The protocol presented here diminishes excessive hypoxic damage during preparation of adult and aging mouse hippocampal slices, thus removing a major obstacle for studying function of mature and aging neural circuits. Introduction of hypothermia in sodium free solutions results in hippocampal slices that are healthy for up to 10 hours after cutting and appropriate for both the long-term field-recordings and patch-clamp studies. This method of preparing hippocampal slices could be particularly relevant for animal models in neurodegenerative diseases that by definition require an aging brain preparation.
The most critical step in this protocol is introducing hypothermia via transcardial profusion with an ice-cold solution. With some practice, researchers will be able to reliably execute this step. Begin by preparing one liter of aCSF solution for the recovery chamber and subsequent recordings.
Then prepare 300 milliliters of NMDG-aCSF for the transcardial profusion and cutting steps. Place the vibrating microtome cutting tray and mounting disc in a minus 20 degrees Celsius freezer. To prepare the recovery chamber, fill it to just above the slice holding mesh and start the bubbler keeping the chamber on the bench at room temperature.
Chill the entire 300 milliliters of NMDG-aCSF in a freezer until ice crystals start to form on the surface and the walls of the bottle. Place the bottle with chilled and NMDG-aCSF on ice and bubble it, keeping the solution between zero and two degrees Celsius. Take the tissue mounting disc out of the freezer and wipe it dry, if needed.
Cut out a block of 5%agar about the size of a mouse brain and glue it in the center of the disc using a thin layer of cyanoacrylate glue. Place the disc with glued agar on ice and cover it with paper towels until ready to use. Take the cutting tray out of the freezer, place it into the microtome, then surround it with ice and load the blade.
Prepare all tools ahead of time for the brain dissection. Set up the peristaltic pump for transcardial perfusion. Insert one side of the pump tubing into the bottle with iced NMDG-aCSF and fit the other side with a 27 gauge needle.
Set the pump speed to approximately 3.5 milliliters per minute. At this speed, the outflow of an NMDG-aCSF is a fast trip, not a continuous flow. Place the mouse on its back on a diaper.
Before continuing, confirmed that the mouse is at a surgical plane of anesthesia by performing a toe pinch. The mouse should be unresponsive, then taped down its front and hind legs so that the chest and abdomen are exposed. Cut out a large patch of the skin on the chest, going from below the sternum to the throat.
Grab the sternum with forceps, lift it gently and start cutting through the rib cage on both sides until the chest cavity is exposed. Cut through the diaphragm, leaving the flap of the rib cage attached via a thin piece of muscle. It should be possible to set it aside without having it fall back onto the exposed chest cavity.
Check that the heart is still beating and ensure that most of the liver is visible. Insert the needle into the left ventricle, which looks lighter in color than the right. To stabilize the needle, drive it through the remaining ribs on the left side of the body.
Locate the dark red-colored right atrium and cut through it with small scissors. The blood should start flowing out. Start the pump and observe the liver, which will change color from red to brown.
Monitor the liver color and continue perfusion until the liver turns pale brown. Run the pump for a few more minutes. The body temperature of the animal should fall to 28 to 29 degrees Celsius and its nose should be cold to the touch.
Decapitate the mouse with large decapitation scissors, then use a scalpel with a number 10 blade to cut open the skin on top of the skull. With small angled scissors, cut the skull at the midline. Next, use the number three forceps to pry away the right and left halves of the skull, being careful to take the dura away with it.
Remove the brain, which should be an off-white color by scooping it out with a spatula and drop it into the NMDG-aCSF solution on ice. Leave it there for up to a minute. Take the brain out of the NMDG-aCSF and place it on a piece of filter paper.
Cut and remove a 60 degree wedge of tissue with a 60 degree tool centered at the midline from the rostral end of the forebrain. Use the cut sides of the mounting surface as it provides the proper angle for hippocampal slices. Separate the hemispheres down the midline with the scalpel and glue them onto the mounting disc.
Take the mounting disc from the ice and wipe it dry, if needed. Then glue each hemisphere in front of the agar block, cut side down. Ensure that the ventral sides of both hemispheres are touching the agar block and that the dorsal sides of both hemispheres are facing the blade.
When glued on the cut side, each hemisphere should be oriented relative to the blade in a way that ensures transverse slices of the dorsal hippocampus in situ. Submerge the disc with the hemispheres into a cutting chamber containing ice-cold carbogenated NMDG-aCSF. Cut a 400 micrometer section, then use a disposable transfer pipette with a cutoff tip to transfer the slice to a recovery chamber containing carbogenated NMDG-aCSF at room temperature.
Continue cutting the rest of the sections and transferring them to the recovery chamber taking no more than 10 minutes to cut a total of eight to 10 slices from the dorsal hippocampus region. Incubate the slices at room temperature for approximately two hours before recording. This protocol was used to generate hippocampal slices from adult mice.
Large numbers of pyramidal cells in the CA1 field and subiculum appear in low contrast, a hallmark of healthy cells, when observed under infrared differential contrast microscopy. Patch-clamp recordings can be obtained from CA1 neurons of mice that are over six months old. In the example experiment here, the frequency of miniature excitatory postsynaptic currents was measured after NMDA-LTD was induced relative to control conditions.
The frequency of miniature excitatory postsynaptic currents was lower in neurons after an NMDA-LTD induction, indicating activity dependent pruning of synopses in CA1. Changes in amplitude were not detected. During mEPSC recordings, CA1 cells were also filled biocytin and intact dendritic arbor and healthy cell habitus of the pyramidal neurons can be seen here.
A robust distribution of the fluorescent dye throughout the cell allows analysis of dendrites and dendritic spines under different conditions. Long-term potentiation, or LTP, of CA3-CA1 synapses of approximately 170%was observed, suggesting that maintenance of signaling cascades needed for LTP is present in slices prepared from adult mice. A robust field excitatory postsynaptic potential signal suggests that network connectivity was also preserved.
Two critical aspects of the protocol are transcardial profusion with an ice-cold solution to induce hypothermia and introduction of an NMDG as a sodium ion substitute in solutions to prevent cytotoxic edema. Using these precautions, acute slices can be prepared from any brain region. Moreover, slices prepared in this manner can be used for calcium and voltage imaging using two-photon or widefield microscopy.
This protocol could serve as a basis for a standardized preparation for acute hippocampal slices from aging animals and thus facilitate comparisons across studies in the context of neurodegenerative disease mechanisms.
