This method can help answer key questions in the neuroscience field, such as the functions of different types of neurons. The main advantage of this technique is that it makes it possible to reliably identify the different types of neurons in vivo with inexpensive and accessible materials. I will be demonstrating the procedure together with Shinji Muramoto, our technician, and Lanlan Ma, our assistant professor from our laboratory.
To begin this procedure, gently pull out the steel tube for pressure control from the barrel of the holder. Broaden the hole of the pin seat side in the barrel of the holder by drilling. Then, place a stainless steel pipe in the hole.
Next, insert a ceramic split mating sleeve for 2.5 millimeter diameter ferrule into the hole. Make a hole in an L-shaped rabbit and attach the barrel of the electrode holder to the rabbit with epoxy adhesive. Subsequently, attach it to a manipulator with screws.
Prepare a custom-built head post by making the columnar circuit board spacer into a cuboidal shape with a milling cutter, and cut it into two head posts with a saw. Then, flatten the bottom of the head post with a milling cutter. Before the surgery, sterilize all the surgical instruments and a head post with an autoclave.
Clean the stereotaxic frame and the heating pad with 70 percent alcohol. Next, anesthetize the mouse and place it in the stereotaxic frame on a heating pad. In this experiment, a transgenic mouse in which the inhibitory neurons expressing channelrhodopsin-II is used.
Place the teeth in the hole of the bite bar and lightly tighten the nose clamp. Subsequently, fix both sides of the head using ear bars. When the head is correctly fixed by the ear bars, tighten the nose clamp.
Apply an ophthalmic ointment to the eyes. Trim the hair over the head with small scissors and clean the scalp with a cotton swab of chlorhexidine gluconate. Then, cut the scalp along the midline and push it aside.
Remove the periosteum with a cotton swab dipped in 70 percent alcohol and let the skull dry. Then, attach the head post to the manipulator. Position the head post flat on the skull around the bregma using the manipulator.
Mix four drops of the monomer, one drop of catalyst, and one small small spoon of polymer on a small dish that is chilled in the refrigerator. Apply dental cement on the frontal and parietal bone and fix the head post to the skull. After the dental cement has hardened, remove the mouse from the stereotaxic frame.
Apply an antibiotic ointment on the exposed skin and inject a cocktail of medetomidine antagonist and antibiotic. Then, place the mouse on the heat pad in a cage until it awakes and return it to a housing cage with enrichments. To perform craniotomy over the target brain region for recording, clean the skull with a cotton swab soaked with chlorhexidine gluconate and 70 percent alcohol.
Carefully scrape the skull over the recording site with a dental drill. When the bone is thin enough, cut it with the tip of the scalpel and remove it. Then apply dental cement around the exposed region in order to strengthen the skull.
Cover the exposed brain area with silicone adhesive after the dental cement has hardened. Then, place the animal on the heat pad in a cage until it awakes, and return it to a housing cage. Allow the animal to recover for at least one day after craniotomy, before performing the electrophysiological recording.
Before performing the electrophysiological recording, make some glass pipettes from borosilicate glass capillaries using an electrode puller. Fill the pipettes with 10 millimolar PBS. In some recordings, two percent neurobiotin is added to the 10 millimolar PBS.
Next, place the animal in the recording chamber. Attach a filled glass pipette to the custom-made optrode holder. Insert a silver chloride coated wire into the glass pipette.
Afterward, connect the pin to the head stage of the recording amplifier. Subsequently, connect the fiber-optic patch cord with a zirconia ferrule to the split mating sleeve on the optrode holder. Push the patch cord into the sleeve until the tip of the ferrule contacts the back of the glass pipette.
The patch cord delivers the light from LED, which is controlled by a single-channel LED driver. Afterward, remove the silicone adhesive over the craniotomy. Apply warm saline on the brain surface periodically to prevent the brain surface from drying during the recording session.
If necessary, remove the dura with sharp tweezers. Here, the spike responses were recorded in the inferior colliculus of the awake VGAT channelrhodopsin-II mice. In the inferior colliculus of the VGAT channelrhodopsin-II mice, the GABAergic neurons specifically express channelrhodopsin-II.
In the awake mice, both the GABAergic and glutamatergic neurons were excited or suppressed respectively when they were given light stimuli. When the light stimulus is delivered through the glass electrode, it evoked spike responses in some neurons. In contrast, in other neurons, the light stimuli suppressed the spike responses evoked by sound stimuli.
These results show that a glass optrode can record the well-isolated single unit activity and reliably stimulate the recorded neurons by light. After the development, this technique paved the way for researchers in the field of neuroscience to explore how different types of neurons work in the neural socket of various brain regions.
