from a one-time, peripheral administration.

Adeno-associated virus (AAV) vectors are a key reagent in the neurosciences for clustered regularly interspaced short palindromic repeats (CRISPR), ...

Adeno-associated Virus-mediated Transgene Expression in Genetically Defined Neurons of the Spinal Cord

Selective manipulation of spinal neuronal subpopulations has mainly been achieved by two different methods: 1) Intersectional genetics, whereby double or triple transgenic mice are generated in order to achieve selective expression of a reporter or effector gene (e.g., from the Rosa26 locus) in the desired spinal population. 2) Intraspinal injection of Cre-dependent recombinant adeno-associated virus (rAAV); here Cre-dependent AAV vectors coding for the reporter or effector gene of choice are injected into the spinal cord of mice expressing Cre recombinase in the desired neuronal subpopulation. This protocol describes how to generate Cre-dependent rAAV vectors and how to transduce neurons in the dorsal horn of the lumbar spinal cord segments L3-L5 with rAAVs. As the lumbar spinal segments L3-L5 are innervated by those peripheral sensory neurons that transmit sensory information from the hindlimbs, spontaneous behavior and responses to sensory tests applied to the hindlimb ipsilateral to the injection side can be analyzed in order to interrogate the function of the manipulated neurons in sensory processing. We provide examples of how this technique can be used to analyze genetically defined subsets of spinal neurons. The main advantages of virus-mediated transgene expression in Cre transgenic mice compared to classical reporter mouse-induced transgene expression are the following: 1) Different Cre-dependent rAAVs encoding various reporter or effector proteins can be injected into a single Cre transgenic line, thus overcoming the need to create several multiple transgenic mouse lines. 2) Intraspinal injection limits manipulation of Cre-expressing cells to the injection site and to the time after injection. The main disadvantages are: 1) Reporter gene expression from rAAVs is more variable. 2) Surgery is required to transduce the spinal neurons of interest. Which of the two methods is more appropriate depends on the neuron population and research question to be addressed.

Intraspinal injection of recombinase dependent recombinant adeno-associated virus (rAAV) can be used to manipulate any genetically labelled cell type in the spinal cord. Here we describe how to transduce neurons in the dorsal horn of the lumbar spinal cord. This technique enables functional interrogation of the manipulated neuron subtype.

Selective manipulation of spinal neuronal subpopulations has mainly been achieved by two different methods: 1) Intersectional genetics, whereby double or ...

Direct Intrathecal Injection of Recombinant Adeno-associated Viruses in Adult Mice

Intrathecal (IT) injection of adeno-associated virus (AAV) has drawn considerable interest in CNS gene therapy by virtue of its safety, noninvasiveness, and excellent transduction efficacy in the CNS.&#160;Previous studies have demonstrated the therapeutic potency of AAV-delivered gene therapy in neurodegenerative disorders by IT administration. However, high rates of unpredictable failure due to the technical limitation of IT administration in small animals have been reported. Here, we established a scoring system to indicate the success extent of lumbar puncture in small animals by adding 1% lidocaine hydrochloride into the injection solution. We further show that the extent of transient weakness following injection can predict the transduction efficiency of AAV. Thus, this IT injection method can be used to optimize therapeutic trials in mouse models of CNS diseases that afflict wide regions of the CNS.

Here we present a direct intrathecal injection technique using 1% lidocaine hydrochloride in a viral solution to ensure efficient adeno-associated virus delivery to small animals and establish a scoring system to predict transduction efficiency in the central nervous system according to the degree of transient weakness induced by lidocaine.

Intrathecal (IT) injection of adeno-associated virus (AAV) has drawn considerable interest in CNS gene therapy by virtue of its safety, noninvasiveness, ...

Stereotaxic Intracranial Delivery of Chemicals, Proteins or Viral Vectors to Study Parkinson's Disease

Parkinson's disease (PD) is a progressive disorder traditionally defined by resting tremor and akinesia, primarily due to loss of dopaminergic neurons in the substantia nigra. Affected brain areas display intraneuronal fibrillar inclusions consisting mainly of alpha-synuclein (asyn) proteins. No animal model thus far has recapitulated all characteristics of this disease. Here, we describe the use of stereotaxic injection to deliver chemicals, proteins, or viral vectors intracranially in order to mimic various aspects of PD. These methods are well-established and widely used throughout the PD field. Stereotaxic injections are incredibly flexible; they can be adjusted in concentration, age of animal used for injection, brain area targeted and in animal species used. Combinations of substances allow for rapid variations to assess treatments or alter severity of the pathology or behavioral deficits. By injecting toxins into the brain, we can mimic inflammation and/or a severe loss of dopaminergic neurons resulting in substantial motor phenotypes. Viral vectors can be used to transduce cells to mimic genetic or mechanistic aspects. Preformed fibrillar asyn injections best recapitulate the progressive phenotype over an extended period of time. Once these methods are established, it can be economical to generate a new model compared to creating a new transgenic line. However, this method is labor intensive as it requires 30 minutes to four hours per animal depending on the model used. Each animal will have a slightly different targeting and therefore create a diverse cohort which on one hand can be challenging to interpret results from; on the other hand, help mimic a more realistic diversity found in patients. Mistargeted animals can be identified using behavioral or imaging readouts, or only after being sacrificed leading to smallercohort size after the study has already been concluded. Overall, this method is a rudimentary but effective way to assess a diverse set of PD aspects.

We describe how to successfully inject solutions into specific brain areas of rodents using a stereotaxic frame. This survival surgery is a well-established method used to mimic various aspects of Parkinson's disease.

Parkinson's disease (PD) is a progressive disorder traditionally defined by resting tremor and akinesia, primarily due to loss of dopaminergic neurons in ...

Targeting the Corticospinal Tract in Neonatal Rats with a Double-Viral Vector using Combined Brain and Spine Surgery

Successfully tackling the obstacles that constrain research on neonatal rats is important for studying the differences in outcomes seen in pediatric spinal cord injuries (SCIs) compared to adult SCIs. In addition, reliably introducing therapies into the target cells of the central nervous system (CNS) can be challenging, and inaccuracies can compromise the efficacy of the study or therapy. This protocol combines viral vector technology with a novel surgical technique&#160;to accurately introduce gene therapies into neonatal rats at postnatal day 5. Here, a virus engineered for retrograde transport (retroAAV2) of Cre is introduced at the axon terminals of corticospinal neurons in the spinal cord, where it is subsequently transported to the cell bodies. A double-floxed&#160;inverted orientation (DIO) designer receptor exclusively activated by designer drug(s) (DREADD) virus is then injected into the somatomotor cortex of the brain. This double-infection technique promotes the expression of the DREADDs only in the co-infected corticospinal tract (CST) neurons.&#160;Thus, the simultaneous co-injection of the somatomotor cortex and cervical CST terminals is a valid method for studying the chemogenetic modulation of recovery following cervical SCI models in neonatal rats.

This protocol demonstrates a novel method for applying gene therapies to subpopulations of cells in neonatal rats at postnatal ages 5-10 days by injecting an anterograde chemogenetic modifier into the somatomotor cortex and a retrogradely transportable Cre recombinase into the cervical spinal cord.

Successfully tackling the obstacles that constrain research on neonatal rats is important for studying the differences in outcomes seen in pediatric ...

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 ...

Direct Vagus Nerve Injection Protocol For Rats

There is a relative abundance of strategies and methodologies to facilitate drug delivery to the central nervous system. However, drug delivery directly to the peripheral nervous system is less common, with fewer detailed methods publications available to aid researchers. Here, we describe a direct nerve injection method for peripheral nervous system drug delivery, using the vagus nerve as a model nerve. This method can be used in the treatment of autonomic nervous system disorders through targeting of the left vagus nerve, although this general injection method can be extrapolated to injection of other nerves with minor modification. This method explains all critical steps involved in the procedure involving microsurgery in anesthetized adult rats under a dissecting microscope. The use of a tracking dye is described to facilitate the monitoring of injection fidelity in real time. Illustrations of successful and failed injections are provided. If carried out properly, direct vagus nerve injections can be conducted in a safe manner that is well-tolerated by the rat without post-delivery complications. For example, once the surgeons were trained in this method, six out of six rats were successfully injected without any complications. This method of direct nerve injection for preclinical rat studies is capable of delivering agents (including but not limited to gene therapy) to peripheral nerves.

We present a protocol for direct vagus nerve injection in rats, enabling drug delivery directly into the nerve without post-injection complications. This method applies to preclinical neurological studies involving autonomic nervous system manipulation. It can be used for direct nerve injection for other nerves in rats and other species, with necessary modifications.

There is a relative abundance of strategies and methodologies to facilitate drug delivery to the central nervous system. However, drug delivery directly ...