Animal experiments were carried out in accordance with the European Communities Council Directive of 22 September 2010 (2010/63/EU) and were reviewed and permitted by Swiss authorities.

1. Setting up Organotypic Entorhino-hippocampal Slice Cultures

1. Prepare entorhino-hippocampal organotypic slice cultures (EHOSCs) from postnatal day 4 (P4) mouse pups using the static incubation method^4,11. Here, use C57Bl6 and CB6F1 mice.
   1. Take approximately 30 min per mouse pup for the preparation of slice cultures, i.e., 3 hr for a litter of 6 mouse pups. Carry out all steps under sterile conditions in a laminar flow workbench with sterilized surgical instruments.
   2. In order to avoid a possible interference from the position of the slice along the septo-temporal axis, use only hippocampal slices of the septal half. If not stated otherwise, carry out all steps at RT.
2. Cut off the head of a P4 mouse pup with surgical scissors. No anesthesia is required for this step.
   1. Remove the skull around the brain and identify the fissure between the caudal end of the cerebral hemispheres and the midbrain. Carefully remove the part of the brain rostral to the midbrain from the skull and place it in ice-cold preparation medium consisting of MEM supplemented with a stabilized form of L-glutamine.
3. Continue the dissection under a stereomicroscope with the brain being immersed in the ice cold preparation medium. Remove remaining meninges from the cortical hemispheres. Identify the hippocampus at the medial caudal surface of the hemisphere and separate it together with the adjacent entorhinal cortex from the remainder of the brain.
   1. Transfer the tissue to a tissue chopper and cut 400 µm thick slices under aseptic conditions transversal to the septo-temporal axis of the hippocampus.
4. Place the slices on cell culture inserts (approximately 6 slices per insert) and incubate them in 6-well plates with 1 ml per well of incubation medium (47 ml MEM, 25 ml Eagle Medium, 25 ml horse serum, 1 ml glutamax I, 1 ml of a sterile 10% glucose solution, pH 7.3) in a humidified atmosphere with 5% CO₂ at 37 °C. Change the medium the next day and then every other day up to 1 week.

2. In Vitro Hypoxia and Oxygen-glucose Deprivation with Reperfusion

1. See Table 1 for an exact time schedule of the procedure. Culture EHOSCs for 5 days (DIV5) before exposure to hypoxia or OGD. At this time point switch the medium from incubation medium containing 25% horse serum to serum free medium with the following composition: Neurobasal medium with 2% B27 supplement, 1% glutamax and additional 0.1% glucose.
2. Prepare the OGD medium as detailed in Reference 7 .The final concentrations of the components will be the following: 0.3 mM CaCl₂, 70 mM NaCl, 5.25 mM NaHCO₃, 70 mM KCl, 1.25 mM NaH₂PO₄, 2 mM MgSO₄, and 10 mM sucrose at pH 6.8. Add 1 ml of OGD medium to each well of two 6-well tissue culture plates.
   1. Perfuse the OGD medium with N₂ for 1 hr in a hypoxia chamber and then seal and keep O/N in an incubator at 37 °C.
   2. Use anaerobic strips as hypoxic indicators. Consider the medium to be oxygen free when the color of the strips changed from pink to white.
   3. For the induction of hypoxia, use serum free medium with glucose instead of OGD medium. Before use, perfuse the medium with N₂ in a hypoxia chamber as detailed above for OGD medium.
3. Before the slices are exposed to OGD, perfuse the OGD medium again with N₂ for 30 min. Add 2 µl of a 1 mg/ml propidium iodide (PI) solution per well (for a final concentration of 2 µg/ml) to the slices for 30 min and select only healthy slices for experimentation as indicated by low numbers of PI positive cells.
4. Use the schedule of OGD exposure and reperfusion indicated in Figure 1. Deprive slices of either oxygen only (hypoxia) or oxygen and glucose (OGD) for 15 min.
   1. For induction of hypoxia or OGD, transfer slices into OGD medium and keep for 15 min in the hypoxia chamber. Ensure that all fluids that remain on the sides of the insert are aspirated away when the cultures are switched from regular medium to OGD medium and back.
   2. After hypoxia or OGD, replace the OGD medium with oxygenated serum free medium and cultures are allowed to recover for 3, 24 or 48 hr in serum free medium in the regular incubator at 37 °C with 5% CO₂. Add PI again prior to fixation for the determination of cell death.

3. Pharmacological Treatments of Entorhino-hippocampal Organotypic Slice Cultures

1. Culture EHOSCs for 5 days (DIV5) in incubation medium before exposure to pharmacological treatments. Start pharmacological treatments either during OGD or immediately after OGD when slices are transferred to serum free medium.
2. Treatments for protection from OGD.
   1. Add 100 µM 6-cyano-7-nitroquinoxaline-2,3-dione (CNQX) or 1 µM tetrodotoxin (TTX) to the culture medium during OGD induction (for 15 min) or immediately after OGD induction for 30 min. Then, incubate in normoxic serum free medium for 48 hr before fixation.
3. Treatments for induction of excitotoxic death.
   1. After 5 days in incubation medium switch cultures to serum free medium containing 100 µM (RS)-α-amino-3-hydroxy-5-methyl-4-isoxazolepropionic acid (AMPA) for 30 min.
   2. Remove the culture medium containing AMPA by aspiration (take care to remove any residual medium on inserts), then transfer the slices to fresh plates with normal serum free medium and keep them for 48 hr until fixation.

4. Fixation and Immunostaining of Entorhino-hippocampal Organotypic Slice Cultures

1. Propidium iodide (PI) staining.
   1. Add PI for 30 min to the culture medium of living slices at a concentration of 2 µg/ml. View live cultures with an inverted microscope equipped with fluorescence optics and select slices for further treatments. For PI staining in combination with immunohistochemistry, add PI solution 30 min prior to fixation of the cultures.
2. Fixation of the slices.
   1. Remove the culture medium from the 6-well plate and add 3 ml of freshly prepared 4% paraformaldehyde (PFA) to each well. Then place the 6-well plate in a refrigerator and fix O/N at 4 °C.
   2. Remove the PFA solution the following day and wash the slices 3-4 times with phosphate buffered saline (PBS).
3. Removal of the slices from the cell culture insert and blocking procedure for immunohistochemistry of free-floating sections.
   1. Carefully detach and remove the slices from the culture inserts using an art paint brush of the size 0 or smaller. Transfer each individual slice into a well of a 96 well plate filled with blocking buffer (PBS with 3% normal goat serum (NGS) and 2% Triton-X100). Keep the slices in the blocking buffer for 2 hr at RT under constant agitation by an orbital shaker.
4. Immunostaining of the slice cultures
   1. For labeling blood vessels use polyclonal anti-laminin at a dilution of 1:200 in PBS containing 1% NGS and 0.5% Triton-X100.
   2. For labeling neurons use anti-microtubule associated protein 2 (MAP2) at a dilution of 1:500 in PBS containing 1% NGS and 0.5% Triton-X100.
   3. Incubate the slices with primary antibodies for 48 hr at 4 °C with constant agitation. After the incubation period, wash them in PBS with 0.5% Triton-X100 for 3-4 times.
   4. Dilute secondary antibodies 1:500 in PBS containing 1% NGS and 0.5% Triton-X100. Incubate the slices for 2 hr at RT in the dark with the Alexa fluorophore conjugated secondary antibodies. Use the appropriate goat anti-rabbit or goat anti-mouse secondary antibody conjugated to Alexa 350 or 488.
   5. Remove the secondary antibody solution and wash the slices three times with Tris-buffered saline (TBS); mount them on glass slides and coverslip the slides with Mowiol.
5. Microscopy of the stained slices
6. View the stained slices either on a standard microscope with epifluorescence equipment or with a confocal microscope. Take images with an appropriate camera. For some images, invert the fluorescence contrast in order to yield darkly stained vessels on a bright background (Figures 5 and 6).

5. Determination of Regional Vessel Density in the Cultures

1. Quantify the vessel density based on vessel crossings as described in Reference 8.
2. Use slices stained for laminin for these measurements. Superimpose recorded images of 10X magnification with a 6 x 6 grid, where each grid is equal to 100 x 100 µm per square, total field of 0.25 mm².
3. Overlay three such grids in the CA1, CA3, DG, and EC region (Figure 2). Count the vessels crossing the lines of the grid.
4. For obtaining results with a good statistical relevance, make measurements in at least 3 independent experiments, each including data from 3 mouse pups with 3 slices per mouse pup.
5. Determine the average value of vessel crossings from the grids for each region and perform a statistical analysis by ANOVA with Bonferroni correction as post hoc test (p<0.05 defined as significant) using appropriate statistics software.