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

Summary

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.

Abstract

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.

Introduction

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 models^2,3. 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's head (anesthetized or awake)^2,4. In the weight drop method, the injury's severity depends on several parameters, including craniotomy performed or not, head fixed or free, and the distance and weight of the falling object^2,4. One disadvantage of this model is the high variability in the severity of the injury and the high mortality rate associated with respiratory depression^5,6. 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 craniotomy^7,8, 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 phenotypes^{11,12,13,14,15}. 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 injury^16,17.

Protocol

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 ˚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.).

1. Use a designated surgical area, such as a hood or dedicated surgical procedure room, to perform the animal surgery.
2. 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).
3. Ensure that the surgeon or personnel involved in the surgery wear a clean lab coat, a face mask, gloves, and a surgical cap.
4. 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.
5. Use an anesthesia induction chamber to prepare the mouse for surgery in a pre-op area.
6. Use heating pads to maintain the animal's temperature, clean post-op mouse holding cages, and timers to record the righting reflex of the mouse after surgery.

2. Pre-surgery procedure

1. Prepare the head support apparatus (see Figure 1B).
   1. Remove the rolled end ridge from a 1 mL latex pipette bulb (inflatable end) (see Figure 1C).
   2. Attach the bulb to the tubing using parafilm (see Figure 1C).
   3. 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'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.
2. Impactor setup.
   1. 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).
   2. 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.
   3. 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 ± 0.2 m/s appears on the display (see Figure 1D).
   4. Set the dwell counter 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.
   5. 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).
3. Prepare the mouse for surgery.
   1. 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 (<20 g) for a 4-month-old mouse.
   2. Anesthetize the mouse with 4%-5% isoflurane in 100% oxygen using an induction chamber placed on a heating pad for 1-2 min.
   3. Shave the fur from the operative site using an electric hair clipper.
   4. 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.
   5. 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.
4. Check one more time that the stereotaxic frame, impactor, and digital stereotaxic display (see Figure 1F) are ready to be used.
5. Return the mouse to the isoflurane induction chamber with 4%-5% isoflurane in 100% oxygen for approximately 3 min.
6. Fix the mouse in the head stage.

3. Surgical procedure

1. 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's respiration to ensure the depth of anesthesia and adjust the level of gas as needed.
2. Apply sterile eye lubricant to the eyes to prevent corneal drying.
3. Sterilize the scalp with povidone-iodine swabs and sterile alcohol pads three times.
4. Ensure the mouse is deeply anesthetized by verifying the lack of a toe-pinch response.
5. Make an approximately 1 cm midline scalp incision between the eyes and neck using a scalpel, exposing the skull (see Figure 1I).
6. Allow the skull to dry for 1-2 min.
7. 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.
8. Place the head support apparatus under the head and inflate the bulb with water until it is pressing against the bottom of the mouse'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.
9. Move the impactor into place over the animal's head.
10. 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.
11. Line up the impactor until it is centered over the bregma (see Figure 1K).
12. Reset the digital stereotaxic x and y coordinates in the stereotaxic reader to 0 (on the touch screen control)
13. 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 = −1.6 mm.
14. Clip the contact sensor to the animal's ear.
    1. Slowly lower the probe tip with the extended probe until first contact with the surface is made. Stop at the beep.
    2. Reset the digital stereotaxic z coordinates in the stereotaxic reader to 0.
15. 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.
16. 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's head until the time of impact.
17. Set the impact depth by adjusting the dorsal-ventral depth to −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.
18. Carefully monitor the mouse'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.
19. 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.
20. 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.
21. 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.
22. Remove the animal from the stereotaxic frame.
23. 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.
24. Apply triple antibiotic ointment with a sterile cotton-tipped applicator to the closed incision.
25. 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'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.
26. 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.
27. 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.
28. 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.
29. 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.

4. Post-surgery care

1. Monitor the animals for 5 days post-surgery.
2. 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.
3. 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).
4. Remove the staples 7-10 days post-surgery under anesthesia and on a heating pad.
   ​NOTE: If non-absorbable sutures are used, they must be removed 7-10 days post-surgery under anesthesia.

5. Cleaning

1. Clean and sterilize the surgical area and tools.
2. 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.

6. Exclusion criteria

1. Exclude animals prior to surgery with a poor health condition, like poor weight <20 g for a 4-month-old mouse, lethargy, and poor coat condition.
2. Exclude animals with complications during surgery like a depressed skull fracture, a visible hemorrhage related to surgery, or ear bleeding.
3. 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.

Results

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...

Discussion

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...

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
9 mm Autoclip Applier	Braintree scientific	ACS- APL	Surgery
9 mm Autoclip Remover	Braintree scientific	ACS- RMV	Surgery
9 mm Autoclip, Case of 1,000 clips	Braintree scientific	ACS- CS	Surgery (Staples)
Aperio ImageScope software	Leica BioSystems	NA	IHC
BladeFLASK Blade Remover	Fisher Scientific	22-444-275	Surgery
Cotton tip applicator	VWR	89031-270	Surgery
Digitial mouse stereotaxic frame	Stoelting	51730D	Surgery
Dumont #7 Forceps	Roboz	RS-5047	Surgery
Ear bars	Stoelting	51649	Surgery
EthoVision XT 11.0	Noldus Information Technology	NA	RAWM
Fiber-Lite	Dolan-Jeffer Industries	UN16103-DG	Surgery
Fisherbrand Bulb for Small Pipets	Fisher Scientific	03-448-21	Head support apparatus
Gemini Avoidance System	San Diego Instruments	NA	Active avoidance
Heating Pad	Sunbeam	732500000U	Surgery prep
HRP conjugated goat anti-rabbit IgG	Jackson Immuno Research laboratories	111-065-144	IHC
Induction chamber	Kent Scientific	VetFlo-0530XS	Surgery prep
Isoflurane, USP	Covetrus	NDC: 11695-6777-2	Surgery
Mouse gas anesthesia head holder	Stoelting	51609M	Surgery
Neuropactor Stereotaxic Impactor	Neuroscience Tools	n/a	Surgery: Formally distributed by Lecia as impact one
NexGen Mouse 500	Allentown	n/a	Post-surgery, holding cage
Parafilm	Bemis	PM992	Head support apparatus
Peanut - Professional Hair Clipper	Whal	8655-200	Surgery prep
Povidone-Iodine Solution USP, 10% (w/v), 1% (w/v) available Iodine, for laboratory	Ricca	3955-16	Surgery
Puralube Vet Oinment,petrolatum ophthalmic ointment, Sterile ocular lubricant	Dechra	17033-211-38	Surgery
Rabbit anti-GFAP	Dako	Z0334	IHC
Rabbit anti-IBA1	Wako	019-19741	IHC
8-arm Radial Arm Water Maze	MazeEngineers	n/a	RAWM
Scale	OHAUS CS series	BAL-101	Surgery prep
Scalpel Handle #7 Solid 6.25"	Roboz	RS-9847	Surgery
Sterile Alcohol Prep Pads (isopropyl alcohol 70% v/v)	Fisher Brand	22-363-750	Surgery prep
SumnoSuite low-flow anesthesia system	Kent Scientific	SS-01	Surgery
10 mL syringe Luer-Lok Tip	BD Bard-Parker	302995	Head support apparatus
Timers	Fisher Scientific	6KED8	Surgery
Topical anesthetic cream	L.M.X 4	NDC 0496-0882-15	Surgery prep
Triple antibiotic ointment	Major	NDC 0904-0734-31	Post-surgery
Tubing	MasterFlex	96410-16	Head support apparatus
Vaporizer Single Channel Anesthesia System	Kent Scientific	VetFlo-1210S	Surgery prep