A Novel and Translational Rat Model of Concussion Combining Force and Rotation with In Vivo Cerebral Microdialysis

The animal protocol for this project obtained the approval from the Animal Care Committee of the Hopital du Sacre-Cœur de Montreal in compliance with the guidelines of the Canadian Council on Animal Care.

NOTE: A schematic outline of the research protocol is presented in Figure 1.

1. Animal Preparation

1. Order Sprague-Dawley rats from a standard laboratory animal supplier to be delivered between 43 and 50 days of age and at a weight between 151 and 200 g.
2. House all rats individually in a cycle of 12:12 h light: darkness, at 24-26 °C with ad libitum access to water and food.
3. During the 2 weeks before starting the protocol, handle the rats for 5 min on a daily basis to facilitate their habituation in contact with researchers. Rats should be aged about 10 weeks old and their weight should be between 295 and 351 g at the time of concussion or sham injury induction.

2. Microdialysis Guide Cannula Implantation Surgery

1. Perform the surgery under sterile conditions. Wear sterile gloves, a hair bonnet and a surgical mask throughout the procedure. Autoclave and sterilize all the materials and surgical instruments beforehand. Clean and disinfect the working area and stereotaxic apparatus thoroughly with a solution of ethanol (70%).
2. Anesthetize the animals by injecting a cocktail of ketamine (70 mg/kg) and xylazine (10 mg/kg) intraperitoneally. Asses anesthetic depth by testing the reflex to a toe pinch.
3. Remove fur from the head of the animal using electric clippers. Clean shaved head using a solution of 2% isopropyl alcohol and 2% chlorhexidine gluconate (3 times). Apply lubricating eye ointment during the anesthesia to prevent dryness.
4. Drape-off the surgical field so that only the head of the animal is exposed. Place the head of the rat in a stereotaxic apparatus, insert the ear bars into the ear canals with great care then tighten the nose clamp. Fix a 26 G stainless-steel guide cannula to the holder arm on the stereotaxic apparatus.
5. Locally inject an anesthetic cocktail of bupivacaine (1.5 mg/kg) and lidocaine (1.5 mg/kg) subcutaneously on the head, 10 min prior to incision.
6. Maintain anesthesia during the whole procedure by delivering sodium isoflurane (2.5%) at 0.5 L/min oxygen flow with a nose cone.
7. Make a midline incision (3 cm) along the scalp with a scalpel. Leave the skull clear by installing 4 clamps around the incision.
8. Scrape firmly the periosteum from the skull with a surgical blade until the Bregma and Lambda sutures are visible. Maintain firm pressure on the skull with a gauze pad or cotton tipped applicator if there is bleeding.
9. Confirm if the skull is correctly aligned on the stereotaxic apparatus by comparing the dorsoventral coordinates of the Bregma and Lambda sutures. Identify the anteroposterior, mediolateral and dorsoventral coordinates of the Bregma suture as the reference points for the coordinates of the guide cannula.
10. Taking the Bregma suture coordinates as references, calculate the coordinates of the guide cannula implantation site in the hippocampus.
    NOTE: The following coordinates were determined according to the rat brain atlas from Paxinos and Watson (anteroposterior: -0.60 cm; mediolateral: ±0.58 cm; dorsoventral: -0.16 cm, Figure 2A)³².
11. Mark the precise implantation site using a marker.
12. Drill a 0.5 mm hole through the cranium at the target site of the guide cannula. Drill 3 other holes approximately 5 mm around this point to thread 3 anchor screws into the skull that will solidify the cannula after acrylic dental cement is applied.
13. Insert the cannula into the hippocampus and fix it with dental cement. This cannula will be used to insert the probe into the region of interest 7 days later during the microdialysis procedure. Be careful as to not spill excess dental cement around the site where the weight will be dropped.
14. Leave the cement to dry for 2 min, then remove the holder arm from the cannula. Insert a stainless-steel removeable obturator into the cannula to avoid cerebrospinal fluid seepage and risks of infection.
15. Remove the 4 clamps, pull back the retracted skin and stitch it with a surgical suture thread 4-0.
16. Remove the rat from the apparatus and inject buprenorphine subcutaneously to treat pain (0.05 mg/kg, after surgery then once per day during the following 2 days). Place the rodent back in its cage with a heating pad under until it becomes conscious, then return it to the animal care facility for a 7 days recovery period under close monitoring.

3. Microdialysis Procedure

1. While performing the microdialysis procedure, wear sterile gloves, a hair bonnet and a surgical mask. Seven days following the cannula implantation surgery, anesthetize the rat with sodium isoflurane (2.5%) at 0.5 L/min oxygen flow.
2. Remove the obturator from the cannula and insert slowly a microdialysis probe, perfused with artificial cerebral spinal fluid (ACSF) (26 mmol/L NaHCO₃, 3 mmol/L NaH₂PO₄, 1.3 mmol/L MgCl₂, 2.3 mmol/L CaCl₂, 3.0 mmol/L KCl, 126 mmol/L NaCl, 0.2 mmol/L L-ascorbic acid), through the cannula into the hippocampus or other region of interest.
   NOTE: Rats need to be anesthetized only while removing the obturator and inserting the microdialysis probe, and during induction of concussion or sham injury. The probes used here are laboratory-constructed, I-shaped, and comprised of fused side-by-side silica inlet-outlet lines [internal diameter (ID): 50 μm] encased in polyethylene tubing (ID: 0.58-0.38 mm). The end of the cannula is secured with a length of regenerated hollow cellulose membrane [molecular weight cut-off: 13 kDa, outer diameter (OD): 216 μm; ID: 200 μm] using cyanoacrylate adhesive and the tip sealed with epoxy. The active membrane measures 2.5 mm for implantation in the hippocampus but can be adjusted according to the depth of the region of interest. The connection of the indwelling cannula of the rat to the probe is secured with a fitted, threaded stainless-steel collar.
3. Fix the probe assembly to a stainless-steel spring tethered to a liquid swivel and counter balance lever arm suspended above the cage with a ring stand and clamps so that the animal can move freely within its cage. Tethered rats spend the entire duration of the microdialysis procedure with ad libitum access to water and food.
4. Use a microinfusion pump to send perfusate to the probes, and collect the dialysate from the fused silica outlet line (dead volume: 0.79 μL).
5. At least 1 h and 30 min before the procedure begins, turn up the probe to its working flow rate (1 μL/min). Verify that the flow rate of the probe is consistent by measuring volume over time with a pipette.
   NOTE: The flow rate can be more or less depending on the neurotransmitters sampled and the brain region of interest. Dialysis samples are taken before, during, and after concussion or sham injury induction. Sampling interval depends on the brain region of interest, neurotransmitters being analyzed, dialysate concentrations of the analyte, and sensitivity of the analytical chemistry equipment used. The collecting phases done here in the hippocampus for glutamate and GABA sampling are as follows:
   1. Baseline: At beginning of experiment, collect dialysis samples at 10 min intervals for 60 min.
   2. Post-concussion or sham injury: After concussion or sham injury, collect samples for an additional 90 min (9 samples).
6. Collect each dialysate sample in a fraction vial preloaded with 1 μL of 0.25 mol/L perchloric acid to prevent analyte degradation. Store the samples at 4 °C for subsequent analysis.
7. Following the collection of the last dialysate sample, re-anesthetize the rat with a nose cone delivering sodium isoflurane (2.5%) at 0.5 L/min oxygen flow.
8. Remove the microdialysis probe from the cannula, re-insert the obturator and then return the rat to the animal care facility.

4. Concussion Apparatus Installation

1. Prior to commencement of procedure, carve a weight to be used to inflict the concussion (19 mm in diameter) from solid brass to obtain a mass of 450 g. Insert a metal loop at the top of the brass weight. Drill holes preliminary at a distance of 1.0 m inside a vertical polyvinyl chloride (PVC) guide tube.
2. Slit an aluminum sheet with a sharp razor blade. The slotted aluminum sheet should support the weight of the rat (295 to 351 g) without interfering with the acceleration of its body after head impact from the brass weight.
3. Tape the slotted aluminum sheet tightly to a U-shaped Plexiglas frame (38 cm long x 27 cm wide x 30 cm deep, Figure 3A,B) that contains a foam cushion (37 cm long x 26 cm wide x 12 cm deep).
4. Position the Plexiglas frame under a PVC guide tube (20 mm diameter x 1.5 m length).
5. Hold the PVC guide tube in place with a clamp stand 3.5 cm above the slotted aluminum.
6. Attach a nylon fly fishing line (capacity of 9.1 kg, 0.46 mm diameter) through the metal loop so that the bottom of the weight hangs 2.5 cm over the slotted aluminum as to prevent multiple hits when the rat is falling on the foam cushion following impact.
7. Attach the nylon fly fishing line to the clamp stand.
8. Pull up the weight through the PVC tube with the nylon fly fishing line then keep it in place by inserting a hex key through the preliminary drilled holes at 1.0 m.

5. Concussion Induction

1. After the baseline phase of the dialysis samples collection, re-anesthetize the rat lightly by placing a nose cone delivering sodium isoflurane (2.5% isoflurane at 0.5 L/min oxygen flow) until it as no response to a toe pinch (as mentioned in section 3.1).
2. Place the animal on its chest on the slotted aluminum sheet so that its head is positioned directly in the path of the brass weight (Figure 3C,D). Maintain anesthesia with the nose cone to make sure that the rat does not move or wake-up before the weight strikes it.
3. Remove the nose cone and pull the hex key. The weight will fall vertically through the PVC tube and impact the head of the rat. The rat will undergo a rapid 180° rotation and land on its back (Figure 3E).
4. Remove the rat from the foam cushion and place it on its back in its cage.
5. Use a digital timer to measure the righting reflex time as a sign of recovery and injury severity. The righting reflex time is the total time from the impact until the rodents wake up and spontaneously right themselves to the prone position from the supine position, or start walking. Note any signs of death, fracture, or bleeding.
   NOTE: The procedure can be repeated on the same subject at different time points for repeated concussions.

6. Sham Induction

1. After the baseline phase of the dialysis samples collection, re-anesthetize the rat lightly by placing a nose cone delivering sodium isoflurane (2.5%) at 0.5 L/min oxygen flow until it as no response to a toe pinch (as mentioned in section 3.1).
2. Place the animal on its chest on the slotted aluminum sheet so that its head lays directly in the path of the brass weight. Maintain anesthesia with the nose cone to make sure the rat does not move or wake-up.
3. Remove the nose cone and remove the animal from the aluminum sheet without pulling the hex key. The rat will not undergo a rapid 180° rotation.
4. Place the rat on its back in its cage.
5. Use a digital timer to measure the righting time as an indicator of neurologic restoration.

7. High-performance Liquid Chromatography

1. Determine neurotransmitter levels (i.e., glutamate and GABA) by precolumn derivatization using high-performance liquid chromatography with rapid separation fluorescence detection, and a system consisting of a rapid separation autosampler and a pump coupled to a 3.0 x 50 mm 5 μm analytical column.
2. Prepare a mobile phase with 100 mmol/L sodium phosphate dibasic (Na₂HPO₄), 3.5% acetonitrile and 20% methanol. Adjust the pH to 6.7 with phosphoric acid (85%) as needed.
3. Set the flow rate to 0.5 mL/min.
4. Prepare fresh daily derivatization reagents and working standards (100 ng/mL) from stock solutions. Load them into a refrigerated (10 °C) rapid separation autosampler with samples.
5. Mix each fraction sequentially into the analytical column with 20 μL of 3-mercaptopropionic acid (0.071 mol/L) diluted with H₂O and 20 μL of o-phthaldehyde (0.0143 mol/L) diluted with 0.1 mol/L sodium tetraborate. Allow 10 min for the mix to react.
6. To prevent contamination of next samples, flush the injection loop with methanol (20%), following each injection.
   NOTE: The glutamate retention time would be of 1 min approximately in this protocol, for a total run time of 30 min for each sample.
7. During analysis of chromatographic peaks, identify unknown peaks using samples matched according to time of retention from known standards. Express levels of analytes as μg/mL.

8. Histology

1. A month following the microdialysis procedure and concussion or sham injury induction, anesthetize the animals by injecting a cocktail of ketamine (70 mg/kg) and xylazine (10 mg/kg) intraperitoneally and euthanize them by paraformaldehyde (4%) and saline intracardiac perfusion.
2. Decapitate the rodents then dissect the brains.
3. Store the brains in paraformaldehyde (4%) and subsequently cryoprotect them in a solution of sucrose (30%).
4. Slice the brains in coronal sections of 50 μm with a cryostat.
5. Stain the brain slices with cresyl violet for histological verification of injury and probe placement (Nissl staining).