1. Cultivation of forebrain neurons from embryonic mice (E13.5)
1.1.1. Coating of cell culture plates for neurons
1. Transfer one coverslip into each well of a 24- or 6- well culture plate.
2. Add 500- 1000 µl of coating solution (20 µg/ml Poly-D-Lysine in DPBS) into each well.
3. Incubate plate at 37°C for at least 2 hours (we recommend overnight incubation), e.g., in a humidified cell culture incubator.
4. Wash twice thoroughly with DPBS (see Note 1).
5. Store plates in the final washing solution for use on the same day.
6. For long-term storage (up to 3 weeks): let plates dry completely at room temperature, store at 4°C.
1.1.2. Dissection of embryonic forebrain
Before starting: Sterilize preparation tools (one large and one small scissors, one large and two small forceps) using ethanol. It is essential to work as fast and sterile as possible, using pre-cooled HBSS + 1x AA for dissection. In addition, it is crucial to work on ice during the whole dissection procedure.
1. Euthanize a pregnant mouse at day 13.5 of gestation, dissect mouse embryos and keep heads on ice.
2. Remove scalp of dissected heads.
3. Extract the entire brain carefully and remove meninges with forceps (thoroughly!); it is crucial to remove the meninges completely to avoid endothelial cell growth.
4. If necessary: Remove olfactory bulbs.
5. Gently divide the brainstem and cortical hemispheres.
6. If necessary: Remove remaining meninges patches.
7. Transfer cortical hemispheres of each animal in different reaction tubes (filled with HBSS) and store them on ice. Note: Cortices from different animals can be pooled to increase yield.
8. Optional: Use other parts of the brain, e.g., brain stem, to culture glial cells.
1.1.3. Cell isolation and plating
Transfer dissected brains to a sterile clean bench and proceed with cell isolation.
1. Carefully remove supernatant and wash twice using pre-warmed HBSS.
2. Add 1x Trypsin and incubate for 7 minutes at 37°C, swirl 2-3 times during incubation.
3. Inactivate Trypsin with 10% FBS in HBSS by doubling the volume.
4. Discard the supernatant and dilute cells with HBSS.
5. Gently but thoroughly triturate cells using a 1 ml pipette.
6. Count cells using Trypan Blue (1450022, BioRad).
7. Dilute cells to desired density using pre-warmed (37°C) NPM (see Notes 2 and 3):
a. For a 24-Well plate: seed 200.000 cells/ well in 500 µl.
b. For a 6-Well plate: seed 800.000 cells/ well in 1 ml.
8. Transfer plate immediately into a humidified cell culture incubator, at 37°C and 5% CO2.
1.1.4. Neuronal maintenance
1. Grow mouse neurons for 4 days after plating in a humidified cell culture incubator at 37°C and 5% CO2
2. Freshly prepare NMM with 1 µg/ml AraC (see Note 4).
3. Double the volume of the cell culture medium by adding NMM including AraC (see also Note 5). Cells can now be grown for at least 4 weeks, depending on the desired analysis (see Note 6).
2. Cultivation of hippocampal and cortical neurons from postnatal mice (P0-P3)
2.1.1. Coating of coverslips for neurons
1. Prepare coverslips for top-down culture: Heat paraffin in a blue-cap glass bottle (VWR, e.g., 215-1514) in a water bath at 70-80°C (CAVE: do not microwave or cook, Paraffin wax is ignitive! Only use a water bath! Loosen blue cap!). Meanwhile, put sterile coverslips side-to-side into a petri dish and add 3 drops of liquid and sterile Paraffin warmed to ~70°C onto each coverslip with a sterile glass Pasteur pipette in a tripod-shape. Let dry under the hood until the drops get white and solid for at least 10 minutes. Store coverslips sealed with parafilm at room temperature.
2. Put five 12 mm coverslips in each well of a 30 mm petri dish.
3. Add 90 µl for 12 mm (200 µl for 18 mm coverslips) of coating solution (1 mg/ml Poly-L-Lysine in borate buffer) to each coverslip.
4. Incubate coverslips at 37°C for at least 2-3 hours (best is overnight), e.g., in a humidified cell culture incubator.
5. Wash twice with sterile ddH2O, once with 1x HBSS (without phenol red!; see Note 1).
6. Dry coverslips for 5 minutes under a sterile hood.
7. Add 4 ml plating medium per petri dish containing the coated coverslips and store them in the incubator at 37°C until needed.
8. For long-term storage (up to 3 weeks): let coverslips dry completely at room temperature under the sterile hood, wrap in aluminum foil, and store at 4°C.
2.1.2. Dissection of hippocampi/ cortices
Before starting: Sterilize preparation tools (scissors (big and small), one big and two small forceps) using ethanol. It is essential to work as fast and sterile as possible, using pre-cooled HBSS + 1x AA for dissection. All dissection steps should be performed on ice. We regularly use the pooled hippocampi of at least three animals per genotype, which yields (in the case of three animals each) sufficient material for ten 12 mm or three 18 mm coverslips.
1. Euthanize the pub by decapitation using a sharp scissor.
2. Place head in a dish filled with HBSS +1x AA kept on ice.
3. Remove scalp of dissected heads.
4. Carefully remove the skull using fine forceps.
5. Extract the entire brain carefully.
6. Optional: To reduce the risk of contamination, transfer brains to a fresh dish filled with HBSS+1x AA.
7. Gently divide the brainstem and the cortical hemispheres.
8. Gently remove the midbrain and thalamic tissue from each hemisphere.
9. Carefully remove meninges (thoroughly!); it is crucial to remove the meninges completely to avoid the growth of glial/endothelial cells.
10. Dissect the hippocampus or cortex and collect in a 15 ml falcon tube (filled with 5 ml HBSS+1xAA, on ice). Hippocampi/ cortices can be pooled or processed individually.
2.1.3. Cell isolation and plating
Transfer dissected hippocampi/ cortices to a sterile clean bench and proceed with cell isolation.
1. Carefully remove supernatant and wash 3 times using 5 ml of HBSS.
2. Add 5 ml of Trypsin solution (maximum of 3 half-cortices or 6 hippocampi) and incubate for 25 minutes at 37°C, swirling several times during incubation.
3. Discard the supernatant and wash tissue three times with 5 ml of HBSS.
4. Add 2 ml NPM and 2 µl DNase I (improves dissociation by reducing shear forces).
5. Gently but thoroughly triturate cells using three fire-polished glass pipettes of decreasing diameter (ranging from original size (but polished) to reduced size that allows still smoothly aspirating fluid, but avoiding pressure differences due to high resistance of too small polished pipettes).
6. Count cells using Trypan Blue (1450022, BioRad).
7. Dilute cells to desired density using pre-warmed (37°C) NPM (see Notes 2 and 3) and plate at the desired density directly onto the coverslips in Petri dishes:
Generally, distribute hippocampal neurons from 3 animals onto ten 12 mm coverslips or three 18 mm coverslips, respectively. Cortical cultures are more complex/sensitive to plating density, so the optimal cell density is between 30.000 to 60.000 cells/12 mm coverslip.
8. Transfer neurons immediately into a humidified cell culture incubator, at 37°C and 5% CO2. Let cells attach for 45 - 60 minutes.
9. Afterwards, flip 12 mm coverslips (neurons facing down) and transfer to a 24-well plate filled with NMM. For 18 mm coverslips use 12-well-plates filled with 1 ml of NMM.
2.1.4. Neuronal maintenance
1. Grow mouse neurons for 2-3 days after plating in a humidified cell culture incubator at 37°C and 5% CO2.
2. Freshly prepare NMM with 2.5 µM AraC (see Note 4).
3. Double the volume of the cell culture medium by adding pre-warmed NMM including AraC (see also Note 5). Cells can now be grown for up to 1 month, depending on the desired analysis (see Note 6).
3. Fixation and staining of primary neurons for endogenous TAU trafficking
3.1.1. Fixation and staining of neurons
At the desired stage of TAU development, neurons can then be treated (see Note 7) and fixed for immunofluorescence analysis.
1. Remove cell culture medium completely and immediately add 3.7% fixation solution (see Note 8).
Alternatively: directly add 7.4% FA onto the cells (with medium still in the wells, doubling the total amount of liquid, thus resulting in a final concentration of 3.7% FA).
2. Incubate for 30 minutes at room temperature (see Note 9).
3. Wash cells once with DPBS.
4. Pause step: It is possible to directly store the cells for up to 2 years after fixation using 60% glycerol diluted in DPBS, store plates at -20°C. Let cells adjust to room temperature before continuing with the staining procedure.
5. Wash coverslips two times with DPBS.
6. Add B+P solution for 5- 10 minutes at room temperature.
Optional: add 1 drop of NucBlue to the B+P solution incubate for 10 minutes at room temperature. Omit NucBlue in the last washing step of step 14.
7. Wash twice with DPBS.
8. Prepare a parafilm-coated lid and add a drop of antibody solution (30 µl for 12 mm coverslip and 80-100 µl for 25 mm coverslip) for each coverslip to be stained.
9. Transfer coverslip with cells facing down to the parafilm-coated plate and place the well-plate lid in a moist chamber (containing wet towels) for incubation.
10. Incubate 16 hours at 4°C.
11. Transfer coverslips back to well-plates (cells facing up) and wash 3 times with DPBS.
12. Proceed with secondary antibody staining (optional: include Phalloidin) as described before (see points 7. and 8.)
13. Incubate for 2 hours at 37°C in the dark in a moist chamber (see Note 10).
14. Transfer coverslips back to well-plates (cells facing up) and wash 3 times with DPBS. In the last washing step: add 1 drop of NucBlue to each well and incubate for 20 minutes.
15. Wash coverslip once with DPBS and twice with ddH2O.
16. Mount coverslips on a microcopy glass slide upside down with a drop (~5 µl) of mounting medium. Take care that coverslips are mounted as plane as possible.
17. Allow mounting medium to polymerize overnight at room temperature before imaging. Microscopy slides can be stored at 4°C in the dark for at least 5 years with only a slight decrease in fluorescence intensity.