Calcium Imaging in Mouse Superior Colliculus

My research focuses on neural coding on the mechanisms for visual processing. We are trying to understand how visual information is encoded in the brain, especially in the superior colliculus under the underlying neuromechanisms. To understand visual processing, scientists combine various approaches, including neuron morphology, anterograde and retrograde tracing, IL sequencing, neural modeling, machine learning, and so on.

The challenger is to remove the screw over the SSA cutting and the lifting the bone in one large piece is likely to fail because the bone is attached to the dura. By combining two photo on a widefield calcium image, our recent work revealed the function architecture of motion direction in the mouse SC at both single cell resolution and the global scale. We found that neurons with similar preference from patches up to 500 micrometer on the global biopsy to the upward on the nasal motion.

With our protocol, we are adjusting two research caps. First, researchers can perform long-term calcium imaging in mouse SC at a single cell resolution without breaking the cortex. Second, researchers can record the neuroactivity across the entire SC using widefield microscopy.

Our protocol provides a laser to image the posterior medial SC at single cell resolution with an intact cortex in wild type mice. Also, we use a biocompatible plug to expose the SC, which reduces effection for chronic imaging. Our results provide a way to study the neural coding of visual information across a large visual field.

Combined with optogenetics, one can study half inputs from different brain regions modulating the neuroactivity in SC.Begin the process of creating a suction cup by depositing a drop of PBS in an acrylic dish. Use a 21-gauge flat needle to fill the tip of the needle with PBS by capillarity. Cover the tip of the needle with a translucent silicone adhesive and set it for approximately 10 minutes.

Cut the silicone adhesive with scissors to a disc with a two millimeter diameter. To prepare the plugs, take a 0.75 millimeter thick plastic shim stock and cut out a one by one centimeter square in the center. Clean the shim stock and two acrylic blocks with alcohol and wipe them with lens paper.

Place the shim stock on one acrylic block and deposit silicone adhesive into the center to fill 90%of the opening. Add another acrylic block. Apply approximately one kilogram of force and wait 20 minutes.

Under a surgical microscope, cut the silicone adhesive with a scalpel into one by 1.5 millimeter triangular prisms. Remove any dust from the triangular prisms with sticky plastic tape and place them on a glass cover slip. Place the cover slip with the triangular prisms under a corona treater and turn it on.

Bring the corona treater closer to the cover slip until lightning appears and hold it in that position for a few minutes until the prisms and cover slip are attached. After placing the plugs in a Petri dish, incubate at 70 degrees Celsius overnight. Begin by securing the anesthetized mouse onto a stereotaxic frame with ear bars.

Apply ophthalmic ointment on the mouse's eyes to prevent drying and maintain the body temperature of the mouse at 37 degrees Celsius with a thermostatic heating pad. After shaving the fur and sterilizing the surgical site, inject lidocaine, tilidine, and meloxicam. Once the skin over the SC is removed, clean the skull with a cotton swab.

Adjust the stereotaxic frame until the skull surface is flat. Using a drill, create a hole 0.5 millimeter lateral and 0.42 millimeter anterior to the lambda until the dura is exposed. Inject an adeno-associated virus, or AAV-expressing GCaMP6m, at depths of one millimeter and 1.6 millimeters from the lambda using a beveled glass micropipette.

After the injection, slowly withdraw the injector and proceed with the implantation of the head plate. After injecting the viral construct into the mouse brain, clean the muscles and tissues over the skull and scratch the skull with a blade. Prepare the dental adhesive resin cement by mixing a three quarter spoonful of polymer, three drops of monomer, and one drop of catalyst in a ceramic bowl on ice.

Apply the mixture to the head plate and the skull. Attach them and wait for the adhesive to set for five minutes. Finally, inject antibiotics to prevent infections.

After removing the mouse from the stereotaxic frame, place it on a heating pad for recovery and return it to the home cage after recovering from anesthesia. Three weeks after the viral infection, anesthetize and fix the mouse with a head plate on a stereotaxic frame. Soak a gel foam in saline to stop potential bleeding.

Use a micro drill to create a three by two millimeter oval centered at 0.5 millimeter posterior to the lambda. Thin the boundary of the oval until it cracks, then thin the skull before the oval recording window to close the cover slip to the SC.Using fine forceps, take off the bone flaps. Make a cut down the dura posterior to the transverse sinus and remove the piece of dura.

Dry the skull and the recording window with gel foam and cotton swabs, then deposit a drop of silicone adhesive into the recording window. Use the suction cup to hold the plug with negative pressure and use a motorized micromanipulator to move the suction cup and lower the plug into the silicone adhesive until the cover slip touches the skull. Once the head plate is cleaned, apply butyl cyanoacrylate and resin cement around the boundary of the plug to fix it to the head plate and proceed with imaging.

After implanting the head plate and plug, secure the animal on a rotating treadmill with its head plate. Place the treadmill beneath the two-photon microscope and adjust the height to an appropriate position. Place a black aluminum cone between the objective and the head plate to prevent light contamination from the monitor for visual stimulation.

To perform two-photon imaging, adjust the laser power at the sample plane between 20 and 80 milliwatts. Scan a 600 by 600 micrometer field of view at 4.8 hertz, with a spatial resolution of 2.4 micrometers per pixel and image neural activity up to 350 micrometers in depth. Using a rotary encoder, record the mouse's locomotion on the treadmill.

Use a camera with a 50-millimeter lens to record its pupil size and position. Lastly, synchronize these recordings and image acquisitions to visual stimulation by recording triggers sent by the stimulation computer. The calcium responses of SC neurons from a wild type mouse and the standard deviation projection of the fluorescence across images are presented.

To begin, inject 100 nanoliters of AAV-expressing GCaMP6M at a depth of 200 micrometers at the center of each side of the SC to the mutant mouse whose posterior cortex has failed to develop. After approximately three weeks, implant a five millimeter diameter cover slip over the SC.After fixing the cover slip, acquire images using a CMOS camera after passing an emission filter at 10 hertz. The calcium responses of SC neurons from a partial cortex mutant mouse were imaged using widefield microscopy.

The acquired image was downsampled to one quarter of the original size, and visually evoked calcium responses are shown.