This work was supported by a Griffith University International Student (PhD) stipend to RR, a Perry Cross Foundation Grant to JE and JSJ, a Clem Jones Foundation Grant to JSJ and JE, and a Motor Accident Insurance Commission of Queensland grant to JSJ and JE.

All procedures were carried out with the approval of the Griffith University Animal Ethics Committee (ESK/04/16 AEC and MSC/04/18 AEC) under the guidelines of the National Health and Medical Research Council of Australia.
1. Animal set-up procedure for the surgery
<ol>
	<li>Anesthetize and stabilize the animal.
	<ol>
		<li>Use 8&#8211;10-week old female C57BL/6 mice. Use 5% isoflurane in 1 L/min oxygen for induction of anesthesia. For maintenance of anesthesia, use 1.5&#8211;2% isoflurane in 1 L/min oxygen. Confirm appropriate anaesthetization by establishing a lack of pain reflex in tail and hind paws.</li>
	</ol>
	</li>
	<li>&#160;Administer buprenorphine (0.03 mg/kg body weight) for analgesia and enrofloxacin (10 mg/kg body weight) for antibiotic cover, subcutaneously according to body weight. Meloxicam (2 mg/kg body weight) can be given for long term analgesia if required.&#160;</li>
	<li>Keep the animal&#8217;s body temperature steady at 37 &#176;C with a heating pad
	<ol>
		<li>Shave the back fur to expose the surgical area over the dorsal spine. Remove the shaved fur from surgical area before sterilizing the incision site. Sterilize the shaved area with sterile cotton swabs soaked in povidone iodine antiseptic liquid and surgical spirit.</li>
	</ol>
	</li>
	<li>Tape the paws of the mouse to the surgical area to stabilize the animal (Figure 1A). Place an oval window drape over mouse (Figure 1B).</li>
</ol>
2. Laminectomy
<ol>
	<li>Make a vertical midline incision at the T10 vertebral level using a surgical scalpel.
	<ol>
		<li>Locate the spinous process of T10 vertebra to determine the site of laminectomy. The body of the vertebra lies slightly cranial to the tip of the spinous process8 (see Figure 2). The tip rests approximately at the midpoint of the T11 vertebra8.</li>
		<li>Make the incision ~2.5 cm long, so that the spine of T10 vertebra is approximately at the middle of the length of the incision.</li>
	</ol>
	</li>
	<li>Reflect the skin and retract with retractors.
	<ol>
		<li>Use the straight forceps to lift the skin from the underlying fascia. This will create space for the retractors to be placed; these will keep the surgical field open.</li>
	</ol>
	</li>
	<li>Perform blunt dissection of subcutaneous tissue and fascia to expose the spinous processes.
	<ol>
		<li>Use the blunt edge of the scalpel to make a small midline incision in the subcutaneous tissue and underlying fascia to expose the spines of T9&#8211;T11 vertebrae. Use the fine tip forceps (non-sharp) to perform blunt dissection and reflect the fascia.</li>
	</ol>
	</li>
	<li>Split and separate the para-spinous muscles to expose the laminae.
	<ol>
		<li>Use the blunt tip of the scalpel to split the dorsal trunk muscles and para-spinous muscles along the spines of the T9&#8211;T11 vertebrae. Use the blunt fine tip forceps to perform blunt dissection of the muscles in layers and expose the laminae of the vertebrae. This should minimize any bleeding. 
		NOTE: If there is any bleeding, use warmed saline (37 &#176;C) irrigation and cotton swabs to control it and clear blood from the surgical field.</li>
		<li>Use the same forceps to make small pockets around the transverse processes of T10 vertebra. Use the curved forceps to stabilize the T10 vertebral body by hooking its prongs under the transverse processes, in the pockets created (Figure 1C).</li>
		<li>Thoroughly rinse the T10 laminae with warm saline and gently wipe clean with cotton swabs to clearly visualize the bony surface. Make sure that no muscle/ligament attachments remain along the surface bilaterally.</li>
	</ol>
	</li>
	<li>Use a drill with a fine tip (0.55 mm diameter, 7 mm length) to break the laminae bilaterally.
	<ol>
		<li>Use the tip of the drill to trace a vertical path from T9&#8211;T10 intervertebral space to T10&#8211;T11 intervertebral space along both the T10 laminae, without turning the drill on. This is to ensure that the drill bit does not catch any tissue (Figure 1D).</li>
		<li>Now turning the drill on, slowly and carefully dig a vertical trench on the right lamina of the T10 vertebra. This part of the laminectomy should create a precise surgical defect throughout the thickness of the bone in a straight vertical line. Maintain the grip with the curved forceps to keep the vertebra stable.</li>
	</ol>
	</li>
	<li>Make sure that the tip of the drill does not penetrate through the bone and injure the spinal cord. Repeat the same process on the left side of the lamina, keeping the vertebra stable with the curved forceps. Irrigate with warm saline to wash away any remaining bone fragments.</li>
	<li>Lift and remove the posterior part of the neural arch (Figure 1F).
	<ol>
		<li>Use the angled fine tip forceps to grip the spinous process and remove the whole dorsal segment of the laminae separated by the bilateral drilling. Irrigate and swab again if there is any bleeding, to clearly visualize the spinal cord exposed under the laminectomy window (Figure 1E).</li>
	</ol>
	</li>
</ol>
3. Transection
<ol>
	<li>Induce the spinal cord injury by transection of the exposed cord with a single slice of the blade.
	<ol>
		<li>Use the narrow, round cutting edged blade to slice the cord at the center of the laminectomy window. Ensure to sweep the lateral recesses of the spinal column to induce a complete transection injury (Figure 1G).</li>
	</ol>
	</li>
	<li>Confirm completeness of the transection injury using the blunt fine tip forceps and remove any remaining connections at the transection site.</li>
	<li>Control the bleeding if any, before closing the surgical layers.</li>
	<li>Use the warm saline to irrigate and clear any bleeding that occurs from the transected cord stumps. Use a cotton swab to apply gentle pressure to achieve hemostasis if needed. Take care not to compress the spinal cord.</li>
</ol>
4. Closure and immediate post-operative care
<ol>
	<li>Bring the muscles together and suture in layer.
	<ol>
		<li>Once hemostasis is achieved at the transection site, release the curved forceps grip on the T10 vertebrae. Bring the edges of dissected muscles together along the midline to achieve good apposition.</li>
		<li>Suture the muscles in a layer using 5-0 polyglactin 910 absorbable sutures. Make sure that the natural curvature of the spine does not cause any tension at the suture line or open up the sutures, exposing laminectomy site.</li>
	</ol>
	</li>
	<li>Close subcutaneous tissue and skin.
	<ol>
		<li>Use 5-0 non-absorbable silk sutures to close the skin incision. Make sure that there is no bleeding, clots or debris remaining under the skin before closure. A final irrigation with warm saline may be necessary at this step.</li>
	</ol>
	</li>
	<li>Stop the anesthesia. Observe the animal for 10&#8211;30 min until recovery. The animal should remain on the heating pad for this duration. Provide water gel and hydrated food in the cage.</li>
	<li>For post-operative care, include buprenorphine (twice daily), enrofloxacin (once daily) for the first two days prophylactically. Additionally, manually empty the bladder at least twice daily, adhering to the animal ethics committee&#8217;s guidelines. 
	NOTE: The animals in this experiment were assessed twice daily for their general health and wellbeing, which included checking for persisting pain (to give additional doses of buprenorphine) or infection (additional enrofloxacin), their nutrition and hydration status (give injectable fluids if dehydrated) and any signs of autotomy (provide wound care if minor autotomy). It is strongly recommended that these aspects of post-operative care, including the euthanasia decisions must be determined with the guidance of the institutional animal ethics committee.</li>
</ol>
5. Evaluation of the injury model
<ol>
	<li>Establishing the loss of motor function.
	<ol>
		<li>Assess motor behavior on the injured mice 2, 7, 14, 21 and 28 days after the injury in an open field using the Basso Mouse Scale (BMS) to determine the loss of function9 (Figure 3C&#8211;E).</li>
	</ol>
	</li>
	<li>Histological confirmation of injury
	<ol>
		<li>Harvest the injured spinal cords with the vertebral columns after euthanasia (done with carbon dioxide in this experiment, as approved by the animal ethics committee).</li>
		<li>Fix the tissue by overnight submersion in 4% paraformaldehyde and decalcify the bones by treatment with 20% ethylene diamine tetraacetic acid (EDTA) over 3 weeks, replacing fresh EDTA every 48 h.</li>
		<li>Prepare the decalcified spines for cryo-sectioning and cut them into 30 &#181;m thick sections.</li>
		<li>Mount the sections on gelatin-coated glass slides for immuno-staining with anti-GFAP antibodies and Hoechst 33342.</li>
		<li>Image the slides on a fluorescent microscope (Figure 3A) and perform measurements of the injury size using image analysis software (e.g., Nikon analysis software - NIS Elements [Figure 3B]).</li>
	</ol>
	</li>
</ol>

<ol>
	<li>Sharif-Alhoseini, 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=Animal+models+of+spinal+cord+injury:+a+systematic+review.">Animal models of spinal cord injury: a systematic review.</a> Spinal Cord. 55 (8), 714-721 (2017).</li><li>Lee, D. H., Lee, J. K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Animal+models+of+axon+regeneration+after+spinal+cord+injury.">Animal models of axon regeneration after spinal cord injury.</a> Neuroscience Bulletin. 29 (4), 436-444 (2013).</li><li>Sharif-Alhoseini, M., Rahimi-Movaghar, V., Dionyssiotis, Y. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Topics in Paraplegia. , (2014).</li><li>Talac, 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=Animal+models+of+spinal+cord+injury+for+evaluation+of+tissue+engineering+treatment+strategies.">Animal models of spinal cord injury for evaluation of tissue engineering treatment strategies.</a> Biomaterials. 25 (9), 1505-1510 (2004).</li><li>Nakae, A., 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=The+animal+model+of+spinal+cord+injury+as+an+experimental+pain+model.">The animal model of spinal cord injury as an experimental pain model.</a> Journal of Biomedicine &amp; Biotechnology. 2011, 939023 (2011).</li><li>Kwon, B., Oxland, T., Tetzlaff, W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Animal+models+used+in+spinal+cord+regeneration+research.">Animal models used in spinal cord regeneration research.</a> Spine. 27, 1504-1510 (2002).</li><li>Kundi, S., Bicknell, R., Ahmed, Z. <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+injury:+current+mammalian+models.">Spinal cord injury: current mammalian models.</a> American Journal of Neuroscience. (4), 1-12 (2013).</li><li>Harrison, 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=Vertebral+landmarks+for+the+identification+of+spinal+cord+segments+in+the+mouse.">Vertebral landmarks for the identification of spinal cord segments in the mouse.</a> Neuroimage. 68, 22-29 (2013).</li><li>Basso, D. 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=Basso+Mouse+Scale+for+locomotion+detects+differences+in+recovery+after+spinal+cord+injury+in+five+common+mouse+strains.">Basso Mouse Scale for locomotion detects differences in recovery after spinal cord injury in five common mouse strains.</a> Journal of Neurotrauma. 23 (5), 635-659 (2006).</li><li>Seitz, A., Aglow, E., Heber-Katz, E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Recovery+from+spinal+cord+injury:+a+new+transection+model+in+the+C57Bl/6+mouse.">Recovery from spinal cord injury: a new transection model in the C57Bl/6 mouse.</a> Journal of Neuroscience Research. 67 (3), 337-345 (2002).</li></ol>

The authors have nothing to disclose.

This method induces a complete transection type injury at the T10 vertebral level in mice, which results in complete paraplegia of the animal, below the level of injury. Overall, this method results in minimal bleeding, negligible collateral damage and a stable, reproducible injury. As compared to previously published methods of transection without laminectomy10, this method offers the benefits in terms of direct visualization without manipulating the curvature of the spine, better control over co...

Spinal cord injury (SCI) is a complex medical problem that results in drastic changes in health and lifestyle. There is no cure for SCI, and the pathophysiology of SCI is not thoroughly understood. Animal SCI models, in particular rodent models, offer an invaluable tool for trialing new treatments, and have been used to explore SCI for decades. To date, over 72% of pre-clinical SCI studies have employed rat models, as compared to a mere 16% that have used mice1. Although rats, due to their larger size and tendency to form cavities akin to human SCIs, have traditionally been the preferred model animals for studying novel therapeutic approaches, mice (including many transgenic mouse models) are now being used more frequently to study cellular and molecular mechanisms of the SCI2. The mouse model offers additional benefits in terms of easier handling, faster reproductive rates and lower costs than rats; mice also exhibit a high degree of genomic similarity with humans1,2,3. The major disadvantage of the mouse model has been identified as the significantly smaller size that creates challenges for surgical interventions for creating and treating spinal cord injuries4,5.
There is a gap in the existing literature that highlights the need for a robust and reproducible surgical protocol to induce stable SCI in the mouse model. Therefore, we provide a novel and precise surgical approach in this protocol to overcome these limitations. This protocol provides in-depth guidelines to induce a transection-type injury in mice, as this injury type has been recognized to be the most appropriate to study regenerative and degenerative changes following an injury6, as well as neuroplasticity, neural circuitry and tissue engineering approaches7. We have chosen to induce the injury in the lower thoracic region, since thoracic level SCI is used most commonly in the literature1.

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		<col />
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		<tr height="21">
			<td height="21" style="height:21px;width:449px;">Baytril injectable 50 mg/mL, 50 mL</td>
			<td style="width:243px;">Provet</td>
			<td style="width:161px;">BAYT I</td>
			<td style="width:184px;">Post-operative care drug</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:449px;">Betadine 500 mL</td>
			<td style="width:243px;">Provet</td>
			<td style="width:161px;">BETA AS</td>
			<td>Consumable</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:449px;">Castroviejo needle holder, locking</td>
			<td style="width:243px;">ProSciTech</td>
			<td style="width:161px;">T149C</td>
			<td>Reusable</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:21px;width:449px;">Ceramic zirconia blade, round with sharp sides, single edge, angled</td>
			<td style="width:243px;">ProSciTech</td>
			<td style="width:161px;">TXD101A-X</td>
			<td>Reusable</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:449px;">Cotton swabs (5pcs)</td>
			<td style="width:243px;">Multigate</td>
			<td style="width:161px;">21-893</td>
			<td>Consumable</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:449px;">Dremel Micro</td>
			<td style="width:243px;">DREMEL</td>
			<td style="width:161px;">8050-N/18</td>
			<td>Cordless rotary tool</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:21px;width:449px;">Dressing forceps fine</td>
			<td style="width:243px;">Multigate</td>
			<td style="width:161px;">06-306</td>
			<td>Single use disposable</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:449px;">Drill bits</td>
			<td style="width:243px;">Kemmer Pr&#228;zision</td>
			<td style="width:161px;">SM 32 M 0550 070</td>
			<td>Reusable</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:449px;">Dumont #7b forceps</td>
			<td style="width:243px;">Fine Science Tools</td>
			<td style="width:161px;">11270-20</td>
			<td>Reusable</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:449px;">Dumont tweezers, style 5</td>
			<td style="width:243px;">ProSciTech</td>
			<td style="width:161px;">T05-822</td>
			<td>Reusable</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:449px;">Fur trimmer</td>
			<td style="width:243px;">WAHL</td>
			<td style="width:161px;">WA9884-312</td>
			<td>Zero Overlap Hair Trimmer</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:449px;">Iris scissors, Ti, sharp tips, straight, 90mm</td>
			<td style="width:243px;">ProSciTech</td>
			<td style="width:161px;">TY-3032</td>
			<td>Reusable</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:449px;">Isoflurane isothesia NXT 250</td>
			<td style="width:243px;">Provet</td>
			<td style="width:161px;">ISOF 00 HS</td>
			<td>Anaesthetic agent</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:449px;">Colibri Retractor - 4cm</td>
			<td style="width:243px;">Fine Science Tools</td>
			<td>17000-04</td>
			<td>Reusable</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:449px;">Scalpel handle</td>
			<td style="width:243px;">ProSciTech</td>
			<td style="width:161px;">T133</td>
			<td>Reusable</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:449px;">Signature latex surgical gloves size 7.5</td>
			<td style="width:243px;">Medline</td>
			<td style="width:161px;">MSG5475</td>
			<td>Consumable</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:449px;">Sodium Chloride 0.9%</td>
			<td style="width:243px;">STS</td>
			<td>PHA19042005</td>
			<td>Consumable</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:449px;">Sterile Dressing Pack</td>
			<td style="width:243px;">Multigate</td>
			<td style="width:161px;">08-709</td>
			<td>Single use disposable</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:449px;">Sterile Fluid Impervious Drape 60x60 cm</td>
			<td style="width:243px;">Multigate</td>
			<td style="width:161px;">29-220</td>
			<td>Single use disposable</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:449px;">Surgical spirit 100 mL</td>
			<td style="width:243px;">Provet</td>
			<td style="width:161px;"># SURG SP</td>
			<td>Consumable</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:449px;">Suture Material - SILK BLK 45CM 5/0 FS-2</td>
			<td style="width:243px;">Johnson &#38; Johnson Medical</td>
			<td style="width:161px;">682G</td>
			<td>Silk Suture</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:449px;">Suture Material - Vicryl 70CM 5-0 S/A FS-2</td>
			<td style="width:243px;">Johnson &#38; Johnson Medical</td>
			<td style="width:161px;">VCP421H</td>
			<td>Vicryl Suture</td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:449px;">Temgesic 0.3 mg in 1 mL, x 5 ampoules (class S8 drug)</td>
			<td style="width:243px;">Provet</td>
			<td style="width:161px;">TEMG I</td>
			<td>Post-operative care drug</td>
		</tr>
	</tbody>
</table>

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.

The resulting method as depicted in Figure 1, involves adequate stabilization of the mouse (Figure 1A) and good visualization of the spine and paraspinous tissue (Figure 1B). Spinous process and laminae can be clearly visualized with minimal muscle dissection and blood loss (Figure 1C, highlighted zone). The fine tip drilling is performed as shown in Figure 1D, to create a ...

Watch this Scientific Journal Video about Induction of Complete Transection-Type Spinal Cord Injury in Mice at JoVE.com

Induction of Complete Transection-Type Spinal Cord Injury in Mice

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, ...

induction-of-complete-transection-type-spinal-cord-injury-in-mice

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8.1K Views.  Griffith University. 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.

Video: Induction of Complete Transection-Type Spinal Cord Injury in Mice

Dissection and Culture of Commissural Neurons from Embryonic Spinal Cord

Commissural neurons have been widely used to investigate the mechanisms underlying axon guidance during embryonic spinal cord development. The cell bodies of these neurons are located in the dorsal spinal cord and their axons follow stereotyped trajectories during embryonic development. Commissural axons initially project ventrally towards the floorplate. After crossing the midline, these axons turn anteriorly and project towards the brain. Each of these steps is regulated by the action of several guidance cues. Cultures highly enriched in commissural neurons are ideally suited for many experiments addressing the mechanisms of axon pathfinding, including turning assays, immunochemistry and biochemistry. Here, we describe a method to dissect and culture commissural neurons from E13 rat dorsal spinal cord. First, the spinal cord is isolated and dorsal strips are dissected out. The dorsal tissue is then dissociated into a cell suspension by trypsinization and mechanical disruption. Neurons are plated onto poly-L-lysine-coated glass coverslips or tissue-culture dishes. After 30 hours in vitro, most neurons have extended an axon. The purity of the culture (Yam et al. 2009), typically over 90%, can be assessed by immunolabeling with the commissural neuron markers DCC, LH2 and TAG1 (Helms and Johnson, 1998). This neuronal preparation is a useful tool for in vitro studies of the cellular and molecular mechanisms of commissural axon growth and guidance during spinal cord development.

This video demonstrates a method to dissect and culture commissural neurons from E13 rat dorsal spinal cord. Dissociated commissural neurons are useful to study the cellular and molecular mechanisms of axon growth and guidance.

Commissural neurons have been widely used to investigate the mechanisms underlying axon guidance during embryonic spinal cord development. The cell bodies ...

Lateral Fluid Percussion: Model of Traumatic Brain Injury in Mice

Traumatic brain injury (TBI) research has attained renewed momentum due to the increasing awareness of head injuries, which result in morbidity and mortality. Based on the nature of primary injury following TBI, complex and heterogeneous secondary consequences result, which are followed by regenerative processes 1,2. Primary injury can be induced by a direct contusion to the brain from skull fracture or from shearing and stretching of tissue causing displacement of brain due to movement 3,4. The resulting hematomas and lacerations cause a vascular response 3,5, and the morphological and functional damage of the white matter leads to diffuse axonal injury 6-8. Additional secondary changes commonly seen in the brain are edema and increased intracranial pressure 9. Following TBI there are microscopic alterations in biochemical and physiological pathways involving the release of excitotoxic neurotransmitters, immune mediators and oxygen radicals 10-12, which ultimately result in long-term neurological disabilities 13,14. Thus choosing appropriate animal models of TBI that present similar cellular and molecular events in human and rodent TBI is critical for studying the mechanisms underlying injury and repair.
Various experimental models of TBI have been developed to reproduce aspects of TBI observed in humans, among them three specific models are widely adapted for rodents: fluid percussion, cortical impact and weight drop/impact acceleration 1. The fluid percussion device produces an injury through a craniectomy by applying a brief fluid pressure pulse on to the intact dura. The pulse is created by a pendulum striking the piston of a reservoir of fluid. The percussion produces brief displacement and deformation of neural tissue 1,15. Conversely, cortical impact injury delivers mechanical energy to the intact dura via a rigid impactor under pneumatic pressure 16,17. The weight drop/impact model is characterized by the fall of a rod with a specific mass on the closed skull 18. Among the TBI models, LFP is the most established and commonly used model to evaluate mixed focal and diffuse brain injury 19. It is reproducible and is standardized to allow for the manipulation of injury parameters. LFP recapitulates injuries observed in humans, thus rendering it clinically relevant, and allows for exploration of novel therapeutics for clinical translation 20.
We describe the detailed protocol to perform LFP procedure in mice. The injury inflicted is mild to moderate, with brain regions such as cortex, hippocampus and corpus callosum being most vulnerable. Hippocampal and motor learning tasks are explored following LFP.

Lateral fluid percussion (LFP), an established model of traumatic brain injury in mice, is demonstrated. LFP fulfills three major criteria for animal models: validity, reliability and clinical relevance. The procedure, consisting of surgical craniotomy, fixation of hub followed by induction of injury, resulting in focal and diffuse injuries, is described.

Traumatic brain injury (TBI) research has attained renewed momentum due to the increasing awareness of head injuries, which result in morbidity and ...

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 ...

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 ...

An Ex Vivo Laser-induced Spinal Cord Injury Model to Assess Mechanisms of Axonal Degeneration in Real-time

Injured CNS axons fail to regenerate and often retract away from the injury site. Axons spared from the initial injury may later undergo secondary axonal degeneration. Lack of growth cone formation, regeneration, and loss of additional myelinated axonal projections within the spinal cord greatly limits neurological recovery following injury. To assess how central myelinated axons of the spinal cord respond to injury, we developed an ex vivo living spinal cord model utilizing transgenic mice that express yellow fluorescent protein in axons and a focal and highly reproducible laser-induced spinal cord injury to document the fate of axons and myelin (lipophilic fluorescent dye Nile Red) over time using two-photon excitation time-lapse microscopy. Dynamic processes such as acute axonal injury, axonal retraction, and myelin degeneration are best studied in real-time. However, the non-focal nature of contusion-based injuries and movement artifacts encountered during in vivo spinal cord imaging make differentiating primary and secondary axonal injury responses using high resolution microscopy challenging. The ex vivo spinal cord model described here mimics several aspects of clinically relevant contusion/compression-induced axonal pathologies including axonal swelling, spheroid formation, axonal transection, and peri-axonal swelling providing a useful model to study these dynamic processes in real-time. Major advantages of this model are excellent spatiotemporal resolution that allows differentiation between the primary insult that directly injures axons and secondary injury mechanisms; controlled infusion of reagents directly to the perfusate bathing the cord; precise alterations of the environmental milieu (e.g., calcium, sodium ions, known contributors to axonal injury, but near impossible to manipulate in vivo); and murine models also offer an advantage as they provide an opportunity to visualize and manipulate genetically identified cell populations and subcellular structures. Here, we describe how to isolate and image the living spinal cord from mice to capture dynamics of acute axonal injury.

An Ex Vivo Laser-induced Spinal Cord Injury Model to Assess Mechanisms of Axonal Degeneration in Real-time

We present a protocol utilizing two-photon excitation time-lapse microscopy to simultaneously visualize the dynamics of axon and myelin injuries in real time. This proposed protocol permits studies of both intrinsic and extrinsic factors which can influence central myelinated axon fate after injury and contribute to permanent clinical disability.

Injured CNS axons fail to regenerate and often retract away from the injury site. Axons spared from the initial injury may later undergo secondary axonal ...

Spinal Cord Neurons Isolation and Culture from Neonatal Mice

We present a protocol for the isolation and culture of spinal cord neurons. The neurons are obtained from neonatal C57BL/6 mice and are isolated on postnatal day 1-3. A mouse litter, usually 4-10 pups born from one breeding pair, is gathered for one experiment, and spinal cords are collected individually from each mouse after euthanasia with isoflurane. The spinal column is dissected out and then the spinal cord is released from the column. The spinal cords are then minced to increase the surface area of delivery for an enzymatic protease that allows for the neurons and other cells to be released from the tissue. Trituration is then used to release the cells into solution. This solution is subsequently fractionated in a density gradient to separate the various cells in solution, allowing for neurons to be isolated. Approximately 1-2.5 x 106 neurons can be isolated from one litter group. The neurons are then seeded onto wells coated with adhesive factors that allow for proper growth and maturation. The neurons take approximately 7 days to reach maturity in the growth and culture medium and can be used thereafter for treatment and analysis.

This study presents a technique for the isolation of neurons from WT neonatal mice. It requires the careful dissection of the spinal cord from the neonatal mouse, followed by the separation of neurons from the spinal cord tissue through mechanical and enzymatic cleavage.

We present a protocol for the isolation and culture of spinal cord neurons. The neurons are obtained from neonatal C57BL/6 mice and are isolated on ...

Imaging Neural Activity in the Primary Somatosensory Cortex Using Thy1-GCaMP6s Transgenic Mice

The mammalian brain exhibits marked symmetry across the sagittal plane. However, detailed description of neural dynamics in symmetric brain regions in adult mammalian animals remains elusive. In this study, we describe an experimental procedure for measuring calcium dynamics through dual optical windows above bilateral primary somatosensory corticies (S1) in Thy1-GCaMP6s transgenic mice using 2-photon (2P) microscopy. This method enables recordings and quantifications of neural activity in bilateral mouse brain regions one at a time in the same experiment for a prolonged period in vivo. Key aspects of this method, which can be completed within an hour, include minimally invasive surgery procedures for creating dual optical windows, and the use of 2P imaging. Although we only demonstrate the technique in the S1 area, the method can be applied to other regions of the living brain facilitating the elucidation of structural and functional complexities of brain neural networks.

Imaging Neural Activity in the Primary Somatosensory Cortex Using Thy1-GCaMP6s Transgenic Mice

We describe an experimental procedure for measuring neuronal activity through dual optical windows above bilateral primary somatosensory corticies (S1) in Thy1-GCaMP6s transgenic mice using 2-photon (2P) microscopy in vivo.

The mammalian brain exhibits marked symmetry across the sagittal plane. However, detailed description of neural dynamics in symmetric brain regions in ...

An In Vivo Duo-color Method for Imaging Vascular Dynamics Following Contusive Spinal Cord Injury

Spinal cord injury (SCI) causes significant vascular disruption at the site of injury. Vascular pathology occurs immediately after SCI and continues throughout the acute injury phase. In fact, endothelial cells appear to be the first to die after a contusive SCI. The early vascular events, including increased permeability of the blood-spinal cord barrier (BSCB), induce vasogenic edema and contribute to detrimental secondary injury events caused by complex injury mechanisms. Targeting the vascular disruption, therefore, could be a key strategy to reduce secondary injury cascades that contribute to histological and functional impairments after SCI. Previous studies were mostly performed on postmortem samples and were unable to capture the dynamic changes of the vascular network. In this study, we have developed an in vivo duo-color two-photon imaging method to monitor acute vascular dynamic changes following contusive SCI. This approach allows detecting blood flow, vessel diameter, and other vascular pathologies at various sites of the same rat pre- and post-injury. Overall, this method provides an excellent venue for investigating vascular dynamics.

An In Vivo Duo-color Method for Imaging Vascular Dynamics Following Contusive Spinal Cord Injury

We introduce an in vivo imaging method using two different fluorescent dyes to track dynamic spinal vascular changes following a contusive spinal cord injury in adult Sprague-Dawley rats.

Spinal cord injury (SCI) causes significant vascular disruption at the site of injury. Vascular pathology occurs immediately after SCI and continues ...

Activity-based Training on a Treadmill with Spinal Cord Injured Wistar Rats

Spinal cord injury (SCI) results in lasting deficits that include both mobility and a multitude of autonomic-related dysfunctions. Locomotor training (LT) on a treadmill is widely used as a rehabilitation tool in the SCI population with many benefits and improvements to daily life. We utilize this method of activity-based task-specific training (ABT) in rodents after SCI to both elucidate the mechanisms behind such improvements and to enhance and improve upon existing clinical rehabilitation protocols. Our current goal is to determine the mechanisms underlying ABT-induced improvements in urinary, bowel, and sexual function in SCI rats after a moderate to severe level of contusion. After securing each individual animal in a custom-made adjustable vest, they are secured to a versatile body weight support mechanism, lowered to a modified three-lane treadmill and assisted in step-training for 58 minutes, once a day for 10 weeks. This setup allows for the training of both quadrupedal and forelimb-only animals, alongside two different non-trained groups. Quadrupedal-trained animals with body weight support are aided by a technician present to assist in stepping with proper hind limb placement as necessary, while forelimb-only trained animals are raised at the caudal end to ensure no hind limb contact with the treadmill and no weight-bearing. One non-trained SCI group of animals is placed in a harness and rests next to the treadmill, while the other control SCI group remains in its home cage in the training room nearby. This paradigm allows for the training of multiple SCI animals at once, thus making it more time-efficient in addition to ensuring that our pre-clinical animal model mimics the clinical representation as close as possible, particularly with respect to the body weight support with manual assistance.

This protocol demonstrates our model of activity-based locomotor treadmill training for rats with spinal cord injury (SCI). Included is both quadrupedal and forelimb-only groups, in addition to two distinct types of non-trained control groups. Investigators are able to assess training effects on SCI rats using this protocol.

Spinal cord injury (SCI) results in lasting deficits that include both mobility and a multitude of autonomic-related dysfunctions. Locomotor training (LT) ...

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

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

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

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