An Indirect Neuron-Astrocyte Co-Culture Technique for Studying Neuron-Glia Interactions

All procedures involving animal models have been reviewed by the local institutional animal care committee and the JoVE veterinary review board.

1. Preparation and Cultivation of Cortical Astrocytes

NOTE: Complete these steps of the protocol at least 7 d before proceeding to the next steps, as the astrocyte cultures should develop into confluent monolayers before the neurons are prepared. Primary astrocytes are derived from mixed glial cultures obtained from mouse pups around postnatal day (P) 0-3. Three brains (6 cortices) per T75 flask are to be used.

1. Prepare the astrocyte medium by supplementing Dulbecco's Modified Eagle Medium (DMEM) with 10% v/v horse serum and 0.1% v/v gentamicin under sterile conditions. Store the medium at 4 °C for up to 2 weeks.
2. Coat 3x T75 flasks with 10 µg/mL poly-D-Lysine (PDL; in cell-culture-grade water) for at least 1 h at 37 °C in the presence of 6% v/v CO₂. After coating, wash the flasks twice with phosphate-buffered Saline (PBS) to remove unbound cations, and add 9 mL astrocyte medium.
3. For the preparation of the brains, add 10 mL Hank's Balanced Salt Solution (HBSS) to 1x 10 cm Petri dish and 1 mL HBSS to 1x 15 mL tube (regardless of the number of brains that will be used). Keep a pair of surgical scissors, 2 fine forceps, and a binocular within reach for the procedure.
4. Decapitate 3 pups quickly with the surgical scissors and collect the heads in an empty 10 cm dish.
   1. Perform the preparation of the cortices under the binoculars. Using two fine forceps, remove the dorsal skin from the skull at the midline, starting from the neck up towards the level of the eyes. Thereafter, the brain with its blood vessels is visible under the skull.
   2. Fix the head at the rostral end with a pair of forceps and incise the midline of the skull, starting at the posterior end. Subsequently, clip off the right and the left halves of the skull to expose the brain.
   3. Close the tip of the forceps and position it ventrally under the brain. Lift the brain out of the skull and transfer it into the HBSS-filled 10 cm Petri dish. Proceed in the same manner with the remaining specimen until all brains are collected in the dish.
5. After collecting the brains, start with the separation of the cortices by pinching off the hindbrain using a pair of forceps like a pair of scissors. Perform a midline incision between the two hemispheres with one pair of forceps. Carefully fix the brain with a forceps and cut off the midbrain and the olfactory bulbs using a second forceps to end up with the cortical halves.
6. Fix one cortical half with one pair of forceps and peel the meninges from the hemispheres by detaching them from the lateral edge and subsequently pulling them from the cortical surface using a second forceps.
7. Flip the cortical half with its surface oriented down towards the dish; carefully dissect and remove the crescent-shaped hippocampus using one pair of forceps. Transfer the single cortices into the conical 15 mL tube using one closed forceps to carry an individual cortex.
8. After collecting all cortices, transit to the sterile laminar flow hood and begin with the preparations for the dissociation of the cortical tissue.
9. Add 1 mL DMEM to a 2 mL reaction tube and add 0.1% w/v papain (30 units). Incubate the suspension in a water bath at 37 °C until a clarified solution develops (about 3 min). Add 0.24 mg/mL L-cysteine and 20 µg/mL DNAse and shake gently.
   NOTE: For digestion, approximately 1 mL of enzyme solution is needed.
10. Filter (0.22 µm pore size) to obtain 1 mL sterile solution before adding it to the cortical tissue. Incubate the tubes for 30 min – 1 h at 37 °C (i.e., in a water bath)_.
    NOTE: Steps 1.11 – 1.14 are critical and should be completed within 15 min.
11. To terminate the digestion reaction, add 1 mL astrocyte medium and carefully triturate the digested tissue using a 1 mL pipette. For trituration, gently move the pipette plunger, thereby aspirating and extruding the cortical tissue suspension.
12. Once the tissue has been dissociated into a single-cell suspension, add 5 mL astrocyte medium and centrifuge for 5 min at 216 x g.
13. Aspirate the supernatant carefully from the resulting pellet and resuspend the cells in 1 mL astrocyte medium.
14. Add the cell suspensions to T75 flasks replenished with 9 mL astrocyte medium (see 1.2).
15. Incubate the cells at 37 °C in the incubator in the presence of 6% v/v CO₂. After 4 d, perform the first complete medium change (10 mL). Thereafter, change the culture medium every 2 – 3 days.
16. After 7 days, check if the cells have formed a confluent monolayer. If not, wait for another 2 – 3 days until complete confluence of the culture is attained.
    NOTE: Astrocytes display large, flattened cell bodies, localize at the bottom of the flask, and do not develop notable cellular processes. They differ from the small phase-contrast bright progenitor cells and microglia residing on top of the monolayer.
17. In order to get rid of the progenitor cells and obtain a pure astrocyte culture, place the T75 flasks on an orbital shaker and shake the cultures O/N at 250 rpm. Ensure that the temperature is adjusted to 37 °C and that the filter of the flask is sealed with laboratory film to prevent evaporation of the CO_2.
18. The next day, aspirate the medium and add 10 mL fresh culture medium replenished with 20 µM cytosine-1-β-D-arabinofuranoside (AraC) to eliminate residual dividing cells.     
    NOTE: incubation with AraC for 2 – 3 d in the incubator at 37 °C, 6% CO₂ will result in a pure astrocyte culture.
19. Monitor the purity of the astrocyte cultures by visual inspection under the light microscope. If residual phase-contrast bright progenitor cells and microglia still reside on top of the astrocytic monolayer (see step 1.16), repeat steps 1.18 and 1.19.
    NOTE: The confluent and pure astrocyte monolayers can be maintained for up to 14 d in the incubator (37 °C, 6% v/v CO₂) before proceeding to the next step. Change half of the medium every 3 d. Do not collect, freeze and thaw the astrocytes, as these cells will lose their neuronal supportive properties through the storage procedure.

2. Preparing Astrocytes for the Indirect Coculture

NOTE: 48 – 72 h before preparing the neurons, transfer astrocytes to the cell-culture inserts. Generally, a single T75 flask delivers a sufficient number of cells to support the neuronal cultures obtained with one preparation.

1. Place as many inserts as needed into 24-well-plates. Coat each insert with 10 µg/mL PDL (dissolved in cell-culture-grade water) for about 1 h at 37 °C, 6% v/v CO₂. After the incubation, wash the inserts twice with PBS. In the meantime, proceed with the preparation of astrocytes.       
   NOTE: The inserts filled with PBS can be kept until the astrocyte cell suspension is ready for use.
2. Aspirate the astrocyte medium (with AraC) and wash the culture one time with 10 mL PBS to ensure any residual serum is removed.
3. Add 3 mL of 0.05% trypsin-EDTA (Ethylenediaminetetraacetic acid) to the flasks and incubate the cells for trypsinization at 37 °C, 6% v/v CO₂ for approximately 10 min.
4. Control the cell detachment by visual inspection using a microscope and facilitate the detachment of the cells by tapping the side of the flasks against the desktop.
   NOTE: Steps 2.5 – 2.8 are critical and should be finished within 15 – 20 min.
5. Following the trypsinization, gently resuspend the cells in 7 mL astrocyte medium and transfer the cell suspension to a 15 mL tube. Centrifuge for 5 min at 216 x g.
6. Carefully aspirate the supernatant and resuspend the cell pellet in 1 mL astrocyte medium.
7. Count the cells using a counting chamber.
8. Fill the bottom of the wells of the 24-well-plate with 500 µL astrocyte medium and transfer 25,000 cells in 500 µL astrocyte medium into each individual insert. Incubate the culture at 37 °C, 6% v/v CO₂. 
   NOTE: After 42 – 72 h the cultures will reach approximately 100% confluence. At this stage the purity of the culture can be verified by immunocytochemistry. The confluent cultures can be maintained for up to 7 d until proceeding to the next step. Change half of the medium every 3 days.

3. Preparation of Primary Hippocampal Neurons

NOTE: Primary mouse hippocampal neurons should be derived from E15.5 – E16 embryos of timed pregnant mice.

1. Prepare neuron medium (MEM with 10 mM sodium pyruvate, 0.1% w/v ovalbumin, 2% v/v B27 medium supplement, and 0.1% v/v gentamicin (sterile filtered)), and preparation medium (HBSS with 0.6% w/v glucose and 10mM HEPES (4-(2-hydroxyethyl)-1-piperazine ethanesulfonic acid)). Pre-warm and pre-equilibrate the neuron medium in filter flasks at 37 °C, 6% v/v CO₂.
2. Place the glass-coverslips (12 mm diameter) into the wells of 24-well-plates (use the special coverslips, which are notched for the use with cell culture inserts).
   1. Coat for at least 1 h with 15 µg/mL Poly-DL-Ornithine in water (cell-culture-grade) at 37 °C. Use at least 500 µL per well to ensure that the coverslips are completely covered. Wash the wells twice with PBS. Leave the PBS in the wells until plating the cells. Ensure that the wells do not dry out.
3. To dissect the hippocampi, prepare 3x 10 cm dishes filled with preparation medium and 1x 2 mL tube with 1 mL preparation medium. Prepare scissors and 2 fine forceps.
4. Sacrifice the pregnant mouse by cervical dislocation, sterilize the abdomen by washing with 70% v/v ethanol, and open the abdomen using sharp scissors. Excise the beaded uterus carefully with scissors and transfer the organ into 1x 10 cm dish filled with preparation medium.
5. Next, remove the embryos from the uterus. Carefully incise the uterus and remove the amnion without harming the embryos by using 2 forceps. Thereafter, decapitate each embryo with a pair of scissors and transfer the heads into a second 10 cm dish supplied with preparation medium.
6. Proceed with the preparation of the heads (similar to the preparation described in 1.4 – 1.7).
   1. Immobilize the head with a pair of forceps and incise the skull and skin in the neck region close to the hindbrain, perpendicular to the neuraxis. Use a second pair of forceps to lift both skull and covering skin, and bend the soft skull cap towards the rostral end of the head, thereby exposing the brain.
   2. With the help of 2 forceps, detach the brain from the cranial cavity. To do this, close 1 pair of forceps and separate the brain from the skull starting at the olfactory bulbs and moving towards the hindbrain. Collect the brains in another 10 cm dish filled with preparation medium.
7. Dissect the hippocampi from the cortex moieties as described above for the astrocyte preparation (step 1.6). Remove the hippocampi from each hemisphere and collect them in the 2 mL tube filled with preparation medium.
8. After completing the dissection, transfer the tube to the sterile laminar flow bench and digest the hippocampal tissue with 1 mL digestion solution containing papain. Prepare the digestion solution as described in step 1.9 – 1.10, but use MEM instead DMEM.
   NOTE: Steps 3.9 – 3.12 are critical and should be completed within 10 – 15 min.
9. After completion of digestion, carefully withdraw the digestion solution by gentle suction with a pipette.
10. Wash the hippocampi 3 times with neuron medium by sequentially adding and thereafter carefully aspirating 1 mL fresh culture medium per wash cycle. Do not centrifuge the cell suspension.
11. After the final washing step, carefully triturate the tissue in 1 mL neuron medium.
12. Count the cells using a counting chamber and plate 35,000 cells in 500 µL neuron medium per a single well of a 24-well-plate (the plate mentioned in 3.2).
    NOTE: Optionally, a cell aliquot can be counterstained with trypan blue to assess the vitality of the cell preparation. Control the plating density by visual inspection using the microscope (typically 90 – 110 cells per visual field of a standard 10X objective).
13. Place neurons in the incubator at 37 °C, 6% v/v CO₂ for 1 h.
14. Post incubation, take the prepared inserts (seeded with confluent astrocyte monolayers) out of the incubator (see step 2.8). Exchange the medium by aspirating the astrocyte medium and replacing it with 500 µl fresh neuron medium.
    NOTE: The inserts should harbor confluent astrocyte monolayers after 2 – 3 DIV (Days In Vitro).
15. Place the inserts with astrocytes carefully into the wells that contain the neuron cultures (step 3.13) using sterile forceps.
16. Place the resulting indirect neuron-astrocyte co-culture back into the incubator at 37 °C, 6% v/v CO₂.
    NOTE: Under these conditions, neurons can be cultivated for up to 4 weeks in a completely defined medium. No medium change is necessary. For long-time cultures, it is recommended to fill empty wells with sterile water in order to reduce the evaporation from the 24-well plate.