performing thoracic spinal cord hemisection surgery in rats

Performing Thoracic Spinal Cord Hemisection Surgery in Rats

After confirming a lack of response to toe pinch in an anesthetized adult rat, make a 2.5-centimeter incision in the shaved skin overlying the T6 to T10 vertebrae, and use blunt scissors to retract the skin and superficial fat. Use blunt dissection scissors and a self-retaining retractor to separate the paravertebral muscles, inserting on the dorsal aspect of the T7 to T9 vertebrae, and use fine forceps and cotton-tipped applicators to debride and clear any remaining tissue to expose the spinous processes and vertebral lamina.
Place the rat under a stereomicroscope, and use delicate bone trimmers to carefully cut the facets bilaterally on the T7 and T8 vertebrae. Use a scalpel to make a 1 millimeter superficial cut in the dorsal connective tissue between the T8 and T9 vertebral laminae, taking care not to injure the underlying cord, and use bone trimmers to remove the spinous process of the T8 vertebra.
With curved hemostatic forceps, carefully clamped on the T7 spinous process, rotate the caudal end of the T8 laminae slightly rostrally approximately 20 degrees, and insert the bone trimmers under the T8 lamina. Make a midline cut extending along the lamina, continuing the laminectomy by repeating the cuts on the left and right side of the vertebral lamina, medial to the transverse processes to expose the spinal cord.
Drip 100 microliters of 2% lidocaine into the exposed spinal canal, and use fine forceps and iridectomy scissors to remove the dura, overlaying the T8 spinal segment. Repeat the lidocaine administration to the exposed cord and identified the midline of the cord by visualization of a center line created between the spinous processes extending between the exposed T7 to T9 vertebra.
Using fine forceps to stabilize the spinal cord, use a dissecting knife to hemisect the spinal cord from the midline toward one side of the animal, taking care not to cut through the anterior spinal artery on the ventral side. Using iridectomy scissors, carefully cut through any remaining tissue on the lesion side of the spinal cord to ensure the ventrolateral quadrant is appropriately transected, and place a sterile, approximately 6 by 2-millimeter saline soaked hemostatic sponge into the exposed cavity above the spinal cord.
Then use 4-0 polyglactin 910 sutures to close muscle layers and the skin around the incision site and place the rat in a warm environment under a heat lamp with monitoring until full recovery.

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All procedures involving animal models have been reviewed by the local institutional animal care committee and the JoVE veterinary review board.
1. Thoracic hemisection surgery
<ol>
	<li>Wear appropriate protective equipment (gloves, mask, and gown) to maintain an aseptic environment for surgery. Clean the surgical area with alcohol wipes, and place sterile surgical drapes over the surgical field. Sterilize surgical tools and place on the surgical field.</li>
	<li>Anesthetize the rat under a mixture of isoflurane gas (3% induction, 0.5&#8722;3% maintenance) and oxygen (1 L/min). Confirm proper surgical anesthetic depth by verifying the absence of toe pinch&#160;and corneal reflex responses. Continuously monitor the rat during the entire procedure and adjust the amount of anesthetic delivery as required to maintain surgical anesthetic depth.</li>
	<li>Shave the dorsal trunk between the hip and the neck, place the rat on the surgical field, disinfect the incision site with alcohol wipes and proviodine solution, and maintain core body temperature at 37 &#176;C using a feedback-controlled heating pad monitored by rectal thermometer.</li>
	<li>Place ophthalmic ointment on the eyes to keep them hydrated and reapply throughout surgery as required.</li>
	<li>Make a 2.5 cm incision in the skin overlaying the T6&#8722;T10 vertebrae with a scalpel. Retract the skin and superficial fat using blunt dissection scissors. 
	NOTE: The T6&#8722;T10 vertebral segments can be identified either rostrally by gentle palpation of the dorsal spinal segments from the base of the skull starting from the noticeable protuberance of the 2nd&#160;thoracic vertebra or caudally by palpation of the most posterior floating rib, which will induce movement in the 13th&#160;thoracic vertebrae.</li>
	<li>Separate the paravertebral muscles inserting on the dorsal aspect of the T7&#8722;T9 vertebrae using blunt dissection scissors and a self-retaining retractor. Debride and clear any remaining tissue using fine forceps and cotton tipped applicators to expose the spinous processes and vertebral laminae. 
	NOTE: This and the following steps are greatly aided by microscopic visualization (~5&#8722;15x).</li>
	<li>Carefully cut the facets (zygapophysial joints) bilaterally on the T7 and T8 vertebrae with delicate bone trimmers. Cut the dorsal connective tissue between the T8 and T9 vertebral laminae superficially with a scalpel (1 mm depth) being careful not to injure the underlying cord.</li>
	<li>Remove the spinous process of the T8 vertebra with bone trimmers. With curved hemostatic forceps carefully clamped on the T7 spinous process, rotate the caudal end of the T8 laminae slightly rostrally (~20&#176;), insert the bone trimmers under the T8 lamina, and make a midline cut extending along the lamina. Continue the laminectomy by repeating the cuts on the left and right side of the vertebral lamina medial to the transverse processes to expose the spinal cord. 
	NOTE: Be careful to remove all bone fragments created from the laminectomy.</li>
	<li>Drip lidocaine (2%, 0.1 mL) in the exposed spinal canal and remove the dura overlaying the T8 spinal segment using fine forceps and iridectomy scissors. Repeat lidocaine administration to the exposed cord and identify the midline of the cord by visualization of a center line created between the spinous processes extending between the exposed T7&#8722;T9 vertebra. 
	NOTE: Along with the spinous processes on T7 and T9, the exposed dorsal root ganglia on T8 can also be used to aid in the identification of the midline. A 30 G needle can be placed in the midline of the cord to aid with the subsequent hemisection.</li>
	<li>Hemisect the spinal cord from the midline towards the one side with a dissecting knife. Be careful not to cut through the anterior spinal artery on the ventral side (do not apply firm pressure to the vertebral body). Using iridectomy scissors, carefully cut through any remaining tissue on the lesioned side of the spinal cord to ensure the ventrolateral quadrant is appropriately transected.</li>
	<li>Place a sterile saline-soaked hemostatic sponge (~6 x 2 mm) in the exposed cavity above the spinal cord and suture the muscle layers (4-0 polyglactin 910). Next, suture the skin around the incision site.</li>
	<li>Provide adequate analgesic (buprenorphine 0.05 mg/kg subcutaneous [s.c.]), antibiotic (enrofloxacin, 10 mg/kg s.c.), and replenish lost fluids with 5 cc lactated ringer&#39;s solution (intraperitoneal [i.p.]) immediately after surgery.</li>
	<li>Remove the rat from anesthesia. Place the rat in a warm environment under a heating pad or lamp (~ 33 &#176;C) until the animal is fully awake.</li>
	<li>Provide supplemental analgesia daily over the first 3 post-surgical days and continually monitor for signs of pain, weight loss, improper micturition, infection, problems with wound healing, or autophagia.</li>
</ol>

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Baytril</td>
			<td>CDMV</td>
			<td>11242</td>
			<td></td>
		</tr>
		<tr>
			<td>Blunt dissection scissors</td>
			<td>World Precision Instruments</td>
			<td>503669</td>
			<td></td>
		</tr>
		<tr>
			<td>Buprenorphine hydrochoride</td>
			<td>CDMV</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Compressed oxygen gas</td>
			<td>Praxair</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Cotton tipped applicators</td>
			<td>CDMV</td>
			<td>108703</td>
			<td></td>
		</tr>
		<tr>
			<td>Delicate bone trimmers</td>
			<td>Fine Science Tools</td>
			<td>16109-14</td>
			<td></td>
		</tr>
		<tr>
			<td>Dissecting knife</td>
			<td>Fine Science Tools</td>
			<td>10055-12</td>
			<td></td>
		</tr>
		<tr>
			<td>Dumont fine forceps (#5)</td>
			<td>Fine Science Tools</td>
			<td>11254-20</td>
			<td></td>
		</tr>
		<tr>
			<td>Ethicon Vicryl 4/0 Violet Braided FS-2&#160; suture (J392H)</td>
			<td>CDMV</td>
			<td>111689</td>
			<td></td>
		</tr>
		<tr>
			<td>Feedback-controlled heating pad</td>
			<td>Harvard Apparatus</td>
			<td>55-7020</td>
			<td></td>
		</tr>
		<tr>
			<td>Female Long-Evans rats</td>
			<td>Charles River Laboratories</td>
			<td>Strain code: 006</td>
			<td>225-250g</td>
		</tr>
		<tr>
			<td>Gelfoam</td>
			<td>CDMV</td>
			<td>102348</td>
			<td></td>
		</tr>
		<tr>
			<td>Curved hemostat forceps</td>
			<td>Fine Science Tools</td>
			<td>13003-10</td>
			<td></td>
		</tr>
		<tr>
			<td>Hot bead sterilizer</td>
			<td>Fine Science Tools</td>
			<td>18000-45</td>
			<td></td>
		</tr>
		<tr>
			<td>Hydrogel</td>
			<td>70-01-5022</td>
			<td>Clear H20</td>
			<td></td>
		</tr>
		<tr>
			<td>Isofluorane</td>
			<td>CDMV</td>
			<td>118740</td>
			<td></td>
		</tr>
		<tr>
			<td>Lactated Ringer&#39;s solution</td>
			<td>CDMV</td>
			<td>116373</td>
			<td></td>
		</tr>
		<tr>
			<td>Lidocaine (2%)</td>
			<td>CDMV</td>
			<td>123684</td>
			<td></td>
		</tr>
		<tr>
			<td>Needle 30 ga</td>
			<td>CDMV</td>
			<td>4799</td>
			<td></td>
		</tr>
		<tr>
			<td>Opthalmic ointment</td>
			<td>CDMV</td>
			<td>110704</td>
			<td></td>
		</tr>
		<tr>
			<td>Physiological saline</td>
			<td>CDMV</td>
			<td>1399</td>
			<td></td>
		</tr>
		<tr>
			<td>Proviodine</td>
			<td>CDMV</td>
			<td>4568</td>
			<td></td>
		</tr>
		<tr>
			<td>Rodent Liquid Diet</td>
			<td>Bioserv</td>
			<td>F1268</td>
			<td></td>
		</tr>
		<tr>
			<td>Scalpal blade #11</td>
			<td>CDMV</td>
			<td>6671</td>
			<td></td>
		</tr>
		<tr>
			<td>Self-retaining retractor</td>
			<td>World Precision Instruments</td>
			<td>14240</td>
			<td></td>
		</tr>
		<tr>
			<td>Vannas iridectomy spring scissors</td>
			<td>Fine Science Tools</td>
			<td>15002-08</td>
			<td></td>
		</tr>
		<tr>
			<td>Veterinary Anesthesia Machine and isofluarane vaporizer</td>
			<td>Dispomed</td>
			<td>975-0510-000</td>
			<td></td>
		</tr>
	</tbody>
</table>

Source: Brown, A. R., et al., <a href="https://app.jove.com/v/59738">Thoracic Spinal Cord Hemisection Surgery and Open-Field Locomotor Assessment in the Rat.</a>&#160;J. Vis. Exp.&#160;(2019).
This video demonstrates the surgical procedure for performing a thoracic spinal cord hemisection in rats. It outlines the steps to expose the T8 spinal cord segment, perform the hemisection to disrupt neural tracts on one side, and provide post-operative care.

Source: Brown, A. R., et al., Thoracic Spinal Cord Hemisection Surgery and Open-Field Locomotor Assessment in the Rat.&#160;J. Vis. Exp.&#160;(2019).
This ...

Secure an anesthetized rat with a shaved dorsal trunk in a stereotaxic frame.
Disinfect the shaved area and cover the rat with a drape.
Make an incision and retract the skin and subcutaneous fat over the thoracic T6 to T10 vertebrae.
Separate the paravertebral muscles to expose the T7 to T9 vertebrae.
Bilaterally cut the facet joints on the T7 and T8 vertebrae.
Superficially cut the dorsal connective tissue between the T8 and T9 vertebrae.
Remove the T8 spinous process and slightly rotate its lamina.
Cut the lamina on both sides, exposing the spinal cord.
Apply anesthetic and remove the dura covering the T8 spinal segment.
Identify the spinal cord midline and cut toward one side, without damaging the anterior spinal artery.
This hemisection disrupts the neural tracts on one side.
Place a saline-soaked absorbent sponge to control bleeding.
Suture the incision and allow the rat to recover.

9 Views. Performing Thoracic Spinal Cord Hemisection Surgery in Rats

Video: Performing Thoracic Spinal Cord Hemisection Surgery in Rats - Experiment

Laminectomy and Spinal Cord Window Implantation in the Mouse

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

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

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

JoVE Journal

Spinal Cord Lateral Hemisection and Asymmetric Behavioral Assessments in Adult Rats

Incomplete spinal cord injury (SCI) often leads to impairments of sensorimotor functions and is clinically the most frequent type of SCI. Human Brown-S&#233;quard syndrome&#160;is a common type of incomplete SCI caused by a lesion to one half of the spinal cord which results in paralysis and loss of&#160;proprioception&#160;on the same (or ipsilesional) side as the injury, and loss of pain and temperature sensation on the opposite (or contralesional) side. Adequate methodologies for producing a spinal cord lateral hemisection (HX) and assessing neurological impairments are essential to establish a reliable animal model of Brown-S&#233;quard syndrome. Although lateral hemisection model plays a pivotal role in basic and translational research, standardized protocols for creating such a hemisection and assessing unilateralized function are lacking. The goal of this study is to describe step-by-step procedures to produce a rat spinal lateral HX at the 9th thoracic (T9) vertebral level. We, then, describe a combined behavior scale for HX (CBS-HX) that provides a simple and sensitive assessment of asymmetric neurological performance for unilateral SCI. The CBS-HX, ranging from 0 to 18, is composed of 4 individual assessments which include unilateral hindlimb stepping (UHS), coupling, contact placing, and grid walking. For CBS-HX, the ipsilateral and contralateral hindlimbs are assessed separately. We found that, after a T9 HX, the ipsilateral hindlimb showed impaired behavior function whereas the contralateral hindlimb showed substantial recovery. The CBS-HX effectively discriminated behavioral functions between ipsilateral and contralateral hindlimbs and detected temporal progression of recovery of the ipsilateral hindlimb. The CBS-HX components can be analyzed separately or in combination with other measures when needed. Although we only provided visual descriptions of the surgical procedures and behavioral assessments of a thoracic HX, the principle may be applied to other incomplete SCIs and at other levels of the injury.

Here we describe surgical procedures to produce a reliable spinal cord lateral hemisection (HX) at the 9th thoracic level in adult rats and neurobehavioral assessments designed for detecting asymmetric deficits after such a unilateral injury.

Incomplete spinal cord injury (SCI) often leads to impairments of sensorimotor functions and is clinically the most frequent type of SCI. Human ...

Medicine

A Minimally Invasive, Fast Spinal Cord Lateral Hemisection Technique for Modeling Open Spinal Cord Injuries in Rats

Open spinal cord injury techniques modeling laceration-like injuries are time-consuming and invasive because they involve laminectomy. This new technique eliminates laminectomy by removing two spinous processes and lifting, then tilting the caudal vertebral arch. The surgical area opens up without the need for laminectomy. Lateral hemisection is then performed with direct visible control under a microscope. The trauma is minimized, requiring only a small bone wound. 
This technique has several advantages: it is faster and, therefore, less of a burden for the animal, and the bone wound is smaller. Because the laminectomy is eliminated, there is less chance for unwanted injury to the spinal cord, and there are no bone splinters that can cause problems (bone splinters embedded in the spinal cord can cause swelling and secondary damage). The vertebral canal remains intact. The main limitation is that the hemisection can only be performed in the intervertebral spaces. 
The results show that this technique can be performed much faster than the traditional surgical approach, using laminectomy (11 min vs. 35 min). This technique can be useful for researchers working with animal models of open spinal cord injury as it is widely adaptable and does not require any additional specialized instrumentation.

Here, we describe a new, fast technique modeling open spinal cord injury in rats that eliminates laminectomy. Lateral hemisection is performed while viewing through a microscope. The technique is versatile and can also be used in the cervical, thoracic, and lumbar regions of the spinal cord of other animals.

Open spinal cord injury techniques modeling laceration-like injuries are time-consuming and invasive because they involve laminectomy. This new technique ...

Performing Thoracic Spinal Cord Hemisection Surgery in Rats

Transcript

Performing Thoracic Spinal Cord Hemisection Surgery in Rats