Name: calibrated forceps model of spinal cord compression injury
Uploaded: 2015-04-24T15:00:00
Duration: 9 min 41 s
Description: Watch this Scientific Journal Video about Calibrated Forceps Model of Spinal Cord Compression Injury at JoVE.com

Funding for this work was provided by Shriners Hospitals and CIRM fellowships to AMC and AM. We would also like to acknowledge Plemel et al.7 for first designing and publishing the calibrated forceps model.

All animal procedures and animal care methods should be approved by the institution&#39;s Institutional Animal Care and Usage Committee (IACUC).&#160;
1. Surgical Preparation
<ol>
	<li>Assemble all necessary surgical tools and reagents: forceps, vana scissors, roungeurs, retractor, scalpels, scissors, sutures, skin staples, Q-tips, sterile saline, surgical sponges, and isoflurane. Prepare and autoclave a complete pack of surgical tools prior to surgery. If performing surgeries on more than one mouse, prepare and autoclave one packet of tools per animal, or autoclave one packet of surgical instruments and sterilize with a tool sterilizer in between surgeries. Generally, one pack of sterilized tools can be used on up to 5 mice if a glass bead tool terilizer is used to clean tools in-between surgeries (after 5 animals the tools must be re-cleaned and sterilized before use). Please check with your local IACUC administrator for institution specific guidelines.</li>
	<li>Sterilize the surgical field with 70% isopropyl alcohol wipes. Set up surgical drapes to ensure a sterile field is maintained during surgery.</li>
	<li>Weigh each mouse prior to surgery. Administer a dose of 0.05 mg/kg body weight of buprenorphine, subcutaneously.</li>
	<li>Anesthetize the animal by administering isoflurane at a dose of 4% in the induction chamber of isoflurane machine.</li>
	<li>Once the animal is anesthetized, apply ointment to the eyes to prevent dehydration, set the animal on a heating pad at 37 &#176;C, and ensure that the mouse&#8217;s head is properly situated in the anesthesia cone. (Note: Use a heating source that will not cause thermal burns; i.e., a circulating water blanket, hot water bottle, or equivalent.)&#160;At this point administer a dose of 2% isoflurane to the animal.</li>
	<li>Shave the dorsal surface of the mouse, approximately 1 cm around the intended incision location.</li>
	<li>Disinfect the incision site by washing with 70% isopropyl alcohol wipes, then with an iodine solution (10% Povidone-iodine, 1% available iodine). Repeat 3 times.</li>
</ol>
2. Dorsal Laminectomy
<ol>
	<li>Before making an incision, ensure that the animal is properly anesthetized by checking for reflexes using the toe or tail pinch method.</li>
	<li>Make an incision along the dorsal spine with scalpel and blade, and then check for reflexes again. Arch the back to help properly visualize landmarks, such as the boundaries between vertebrae.</li>
	<li>Cut through the skin. Insert retractor to hold skin and fascia back from the spinal cord. Clear the tissue on either side of the spinal cord to expose the muscles covering the spine. 
	NOTE: The surgeon should be familiar with the anatomical landmarks. For example, the inferior angle of the scapula corresponds with T7. The top of the natural curve of the mouse&#8217;s spinal is T12 and may be used as a reference point.</li>
	<li>With proper lighting, determine the space between the vertebrae. Find the posterior end of T10 and cut muscles and fascia perpendicular to the intervertebral disk space. Cut just enough to expose the spinous process and posterior lamina of T10.</li>
	<li>Using a pair of fine tipped Dumont #5 forceps, remove some of the tissue from the lamina and spinous process to expose a small sliver of the spinal cord. When necessary, use tissue forceps to stabilize the spinal column.</li>
	<li>Perform the laminectomy by inserting one side of a pair of small vana scissors along the dorsolateral side of the vertebra and just beneath the lamina.
	<ol>
		<li>Make small, careful snips to cut through the lateral side of the vertebral lamina. Ensure that no pressure is applied to the spinal cord.</li>
		<li>Repeat on the other side.</li>
		<li>Apply gentle pressure to halt bleeding as necessary with a Q-tip or surgical sponge, taking care to not apply pressure to the spinal cord. 
		NOTE: Prepare gel foam soaked in sterile saline in the event that bleeding needs to be controlled.</li>
		<li>Once the incisions have been made, lift off the dorsal aspect of the vertebra and gently clear any tissue attachments. Use appropriate means to control bleeding if necessary.</li>
	</ol>
	</li>
</ol>
3. Spinal Cord Compression
<ol>
	<li>Using roungeurs or laminectomy forceps, ensure that the lateral sides of the spinal cord are free of vertebral bone so that the calibrated forceps for the compression injury can situate on either side of the spinal cord. The arms of the forceps must be able to be placed within the epidural space on adjacent sides of the spinal cord and the tips of the forceps must be able to reach the floor of the vertebral canal.
	<ol>
		<li>Ensure that visibility of the spinal cord is good.</li>
	</ol>
	</li>
	<li>Situate the calibrated Dumont #5 forceps approximately in the middle of the exposed segment of the spinal cord. Recall that the arms of the forceps must be placed within the epidural space on adjacent sides and the tips must touch the floor of the vertebral canal to generate reproducible injuries.</li>
	<li>Carefully compress the spinal cord until the spacers connect. Hold in place for 15 sec.</li>
	<li>Gently release the compressive force and remove the calibrated forceps from the spinal cord. Sterile saline should be used to regain homeostasis before the wound is closed.</li>
</ol>
4. Wound Closing
<ol>
	<li>Carefully suture the muscle layer over the spinal cord, taking care to not disrupt or apply pressure to the spinal cord.</li>
	<li>Use either sutures or staples to close the skin over the wound.</li>
	<li>If using gas anesthetic, begin to taper/turn off the anesthetic.</li>
	<li>Administer 0.1 ml of lactated ringers per 10 g body weight to help account for dehydration during the surgery and after the surgery when the animal is lethargic and recovering from the injury. The solution should be warmed to body temperature prior to injection.</li>
	<li>Place the mouse in a bedding-free cage. The cage should&#160;rest&#160;on top of a heating pad (as described in 1.1.5) in a manner that allows half of the cage area to be on the pad, while the other half is resting on a RT counter, in order to give the mouse climate options once it is ambulatory. Take care to ensure that the &#8220;recovery cage&#8221; is located in a quiet environment. Closely monitor the animal until it has regained consciousness, at which time the mouse can be transferred to a regular cage with bedding.</li>
</ol>
5. Post-operative Care
<ol>
	<li>After the surgery, administer a dose of 0.05 mg/kg body weight buprenorphine subcutaneously every 12 hr for the first 3 days after surgery, and then as needed to manage symptoms of pain.</li>
	<li>Administer a dose of lactated ringers (0.1 ml per 10 g body weight subcutaneously) for the first 3 - 5 days after surgery. Give this dose if/when the animal begins to exhibit signs of dehydration outside of this initial time period.</li>
	<li>Manually express the bladder of the animals using the Crede maneuver twice daily. Gently palpate the animal&#8217;s abdomen to locate the bladder, and then apply gentle downward pressure until the bladder is empty.
	<ol>
		<li>If the bladder does not empty or the urine is bloody or cloudy, administer 50 mg/kg by body weight of Baytril to the animal via interperitoneal injection for 10 days.</li>
	</ol>
	</li>
	<li>Monitor animals for signs of autophagy, dehydration, and excessive weight loss (greater than 20% of body weight). If an animal experiences any of these symptoms please consult with a veterinarian immediately with regards to treatment options, or euthanize the animal in a humane manner following IACUC guidelines.
	<ol>
		<li>Given that mobility may be limited right after surgery, take the necessary steps to make sure animals have access to food and water. Pre-packaged wet food, as well as hydrogel, can be made available to animals in these instances.</li>
	</ol>
	</li>
</ol>
6. Assess Tissue Damage Resulting from Compression Injury
<ol>
	<li>Anesthetize animal as described in steps 1.4 and 1.5. Check depth of anesthesia by toe pinch and monitoring corneal blink reflexes. When the animal is insensitive to stimuli, proceed to step 6.2.</li>
	<li>Perform an intracardial perfusion10.
	<ol>
		<li>Expose the chest cavity and insert a needle into the left ventricle. Flush out existing fluids with 20 - 30 ml ice-cold phosphate buffered saline (PBS) followed by 15 - 25 ml of ice-cold 4% paraformaldehyde (PFA).</li>
	</ol>
	</li>
	<li>Remove the spinal cord.
	<ol>
		<li>Cut the skin dorsal to the spinal cord and clear away any excess tissue surrounding the length of the spinal column.</li>
		<li>Excise spinal column and cut away any remaining tissue. The actual level of the laminectomy and injury can be confirmed by counting ribs.</li>
		<li>Use vanna scissors and forceps to displace small sections of the spinal column in a caudal-to-rostral direction. Continue to cut until the cord is exposed enough to allow safe removal. Visualization of the cutting location and process can be facilitated through the use of a stereoscope.</li>
	</ol>
	</li>
	<li>Place spinal cord tissue in 4% PFA. Allow tissue to post-fix in this solution for 24 hr at 4 &#176;C.</li>
	<li>Cryoprotect tissue by incubating in 30% sucrose for 24 hr at 4 &#176;C.</li>
	<li>Embed tissue in OCT. Briefly, take tissue from the 30% sucrose incubation and remove any excess solution. Place the tissue into a cryomold filled with OCT and incubate for 1 hr at 4 &#176;C.
	<ol>
		<li>Remove mold from 4 &#176;C, confirm directional orientation of the tissue, put the mold in a shallow dish of 2-methylbutane (pre-chilled for 1 hr on dry ice) and allow OCT to completely solidify. Keep on dry ice if using immediately or store at -80 &#176;C.</li>
	</ol>
	</li>
	<li>Cut the tissue into 20 &#181;m sagittal sections using a cryostat. Mount tissue directly onto slides. Store at -20 &#176;C until use.</li>
	<li>Perform Hematoxylin and Eosin (H&#38;E) stain.
	<ol>
		<li>Briefly, rehydrate the tissue (5 min, 2 times), stain with Hematoxylin (2.5 min), and wash with water (1 min, 2 times).</li>
		<li>Incubate tissue in 50% and then 70% ethanol (3 min each), stain with Eosin (45 sec), and dehydrate by incubation in 90% (5 sec), 95% (5 sec), 100% ethanol (2 min), and isopropanol (2 min).</li>
		<li>Clear with xylene (5 min, 3 times). 
		NOTE: H&#38;E staining will vary with specific tissue thickness and conditions. Therefore, standardization is required before proceeding with experimental tissues samples.</li>
	</ol>
	</li>
	<li>Cover tissue with a thin strip of Permount (~100 &#181;m) and coverslip. Press down on all sides of coverslip to ensure proper distribution of fluid. Let slides dry O/N.</li>
	<li>Visualize tissue sections using a digital microscope and capture images using the accompanying software.</li>
</ol>

<ol>
	<li>McDonough, A., Martinez-Cerdeno, V. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Endogenous+proliferation+after+SCI+in+animal+models.">Endogenous proliferation after SCI in animal models.</a> Stem Cells Int. 2012, 387513 (2012).</li><li>Onifer, S. M., Rabchevsky, A. G., Scheff, S. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Rat+models+of+traumatic+SCI+to+assess+motor+recovery.">Rat models of traumatic SCI to assess motor recovery.</a> ILAR J. 48 (4), 385-395 (2007).</li><li>Bunge, R. P., Puckett, W. R., Becerra, J. L., Marcillo, A., Quencer, 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=Observations+on+the+pathology+of+human+SCI.+A+review+and+classification+of+22+new+cases+with+details+from+a+case+of+chronic+cord+compression+with+extensive+focal+demyelination.">Observations on the pathology of human SCI. A review and classification of 22 new cases with details from a case of chronic cord compression with extensive focal demyelination.</a> Adv Neurol. 59, 75-89 (1993).</li><li>Beattie, M. S., 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=Endogenous+repair+after+spinal+cord+contusion+injuries+in+the+rat.">Endogenous repair after spinal cord contusion injuries in the rat.</a> Exp Neurol. 148 (2), 453-463 (1997).</li><li>Basso, D. M., Beattie, M. S., Bresnahan, J. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Graded+histological+and+locomotor+outcomes+after+spinal+cord+contusion+using+the+NYU+weight-drop+device+versus+transection.">Graded histological and locomotor outcomes after spinal cord contusion using the NYU weight-drop device versus transection.</a> Experimental Neurology. 139 (2), 244-256 (1996).</li><li>Krishna, V., 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=A+contusion+model+of+severe+SCI+in+rats.">A contusion model of severe SCI in rats.</a> J Vis Exp. 78, (2013).</li><li>Plemel, J. 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=A+graded+forceps+crush+SCI+model+in+mice.">A graded forceps crush SCI model in mice.</a> J Neurotrauma. 25 (4), 350-370 (2008).</li><li>Wu, D., Shibuya, S., Miyamoto, O., Itano, T., Yamamoto, T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Increase+of+NG2-positive+cells+associated+with+radial+glia+following+traumatic+SCI+in+adult+rats.">Increase of NG2-positive cells associated with radial glia following traumatic SCI in adult rats.</a> J Neurocytol. 34 (6), 459-469 (2005).</li><li>Namiki, J., Tator, C. H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cell+proliferation+and+nestin+expression+in+the+ependyma+of+the+adult+rat+spinal+cord+after+injury.">Cell proliferation and nestin expression in the ependyma of the adult rat spinal cord after injury.</a> Journal of Neuropathology and Experimental Neurology. 58 (5), 489-498 (1999).</li><li>Teletin, 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=Histopathology+in+Mouse+Metabolic+Investigations.">Histopathology in Mouse Metabolic Investigations.</a> Current Protocols in Molecular Biology. 29, (2007).</li><li>McDonough, A., Hoang, A. N., Monterrubio, A. M., Greenhalgh, S., Martinez-Cerdeno, V. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Compression+injury+in+the+mouse+spinal+cord+elicits+a+specific+proliferative+response+and+distinct+cell+fate+acquisition+along+rostro-caudal+and+dorso-ventral+axes.">Compression injury in the mouse spinal cord elicits a specific proliferative response and distinct cell fate acquisition along rostro-caudal and dorso-ventral axes.</a> Neuroscience. 254, 1-17 (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+SCI+in+five+common+mouse+strains.">Basso Mouse Scale for locomotion detects differences in recovery after SCI in five common mouse strains.</a> J Neurotrauma. 23 (5), 635-659 (2006).</li><li>Chaplan, S. 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The authors have nothing to disclose.

The choice of an SCI model is important in designing experiments to determine the efficacy of treatments for human cases of SCI. Such experiments require an animal model that is highly reproducible to limit variability that may result in inconclusive data. They should also be of clinical relevance to accurately evaluate the human condition they are modeling. To that end, choosing a compressive or contusive injury over a transection is more clinically relevant3. However, impactors and weight drop apparatuses for contusion injuries require the use of expensive and complicated machinery. In contrast, the calibrated forceps model of SCI utilizes modified forceps that are easy to assemble from common laboratory materials, and the surgery requires only one additional step after a standard dorsal laminectomy to expose the spinal cord. However, one drawback of using this method is that the compressive force is always applied laterally rather than dorsally, as is most often seen in human clinical cases of SCI9, and compressive injuries generated using the method affect a greater rostral-caudal extent of tissue than contusion models1,2. This model has been demonstrated by the originators of the technique, and us, to generate reproducible SCI7,11, and is well suited to the size of mice. Furthermore, this injury model allows for animals to be evaluated after surgery and therapeutic treatments using a multitude of behavioral tests, such as the Basso Mouse Scale for Locomotion and the von Frey hair test, to verify that a cohort of animals share the same injury severity and neurological deficits7,11-13. These same techniques can also be used to evaluate the efficacy of treatments administered to animals during investigative studies, fulfilling the general criteria for animal models used to evaluate therapies for SCI2,7.
The method of producing the calibrated forceps for the injury model is simple and can be accomplished with a variety of different methods. We have used the spacer method11, as published by Plemel7, and have also modified forceps using a small screw, which not only provides an easier method for creating the compression device, but also allows for versatility in adjusting the final compression width, of benefit for comparative studies. The range of options in creating the forceps is nearly unlimited so long as the spacer(s) provide a stable means of always closing the forceps to the same distance and can withstand autoclaving and sterilization. The surgical methods described within this video are highly reproducible across users, however it is necessary that care be taken when performing the laminectomy and suturing the animal after the procedure has been performed so that the spinal cord does not suffer any additional compressive forces that may increase the injury severity and confound future experiments.
With proper training and practice, the calibrated forceps model of compression injury is well suited for performing SCI in mice that mimic clinical cases observed in humans2,3,7. Due to the ease of creating forceps, producing mice of differing degrees of injury severity can be easily done. This will be of great benefit for observing genetic effects on SCI of differing degrees of severity in transgenic mice as well as evaluating the efficacy of stem cell transplantations in mice. The majority of studies in the literature have been performed on rats due to their size, which generally makes surgeries easier to perform. However the method published by Plemel et al.7 and described by us in this video should enable SCI to be performed on mice with great ease and reproducibility.

Animal models of SCI are valuable tools for assessing the efficacy of therapeutic paradigms designed to mitigate damage as a consequence of trauma to the spinal cord. Out of experimental necessity, these models must provide reproducible deficits in locomotor and sensory behaviors, be adjustable to produce injuries of differing severity, and demonstrate that injury severity correlates with the degree of neurological deficit observed. There are three main types of SCI with distinct features of injury: transection, contusion, and compression1. Briefly, a transection injury is a laceration to the spinal cord, a contusion injury arises from a brief, focal force applied to the dorsal spinal cord, and a compression injury occurs when an injurious force is applied to the spinal cord, and may also be referred to as a crush injury2.
Complete transection injuries are clinically rare in humans3, while contusion and compression injuries are more common. The compression injury produces an outcome similar to what is found in human SCI caused by, for example, tumor compression or other injurious compressive forces, and can be performed using a simple array of tools. Contusion and compression injuries are similar in that both are a compressive force and both have similar pathological features, such as cytoarchitectonic disorganization, and evoke similar endogenous responses to injury1,4. The contusion injury model usually applies this force to the dorsal spinal cord using a special apparatus in a manner similar to human cases of SCI resulting from an impaction of the spinal column2,5,6. In contrast, compression injuries can be generated by a variety of methods applying force dorsally or laterally. Methods of a compression injury include calibrated forceps7, aneurysm clips2, or placing a weight directly onto the spinal cord8. An advantage of the aneurysm clips is that they are able to provide different amounts of force9. The method of adding weights to the dorsal surface of the spinal cord directly8 requires the weight to be in place for 10 min, drastically increasing the length of surgery and resulting in inconsistencies due to placement of the weight and movement due to the respiration of the animal. Due to the small size of mice, situating animals in specialized apparatuses designed for use in rats, such as impactors for contusion injuries, may be difficult or result in inconsistent injuries7. However, mice are available in a wide range of transgenic strains, unlike larger animals such as rats or rabbits that are very useful for SCI research.
The Plemel method of using calibrated forceps to compress the spinal cord generates a reproducible SCI with a high degree of correlation between injury severity and neurological deficit7. This surgical SCI model is generated using a pair of No. 5 Dumont forceps modified to be held apart at a defined distance by either metal epoxy or some other obstruction to prevent complete closure. This engineered spacing ensures that the forceps will always close to a certain width in multiple surgeries and by different users. The advantage of the Plemel method is that the materials to produce the calibrated forceps can be easily purchased and assembled in the laboratory without need of specialized equipment. These forceps can withstand multiple rounds of autoclaving and sterilization, and the lack of a separate, bulky apparatus streamlines surgeries.
In this video we demonstrate the surgical use of a pair of calibrated forceps on the mouse spinal cord to generate a compression injury. We also address unique concerns related to the care of spinal cord injured laboratory animals to improve their quality of life post-operatively and reduce mortality.

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	<tbody>
		<tr height="22">
			<td height="22" style="height:22px;width:101px;">Name</td>
			<td style="width:127px;">Company</td>
			<td style="width:123px;">Catalog No.</td>
		</tr>
		<tr height="43">
			<td height="43" style="height:43px;width:101px;">Isoflurane Machine</td>
			<td style="width:127px;">Smiths Medical PM, Inc</td>
			<td style="width:123px;">VCT302</td>
		</tr>
		<tr height="43">
			<td height="43" style="height:43px;width:101px;">Isoflurane</td>
			<td style="width:127px;">Phoenix Pharmaceutical</td>
			<td style="width:123px;">NDC: 66794-013-25</td>
		</tr>
		<tr height="43">
			<td height="43" style="height:43px;width:101px;">Dissecting Scope</td>
			<td style="width:127px;">Seiler Precision Microscopes</td>
			<td style="width:123px;">SSI 202/402</td>
		</tr>
		<tr height="64">
			<td height="64" style="height:64px;width:101px;">Germinator-500 (tool sterilizer)</td>
			<td style="width:127px;">Thomas Scientific</td>
			<td style="width:123px;">3885A20</td>
		</tr>
		<tr height="43">
			<td height="43" style="height:43px;width:101px;">Puralube (Eye Ointment)</td>
			<td style="width:127px;">Dechra</td>
			<td style="width:123px;">NDC 17033-211-38</td>
		</tr>
		<tr height="43">
			<td height="43" style="height:43px;width:101px;">Scalpel Handle (#3)</td>
			<td style="width:127px;">Fine Science Tools</td>
			<td style="width:123px;">10003-12</td>
		</tr>
		<tr height="43">
			<td height="43" style="height:43px;width:101px;">Scalpel Blade (#11)</td>
			<td style="width:127px;">Fisher Scientific</td>
			<td style="width:123px;">&#160;08-914B</td>
		</tr>
		<tr height="43">
			<td height="43" style="height:43px;width:101px;">Retractor (Colibri )</td>
			<td style="width:127px;">Fine Science Tools</td>
			<td style="width:123px;">17000-03</td>
		</tr>
		<tr height="64">
			<td height="64" style="height:64px;width:101px;">Friedman Pearson Roungeur</td>
			<td style="width:127px;">Fine Science Tools</td>
			<td style="width:123px;">16021-14</td>
		</tr>
		<tr height="64">
			<td height="64" style="height:64px;width:101px;">Vanna (Castroviejo) Scissors</td>
			<td style="width:127px;">Roboz</td>
			<td style="width:123px;">RS-5658</td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:101px;">Tissue Forceps</td>
			<td style="width:127px;">Fine Science Tools</td>
			<td style="width:123px;">11029-14</td>
		</tr>
		<tr height="64">
			<td height="64" style="height:64px;width:101px;">Laminectomy Forceps (Dumont #2)&#160;</td>
			<td style="width:127px;">Fine Science Tools</td>
			<td style="width:123px;">11223-20</td>
		</tr>
		<tr height="43">
			<td height="43" style="height:43px;width:101px;">Dumont #5 Forceps</td>
			<td style="width:127px;">Fine Science Tools</td>
			<td style="width:123px;">11252-20</td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:101px;">Stapler</td>
			<td style="width:127px;">Fine Science Tools</td>
			<td style="width:123px;">12031-07</td>
		</tr>
		<tr height="43">
			<td height="43" style="height:43px;width:101px;">Staples&#160;&#160;&#160;&#160;&#160;&#160; (wound clips)</td>
			<td style="width:127px;">Reflex7</td>
			<td style="width:123px;">203-1000</td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:101px;">Sutures</td>
			<td style="width:127px;">Henry Schein</td>
			<td style="width:123px;">101-2636</td>
		</tr>
		<tr height="43">
			<td height="43" style="height:43px;width:101px;">Needles (30 G x &#189;)</td>
			<td style="width:127px;">BD Biomedical</td>
			<td style="width:123px;">305106</td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:101px;">Syringe (1 ml)</td>
			<td style="width:127px;">BD Biomedical</td>
			<td style="width:123px;">309659</td>
		</tr>
		<tr height="43">
			<td height="43" style="height:43px;width:101px;">Baytril (enrofloxacin)</td>
			<td style="width:127px;">Bayer</td>
			<td style="width:123px;">NADA 140-913</td>
		</tr>
		<tr height="64">
			<td height="64" style="height:64px;width:101px;">Buprenex (buprenorphine)</td>
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calibrated forceps model of spinal cord compression injury

Compression injuries of the murine spinal cord are valuable animal models for the study of spinal cord injury (SCI) and spinal regenerative therapy. The calibrated forceps model of compression injury is a convenient, low cost, and very reproducible animal model for SCI. We used a pair of modified forceps in accordance with the method published by Plemel et al. (2008) to laterally compress the spinal cord to a distance of 0.35 mm. In this video, we will demonstrate a dorsal laminectomy to expose the spinal cord, followed by compression of the spinal cord with the modified forceps. In the video, we will also address issues related to the care of paraplegic laboratory animals. This injury model produces mice that exhibit impairment in sensation, as well as impaired hindlimb locomotor function. Furthermore, this method of injury produces consistent aberrations in the pathology of the SCI, as determined by immunohistochemical methods. After watching this video, viewers should be able to determine the necessary supplies and methods for producing SCI of various severities in the mouse for studies on SCI and/or treatments designed to mitigate impairment after injury.

We performed a laminectomy on 12 mice (25 - 30 grams) as described above, followed by spinal cord compressions at 0.25 mm (n = 4), 0.35 mm (n = 4) and 0.55 mm (n = 4). We sacrificed animals at three (n = 6) and seven (n = 6) days post injury by intracardiac perfusion. The spinal cord was removed from the spinal column, and the tissue was prepared and processed as described above. Images of whole spinal cord were taken with a Leica EZ4 digital microscope and accompanying software. Images of spinal cord sections were taken at 2X magnification using an Olympus digital microscope and accompanying software.
We found that spinal cord compression produces an injury with the epicenter at the site of the compression (Figure 1). The effects of the injury extend several millimeters in the rostral and caudal directions. The severity of the injury increased as the distance between the spacers decreased (0.25 mm &#62; 0.35 mm &#62; 0.55 mm, Figure 2). Three days after compression there was blood at the epicenter of the injury and surrounding regions that was not present 7 days after injury. The 0.25 mm and 0.35 mm compressions produced a cavity, but not the 0.55 mm model. After 7 days, the dorsal and ventral white matter largely decreased in size at the epicenter, the grey matter organization was highly distorted, and cavitation was persistent. These cytoarchitectonical alterations are translated into motor and sensory alterations in animal behavior evaluated using appropriate tests such as the Basso Mouse Scale for Locomotor Function and the von Frey hair and ethyl chloride tests for sensory function as we demonstrated in previous publications8.
<img alt="Figure 1" src="/files/ftp_upload/52318/52318fig1highres.jpg" width="500" /> 
Figure 1. Representative images of the intact spinal cord both before and after injury. (A) Intact spinal cord. (B) Spinal cord after 0.35 mm compression. Arrows indicate border of injury. Asterisk indentifies epicenter of injury. D = Dorsal, L = Lateral. Scale bar: 0.50 mm. <a href="https://www.jove.com/files/ftp_upload/52318/52318fig1highres.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" src="/files/ftp_upload/52318/52318fig2highres.jpg" width="650" /> 
Figure 2. Representative images of mouse spinal cord before and after compression injury at varying compression widths. (A) Sagittal section of control spinal cord. (B) Coronal section at epicenter of a 0.35 mm SCI 7 days post compression injury (dpi). (C,E,G) Sagittal sections of spinal cords 3 dpi to a width of 0.25, 0.35 or 0.55 mm. (D,F,H) Sagittal sections of H&#38;E stained spinal cords 7 dpi to a width of 0.25, 0.35 or 0.55 mm. Asterisk indentifies epicenter of injury. All sections stained with H&#38;E. D = Dorsal, L = Lateral. Scale bar: 1.25 mm. <a href="https://www.jove.com/files/ftp_upload/52318/52318fig2highres.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Calibrated Forceps Model of Spinal Cord Compression Injury at JoVE.com

Calibrated Forceps Model of Spinal Cord Compression Injury

Spinal cord injury models should be highly reproducible. We demonstrate that the calibrated forceps compression model of spinal cord injury is an easy to use surgical method for generating reproducible injuries to the murine spinal cord.

Compression injuries of the murine spinal cord are valuable animal models for the study of spinal cord injury (SCI) and spinal regenerative therapy. The ...

calibrated-forceps-model-of-spinal-cord-compression-injury

Research

JoVE Journal

Medicine

Acute and Chronic Tactile Sensory Testing after Spinal Cord Injury in Rats

Spinal cord injury (SCI) impairs sensory systems causing allodynia1-8. To identify cellular and molecular causes of allodynia, sensitive and valid sensory testing in rat SCI models is needed. However, until recently, no single testing approach had been validated for SCI so that standardized methods have not been implemented across labs. Additionally, available testing methods could not be implemented acutely or when severe motor impairments existed, preventing studies of the development of SCI-induced allodynia3. Here we present two validated sensory testing methods using von Frey Hair (VFH) monofilaments which quantify changes in tactile sensory thresholds after SCI4-5. One test is the well-established Up-Down test which demonstrates high sensitivity and specificity across different SCI severities when tested chronically5. The other test is a newly-developed dorsal VFH test that can be applied acutely after SCI when allodynia develops, prior to motor recovery4-5. Each VFH monofilament applies a calibrated force when touched to the skin of the hind paw until it bends. In the up-down method, alternating VFHs of higher or lower forces are used on the plantar L5 dermatome to delineate flexor withdrawal thresholds. Successively higher forces are applied until withdrawal occurs then lower force VFHs are used until withdrawal ceases. The tactile threshold reflects the force required to elicit withdrawal in 50% of the stimuli. For the new test, each VFH is applied to the dorsal L5 dermatome of the paw while the rat is supported by the examiner. The VFH stimulation occurs in ascending order of force until at least 2 of 3 applications at a given force produces paw withdrawal. Tactile sensory threshold is the lowest force to elicit withdrawal 66% of the time. Acclimation, testing and scoring procedures are described. Aberrant trials that require a retest and typical trials are defined. Animal use was approved by Ohio State University Animal Care and Use Committee.

We describe two tactile sensory testing methods for acute or chronic periods of spinal cord injury in rats. These validated procedures can detect the development and maintenance of allodynia-like sensations.

Spinal cord injury (SCI) impairs sensory systems causing allodynia1-8. To identify cellular and molecular causes of allodynia, sensitive and valid sensory ...

A Contusive Model of Unilateral Cervical Spinal Cord Injury Using the Infinite Horizon Impactor

While the majority of human spinal cord injuries occur in the cervical spinal cord, the vast majority of laboratory research employs animal models of spinal cord injury (SCI) in which the thoracic spinal cord is injured. Additionally, because most human cord injuries occur as the result of blunt, non-penetrating trauma (e.g. motor vehicle accident, sporting injury) where the spinal cord is violently struck by displaced bone or soft tissues, the majority of SCI researchers are of the opinion that the most clinically relevant injury models are those in which the spinal cord is rapidly contused.1 Therefore, an important step in the preclinical evaluation of novel treatments on their way to human translation is an assessment of their efficacy in a model of contusion SCI within the cervical spinal cord. Here, we describe the technical aspects and resultant anatomical and behavioral outcomes of an unilateral contusive model of cervical SCI that employs the Infinite Horizon spinal cord injury impactor.
Sprague Dawley rats underwent a left-sided unilateral laminectomy at C5. To optimize the reproducibility of the biomechanical, functional, and histological outcomes of the injury model, we contused the spinal cords using an impact force of 150 kdyn, an impact trajectory of 22.5&deg; (animals rotated at 22.5&deg;), and an impact location off of midline of 1.4 mm. Functional recovery was assessed using the cylinder rearing test, horizontal ladder test, grooming test and modified Montoya's staircase test for up to 6 weeks, after which the spinal cords were evaluated histologically for white and grey matter sparing.
The injury model presented here imparts consistent and reproducible biomechanical forces to the spinal cord, an important feature of any experimental SCI model. This results in discrete histological damage to the lateral half of the spinal cord which is largely contained to the ipsilateral side of injury. The injury is well tolerated by the animals, but does result in functional deficits of the forelimb that are significant and sustained in the weeks following injury. The cervical unilateral injury model presented here may be a resource to researchers who wish to evaluate potentially promising therapies prior to human translation.

A reliable and repeatable way to produce a cervical unilateral spinal cord injury using the Infinite Horizon impactor is described. The method takes advantage of a custom designed frame and clamp to stabilize the spine. The standardized procedure and biomechanical injury parameters result in sufficient and sustained injuries.

While the majority of human spinal cord injuries occur in the cervical spinal cord, the vast majority of laboratory research employs animal models of ...

A Contusion Model of Severe Spinal Cord Injury in Rats

The translational potential of novel treatments should be investigated in severe spinal cord injury (SCI) contusion models. A detailed methodology is described to obtain a consistent model of severe SCI. Use of a stereotactic frame and computer controlled impactor allows for creation of reproducible injury. Hypothermia and urinary tract infection pose significant challenges in the post-operative period. Careful monitoring of animals with daily weight recording and bladder expression allows for early detection of post-operative complications. The functional results of this contusion model are equivalent to transection models. The contusion model can be utilized to evaluate the efficacy of both neuroprotective and neuroregenerative approaches.

A contusion model of severe spinal cord injury is described. Detailed pre-operative, operative and post-operative steps are described to obtain a consistent model.

The translational potential of novel treatments should be investigated in severe spinal cord injury (SCI) contusion models. A detailed methodology is ...

Evaluation of Respiratory Muscle Activation Using Respiratory Motor Control Assessment (RMCA) in Individuals with Chronic Spinal Cord Injury

During breathing, activation of respiratory muscles is coordinated by integrated input from the brain, brainstem, and spinal cord. When this coordination is disrupted by spinal cord injury (SCI), control of respiratory muscles innervated below the injury level is compromised1,2 leading to respiratory muscle dysfunction and pulmonary complications. These conditions are among the leading causes of death in patients with SCI3. Standard pulmonary function tests that assess respiratory motor function include spirometrical and maximum airway pressure outcomes: Forced Vital Capacity (FVC), Forced Expiratory Volume in one second (FEV1), Maximal Inspiratory Pressure (PImax) and Maximal Expiratory Pressure (PEmax)4,5. These values provide indirect measurements of respiratory muscle performance6. In clinical practice and research, a surface electromyography (sEMG) recorded from respiratory muscles can be used to assess respiratory motor function and help to diagnose neuromuscular pathology. However, variability in the sEMG amplitude inhibits efforts to develop objective and direct measures of respiratory motor function6. Based on a multi-muscle sEMG approach to characterize motor control of limb muscles7, known as the voluntary response index (VRI)8, we developed an analytical tool to characterize respiratory motor control directly from sEMG data recorded from multiple respiratory muscles during the voluntary respiratory tasks. We have termed this the Respiratory Motor Control Assessment (RMCA)9. This vector analysis method quantifies the amount and distribution of activity across muscles and presents it in the form of an index that relates the degree to which sEMG output within a test-subject resembles that from a group of healthy (non-injured) controls. The resulting index value has been shown to have high face validity, sensitivity and specificity9-11. We showed previously9 that the RMCA outcomes significantly correlate with levels of SCI and pulmonary function measures. We are presenting here the method to quantitatively compare post-spinal cord injury respiratory multi-muscle activation patterns to those of healthy individuals.

Evaluation of Respiratory Muscle Activation Using Respiratory Motor Control Assessment RMCA in Individuals with Chronic Spinal Cord Injury

The purpose of this publication is to present our original work on a multi-muscle surface electromyographic approach to quantitatively characterize respiratory muscle activation patterns in individuals with chronic spinal cord injury using vector-based analysis.

During breathing, activation of respiratory muscles is coordinated by integrated input from the brain, brainstem, and spinal cord. When this coordination ...

A Murine Model of Cervical Spinal Cord Injury to Study Post-lesional Respiratory Neuroplasticity

A cervical spinal cord injury induces permanent paralysis, and often leads to respiratory distress. To date, no efficient therapeutics have been developed to improve/ameliorate the respiratory failure following high cervical spinal cord injury (SCI). Here we propose a murine pre-clinical model of high SCI at the cervical 2 (C2) metameric level to study diverse post-lesional respiratory neuroplasticity. The technique consists of a surgical partial injury at the C2 level, which will induce a hemiparalysis of the diaphragm due to a deafferentation of the phrenic motoneurons from the respiratory centers located in the brainstem. The contralateral side of the injury remains intact and allows the animal recovery. Unlike other SCIs which affect the locomotor function (at the thoracic and lumbar level), the respiratory function does not require animal motivation and the quantification of the deficit/recovery can be easily performed (diaphragm and phrenic nerve recordings, whole body ventilation). This pre-clinical C2 SCI model is a powerful, useful, and reliable pre-clinical model to study various respiratory and non-respiratory neuroplasticity events at different levels (molecular to physiology) and to test diverse putative therapeutic strategies which might improve the respiration in SCI patients.

Respiratory failure is the leading cause of death following a cervical spinal cord injury. Having a reproducible, quantifiable, and reliable pre-clinical animal model of respiratory failure induced by a partial cervical injury will help to understand the subsequent respiratory and non-respiratory neuroplasticity and allow testing putative repair strategies.

A cervical spinal cord injury induces permanent paralysis, and often leads to respiratory distress. To date, no efficient therapeutics have been developed ...

Controlled Cortical Impact Model for Traumatic Brain Injury

Every year over a million Americans suffer a traumatic brain injury (TBI). Combined with the incidence of TBIs worldwide, the physical, emotional, social, and economical effects are staggering. Therefore, further research into the effects of TBI and effective treatments is necessary. The controlled cortical impact (CCI) model induces traumatic brain injuries ranging from mild to severe. This method uses a rigid impactor to deliver mechanical energy to an intact dura exposed following a craniectomy. Impact is made under precise parameters at a set velocity to achieve a pre-determined deformation depth. Although other TBI models, such as weight drop and fluid percussion, exist, CCI is more accurate, easier to control, and most importantly, produces traumatic brain injuries similar to those seen in humans. However, no TBI model is currently able to reproduce pathological changes identical to those seen in human patients. The CCI model allows investigation into the short-term and long-term effects of TBI, such as neuronal death, memory deficits, and cerebral edema, as well as potential therapeutic treatments for TBI.

Traumatic brain injuries (TBIs) remain a serious health problem. Using the controlled cortical impact surgery model, research on the effects of TBI and possible treatment methods may be performed.

Every year over a million Americans suffer a traumatic brain injury (TBI).  Combined with the incidence of TBIs worldwide, the physical, emotional, ...

A Radio-telemetric System to Monitor Cardiovascular Function in Rats with Spinal Cord Transection and Embryonic Neural Stem Cell Grafts

High thoracic or cervical spinal cord injury (SCI) can lead to cardiovascular dysfunction. To monitor cardiovascular parameters, we implanted a catheter connected to a radio transmitter into the femoral artery of rats that underwent a T4 spinal cord transection with or without grafting of embryonic brainstem-derived neural stem cells expressing green fluorescent protein. Compared to other methods such as cannula insertion or tail-cuff, telemetry is advantageous to continuously monitor blood pressure and heart rate in freely moving animals. It is also capable of long term multiple data acquisitions. In spinal cord injured rats, basal cardiovascular data under unrestrained condition and autonomic dysreflexia in response to colorectal distension were successfully recorded. In addition, cardiovascular parameters before and after SCI can be compared in the same rat if a transmitter is implanted before a spinal cord transection. One limitation of the described telemetry procedure is that implantation in the femoral artery may influence the blood supply to the ipsilateral hindlimb.

We present a protocol for using a radio-telemetric system to record cardiovascular parameters in T4 spinal cord transected rats eight weeks after embryonic brainstem neural stem cell grafting into the lesion site. Telemetry is an advanced technique to accurately evaluate cardiovascular function in conscious freely moving spinal cord injured rats.

High thoracic or cervical spinal cord injury (SCI) can lead to cardiovascular dysfunction. To monitor cardiovascular parameters, we implanted a catheter ...

Neural Stem Cell Transplantation in Experimental Contusive Model of Spinal Cord Injury

Spinal cord injury is a devastating clinical condition, characterized by a complex of neurological dysfunctions. Animal models of spinal cord injury can be used both to investigate the biological responses to injury and to test potential therapies. Contusion or compression injury delivered to the surgically exposed spinal cord are the most widely used models of the pathology. In this report the experimental contusion is performed by using the Infinite Horizon (IH) Impactor device, which allows the creation of a reproducible injury animal model through definition of specific injury parameters. Stem cell transplantation is commonly considered a potentially useful strategy for curing this debilitating condition. Numerous studies have evaluated the effects of transplanting a variety of stem cells. Here we demonstrate an adapted method for spinal cord injury followed by tail vein injection of cells in CD1 mice. In short, we provide procedures for: i) cell labeling with a vital tracer, ii) pre-operative care of mice, iii) execution of a contusive spinal cord injury, and iv) intravenous administration of post mortem neural precursors. This contusion model can be utilized to evaluate the efficacy and safety of stem cell transplantation in a regenerative medicine approach.

Spinal cord injury is a traumatic condition that causes severe morbidity and high mortality. In this work we describe in detail a contusion model of spinal cord injury in mice followed by a transplantation of neural stem cells.

Spinal cord injury is a devastating clinical condition, characterized by a complex of neurological dysfunctions. Animal models of spinal cord injury can ...

Contrast Enhanced Ultrasound Imaging for Assessment of Spinal Cord Blood Flow in Experimental Spinal Cord Injury

Reduced spinal cord blood flow (SCBF) (i.e., ischemia) plays a key role in traumatic spinal cord injury (SCI) pathophysiology and is accordingly an important target for neuroprotective therapies. Although several techniques have been described to assess SCBF, they all have significant limitations. To overcome the latter, we propose the use of real-time contrast enhanced ultrasound imaging (CEU). Here we describe the application of this technique in a rat contusion model of SCI. A jugular catheter is first implanted for the repeated injection of contrast agent, a sodium chloride solution of sulphur hexafluoride encapsulated microbubbles. The spine is then stabilized with a custom-made 3D-frame and the spinal cord dura mater is exposed by a laminectomy at ThIX-ThXII. The ultrasound probe is then positioned at the posterior aspect of the dura mater (coated with ultrasound gel). To assess baseline SCBF, a single intravenous injection (400 &#181;l) of contrast agent is applied to record its passage through the intact spinal cord microvasculature. A weight-drop device is subsequently used to generate a reproducible experimental contusion model of SCI. Contrast agent is re-injected 15 min following the injury to assess post-SCI SCBF changes. CEU allows for real time and in-vivo assessment of SCBF changes following SCI. In the uninjured animal, ultrasound imaging showed uneven blood flow along the intact spinal cord. Furthermore, 15 min post-SCI, there was critical ischemia at the level of the epicenter while SCBF remained preserved in the more remote intact areas. In the regions adjacent to the epicenter (both rostral and caudal), SCBF was significantly reduced. This corresponds to the previously described &#8220;ischemic penumbra zone&#8221;. This tool is of major interest for assessing the effects of therapies aimed at limiting ischemia and the resulting tissue necrosis subsequent to SCI.

Contrast Enhanced Ultrasound imaging is a reliable in-vivo tool for quantifying spinal cord blood flow in an experimental rat spinal cord injury model. This paper contains a comprehensive protocol for application of this technique in association with a contusion model of thoracic spinal cord injury.

Reduced spinal cord blood flow (SCBF) (i.e., ischemia) plays a key role in traumatic spinal cord injury (SCI) pathophysiology and is accordingly an ...

Photothrombosis-induced Focal Ischemia as a Model of Spinal Cord Injury in Mice

Spinal cord injury (SCI) is a devastating clinical condition causing permanent changes in sensorimotor and autonomic functions of the spinal cord (SC) below the site of injury. The secondary ischemia that develops following the initial mechanical insult is a serious complication of the SCI and severely impairs the function and viability of surviving neuronal and non-neuronal cells in the SC. In addition, ischemia is also responsible for the growth of lesion during chronic phase of injury and interferes with the cellular repair and healing processes. Thus there is a need to develop a spinal cord ischemia model for studying the mechanisms of ischemia-induced pathology. Focal ischemia induced by photothrombosis (PT) is a minimally invasive and very well established procedure used to investigate the pathology of ischemia-induced cell death in the brain. Here, we describe the use of PT to induce an ischemic lesion in the spinal cord of mice. Following retro-orbital sinus injection of Rose Bengal, the posterior spinal vein and other capillaries on the dorsal surface of SC were irradiated with a green light resulting in the formation of a thrombus and thus ischemia in the affected region. Results from histology and immunochemistry studies show that PT-induced ischemia caused spinal cord infarction, loss of neurons and reactive gliosis. Using this technique a highly reproducible and relatively easy model of SCI in mice can be achieved that would serve the purpose of scientific investigations into the mechanisms of ischemia induced cell death as well as the efficacy of neuroprotective drugs. This model will also allow exploration of the pathological changes that occur following SCI in live mice like axonal degeneration and regeneration, neuronal and astrocytic Ca2+ signaling using two-photon microscopy.

Photothrombosis is a minimally invasive and highly reproducible procedure to induce focal ischemia in the spinal cord and serves as a model of spinal cord injury in mice.

Spinal cord injury (SCI) is a devastating clinical condition causing permanent changes in sensorimotor and autonomic functions of the spinal cord (SC) ...