Inducing Post-Traumatic Epilepsy in a Mouse Model of Repetitive Diffuse Traumatic Brain Injury

Summary

This systematic protocol describes a new animal model of post-traumatic epilepsy after repetitive mild traumatic brain injury. The first part details steps for traumatic brain injury induction using a modified weight drop model. The second part provides instructions on the surgical approach for single- and multi-channel electroencephalographic data acquisition systems.

Abstract

Traumatic brain injury (TBI) is a leading cause of acquired epilepsy. TBI can result in a focal or diffuse brain injury. Focal injury is a result of direct mechanical forces, sometimes penetrating through the cranium, creating a direct lesion in the brain tissue. These are visible during brain imaging as areas with contusion, laceration, and hemorrhage. Focal lesions induce neuronal death and glial scar formation and are present in 20%−25% of all people who incur a TBI. However, in the majority of TBI cases, injury is caused by acceleration-deceleration forces and subsequent tissue shearing, resulting in nonfocal, diffuse damage. A subpopulation of TBI patients continues to develop post-traumatic epilepsy (PTE) after a latency period of months or years. Currently, it is impossible to predict which patients will develop PTE, and seizures in PTE patients are challenging to control, necessitating further research. Until recently, the field was limited to only two animal/rodent models with validated spontaneous post-traumatic seizures, both presenting with large focal lesions with massive tissue loss in the cortex and sometimes subcortical structures. In contrast to these approaches, it was determined that diffuse TBI induced using a modified weight drop model is sufficient to initiate development of spontaneous convulsive and non-convulsive seizures, even in the absence of focal lesions or tissue loss. Similar to human patients with acquired post-traumatic epilepsy, this model presents with a latency period after injury before seizure onset. In this protocol, the community will be provided with a new model of post-traumatic epilepsy, detailing how to induce diffuse non-lesional TBI followed by continuous long-term video-electroencephalographic animal monitoring over the course of several months. This protocol will detail animal handling, the weight drop procedure, the electrode placement for two acquisition systems, and the frequent challenges encountered during each of the steps of surgery, postoperative monitoring, and data acquisition.

Introduction

Every year TBI affects an estimated 60 million people worldwide. Impacted individuals are at higher risk of developing epilepsy, which can manifest years after the initial injury. Though severe TBIs are associated with a higher risk of epilepsy, even mild TBI increases an individual’s chance of developing epilepsy^1,2,3,4. All TBIs can be classified as focal, diffuse, or a combination of both. Diffuse brain injury, present in many if not all TBIs, is a result of brain tissues of different densities shearing against each other due to acceleration-deceleration and rotational forces. By definition, diffuse injury only occurs in isolation in mild/concussive non-penetrating brain injury, in which no brain lesions are visible on computed tomography scans⁵.

There are currently two critical problems in the management of patients who have, or are at risk of, developing post-traumatic epilepsy (PTE). The first is that once PTE has manifested, seizures are resistant to available anti-epileptic drugs (AEDs)⁶. Secondly, AEDs are equally ineffective at preventing epileptogenesis, and there are no effective alternative therapeutic approaches. In order to address this deficit and find better therapeutic targets and candidates for treatment, it will be necessary to explore new cellular and molecular mechanisms at the root of PTE⁶.

One of the prominent features of post-traumatic epilepsy is the latent period between the initial traumatic event and the onset of spontaneous, unprovoked, recurrent seizures. The events that occur within this temporal window are a natural focus for researchers, because this time window might allow treatment and prevention of PTE altogether. Animal models are most commonly used for this research because they offer several distinct benefits, not the least of which is that continuous monitoring of human patients would be both impractical and costly over such potentially long spans of time. Additionally, cellular and molecular mechanisms at the root of epileptogenesis can only be explored in animal models.

Animal models with spontaneous post-traumatic seizures and epilepsy are preferred over models where seizures are induced after TBI by less physiologically relevant means, such as by chemoconvulsants or electric stimulation acutely, chronically, or by kindling. Spontaneous post-traumatic seizure models test how TBI modifies the healthy brain network leading to epileptogenesis. Studies using additional stimulation after TBI assess how exposure to TBI reduces seizure threshold and affects susceptibility to seizures. The advantages of animal models with seizures induced chemically or with electric stimulation are in testing the specific mechanisms of refractoriness to AEDs and the efficacy of existing and novel AEDs. Yet, the degree of relevance and translation of these data to humans may be ambiguous⁷ due to the following: 1) seizure mechanisms may be different from those induced by TBI alone; 2) not all of these models lead to spontaneous seizures⁷; 3) lesions created by the convulsant agent itself, with the cannula required for its delivery, or by stimulating electrode placement in depth structures (e.g., the hippocampus or amygdala) can already cause increased seizure susceptibility and even hippocampal epileptiform field potentials⁷. Furthermore, some convulsant agents (i.e., kainic acid) produce direct hippocampal lesions and sclerosis, which is not typical after diffuse TBI.

Until recently, only two animal models of post-traumatic epilepsy existed: controlled cortical impact (CCI, focal) or fluid percussion injury (FPI, focal and diffuse)⁸. Both models result in large focal lesions alongside tissue loss, hemorrhage, and gliosis in rodents⁸. These models mimic post-traumatic epilepsy induced by large focal lesions. A recent study demonstrated that repeated (3x) diffuse TBI is sufficient for the development of spontaneous seizures and epilepsy in mice even in the absence of focal lesions⁹, adding a third rodent PTE model with confirmed spontaneous recurrent seizures. This new model mimics cellular and molecular changes induced by diffuse TBI, better representing the human population with mild, concussive TBIs. In this model, the latent period of three weeks or more before seizure onset and the emergence of late, spontaneous, recurrent seizures allows for investigating the root causes of post-traumatic epileptogenesis, testing the efficacy of preventive approaches and new therapeutic candidates after seizure onset, and has potential for the development of biomarkers of post-traumatic epileptogenesis because approximately half of the animals develop post-traumatic epilepsy.

The choice of animal model for the study of post-traumatic epilepsy depends on the scientific question, the type of brain injury investigated, and what tools will be used to determine the underlying cellular and molecular mechanisms. Ultimately, any model of post-traumatic epilepsy must demonstrate both the emergence of spontaneous seizures after TBI and an initial latency period in a subset of TBI animals, because not all patients who incur a TBI go on to develop epilepsy. To do this, electroencephalography (EEG) with simultaneous video acquisition is used in this protocol. Understanding the technical aspects behind data acquisition hardware and approaches is critical for accurate data interpretation. The critical hardware aspects include the type of recording system, type of electrodes (screw or wire lead) and material they are made of, synchronized video acquisition (as part of the EEG system or third party), and properties of the computer system. It is imperative to set the appropriate acquisition parameters in any type of system depending on study goal, EEG events of interest, further analysis method, and sustainability of data storage. Lastly, the method of electrode configuration (montage) must be considered, as each has advantages and disadvantages and will affect the data interpretation.

This protocol details how to use the modified Marmarou weight drop model^10,11 to induce diffuse injury resulting in spontaneous, unprovoked, recurrent seizures in mice, describes surgical approaches to acquire a single- and multi-channel continuous, and synchronized video EEG using monopolar, bipolar, or mixed montage.

Protocol

All animal procedures described in this protocol were performed in accordance with the Institutional Animal Care and Use Committee (IACUC) of Virginia Tech and in compliance with the National Institutes of Health's 'Guide for the Care and Use of Laboratory Animals'.

1. Animal handling protocol

NOTE: This protocol is intended to habituate animals ordered from a vendor to the facility after arrival and to condition them to being handled by the experimenter. This improves animal well-being by reducing stress and anxiety and simplifies certain procedures that require handling animals, including inducing the TBI, post-operative monitoring, and connecting the animal to the acquisition system.

1. When many animals are received from the vendor, ear-tag and randomly assign them to an experimental group (TBI) or control group (sham surgery) while combining them in cages of 2−5 animals. House TBI animals separately from sham animals because sham mice occasionally act aggressively toward mice that underwent TBI.
2. Handling day 1 (24−48 h after ear-tagging): Prepare a chart for logging animal ear tags, date of birth, dates of handling, animal weight on the handling days, duration of the handling, and a section for comments and observations.
3. Gently cup the animal using both hands. Do not grab the animal by the tail as it induces defense mechanisms and a stress response.
4. Check and record the ear tag of the animal.
5. Place the animal in the container on the weight scale and record the weight.
6. Gently cup the animal with both hands again and handle it for 1 min, allowing it to move and explore within the hands. Perform this over a bench in the procedure room and be careful to not drop the animal on the floor.
7. After 1 min of handling, place the animal back in its cage.
8. Repeat steps 1.3−1.7 for the other animals in the cage.
9. Handling day 2 (the following day): Repeat steps 1.2−1.5.
10. Gently cup the animal with both hands again and handle it for 2 min, allowing it to move and explore within the hands. Perform this over a bench in the procedure room and be careful to not drop the animal on the floor.
11. After 2 min of handling, place the animal back in its cage.
12. Repeat steps 1.10−1.11 for the other animals in the cage.
13. Handling day 3 (the following day): Repeat steps 1.2−1.5.
14. Gently cup the animal with both hands again and handle it for 4 min, allowing it to move and explore within the hands. Perform this over a bench in the procedure room and be careful to not drop the animal on the floor.
15. After 4 min of handling, place the animal back in its cage.
16. Repeat steps 1.14−1.15 for the other animals in the cage.
17. Handling day 4 (control day, 1 week from handling day 1): Repeat steps 1.2−1.5.
18. Gently cup the animal with both hands again and handle it for 4 min, allowing it to move and explore within the hands. Perform this over a bench in the procedure room and be careful to not drop the animal on the floor.
19. After 4 min handling, place the animal back in its cage.
20. Repeat steps 1.18-1.19 for the other animals in the cage.
    NOTE: The control handling day tests the retention of the calm behavior after a three-day handling protocol.

2. Weight drop procedure

1. Place the mouse in an induction chamber. Set the flow of oxygen and vacuum both to 1 L/min and the level of isoflurane gas to 3%−5%. Anesthetize the mouse for 5 min.
2. Remove the mouse from the induction chamber and place it on a foam pad. Test for the absence of a response to a toe or tail pinch.
3. Administer an analgesic (0.1 mg/kg buprenorphine) subcutaneously. If the EEG surgery is performed that same day, administer the buprenorphine subcutaneously in combination with the non-steroidal anti-inflammatory carprofen (5 mg/kg).
4. Administer the sodium lactate solution (3 µL per gram of the animal's weight) subcutaneously before or after the last impact. The sodium lactate solution can be mixed with the analgesics for quick administration in a single injection.
   NOTE: The sodium lactate solution contains a mixture of sodium chloride, potassium chloride, calcium chloride, and sodium lactate in water. This step helps to replace fluids and electrolytes, aiding recovery.
5. Position the head of the mouse under the weight drop tube (Figure 1A) and place a flat stainless steel disc (1.3 cm diameter, 1 mm thick, and 880 mg weight) in the center of the head, between the line of the eyes and ears.
   NOTE: This disc diffuses the impact across the surface of the skull (Figure 1B).
6. Remove the pin in the weight drop tube to release the 100 g weight rod from a height of 50 cm. To induce the sham injury for the control mice, remove the weight rod from the tube to prevent accidental release of the pin and weight drop.
   NOTE: The animal's head must be positioned flat, so that the rod free-falls on the entire surface of the disc.
7. Place the unconscious animal on its back for recovery on a heating pad covered with a sterile polylined absorbent towel. The righting reflex recovery time (i.e., the time it takes the mouse to right itself from its back) can be measured as a readout for the time spent unconscious.
8. When the animal regains consciousness, place it in a clean cage that has been warmed on a heating pad, with recovery gel and a few moistened chow pieces to recover for 45 min. Make sure there is sufficient litter so the cage does not get overheated. Overheating the animal can prove just as great an obstacle to recovery as allowing the mouse to become too cold.
9. After 45 min, repeat steps 2.1−2.8 twice, omitting step 2.3 (i.e., administration of analgesics and anti-inflammatory drugs).
10. Allow the animals to recover for 1−2 h if EEG electrode implantation surgery is performed on the same day.

3. Surgical field preparation for implantation of EEG electrodes

NOTE: Autoclave the surgical tools and screws prior to surgery. Clean the surgical gloves by spraying and rubbing with 70% ethanol before and after touching the animal, non-sterile materials, and in between handling the animals. Sterilize the surgical tools for 2−3 min in the bead sterilizer (see Table of Materials) between animals. Change the sterile drape before placing a new animal into the stereotactic apparatus. Ensure that the surgical field contains all the necessary components for the surgery (Figure 2). The absence of an invasive surgical procedure to induce the TBI in this model has several advantages: 1) implantation of the electrodes is flexible and may be performed on the same day as TBI or after a defined period of time; 2) the animal's recovery time is faster; 3) the cranium remains intact, allowing more surface area and flexibility for implanting electrodes.

1. Anesthetize the mouse in 3%−5% isoflurane gas in an induction chamber for 5 min.
2. Transfer the mouse from the induction chamber to the stereotactic apparatus and place it on a sterile drape on a heating pad with isoflurane gas and vacuum tubes connected to the nose cone.
3. Maintain the body temperature at 37 °C over the course of the surgery. Place the temperature sensor so that it makes contact with the chest or abdominal wall of the mouse.
4. Fix the animal's head in place using the ear bars.
5. Maintain the anesthesia at 1.5%−3.5% isoflurane or at ~60 breaths/min in the surgical plane (with no response to toe or tail pinch).
6. Apply an eye ointment to the animal's eyes to keep them lubricated throughout the surgery.
7. Administer a mixture of analgesics (0.1 mg/kg buprenorphine) and the non-steroidal anti-inflammatory drug (5 mg/kg carprofen) in a single injection subcutaneously unless the TBI was performed earlier during the day, in which case the animal already received analgesics and anti-inflammatories.
   NOTE: Buprenorphine should be administered again if the time between the first TBI and EEG placement surgery exceeds 8 h or if the animal displays signs of pain 8 h after the first administration, but it should be given without the addition of carprofen.
8. Administer sodium lactate solution (3 µL per gram of the animal's weight) subcutaneously to replace fluids and electrolytes in the animal.
   NOTE: If surgery is performed immediately after the TBI, this step has to be timed properly. Sodium lactate solution should be administered every 2 h while the animal undergoes the procedures and once after the surgery, 2 h from the previous injection.
9. Remove the hair from the scalp using a hair removal cream.
10. Before making the incision, disinfect the skin of the scalp with povidone-iodine surgical antiseptic solution and 70% ethanol in alternating swabs with sterile gauze pads in a circular motion 3x (20 s per solution each time).
11. Using a scalpel, make a rostral-caudal incision on the scalp midline from just above the eyes to the back of the head. This method of scalp opening is preferred over cutting the scalp off, as skin flaps can be sealed over or around the EEG-cap providing more stability.
    NOTE: When preparing the skull for implantation of the 3-EEG headmount, cutting the scalp off is required, as the size of the headmount will not allow for closure of the skin flaps over the headmount.
12. Expand the area of incision by applying small hemostats on the opened skin borders. If any bleeding occurs after the incision, clean with a sterile cotton gauze or swab.
13. Gently remove the periosteum (i.e., the thin membrane over the cranial bone) with a scalpel blade. If any bleeding occurs during this step, press on bleeding site with a sterile cotton swab until it stops.
14. Use sterile cotton swabs to clean the cranium with hydrogen peroxide, but avoid touching the soft tissue surrounding the exposed cranial area. Repeat this step until the cranium is cleaned from any soft tissue and has a whitish appearance.
15. Dry the cranium with a sterile gauze or cotton swab.
    NOTE: Steps 3.12−3.15 are important for the proper fixation of the electrodes and dental cement. Any soft tissue, non-cauterized bleeding, and debris can cause infection, unstable headmount fixation, distorted or absent signal, and loss of the implant within several days or weeks after surgery.

4. Electrode placement

1. Implant the single EEG (1EEG) channel headmount.
   NOTE: Abbreviations in the stereotactic coordinates represent spatial relationships and specify the distance in millimeters of the target from the bregma at a given orientation on the animal's head: anterior-posterior (AP) and medial-lateral (ML). Dorsal-ventral is not applicable in this protocol because all electrodes are placed into the epidural space rather than in a certain structure within the brain (Figure 3). Vin+ is an active electrode and Vin- is its reference electrode.
   1. Use a high-speed drill with a steel bit (0.5 mm, round, ¼ in.) at ~5,000−6,000 rounds per min (rpm) to create six burr holes (three for stability screws and three for electrodes) using the provided stereotactic coordinates¹². For the two anterior screws: AP = +1.5 mm, ML = ±1.5 mm; for the one posterior screw: AP = -5.2 mm, ML = -1.5 mm; for the ground electrode: AP = -5.2 mm, ML = +1.5 mm; for the recording electrodes: AP = -2.3 mm, ML = ±2.7 mm, with Vin+ to the right and Vin- to the left.
   2. Add three screws for enhanced stability of the head stage. Using a screwdriver, turn screws 1−1.5 x each to be fixed stably in the cranium.
      NOTE: Placing the screws deeper will damage the brain.
   3. Insert the 1EEG headmount into a stereotactic holder arm and position the headmount so that the three electrodes are located along the cranial midline. In this configuration the ground electrode and its respective opening on top of the headmount is in the back, the Vin+ electrode in the middle, and the Vin- electrode in the front. A mark can be made on the headmount with a permanent marker.
   4. Bend each electrode 90° so that the end of each wire is bent downwards and is positioned above the corresponding burr hole. Then, measure out 1 mm length of the portion of the wire that is now perpendicular to the burr hole and trim the excess off (Figure 3). This will ensure epidural placement of the electrodes. The electrodes should be barely touching the dura mater surface.
   5. Lower the headmount and adjust all three electrodes to match the respective burr hole. For epidural recording, the electrodes must be placed above or barely touching the dura mater.
   6. Prepare dental cement for application by mixing a ½ scoop of powder with several drops of solvent. Use a mixing spatula and stir until the final mixture is putty-like, tacky but malleable, and stiff enough to be properly condensed when placed on the animal's cranium.
   7. Apply dental cement mixture covering all screws and electrodes and wait ~3−5 min for it to solidify. Make sure not to cover the plastic pedestal with dental cement, because it will make it impossible to connect the animal to the commutator with a tether.
   8. Release the hemostats holding the skin flaps and close the incision by connecting the skin flaps around the plastic pedestal. Apply several drops of tissue adhesive (see Table of Materials) to seal the skin flaps.
   9. Apply chlorhexidine antiseptic to the area around the implant to avoid infection. If the animal is under anesthesia for longer than 2 h after the previous injection of sodium lactate solution, given during the TBI induction, administer another injection subcutaneously. To maintain proper hydration of the animal, repeat the injection every 2 h that the animal spends under anesthesia.
   10. After the surgery, give a final injection of sodium lactate solution 2 h after the previous injection. If the surgery is less than 2 h long, administer the final recovery dose of the sodium lactate solution 2 h from the first injection.
   11. Remove the animal from the stereotactic apparatus and measure the animal's weight after the EEG surgery as a reference for future monitoring. Due to the implant, the animal's weight will be greater than before surgery.
   12. Place the animal in a clean cage on a warm heating pad for recovery.
2. Implant the two EEG and one EMG (2EEG/1EMG) channels headmount.
   1. Use the bregma as a landmark for placement of the headmount. Apply a small amount of tissue adhesive (see Table of Materials) to the bottom side of the 2EEG/1EMG headmount, avoiding the four screw holes and place the 2EEG/1EMG headmount on the surface of the cranium.
      NOTE: There are no specific coordinates for placement of this headmount. The headmount is 8 mm long and 5 mm wide, which covers most of the cranial surface. Positioning the headmount with its front edge ῀3.0 mm anterior to the bregma is optimal and provides good signal quality. Quick manual placement is necessary before the drop of tissue adhesive cures. Allow approximately 5 min for tissue glue to cure completely.
   2. Use a sterile 23 G needle to create pilot holes for the screws through the four openings in the headmount. To accomplish this, gently push the needle and slowly rotate until the tip of the needle penetrates the skull without damaging the brain. Remove any bleeding from the pilot holes using a sterile cotton swab.
   3. Insert the 0.10 in screws in the pilot holes and rotate them until each is fixed in the skull. This can be up to half of the screw length, but not the full length, as this would damage the dura mater and cortex. If the headmount is positioned so that there is a gap between the skull surface and the rear end of the headmount use two 0.12 in screws in the posterior part.
   4. Make small opening on the sides of the two-component epoxy (silver-epoxy) twin-pack pouch. Take a double-sided spatula and use each side to scoop a small and equal amount of each component from the pouch and mix them together. Use only a small amount sufficient for a single surgery, because the mixture solidifies within 20 min. Seal the sides of the pouch to prevent drying.
      NOTE: The silver-epoxy allows for proper electrical contact between the screw and headmount and enhances the stability of the screws.
   5. Apply a small amount of this mixture between screwhead and screw hole, then tighten each screw until its head rests on the base of the implant. Ensure that no silver-epoxy is making contact between the two screws because each screw serves as an individual electrode and, to ensure an accurate signal, it should not make contact with the other screw.
   6. If the silver-epoxy mixture was misplaced, there is a few second time window to carefully scoop out the excess to separate the connection. Carefully bend both EMG leads from the posterior edge of the headmount to follow the contour of the animal's head and neck, and then insert them into the nuchal muscles.
   7. Prepare dental cement for application by mixing a ½ scoop of powder with several drops of solvent. Use a mixing spatula and stir until the final mixture is putty-like, tacky but malleable, and stiff enough to be properly condensed when placed on the animal's cranium.
   8. Apply dental cement mixture covering the entire headmount while avoiding covering the six pin holes, as this will make it impossible to connect the pre-amplifier. Wait ~3−5 min for the cement to solidify. Make sure that the skin is not sealed to the headmount with dental cement.
   9. Release the hemostats holding the skin flaps and close the incision by connecting the skin flaps around the plastic pedestal. Apply several drops of tissue adhesive to seal the skin flaps.
      NOTE: If the skin incision was made longer to allow for straightening of the EMG wire leads, the skin can be sealed with tissue adhesive or sutured. Sealing the skin with tissue adhesive is usually sufficient. However, if during post-operative monitoring opening of the incision is observed, sutures are recommended instead.
   10. Apply chlorhexidine antiseptic to the area around the implant to avoid infection. Administer sodium lactate solution (3 µL per gram of the animal's weight) subcutaneously to replace fluids and electrolytes if the animal is under anesthesia for longer than 2 h after the previous injection.
   11. Remove the animal from the stereotactic apparatus and measure the animal's weight after the EEG surgery as a reference for future monitoring. Due to the implant, the animal's weight will be greater than before surgery.
   12. Place the animal in a clean cage on a warm heating pad, with recovery gel and a few moistened chow pieces for recovery.
3. Implant a three EEG channels (3EEG) headmount.
   1. Use high-speed drill with a steel bit (0.5 mm, round, ¼) at ~5,000−6,000 rpm to create six burr holes (three for stability screws and three for electrodes) using the provided stereotactic coordinates¹². For ground and common reference for EEG1 and EEG2: AP = 5.2 mm, ML = ±1.5 mm; for EEG1 and EEG2: AP = -3.0 mm, ML = ±3.0 mm; for independent EEG3: AP =-1.4 mm, ML = ±1.5 mm.
   2. Place the six screw electrodes into the burr holes.
      NOTE: Placing the screws deeper will create significant damage to the brain. Screw electrodes provide better stability of the headmount.
   3. Prepare dental cement for application by mixing a ½ scoop of powder with several drops of solvent. Use a mixing spatula and stir until the final mixture is putty-like, tacky but malleable, and stiff enough to be properly condensed when placed on the animal's cranium.
   4. Apply dental cement mixture covering the entire exposed surface of the cranium and each screw electrode. Make sure that skin is not sealed to the headmount with dental cement. Wait ~1−2 min for the cement to mildly solidify. There is no need to wait until full solidification before proceeding to the next step.
   5. Turn on the soldering iron to heat it up. Place the 3EEG headmount in a stereotactic holder arm.
      NOTE: Position the headmount so that the six wire lead positions match the position of the wire leads of each screw electrode.
   6. Lower the headmount so that its ventral part rests on top of the dental cement.
   7. Twist the wire of each lead from each of the screw electrodes with the corresponding wire lead of the headmount.
      NOTE: Twisting the wrong wire leads will make data interpretation complicated or impossible.
   8. Carefully trim the excess wire off using scissors. Solder each twisted pair of wire for proper signal conduction.
      NOTE: Each pair of wires must make contact with another pair, otherwise signal quality and data interpretation will be compromised.
   9. Bend each soldered pair of wire leads around the headmount, avoiding contact between each pair.
      NOTE: If the wire leads are not trimmed short enough it can be difficult to bend them around the headmount without touching another wire. In this case, bend one pair first, cover it with dental cement mixture, wait ~1−2 min to solidify, then proceed with the next pair in the same fashion.
   10. Finish covering all the wire with dental cement leaving only the black portion of the headmount exposed.
       NOTE: Be careful to not apply any dental cement powder or mixture to the top of the exposed portion of the headmount as any debris or cement in the holes will block the contact and will lead to either signal absence or noise.
   11. Release the hemostats holding the skin flaps. Apply chlorhexidine antiseptic to the area around the implant to avoid infection.
   12. Administer sodium lactate solution (3 µL per gram of the animal's weight) subcutaneously to replace fluids and electrolytes if the animal has been under anesthesia for longer than 2 h after the previous injection.
   13. Remove the animal from the stereotactic apparatus and measure the animal's weight after the EEG surgery as a reference for future monitoring. Due to the implant, the animal's weight will be greater than before surgery.
   14. Place the animal in a clean cage on a warm heating pad, with recovery gel and a few moistened chow pieces for recovery.
       NOTE: Hydrogen peroxide aids in removing of the any remaining soft tissue from the cranium.

5. Connecting animals to the acquisition system

1. Cup the animal with both hands to remove it from the acquisition cage and transfer it to a clean area with a flat surface, like an Animal Transfer Station (ATS).
2. Gently grab the mouse by the skin of its back. Do not grab the animal by the tail, as this causes distress.
3. Identify the opening in the EEG headmount corresponding to the ground electrode and match the respective pin of the tether for proper connection.
   NOTE: Reverse connection of the tether from the commutator to the animal headmount will result in a different reading from the electrodes and potentially distorted waveforms.
4. Return the animal to the acquisition cage and connect the other end of the tether (EEG System 1) or pre-amplifier (EEG System 2) to the commutator.
   NOTE: When connecting the pre-amplifier (EEG System 2) to the tether from the commutator, match the white marks on the ends of both tethers. Reverse connection will result in permanent damage of the amplifier and requires repairs by the manufacturer, which are expensive.
5. Gently rotate the tether connecting the animal to the commutator to ensure the mechanism works properly and the animal can move freely.

6. EEG data acquisition settings

1. Set EEG System 1 acquisition parameters.
   1. Set sampling rate to 500 Hz; gain 5,000; mode Norm 35 Hz; LPN off. Set high pass filter to 0.5 Hz.
      NOTE: 100 Hz (low pass) is built-in and does not require manual input.
2. Set EEG System 2 acquisition parameters.
   1. Set sampling rate to 600 Hz; preamp gain 100; gain 1 (EEG1,2). Set low pass filter to 100 Hz.
      NOTE: 1 Hz (high pass) is built-in and does not require manual input.

7. Video data acquisition settings

1. Set acquisition parameters for EEG System 1.
   NOTE: A third party video acquisition system is needed for obtaining simultaneous video data.
   1. Set frame rate between 15 (minimum recommended) and 30 (maximum available) for appropriate video quality. Set the resolution to 640 x 640 pixels. Set type of compression to H.264H.
2. Set acquisition parameters for EEG System 2.
   NOTE: This EEG system offers a video system and software which synchronize video and EEG data together in a single file for up to four animals (see Table of Materials).
   1. Set frame rate between 15 (minimum recommended) and 30 (maximum available) for appropriate video quality. Set the resolution to 640 x 480 pixels. Set the type of compression to the WebM file format.

Results

The protocol outlined here describes the method for induction of a diffuse injury in isolation (e.g., in the absence a focal lesion) using a mouse model of repetitive diffuse TBI (Figure 1). Figure 1A depicts the weight drop device and its components (Figure 1A, a1−a5) used for induction of TBI in this model and crucial steps during the procedure (Figure 1...

Discussion

In contrast to CCI and FPI models inducing either focal or combination of focal and diffuse injury, the model of repetitive diffuse TBI described in this protocol allows for the induction of diffuse injury in the absence of focal brain injury and does not require scalp or cranial openings and the associated inflammation. An added benefit of the absence of craniectomy in this model is that it allows to not only implant the electrodes for chronic continuous EEG recording, but also the creation of a thinned-skull cranial wi...

Materials

Name	Company	Catalog Number	Comments
0.10" screw	Pinnacle Technology Inc., KS, USA	8209	0.10 inch long stainless steel
0.10" screw	Pinnacle Technology Inc., KS, USA	8403	0.10 inch long with pre-soldered wire lead
0.12" screw	Pinnacle Technology Inc., KS, USA	8212	0.12 inch long stainless steel
1EEG headmount	Invitro1 (subsidiary of Plastics One), VA, USA	MS333/8-A/SPC	3 individually Teflon-insulated platinum iridium wire electrodes (twisted or untwisted, 0.005 inch diameter) extending below threaded plastic pedestal
2EEG/1EMG headmount	Pinnacle Technology Inc., KS, USA	8201	2EEG/1EMG channels
3% hydrogen peroxide			Pharmacy
3EEG headmount	Pinnacle Technology Inc., KS, USA	8235-SM-C	custom 6-Pin Connector for 3EEG channels
Buprenorphine	Par Pharmaceuticals, Cos. Inc., Spring Valley, NY, USA	060969
Buprenorphine	Par Pharmaceuticals, Cos. Inc., Spring Valley, NY, USA	060969
C57BL/6 mice	Harlan/Envigo Laboratories Inc		male, 12-16 weeks old
C57BL/6 mice	The Jackson Laboratory		male, 12-16 weeks old
Carprofen	Zoetis Services LLC, Parsippany, NJ, USA	026357	NOTE: this drug is added during weight drop only if stereotactic electrode implantation will be performed on the same day
Chlorhexidine antiseptic			Pharmacy
Dental cement and solvent kit	Stoelting Co., USA	51459
Drill	Foredom	HP4-917
Drill bit	Meisinger USA, LLC, USA	HM1-005-HP	0.5 mm, Round, 1/4, Steel
Dry sterilizer	Cellpoint Scientific, USA		Germinator 500
EEG System 1	Biopac Systems, CA, USA
EEG System 2	Pinnacle Technology Inc., KS, USA
Ethanol ≥70%	VWR, USA	71001-652	KOPTEC USP, Biotechnology Grade (140 Proof)
Eye ointment	Pro Labs Ltd, USA		Puralube Vet Ointment Sterile Ocular Lubricant available in general online stores and pharmacies
Fluriso liquid for inhalation anesthesia	MWI Veterinary Supply Co., USA	502017
Hair removal product	Church & Dwight Co., Inc., USA		Nair cream
Isoflurane	MWI Veterinary Supply Co., USA	502017
Povidone-iodine surgical solution	Purdue Products, USA	004677	Betadine
Rimadyl/Carprofen	Zoetis Services LLC, Parsippany, NJ, USA	026357
Solder			Harware store
Soldering iron	Weller, USA	WP35	ST7 tip, 0.8mm
Stainless steel disc			Custom made
Sterile cotton swabs
Sterile gauze pads	Fisher Scientific, USA	22362178
Sterile poly-lined absorbent towels pads	Cardinal Health, USA	3520
Tissue adhesive	3M Animal Care Products, USA	1469SB