This work was supported in part by the National Institutes of Health under award numbers R01NS120882, RF1NS119165, and R01NS103785 and the Department of Defense award number AZ190017. The content is solely the responsibility of the authors and does not represent the official views of the National Institutes of Health or the Department of Defense.

The experiments performed were approved by the Institutional Animal Care and Use Committee (IACUC) of the University of Kentucky, and both the ARRIVE and the Guide for the Care and Use of Laboratory Animals guidelines were followed during the study.
1. Surgical setup
NOTE: Mice are housed in groups of 4-5/cage, humidity in the housing room is maintained at 43%-47%, and the temperature is maintained at 22-23 &#730;C. Mice are given ad libitum access to food and water and exposed to a 12 h/12 h light/dark cycle (7 a.m./7 p.m.).
<ol>
	<li>Use a designated surgical area, such as a hood or dedicated surgical procedure room, to perform the animal surgery.</li>
	<li>Ensure that the surgical area includes a heating pad, a stereotaxic frame equipped with an electromagnetic impactor, and an anesthesia mask designed to administer isoflurane gas (see Figure 1A).</li>
	<li>Ensure that the surgeon or personnel involved in the surgery wear a clean lab coat, a face mask, gloves, and a surgical cap.</li>
	<li>Use sterile surgical tools, sterile cotton-tipped applicators, and gauze pads. Use a hot bead sterilizer to sterilize the instruments between mice during the day of surgery.</li>
	<li>Use an anesthesia induction chamber to prepare the mouse for surgery in a pre-op area.</li>
	<li>Use heating pads to maintain the animal&#39;s temperature, clean post-op mouse holding cages, and timers to record the righting reflex of the mouse after surgery.</li>
</ol>
2. Pre-surgery procedure
<ol>
	<li>Prepare the head support apparatus (see Figure 1B).
	<ol>
		<li>Remove the rolled end ridge from a 1 mL latex pipette bulb (inflatable end) (see Figure 1C).</li>
		<li>Attach the bulb to the tubing using parafilm (see Figure 1C).</li>
		<li>Connect the tubing to a 10 mL syringe using a stopcock. Fill the syringe with water (see Figure 1C). 
		NOTE: The 1 mL latex pipette bulb will be placed under the mouse&#39;s head to displace the impact force away from the ears. Try to remove as much air as possible from the bulb before use so that the bulb is filled with mostly water and not air.</li>
	</ol>
	</li>
	<li>Impactor setup.
	<ol>
		<li>Select the 5 mm probe tip, screw it to the piston at the bottom center of the actuator (inside the larger cylinder), and gently tighten the probe without applying excessive force. Re-tighten the tip between impacts (see Figure 1B).</li>
		<li>Before turning on the impactor, ensure that the Extend/Retract switch is positioned in the center Off position. Then, connect the cable on the actuator to the jack on the front panel of the impactor control box and the sensor cable to the jack on the front panel. Then, turn on the power switch on the back panel (see Figure 1D). 
		NOTE: The Extend/Retract toggle switch needs to remain in the center Off position when not in use.</li>
		<li>Set up the impact velocity by rotating the large knob on the left side of the control box until an impact velocity of 5.0 &#177; 0.2 m/s appears on the display (see Figure 1D).</li>
		<li>Set the dwell counter&#160;to 100 ms by turning the dials until the dwell reads 0.01 (see Figure 1D). 
		NOTE: The dwell is the time of contact before automatic retraction occurs.</li>
		<li>Place the impactor actuator on an ice pack to prevent the plastic cylinder from expanding, which locks the cylinder in place, preventing the movement of the cylinder and the delivery of future impacts (see Figure 1E).</li>
	</ol>
	</li>
	<li>Prepare the mouse for surgery.
	<ol>
		<li>Visually inspect the mouse before surgery and eliminate the mouse from the study if one of the following conditions is observed: poor coat condition, lethargy, or poor weight (&#60;20 g) for a 4-month-old mouse.</li>
		<li>Anesthetize the mouse with 4%-5% isoflurane in 100% oxygen using an induction chamber placed on a heating pad for 1-2 min.</li>
		<li>Shave the fur from the operative site using an electric hair clipper.</li>
		<li>Clean the head with sterile alcohol prep pads and apply a topical anesthetic to the shaved scalp at least 15 min before the start of the surgery.</li>
		<li>Return the mouse to a clean holding cage before the surgery. Start the surgery after at least 15 min of topical anesthetic application (induction time). 
		NOTE: The time for anesthesia could vary depending on the anesthetic used in the procedure.</li>
	</ol>
	</li>
	<li>Check one more time that the stereotaxic frame, impactor, and digital stereotaxic display (see Figure 1F) are ready to be used.</li>
	<li>Return the mouse to the isoflurane induction chamber with 4%-5% isoflurane in 100% oxygen for approximately 3 min.</li>
	<li>Fix the mouse in the head stage.</li>
</ol>
3. Surgical procedure
<ol>
	<li>Secure the mouse into the stereotaxic frame using lightweight acetal resin tapered point ear bars, a bite bar, and a mouse anesthesia mask (see Figure 1G,H). The isoflurane gas is delivered at 2%-3% in room air at 100-200 mL/min. Carefully monitor the mouse&#39;s respiration to ensure the depth of anesthesia and adjust the level of gas as needed.</li>
	<li>Apply sterile eye lubricant to the eyes to prevent corneal drying.</li>
	<li>Sterilize the scalp with povidone-iodine swabs and sterile alcohol pads three times.</li>
	<li>Ensure the mouse is deeply anesthetized by verifying the lack of a toe-pinch response.</li>
	<li>Make an approximately 1 cm midline scalp incision between the eyes and neck using a scalpel, exposing the skull (see Figure 1I).</li>
	<li>Allow the skull to dry for 1-2 min.</li>
	<li>Identify bregma (the intersection point of the coronal and sagittal sutures) and lambda (the intersection of the sagittal and lambdoid sutures) (see Figure 1J). 
	NOTE: A mouse brain atlas could be used for reference.</li>
	<li>Place the head support apparatus under the head and inflate the bulb with water until it is pressing against the bottom of the mouse&#39;s head but not lifting the head away from the bite bar. 
	NOTE: This step is essential to reduce possible ear issues from the CHI. Any animal with damage to the ear from the ear bars, resulting in rolling or bleeding, should be eliminated from the study and euthanized.</li>
	<li>Move the impactor into place over the animal&#39;s head.</li>
	<li>Extend the impactor by placing the Extend/Retract toggle switch (on the impactor control box) on Extend. 
	NOTE: Be sure to check that the tip is fully extended by pulling down on the tip.</li>
	<li>Line up the impactor until it is centered over the bregma (see Figure 1K).</li>
	<li>Reset the digital stereotaxic x and y coordinates in the stereotaxic reader to 0 (on the touch screen control)</li>
	<li>Align the probe over the impact location by moving the probe from the bregma over to the target coordinates: medial-lateral = 0.0 mm, anterior-posterior = &#8722;1.6 mm.</li>
	<li>Clip the contact sensor to the animal&#39;s ear.
	<ol>
		<li>Slowly lower the probe tip with the extended probe until first contact with the surface is made. Stop at the beep.</li>
		<li>Reset the digital stereotaxic z coordinates in the stereotaxic reader to 0.</li>
	</ol>
	</li>
	<li>Carefully inspect if the tip is flush with the skull (medial-lateral and anterior-posterior planes). 
	NOTE: Positioning the probe tip is the most crucial step of this process to prevent skull fractures and ear damage.</li>
	<li>Retract the impactor by placing the toggle switch on the control box in the Retract position. The tip withdraws and has no more contact with the animal&#39;s head until the time of impact.</li>
	<li>Set the impact depth by adjusting the dorsal-ventral depth to &#8722;1.2 mm. 
	NOTE: The depth of the impact affects the severity of the injury. The depth should be titered for different ages, weights, and strains of mice to the desired injury severity. The depth may need to be adjusted/re-titered over time to maintain a consistent injury severity. The severity can be assessed neuropathologically: microglia and astrocytes (IHC), and behaviorally: the radial arm water maze and the active avoidance test.</li>
	<li>Carefully monitor the mouse&#39;s respiration to ensure the anesthesia depth and adjust the gas level as needed. 
	NOTE: Often, the percentage of isoflurane gas should be lowered or shut off for 10-20 s prior to impact. Watch closely for the respiration to accelerate slightly. If respiration is too slow at the time of impact, the animal may die within the first 60 s after the impact from apnea. This can be prevented by adjusting the depth of anesthesia in the seconds before the impact.</li>
	<li>Induce the impact by pressing the right toggle switch to impact. The probe tip goes down at the displayed velocity, and then stays down for the set dwell time and retracts. 
	NOTE: Sham mice receive identical handling to the CHI mice, but the impact is not delivered.</li>
	<li>Start the timer immediately after the CHI impact is delivered to record the righting times (time to return from the side position to the prone position) or start the timer when the mouse is removed from the stereotaxic frame for the sham mice. The average righting reflex time is 5-15 min. 
	NOTE: The righting reflex times may vary based on mouse strain and age.</li>
	<li>Evaluate the mice for visible skull fractures, hemorrhages, and apnea. Exclude the mice with a depressed skull fracture or visible hemorrhage from the study. 
	NOTE: There are graded levels of skull fractures. Animals with decompressed skull fractures, where the bone is observably pressing into the brain tissue, are euthanized (CO2 first, and decapitation used as a secondary method). If the impactor tip is set correctly, these types of skull fractures are exceedingly rare. If a skull fracture does occur, the more common presentation is a small drop of blood on the skull and a slight tactile roughing of the skull, often along the suture connecting the posterior tip of the nasal bone. These mice are noted as possible skull fracture in the records but are not normally excluded from the study.</li>
	<li>Remove the animal from the stereotaxic frame.</li>
	<li>Close the scalp by stapling the skin together. 
	NOTE: Absorbable or non-absorbable sutures could be used to close the scalp as an alternative to staples.</li>
	<li>Apply triple antibiotic ointment with a sterile cotton-tipped applicator to the closed incision.</li>
	<li>Return the mouse to a clean holding cage for recovery. Half of the recovery cage is on a heating pad (low setting), providing the ability to move away from the heat when awake and maintaining the animal&#39;s temperature while unconscious (see Figure 1L). 
	NOTE: The mouse is placed on its side in the recovery cage. To prevent suffocation, place the animal in a recovery cage without bedding or on a tissue if bedding is in the cage.</li>
	<li>Return the Extend/Retract toggle switch to the Center/Off position. 
	NOTE: The current will continue running if the switch is left in either the extend or retract position, causing the piston to swell. The impactor will then not be functional until the piston cools down.</li>
	<li>Remove the impactor from its holder and gently place it on the ice pack. 
	NOTE: Keeping the impactor on an ice pack helps reduce the potential swelling of the impactor.</li>
	<li>Monitor the animal until the righting reflex occurs and document the time until righting (see Figure 1M). 
	NOTE: The righting reflex is defined as the moment the mouse returns to a prone position. The cage needs to be left undisturbed; the mouse could right if the cage is touched, moved, or exposed to some noises.</li>
	<li>Return the mice to their home cage when they are awake and alert. Usually, within 1 h after injury, animals are fully conscious and ambulating. Also, add some moist food at the bottom of the cage.</li>
</ol>
4. Post-surgery care
<ol>
	<li>Monitor the animals for 5 days post-surgery.</li>
	<li>Record their weight and any physical/ behavioral changes like breathing rate (qualitative respiratory function), gait, body and hair coat condition, eating, drinking, defecation, and urination.</li>
	<li>Observe the mouse for any sign of discomfort and the surgical wound for swelling, exudates or red edges, ordehiscence. Contact a veterinarian if the animal shows signs of pain and discomfort (vocalizations, not moving, hypothermia, does not drink or eat).</li>
	<li>Remove the staples 7-10 days post-surgery under anesthesia and on a heating pad. 
	&#8203;NOTE: If non-absorbable sutures are used, they must be removed 7-10 days post-surgery under anesthesia.</li>
</ol>
5. Cleaning
<ol>
	<li>Clean and sterilize the surgical area and tools.</li>
	<li>Clean the probe tip after each use and at the end of the day with alcohol prep pads. 
	NOTE: The impactor is calibrated at the factory and is reported to be stable over time and use. No routine calibration is needed. However, the impactor and stereotaxic frame should be routinely inspected. Also, the model endpoint perimeters such as righting reflex time, mortality, and neuropathology should be monitored to evaluate possible experimental drift.</li>
</ol>
6. Exclusion criteria
<ol>
	<li>Exclude animals prior to surgery with a poor health condition, like poor weight &#60;20 g for a 4-month-old mouse, lethargy, and poor coat condition.</li>
	<li>Exclude animals with complications during surgery like a depressed skull fracture, a visible hemorrhage related to surgery, or ear bleeding.</li>
	<li>Exclude animals from the study with the following post-surgery symptoms: failure to eat and/or to move normally, unusual vocalizations, weight loss, or failure of the wound to heal normally after surgery. 
	NOTE: This model could be used as a repetitive model of mild TBI. If the mice receive the second surgery 24 h apart from the first one, the staples or suture could be removed, and the same incision could be used to expose the skull. A new incision needs to be made if a longer time elapses between the surgeries.</li>
</ol>

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The authors have nothing to disclose.

Several steps are involved in recreating a consistent injury model using the described model. First, it is critical to correctly secure the animal into the stereotaxic frame. The animal's head should not be able to move laterally, and the skull should be completely flat with bregma and lambda reading the same coordinates. Correctly placing the ear bars is the most difficult aspect of this surgery, and this can only be learned with practice. If the skull is not level, the head should be adjusted before inducing CHI. Failu...

Traumatic brain injury (TBI) is caused by an external force on the brain, often associated with falls, sports injuries, physical violence, or road accidents. In 2014, the Centers for Disease Control and Prevention determined that 2.53 million Americans visited the emergency department to seek medical help for TBI-related accidents1. Since mild TBI (mTBI) represents the majority of TBI cases, over the past several decades, multiple models of mTBI have been adopted, which include weight drop, piston-driven closed head injury and controlled cortical impact, rotational injury, mild fluid percussion injury, and blast injury models2,3. The heterogeneity of the mTBI models is useful to address the different features associated with mTBI seen in people and to help evaluate the cellular and molecular mechanisms associated with brain injury.
Of the commonly used models of closed head injury, one of the first and most widely used models is the weight drop method, where an object is dropped from a specific height onto the animal&#39;s head (anesthetized or awake)2,4. In the weight drop method, the injury&#39;s severity depends on several parameters, including craniotomy performed or not, head fixed or free, and the distance and weight of the falling object2,4. One disadvantage of this model is the high variability in the severity of the injury and the high mortality rate associated with respiratory depression5,6. A common alternative is to deliver the impact using a pneumatic or electromagnetic device, which can be done directly on the exposed dura (controlled cortical impact: CCI) or closed skull (closed head injury: CHI). One of the strengths of the piston-driven injury is its high reproducibility and low mortality. However, CCI requires craniotomy7,8, and a craniotomy itself induces inflammation9. Instead, in the CHI model, there is no need for craniotomy. As already stated, each model has limitations. One of the limitations of the CHI model described in this paper is that the surgery is performed using a stereotaxic frame, and the head of the animal is immobilized. While the full head immobilization assures reproducibility, it does not account for movement after the impact that could contribute to the injury associated with a mTBI.
This protocol describes a basic method to perform a CHI impact with a commercially available electromagnetic impactor device10 in a mouse. This protocol details the exact parameters involved to achieve a highly reproducible injury. In particular, the investigator has precise control over the parameters (depth of injury, dwell time, and velocity of impact) to precisely define the injury severity. As described, this CHI model produces an injury that results in bilateral pathology, both diffuse and microscopic (i.e., chronic activation of glia, axonal and vascular damage), and behavioral phenotypes11,12,13,14,15. In addition, the described model is considered mild as it does not induce structural brain lesions based on MRI or gross lesions on pathology even 1 year after the injury16,17.

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electromagnetic controlled closed-head model of mild traumatic brain injury in mice

Highly reproducible animal models of traumatic brain injury (TBI), with well-defined pathologies, are needed for testing therapeutic interventions and understanding the mechanisms of how a TBI alters brain function. The availability of multiple animal models of TBI is necessary to model the different aspects and severities of TBI seen in people. This manuscript describes the use of a midline closed head injury (CHI) to develop a mouse model of mild TBI. The model is considered mild because it does not produce structural brain lesions based on neuroimaging or gross neuronal loss. However, a single impact creates enough pathology that cognitive impairment is measurable at least 1 month after injury. A step-by-step protocol to induce a CHI in mice using a stereotaxically guided electromagnetic impactor is defined in the paper. The benefits of the mild midline CHI model include the reproducibility of the injury-induced changes with low mortality. The model has been temporally characterized up to 1 year after the injury for neuroimaging, neurochemical, neuropathological, and behavioral changes. The model is complementary to open skull models of controlled cortical impact using the same impactor device. Thus, labs can model both mild diffuse TBI and focal moderate-to-severe TBI with the same impactor.

This stereotaxic electromagnetic impactor device is versatile. It is used for both an open skull controlled cortical impact (CCI) or a closed head injury (CHI) surgery. Furthermore, the injury severity can be modulated by changing the injury parameters such as impact velocity, dwell time, impact depth, impactor tip, and injury target. Herein is described a CHI surgery using a 5.0 mm steel tip impactor. This injury is considered mild because there are no structural brain lesions. The mortality rate in adult mice is less t...

Watch this Scientific Journal Video about Electromagnetic Controlled Closed-Head Model of Mild Traumatic Brain Injury in Mice at JoVE.com

Electromagnetic Controlled Closed-Head Model of Mild Traumatic Brain Injury in Mice

The protocol describes mild traumatic brain injury in a mouse model. In particular, a step-by-step protocol to induce a mild midline closed head injury and the characterization of the animal model is fully explained.

Highly reproducible animal models of traumatic brain injury (TBI), with well-defined pathologies, are needed for testing therapeutic interventions and ...

electromagnetic-controlled-closed-head-model-mild-traumatic-brain

Research

JoVE Journal

Neuroscience

2.0K Views.  University of Kentucky. The protocol describes mild traumatic brain injury in a mouse model. In particular, a step-by-step protocol to induce a mild midline closed head injury and the characterization of the animal model is fully explained.

Video: Electromagnetic Controlled Closed-Head Model of Mild Traumatic Brain Injury in Mice

Lateral Fluid Percussion: Model of Traumatic Brain Injury in Mice

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

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

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

A Method to Inflict Closed Head Traumatic Brain Injury in Drosophila

Traumatic brain injury (TBI) affects millions of people each year, causing impairment of physical, cognitive, and behavioral functions and death. Studies using Drosophila have contributed important breakthroughs in understanding neurological processes. Thus, with the goal of understanding the cellular and molecular basis of TBI pathologies in humans, we developed the High Impact Trauma (HIT) device to inflict closed head TBI in flies. Flies subjected to the HIT device display phenotypes consistent with human TBI such as temporary incapacitation and progressive neurodegeneration. The HIT device uses a spring-based mechanism to propel flies against the wall of a vial, causing mechanical damage to the fly brain. The device is inexpensive and easy to construct, its operation is simple and rapid, and it produces reproducible results. Consequently, the HIT device can be combined with existing experimental tools and techniques for flies to address fundamental questions about TBI that can lead to the development of diagnostics and treatments for TBI. In particular, the HIT device can be used to perform large-scale genetic screens to understand the genetic basis of TBI pathologies.

A Method to Inflict Closed Head Traumatic Brain Injury in Drosophila

Here we describe a method to inflict closed head traumatic brain injury (TBI) in Drosophila. This method provides a gateway to investigate the cellular and molecular mechanisms that underlie TBI pathologies using the vast array of experimental tools and techniques available for flies.

Traumatic brain injury (TBI) affects millions of people each year, causing impairment of physical, cognitive, and behavioral functions and death. Studies ...

Advanced Diffusion Imaging in The Hippocampus of Rats with Mild Traumatic Brain Injury

Mild traumatic brain injury (mTBI) is the most common type of acquired brain injury. Since patients with traumatic brain injury show a tremendous variability and heterogeneity (age, gender, type of trauma, other possible pathologies, etc.), animal models play a key role in unraveling factors that are limitations in clinical research. They provide a standardized and controlled setting to investigate the biological mechanisms of injury and repair following TBI. However, not all animal models mimic the diffuse and subtle nature of mTBI effectively. For example, the commonly used controlled cortical impact (CCI) and lateral fluid percussion injury (LFPI) models make use of a craniotomy to expose the brain and induce widespread focal trauma, which are not commonly seen in mTBI. Therefore, these experimental models are not valid to mimic mTBI. Thus, an appropriate model should be used to investigate mTBI. The Marmarou weight drop model for rats induces similar microstructural alterations and cognitive impairments as seen in patients who sustain mild trauma; therefore, this model was selected for this protocol. Conventional computed tomography and magnetic resonance imaging (MRI) scans commonly show no damage following a mild injury, because mTBI induces often only subtle and diffuse injuries. With diffusion weighted MRI, it is possible to investigate microstructural properties of brain tissue, which can provide more insight into the microscopic alterations following mild trauma. Therefore, the goal of this study is to obtain quantitative information of a selected region-of-interest (i.e., hippocampus) to follow up disease progression after obtaining a mild and diffuse brain injury.

The overall goal of this procedure is to obtain quantitative microstructural information of the hippocampus in a rat with mild traumatic brain injury. This is done using an advanced diffusion-weighted magnetic resonance imaging protocol and region-of-interest based analysis of parametric diffusion maps.

Mild traumatic brain injury (mTBI) is the most common type of acquired brain injury. Since patients with traumatic brain injury show a tremendous ...

Murine Model of Controlled Cortical Impact for the Induction of Traumatic Brain Injury

The Centers for Disease Control and Injury Prevention estimate that almost 2 million people sustain a traumatic brain injury (TBI) every year in the United States. In fact, TBI is a contributing factor to over a third of all injury-related mortality. Nonetheless, the cellular and molecular mechanisms underlying the pathophysiology of TBI are poorly understood. Thus, preclinical models of TBI capable of replicating the injury mechanisms pertinent to TBI in human patients are a critical research need. The controlled cortical impact (CCI) model of TBI utilizes a mechanical device to directly impact the exposed cortex. While no model can full recapitulate the disparate injury patterns and heterogeneous nature of TBI in human patients, CCI is capable of inducing a wide range of clinically applicable TBI. Furthermore, CCI is easily standardized allowing investigators to compare results across experiments as well as across investigative groups. The following protocol is a detailed description of applying a severe CCI with a commercially available impacting device in a murine model of TBI.

Here we describe a protocol for the induction of murine traumatic brain injury via an open-head controlled cortical impact.

The Centers for Disease Control and Injury Prevention estimate that almost 2 million people sustain a traumatic brain injury (TBI) every year in the ...

Low-intensity Blast Wave Model for Preclinical Assessment of Closed-head Mild Traumatic Brain Injury in Rodents

Traumatic brain injury (TBI) is a large-scale public health problem. Mild TBI is the most prevalent form of neurotrauma and accounts for a large number of medical visits in the United States. There are currently no FDA-approved treatments available for TBI. The increased incidence of military-related, blast-induced TBI further accentuates the urgent need for effective TBI treatments. Therefore, new preclinical TBI animal models that recapitulate aspects of human blast-related TBI will greatly advance the research efforts into the neurobiological and pathophysiological processes underlying mild to moderate TBI as well as the development of novel therapeutic strategies for TBI.
Here we present a reliable, reproducible model for the investigation of the molecular, cellular, and behavioral effects of mild to moderate blast-induced TBI. We describe a step-by-step protocol for closed-head, blast-induced mild TBI in rodents using a bench-top setup consisting of a gas-driven shock tube equipped with piezoelectric pressure sensors to ensure consistent test conditions. The benefits of the setup that we have established are its relative low-cost, ease of installation, ease of use and high-throughput capacity. Further advantages of this non-invasive TBI model include the scalability of the blast peak overpressure and the generation of controlled reproducible outcomes. The reproducibility and relevance of this TBI model has been evaluated in a number of downstream applications, including neurobiological, neuropathological, neurophysiological and behavioral analyses, supporting the use of this model for the characterization of processes underlying the etiology of mild to moderate TBI.

We present here a protocol of a blast wave model for rodents to investigate neurobiological and pathophysiological effects of mild to moderate traumatic brain injury. We established a gas-driven, bench-top setup equipped with pressure sensors allowing for reliable and reproducible generation of blast-induced mild to moderate traumatic brain injury.

Traumatic brain injury (TBI) is a large-scale public health problem. Mild TBI is the most prevalent form of neurotrauma and accounts for a large number of ...

Assessing Changes in Synaptic Plasticity Using an Awake Closed-Head Injury Model of Mild Traumatic Brain Injury

Mild traumatic brain injuries (mTBIs) are a prevalent health issue in North America. There is increasing pressure to utilize ecologically valid models of closed-head mTBI in the preclinical setting to increase translatability to the clinical population. The awake closed-headed injury (ACHI) model uses a modified controlled cortical impactor to deliver closed-headed injury, inducing clinically relevant behavioral deficits without the need for a craniotomy or the use of an anesthetic.
This technique does not normally induce fatalities, skull fractures, or brain bleeds, and is more consistent with being a mild injury. Indeed, the mild nature of the ACHI procedure makes it ideal for studies investigating repetitive mTBI (r-mTBI). Growing evidence indicates that r-mTBI can result in a cumulative injury that produces behavioral symptoms, neuropathological changes, and neurodegeneration. r-mTBI is common in youths playing sports, and these injuries occur during a period of robust synaptic reorganization and myelination, making the younger population particularly vulnerable to the long-term influences of r-mTBI.
Further, r-mTBI occurs in cases of intimate partner violence, a condition for which there are few objective screening measures. In these experiments, synaptic function was assessed in the hippocampus in juvenile rats that had experienced r-mTBI using the ACHI model. Following the injuries, a tissue slicer was utilized to make hippocampal slices to evaluate bidirectional synaptic plasticity in the hippocampus at either 1 or 7 days following the r-mTBI. Overall, the ACHI model provides researchers with an ecologically valid model to study changes in synaptic plasticity following mTBI and r-mTBI.

Here, it is demonstrated how an awake closed-head injury model can be used for examining the effects of repeated mild traumatic brain injury (r-mTBI) on synaptic plasticity in the hippocampus. The model replicates important features of r-mTBI in patients and is used in conjunction with in vitro electrophysiology.

Mild traumatic brain injuries (mTBIs) are a prevalent health issue in North America. There is increasing pressure to utilize ecologically valid models of ...

Validating Injury Response and Behavioral Assessments in an mTBI Rat Model

Rat Model of Closed-Head Mild Traumatic Injury and its Validation

Animal models are crucial for advancing our understanding of mild traumatic brain injury (mTBI) and guiding clinical research. To achieve meaningful insights, developing a stable and reproducible animal model is essential. In this study, we report a detailed description of a closed-head mTBI model and a representative validation method using Sprague-Dawley rats to verify the modeling effect. The model involves dropping a 550 g mass weight from a height of 100 cm directly onto the head of a rat on a destructible surface, followed by a 180-degree turn. To assess the injury, rats underwent a series of neurobehavioral assessments 10 min post-injury, including time of loss of consciousness, first seeking-behavior time, escape ability, and beam balance ability test. During the acute and subacute stages following the injury, behavioral tests were conducted to assess motor coordination ability (Beam task), anxiety (Open Field test), and learning and memory abilities (Morris Water Maze test). The closed-head mTBI model produced a consistent injury response with minimal mortality and replicated real-life situations. The validation method effectively verified the model development and ensured the stability and consistency of the model.

Author Spotlight: Advancing Traumatic Brain Injury Research - A Closed-Head Model for Accurate Replication and Rapid Assessment 

Here, we present a closed-head mild traumatic brain injury (mTBI) rat model and its validation exhibiting remarkable similarity to human mTBI concerning behavioral manifestations during the acute and subacute stages.

Animal models are crucial for advancing our understanding of mild traumatic brain injury (mTBI) and guiding clinical research. To achieve meaningful ...