Assessing Changes in Synaptic Plasticity Using an Awake Closed-Head Injury Model of Mild Traumatic Brain Injury

This protocol provides a clear-cut method to produce high-quality transverse hippocampal slices From r-mTBI-administered animals with the ACHI model. The ACHI model produces mild injuries. It does not involve skull fractures or bleeds, craniotomies, or the use of anesthetic.

Additionally, it allows for stable electrophysiological recordings. These techniques allow for the investigation of the changes in synaptic plasticity following repeated MTBI, for which the etiology is vastly unknown, with the potential to inform therapeutic avenues. Demonstrating the procedure will be Allyson Gross, a master's student for my laboratory, and Dr.Eric Eyolfson, a postdoctoral fellow.

Begin with placing the rat in a restraint cone, ensuring that the snout and nostrils are close to the cones small opening to allow adequate ventilation. Prevent the rat movement by holding the cone closed at the caudal end with a plastic hair clip. Manually position the helmet over the midline of the restrained rat with the targeting disc over the left parietal lobe.

Next, place the rat on the foam pad before manually setting the impactor to the extend position. Lower the impactor tip manually to contact the target disc on the helmet. Then set the impactor to the retract position to make the impactor withdraw 10 millimeters above the helmet.

Using the dial on the stereotaxic arm, lower the impact tip by 10 millimeters to touch the targeting disc on the helmet again. Then, flip the impact switch so that the animal's head is rapidly accelerated at six meters per second for 10 millimeters. Once the device is activated, remove the animal from the restraint cone immediately to perform an immediate neurological assessment protocol, or NAP.

After euthanizing the mouse, dissect the brain and place it on a wetted filter paper on an upside down Petri dish. Using a sharp scalpel, remove the cerebellum and prefrontal cortex to blot the brain. Separate the two hemispheres by cutting down the brain midline.

To create transverse hippocampal slices, place one hemisphere on the medial surface, tilt the scalpel at approximately 30 degrees inward, and remove a thin slice from the dorsal surface of the brain to provide a flat surface for mounting on the piston. Flip the brain onto the dorsal surface and gently dab the brain tissue on dry filter paper to remove excess ACSF. Then, attach the dorsal surface of the brain to the piston using cyanoacrylate glue, leaving the ventral surface upright.

Extend the outer tube of the piston over the brain and pour the liquid agarose into the tube until the brain is completely covered. Quickly solidify the agarose by clamping a chilling block over the piston tube. Position the piston into the chamber of the slicer and secure the chamber with a screw.

After securing the blade, add ice-cold oxygenated ACSF to the slicer chamber. On the slicer, set the cutting speed to four, oscillation to six, and toggle the continuous single slicing switch to Continuous. Then, push Start to begin sectioning the brain at 400 micrometers.

As the slicer sections the brain, use a large-diameter Pasteur pipette to transfer each slice to the recovery bath of oxygenated ACSF. Let the slices recover at 32 degrees Celsius for 30 minutes and then leave them to recover for an additional 30 minutes at room temperature. Repeat these steps to create slices from the second hemisphere.

Using a commercially available micropipette puller, pull one-to two-megaohm recording electrodes from 10-centimeter borosilicate glass capillaries with an outer diameter of 1.5 millimeters and an inner diameter of 1.1 millimeters. Use a Pasteur pipette and transfer a hippocampal slice from the recovery bath to the chamber being perfused with carbogenated ACSF and maintained at 30 degrees Celsius. Orientate the brain slice so that the dentate gyrus, and granule cell layer are visible in the field-of-view.

Use an upright microscope to visualize the dentate gyrus with oblique optics. Position a concentric bipolar stimulating electrode to activate the medial perforent path, or MPP fibers, in the middle third of the molecular layer. Then, position a glass micropipette filled with ACSF in the MPP.

Begin with the electrodes further apart, as touching the tissue will damage the fibers. Once the stimulating and recording electrodes are positioned, visualize the evoked field responses using an amplifier, a digitizer, and recording software. Find a suitable field-excitatory postsynaptic potential, or field EPSP, by stimulating the tissue with current pulses and ensuring a minimum amplitude of 0.7 millivolts with a clear fiber volley smaller than the field EPSP.

Increase and set the simulating intensity until the field EPSP is at 70%of the maximum amplitude. Next, establish a stable preconditioning baseline for 20 minutes with 0.12-millisecond pulses delivered at 0.067 hertz. For a stable slice, the initial slope of the fEPSP should be less than 10%variability and the slope of the line of best fit through the plotted field EPSP slopes should be less than 0.5.

Determine the changes in basic synaptic properties using paired-pulse stimuli and constructing stimulus/response input/output curves. For the paired-pulse test, apply a series of paired pulses with an interpulse interval of 50 milliseconds at 0.033 hertz. For the input/output curves, apply a series of increasing stimulus intensities from 0.0 to 0.24 milliseconds at 0.033 hertz to plot the field EPSP response size change.

To study long-term depression primarily dependent on the activation of the CB1 receptor, employ 6, 000 pulses at 10 hertz. For post conditioning recordings, resume for an additional 60 minutes using single-pulse stimulations of 0.12 milliseconds at a frequency of 0.067 hertz. Following the post conditioning recording, administer the paired-pulse stimuli followed by an input/output curve.

Compare these to baseline recordings to observed alterations in presynaptic release properties and assess the health of the slice for long-term recordings. During analysis, adhere to the exclusion criteria for determining the data retention from individual slices in the presynaptic plasticity dataset. Exclude slices displaying a large slope in a line of best fit of field EPSP slopes during the pre-conditioning baseline, instability in the pre-conditioning baseline, and instability in the post-condition period.

At baseline, the neurological assessment protocol, or NAP scores, showed no difference between sham and repeated mTBI rats. Following all awake closed-head injury sessions, the repeated mTBI rats showed significant impairments within the NAP tasks compared to shams, indicating that subtle yet significant behavioral deficits were produced. A series of paired pulses were administered and the ratio of the size of the second field EPSP was calculated relative to the first field EPSP.

The paired-pulse ratios did not differ between sham and repeated mTBI rats, revealing that repeated mTBI rats did not alter basic synaptic physiology in the MPP input to the dentate gyrus. When a 10 hertz protocol was administered to induce a long-term depression, there was a temporary but significant decrease in the capacity of the MPP input to the dentate gyrus to sustain long-term depression on post-injury day one. However, by post-injury day seven, slices from sham and repeated mTBI animals displayed equivalent long-term depression, although there was an indication of a slight trend for repeated mTBI animals to exhibit an increase in long-term depression.

Planning the dissection and cutting process in advance is critical for successfully creating viable hippocampal slices and obtaining stable electrophysiological recordings. This procedure creates a stable experimental platform that can be used to examine TBI pathophysiology and develop new therapeutic avenues. The movement from removing the brain to positioning it to cut in the compressed film for transverse hippocampal slices is important, as is the accurate placement of electrodes in the dentate gyrus.