Low-Density Primary Hippocampal Neuron Culture

All experiments and protocols using laboratory animals were approved by the University of Manitoba animal ethics committee and were compliant with the guidelines of the Canadian Council on Animal Care.

1. Preparation of Instruments, Buffers and Solutions

1. Sterilize by autoclaving all dissection equipment, glass Pasteur pipettes, pipette tips, filter apparatus, and deionized water.
2. Prepare a 20% (w/v; 1.1 M) stock glucose solution in deionized water and filter-sterilize. Store at 4 °C.
3. Prepare a 100 mM sodium pyruvate solution in deionized water and filter-sterilize. Store at 4 °C.
4. Prepare calcium- and magnesium-free Hank's Balanced Salt Solution (CMF-HBSS) by making a solution of 10% HBSS (10x), 10 mM HEPES, and 100 U/mL penicillin-streptomycin in deionized water. Store at 4 °C for no more than 1 to 2 months.
5. Prepare 10 mg/mL deoxyribonuclease I (DNase) in CMF-HBSS, and then filter-sterilize and store at -20 °C.
6. Prepare glial medium by making a solution of 30 mM glucose (from the sterile solution), 95 U/mL penicillin-streptomycin, and 10-15% bovine growth serum (BGS) in Minimum Essential Medium (MEM).
   NOTE: As indicated in the protocol, a solution of 5% BGS can be used if required to slow cell proliferation. Horse Serum may be used instead of BGS, but note that each batch must be tested before use due to inter-batch variation. Store the prepared solution at 4 °C for no more than 1 to 2 months. Though many laboratories use FBS, we consistently observed that glial growth in BGS is as robust as in FBS. BGS is derived from fetal calf serum which is supplemented with chemically defined components. In addition, BGS is substantially more economical than FBS.
7. Prepare the neuron growth and maintenance medium (neurobasal-B27, NBG) by making a solution of 1x B27 supplement and 0.5 mM L-Glutamine in 500 mL Neurobasal Medium. Store at 4 °C for no more than 1 to 2 months. L-Glutamine can be substituted with GlutaMAX, which has been suggested to be more stable in the growth medium.
8. Prepare the neuron plating medium by making a solution of 30 mM glucose (from the sterile solution), 1 mM sodium pyruvate (from the sterile solution), and 10% BGS in MEM. Store at 4 °C for no more than 1 to 2 months.
9. Prepare a solution of 1 mM Cytarabine (Ara-C) and filter-sterilize. Aliquot into 1 mL portions and store at -20 °C.
10. Prepare a solution of 100 mM APV in 100 mN filter-sterilized NaOH. Store at -20 °C.
11. Dissolve 1 mg/mL poly-L-lysine in borate buffer, pH 8.5 (50 mM boric acid and 12 mM borax) and filter-sterilize. Aliquot and store at -20 °C. Instead of poly-L-lysine, one may use poly-L-ornithine which may be less toxic to cells.

2. Glia Culture Preparation

1. Prepare cortical astroglia cell culture
   1. Obtain rat pups within 24 h of birth. Place the pups on ice to anaesthetize them.
   2. Spray the pups with 70% ethanol and decapitate them.
   3. Place the heads in a 100 mm dish on ice containing 15 mL CMF-HBSS.
   4. Dissect out each brain by cutting the scalp, opening the skull, severing the brainstem, and then transferring the brain to 60 mm Petri dishes on ice containing 3 mL CMF-HBSS.
   5. Separate the cerebral hemispheres from the hippocampi and then strip away the meninges surrounding them. Exercise adequate precaution to ensure minimal contamination from meningeal fibroblasts. Meningeal fibroblasts in the culture can compete with glia for growth and be detrimental for neuron health. Collect all the cerebral hemispheres in a 60 mm Petri dish on ice containing 5 mL CMF-HBSS.
   6. Remove the CMF-HBSS and then mince the collected cortical tissue as finely as possible using micro-dissecting spring-scissors.
   7. Add sufficient CMF-HBSS to the chopped tissue. Using a glass Pasteur pipette, transfer the tissue pieces to a 15 mL tube. Bring the total volume to 12 mL by adding CMF-HBSS.
   8. Add 1.5 mL each of 2.5% trypsin and 10 mg/mL DNase solution. Incubate the resulting mixture at 37 °C in a water bath for 12 min.
   9. Triturate the tissue 10-15 times using a glass Pasteur pipette, and further incubate it in a 37 °C water bath for 3 min.
   10. Filter the supernatant through lens paper or a filter mesh into a 50 mL tube containing 5 mL of BGS. This is done to remove chunks of undissociated tissue. Alternatively, use commercially available cell strainers.
   11. Add 13.5 mL of CMF-HBSS and 1.5 mL of 2.5% trypsin to the tube containing the remaining tissue pieces and further incubate it at 37 °C in a water bath for an additional 10 min.
   12. Triturate the remaining tissue once more, and then filter the suspension again into the 50 mL tube containing the initial filtrate. Next, rinse the lens paper with 3 mL of BGS. The final volume should be approximately 38 mL.
   13. Centrifuge the suspension containing the dissociated glial cells for 6 min at 130 x g. Carefully discard the supernatant using a glass Pasteur pipette. Ensure that the pelleted cells at the bottom are not disturbed. The bottom of the pellet appears slightly reddish containing blood components.
   14. Transfer the glial cells in 1 mL of glial medium into a fresh tube, taking care not to disturb the reddish portion of the pellet.
   15. Add up to 2 mL of glial medium per pup to resuspend the combined dissociated glial cells. For example, if 10 pups were used, and then the total resuspension volume would be 20 mL.
   16. Prepare one 75 cm² cell culture flask per pup. Add 18 mL pre-warmed glial medium to each flask.
   17. Transfer 2 mL of cell suspension to each flask and incubate in a 37 °C, 5% CO₂ incubator.
   18. After one day, replace with 20 mL glial medium. At this stage, the culture is a mix of glial and other unwanted cell-types.
   19. Four days after seeding the cells, vigorously shake the flasks to dislodge non-astrocyte cells. To do this, rinse the flasks once with CMF-HBSS, and then add 10 mL fresh CMF-HBSS to each flask. Shake the flasks vigorously 5-10 times, and then aspirate off the CMF-HBSS. Rinse twice with CMF-HBSS, and then add 20 mL glial medium.
       NOTE: This step is crucial to ensure the removal of non-target cell types, and should not result in the loss of the desired astrocytes.
   20. Replace with fresh glial medium twice a week until the cells are confluent (Figure 1).
2. Freezing glia
   1. Once the cultures are confluent, aspirate off the medium and wash the cell monolayer with 10 mL CMF-HBSS.
   2. Add 10 mL of 0.25% trypsin/EDTA to each flask and incubate at 37 °C for 3 min.
   3. Tap flasks gently to dislodge the cells. Add 1 mL BGS to each flask and transfer the cell suspensions into 15 mL tubes.
   4. Centrifuge the tubes at 130 x g for 6 min. Discard the resulting supernatant.
   5. Resuspend the cell pellet in 2 mL glial medium.
   6. Prepare the glial freezing medium (1-part dimethyl sulfoxide and 4-parts BGS). Add 0.5 mL of this solution to each of 4 cryogenic vials per flask.
   7. Transfer 0.5 mL of cell suspension to each vial, and then freeze the cells slowly by placing the vials in an insulated container in a -80 °C freezer overnight. After one day, transfer the vials to a liquid nitrogen freezer for long-term storage.
3. Plating revived glia
   1. Plate glia 1-2 weeks before the neuron culture day.
   2. Prepare a 50 mL tube with 10 mL warmed glial medium.
   3. Thaw 1 frozen vial of glia in a 37 °C water bath for 2-3 min.
   4. Transfer the contents of the vial into the 50 mL tube containing 10 mL glial medium.
   5. Centrifuge the tube for 5 min at 130 x g and aspirate off the supernatant.
   6. Resuspend the pellet in 20 mL glial medium.
   7. Add 3 mL fresh glial medium to each of the culture dishes (60 mm), and then transfer 1 mL of the cell suspension to each dish. Incubate the cultures in a 37 °C, 5% CO₂ incubator.
   8. Feed the cells twice a week. If the cell density reaches 50% confluence well before the neuron culture day, switch to glial medium containing 5% BGS to slow the glial growth rate. The desired confluence of the glial monolayer for neuron culture is 50% – 70%.
   9. Replace the glial medium with NBG medium (6 mL/dish) 1-2 days before the neuron culture.

3. Coverslip Preparation

1. Cleaning and Preparation of Paraffin Beads
   1. Submerge coverslips in concentrated nitric acid (70% wt/wt; CAUTION: highly corrosive) and sonicate them for 30 min at room temperature. Other laboratories have found it beneficial to incubate the coverslips in 70% nitric acid in ceramic racks for 12-24 h at room temperature instead.
   2. Carefully discard the nitric acid in a designated chemical fume hood in accordance to institutional guidelines. Rinse the coverslips 3-5 times with deionized water.
   3. Submerge the coverslips in deionized water and sonicate them again for 30 min (skip if following the alternative method).
   4. Remove the deionized water and rinse the coverslips another three times.
   5. Submerge the coverslips in 50 mL deionized water and sterilize them by autoclaving. Alternatively, sterilize the coverslips at 225 °C in an oven for 6 h. If using this alternative approach, skip to step 3.1.9.
   6. Once autoclaved, transfer the coverslips to a sterile laminar flow hood for the following steps.
   7. Remove the deionized water and rinse with 20-30 mL of 100% ethanol.
   8. Add 20-30 mL of 100% ethanol and transfer the coverslips and ethanol to a 100 mm Petri dish.
   9. Use blunt tweezers to transfer coverslips to 60 mm Petri dishes, 4-5 coverslips per dish, and then allow them to air-dry by leaning the coverslips against the edge of the dish. Once the ethanol has evaporated, lay the coverslips flat on the surface.
   10. Transfer 10 g paraffin to a suitable heat-resistant glass bottle. Melt the paraffin and maintain it at about 150-200 °C on a hot plate for at least 1 h. The ideal temperature to which the paraffin should be heated may take some tweaking. Non-ideal temperatures can lead to difficulty producing the correct size of the paraffin beads. The waiting period after melting helps to ensure consistent temperature throughout the liquid.
   11. Dip a Pasteur pipette into the paraffin, and then quickly touch it to three spots near the edge of each coverslip (each spot being approximately 120° apart). The drops will solidify into beads approximately 0.3-0.5 mm high and 1.0-2.0 mm in diameter and will function to provide a space between the neurons on the coverslip and the glial monolayer below (Figure 2).
   12. Sterilize the dishes under UV light for 30 min, and then cover them with aluminum foil and store at room temperature. Prepared coverslips can be stored and used for up to one month. Paraffin beads should be allowed to adhere to the coverslips for at least one week before use. This will prevent the paraffin from detaching during the remaining steps of the protocol.
2. Coating Coverslips with Poly-L-Lysine
   1. Obtain Petri dishes containing previously prepared coverslips with the paraffin beads.
   2. Add 200 µL of poly-L-lysine in borate buffer to the surface of each coverslip and let the solution sit on the coverslips overnight, covered, at room temperature. Coat the same surface as the one on which the paraffin beads are situated. Note that if properly cleaned, the poly-L-lysine should spread easily to cover the surface of the coverslip.
   3. After one day, wash the coverslips twice by soaking them in sterile deionized water for 2 h each time. After the final wash, replace the water with 4 mL of neuronal plating medium per dish. Note that it is important that the surface of the coverslips not be allowed to dry after having been coated with poly-L-lysine.
   4. Place the coverslip-containing dishes in a 37 °C, 5% CO₂ incubator. The coated coverslips should be incubated for at least 24 h before the neuron culture. This is done to prime the medium for neuron culture.

4. Dissection of Hippocampus and Plating Neurons

1. Obtain a pregnant rat (embryonic day 18-19, as measured from the day of vaginal plug discovery) and euthanize by isoflurane-anesthetized decapitation or other approved method.
2. Spray the underside of the rat with 70% ethanol, and then make an incision from the pubic region through to the end of the abdominal cavity. To avoid contamination, take care to cut only through the skin at this step.
3. Rinse the dissection instruments again with 70% ethanol, and then cut through the abdominal wall to expose the uterus. Remove the two uterine horns and place them in a sterile 100 mm Petri dish.
4. Perform the remaining steps in a laminar flow hood. Remove the uterine membrane surrounding the fetuses. Decapitate them and place the heads in a 100 mm Petri dish containing 20 mL CMF-HBSS.
5. Dissect out the brains and immediately place them in a 60 mm Petri dish containing 2 mL CMF-HBSS on ice. Collect only 2-3 brains per dish to allow sufficient space for the next step.
6. Under a dissecting microscope strip away the meninges from the midline of the cerebral hemispheres, and then dissect out the hippocampi and place them in a 60 mm Petri dish containing 4.5 mL CMF-HBSS. Note that the hippocampus can be distinguished as a crescent-shaped structure in the medial surface of the cortical hemisphere, and is easy to remove as it is bordered laterally by a ventricle.
7. Transfer the collected hippocampal tissue and CMF-HBSS to a 15 mL tube. Add 0.5 mL of 2.5% trypsin, and then incubate at 37 °C for 10 min. Papain may be used instead of trypsin because it is gentler and may increase yield.
8. Gently remove the trypsin solution by Pasteur pipette, leaving the hippocampal tissue undisturbed at the bottom of the tube.
9. Gently wash the tissue twice with 5 mL CMF-HBSS while incubating it at 37 °C for 5 min. After the second wash, bring the total volume to at least 5 mL by adding a sufficient volume of CMF-HBSS to the tissue. If there are hippocampi from more than 10 pups, up to 8 mL CMF-HBSS can be added.
10. Prepare two variants of fire-polished pipettes for tissue dissociation. The tip of one variant is of the normal diameter of the pipette but with fire-polished smoothened edges. The other variant is one with the tip diameter reduced by half. Briefly expose the tip of a glass Pasteur pipette diagonally to a flame source, and then examine the tips of the pipettes.
    1. Repeat this step until the tip edge had been smoothened or the diameter has been reduced. It is important to practice to find the ideal diameter to ensure that the tissue is dissociated to yield single cells without damaging these cells.
11. Triturate the hippocampal tissue first by about 10 passes through a smooth-edged normal glass Pasteur pipette, and then a further 5 to 10 passes through the pipette with the reduced diameter. The goal is to obtain a homogenous solution of cells with no or very few tissue clumps.
12. Determine the cell density using a hemocytometer or an automated cell counter. Typically, 0.8 to 1.0 million cells per pup are obtained when hippocampi from the two hemispheres are combined.
13. Transfer a sufficient volume of the cell suspension to dishes containing poly-L-lysine-coated coverslips in 4 mL plating medium to obtain a plating density of 300,000 cells per 60 mm dish. The density of the cells can vary from 100,000 to 500,000, depending upon the intended experiment.
14. After 3-4 h, transfer the coverslips with neurons attached to dishes containing glial cells in NBG medium. This should be done such that the side on which the neurons were plated is facing down towards the glial cell layer. Add a total of 4-5 coverslips per glia dish.
15. Two or three days after plating, add the Ara-C solution to a final concentration of 5 µM per dish to arrest glial growth.
16. Feed the cells once a week by replacing 2 mL of the old medium with 2.5 mL of fresh medium at each feeding. The extra medium added is to compensate for evaporation over time, and only a portion of the medium is changed to ensure consistent conditioning of the medium by the glial cells. These cultures are typically healthy for at least one month (Figure 3). Neuron survival can be improved by adding APV to the culture medium to a final concentration of 100 µM. APV is an NMDA receptor antagonist and promotes survival by reducing glutamate excitotoxicity.