This work was supported by grants from the Christopher and Dana Reeve Foundation, the Medical Research Council, the European Research Council ECMneuro, and the Cambridge NHMRC Biomedical Research Center. We would like to express our deepest gratitude to Heleen Merel van &#8217;t Spijker and Justyna Barratt for their technical assistance during the filming. We would like to thank Dr. Elizabeth Moloney and Professor Joost Verhaagen (Netherlands Institute for Neuroscience) for assisting in AAV production.

All the following animal procedures were conducted in accordance with the United Kingdom Animals (Scientific Procedures) Act 1986. If unfamiliar with these procedures, please check with local/national regulations and seek veterinary advice before starting the protocol.
1. Choosing a Suitable Strain of Animals
NOTE: A dorsal root crush injury results in the loss of sensation and paw deafferentation. Common adverse effects of forepaw deafferentation can include over-grooming, self-mutilation, and paw autotomy.
<ol>
	<li>Obtain animals for the procedure. 
	NOTE: For rats, the more active strains, such as Lister-Hooded and Wistar, have a higher incidence of paw autotomy after deafferentation as compared to the more docile strains, such as Lewis and Sprague-Dawley. If possible, the most docile strain, Lewis, should always be considered first. Should a particularly active strain be necessary for the experiment due to genetic modification or specific behavioral assessment requirements, veterinary treatment should be in place to address expected adverse effects.</li>
</ol>
2. Preparing the Virus for Injection
CAUTION: Handle all viruses in accordance with biological and laboratory safety regulations.
<ol>
	<li>Dilute or concentrate the virus to a titer of 2 x 1012 genome copy (GC)/mL following individual virus packaging protocols9. 
	NOTE: Here, the virus was diluted in Dulbecco&#39;s phosphate-buffered saline (D-PBS) with 5% sucrose. The titer may have to be optimized depending on the type of virus, promoter, construct size, and targeted cell type.</li>
	<li>Optional: Add 1% colored dye (e.g., Fast Green) to the virus solution for easy visualization of the injection later on. Mix the solution very gently.</li>
	<li>Keep the solution on ice for immediate use (within 4 h) or in a -80 &#176;C freezer for long-term storage.</li>
</ol>
3. Performing the Preoperative Preparation of the Animal
NOTE: Due to the extreme invasiveness of the surgery, aseptic techniques should be used at all times.
<ol>
	<li>Prepare and sterilize all surgical tools and the spinal stereotaxic frame prior to starting the surgery. For example, sterilize the tools in an autoclave at 134 &#176;C. Use a fresh set of sterilized tools for each animal.</li>
	<li>Anesthetize the animal. Confirm proper anesthetization by pinching the paw to test for the withdrawal reflex or by touching the cornea for the eyelid reflex.
	<ol>
		<li>For adult rats using inhalation anesthesia such as isoflurane, use 4% isoflurane in 2.0 L/min of oxygen for induction. For maintenance throughout the surgery, use 2 - 3% isoflurane in 1.5 L/min of oxygen, varying the isoflurane concentration according to the breathing pattern of the animal to avoid over- or under-dosing.</li>
	</ol>
	</li>
	<li>Once the animal is initially anesthetized, record the preoperative weight of the animal.</li>
	<li>For the cervical DRG injection, shave off the fur on the neck from between the ears to just distal to the scapulae. Disinfect the shaved area with an antiseptic product.</li>
	<li>Inject 2 mL of saline subcutaneously on the flank of the animal, along with an appropriate dose of analgesic (e.g., 5 mg/kg carprofen) and antibiotic (e.g., 5 mg/kg enrofloxacin).</li>
	<li>Apply eye ointment to both the eyes to prevent corneal drying during the procedure.</li>
	<li>Proceed to surgery on a spinal stereotaxic frame with a heating pad at 37 &#176;C placed underneath the animal.</li>
</ol>
4. Injecting DRG and Performing a Dorsal Root Crush Injury
NOTE: This is an extremely delicate surgery. It is advisable to practice on a few dead animals first to familiarize with the anatomy before advancing to live animal surgery.
<ol>
	<li>Locate the prominent C2 and T2 spinous processes over the skin. Make a skin incision between the C2 and T2 spinous processes with a no.10 scalpel blade (once the skin is opened, a white, fibrous-tissue midline should be visible on the first layer of muscle; the first layer of muscle has a &#34;jelly-like&#34; texture).</li>
	<li>Make a similar-sized incision on the first layer of muscle along the white midline using a scalpel blade. Do not go beyond the prominent T2 spinous process,&#160;as there is a major arterial branch from the descending aorta located near there. 
	NOTE: Heavy bleeding can occur anywhere from this step onwards. Always stop the bleeding and remove the blood by using sterilized cotton buds or surgical absorbable sponges to allow clear visualization at all times. Proceeding with the procedure &#34;blindly&#34; may result in unwanted damage to the spinal cord, DRG, or dorsal root, leading to unexpected adverse effects in the animal later on.</li>
	<li>Retract the first layer of muscle using two retractors, one placed rostrally and one caudally; the second layer of muscle, with a striated appearance, should be visible.</li>
	<li>Locate the midline of the second layer of muscle (where two longitudinal muscles can be observed connected by a thin, membranous tissue). Dissect the membranous tissue using a pair of microscissors to separate the two longitudinal muscles. Avoid using a scalpel blade if possible, as this may result in unnecessary muscle damage and bleeding.</li>
	<li>Adjust the retractors accordingly to expose the third layer of thin muscle covering the spine. The spinous processes can be felt by lightly touching with a pair of forceps over the third layer of muscle.</li>
	<li>Make a small incision on the third layer of muscle using microscissors and gently scrape off the muscle from the bone using a curette or scalpel in a sideway manner to clearly expose the vertebrae. If required, cut some of the muscle or tendon away to make the laminectomy easier.</li>
	<li>To expose the left C5 - C8 DRGs, perform a left hemi-laminectomy on the C4 - T1 vertebrae by carefully removing part of the lamina and pedicle using a pair of fine rongeurs. The DRG is located near the transverse foramen of the vertebrae. 
	NOTE: C3 - C7 vertebrae do not have a prominent spinous process. The C5 DRG is located between the C4 and C5 vertebrae, the C6 DRG is between the C5 and C6 vertebrae, and so on, while the C8 DRG is between the C7 and T1 vertebrae. There is no C8 vertebra, despite the presence of a C8 DRG and a C8 spinal cord segment.</li>
	<li>Once enough of the DRG has been exposed for injection, prepare the syringe by placing the virus-filled microliter syringe fitted with a custom-made, 33-gauge blunt needle onto the stereotaxic syringe holder. Before injection, use a 30-gauge beveled needle to make a small superficial opening on each of the targeted DRG to assist with the insertion of the injection needle.</li>
	<li>Insert the 33-gauge needle into the center of the DRG by turning the knobs gently to adjust the stereotaxic coordinates. Do not over-insert the needle, as this may cause fluid to leak out from the ventral side of the DRG. Should leakage occur, adjust the position of the needle immediately. 
	NOTE: The numeric stereotaxic coordinates are not used; however, it is helpful to utilize the frame to hold the needle for injection.</li>
	<li>Inject 1 &#181;L of the virus into each DRG at 0.2 &#181;L/min using an infusion syringe pump. Wait for an additional three minutes before withdrawing the needle. During the injection, the DRG will slowly change color if the virus solution contains a colored dye.</li>
	<li>To perform a concurrent C5 - C8 dorsal root crush injury, crush each root three times for 10 s each using a pair of fine-tipped forceps (Bonn Micro), opposing the ends of the forceps completely; a white line in the tissue should appear at the crush site. Do not go deeper than required with the forceps, as this may cause damage to the ventral root.</li>
	<li>Following the injection and/or crush, make sure that there is no bleeding or small pieces of bone fragments left at the incision site before closing up the animal. If preferred, place a small piece of surgical absorbable sponge on top of the exposed spinal cord and DRG.</li>
	<li>Allow the third layer of the muscle to retract back naturally onto the spine without suturing. Loosely suture the two longitudinal muscles on the second layer (&#8776; 3 interrupted sutures) with absorbable 6-0 suture material. Suture the first layer of muscle (&#8776; 5 interrupted sutures) with absorbable 6-0 suture material.</li>
	<li>Suture the skin with absorbable 5-0 suture material (&#8776; 10 interrupted sutures). 
	NOTE: Make sure that the sutures are not too tight. Bulging of the skin is a sign of over-tightening and may cause discomfort to the animal.</li>
	<li>If heavy bleeding occurred during the surgery, subcutaneously inject 1 - 2 mL of saline to replenish the loss of fluid from the animal as permitted under local regulations.</li>
	<li>Provide edible hydrating gel and allow the animal to recover fully from anesthesia, performing regular monitoring for at least 1 h before returning the animal back to the holding area.</li>
	<li>Keep the animal for at least 3 weeks for optimal transgene expression to assess sensory axon regeneration.</li>
</ol>
5. Performing Post-operative Care of the Animal
<ol>
	<li>Provide food mash and soft cotton bedding for the animal during the first week post-surgery. If required, administer additional doses of analgesic and antibiotic to aid with the recovery in accordance with the local/national regulations and veterinary advice.</li>
	<li>Remove the sutures on the skin after 7 - 10 days. 
	NOTE: Common adverse effects of the surgery include the formation of seroma or hematoma at the incision site, scratch marks on the skin due to the itchiness caused by the internal absorbable sutures, subtle sensory or locomotion deficits, and signs of self-mutilation on the deafferented paw after the dorsal root crush injury. For any welfare issues that are substantially more severe than expected, seek immediate veterinary advice.</li>
</ol>
6. Performing an Anterograde CTB Injection for Axonal Tracing
NOTE: It is recommended to perform cholera toxin B subunit (CTB) axonal tracing a week prior to tissue collection.
<ol>
	<li>Prepare 1% CTB solution as per the manufacturer&#39;s instruction.</li>
	<li>Optional: Add 1% colored dye to the solution for easy visualization of the injection later on. Mix the solution very gently.</li>
	<li>Anesthetize the animal (see step 3.2.1) and stabilize the left forepaw by taping the limb to the table.</li>
	<li>Manually and slowly inject 1 &#181;L of 1% CTB subcutaneously into the center of the glabrous footpad and four digits using a microliter syringe fitted with a custom-made, 33-gauge short needle. 
	NOTE: Before injection, it helps to first use a 30-gauge beveled needle to make a small superficial opening on the skin, which assists with the insertion of the injection needle.</li>
	<li>Allow the animal to recover fully from anesthesia before returning the animal to the holding area.</li>
</ol>
7. Collecting Tissue
<ol>
	<li>To collect the virus-injected DRG and spinal cord for immunohistochemistry, administer an overdose of anesthetic and transcardially perfuse the animal with phosphate-buffered saline followed by cold 4% paraformaldehyde. Carefully perform a full laminectomy on the spine to collect the fixed tissue under a microscope. Proceed with tissue preparation, sectioning, and processing as required for analysis4,5,6. 
	NOTE: The recovered incision site should be apparent, as observed by the presence of scar tissue.</li>
	<li>Alternatively, to collect the virus-injected DRG for in vitro culture, sacrifice the animal humanely with an approved method, such as a rising concentration of carbon dioxide. Carefully dissect the DRG under a microscope, ensuring aseptic conditions whenever possible. Proceed with the tissue culture as desired4,5,6,10.</li>
</ol>

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M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Characterization+of+rat+forepaw+function+in+two+models+of+cervical+dorsal+root+injury.">Characterization of rat forepaw function in two models of cervical dorsal root injury.</a> J Neurotrauma. 26 (1), 17-29 (2009).</li><li>Hocquemiller, M., Giersch, L., Audrain, M., Parker, S., Cartier, N. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Adeno-Associated+Virus-Based+Gene+Therapy+for+CNS+Diseases.">Adeno-Associated Virus-Based Gene Therapy for CNS Diseases.</a> Hum Gene Ther. 27 (7), 478-496 (2016).</li><li>Cheah, M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Expression+of+an+Activated+Integrin+Promotes+Long-Distance+Sensory+Axon+Regeneration+in+the+Spinal+Cord.">Expression of an Activated Integrin Promotes Long-Distance Sensory Axon Regeneration in the Spinal Cord.</a> J Neurosci. 36 (27), 7283-7297 (2016).</li><li>Andrews, M. R., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Alpha9+integrin+promotes+neurite+outgrowth+on+tenascin-C+and+enhances+sensory+axon+regeneration.">Alpha9 integrin promotes neurite outgrowth on tenascin-C and enhances sensory axon regeneration.</a> J Neurosci. 29 (17), 5546-5557 (2009).</li><li>Tan, C. L., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Kindlin-1+enhances+axon+growth+on+inhibitory+chondroitin+sulfate+proteoglycans+and+promotes+sensory+axon+regeneration.">Kindlin-1 enhances axon growth on inhibitory chondroitin sulfate proteoglycans and promotes sensory axon regeneration.</a> J Neurosci. 32 (21), 7325-7335 (2012).</li><li>McCarty, D. M., Young, S. M., Samulski, R. 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F., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Grafted+neural+progenitors+integrate+and+restore+synaptic+connectivity+across+the+injured+spinal+cord.">Grafted neural progenitors integrate and restore synaptic connectivity across the injured spinal cord.</a> J Neurosci. 31 (12), 4675-4686 (2011).</li><li>Wang, R., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Persistent+restoration+of+sensory+function+by+immediate+or+delayed+systemic+artemin+after+dorsal+root+injury.">Persistent restoration of sensory function by immediate or delayed systemic artemin after dorsal root injury.</a> Nat Neurosci. 11 (4), 488-496 (2008).</li><li>Wong, L. E., Gibson, M. E., Arnold, H. M., Pepinsky, B., Frank, E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Artemin+promotes+functional+long-distance+axonal+regeneration+to+the+brainstem+after+dorsal+root+crush.">Artemin promotes functional long-distance axonal regeneration to the brainstem after dorsal root crush.</a> Proc Natl Acad Sci U S A. 112 (19), 6170-6175 (2015).</li><li>Tanguy, Y., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Systemic+AAVrh10+provides+higher+transgene+expression+than+AAV9+in+the+brain+and+the+spinal+cord+of+neonatal+mice.">Systemic AAVrh10 provides higher transgene expression than AAV9 in the brain and the spinal cord of neonatal mice.</a> Front Mol Neurosci. 8, 36 (2015).</li><li>Foust, K. D., Poirier, A., Pacak, C. A., Mandel, R. J., Flotte, T. 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J., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Directed+evolution+of+a+novel+adeno-associated+virus+(AAV)+vector+that+crosses+the+seizure-compromised+blood-brain+barrier+(BBB).">Directed evolution of a novel adeno-associated virus (AAV) vector that crosses the seizure-compromised blood-brain barrier (BBB).</a> Mol Ther. 18 (3), 570-578 (2010).</li><li>Fagoe, N. D., Attwell, C. L., Kouwenhoven, D., Verhaagen, J., Mason, M. 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James Fawcett is a paid consultant for Acorda Therapeutics and Novartis.

In this article, we present a step-by-step guide to perform a DRG injection and dorsal root crush injury in the lower cervical spinal cord of an adult rat. As this is an extremely invasive and delicate surgery, we strongly recommend that all potential users obtain sufficient training and practice before advancing to live animal surgery. The users should be familiar not only with spinal cord anatomy, but also with the surrounding muscle tissues, vertebral bone structure, and vasculature. Ideally, a competent user should b...

Achieving axon regeneration after nervous system injury is a challenging task1. To study the failure of axon regeneration in the central nervous system (CNS), researchers have used a plethora of nerve injury models. As regions of the CNS differ, it is important to use an anatomically appropriate model to study axon regeneration. By using the appropriate model, researchers 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, as opposed to a &#34;one-for-all&#34; treatment strategy.
In spinal cord injury, for example, the most debilitating symptoms stem from the loss of sensation and locomotion. Loss of sensation is caused by damage to the ascending sensory pathways, while the loss of locomotion is caused by damage to the descending motor pathways. Due to cellular and anatomical differences between these two pathways, many targeted axon regeneration studies only focus on one or the other pathway, with the rationale that successful recovery of either would be of tremendous benefit to patients. In this article, we present a protocol that uses a direct dorsal root ganglia (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.
DRG sensory neurons are responsible for relaying sensory information, such as tactile sensation and pain, from the periphery to the CNS. The long axonal projections of sensory neurons in the spinal cord serve as a good model to study long-distance axon regeneration. In addition, as rodents can survive a sensory pathway lesion such as a dorsal root crush injury with minimal welfare complications, researchers can study CNS axon regeneration without the need to completely lesion the spinal cord. A quadruple C5 - C8 (cervical level 5 - 8) dorsal root crush injury has been shown to be a useful model for forepaw deafferentation2. Additionally, a dorsal root crush injury provides a &#34;cleaner&#34; model to study axon regeneration than a direct spinal cord injury because it is uncomplicated by other factors such as glial scar formation.
The use of viral gene therapy to reprogram neurons into a regenerative state has been increasingly regarded as a promising treatment strategy for many neurological conditions3. Studies have shown the application of an adeno-associated virus (AAV) vector carrying the transgene of a growth-promoting protein can achieve robust axon regeneration with behavioral recovery4,5,6. The apparent low pathogenicity of AAV in eliciting an immune response and the ability to transduce non-dividing cells, such as neurons, make it the optimal vector for gene therapy. Additionally, the recombinant AAV form is used for therapy. In this form, it is incapable of integrating its viral genome into the host genome7, reducing the risk of insertional mutagenesis as compared to other viral vectors, such as lentivirus. This makes AAV a safe choice for gene therapy applications.
As a DRG contains the cell bodies of sensory neurons, it is the most appropriate anatomical target for the administration of virus for gene therapy to study and/or promote sensory axon regeneration. In a study comparing different AAV serotypes and lentivirus, AAV serotype 5 (AAV5) was shown to be the most efficient in transducing DRG neurons over a time course of at least 12 weeks when injected directly into the DRG8. Additionally, AAV can achieve more than 40% transduction efficiency, transducing all DRG neuronal subtypes, such as the large-diameter neurofilament 200 kDa (NF200)-positive neurons and the small-diameter calcitonin gene-related peptide (CGRP)- or isolectin b4 (IB4)-positive neurons4,8.
As the surgical procedure of DRG injection and dorsal root crush injury is extremely invasive and delicate, we believe that this article will help new users to learn the procedure in a very efficient manner. In this article, we show representative results from adult rats four weeks after the injection of a control virus AAV5-GFP (green fluorescent protein) into C6 - C7 DRGs with a concurrent C5 - C8 dorsal root crush injury. This model is especially suitable for researchers who are investigating the use of viral gene therapy to promote sensory axon regeneration.

<table><tbody><tr><td>Fast Green FCF dye</td><td>Sigma-Aldrich</td><td>F7258</td><td>For visualizing colorless solution. Recommended concentration: 1%</td></tr><tr><td>Cholera Toxin B subunit</td><td>List Biological Laboratories</td><td>104</td><td>For anterograde axonal tracing. Recommended concentration: 1%</td></tr><tr><td>IsoFlo</td><td>Zoetis</td><td>115095</td><td>Inhalation anesthetic (active ingredient: isoflurane)</td></tr><tr><td>Baytril 2.5% injectable</td><td>Bayer</td><td>05032756093017</td><td>Antibiotic (active ingredient: enrofloxacin). Manufacturer&#039;s recommended dosage: 10 mg/kg</td></tr><tr><td>Carprieve 5.0% w/v</td><td>Norbrook</td><td>02000/4229</td><td>Analgesic (active ingredient: carprofen). Manufacturer&#039;s recommended dosage: 4 mg/kg</td></tr><tr><td>Lacri-Lube</td><td>Allergan</td><td>PL 00426/0041</td><td>Eye ointment</td></tr><tr><td>Olsen-Hegar Needle Holder</td><td>Fine Science Tools</td><td>FST 12502-12</td><td></td></tr><tr><td>Friedman Pearson Rongeur Curved 0.7 mm Cup</td><td>Fine Science Tools</td><td>FST 16121-14</td><td></td></tr><tr><td>Bonn Micro Forceps</td><td>Fine Science Tools</td><td>FST 11083-07</td><td>For performing dorsal root crush injury</td></tr><tr><td>Tissue Separating Scissors</td><td>Fine Science Tools</td><td>FST 14072-10</td><td></td></tr><tr><td>Fine Scissors</td><td>Fine Science Tools</td><td>FST 14058-11</td><td></td></tr><tr><td>Micro-Adson Forceps</td><td>Fine Science Tools</td><td>FST 11018-12</td><td></td></tr><tr><td>Goldstein Retractor</td><td>Fine Science Tools</td><td>FST 17003-03</td><td></td></tr><tr><td>Vannas Spring Scissors (straight)</td><td>Fine Science Tools</td><td>FST 15018-10</td><td></td></tr><tr><td>SURGIFOAM Absorbable Gelatin Sponge</td><td>Ethicon</td><td>1972</td><td>For bleeding control</td></tr><tr><td>Microliter Syringe RN701 (10 &amp;mu;L)</td><td>Hamilton</td><td>80330</td><td></td></tr><tr><td>Custom-made Removable Needle (for DRG injection)</td><td>Hamilton</td><td>7803-05</td><td>33 gauge, 38 mm, point style 3</td></tr><tr><td>Custom-made Removable Needle (for CTB injection)</td><td>Hamilton</td><td>7803-05</td><td>33 gauge, 10 mm, point style 3</td></tr><tr><td>UltraMicroPump with SYS-Micro4 Controller</td><td>World Precision Instruments</td><td>UMP3-1</td><td></td></tr></tbody></table>

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.

As a representation, a transverse spinal cord section with the attached DRG is presented to show the effectiveness of this protocol in transducing DRG neurons and tracing sensory axons in the spinal cord four weeks after injecting a control virus, AAV5-GFP, directly into the C7 DRG without a dorsal root crush injury (Figure 1A). Axons in both the dorsal column and the dorsal horn of the spinal cord express GFP (Figure 1B), as well as the cell bodies and a...

Watch this Scientific Journal Video about Dorsal Root Ganglion Injection and Dorsal Root Crush Injury as a Model for Sensory Axon Regeneration at JoVE.com

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

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 ...

dorsal-root-ganglion-injection-dorsal-root-crush-injury-as-model-for

Research

JoVE Journal

Neuroscience

17.7K Views.  University of Cambridge. 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.

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

Chromatin Immunoprecipitation from Dorsal Root Ganglia Tissue following Axonal Injury

Axons in the central nervous system (CNS) do not regenerate while those in the peripheral nervous system (PNS) do regenerate 
to a limited extent after injury (Teng et al., 2006). It is recognized that transcriptional programs essential for neurite and axonal outgrowth are 
reactivated upon injury in the PNS (Makwana et al., 2005). However the tools available to analyze neuronal gene regulation in vivo are limited and 
often challenging.
The dorsal root ganglia (DRG) offer an excellent injury model system because both the CNS and PNS are innervated by a 
bifurcated axon originating from the same soma. The ganglia represent a discrete collection of cell bodies where all transcriptional events occur, 
and thus provide a clearly defined region of transcriptional activity that can be easily and reproducibly removed from the animal. Injury of nerve 
fibers in the PNS (e.g. sciatic nerve), where axonal regeneration does occur, should reveal a set of transcriptional programs that are distinct from 
those responding to a similar injury in the CNS, where regeneration does not take place (e.g. spinal cord). Sites for transcription factor binding, 
histone and DNA modification resulting from injury to either PNS or CNS can be characterized using chromatin immunoprecipitation (ChIP). 
Here, we describe a ChIP protocol using fixed mouse DRG tissue following axonal injury. This powerful combination provides a means for characterizing the pro-regeneration chromatin environment necessary for promoting axonal regeneration.

We present a method for chromatin immunoprecipitation from dorsal root ganglia tissue following axonal injury. The approach can be used to identify specific transcription factor binding sites and epigenetic modification of histone and DNA important for the regeneration of injured axons in both the peripheral and central nervous system.

Axons in the central nervous system (CNS) do not regenerate while those in the peripheral nervous system (PNS) do regenerate 
to a limited extent after ...

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. ...

Using Microfluidics Chips for Live Imaging and Study of Injury Responses in Drosophila Larvae

Live imaging is an important technique for studying cell biological processes, however this can be challenging in live animals. The translucent cuticle of the Drosophila larva makes it an attractive model organism for live imaging studies. However, an important challenge for live imaging techniques is to noninvasively immobilize and position an animal on the microscope. This protocol presents a simple and easy to use method for immobilizing and imaging Drosophila larvae on a polydimethylsiloxane (PDMS) microfluidic device, which we call the 'larva chip'. The larva chip is comprised of a snug-fitting PDMS microchamber that is attached to a thin glass coverslip, which, upon application of a vacuum via a syringe, immobilizes the animal and brings ventral structures such as the nerve cord, segmental nerves, and body wall muscles, within close proximity to the coverslip. This allows for high-resolution imaging, and importantly, avoids the use of anesthetics and chemicals, which facilitates the study of a broad range of physiological processes. Since larvae recover easily from the immobilization, they can be readily subjected to multiple imaging sessions. This allows for longitudinal studies over time courses ranging from hours to days. This protocol describes step-by-step how to prepare the chip and how to utilize the chip for live imaging of neuronal events in 3rd instar larvae. These events include the rapid transport of organelles in axons, calcium responses to injury, and time-lapse studies of the trafficking of photo-convertible proteins over long distances and time scales. Another application of the chip is to study regenerative and degenerative responses to axonal injury, so the second part of this protocol describes a new and simple procedure for injuring axons within peripheral nerves by a segmental nerve crush.

Using Microfluidics Chips for Live Imaging and Study of Injury Responses in Drosophila Larvae

Drosophila larvae are an attractive model system for live imaging due to their translucent cuticle and powerful genetics. This protocol describes how to utilize a single-layer PDMS device, called the &#39;larva chip&#39; for live imaging of cellular processes within neurons of 3rd instar Drosophila larvae.

Live imaging is an important technique for studying cell biological processes, however this can be challenging in live animals. The translucent cuticle of ...

Bioengineering

Reproducible Mouse Sciatic Nerve Crush and Subsequent Assessment of Regeneration by Whole Mount Muscle Analysis

Regeneration in the peripheral nervous system (PNS) is widely studied both for its relevance to human disease and to understand the robust regenerative response mounted by PNS neurons thereby possibly illuminating the failures of CNS regeneration1. Sciatic nerve crush (axonotmesis) is one of the most common models of peripheral nerve injury in rodents2. Crushing interrupts all axons but Schwann cell basal laminae are preserved so that regeneration is optimal3,4. This allows the investigator to study precisely the ability of a growing axon to interact with both the Schwann cell and basal laminae4. Rats have generally been the preferred animal models for experimental nerve crush. They are widely available and their lesioned sciatic nerve provides a reasonable approximation of human nerve lesions5,4. Though smaller in size than rat nerve, the mouse nerve has many similar qualities. Most importantly though, mouse models are increasingly valuable because of the wide availability of transgenic lines now allows for a detailed dissection of the individual molecules critical for nerve regeneration6, 7. Prior investigators have used multiple methods to produce a nerve crush or injury including simple angled forceps, chilled forceps, hemostatic forceps, vascular clamps, and investigator-designed clamps8,9,10,11,12. Investigators have also used various methods of marking the injury site including suture, carbon particles and fluorescent beads13,14,1. We describe our method to obtain a reproducibly complete sciatic nerve crush with accurate and persistent marking of the crush-site using a fine hemostatic forceps and subsequent carbon crush-site marking. As part of our description of the sciatic nerve crush procedure we have also included a relatively simple method of muscle whole mount we use to subsequently quantify regeneration.

In this report we describe a method to crush mouse sciatic nerve. This method uses readily available hemostatic forceps and easily and reproducibly produces complete sciatic nerve crush. In addition, we describe a method to prepare muscle whole mounts suitable for analysis of nerve regeneration after sciatic nerve crush.

Regeneration in the peripheral nervous system (PNS) is widely studied both for its relevance to human disease and to understand the robust regenerative ...

Genetic Study of Axon Regeneration with Cultured Adult Dorsal Root Ganglion Neurons

It is well known that mature neurons in the central nervous system (CNS) cannot regenerate their axons after injuries due to diminished intrinsic ability to support axon growth and a hostile environment in the mature CNS1,2. In contrast, mature neurons in the peripheral nervous system (PNS) regenerate readily after injuries3. Adult dorsal root ganglion (DRG) neurons are well known to regenerate robustly after peripheral nerve injuries. Each DRG neuron grows one axon from the cell soma, which branches into two axonal branches: a peripheral branch innervating peripheral targets and a central branch extending into the spinal cord. Injury of the DRG peripheral axons results in substantial axon regeneration, whereas central axons in the spinal cord regenerate poorly after the injury. However, if the peripheral axonal injury occurs prior to the spinal cord injury (a process called the conditioning lesion), regeneration of central axons is greatly improved4. Moreover, the central axons of DRG neurons share the same hostile environment as descending corticospinal axons in the spinal cord. Together, it is hypothesized that the molecular mechanisms controlling axon regeneration of adult DRG neurons can be harnessed to enhance CNS axon regeneration. As a result, adult DRG neurons are now widely used as a model system to study regenerative axon growth5-7.
Here we describe a method of adult DRG neuron culture that can be used for genetic study of axon regeneration in vitro. In this model adult DRG neurons are genetically manipulated via electroporation-mediated gene transfection6,8. By transfecting neurons with DNA plasmid or si/shRNA, this approach enables both gain- and loss-of-function experiments to investigate the role of any gene-of-interest in axon growth from adult DRG neurons. When neurons are transfected with si/shRNA, the targeted endogenous protein is usually depleted after 3-4 days in culture, during which time robust axon growth has already occurred, making the loss-of-function studies less effective. To solve this problem, the method described here includes a re-suspension and re-plating step after transfection, which allows axons to re-grow from neurons in the absence of the targeted protein. Finally, we provide an example of using this in vitro model to study the role of an axon regeneration-associated gene, c-Jun, in mediating axon growth from adult DRG neurons9.

An in vitro model for genetic study of axon regeneration using cultured adult mouse dorsal root ganglion neurons is described. The method includes a re-suspension/re-plating step to allow axon re-growth from neurons undergoing genetic manipulation. This approach is especially useful for loss-of-function studies of axon regeneration using RNAi-based protein knockdown.

It is well known that mature neurons in the central nervous system (CNS) cannot regenerate their axons after injuries due to diminished intrinsic ability ...

Combining Peripheral Nerve Grafting and Matrix Modulation to Repair the Injured Rat Spinal Cord

Traumatic injury to the spinal cord (SCI) causes death of neurons, disruption of motor and sensory nerve fiber (axon) pathways and disruption of communication with the brain. One of the goals of our research is to promote axon regeneration to restore connectivity across the lesion site. To accomplish this we developed a peripheral nerve (PN) grafting technique where segments of sciatic nerve are either placed directly between the damaged ends of the spinal cord or are used to form a bridge across the lesion. There are several advantages to this approach compared to transplantation of other neural tissues; regenerating axons can be directed towards a specific target area, the number and source of regenerating axons is easily determined by tracing techniques, the graft can be used for electrophysiological experiments to measure functional recovery associated with axons in the graft, and it is possible to use an autologous nerve to reduce the possibility of graft rejection. In our lab we have performed both autologous (donor and recipient are the same animal) and heterologous (donor and recipient are different animals) grafts with comparable results. This approach has been used successfully in both acute and chronic injury situations. Regenerated axons that reach the distal end of the PN graft often fail to extend back into the spinal cord, so we use microinjections of chondroitinase to degrade inhibitory molecules associated with the scar tissue surrounding the area of SCI. At the same time we have found that providing exogenous growth and trophic molecules encourages longer distance axonal regrowth into the spinal cord. Several months after transplantation we perform a variety of anatomical, behavioral and electrophysiological tests to evaluate the recovery of function in our spinal cord injured animals. This experimental approach has been used successfully in several spinal cord injury models, at different levels of injury and in different species (mouse, rat and cat). Importantly, the peripheral nerve grafting approach is effective in promoting regeneration by acute and chronically injured neurons.

Traumatic injury to the spinal cord disrupts communication with the brain. To restore lost connectivity we utilize a peripheral nerve graft to provide a substratum for regenerating fibers in combination with neurotrophic factors and matrix-modulating enzymes to remove inhibitory molecules to promote long distance growth.

Traumatic injury to the spinal cord (SCI) causes death of neurons, disruption of motor and sensory nerve fiber (axon) pathways and disruption of ...

Biology

Dorsal Root Ganglia Neurons and Differentiated Adipose-derived Stem Cells: An In Vitro Co-culture Model to Study Peripheral Nerve Regeneration

Dorsal root ganglia (DRG) neurons, located in the intervertebral foramina of the spinal column, can be used to create an in vitro system facilitating the study of nerve regeneration and myelination. The glial cells of the peripheral nervous system, Schwann cells (SC), are key facilitators of these processes; it is therefore crucial that the interactions of these cellular components are studied together. Direct contact between DRG neurons and glial cells provides additional stimuli sensed by specific membrane receptors, further improving the neuronal response. SC release growth factors and proteins in the culture medium, which enhance neuron survival and stimulate neurite sprouting and extension. However, SC require long proliferation time to be used for tissue engineering applications and the sacrifice of an healthy nerve for their sourcing. Adipose-derived stem cells (ASC) differentiated into SC phenotype are a valid alternative to SC for the set-up of a co-culture model with DRG neurons to study nerve regeneration. The present work presents a detailed and reproducible step-by-step protocol to harvest both DRG neurons and ASC from adult rats; to differentiate ASC towards a SC phenotype; and combines the two cell types in a direct co-culture system to investigate the interplay between neurons and SC in the peripheral nervous system. This tool has great potential in the optimization of tissue-engineered constructs for peripheral nerve repair.

Dorsal Root Ganglia Neurons and Differentiated Adipose-derived Stem Cells: An In Vitro Co-culture Model to Study Peripheral Nerve Regeneration

Dorsal root ganglia (DRG) are structures containing the sensory neurons of the peripheral nervous system. When dissociated, they can be co-cultured with SC-like adipose-derived stem cells (ASC), providing a valuable model to study in vitro nerve regeneration and myelination, mimicking the in vivo environment at the injury site.

Dorsal root ganglia (DRG) neurons, located in the intervertebral foramina of the spinal column, can be used to create an in vitro system facilitating the ...

Investigating Mammalian Axon Regeneration: In Vivo Electroporation of Adult Mouse Dorsal Root Ganglion

Electroporation is an essential non-viral gene transfection approach to introduce DNA plasmids or small RNA molecules into cells. A sensory neuron in the dorsal root ganglion (DRGs) extends a single axon with two branches. One branch goes to the peripheral nerve (peripheral branch), and the other branch enters the spinal cord through the dorsal root (central branch). After the neural injury, the peripheral branch regenerates robustly whereas the central branch does not regenerate. Due to the high regenerative capacity, sensory axon regeneration has been widely used as a model system to study mammalian axon regeneration in both the peripheral nervous system (PNS) and the central nervous system (CNS). Here, we describe a previously established approach protocol to manipulate gene expression in mature sensory neurons in vivo via electroporation. Based on transfection with plasmids or small RNA oligos (siRNAs or microRNAs), the approach allows for both loss- and gain-of-function experiments to study the roles of genes-of-interests or microRNAs in regulation of axon regeneration in vivo. In addition, the manipulation of gene expression in vivo can be controlled both spatially and temporally within a relatively short time course. This model system provides a unique tool to investigate the molecular mechanisms by which mammalian axon regeneration is regulated in vivo.

Investigating Mammalian Axon Regeneration: In Vivo Electroporation of Adult Mouse Dorsal Root Ganglion

Electroporation is an effective approach to deliver genes of interests into cells. By applying this approach in vivo on the neurons of adult mouse dorsal root ganglion (DRG), we describe a model to study axon regeneration in vivo.

Electroporation is an essential non-viral gene transfection approach to introduce DNA plasmids or small RNA molecules into cells. A sensory neuron in the ...

A Drosophila In Vivo Injury Model for Studying Neuroregeneration in the Peripheral and Central Nervous System

The regrowth capacity of damaged neurons governs neuroregeneration and functional recovery after nervous system trauma. Over the past few decades, various intrinsic and extrinsic inhibitory factors involved in the restriction of axon regeneration have been identified. However, simply removing these inhibitory cues is insufficient for successful regeneration, indicating the existence of additional regulatory machinery. Drosophila melanogaster, the fruit fly, shares evolutionarily conserved genes and signaling pathways with vertebrates, including humans. Combining the powerful genetic toolbox of flies with two-photon laser axotomy/dendriotomy, we describe here the Drosophila sensory neuron &#8211; dendritic arborization (da) neuron injury model as a platform for systematically screening for novel regeneration regulators. Briefly, this paradigm includes a) the preparation of larvae, b) lesion induction to dendrite(s) or axon(s) using a two-photon laser, c) live confocal imaging post-injury and d) data analysis. Our model enables highly reproducible injury of single labeled neurons, axons, and dendrites of well-defined neuronal subtypes, in both the peripheral and central nervous system.

A Drosophila In Vivo Injury Model for Studying Neuroregeneration in the Peripheral and Central Nervous System

Here, we present a protocol using the Drosophila sensory neuron - dendritic arborization (da) neuron injury model, which combines in vivo live imaging, two-photon laser axotomy/dendriotomy, and the powerful fly genetic toolbox, as a platform for screening potential promoters and inhibitors of neuroregeneration.

The regrowth capacity of damaged neurons governs neuroregeneration and functional recovery after nervous system trauma. Over the past few decades, various ...

Medicine

Sheath-Preserving Optic Nerve Transection in Rats to Assess Axon Regeneration and Interventions Targeting the Retinal Ganglion Cell Axon

Retinal ganglion cell (RGC) axons converge at the optic nerve head to convey visual information from the retina to the brain. Pathologies such as glaucoma, trauma, and ischemic optic neuropathies injure RGC axons, disrupt transmission of visual stimuli, and cause vision loss. Animal models simulating RGC axon injury include optic nerve crush and transection paradigms. Each of these models has inherent advantages and disadvantages. An optic nerve crush is generally less severe than a transection and can be used to assay axon regeneration across the lesion site. However, differences in crush force and duration can affect tissue responses, resulting in variable reproducibility and lesion completeness. With optic nerve transection, there is a severe and reproducible injury that completely lesions all axons. However, transecting the optic nerve dramatically alters the blood brain barrier by violating the optic nerve sheath, exposing the optic nerve to the peripheral environment. Moreover, regeneration beyond a transection site cannot be assessed without reapposing the cut nerve ends. Furthermore, distinct degenerative changes and cellular pathways are activated by either a crush or transection injury.
The method described here incorporates the advantages of both optic nerve crush and transection models while mitigating the disadvantages. Hydrostatic pressure delivered into the optic nerve by microinjection completely transects the optic nerve while maintaining the integrity of the optic nerve sheath. The transected optic nerve ends are reapposed to allow for axon regeneration assays. A potential limitation of this method is the inability to visualize the complete transection, a potential source of variability. However, visual confirmation that the visible portion of the optic nerve has been transected is indicative of a complete optic nerve transection with 90-95% success. This method could be applied to assess axon regeneration promoting strategies in a transection model or investigate interventions that target the axonal compartments.

This protocol describes an optic nerve transection that preserves the optic nerve sheath in rats. Hydrostatic pressure from microinjections into the optic nerve generate a complete transection, allowing for sutureless reapposition of the transected optic nerve ends and direct targeting of the axonal compartment in a transection model.

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

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Chapters in this video

Chromatin Immunoprecipitation from Dorsal Root Ganglia Tissue following Axonal Injury

Live Imaging of Dorsal Root Axons after Rhizotomy

Using Microfluidics Chips for Live Imaging and Study of Injury Responses in Drosophila Larvae

Reproducible Mouse Sciatic Nerve Crush and Subsequent Assessment of Regeneration by Whole Mount Muscle Analysis

Genetic Study of Axon Regeneration with Cultured Adult Dorsal Root Ganglion Neurons

Combining Peripheral Nerve Grafting and Matrix Modulation to Repair the Injured Rat Spinal Cord

Dorsal Root Ganglia Neurons and Differentiated Adipose-derived Stem Cells: An In Vitro Co-culture Model to Study Peripheral Nerve Regeneration

Investigating Mammalian Axon Regeneration: In Vivo Electroporation of Adult Mouse Dorsal Root Ganglion

A Drosophila In Vivo Injury Model for Studying Neuroregeneration in the Peripheral and Central Nervous System

Sheath-Preserving Optic Nerve Transection in Rats to Assess Axon Regeneration and Interventions Targeting the Retinal Ganglion Cell Axon