Generation and On-Demand Initiation of Acute Ictal Activity in Rodent and Human Tissue

All research involving patients was performed under a protocol approved by the University Health Network Research Ethics Board in accordance with the Declaration of Helsinki. Procedures involving animals were in accordance with guidelines by the Canadian Council on Animal Care and approved by the Krembil Research Institute Animal Care Committee.

1. Protocol I: Acute In vitro Seizure Model

1. Preparation of dissection solutions and artificial cerebrospinal fluid
   1. Carbogenate ultrapure water (which is water filtered with a resistivity of 18.2 MΩ·cm) with carbogen (95% O₂/5% CO₂) for 5 min using air stone bubble diffusers (aerators for aquariums).
      Note: Air stones can be connected to the carbogen tank’s regulator via standard silicone tubing. If air stones are not available, seal shut the end of the silicone tubing and poke small holes in the seal to allow the carbogen gas to bubble out and carbogenate the solution.
   2. Prepare a dissection solution by dissolving the solutes from Table 1 in ultrapure water. Add CaCl₂ to a solution that has been carbogenated for at least 5 min to help it dissolve.
      Note: Alternatively, add CaCl₂ first, so that it can dissolve easily in an unsaturated solution. The solution, in liquid form, should be 300 – 320 mOsm/L and have a pH of 7.4 after being saturated with carbogen.
      1. Chill the dissection solution to 0 – 4 °C. Leave the solution in the fridge overnight or place the solution in the freezer for 1 h and continually monitor the temperature.
         Note: The dissection solution can be stored for up to 3 nights; discard afterward.
      2. Carbogenate the dissection solution for 5 – 10 min before use.
         Note: Ideally, the solution should either appear as slush or be transitioning to slush prior to use.
   3. Prepare rodent artificial cerebrospinal fluid (ACSF) by dissolving the solutes from Table 2 (or Table 3 for human ACSF) in ultrapure water. Carbogenate the ACSF solution while it is in a water bath heated to 35 °C until use. Add CaCl₂ to a solution that has been carbogenated for at least 5 min to help it dissolve.
      Note: Alternatively, add CaCl₂ first so that it can dissolve easier in an unsaturated solution. The solution, in liquid form, should be 290 – 320 mOsm/L and have a pH of 7.4 after being saturated with carbogen. Solutions should be made in 1, 2, or 4 L volumes depending on the duration of the experiments. Make 4 L for a full day (8 h) of experiments. The ACSF should be made fresh each day; do not store it for longer than 1 d.
2. Mouse dissection to collect brain tissue
   Note: For human tissue, proceed to the next step (step 1.3) on making brain slices with the vibratome. The cube-shaped blocks (1 cm³) of brain tissue should be obtained from the neurosurgeon using procedures previously described^26,27.
   1. Collect all the tools and materials necessary for animal dissections. Set up the workspace to prepare for animal dissections.
      1. Check the carbogen tank (95%O₂/5%CO₂) to ensure it is not empty. Replace the gas cylinder prior to experiments if there is less than 500 psi of pressure remaining.
      2. Calibrate the vibratome (vibrating blade tissue slicer) according to the manufacturer’s instruction manual. Adjust the vibratome setting to have a cutting amplitude of 1 mm and a cutting speed of 0.12 mm/s.
         Note: The optimal settings may differ for each individual vibrating tissue blade cutter; however, in general, the amplitude should be high and the cutting speed should be low.
      3. Fill the brain slice incubation chamber (i.e., the brain slice keeper) with ACSF and bubble it with carbogen. Then place the incubation chamber in a water bath set at 35 °C.
         Note: Use rodent ACSF for rodent brain slices and human ACSF for human brain slices.
   2. Obtain a juvenile mouse of either sex, between the age of 13 d (p13) to 21 d (p21, where p0 is the date of birth).
   3. Anesthetize the mouse with an intraperitoneal injection of sodium pentobarbital (55 mg/kg of body weight). Once the mouse is deeply anesthetized, as indicated by the absence of the toe pinch reflex, promptly use a vet-approved guillotine instrument to decapitate the mouse in one swift movement.
      Note: Prepare the sodium pentobarbital injection according to the procedures recommended by institutional guidelines.
   4. Hold the mouse’s nose to stabilize the decapitated head and gently make a midline incision starting between the mouse’s eyes and along the entire length of the scalp to expose the skull. Ensure the skull is not compressed by the pressure applied by the razor. Use the corner of the razor’s edge to increase cut precision.
   5. Spread apart the flaps of the mouse’s scalp using fingers to expose the skull. Then, use the fresh corner of a razor’s edge to make an incision along the midline of the skull; pay careful attention to avoid any contact with the cerebral cortex.
   6. Insert splinter forceps into the mouse’s eye sockets to stabilize the decapitated head and place it in a Petri dish filled with cold (0 – 4 °C) dissection solution. Ensure that the head is fully submerged in the dissection solution.
   7. Use a second set of splinter forceps to gently peel away the mouse’s scalp, remove the nose bone by peeling it, and remove the back (caudal side) of the skull.
      Note: The nose bone of juvenile mice is very soft and easily broken.
   8. Use a micro spatula to cut the optic nerves, trigeminal nerves, and spinal cord. Then, use the micro spatula to gently separate the brain from the cranium. Leave the brain fully submerged in the Petri dish filled with dissection solution.
3. Preparation of cortical slices from the somatosensory-motor cortex
   1. Fill the vibratome’s buffer tray with ~150 mL of cold (0 – 4°C) dissection solution.
      Note: Optionally, keep the buffer tray in the freezer until needed to maintain a low temperature during the slicing procedure.
   2. Glue the brain tissue from mouse or human onto the vibratome stage (specimen holder/tray) using instant adhesive glue. For mice, cut off a small caudal portion of the brain (i.e., the cerebellum) so that it can be easily glued flat onto the specimen holder (allowing the rostral side of the brain to face the ceiling).
      Note: Alternatively, horizontal mouse brain slices can be prepared by gluing the dorsal side of the brain onto the specimen holder (the ventral side of the brain should face the ceiling)²⁸.
   3. Gently place the specimen holder (with brain tissue) into the buffer tray. Ensure that the dorsal portion of the brain is facing the vibratome’s blade.
      Note: It is critical to minimize the amount of time the brain is exposed to air.
4. Brain tissue sectioning and collection
   1. Slice the brain into 450 μm thick slices using the vibratome in the dorsal to ventral direction. Ensure that the brain remains completely anchored to the specimen tray while slicing it.
      1. Make the first cut in the mouse brain to remove the olfactory bulb. Then, make subsequent cuts until the somatosensory-motor area is observed (located approximately halfway between the olfactory bulb and bregma).
         Note: Optionally, slice the brain in thinner (200 μm) slices until the corpus callosum appears in the coronal view of the brain, which indicates that the somatosensory-motor area is close.
   2. Use a wide-bore transfer pipette to collect coronal slices (450 μm) that contain the somatosensory-motor area and submerge them in a Petri dish containing cold (0 – 4 °C) dissection solution.
   3. Use a new razor blade and cut off any excess tissue from the slices. For coronal slices from mice, perform a transverse cut just below the neocortical commissure (i.e., corpus callosum). Do not cut in a sawing motion; simply apply pressure on the blade into the tissue and use a detailing brush to gently separate the tissue. Make sure to minimize the movement of the coronal slice.
   4. Use a wide-bore transfer pipette to transfer the dorsal portion of the coronal slices that contain the neocortex (layer 1 – 6) to a second Petri dish filled with warm (35 °C) ACSF for a moment (~1 s). Then, promptly transfer the slices to an incubation chamber containing warm (35 °C) carbogenated ACSF.
      Note: The purpose of the transfer into the Petri dish with ACSF is to minimize the transfer of dissection solution to the incubation chamber while transferring brain slices with the wide-bore pipette. Utilize the incubation chamber’s multiple wells to keep different brain slices organized.
   5. Discard the rest of the brain and animal carcass according to the institutional guidelines.
5. Incubation and maintenance
   1. Leave the brain slices slightly submerged in the incubation chamber at 35 °C for 30 min. Then, remove the incubation chamber from the water bath and allow it to return to room temperature (20 – 25 °C). Wait 1 h for the brain slices to recover before performing electrophysiological recordings.
      Note: Ensure there are no air bubbles that collect underneath the brain slices in the incubation chamber. Air bubbles are air interfaces that cause tissue damage. The mouse brain slices can be maintained for 6 – 8 h, while human brain tissue can be stored for up to 24 h in a well-carbogenated incubation chamber with the appropriate ACSF.
6. Electrophysiological recordings of the superficial cortical layer
   Note: Ictal events are observed in the extracellular local field potential (LFP) recordings from brain slices. The LFP of the brain slice can be observed and recorded with either an interface type chamber²⁹ or a submerged multi-electrode array (MEA) system¹⁶. The procedure has been previously described¹⁶ and additional details are described below.
   1. Use a wide-bore transfer pipette or a detailing brush to move a brain slice onto slightly larger pre-cut lens paper that is held in place using a dental tweezer. Transfer the lens paper (that the brain slice is resting upon) to the recording chamber and secure it in position with a harp screen.
   2. Run warm (35 °C), carbogenated ACSF perfusate through the recording chamber over the brain slice at a rate of 3 mL/min (~1 drip/s). Use a digital thermometer to ensure the recording chamber is 33 – 36°C.
   3. Pull glass electrodes with an impedance of 1 – 3 M‎Ω from borosilicate glass tubing (with an outer diameter of 1.5 mm) using a puller. Backfill the glass electrodes with ACSF (~10 µL) using a Hamilton syringe. Immediately discard the electrode if the tip is damaged.
      Note: Bleach the silver wire (5 min) and only allow a minimal portion (i.e., the tip) of the silver wire to be submerged in the ACSF back-filled glass electrodes to minimize noise and drift during the recordings. Remove any excess ACSF from the glass electrode with a Hamilton syringe if necessary.
   4. Use a 20X stereo microscope to accurately guide the recording glass electrode into the superficial cortical layer (2/3) using manual manipulators. Record/view the electrical activity of the brain slice on a computer with standard software.
      Note: Viable (high-quality) brain slices will exhibit a robust evoked potential in response to the applied electrical stimuli (100 µs, 30 – 300 μA) or light pulses (30 ms, 10 mW/mm²) for optogenetic tissue.
7. Induction of seizure-like activities
   1. Perfuse ACSF containing 100 µM 4-aminopyrimidine (4-AP) over the brain slice. Dissolve 80 mg of 4-AP in 8.5 mL of water to make a stock solution of 100 mM 4-AP. Add 100 µL of 100 mM 4-AP stock solution to 100 mL of ACSF to achieve an ACSF perfusate with 100 µM 4-AP.
      Note: Alternatively, use Zero-Mg²⁺ ACSF (Table 4), a modified ACSF solution containing no added Mg²⁺, to perfuse the brain slice. For human tissue, use a combination of 100 µM 4-AP in Zero-Mg²⁺ Human ACSF (Table 5) to achieve optimal results. The average time for ictal events to appear is 15 min; however, it may take up to 40 min for some brain slices.
8. On-demand seizure generation: an optogenetic strategy for optogenetic mice
   1. Apply a brief (30 ms) pulse of blue (470 nm) light (with a minimum 1 mW/mm² output intensity) to initiate an ictal event. Use a manual manipulator to position a 1,000 µm core diameter optical fiber (0.39 NA) directly above the recording region.
      Note: Set the rate of the photo-stimulation to match the desired rate of the ictal event occurrence (i.e., 1 pulse every 50 s). However, the rate cannot be overly exaggerated from the intrinsic rate at which ictal events occur.
9. On-demand seizure generation: a non-optogenetic strategy for human tissue
   1. Position the brain slice so that the superficial layers are upstream, and the deep layers are downstream to the flow of the perfusate through the recording chamber.
   2. Pull a glass electrode (the same type as used for the LFP recordings) and gently touch its tip with a delicate task wiper to create a small opening. Backfill the electrode with ~25 µL of 100 mM GABA (dissolved in water) and attach it to the picospritzer. Alternatively, backfill the electrode with 200 µM glutamate (dissolved in water).
      Note: To estimate the volume being puffed from the picospritzer (~50 µL), apply a test puff onto a plastic plate (preferably gridded). Then, apply a droplet of known volumes (i.e., 10 µL, 20 µL, 50 µL, or 100 µL) with a pipette for a comparison with the test puff.
   3. Apply puffs of the neurotransmitter (GABA or glutamate) to the human tissue with a picospritzer (10 – 20 psi for 30 – 100 ms) to initiate an ictal event. Apply a single puff of 100 mM GABA onto the deep layer (5) of the brain tissue to generate ictal events in the superficial layer (2/3). Alternatively, apply a single puff of 200 µM glutamate directly onto the superficial layer to generate ictal events in the superficial layer.
      Note: Set the rate of the picospritzer puff to match the desired rate of the ictal event occurrence (i.e., 1 pulse every 50 s). However, the rate cannot be overly exaggerated from the intrinsic rate at which ictal events occur.

2. Protocol II: Acute In vivo Seizure Model

1. Surgical preparation and surgery
   1. Obtain an adult mouse (p35 – p60) of either sex.
   2. Deeply anesthetize mouse with Ketamine (95 mg/kg) and Xylazine (5 mg/kg). Perform the toe-pinch reflex test to ensure the mouse is sedated. Alternatively, use isoflurane (4% for induction, 1.5 – 2% for surgery) to anesthetize the mouse.
      Note: The choice of anesthetics can impact the seizure activity^30,31.
   3. Mount the mouse into a stereotaxic frame by securing its head with ear bars. Place a heated pad underneath the mouse for the duration of the surgery.
   4. Shave the top of the mouse’s head using a rodent trimmer to expose the scalp. Inject 0.3 mL of lidocaine with a 25 G 5/8 in needle into the periosteum to avoid excessive bleeding or pain. Apply eye ointment to the mouse’s eyes to prevent them from drying out during the experiment.
   5. Pinch and lift the center of the mouse’s scalp to assess if its reflexes are gone. Subsequently, perform a horizontal cut to expose the bregma to lambda region of the skull. Gently scrape the entire exposed area of the skull with a cotton swab to create a dry surface.
   6. Perform a craniotomy of 4 mm in diameter at coordinates 2.0 mm lateromedial and -2.0 mm rostrocaudal to expose the somatosensory cortex. Mark the location with a circle (4 mm in diameter) using a permanent marker. Drill around the circle with a pneumatic dental drill until the remaining layer of the craniotomy bone easily gives way when pushed down. Apply saline solution onto the layer of bone before removing the craniotomy with splinter forceps. Subsequently, apply warm saline solution to the exposed region.
2. Recording methods and chemical induction of seizures
   1. Pull glass electrodes with an impedance of 1 – 3 M‎Ω from borosilicate glass tubing (with an outer diameter of 1.5 mm) using a puller. Backfill the glass electrodes with ~10 µL of ACSF or saline using a Hamilton syringe. Immediately discard the electrode if the tip is damaged.
      Note: Bleach the silver wire (5 min) and only allow a minimal portion (i.e., the tip) of the silver wire to be submerged in the ACSF back-filled glass electrodes to minimize noise and drift during the recordings. Remove any excess ACSF from the glass electrode with a Hamilton syringe if necessary.
   2. Use a manual manipulator to guide the glass electrode into the superficial cortical layer (2/3) in the somatosensory-motor area. Record/view the electrical activity of the brain slice on a computer with standard software.
      Note: Layer 2/3 is approximately 0.3 mm deep from the surface³².
   3. Topically apply ~0.5 mL 1.5 mM 4-AP saline onto the exposed cortex of the mouse with a syringe until it completely covers the craniotomy. Dissolve 14 mg of 4-AP in 100 mL of isotonic saline to make a stock solution of 1.5 mM 4-AP saline.
      Note: Prepare the 4-AP saline before the experiment begins. On average, ictal events appear 15 – 30 min after topical application.
3. On-demand generation of seizures in optogenetic mice
   1. Apply a brief (30 ms) pulse of blue (470 nm) light (with a minimum 10 mW/mm² output intensity) to initiate an ictal event. Use a manual manipulator to position a 1,000 µm core diameter optical fiber (0.39 NA) directly above the recording region.
      Note: Set the rate of the photo-stimulation to match the desired rate of the ictal event occurrence (i.e., 1 pulse every 300 s). However, the rate cannot be overly exaggerated from the intrinsic rate at which ictal events occur.