1. Before Starting the Preparation of Hippocampal Slices.

1. Prepare the tissue slicer by placing a piece of Teflon sheet and mounting a new blade.
2. Wipe the tissue culture (TC) hood with 70% ethanol and set the dissecting microscope inside. Sterilize the hood, microscope, tissue slicer and all dissecting instruments for 15 minutes with UV light.
3. Prepare six well TC plates. Add 750 μL slice culture media (SCM) per well and place cell culture inserts in each well. Make sure the membranes are thoroughly wet with no bubbles underneath. Place the plates in incubator at 35°C gassed with 5% CO₂ until needed.
4. Pour 50 mL low Na⁺ ACSF (dissecting solution) into a 100 mL beaker and place it on ice-salt mix. Bubble the low Na⁺ ACSF with 5% CO₂ / 95% O₂ until color changes and ACSF forms a slurry mix of frozen and liquid solution (10-20 min).
5. Get a P5-P7 rat pup. Up to three pups can be used.

2. Hippocampal Slices Preparation.

1. Cut the head of the animal with sharp utility scissors. Cut the skin and expose the skull. Open the skull by cutting from side to side along the interaural line and then along the sagittal suture with small scissors. An optional cut from side to side in the front can be made to facilitate removing the bones and exposure of the brain. Scoop out the brain quickly with a rounded spoon micro spatula and place it in the slurry of dissecting solution to chill for ~ 1 minute. Pour ~10 mL of ice cold dissecting solution onto a 60 mm dish and transfer the brain from the beaker to the dish. The brain should be covered with dissecting solution.
2. Under the dissecting microscope: Place the brain and hold it at the midline with the dissecting forceps pressed to the bottom of the 60 mm dish. Use the hippocampus dissecting tool to separate the hemispheres leaving out the midbrain. The hippocampi are then exposed on each hemisphere. Then gently scoop the hippocampus out with the hippocampus dissecting tool. Use the dissecting needle to completely isolate the hippocampus and clean it as much as possible.
3. Using a snipped tip of a P1000 filter pipette tip, gently aspirate the hippocampus and transfer it to the Teflon sheet on the tissue slicer. Position the hippocampus on its concave side.
4. Align the hippocampi perpendicular to the blade to obtain coronal sections and drain excess of liquid.
5. Slice the hippocampi every 400 μm.
6. Pour ~10 mL cold SCM into a 60 mm dish and transfer sliced hippocampi from the slicer using another snipped P1000 filter tip and cold SCM. Avoid making bubbles.
7. With the help of an iris spatula and a straight spatula gently separate the slices from each other.
8. Separate well defined and undamaged slices from damaged slices.

3. Hippocampal Slices Culture

1. Bring the six-well plates with SCM and cell culture inserts from the incubator. With the help of another snipped P1000 filter tip, transfer individual slices onto the membrane. Place 4-5 slices per membrane. Be careful not to place the slices either close to the insert wall or close to each other. When necessary, use iris spatula to separate slices. Remove excess medium. Touch slices as little as possible once they are on the membrane.
2. Move plate back to incubator and culture at 35°C and 5% CO₂.
3. Change SCM every 48 hours inside the TC hood by aspirating the SCM with a Pasteur pipette. Add 750 μL of fresh pre-warmed SCM per well. Make sure no bubbles are formed under the membrane.

4. Solutions

1. Low Na⁺ ACSF – Dissecting Solution for slice cultures
   To deionized and sterile H₂O add:
   

   	For 500 mL	For 1000 mL	Final Concentration
   CaCl2 (1 M)	0.5  mL	1 mL	1 mM
   D-Glucose	0.901 g	1.802 g	10 mM
   KCl	0.149 g	0.298 g	4 mM
   MgCl2 (1 M)	2.5 mL	5 mL	5 mM
   NaHCO3	1.092 g	2.184 g	26 mM
   Sucrose	40 g	80 g	234 mM
   Phenol Red Solution 0.5% in DPBS	0.5 mL	1 mL	0.1% v/v

   
   Mix ~30 min
   Sterilize by passage through 0.22μm filter
   Make 50 mL aliquots and store at 4 °C no longer than 2 months.
2. Slice Culture Medium (SCM)
   

   	For 500 mL	For 1000 mL	Final Concentration
   MEM Eagle medium	4.2 g	8.4 g	8.4 g/l
   Horse serum heat inactivated	100 mL	200 mL	20%
   L-Glutamine (200 mM)	2.5 mL	5 mL	1 mM
   CaCl2 (1 M)	0.5 mL	1 mL	1 mM
   MgSO4 (1 M)	1 mL	2 mL	2 mM
   Insulin (1 mg/ mL), dissolved in HCl 0.01 N	0.5 mL	1 mL	1 mg/l
   Ascorbic Acid, solution (25% w/v)	0.024 mL	0.048 mL	0.00125%
   D-Glucose	1.16g	2.32g	13 mM
   NaHCO3	0.22g	0.44g	5.2 mM
   Hepes	3.58g	7.16g	30 mM

   
   Mix until thoroughly dissolved and bring to room temperature.
   Adjust pH to 7.27-7.28 with 1 N NaOH
   Measure osmolarity. Adjust to 320 mmol/kg with deionized and sterilized H₂O. Expect to add approximately 25-40 mL. Check osmolarity again.
   ***pH may change slightly while adjusting osmolarity, this is ok, it is more important that the osmolarity is in the correct range (317-323).
   Sterilize by passage through 0.22 μm filter.
   Make 20 mL aliquots and store for up to two-three weeks at 4°C.
   Gluatamine stock: prepare at a concentration of 200 mM and store at -20 °C in aliquots of 2.5 mL.
   Ascorbic acid stock: prepare at a concentration of 25% (w/v) and stored at -20 °C in aliquots of 100 μL.

5. Representative Results:

Slices should look white under a dissecting scope without black spots and well defined and undamaged CA1, CA3, and Dentate gyrus regions. Bacterial contamination is easily seen as moving black specks in the medium or turbidity of the SCM. When placed under the microscope, the surface of the slice should look clean after 4 days in culture with clear and discernibly cell bodies. If no clear cell bodies are seen and much debris covers the surface after 4 days, then is not a healthy slice.