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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. Induce anesthesia
<ol>
	<li>Anesthetize the mouse with ketamine (125 mg/kg) and xylazine (10 mg/kg) injected intraperitoneally.</li>
</ol>
2. Vital signs monitoring every 15 min
<ol>
	<li>Monitor temperature, respiratory rate, and skin color. The mouse should feel warm to the touch. The skin should appear pink and well perfused. The respiratory rate should range 50&#8211;70 breaths per minute.</li>
</ol>
3. Pre-surgical procedures
<ol>
	<li>Weigh all the mice on the day prior to injury induction.</li>
	<li>Sterilize one set of surgical instruments by autoclaving for each experimental subject. Sterilize the impacting device before use.&#160;</li>
	<li>Prepare a recovery cage by placing a clean cage over an electric heating pad set to &#34;low&#34; setting and positioned in a manner such that the mice can move away from the heat once ambulatory.</li>
	<li>Set up the operating theater within a sterilized laminar flow hood.
	<ol>
		<li>Position the stereotaxic operating frame.</li>
		<li>Attach the impacting device to the stereotaxic frame.</li>
		<li>Set the actuating device with the desired biomechanical parameters for velocity and dwell time. 
		NOTE: In this protocol a severe brain injury is described utilizing a 3 mm diameter impact tip via a 5 mm diameter craniectomy with the velocity set at 2.5 m/s and a dwell time of 0.1 s. A wide range of biomechanical parameters may be used to induce the full spectrum of TBI.</li>
	</ol>
	</li>
	<li>Don new personal protective equipment and sterile gloves.</li>
	<li>Shave the fur from the operative site using electric clippers.</li>
	<li>Apply protective opthalmic ointment to the eyes of the mouse to prevent corneal injury and drying.</li>
	<li>Place the mouse into the operating theater.</li>
	<li>Prep the skin with an iodine based surgical scrub alternated with alcohol three times.</li>
</ol>
4. Application of controlled cortical impact
<ol>
	<li>Incise the scalp 1 cm in the midline with a scalpel exposing the skull.</li>
	<li>Position the mouse within a stereotaxic operating frame by securing the bilateral temporal bones between miniature ear bars and locking the incisors within an incisor clamp creating a stable three-point-hold on the mouse head.</li>
	<li>Retract the scalp away from the operative site with a hemostat or locking forceps to ensure the scalp does not come in contact with the drill bit during craniectomy.</li>
	<li>Identify the sagittal and coronal sutures on the exposed skull. 
	NOTE: This protocol centers the craniectomy 2 mm left of the sagittal suture and 2 mm rostral to the coronal suture.</li>
	<li>Perform a craniectomy using a drill with a trephine drill bit.
	<ol>
		<li>To perform the craniectomy, first activate the drill at maximum speed and then apply the trephine drill bit perpendicular to the skull at the site of craniectomy.</li>
		<li>Apply gentle, even pressure to the drill once contact is made with the skull. A slight &#34;give&#34; will be felt once the drill penetrates through the skull. Do not penetrate the underlying dura. 
		NOTE: This protocol utilizes a 5 mm trephine drill bit to perform the craniectomy.</li>
	</ol>
	</li>
	<li>Use forceps and a small gauge hypodermic needle to remove the bone flap, fully exposing the underlying dura mater.</li>
	<li>Rotate the impactor tip into the operative field and lower it until it makes contact with the exposed dura mater. Once contact is made the instrument&#39;s contact sensor will make an audible tone to alert the surgeon that contact has been made. This will mark the zero point from which the deformation depth is set. 
	NOTE: This protocol utilizes a 3 mm impacting tip to generate a severe injury. Tips as small as 1 mm may be used to apply more localized injury.</li>
	<li>Retract the impacting tip and set the desired impact depth by lowering the impactor position on the stereotaxic frame. 
	NOTE: In this protocol we describe a severe injury by setting the deformation depth to 2 mm.</li>
	<li>Apply the injury by activating impactor on the actuating device.</li>
	<li>Rotate the impact device out of the field and remove the animal from the stereotaxic frame.</li>
</ol>
5. Surgical site closure
<ol>
	<li>Control bleeding from the skull and injured cortical surface with direct pressure from a sterile cotton tipped applicator.</li>
	<li>Dry the skull with a sterile cotton tipped applicator.</li>
	<li>Close the scalp over the craniectomy using a commercially available surgical adhesive or monofilament suture. 
	NOTE: In this protocol a veterinary surgical adhesive is used to close the scalp. The bone flap is not replaced and is discarded.</li>
</ol>
6. Post-operative care and monitoring
<ol>
	<li>Administer post-operative analgesia (e.g., sustained release buprenorphine 0.1&#8211;0.5 mg/kg administered subcutaneously providing 72 h of sustained analgesia).</li>
	<li>Place the animal in the lateral decubitus recovery position in a clean pre-warmed cage.</li>
	<li>Observe the animals until awake and mobile, then return each mouse to its home cage.</li>
	<li>Ensure free access to food and water. Normal food and water intake typically resume within one to two hours after injury.</li>
	<li>Measure body weight every three days throughout the experiment.</li>
</ol>

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
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			<td>AnaSed Injection Xylazine Sterile Solution</td>
			<td>LLOYD, Inc.</td>
			<td>5939911020</td>
			<td></td>
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			<td>Buprenorphine SR Lab 0.5mg/mL</td>
			<td>Zoopharm-Wildlife Pharmaceuticals USA</td>
			<td>BSRLAB0.5-182012</td>
			<td></td>
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			<td>High Speed Rotary Micromotor KiT0</td>
			<td>Foredom Electric Company</td>
			<td>K.1070</td>
			<td></td>
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			<td>Imapact one for Stereotaxix CCI</td>
			<td>Leica Biosystems Nussloch GmbH</td>
			<td>39463920</td>
			<td></td>
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			<td>Ketathesia Ketamine HCl Injection USP</td>
			<td>Henry Schein, Inc</td>
			<td>56344</td>
			<td></td>
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			<td>Mouse Specific Stereotaxic Base</td>
			<td>Leica Biosystems Nussloch GmbH</td>
			<td>39462980</td>
			<td></td>
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			<td>Trephines for Micro Drill</td>
			<td>Fine Science Tools, Inc</td>
			<td>18004-50</td>
			<td></td>
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	</tbody>
</table>

generating a mouse model of traumatic brain injury using an impactor device

Source: Schwulst, S. J. et al. <a href="https://app-jove-com.remotexs.ntu.edu.sg/v/60027">Murine Model of Controlled Cortical Impact for the Induction of Traumatic Brain Injury.</a> J. Vis. Exp. (2019)
This video demonstrates the procedure for inducing traumatic brain injury (TBI) in a mouse model. An anesthetized mouse with an exposed skull is taken to create an opening in the skull, and then a controlled cortical injury is induced using an impact device. The injury damages the brain tissue, causing impaired cognitive function and motor coordination.

Generating a Mouse Model of Traumatic Brain Injury Using an Impactor Device

All procedures involving animal models have been reviewed by the local institutional animal care committee and the JoVE veterinary review board.
1. Induce ...

Take an anesthetized mouse with an exposed skull.
Identify the sagittal and coronal sutures, which are skull joints, to locate the target site for injury.
Drill through the skull and remove the bone flap to expose the dura mater, the protective layer over the brain&#39;s cortex.
Secure the mouse in a stereotaxic frame with an attached impact device.
Position the impactor tip over the exposed area to inflict traumatic brain injury.
Lower the tip to contact the dura mater, retract it to set an impact depth, and apply the injury.
Withdraw the impactor and remove the mouse from the frame.
Control bleeding from the injury site, then close the scalp using surgical adhesive.
Administer an analgesic subcutaneously to alleviate pain and allow the mouse to recover.
The injury causes progressive damage to the cortex and underlying hippocampus and thalamus regions, along with fluid accumulation, impairing cognitive functions and motor coordination.

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42 Views. Source: Schwulst, S. J. et al. Murine Model of Controlled Cortical Impact for the Induction of Traumatic Brain Injury. J. Vis. Exp. (2019) This video demonstrates the procedure for inducing traumatic brain injury (TBI) in a mouse model. An anesthetized mouse with an exposed skull is taken to create an opening in the skull, and then a controlled cortical injury is induced using an impact device. The injury damages the brain tissue, causing impaired cognitive function and motor coordination. 

Video: Generating a Mouse Model of Traumatic Brain Injury Using an Impactor Device - Concept

Detecting Behavioral Deficits in Rats After Traumatic Brain Injury

With the increasing incidence of traumatic brain injury (TBI) in both civilian and military populations, TBI is now considered a chronic disease; however, few studies have investigated the long-term effects of injury in rodent models of TBI. Shown here are behavioral measures that are well-established in TBI research for times early after injury, such as two weeks, until two months. Some of these methods have previously been used at later times after injury, up to one year, but by very few laboratories. The methods demonstrated here are a short neurological assessment to test reflexes, a Beam-Balance to test balance, a Beam-Walk to test balance and motor coordination, and a working memory version of the Morris water maze that can be sensitive to deficits in reference memory. Male rats were handled and pre-trained to neurological, balance, and motor coordination tests prior to receiving parasagittal fluid percussion injury (FPI) or sham injury. Rats can be tested on the short neurological assessment (neuroscore), the beam-balance, and the Beam-Walk multiple times, while testing on the water maze can only be done once. This difference is because rats can remember the task, thus confounding the results if repeated testing is attempted in the same animal. When testing from one to three days after injury, significant differences are detected in all three non-cognitive tasks. However, differences in the Beam-Walk task were not detectable at later time points (after 3 months). Deficits were detected at 3 months in the Beam-Balance and at 6 months in the neuroscore. Deficits in working memory were detected out to 12 months after injury, and a deficit in a reference memory first appeared at 12 months. Thus, standard behavioral tests can be useful measures of persistent behavioral deficits after FPI.

The goal of the behavioral tests presented here is to detect functional deficits in rats after traumatic brain injury. Four specific tests are presented that detect deficits in behaviors to reflect the damage to specific brain areas at times extending to one year after injury.

With the increasing incidence of traumatic brain injury (TBI) in both civilian and military populations, TBI is now considered a chronic disease; however, ...

JoVE Journal

Behavior

A Novel In Vitro Model of Blast Traumatic Brain Injury

Traumatic brain injury is a leading cause of death and disability in military and civilian populations. Blast traumatic brain injury results from the detonation of explosive devices, however, the mechanisms that underlie the brain damage resulting from blast overpressure exposure are not entirely understood and are believed to be unique to this type of brain injury. Preclinical models are crucial tools that contribute to better understand blast-induced brain injury. A novel in vitro blast TBI model was developed using an open-ended shock tube to simulate real-life open-field blast waves modelled by the Friedlander waveform. C57BL/6N mouse organotypic hippocampal slice cultures were exposed to single shock waves and the development of injury was characterized up to 72 h using propidium iodide, a well-established fluorescent marker of cell damage that only penetrates cells with compromised cellular membranes. Propidium iodide fluorescence was significantly higher in the slices exposed to a blast wave when compared with sham slices throughout the duration of the protocol. The brain tissue injury is very reproducible and proportional to the peak overpressure of the shock wave applied.

A Novel In Vitro Model of Blast Traumatic Brain Injury

This paper describes a novel model of primary blast traumatic brain injury. A compressed-air driven shock tube is used to expose in vitro mouse hippocampal slice cultures to a single shock wave. This is a simple and rapid protocol generating a reproducible brain tissue injury with a high throughput.

Traumatic brain injury is a leading cause of death and disability in military and civilian populations. Blast traumatic brain injury results from the ...

Generating a Mouse Model of Traumatic Brain Injury Using an Impactor Device

Transcript

Generating a Mouse Model of Traumatic Brain Injury Using an Impactor Device

Detecting Behavioral Deficits in Rats After Traumatic Brain Injury

A Novel In Vitro Model of Blast Traumatic Brain Injury