A Procedure for Implanting Organized Arrays of Microwires for Single-unit Recordings in Awake, Behaving Animals

The overall goal of this procedure is to accurately implant microwire arrays into target brain regions. This is accomplished by first precisely measuring and marking coordinates on the skull for a craniotomy. The second step of the procedure is to slowly remove layers from the skull using a dental drill to accurately produce the craniotomy.

The third step is to position the microwire array so that it will squarely pass the confines of the skull and be level with the skull. The final step is to slowly lower the array until it reaches its target. Ultimately, results can show that microwire arrays implanted.

Using this procedure can target discrete sub regions of the brain and produce a high neural yield, which can be maintained over longitudinal recordings. Microwire arrays allow neuroscientists to examine the firing patterns of single neurons with high spatial and temporal specificity, whether used alone or in combination with other techniques. Single unit recordings from Microwire arrays provide insight into the brain's information processing.

Recording neural signals during behavior is crucial for understanding brain function. My lab has used single unit electrophysiology to study normal processes such as somatotopic mapping, motor learning, as well as disease processes such as Parkinson's disease and drug addiction. In vivo electrophysiology is a highly specialized skill that takes years to master.

Visual demonstration of this technique reduces the time needed to adequately perform the surgery. After anesthetizing the animal with pentobarbital, administer atropine, methyl nitrate, and penicillin to maintain respiratory function and prevent infection. Verify the anesthetized state using the tail pinch test before proceeding as necessary.

Give alternating injections of ketamine, hydrochloride and sodium pentobarbital to maintain anesthesia throughout the surgery. Next, shave the scalp using a number 22 scalpel blade, followed up with povidone iodine. Use four subcutaneous injections of bupivacaine to locally anesthetize the scalp.

Allow five to 10 minutes for the local anesthetic to take effect. Next, apply an ophthalmic lubricant to the eyes and secure the animal into the ear bars and nose clamp of a stereotaxic apparatus by eye. Bring the animal's head to an approximately level position.

Then use a scalpel to make an incision along the midline of the scalp. The incision must extend from the posterior portion of the nasal bone to just behind the ears. Using a dissection spatula, clear the skull of all remaining tissue until both lateral skull ridges and the posterior skull ridge have been reached.

Pull back the skin around the incision using about six hemostats. Then clean any blood off the skull and allow it to dry so dental acrylic can adhere to it later. Next, using a dissecting microscope, mark the BMA and lambda by interpolating the intersection of the skull sutures.

Attach a small pointed item like a dental drill bit to a stereotaxic arm and lower it to determine the dorsal ventral coordinate of bgma and lambda. Next, adjust the nose clamp until the DV coordinates for bgma and lambda are within 100 microns of each other. Once leveled, record the coordinates for bgma along with the position of the nose clamp.

Double check these coordinates the surgery depends on them. Now use the measured coordinates to calculate the media lateral and anterior posterior coordinates for the four corners of the craniotomy relative torema. Using the calculations, mark the corners on the skull with a pointed stereotaxic attachment that has been coated in black ink.

Now, drill out the skull window by removing the bone in a series of small layers. Start by drilling the marked corners of the window where the marks have been placed. Next, connect the corners and outline the window.

Finally, clear out the area within the outline down to the depth of the dura mater. The hole must be beveled widening below the superficial layers of the skull. Using micro forceps carefully remove any remaining bone chips, debris, or dura mater inside the skull window.

This is extremely important as debris can compromise the integrity of the array during implantation. Once cleared, the window must be kept moist with bacterial static saline. For the remainder of the surgery, begin the implantation by marking the location for placing five skull screws and a ground wire.

Ideally, each screw should be in a different skull bone. However, the locations cannot interfere with the array placement. Next, drill the holes for the skull screws and secure the screws in place.

Lower the screws until three or four threads show deep enough to traverse the thickness of the skull After placement, clean off the screw threads. Now drill a hole for the ground wire and lower the wire slowly over one to two minutes into the targeted DV coordinate. Fix the wire in place using dental acrylic.

Next, add acrylic around the threads of the skull screws before it dries. Remove any excess acrylic from the ground wire or skull screws. Allow 15 minutes for the dental acrylic to dry.

Meanwhile, attach the array to the stereotaxic arm. Level the array and orient it so that it will squarely pass through the confines of the skull window. When the acrylic is dry, completely fill the skull window with biological saline.

Then lower the array until it creates a dimple in the saline. This coordinate is used as the skull level for the array. Use this value to calculate the final DV coordinate of the array.

Lower the array slowly to the final dv. Coordinate stopping and awaiting a few minutes. With each millimeter traveled, the sterile saline will dissolve the polyethylene glycol on the array that temporarily keeps the wires in their confirmation.

When the array is just one millimeter from the target, lower it more slowly to the final destination At one 10th millimeter intervals. When the array is in place, allow the tissue to rest for five minutes. After the rest, dissolve the remaining peg by applying more saline.

Then use dental acrylic to cement the micro wires in place. Add multiple layers of acrylic and allow them 15 to 20 minutes to harden. Next, build up the remainder of the animal's head stage with dental acrylic in case the skull screws ground wire, and connector for the micro wires.

See the text protocol for postoperative procedures. Next, remove the animal from the ear bars and place it in a clean area. Observe the animal frequently until thermal regulation and locomotion have recovered.

Once recovered from anesthesia, move the animal into single housing. Monitor the animal's recovery for seven or more days and provide injections of carprofen and enro. Floxacin as recommended by the attending veterinarian, may measured neural discharge from an implanted microwire array was correlated with behavioral events.

For example, tonic firing of this cell in the nucleus accumbens shown in 32nd bins was measured while the animal self-administered cocaine. Notice that the cell becomes inhibited as the animal's body level of cocaine increases. However the cell returns to its original firing rate.

As the cocaine falls after the self-administration contingencies have ended. While attempting this procedure, it's important to remember to check the precision of the skull window and verify the alignment of the array to ensure target accuracy. Don't forget that micro I arrays are particularly delicate and that the integrity of the array is critical to proper implantation.

Patients and care should be taken when handling the erase Single unit. Recording can be combined with many other techniques such as immunohistochemistry or optogenetics. When these combinations are coupled with an appropriate behavioral paradigm, they can reveal deep insights into the complexities of brain function.