This work was funded by research grants from: PG Slater: FONDECYT N&#176; 3190820; J. Larra&#237;n: FONDECYT N&#176; 1180429, CARE Chile UC-Centro de Envejecimiento y Regeneraci&#243;n (PFB 12/2007).

This protocol provides enough information to successfully perform spinal cord transection. Of note, there are excellent detailed protocols of these techniques published elsewhere14, which can complement the one presented here. All animal procedures have been approved by the Committee on Bioethics and Biosafety from the Faculty of Biological Sciences, Pontificia Universidad Cat&#243;lica de Chile.
1. Natural mating of frogs
<ol>
	<li>Three to five days prior to mating, subcutaneously preinject male and female frogs with 50 units of human chorionic gonadotropin (hCG). Use the &#34;iron claw&#34; technique for restraining the frogs; as the frogs are slippery, use a net to surround the frog if necessary. Insert the tip of a 26 G x &#189;&#34; needle posterior to the lateral line, pushing it dorsally to a depth of 1 cm, between the skin and the muscle.</li>
	<li>Before mating, inject the male with 300 units and the female with 700 units of hCG.</li>
	<li>For mating to occur, place the male and the female in 2 L of 0.1x Barth solution immediately after chilling the solution at 4 &#176;C for 15 min to resemble spring conditions and leave overnight at 18 &#176;C.</li>
	<li>Sixteen hours later, carefully collect the embryos with the help of a plastic Pasteur pipette, with the tip cut off, and place them in 10 cm diameter Petri dishes. Remove the embryonic jelly coat by incubating the embryos with 25 mL of 2% cysteine in distilled water (pH 7.8; ensure the solution covers the embryos) for 5 min with slight agitation. Wash 3 times with distilled water and 3 times with 0.1x Barth solution (8.9 mM NaCl; 102 &#181;M KCl; 238.1 &#181;M NaHCO3; 1 mM 4-(2- hydroxyethyl)-1-piperazine ethanesulfonic acid (HEPES); 81.14 &#181;M MgSO4; 33.88 &#181;M Ca(NO3)2; 40.81 &#181;M CaCl2, pH 7.6).</li>
	<li>Select healthy embryos that have a brownish color and symmetrically dividing blastomeres. Place the embryos in 10 cm diameter Petri dishes with 50 mL of 0.1x Barth solution at a density of no more than 100 embryos per dish.</li>
</ol>
2. Husbandry
<ol>
	<li>During the first week, maintain the embryos at 18 &#176;C until they get off the vitelline sac. During this time, change the Barth solution every day, and remove whitish dead embryos and tadpoles presenting any visible anatomical alteration or tadpoles without any swimming movement.</li>
	<li>After the first week, transfer tadpoles to chlorine-free water in plastic tanks at a density of 10 animals per liter. Grow tadpoles at 20-21 &#176;C with a 12-h light/12-h dark cycle, with oxygen stones available in each tank to aerate the water and fed once a day with 0.5 mg per animal. Replace water once a week and check for accumulated waste and dead animals daily24.</li>
</ol>
3. Staging
<ol>
	<li>Three to four weeks after fertilization, place the animals in a Petri dish; then, one by one, check the morphology and appearance of forelimbs and hindlimbs. If necessary, anesthetize the animals by placing the animals in a Petri dish with 50 mL of 0.02% tricaine mesylate in 0.1x Barth solution for better manipulation. After no more than 2 min, place the animals in 0.1x Barth solution for recovery from the anesthesia.</li>
	<li>Look for the following anatomical characteristics of stage 50 animals25: forelimbs that are just appearing and are spherical (Figure 1); hindlimbs that are protruding and are spherical (Figure 1). 
	NOTE: Animals from stages 49 to 51 can be used for this procedure (Figure 1); for more information about stages, refer to Nieuwkoop and Faber&#39;s Normal Table of Xenopus laevis25.</li>
</ol>
4. Surgery: spinal cord transection and sham-operated animals
<ol>
	<li>Anesthetize stage 50 tadpoles by placing them in a Petri dish with 50 mL of 0.02% tricaine mesylate in 0.1x Barth solution for 2 min.</li>
	<li>With the help of a tablespoon and forceps, place the tadpole, dorsal side-up, on a wet piece of gauze in the upper half of a glass Petri dish.</li>
	<li>Perform an incision of the skin and dorsal muscles at the mid-thoracic level (Figure 2A, B) using microdissection spring scissors.
	<ol>
		<li>For control sham animals, ensure that the incision size is only ~0.2 mm (Figure 2C); do not damage the spinal cord (Figure 2D,D&#180;).</li>
		<li>For transected animals, perform a second incision of ~0.2 mm (Figure 2C) to fully transect the spinal cord (Figure 2E,E&#180;).</li>
	</ol>
	</li>
</ol>
5. Postsurgery care
<ol>
	<li>After surgery, transfer the tadpoles to a tank containing 0.5 L of 0.1x Barth solution with 1x Penicillin-streptomycin, at a density of 10-12 animals per tank. Maintain the transected and control sham animals in separate tanks. 
	NOTE: Tadpoles will recover from the anesthesia in a couple of minutes.</li>
	<li>Maintain the tadpoles with aeration at a temperature of 20-21 &#176;C.</li>
	<li>Change the Barth solution with antibiotics every other day until the end of the experiment.</li>
	<li>Start feeding the animals one day after surgery, once per day.</li>
	<li>Eliminate dead animals.</li>
</ol>
6. Swimming assay
<ol>
	<li>Obtain a box with LED illumination from the inside, covered with a transparent polystyrene sheet, which allows the light to pass through.</li>
	<li>Install a camera over the LED box.</li>
	<li>Place a 15 cm diameter Petri dish on top of the box, filled with 100 mL of 0.1x Barth solution.</li>
	<li>One day post transection, place a tadpole in the Petri dish and leave for a 5 min adaptation period.</li>
	<li>After adaptation, start video-tracking the free-swimming behavior using the referenced software (see the Table of Materials) for 5 min.</li>
	<li>After the video is completed, transfer the tadpole back to its tank.</li>
	<li>Repeat the video tracking 5, 10, 15, and 20 days post transection (Figure 3).</li>
</ol>
7. Bioethical considerations
NOTE: The mortality of animals after sham surgery and transection is 13% and 30%, respectively. Additionally, a minimum of 15-20 animals per group is necessary for statistical analysis. Therefore, start with 23 sham and 26 transected animals.
<ol>
	<li>Anesthetize the animals with 0.02% tricaine mesylate for 2 min to assure reduction in neuronal and motor activity and pain before surgery.</li>
	<li>After surgery, check the animals for recovery from anesthesia. Additionally, feed and check the animals daily.</li>
	<li>After finishing the swimming assay, sacrifice the animals with an overdose of tricaine mesylate (1% tricaine mesylate prepared in 30 mM sodium bicarbonate solution).</li>
</ol>

<ol>
	<li>International perspectives on spinal cord injury. World Health Organization Available from: <a target="_blank" href="https://www.who.int/publications/I/item/international-perspectives-on-spinal-cord-injury">https://www.who.int/publications/I/item/international-perspectives-on-spinal-cord-injury</a> (2013)</li><li>Quiroz, J. F. D., Echeverri, K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Spinal+cord+regeneration:+Where+fish,+frogs+and+salamanders+lead+the+way,+can+we+follow.">Spinal cord regeneration: Where fish, frogs and salamanders lead the way, can we follow.</a> Biochemical Journal. 451 (3), 353-364 (2013).</li><li>Lee-Liu, D., M&#233;ndez-Olivos, E. E., Mu&#241;oz, R., Larra&#237;n, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+African+clawed+frog+Xenopus+laevis:+A+model+organism+to+study+regeneration+of+the+central+nervous+system.">The African clawed frog Xenopus laevis: A model organism to study regeneration of the central nervous system.</a> Neuroscience Letters. 652, 82-93 (2017).</li><li>Phipps, L. S., Marshall, L., Dorey, K., Amaya, E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Model+systems+for+regeneration:+Xenopus.">Model systems for regeneration: Xenopus.</a> Development. 147 (6), (2020).</li><li>Lee-Liu, D., Edwards-Faret, G., Tapia, V. S., Larra&#237;n, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Spinal+cord+regeneration:+Lessons+for+mammals+from+non-mammalian+vertebrates.">Spinal cord regeneration: Lessons for mammals from non-mammalian vertebrates.</a> Genesis. 51 (8), 529-544 (2013).</li><li>Beck, C. W., Christen, B., Slack, J. M. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Molecular+pathways+needed+for+regeneration+of+spinal+cord+and+muscle+in+a+vertebrate.">Molecular pathways needed for regeneration of spinal cord and muscle in a vertebrate.</a> Developmental Cell. 5 (3), 429-439 (2003).</li><li>Love, N. 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=Genome-wide+analysis+of+gene+expression+during+Xenopus+tropicalis+tadpole+tail+regeneration.">Genome-wide analysis of gene expression during Xenopus tropicalis tadpole tail regeneration.</a> BMC Developmental Biology. 11, 70 (2011).</li><li>Love, N. 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=Amputation-induced+reactive+oxygen+species+are+required+for+successful+Xenopus+tadpole+tail+regeneration.">Amputation-induced reactive oxygen species are required for successful Xenopus tadpole tail regeneration.</a> Nature Cell Biology. 15 (2), 222-228 (2013).</li><li>Gargiolo, C., Slack, J. M. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cell+lineage+tracing+during+Xenopus+tail+regeneration.">Cell lineage tracing during Xenopus tail regeneration.</a> Development. 131 (11), 2669-2679 (2004).</li><li>Lin, G., Chen, Y., Slack, J. M. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Regeneration+of+neural+crest+derivatives+in+the+Xenopus+tadpole+tail.">Regeneration of neural crest derivatives in the Xenopus tadpole tail.</a> BMC Developmental Biology. 7, 56 (2007).</li><li>Filoni, S., Bosco, L., Cioni, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Reconstitution+of+the+spinal+cord+after+ablation+in+larval+Xenopus+laevistle.">Reconstitution of the spinal cord after ablation in larval Xenopus laevistle.</a> Acta Embryologiae et Morphologiae Experimentalis. 5 (2), 109-129 (1984).</li><li>Gaete, 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=Spinal+cord+regeneration+in+Xenopus+tadpoles+proceeds+through+activation+of+Sox2-positive+cells.">Spinal cord regeneration in Xenopus tadpoles proceeds through activation of Sox2-positive cells.</a> Neural Development. 7, 13 (2012).</li><li>Mu&#241;oz, 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=Regeneration+of+Xenopus+laevis+spinal+cord+requires+Sox2/3+expressing+cells.">Regeneration of Xenopus laevis spinal cord requires Sox2/3 expressing cells.</a> Developmental Biology. 408 (2), 229-243 (2015).</li><li>Edwards-Faret, G., 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=Spinal+cord+regeneration+in+Xenopus+laevis.">Spinal cord regeneration in Xenopus laevis.</a> Nature Protocols. 12 (2), 372-389 (2017).</li><li>M&#233;ndez-Olivos, E. E., Larra&#237;n, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cell+transplantation+as+a+method+to+investigate+spinal+cord+regeneration+in+regenerative+and+nonregenerative+xenopus+stages.">Cell transplantation as a method to investigate spinal cord regeneration in regenerative and nonregenerative xenopus stages.</a> Cold Spring Harbor Protocols. 2018 (12), 943-947 (2018).</li><li>M&#233;ndez-Olivos, E. E., Mu&#241;oz, R., Larra&#237;n, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Spinal+cord+cells+from+pre-metamorphic+stages+differentiate+into+neurons+and+promote+axon+growth+and+regeneration+after+transplantation+into+the+injured+spinal+cord+of+non-regenerative+Xenopus+laevis+froglets.">Spinal cord cells from pre-metamorphic stages differentiate into neurons and promote axon growth and regeneration after transplantation into the injured spinal cord of non-regenerative Xenopus laevis froglets.</a> Frontiers in Cellular Neuroscience. 11, 398 (2017).</li><li>de Vidts, S., M&#233;ndez-Olivos, E., Palacios, M., Larra&#305;n, J., Mery, D. <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+spinal+cord+damage+based+on+automatic+video+analysis+of+froglet+swimming.">Characterization of spinal cord damage based on automatic video analysis of froglet swimming.</a> Biology Open. 8 (12), 2-11 (2019).</li><li>Slater, P. G., Palacios, M., Larra&#237;n, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Xenopus,+a+model+to+study+wound+healing+and+regeneration:+Experimental+approaches.">Xenopus, a model to study wound healing and regeneration: Experimental approaches.</a> Cold Spring Harbor Protocols. 2021 (8), 100966 (2021).</li><li>Lee-Liu, D., 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=Genome-wide+expression+profile+of+the+response+to+spinal+cord+injury+in+Xenopus+laevis+reveals+extensive+differences+between+regenerative+and+non-regenerative+stages.">Genome-wide expression profile of the response to spinal cord injury in Xenopus laevis reveals extensive differences between regenerative and non-regenerative stages.</a> Neural Development. 9, 12 (2014).</li><li>Lee-Liu, D., Sun, L., Dovichi, N. J., Larra&#237;n, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Quantitative+proteomics+after+spinal+cord+injury+(SCI)+in+a+regenerative+and+a+nonregenerative+stage+in+the+frog+Xenopus+laevis.">Quantitative proteomics after spinal cord injury (SCI) in a regenerative and a nonregenerative stage in the frog Xenopus laevis.</a> Molecular and Cellular Proteomics. 17 (4), 592-606 (2018).</li><li>Pe&#241;ailillo, 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=Analysis+of+the+early+response+to+spinal+cord+injury+identi+fi+ed+a+key+role+for+mTORC1+signaling+in+the+activation+of+neural+stem+progenitor+cells.">Analysis of the early response to spinal cord injury identi fi ed a key role for mTORC1 signaling in the activation of neural stem progenitor cells.</a> NPJ Regenerative Medicine. 6 (1), 68 (2021).</li><li>Edwards-Faret, G., 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=Cellular+response+to+spinal+cord+injury+in+regenerative+and+non-regenerative+stages+in+Xenopus+laevis.">Cellular response to spinal cord injury in regenerative and non-regenerative stages in Xenopus laevis.</a> Neural Development. 16 (1), 2 (2021).</li><li>Tapia, V. S., Herrera-Rojas, M., Larrain, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=JAK-STAT+pathway+activation+in+response+to+spinal+cord+injury+in+regenerative+and+non-regenerative+stages+of+Xenopus+laevis.">JAK-STAT pathway activation in response to spinal cord injury in regenerative and non-regenerative stages of Xenopus laevis.</a> Regeneration. 4 (1), 21-35 (2017).</li><li>Ishibashi, S., Amaya, E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=How+to+grow+Xenopus+laevis+tadpole+stages+to+adult.">How to grow Xenopus laevis tadpole stages to adult.</a> Cold Spring Harbor Protocols. 2021 (3), (2021).</li><li>Nieuwkoop, P. D., Faber, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Normal table of Xenopus laevis (Daudin).: A systematical and chronological survey of the development from the fertilized egg till the end of metamorphosis. , (1994).</li><li>Williams, M. C., Patel, J. H., Kakebeen, A. D., Wills, A. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Nutrient+availability+contributes+to+a+graded+refractory+period+for+regeneration+in+Xenopus+tropicalis.">Nutrient availability contributes to a graded refractory period for regeneration in Xenopus tropicalis.</a> Developmental Biology. 473, 59-70 (2021).</li><li>Vleminckx, K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Xenopus: Methods and protocols. , (2018).</li><li>Sive, H. L., Grainger, R. M., Harland, R. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Early development of Xenopus laevis: A laboratory manual. , (2000).</li></ol>

The authors have no conflicts of interest to declare.

The protocol described herein is an excellent method to perform SCI and evaluate functional recovery. For reproducibility, it is essential to grow healthy tadpoles and choose animals that are similar in size. Lack of proper feeding generates nutrient stress, which results in poor regenerative capacities26; therefore, special attention should be paid to tadpole feeding. As tadpoles reach stage 50 after 3-4 weeks, they can be reared at higher temperatures to accelerate the growth process, 18-25 &#176;C being optimal27. Water quality is important, as animals are sensitive to water conditions and chemical products. The optimal water conditions include using carbon filtered, chlorine-free water with the following parameters: pH (6.5-7.5), chloride (&#60;0.02 mg/L), conductivity of water (1.0 mS/cm &#177; 0.1 units), copper (&#60;0.3 mg/L); carbonate hardness (KH: 5-10 dKH); general hardness (GH: 6-16 dGH); nitrate (NO3: &#60;20 mg/L); and nitrite (NO2: &#60;0.1 mg/L)14,27,28. Additionally, to avoid contamination, plastic tanks should be cleaned once a week for rearing animals or every other day after surgery by washing thoroughly with chloride-free water and a sponge; detergent must be avoided.
For a better survival rate after surgery, tadpoles must not be exposed to anesthesia for long periods (no longer than 2 min). Moreover, it is recommended to anesthetize one tadpole at a time. As the animals need to stay hydrated, keep the animals immersed in solution all the time before and after surgery, and pour the solution with a spoon on top of the tadpole before beginning the surgery. Ensure that the damage is extensive enough to cover the whole spinal cord but not too extensive as it can induce poor functional recovery or death. If the notochord is damaged, the animal will be bent, and the functional recovery will be affected. If the damage extends beyond the notochord, the probability of death increases14. During the swimming assay, recording is considered correct if the software identifies each animal with a blue shadow; otherwise, the recording should be repeated. It is important to avoid movement and air or light changes during the recording process to prevent recording mistakes.
There are still many open questions about the cellular and molecular mechanisms underlying spinal cord damage and regeneration. The protocol described in this work can be used to study the contribution of different cellular events, gene expression, and treatments on functional recovery, determined by measuring swimming capacities. Additionally, many other techniques can be applied to the operated animals. The spinal cord can be isolated to perform protein and/or mRNA extraction14 to study protein and gene expression profiles after damage and treatment19,20. This surgery has also been the basis for studying the spinal cord cellular response22 and the behavior of neural stem progenitor cells12,13,22 after spinal cord injury. Signaling cascades involved in spinal cord regeneration have also been studied using the spinal cord damage paradigm described herein23. In summary, the protocol described here is an excellent model to study spinal cord injury and regeneration and has been used for many studies that have contributed to the existing knowledge about the subject.

Spinal cord injury (SCI) is an affliction that affects approximately 250,000-500,000 people worldwide every year1. In addition to this high prevalence, SCI affects sensory and motor nerves, generating paralysis beneath the injury site and disconnection of some internal organs from the control of the CNS. The spinal cord, a part of the CNS, cannot regenerate, and due to the complexity of the affliction and the lack of complete understanding of all the involved processes, there are still no efficient therapies allowing functional recovery.
Non-mammalian organisms, such as teleost fishes, urodele amphibians, and larval stages of anuran amphibians, which can regenerate the spinal cord after severe SCI2,3,4, are excellent model organisms for studying the processes that govern a successful regenerative event and understanding the failure of mammalian regeneration. This understanding is of great interest as it could provide original insights to develop new therapeutic targets and possible therapies for SCI.
The anuran frog, Xenopus laevis, is an excellent model organism to study SCI. It has excellent regenerative capacities during the tadpole stages, which are progressively lost during metamorphosis, allowing experimentation in the regenerative and nonregenerative stages3,5. The established injury method for studying SCI in Xenopus laevis tadpoles consists of tail amputation, where the entire tail is removed, including tissues such as muscle, notochord, and spinal cord6. This approach has been instrumental in the understanding of general mechanisms of regenerative processes4,7,8,9,10.
As tail amputation involves multiple tissues in addition to the spinal cord, which is different from what happens after human SCI, a more relevant injury paradigm is needed for the study of SCI. We have relied on studies used in the past11 for generating comprehensive descriptions of injury paradigms5,12,13,14 and different methods for the study of SCI12,13,14,15,16,17,18. After spinal cord transection, the caudal portion of the spinal cord can be isolated for RNA and protein expression and high-throughput analyses14,19,20,21. Additionally, intracelomic injections of drugs and small molecules, as well as electroporation of cDNA, RNA, or morpholinos, before or after spinal cord transection, allow the study of the effects of these molecules in the prevention or treatment of SCI or of specific events occurring after SCI and spinal cord regeneration13,14. Further, injury evolution and the regenerative processes can be studied at different timings after injury using biochemical, molecular, histological, and functional approaches12,13,14,17,19,20,21,22,23.
Finally, all the aforementioned techniques can be used in non-regenerative stages, highlighting one of the most important advantages of using Xenopus laevis as a model organism to study SCI, the comparative studies of regenerative and non-regenerative mechanisms in the same species13,19,20,21,22. This paper presents a protocol for Xenopus laevis tadpole spinal cord transection, starting with the staging and selection of regenerative Nieuwkoop and Faber (NF) stage 50 tadpoles. This is followed by the description of the procedures for spinal cord surgery to produce sham and transected animals, postsurgical care, and finally the analysis of functional recovery by the measurement of free tadpole swimming distance.

<table><tbody><tr><td>Air pump</td><td>Regent CALM</td><td>RC-006</td><td>For oxygen diffuser stones function</td></tr><tr><td>ANY-maze software</td><td>Stoelting</td><td></td><td>Swimming behavior test</td></tr><tr><td>Ca(NO&lt;sub&gt;3&lt;/sub&gt;)&lt;sub&gt;2&lt;/sub&gt;&amp;middot;4H&lt;sub&gt;2&lt;/sub&gt;O</td><td>Sigma-Aldrich</td><td>237124</td><td></td></tr><tr><td>CaCl&lt;sub&gt;2&lt;/sub&gt;&amp;middot;2H&lt;sub&gt;2&lt;/sub&gt;O</td><td>Sigma-Aldrich</td><td>223506</td><td></td></tr><tr><td>Camera</td><td>Stoelting</td><td>60528</td><td>Swimming behavior test</td></tr><tr><td>Computer</td><td></td><td></td><td>Swimming behavior test (minimum recommended specifications: PC, Windows 7, Intel Core i3, 2 GB RAM, 10-GB drive disk,&lt;br/&gt; 1 available USB port, 1,366 &amp;times; 768 monitor)</td></tr><tr><td>Cysteine</td><td>Sigma-Aldrich</td><td>C7352</td><td></td></tr><tr><td>Dissecting stereomicroscope</td><td>Nikon</td><td>SMZ745T</td><td>Surgery / staging</td></tr><tr><td>Glass Petri dishes</td><td></td><td></td><td>100 x 20 mm</td></tr><tr><td>HEPES</td><td>Gibco</td><td>11344-041</td><td></td></tr><tr><td>Human chorionic gonadotropin</td><td></td><td></td><td>It can be found in different formats in the pharmacy</td></tr><tr><td>KCl</td><td>Merck Millipore</td><td>104936</td><td></td></tr><tr><td>LED light box</td><td>custom made</td><td></td><td>wood box: 55-cm length, 34-cm width, 9-cm height, LED lights, transparent polystyrene sheet)</td></tr><tr><td>MgSO&lt;sub&gt;4&lt;/sub&gt;&amp;middot;7H&lt;sub&gt;2&lt;/sub&gt;O</td><td>Merck Millipore</td><td>105886</td><td></td></tr><tr><td>Microdissection scissors for transection</td><td>Fine Science Tools</td><td>15003-08</td><td>Spring Scissors for surgery</td></tr><tr><td>MS-222</td><td>Sigma-Aldrich</td><td>E10521</td><td>Anesthetic; tricaine mesylate</td></tr><tr><td>NaCl</td><td>Merck Millipore</td><td>106404</td><td></td></tr><tr><td>NaHCO&lt;sub&gt;3&lt;/sub&gt;</td><td>Sigma-Aldrich</td><td>S6014</td><td></td></tr><tr><td>Nasco Frog Brittle for Tadpole Xenopus</td><td>Nasco</td><td>SB09480(LM)MX</td><td>Food for Xenopus tadpoles stage&amp;nbsp; 44 to 60</td></tr><tr><td>Oxygen diffuser stones</td><td>Pentair</td><td>AA1</td><td>Mantainance of animals</td></tr><tr><td>Pair of forceps</td><td>Fine Science Tools</td><td>Dumont n&amp;deg; 5 SF forceps</td><td>For surgery</td></tr><tr><td>Penicillin</td><td>Sigma-Aldrich</td><td>P7794</td><td></td></tr><tr><td>pH meter</td><td></td><td></td><td></td></tr><tr><td>Plastic Pasteur pipette</td><td>Sigma-Aldrich</td><td>Z331740</td><td>For collecting embryos after mating</td></tr><tr><td>Plastic Petri dishes</td><td>Sigma-Aldrich</td><td>P5981</td><td>150 x 15 mm</td></tr><tr><td>Plastic tank/box with lid</td><td></td><td></td><td>4.5 liter capacity; 20 cm &amp;times; 17 cm &amp;times; 15 cm or similar</td></tr><tr><td>Sterilized gauze</td><td></td><td></td><td></td></tr><tr><td>Streptomycin</td><td>Sigma-Aldrich</td><td>S1277</td><td></td></tr><tr><td>Tablespoon</td><td></td><td></td><td></td></tr><tr><td>&lt;em&gt;Xenopus laevis&lt;/em&gt;&lt;br/&gt; specialized strains and lines</td><td>National Xenopus Resource&lt;br/&gt; European Xenopus Resource Centre&lt;br/&gt; Xenopus laevis Research Resource Centre</td><td></td><td>http://www.mbl.edu/xenopus&lt;br/&gt; https://xenopusresource.org/&lt;br/&gt; https://www.urmc.rochester.edu/microbiology-immunology/xenopus-laevis.aspx</td></tr><tr><td>&lt;em&gt;Xenopus laevis&lt;/em&gt; wild type</td><td>Xenopus 1&lt;br/&gt; Xenopus Express</td><td></td><td>https://xenopus1.com&lt;br/&gt; http://www.xenopus.com</td></tr></tbody></table>

spinal cord transection in xenopus laevis tadpoles

Spinal cord injury (SCI) is a permanent affliction, which affects the central nervous system (CNS) motor and sensory nerves, resulting in paralysis beneath the injury site. To date, there is no functional recovery therapy for SCI, and there is a lack of clarity regarding the many complexes and dynamic events occurring after SCI. Many non-mammalian organisms can regenerate after severe SCI, such as teleost fishes, urodele amphibians, and larval stages of anuran amphibians, including Xenopus laevis tadpoles. These are bona fide model organisms to study and understand the response to SCI and the mechanisms underlying successful regenerative processes. This type of research can lead to the identification of potential targets for SCI therapeutic intervention. This article describes how to perform Xenopus laevis tadpole spinal cord transection, including husbandry, surgery, postsurgery care, and functional test evaluation. This injury method can be applied for elucidating the different steps of spinal cord regeneration by studying the cellular, molecular, and genetic mechanisms, as well as histological and functional evolution after SCI and during spinal cord regeneration.

The protocol described herein allows the study of spinal cord regeneration in Xenopus laevis. The effects of specific pharmacological treatments and the contribution of specific gene expression in spinal cord regeneration can be evaluated by measuring their effects on swimming recovery. The total swimming distance is plotted against the days after injury to compare control and treated animals at a specific time point or over a specified period. The recovery of motor function through time is exemplified in Figure 3, showing the swimming distance at 5, 10, 15, and 20 days post transection. At 5 days post transection, animals swam an average of 0.7 m in 5 min, showing a reduced swimming capacity. This capacity increased with the passing days, as an average of 2.1 and 3.1 m/5 min was observed after 10 and 15 days post transection, respectively, and complete recovery of swimming capacities was observed at 20 days post transection, with an average of 5.7 m/5 min.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/63276/63276fig01.jpg" /> 
Figure 1: Xenopus tadpole staging. Representative images of stages 49-51, showing fore- and hindlimbs for animal staging reference. Scale bars = 2 mm. Magnifications of the boxed region are shown in the lower-right of each image.Scale bars = 1 mm. In stage 49, forelimbs are not observed, while hindlimbs are just appearing, showing a spherical shape. Stage 50 presents forelimbs that are just appearing, showing a spherical shape and hindlimbs protruding with a spherical shape. In stage 51, forelimbs present a protruding spherical shape and hindlimbs a protruding elongated shape. Dashed outlines show fore- and hindlimbs. <a href="https://www.jove.com/files/ftp_upload/63276/63276fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/63276/63276fig02.jpg" /> 
Figure 2: Spinal cord transection. (A) Representative image showing the correct positioning of the animal, dorsal side up, for performing the surgery. Scale bar = 2 mm. (B) Magnification of A shows the location and extent of injury. The red cross shows the exact location of the injury site at the thoracic level of the spinal cord, and the dashed line shows the extent of the injury. Scale bar = 1 mm. (C) Representative image showing a lateral view of the thoracic level of the spinal cord. The extension of the sham incision and transection are shown. Dashed lines delineate the limits of the spinal cord. Scale bar = 1 mm. (D) Representative image showing a sham animal with an intact spinal cord. Scale bars = 1 mm. (E) Representative image showing a transected animal with an interrupted spinal cord. Scale bars = 1 mm. Magnifications of the boxed region are shown in the lower-right of each image (D&#180; and E&#180;).Scale bars = 1 mm. Abbreviations: S = sham incision; T = transection. <a href="https://www.jove.com/files/ftp_upload/63276/63276fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/63276/63276fig03.jpg" /> 
Figure 3: Swimming function recovery over time. Representative dot plot of the swimming distance covered by transected animals in 5 min at 5, 10, 15, and 20 days post transection. Samples of swimming trajectories are shown on top. Data presented as mean &#177; SEM from 10 tadpoles. Abbreviations: dpT = days post transection; SEM = standard error of the mean. <a href="https://www.jove.com/files/ftp_upload/63276/63276fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Spinal Cord Transection In Xenopus laevis Tadpoles at JoVE.com

Spinal Cord Transection In Xenopus laevis Tadpoles

Xenopus laevis tadpole spinal cord transection is a relevant injury method to study spinal cord injury and regeneration by making a transverse cut that completely severs the spinal cord at the thoracic level.

Spinal Cord Transection In Xenopus laevis Tadpoles

Spinal cord injury (SCI) is a permanent affliction, which affects the central nervous system (CNS) motor and sensory nerves, resulting in paralysis ...

spinal-cord-transection-in-xenopus-laevis-tadpoles

Research

JoVE Journal

Biology

3.7K Views.  P. Universidad Católica de Chile. Dit protocol maakt de studie van spinale regeneratie bij kikkers mogelijk. De opgedane kennis kan nieuwe inzichten opleveren om te begrijpen hoe mensen reageren op dwarslaesie. Het belangrijkste voordeel van dit protocol is het gemak bij het uitvoeren van de verwonding, en ook dat in vergelijking met het paradigma van staartamputatie, deze techniek beter nabootst wat er bij mensen gebeurt.Deze techniek maakt het mogelijk om het regeneratieve proces uit te dagen om de deelname van versc...

Video: Spinal Cord Transection In Xenopus laevis Tadpoles

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 ...

Neuroscience

Complete Spinal Cord Injury and Brain Dissection Protocol for Subsequent Wholemount In Situ Hybridization in Larval Sea Lamprey

After a complete spinal cord injury, sea lampreys at first are paralyzed below the level of transection. However, they recover locomotion after several weeks, and this is accompanied by short distance regeneration (a few mm) of propriospinal axons and spinal-projecting axons from the brainstem. Among the 36 large identifiable spinal-projecting neurons, some are good regenerators and others are bad regenerators. These neurons can most easily be identified in wholemount CNS preparations. In order to understand the neuron-intrinsic mechanisms that favor or inhibit axon regeneration after injury in the vertebrates CNS, we determine differences in gene expression between the good and bad regenerators, and how expression is influenced by spinal cord transection. This paper illustrates the techniques for housing larval and recently transformed adult sea lampreys in fresh water tanks, producing complete spinal cord transections under microscopic vision, and preparing brain and spinal cord wholemounts for in situ hybridization. Briefly, animals are kept at 16&#160;&#176;C and anesthetized in 1% Benzocaine in lamprey Ringer. The spinal cord is transected with iridectomy scissors via a dorsal approach and the animal is allowed to recover in fresh water tanks at 23 &#176;C. For in situ hybridization, animals are reanesthetized and the brain and cord removed via a dorsal approach.

Complete Spinal Cord Injury and Brain Dissection Protocol for Subsequent Wholemount In Situ Hybridization in Larval Sea Lamprey

Lampreys recover locomotion after a complete spinal cord injury. However, some spinal-projecting neurons are good regenerators and others are not. This paper illustrates the techniques for housing sea lamprey larvae (and recently transformed adults), producing complete spinal cord transections and preparing wholemount brains and spinal cords for in situ hybridization.

After a complete spinal cord injury, sea lampreys at first are paralyzed below the level of transection. However, they recover locomotion after several ...

Experimental Strategies to Bridge Large Tissue Gaps in the Injured Spinal Cord after Acute and Chronic Lesion

After a spinal cord injury (SCI) a scar forms in the lesion core which hinders axonal regeneration. Bridging the site of injury after an insult to the spinal cord, tumor resections, or tissue defects resulting from traumatic accidents can aid in facilitating general tissue repair as well as regenerative growth of nerve fibers into and beyond the affected area. Two experimental treatment strategies are presented: (1) implantation of a novel microconnector device into an acutely and completely transected thoracic rat spinal cord to readapt severed spinal cord tissue stumps, and (2) polyethylene glycol filling of the SCI site in chronically lesioned rats after scar resection. The chronic spinal cord lesion in this model is a complete spinal cord transection which was inflicted 5 weeks before treatment. Both methods have recently achieved very promising outcomes and promoted axonal regrowth, beneficial cellular invasion and functional improvements in rodent models of spinal cord injury.
The mechanical microconnector system (mMS) is a multi-channel system composed of polymethylmethacrylate (PMMA) with an outlet tubing system to apply negative pressure to the mMS lumen thus pulling the spinal cord stumps into the honeycomb-structured holes. After its implantation into the 1 mm tissue gap the tissue is sucked into the device. Furthermore, the inner walls of the mMS are microstructured for better tissue adhesion.
In the case of the chronic spinal cord injury approach, spinal cord tissue - including the scar-filled lesion area - is resected over an area of 4 mm in length. After the microsurgical scar resection the resulting cavity is filled with polyethylene glycol (PEG 600) which was found to provide an excellent substratum for cellular invasion, revascularization, axonal regeneration and even compact remyelination in vivo.

Severe spinal cord injuries often result in tissue defects. Two possibilities are described to successfully bridge such gaps to promote tissue adaptation, regenerative responses and functional improvement in rats via implantation of a mechanical microconnector system after acute injury and five weeks after complete spinal cord transection.

After a spinal cord injury (SCI) a scar forms in the lesion core which hinders axonal regeneration. Bridging the site of injury after an insult to the ...

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 ...

Transplantation of Schwann Cells Inside PVDF-TrFE Conduits to Bridge Transected Rat Spinal Cord Stumps to Promote Axon Regeneration Across the Gap

Among various models for spinal cord injury in rats, the contusion model is the most often used because it is the most common type of human spinal cord injury. The complete transection model, although not as clinically relevant as the contusion model, is the most rigorous method to evaluate axon regeneration. In the contusion model, it is difficult to distinguish regenerated from sprouted or spared axons due to the presence of remaining tissue post injury. In the complete transection model, a bridging method is necessary to fill the gap and create continuity from the rostral to the caudal stumps in order to evaluate the effectiveness of the treatments. A reliable bridging surgery is essential to test outcome measures by reducing the variability due to the surgical method. The protocols described here are used to prepare Schwann cells (SCs) and conduits prior to transplantation, complete transection of the spinal cord at thoracic level 8 (T8), insert the conduit, and transplant SCs into the conduit. This approach also uses in situ gelling of an injectable basement membrane matrix with SC transplantation that allows improved axon growth across the rostral and caudal interfaces with the host tissue.

This article describes a technique to insert a hollow conduit between the spinal cord stumps after complete transection and fill with Schwann cells (SCs) and injectable basement membrane matrix in order to bridge and promote axon regeneration across the gap.

Among various models for spinal cord injury in rats, the contusion model is the most often used because it is the most common type of human spinal cord ...

Medicine

Preparations and Protocols for Whole Cell Patch Clamp Recording of Xenopus laevis Tectal Neurons

The Xenopus tadpole retinotectal circuit, comprised of the retinal ganglion cells (RGCs) in the eye which form synapses directly onto neurons in the optic tectum, is a popular model to study how neural circuits self-assemble. The ability to carry out whole cell patch clamp recordings from tectal neurons and to record RGC-evoked responses, either in vivo or using a whole brain preparation, has generated a large body of high-resolution data about the mechanisms underlying normal, and abnormal, circuit formation and function. Here we describe how to perform the in vivo preparation, the original whole brain preparation, and a more recently developed horizontal brain slice preparation for obtaining whole cell patch clamp recordings from tectal neurons. Each preparation has unique experimental advantages. The in vivo preparation enables the recording of the direct response of tectal neurons to visual stimuli projected onto the eye. The whole brain preparation allows for the RGC axons to be activated in a highly controlled manner, and the horizontal brain slice preparation allows recording from across all layers of the tectum.

Preparations and Protocols for Whole Cell Patch Clamp Recording of Xenopus laevis Tectal Neurons

In this paper, we discuss three brain preparations used for whole cell patch clamp recording to study the retinotectal circuit of Xenopus laevis tadpoles. Each preparation, with its own specific advantages, contributes to the experimental tractability of the Xenopus tadpole as a model to study neural circuit function.

The Xenopus tadpole retinotectal circuit, comprised of the retinal ganglion cells (RGCs) in the eye which form synapses directly onto neurons in the optic ...

Functional Evaluation of Olfactory Pathways in Living Xenopus Tadpoles

Xenopus tadpoles offer a unique platform to investigate the function of the nervous system. They provide multiple experimental advantages, such as accessibility to numerous imaging approaches, electrophysiological techniques and behavioral assays. The Xenopus tadpole olfactory system is particularly well suited to investigate the function of synapses established during normal development or reformed after injury. Here, we describe methodologies to evaluate the processing of olfactory information in living Xenopus larvae. We outline a combination of in vivo measurements of presynaptic calcium responses in glomeruli of the olfactory bulb with olfactory-guided behavior assays. Methods can be combined with the transection of olfactory nerves to study the rewiring of synaptic connectivity. Experiments are presented using both wild-type and genetically modified animals expressing GFP reporters in central nervous system cells. Application of the approaches described to genetically modified tadpoles can be useful for unraveling the molecular bases that define vertebrate behavior.

Functional Evaluation of Olfactory Pathways in Living Xenopus Tadpoles

Xenopus tadpoles offer a unique platform to investigate the function of the nervous system in vivo. We describe methodologies to evaluate the processing of olfactory information in living Xenopus larvae in normal rearing conditions or after injury.

Xenopus tadpoles offer a unique platform to investigate the function of the nervous system. They provide multiple experimental advantages, such as ...

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 ...

Induction of Complete Transection-Type Spinal Cord Injury in Mice

Spinal cord injury (SCI) largely leads to irreversible and permanent loss of function, most commonly as a result of trauma. Several treatment options, such as cell transplantation methods, are being researched to overcome the debilitating disabilities arising from SCI. Most pre-clinical animal trials are conducted in rodent models of SCI. While rat models of SCI have been widely used, mouse models have received less attention, even though mouse models can have significant advantages over rat models. The small size of mice equates to lower animal maintenance costs than for rats, and the availability of numerous transgenic mouse models is advantageous for many types of studies. Inducing repeatable and precise injury in the animals is the primary challenge for SCI research, which in small rodents requires high-precision surgery. The transection-type injury model has been a commonly used injury model over the last decade for transplantation-based therapeutic research, however a standardized method for inducing a complete transection-type injury in mice does not exist. We have developed a surgical protocol for inducing a complete transection type injury in C57BL/6 mice at thoracic vertebral level 10 (T10). The procedure uses a small tip drill instead of rongeurs to precisely remove the lamina, after which a thin blade with rounded cutting edge is used to induce the spinal cord transection. This method leads to reproducible transection-type injury in small rodents with minimal collateral muscle and bone damage and therefore minimizes confounding factors, specifically where behavioral functional outcomes are analyzed.

This protocol describes how to create a precise laminectomy for induction of stable transection-type spinal cord injury in the mouse model, with minimal collateral damage for spinal cord injury research.

Spinal cord injury (SCI) largely leads to irreversible and permanent loss of function, most commonly as a result of trauma. Several treatment options, ...

Contusion Spinal Cord Injury via a Microsurgical Laminectomy in the Regenerative Axolotl

The purpose of this study is to establish a standardized and reproducible regenerative blunt spinal cord injury model in the axolotl (Ambystoma mexicanum). Most clinical spinal cord injuries occur as high energy blunt traumas, inducing contusion injuries. However, most studies in the axolotl spinal cord have been conducted with sharp traumas. Hence, this study aims to produce a more clinically relevant regenerative model. Due to their impressive ability to regenerate almost any tissue, axolotls are widely used as models in regenerative studies and have been used extensively in spinal cord injury (SCI) studies. In this protocol, the axolotls are anesthetized by submersion in a benzocaine solution. Under the microscope, an angular incision is made bilaterally at a level just caudal to the hind limbs. From this incision, it is possible to dissect and expose the spinous processes. Using forceps and scissors, a two-level laminectomy is performed, exposing the spinal cord. A custom trauma device consisting of a falling rod in a cylinder is constructed, and this device is used to induce a contusion injury to the spinal cord. The incisions are then sutured, and the animal recovers from anesthesia. The surgical approach is successful in exposing the spinal cord. The trauma mechanism can produce contusion injuries to the spinal cord, as confirmed by histology, MRI, and neurological examination. Finally, the spinal cord regenerates from the injury. The critical step of the protocol is removing the spinous processes without inflicting damage to the spinal cord. This step requires training to ensure a safe procedure. Furthermore, wound closure is highly dependent on not inflicting unnecessary damage to the skin during incision. The protocol was performed in a randomized study of 12 animals.

This manuscript presents protocols for surgically inflicting controlled blunt and sharp spinal cord injuries to a regenerative axolotl (Ambystoma mexicanum).

Spinal Cord Transection In Xenopus laevis Tadpoles

Samenvatting

Meer video's verkennen