Name: direct injection of a lentiviral vector highlights multiple motor pathways in the rat spinal cord
Uploaded: 2019-03-15T14:30:00
Description: Watch this Scientific Journal Video about Direct Injection of a Lentiviral Vector Highlights Multiple Motor Pathways in the Rat Spinal Cord at JoVE.com

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 ...

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&#39;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&#39;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. ...

Viral vectors can introduce genetic material into cells to compensate for defective genes, to up-regulate important growth proteins, or to manufacture markers that can trace neural circuits. Introducing viral vectors into the nervous system is challenging due to innate biological barriers. Direct injection into spinal cord tissue bypasses these barriers, allowing for access to cell bodies or synapses.Correctly targeting the L1 to L4 spinal levels can be difficult. It's important to use landmarks to verify the target and remember that these segments line to the T11 to T13 vertebrae. On the day of the procedure, plug the injector into the micro-pump and place the micro-pump into a micromanipulator with a vernier scale.Use a syringe with a flexible needle to carefully load a colored dye into one of the glass needles and insert the glass needle into the injector. Then confirm that the needle is seeded correctly into the washers, and that the injector cap is screwed on tight, and that the steel injector needle is extended approximately three-quarters of the length of the glass needle. Before beginning the procedure, thaw at least one microliter, plus a small volume of extra virus per injection from freezer storage, and confirm a lack of response of toe pinch in an anesthetized rat.Shave along the dorsal midline from the hips to the inferior angle of the scapulae of the animal, pulling the skin taut for an easier and more precise shave, and disinfect the exposed skin with sequential 5%iodine and 70%ethanol scrubs. Apply ophthalmic ointment to both eyes. Then place the animal on a sterile cloth and place gauze underneath the bladder to collect any urine.To identify the site of the skin incision, press the fingers gently at the last rib to locate the L1 vertebra. Using this vertebra as a landmark and holding the skin taut by gentle spreading while pressing firmly with the scalpel, use a number 10 surgical blade to make a three to four-centimeter skinned incision ending just inferior to L1 to expose the muscle. Use forceps and scissors to feel for the spinous processes.Make small rostral cuts to allow room to grab securely onto an upper process with rat tooth forceps. And make two long, deep cuts as close to the processes as possible. After securing the lateral muscles in place, clear the muscle around the processes to determine the shape of their heads.Then visualize the shapes of the heads of the spinous processes in order to identify the laminectomy site. The T11 and the joining T12 and T13 processes will be the site of the laminectomy. Once the target area has been correctly identified, gently spread the vertebrae to reveal the intervertebral ligaments, and use rongeurs to take small bites of the spinous processes and the dorsal aspect of the vertebrae.Clear the bone away from the midline so that the midline blood vessel can be observed, leaving a window that clearly reveals the spinal cord tissue and is free of debris. Then fasten stabilizing forceps to the spinous processes rostral and caudal to the laminectomy window to secure the animal in a spinal holder. And raise the abdomen of the animal to negate the effect of breathing movements.To load the virus into the injector, pipette approximately five microliters of the thawed stock onto a piece of Parafilm, and position the glass needles so that the tip is inside the drop. Use the micro-pump to withdraw up to four microliters of virus at a rate of 20 to 100 nanoliters per second, and set the controller to inject before releasing a small amount of virus from the needle tip to ensure that it is not blocked. Wipe off any excess virus with a laboratory wipe and position the micromanipulators so that the vernier scale is visible.Place the needle at the midline of the spinal cord, and use the vernier scale on the micromanipulator to direct the needle laterally 0.8 millimeters. Then lower the needle to the spinal chord until it is indenting, but not puncturing, the dura, and use a quick twisting motion to puncture the dura with the needle until the needle tip has sunk to a depth of 1.5 millimeters. Once the needle is in place, program the injector to deliver the virus at a rate of 400 nanoliters per minute, and confirm that virus is entering the spinal chord by observing the progress of the dye front.Once the injection is finished, allow the needle to rest in the spinal cord for two to five minutes to facilitate the diffusion of the virus, before slowly withdrawing the needle for positioning at the next injection site. After the last injection, remove the animal from the spinal holder to allow the removal of any devices used to spread the lateral muscle. Then close the muscle with a 4.0 chromic catgut suture and staple the skin with nine-millimeter wound clips.Four weeks after a successful injection, significant GFP expression is observed in the neurons within the gray matter of the thoracic spinal cord on the side, ipsilateral to the injection. In the white matter, GFP expression is observed in the axons in the ipsilateral cord, especially in areas typical to the propriospinal axons. A higher magnification of the gray matter neurons reveals an example of a typical signal expression in neuronal cell bodies and dendrites.GFP expression in neurons can also be observed in brainstem nuclei, such as within the pontine reticular formation. The key factors for a successful spinal cord injection procedure are correct vertebral targeting and ensuring that virus is entering the tissue If a rostral spinal cord injury is performed prior to this procedure, direct injection can be used to trace neurons that reestablish connections around the injury.

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A Functional Motor Unit in the Culture Dish: Co-culture of Spinal Cord Explants and Muscle Cells

Human primary muscle cells cultured aneurally in monolayer rarely contract spontaneously because, in the absence of a nerve component, cell differentiation is limited and motor neuron stimulation is missing1. These limitations hamper the in vitro study of many neuromuscular diseases in cultured muscle cells. Importantly, the experimental constraints of monolayered, cultured muscle cells can be overcome by functional innervation of myofibers with spinal cord explants in co-cultures.
Here, we show the different steps required to achieve an efficient, proper innervation of human primary muscle cells, leading to complete differentiation and fiber contraction according to the method developed by Askanas2. To do so, muscle cells are co-cultured with spinal cord explants of rat embryos at ED 13.5, with the dorsal root ganglia still attached to the spinal cord slices. After a few days, the muscle fibers start to contract and eventually become cross-striated through innervation by functional neurites projecting from the spinal cord explants that connecting to the muscle cells. This structure can be maintained for many months, simply by regular exchange of the culture medium.
The applications of this invaluable tool are numerous, as it represents a functional model for multidisciplinary analyses of human muscle development and innervation. In fact, a complete de novo neuromuscular junction installation occurs in a culture dish, allowing an easy measurement of many parameters at each step, in a fundamental and physiological context.
Just to cite a few examples, genomic and/or proteomic studies can be performed directly on the co-cultures. Furthermore, pre- and post-synaptic effects can be specifically and separately assessed at the neuromuscular junction, because both components come from different species, rat and human, respectively. The nerve-muscle co-culture can also be performed with human muscle cells isolated from patients suffering from muscle or neuromuscular diseases3, and thus can be used as a screening tool for candidate drugs. Finally, no special equipment but a regular BSL2 facility is needed to reproduce a functional motor unit in a culture dish. This method thus is valuable for both the muscle as well as the neuromuscular research communities for physiological and mechanistic studies of neuromuscular function, in a normal and disease context.

Cultured muscle cells are an inadequate model to recapitulate innervated muscle in vivo. A functional motor unit can be reproduced in vitro by innervation of differentiated human primary muscle cells using rat embryo spinal cord explants. This article describes how co-cultures of spinal cord explants and muscle cells are established.

Human primary muscle cells cultured  aneurally in monolayer rarely contract spontaneously because, in the absence of  a nerve component, cell ...

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 ...

Spinal Cord Transection in the Larval Zebrafish

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

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

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

Diffusion Imaging in the Rat Cervical Spinal Cord

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

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

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

Zebrafish In Situ Spinal Cord Preparation for Electrophysiological Recordings from Spinal Sensory and Motor Neurons

Zebrafish, first introduced as a developmental model, have gained popularity in many other fields. The ease of rearing large numbers of rapidly developing organisms, combined with the embryonic optical clarity, served as initial compelling attributes of this model. Over the past two decades, the success of this model has been further propelled by its amenability to large-scale mutagenesis screens and by the ease of transgenesis. More recently, gene-editing approaches have extended the power of the model.
For neurodevelopmental studies, the zebrafish embryo and larva provide a model to which multiple methods can be applied. Here, we focus on methods that allow the study of an essential property of neurons, electrical excitability. Our preparation for the electrophysiological study of zebrafish spinal neurons involves the use of veterinarian suture glue to secure the preparation to a recording chamber. Alternative methods for recording from zebrafish embryos and larvae involve the attachment of the preparation to the chamber using a fine tungsten pin1,2,3,4,5. A tungsten pin is most often used to mount the preparation in a lateral orientation, although it has been used to mount larvae dorsal-side up4. The suture glue has been used to mount embryos and larvae in both orientations. Using the glue, a minimal dissection can be performed, allowing access to spinal neurons without the use of an enzymatic treatment, thereby avoiding any resultant damage. However, for larvae, it is necessary to apply a brief enzyme treatment to remove the muscle tissue surrounding the spinal cord. The methods described here have been used to study the intrinsic electrical properties of motor neurons, interneurons, and sensory neurons at several developmental stages6,7,8,9.

Zebrafish In Situ Spinal Cord Preparation for Electrophysiological Recordings from Spinal Sensory and Motor Neurons

This manuscript describes methods for electrophysiological recordings from spinal neurons of zebrafish embryos and larvae. The preparation maintains neurons in situ and often involves minimum dissection. These methods allow for the electrophysiological study of a variety of spinal neurons, from the initial electrical excitability acquisition through the early larval stages.

Zebrafish, first introduced as a developmental model, have gained popularity in many other fields. The ease of rearing large numbers of rapidly developing ...

Laminectomy and Spinal Cord Window Implantation in the Mouse

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

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

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

Thoracic Spinal Cord Hemisection Surgery and Open-Field Locomotor Assessment in the Rat

Spinal cord injury (SCI) causes disturbances in motor, sensory, and autonomic function below the level of the lesion. Experimental animal models are valuable tools to understand the neural mechanisms involved in locomotor recovery after SCI and to design therapies for clinical populations. There are several experimental SCI models including contusion, compression, and transection injuries that are used in a wide variety of species. A hemisection involves the unilateral transection of the spinal cord and disrupts all ascending and descending tracts on one side only. Spinal hemisection produces a highly selective and reproducible injury in comparison to contusion or compression techniques that is useful for investigating neural plasticity in spared and damaged pathways associated with functional recovery. We present a detailed step-by-step protocol for performing a thoracic hemisection at the T8 vertebral level in the rat that results in an initial paralysis of the hindlimb on the side of the lesion with graded spontaneous recovery of locomotor function over several weeks. We also provide a locomotor scoring protocol to assess functional recovery in the open-field. The locomotor assessment provides a linear recovery profile and can be performed both early and repeatedly after injury in order to accurately screen animals for appropriate time points in which to conduct more specialized behavioral testing. The hemisection technique presented can be readily adapted to other transection models and species, and the locomotor assessment can be used in a variety of SCI and other injury models to score locomotor function.

The rat thoracic spinal hemisection is a valuable and reproducible model of unilateral spinal cord injury to investigate the neural mechanisms of locomotor recovery and treatment efficacy. This article includes a detailed step-by-step guide to perform the hemisection procedure and to assess locomotor performance in an open-field arena.

Spinal cord injury (SCI) causes disturbances in motor, sensory, and autonomic function below the level of the lesion. Experimental animal models are ...

3D Kinematic Analysis for the Functional Evaluation in the Rat Model of Sciatic Nerve Crush Injury

Compared to the Sciatic Functional Index (SFI), kinematic analysis is a more reliable and sensitive method for performing functional evaluations of sciatic nerve injury rodent models. In this protocol, we describe a novel kinematic analysis method that uses a three-dimensional (3D) motion capture apparatus for functional evaluations using a rat sciatic nerve crush injury model. First, the rat is familiarized with treadmill walking. Markers are then attached to the designated bone landmarks and the rat is made to walk on the treadmill at the desired speed. Meanwhile, the posterior limb movements of the rat are recorded using four cameras. Depending on the software used, marker tracings are created using both automatic and manual modes and the desired data are produced after subtle adjustments. This method of kinematic analysis, which uses a 3D motion capture apparatus, offers numerous advantages, including superior precision and accuracy. Many more parameters can be investigated during the comprehensive functional evaluations. This method has several shortcomings that require consideration: The system is expensive, can be complicated to operate, and may produce data deviations due to skin shifting. Nevertheless, kinematic analysis using a 3D motion capture apparatus is useful for performing functional anterior and posterior limb evaluations. In the future, this method may become increasingly useful for generating accurate assessments of various traumas and diseases.

We introduce a kinematic analysis method that uses a three-dimensional motion capture apparatus containing four cameras and data processing software for performing functional evaluations during fundamental research involving rodent models.

Compared to the Sciatic Functional Index (SFI), kinematic analysis is a more reliable and sensitive method for performing functional evaluations of ...

Intra-Arterial Delivery of Neural Stem Cells to the Rat and Mouse Brain: Application to Cerebral Ischemia

Neural stem cell (NSC) therapy is an emerging innovative treatment for stroke, traumatic brain injury and neurodegenerative disorders. As compared to intracranial delivery, intra-arterial administration of NSCs is less invasive and produces a more diffuse distribution of NSCs within the brain parenchyma. Further, intra-arterial delivery allows the first-pass effect in the brain circulation, lessening the potential for trapping of cells in peripheral organs, such as liver and spleen, a complication associated with peripheral injections. Here, we detail the methodology, in both mice and rats, for delivery of NSCs through the common carotid artery (mouse) or external carotid artery (rat) to the ipsilateral hemisphere after an ischemic stroke. Using GFP-labeled NSCs, we illustrate the widespread distribution achieved throughout the rodent ipsilateral hemisphere at 1 d, 1 week and 4 weeks after postischemic delivery, with a higher density in or near the ischemic injury site. In addition to long-term survival, we show evidence of differentiation of GFP-labeled cells at 4 weeks. The intra-arterial delivery approach described here for NSCs can also be used for administration of therapeutic compounds, and thus has broad applicability to varied CNS injury and disease models across multiple species.

A method for delivering neural stem cells, adaptable for injecting solutions or suspensions, through the common carotid artery (mouse) or external carotid artery (rat) after ischemic stroke is reported. Injected cells are distributed broadly throughout the brain parenchyma and can be detected up to 30 d after delivery.

Neural stem cell (NSC) therapy is an emerging innovative treatment for stroke, traumatic brain injury and neurodegenerative disorders. As compared to ...

In Vivo Intracellular Recording of Type-Identified Rat Spinal Motoneurons During Trans-Spinal Direct Current Stimulation

Intracellular recording of spinal motoneurons in vivo provides a &#8220;gold standard&#8221; for determining the cells&#8217; electrophysiological characteristics in the intact spinal network and holds significant advantages relative to classical in vitro or extracellular recording techniques. An advantage of in vivo intracellular recordings is that this method can be performed on adult animals with a fully mature nervous system, and therefore many observed physiological mechanisms can be translated to practical applications. In this methodological paper, we describe this procedure combined with externally applied constant current stimulation, which mimics polarization processes occurring within spinal neuronal networks. Trans-spinal direct current stimulation (tsDCS) is an innovative method increasingly used as a neuromodulatory intervention in rehabilitation after various neurological injuries as well as in sports. The influence of tsDCS on the nervous system remains poorly understood and the physiological mechanisms behind its actions are largely unknown. The application of the tsDCS simultaneously with intracellular recordings enables us to directly observe changes of motoneuron membrane properties and characteristics of rhythmic firing in response to the polarization of the spinal neuronal network, which is crucial for the understanding of tsDCS actions. Moreover, when the presented protocol includes the identification of the motoneuron with respect to an innervated muscle and its function (flexor versus extensor) as well as the physiological type (fast versus slow) it provides an opportunity to selectively investigate the influence of tsDCS on identified components of spinal circuitry, which seem to be differently affected by polarization. The presented procedure focuses on surgical preparation for intracellular recordings and stimulation with an emphasis on the steps which are necessary to achieve preparation stability and reproducibility of results. The details of the methodology of the anodal or cathodal tsDCS application are discussed while paying attention to practical and safety issues.

This protocol describes in vivo intracellular recording of rat lumbar motoneurons with simultaneous trans-spinal direct current stimulation. The method enables us to measure membrane properties and to record rhythmic firing of motoneurons before, during and after anodal or cathodal polarization of the spinal cord.