Name: induction of traumatic brain injury in a rat model using a lateral fluid percussion device
Uploaded: 2025-03-31T15:00:50.000+00:00
Duration: 1 min 12 s
Description: Source: McGuire, M. J., et al. Use of a Wireless Video-EEG System to Monitor Epileptiform Discharges Following Lateral Fluid-Percussion Induced Traumatic ...

induction of traumatic brain injury in a rat model using a lateral fluid percussion device

Induction of Traumatic Brain Injury in a Rat Model Using a Lateral Fluid Percussion Device

Make a 1.5 to 2.5-centimeters midline incision through the skin and muscle of the scalp using a number 10 scalpel blade. Retract the skin and muscle to expose the skull and provide a clear surgical field. Electric cautery is useful for achieving quick hemostasis.
Next, shave down the lateral ridge of the left parietal bone, using a surgical curette to produce a smooth, flat surface so that the base of the female-female Luer Lock hub can rest flush with the skull. Irrigate the skull surface and surrounding tissues with 2.0 mg per milliliter gentamicin solution in sterile saline, and blot excess solution with sterile swabs. Then apply 3% hydrogen peroxide to the skull to dry the bone.
At this point, create a 5-millimeter diameter craniectomy site through the left parietal bone. Then, remove the bone flap with the surgical curette and smooth tissue forceps.
Next, using a stereomicroscope, gently remove the thin rim of bone remaining with smooth tissue forceps taking care not to rupture the dura. Then swab the skull with 70% ethanol to remove any bone dust, and to dry the skull. Apply a thin layer of cyanoacrylate glue around the bottom edge of the Luer Lock hub, and secure it to the skull over the craniectomy without obstructing the opening and without allowing the glue to contact the dura. Then seal the Luer lock in place with a thin layer of glue around the outside base of the hub.
Next, prepare a slurry of dental cement and apply this to the surface of the skull around and over the base of the Luer Lock hub to secure it in place. Then fill the Luer Lock hub with a sterile, preservative-free solution containing multiple electrolytes using a syringe and needle. A convex bolus of saline should be seen above the top of the rim. Once the dental cement is completely cured, discontinue gas anesthesia and remove the rat from the stereotaxic frame.
Place the rat on a platform next to the fluid percussion injury device in sternal recumbency. Then, secure a 12-centimeter length of pressure tubing to the end of the device&#39;s curved tip, with the opposite end terminating in a 2-centimeter male Luer Lock twist connector. Secure the rat to the device by connecting the female end of the hub on the rat skull to the male connector.
Repeatedly check the animal for return of withdrawal reflex. As soon as the rat regains withdrawal reflex, but is still sedated, release the pendulum of the device to cause a single 20-millisecond pressure pulse and induce injury. Then immediately disconnect the rat from the FPI device, place it in sternal recumbency, and provide supplemental oxygen via a nose cone until spontaneous breathing returns.
Note that apnea is an anticipated consequence of the injury. If necessary, provide periodic manual breaths via a bag valve mask until the rat begins to spontaneously breathe on its own. Monitor continuously and record the time of return of righting reflex.

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1. Fluid percussion injury
<ol>
	<li>Wear a lab coat or surgical gown, surgical mask, surgical gloves, and head covering and sterilize all tools and materials that contact the surgical site.</li>
	<li>Anesthetize a 10-12-week-old, male, Wistar rat (350-400 g) with 3% isoflurane and 1 L/min oxygen in an induction chamber of appropriate size for rats. Remove the rat from the induction chamber and move it to the prep area once it is unconscious. Put the sterile ophthalmic ointment into both eyes.</li>
	<li>Shave the hair&#160;on the rat&#39;s head with electric clippers with a #40 blade from just above the eyes to the caudal base of the ears to produce enough surgical field. Remove any loose, clipped hair from the site.</li>
	<li>Clean the surgical site by applying 2% chlorohexidine scrub to the shaved scalp followed by 70% ethanol. Start at the center and move outward in concentric circles away from the incision site. Repeat this process 3 times. Apply Betadine solution to the site in the same fashion and allowed to dry.&#160;</li>
	<li>Place the anesthetized rat into the stereotaxic frame and maintain anesthesia at 2-3% isoflurane-1 L/min oxygen via nosecone. Check for loss of withdrawal reflex of hindlimb and loss of palpebral reflex to ensure the rat is in a surgical plane of anesthesia.</li>
	<li>Monitor the respiratory rate, heart rate, body temperature and oxygen saturation throughout the surgery. Maintain heart rate between 300-400 bpm, and SpO2&#160;above 90%. 
	NOTE: A pulse-oximeter attached to a rear foot can be used to provide the constant read out of heart rate and SpO2. A heart rate above 400 bpm indicates the rat is not sufficiently anesthetized. A self-regulating warming pad, coupled to a rectal thermometer, set at 37 &#176;C, can be positioned under the rat throughout the surgery to maintain body temperature. A stereomicroscope with a light source in combination with an optic fiber lamp are helpful for visualizing the procedure.</li>
	<li>Use a 23 g needle to inject 0.5% bupivacaine hydrochloride intradermally into the scalp at the incision site for local analgesia 10 -&#160;15 minutes prior to making an incision.&#160;</li>
	<li>Make a 1.5-2.5 cm midline incision through the skin and muscle of the scalp using a #10 scalpel blade. Retract the skin and muscle to expose the skull and provide a clear surgical field. Reflect the underlying fascia and fatty tissue away from the bone with sterile cotton swabs. 
	NOTE: An electric cautery unit is useful for achieving&#160;quick hemostasis.</li>
	<li>Shave down the lateral ridge of the left parietal bone using a surgical curette to produce a smooth flat surface so that the base of the female-female Luer lock hub can rest flush with the skull.</li>
	<li>Irrigate the skull surface and surrounding tissues with 2.0 mg/mL gentamicin solution in sterile saline. Blot excess solution with a sterile swab.</li>
	<li>Apply 3% hydrogen peroxide to the skull to dry the bone.&#160; 
	NOTE: If the bone is not sufficiently dry the dental cement will not adhere properly and form a solid seal.</li>
	<li>Create a 5 mm diameter craniectomy site through the left parietal bone.&#160; 
	NOTE: A trephine bit placed into a power drill attached to the stereotactic frame can be helpful to initiate the craniectomy. Use a hand drill with a 5 mm diameter trephine to slowly finish the craniectomy through the remaining bone. When close to completing the craniectomy, rotate the trephine in reverse to prevent rupture of the underlying dura mater. There will be a thinning of the skull around the perimeter of the disk and the skull flap will feel loose when pressed lightly.</li>
	<li>Remove the bone flap with the surgical curette and smooth tissue forceps. 
	NOTE: Some bleeding may occur, but hemostasis can be quickly achieved by applying gentle pressure with sterile cotton swabs.</li>
	<li>Use a stereomicroscope and illumination to visually inspect the dura for any signs of rupture.&#160;A thin rim of bone will remain around the circumference of the craniectomy site. &#160;Gently remove this rim with smooth tissue forceps taking care not to rupture the dura.&#160;</li>
	<li>Swab the skull with 70% ethanol to remove any bone dust and to dry the skull.</li>
	<li>Apply a thin layer of cyanoacrylate gel glue around the bottom edge of the Luer lock hub and secure it to the skull over the craniectomy without obstructing the opening. Use caution not to bring glue in contact with the dura. Further, seal the Luer lock in place with an additional thin layer of glue around the outside base of the hub.</li>
	<li>Prepare a slurry of dental cement. Apply the cement to the surface of the skull around and over the base of the Luer lock hub to secure it in place.</li>
	<li>Fill the Luer lock hub with a sterile preservative free artificial cerebrospinal fluid (CSF) solution (pH 7.4) using a syringe and needle so that a convex bolus of saline can be seen above the top of the rim. 
	NOTE: The solution will keep the dura moist as the dental cement dries as well as serves as an indication of the integrity of the seal. If the solution level falls at all, that is an indication of a leak in the system and the Luer lock must be removed and replaced.</li>
	<li>Once the dental cement is completely cured, discontinue gas anesthesia and remove the rat from the stereotaxic frame.</li>
	<li>Place the rat on a platform next to the FPI device.</li>
	<li>The FPI device has a curved metal tip that extends from the pressure transducer at the end of the fluid reservoir. Secure a 12 cm length of pressure tubing to the end of the curved tip with the opposite end terminating in a 2 cm male Luer lock twist connector. Secure the rat to the FPI device by connecting the female end of the hub on the rat&#39;s skull to the male connector. 
	NOTE: Ensure the connection is tightly secured and that all air bubbles have been removed from the system.</li>
	<li>Place the animal in sternal recumbency and repeatedly check for return of withdrawal reflex. As soon as the rat regains withdrawal reflex but is still sedated, release the pendulum of the FPI device to cause a single 20 ms pressure pulse and induce injury. 
	NOTE: It is important to not induce the injury while the animal is deeply anesthetized as this tends to cause increased mortality due to neurogenic-induced pulmonary edema. All devices show variability. However, on the device used for this experiment, a 17&#176; angle placement of the hammer produces a 2.2 - 2.3 atmospheric pressure pulse. Uninjured, sham animals undergo all the same procedures with the exception of the actual fluid pulse to the induce injury.</li>
	<li>Immediately disconnect the rat from the FPI device after injury, place it in sternal recumbency, and provide supplemental oxygen (1 L/min) via a nose cone until spontaneous breathing returns. Apnea is an anticipated consequence of the injury. If necessary, provide periodic manual breaths via a bag valve mask until the rat begins to spontaneously breathe on its own. 
	NOTE: Typically, apnea lasts less than 2 min. A transient rapid rise in heart rate (&#62;500 bpm) is observed immediately after the administration of the pressure pulse due to a catecholamine burst. This can be monitored with a pulse oximeter attached to the rat&#39;s foot and can serve as a possible indicator that a severe injury has occurred.</li>
	<li>Monitor the rat continuously and record the time of return of righting reflex (stable ambulation on all four limbs).</li>
	<li>The magnitude of the atmospheric pressure pulse for each rat should be within &#177; 0.05 atmospheres of each other. Confirm that each of the pressure pulses produces a smooth signal on the oscilloscope with consistent amplitude and duration. 
	NOTE: A noisy signal may indicate air bubbles in the system that must be removed prior to delivering the injury pulse. Atmospheric pressure pulses that produce a severe injury, in this experiment, are those that typically result in animal righting times of 30-60 min. This range of righting times are associated with a mortality rate of approximately 40-50%).</li>
	<li>Administer 10 mL of prewarmed saline subcutaneously as supportive care.</li>
	<li>Return the rat to its home cage and allow it to recover for at least 4 h. 
	NOTE: Increased mortality has been observed when the rats are placed immediately back under anesthesia.</li>
</ol>

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>1.00 mm Drill Bits</td>
			<td>Drill Bit City: New Carbide Tools</td>
			<td>05M200</td>
			<td></td>
		</tr>
		<tr>
			<td>3M ESPE Durelon Carboxylate Cement</td>
			<td>3M , Neuss Germany</td>
			<td>38019</td>
			<td>Dental Cement</td>
		</tr>
		<tr>
			<td>4-0 Suture</td>
			<td>Ethicon, Sommerville, NJ</td>
			<td>K831H</td>
			<td>4-0 Ethicon Perma-hand Silk, 26mm 1/2c Taperpoint, 30&#34; (75cm), Black Braided non-absorbable suture</td>
		</tr>
		<tr>
			<td>5 mm outer diameter trephine</td>
			<td>Fine Science Tools</td>
			<td>18004-50</td>
			<td></td>
		</tr>
		<tr>
			<td>Dumont #6 Forceps</td>
			<td>Fine Science Tools</td>
			<td>11260-20</td>
			<td></td>
		</tr>
		<tr>
			<td>Dumont #7b Forceps</td>
			<td>Fine Science Tools</td>
			<td>11270-20</td>
			<td></td>
		</tr>
		<tr>
			<td>ecgAUTO</td>
			<td>EMKA Technologies, Falls Church, VA</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Female Luer Thread Style Coupler Clear Polycarbonare</td>
			<td>Cole-Palmer instrument</td>
			<td>SKO#45501-22</td>
			<td>Order lot #214271</td>
		</tr>
		<tr>
			<td>Foot Power Drill</td>
			<td>Grobet USA, Carlstadt, NJ</td>
			<td>Model C-300</td>
			<td></td>
		</tr>
		<tr>
			<td>GentaMax 100 (Gentamicin, Sulfate Solution)</td>
			<td>Phoenix, Manufactured by Clipper Distributing Company LLC, St. 
			Joseph, MO</td>
			<td></td>
			<td>NDC 57319-520-05</td>
		</tr>
		<tr>
			<td>Hill&#39;s Prescription Diet a/d Canine/ Feline</td>
			<td>Hill&#39;s Pet Nutrition, Inc. , Topeka, KS</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>IOX2 Software</td>
			<td>EMKA Technologies, Falls Church, VA</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Isoflorane, USP</td>
			<td>Piramal Enterprise Limited, Andhra, India</td>
			<td></td>
			<td>NDC 66794-013-25</td>
		</tr>
		<tr>
			<td>IsoTech Anesthesia machine</td>
			<td>SurgiVet</td>
			<td>WWV9000</td>
			<td></td>
		</tr>
		<tr>
			<td>Lateral FPI device</td>
			<td>AmScien</td>
			<td>302</td>
			<td>curved tip, with pressure tubing extension. connected via screw lock connector (Cole-Palmer; #4550-22)</td>
		</tr>
		<tr>
			<td>Leica A60 Stereomicroscope</td>
			<td>Leica Biosystems, Richmond, VA</td>
			<td>PN: 10 450 488</td>
			<td></td>
		</tr>
		<tr>
			<td>Marcaine (0.5%) Bupivacaine hcl injection usp 5 mg/mL</td>
			<td>Hospira, Lake Forest, IL</td>
			<td>CA-3627</td>
			<td>50mL multiple dose vial; NDC 0409-1610-50</td>
		</tr>
		<tr>
			<td>Micro-Adson Forceps</td>
			<td>Fine Science Tools</td>
			<td>11018-12</td>
			<td></td>
		</tr>
		<tr>
			<td>Olsen-Hegar Needle Holders with Suture Cutters</td>
			<td>Fine Science Tools</td>
			<td>12002-14</td>
			<td></td>
		</tr>
		<tr>
			<td>PALACOS R+G bone cement with gentamicin</td>
			<td>Heraeus,</td>
			<td>REF: 5036964</td>
			<td>Radiopaque bone cement containing 1 x 0.5g Gentamicin</td>
		</tr>
		<tr>
			<td>Povidone-iodine solution</td>
			<td></td>
			<td></td>
			<td>Betadine</td>
		</tr>
		<tr>
			<td>Puralube Vet Ointment</td>
			<td>Dechra Veterinary Products, Overland Park KS</td>
			<td></td>
			<td>NDC 17033-211-38</td>
		</tr>
		<tr>
			<td>Scalpel blade (#10) and holder</td>
			<td>Integra Miltex, York, PA</td>
			<td>REF: 4-110</td>
			<td></td>
		</tr>
		<tr>
			<td>Scalpel Handle - #4</td>
			<td>Fine Science Tools</td>
			<td>10004-13</td>
			<td></td>
		</tr>
		<tr>
			<td>Sickle Knife</td>
			<td>Bausch + Lomb Storz Instruments</td>
			<td>N1705 HM</td>
			<td>5mm curved blade. Round handle. Overall length 168mm, 6.6 inches.</td>
		</tr>
		<tr>
			<td>Silverstein Micro Mirror</td>
			<td>Bausch + Lomb Storz Instruments</td>
			<td>N1706 S8</td>
			<td>3mm diameter. Angled 45 degrees. Overall length 180mm, 7.2 inches</td>
		</tr>
		<tr>
			<td>Thermal Cautery Unit</td>
			<td>Geiger Medical Technology, Delasco Council Bluffs, IA</td>
			<td>Model NO: 150</td>
			<td></td>
		</tr>
		<tr>
			<td>Vetivex</td>
			<td>Dechra Veterinary Products, Overland Park KS</td>
			<td></td>
			<td>Veterinary pHyLyteTM Injection pH 
			7.4 (Multiple Electrolytes Injection, Type 1, USP)</td>
		</tr>
		<tr>
			<td>Wistar IGS rats</td>
			<td>Charles River</td>
			<td>strain code 003</td>
			<td>12 wk old at the time of injury</td>
		</tr>
		<tr>
			<td>Wullstein Retractor</td>
			<td>Fine Science Tools</td>
			<td>17018-11</td>
			<td></td>
		</tr>
	</tbody>
</table>

Source: McGuire, M. J., et al. <a href="https://app.jove.com/v/59637">Use of a Wireless Video-EEG System to Monitor Epileptiform Discharges Following Lateral Fluid-Percussion Induced Traumatic Brain Injury.</a> J. Vis. Exp. (2019)
This video demonstrates the induction of traumatic brain injury (TBI) in a rat using a lateral fluid percussion injury (FPI) device. The procedure involves creating a hole in the skull of an anesthetized rat to expose the dura mater, onto which a cannula is attached. The rat is then connected to the device, which generates a pressure wave that reaches the brain, mimicking TBI by causing neural cell death and vascular damage

Source: McGuire, M. J., et al. Use of a Wireless Video-EEG System to Monitor Epileptiform Discharges Following Lateral Fluid-Percussion Induced Traumatic ...

Place an anesthetized rat on a stereotaxic frame.
Make a midline incision and retract the skin to expose the skull.
Disinfect the skull, dry the surface, and scrape it to flatten the target area for cannula placement.
Drill a hole and remove the bone flap to expose the dura mater, the brain&#39;s outer covering.
Clean the skull with a swab.
Place a cannula over the hole and secure it with dental cement.
Fill it with artificial cerebrospinal fluid to prevent the dura mater from drying.
Connect the rat to the tubing of the lateral fluid percussion device, which consists of a pendulum and a fluid-filled reservoir.
As the rat regains the withdrawal reflex, release the pendulum from a calibrated angle to strike the reservoir, generating a pressure wave that hits the brain.
The pressure damages blood vessels and causes neural cell death, leading to traumatic brain injury.
Allow the rat to recover to assess the injury-induced neurodegeneration.

8 Views. Induction of Traumatic Brain Injury in a Rat Model Using a Lateral Fluid Percussion Device

Video: Induction of Traumatic Brain Injury in a Rat Model Using a Lateral Fluid Percussion Device - Experiment

Assessing Dominant-Submissive Behavior in Adult Rats Following Traumatic Brain Injury

Competition over resources such as food, territory, and mates significantly influences relationships within animal species and is mediated through social hierarchies that are often based on dominant-submissive relationships. The dominant-submissive relationship is a normal behavioral pattern among the individuals of a species. Traumatic brain injury is a frequent cause of social interaction impairment and the reorganization of dominant-submissive relationships in animal pairs. This protocol describes submissive behavior in adult male Sprague-Dawley rats after the induction of traumatic brain injury using a fluid-percussion model compared to naive rats through a series of dominant-submissive tests performed between 29 days and 33 days after induction. The dominant-submissive behavior test shows how brain injury can induce submissive behavior in animals competing for food. After traumatic brain injury, the rodents were more submissive, as indicated by them spending less time at the feeder and being less likely to arrive first at the trough compared to the control animals. According to this protocol, submissive behavior develops after traumatic brain injury in adult male rats.

The present protocol describes a rat model of fluid percussion-induced traumatic brain injury followed by a series of behavioral tests to understand the development of dominant and submissive behavior. Using this model of traumatic brain injury in conjunction with specific behavioral tests enables the study of social impairments following brain injury.

Competition over resources such as food, territory, and mates significantly influences relationships within animal species and is mediated through social ...

JoVE Journal

Behavior

Investigating Alterations in Caecum Microbiota After Traumatic Brain Injury in Mice

Increasing evidence shows that the microbiota-gut-brain axis plays an important role in the pathogenesis of brain diseases. Several studies also demonstrate that traumatic brain injuries cause changes to the gut microbiota. However, mechanisms underlying the bidirectional regulation of the brain-gut axis remain unknown. Currently, few models exist for studying the changes in gut microbiota after traumatic brain injury. Therefore, the presented study combines protocols for inducing traumatic brain injury using a lateral fluid percussion device and analysis of caecum samples following injury for investigating alterations in the gut microbiome. Alterations of the gut microbiota composition after traumatic brain injury are determined using 16S-rDNA sequencing. This protocol provides an effective method for studying the relationships between enteric microorganisms and traumatic brain injury.

Presented here is a protocol to induce diffuse traumatic brain injury using a lateral fluid percussion device followed by the collection of the caecum content for gut microbiome analysis.

Induction of Traumatic Brain Injury in a Rat Model Using a Lateral Fluid Percussion Device

Transcript

Induction of Traumatic Brain Injury in a Rat Model Using a Lateral Fluid Percussion Device