A Method to Make a Craniotomy on the Ventral Skull of Neonate Rodents

The overall goal of this procedure is to perform a craniotomy on the ventral skull of rat pups for in vivo electrophysiology, or two photon imaging studies. This is accomplished by first anesthetizing, the rat pop on a heated pad and using a ventilator to control its breathing. The surgery begins with exposing the ventral side of the skull, followed by a craniotomy to expose the brainstem surface.

This procedure sets up the use of in vivo electrophysiology and two photon microscopy to analyze structural and functional aspects of neural structures. The main advantage of this technique over other methods, such as acute or organotypic brain stem slices, is that it provides a system to study the dynamics of the bare cellular processes in the mammalian brainstem. In vivo, Individuals new to this method, they will struggle because the rhythm pops are small and the methods assume some proficiency with surgical technique.

This surgery requires mammalian ringer solution, the proper tools, a heating pad, and a small animal ventilator. Take a confirmed anesthetized animal position. It dorsally over a heat pad set to 37 degrees Celsius and attach the nose cone to sustain the anesthesia, tape down the limbs and tail and then clean the skin using saline or disinfectant solution.

Now make a longitudinal cut through the neck epidermis. Then make four lateral incisions, bluntly, dissect the skin and move it aside. With forceps, it can then be secured with tape.

Be careful to keep the carotid arteries away from the trachea and to not puncture the arteries. With spring scissors, dissect the longitudinal muscles covering the trachea, and then under the trachea, next position, suture threads around the trachea. Position one thread to secure the ventilation tube and a second to close the trachea rostral, where the ventilation tube will be inserted.

Now make an incision through a tracheal ring between the ties. No fluid should be allowed to enter through the hole. Insert the tube through the hole and tighten and tie the threads to secure the tube first.

Then to close the trachea. Make certain that the tube is able to move without imposing strain on the preparation. Then promptly switch the isof fluorine supply to the ventilator.

The stroke volume and ventilation rate should be set according to the animal's age. Finish this portion of the procedure by sealing the exposed surrounding tissue with an elastomer. First, cut the trachea adjacent to the ventilation tube.

Using two more cuts along the muscle wall of the buccal cavity. Expose the coddle end of the palate After cleaning out fat and muscle tissue from the exposed area, the space between the basi occipital bone and the last vertebra becomes exposed. Locate the medial wall of the bullah using the micro drill.

Thin the skull using a V-shaped motion until the underlying arteries can be seen. Next, gently break the skull using a dissecting chisel and clear the bone chips. Continue drilling and chiseling until the desired size for the craniotomy is achieved.

Clean the area several times with wringers and keep it moist throughout the experiment. Under the dissection scope, prepare the animal for the experiment. Begin by locating the right or left carotid.

Then bluntly, dissect the carotid artery from the surrounding tissues. With a hemostat, pick and hold the dissected artery and tie three pieces of suture around it. Tighten the thread closest to the heart to stop the blood flow.

Then make a small 45 degree cut on the artery between the tide and untied threads. Mop up the blood as needed. Now cannulate the artery with a tube filled with trissy dextran solution.

Insert the tube five millimeters towards the pop's head. Secure the cannula by tightening the other two threads. Trim excess thread, and apply an elastomer to make the cannula more secure.

If the cannula is not sealed, the fluorescent dye will leak into the craniotomy. Now, attach the syringe to the pump. Then set the speed of the pump to the blood flow rate of the animal.

Under a two photon microscope, view the trissy dextran. The microscope should be fully warmed up with the animal in position and the area of interest in focus through the five x air objective, inject the fluorescent D strand into the bloodstream. Next, increase the strength of the objective to 20 or 40 x and focus the scope.

Start imaging the area of interest and adjust the parameters to optimally view the fluorescence. Then collect a time series of images at a fixed focal plane for analysis using patch clamp pipettes. Neural tracer was electroporated at the red dot near the midline, hence labeling decking afferent axons in the medial nucleus of the trapezoid body, or MNTB, which is boxed out.

These cells were accessed using the described craniotomy on P one to P five pups, A two photon microscope equipped with a high numerical aperture water immersion Objective was used to image the fibers reaching the MNTB, including very fine collateral branches labeled by arrows. Neural tracers were also directly delivered to the MNTB, which resulted in labeling of MNTB cells and afferent axons. This was better appreciated from a wider field of view.

Taken at lower magnification individual MNTB cells could be clearly seen. Two photon imaging of vascular permeability was measured using Trixie dextran, a fluorescent solute injected into the cerebral circulation via a cannula inserted in the carotid artery. The pups injected were at the P nine to P 10 stage.

After a brief pulse of labeled solute into the bloodstream, it was possible to obtain a time-lapse sequence of a region of interest. The total fluorescence intensity in the region was measured offline, and a model was used to determine the blood-brain barrier permeability to fluorescently labeled solutes. Once mastered, this technique can be done in an hour if it is performed properly.