Electrocorticographic Recording of Cerebral Cortex Areas Manipulated Using an Adeno-Associated Virus Targeting Cofilin in Mice

This protocol can be used to identify the role of specific individual molecular targets in the regulation of electrocorticographic activity during different Vigilin states. Among the benefits of using adeno-associated virus is the ability to precisely target a given brain region and a specific cell type. Demonstrating the protocol will be Julien Dufort-Gervais, a research associate from my team.

After confirming a lack of response to toe pinch reflex in an anesthetized 12-week-old mouse, use a hair trimmer to shave the hair from the back of the ears to the front of the head between the eyes. Add a generous drop of ophthalmic ointment to each eye to prevent dehydration and use the ear bars to carefully fix the head of the mouse onto a stereotaxic apparatus. Gently pull the tongue out of the mouth to avoid suffocation.

Fix the nose of the mouse to the apparatus and use 70%ethanol to sterilize the exposed skin on the head. Holding the skin with extra-fine Graefe forceps, use tissue scissors to cut the skin from the base of the ears to the level of the eyes and place two surgical clamps on each side of the incision to stretch the skin and to expose the skull. Avoiding brain sutures, use a scissors tip to scratch the skull surface to remove the periosteum and to create overlapping streaks in two or more directions.

Use 70%ethanol to remove the bone fragments and to disinfect the skull. With a previously prepared cannula fixed to the stereotaxic arm, identify the location of the bregma and lambda and note the stereotaxic coordinates of each. Use the stereotaxic arm and a pen to mark the position of the cannula on the skull 1.5 millimeters lateral right to the midline and 1.5 millimeters anterior to the bregma.

Using a 0.7 millimeter drill bit, carefully pierce the skull at the marked position perpendicular to the skull surface and aligned with the vertical axis and wash the skull with a sterile cotton tip soaked in a 10%betadine povidone iodine solution, then retract the plunger of a 10 microliter syringe by one microliter to load the cannula with a one microliter air bubble. Next, mix the AAV mixture of interest with slow pipetting and add 1.7 microliters of the mixture to a sterile Petri dish. Use this syringe to load 1.5 microliters of the solution into the cannula and mark the position of the air bubble on the connected PE50 tube.

Vertically align the cannula with the hole in the skull such that the cannula reaches the upper edge of the bone and mark the Z coordinate of the skull surface. Slowly lower the cannula until the tip reaches 1.5 millimeters below the skull surface and layer five of the motor cortex and start the syringe pump at a 0.025 microliter per minute flow rate to deliver one microliter of AAV over 40 minutes. Use the air bubble in the tube to track the injection, making any adjustments as necessary.

When the entire volume of virus has been delivered, leave the cannula in place for five minutes to ensure a sufficient diffusion and to avoid backflow before slowly and carefully lifting the stereotaxic arm to remove the cannula from the cortex. For electrode implantation, use straight Kelly forceps to slowly screw one electrocorticographic electrode with a straight gold wire into the vertical axis of the hole in which the AAV was injected, leaving at least 2.5 millimeters of screw outside of the skull to minimize the damage to the dura and the cerebral cortex. Use a pen to mark the position of the reference electrode 2.6 millimeters lateral right to the midline and 0.7 millimeters posterior to bregma and the position of the posterior electrocardiographic electrode at 1.5 millimeters lateral right to the midline and 1.5 millimeters anterior to lambda and the positions of three maintenance screws on the left hemisphere with no specific coordinates, but as distant as possible from each other and from the electrocorticographic electrodes.

Use the drill to carefully pierce the skull perpendicular to the skull surface at the marked positions of the other electrodes and screws and wash the pierced skull with a 10%povidone iodine solution. Block the holes with small rolled pieces of delicate task wipe before installing the screws to prevent bleeding and contamination and use straight Kelly forceps to insert the screws into the drill holes at the same angle that the holes were pierced. Place a few small drops of dental cement into the center of the ring-like space inside the screws and use extra-fine Graefe forceps to lift the skin above the neck muscles.

Using Dumont number five forceps, hold the curved extremity of one electromyographic electrode and insert it approximately one to two millimeters into the muscles. Place the curved side and folding point of the electrode into the dental cement and insert the second electromyographic electrode in the same manner. After covering the eyes of the animal, apply light for three to five minutes to solidify the cement and use additional dental cement to cover the base of the electrocorticographic electrodes and the anchor screws to form a crown-shaped contour.

After applying three to five more minutes of light, fill the center of the montage with acrylic cement and remove the surgical clamps. Use a synthetic absorbable monofilament suture to close the skin at the front and back of the montage so that the skull is not exposed and use curved forceps to hold the connector above the montage to carefully align the gold wires of the electrodes with the connector pins, then quickly solder each electrode extremity to a single corresponding connector pin. After removing the mouse from the frame, cover the empty space between the connector and the head with the acrylic cement.

And after weighing, place the mouse in a clean cage with a non-meshed lid on a heat pad. Two weeks after the surgery, connect the mice to recording cables and at least one week later, record the electrocorticographic and electromyographic signals for 24 hours or more. A successful infection is confirmed by HA staining of the neurons within the motor cortex surrounding the injection site, indicating the presence of cofilin S3D HA.Co-staining with the excitatory neuron marker CaMKII alpha indicates clear cofilin S3D HA and CaMKII alpha co-expression under high magnification.

Log transformed relative power spectra for wakefulness, slow wave sleep, and paradoxical sleep show state-specific differences in spectral activity under cofilin inactivation. The combination of electrocorticographic recording and AAV-mediated manipulation of precise molecular targets is also applicable to multiple brain regions and to other neuroscience sub-fields such as research on epilepsy or memory in addition to sleep.