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All procedures involving animal samples have been reviewed and approved by the appropriate animal ethical review committee.
1. Primary Hippocampal Cell Culture
- Prepare the dissociated cell culture of the rat hippocampus on embryonic day 19. Plate the cells on 12 mm coverslips coated with polyethyleneimine (PEI) in 24-well dishes at a density of 50,000 - 60,000 cells/well. Check the density using a cell counting chamber and phase contrast optics.
- Culture the neurons for 3 days (day in vitro (DIV) 3) in a 24-well plate in the incubator at 37 °C with 5% CO2.
- Assess the coverslips for indicators of cell health using transmitted light microscopy (e.g., phase contrast optics at a magnification of 10 - 20X). Check for the following indicators of good health: a clear phase contrast halo, neurites without beaded structures, and no soma clustering or neurite bundling.
2. Transfection
NOTE: The following protocol refers to a double-transfection for 3 wells. However, the protocol works best when amounts sufficient for 4 wells are prepared.
- Prepare 500 mL of transfection buffer (274 mM sodium chloride, NaCl, 10 mM potassium chloride, KCl, 1.4 mM disodium hydrogen phosphate, Na2HPO4, 15 mM glucose, 42 mM 4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid, HEPES) in an Erlenmeyer flask.
- Dissolve 8.0 g of NaCl, 0.37 g of KCl, 0.095 g of Na2HPO4, 1.35 g of glucose, and 5.0 g of HEPES in 400 mL of distilled water in an Erlenmeyer flask.
- Adjust the pH to 6.95 with 1 M sodium hydroxide, NaOH using a pH meter.
- Adjust the volume with distilled water to 500 mL and check the pH using a pH meter.
- Make 20 - 30 mL aliquots of transfection buffer with the following pH values by pipetting 1 M NaOH to the transfection buffer: 6.96, 6.97, 6.98, 6.99, 7.00, 7.01, 7.02, 7.03, 7.04, 7.05, 7.06, 7.07, 7.08, 7.09, 7.11.
NOTE: The pH of the transfection buffer is crucial for the transfection efficacy.
- To test which transfection buffer leads to the highest number of transfected cells, test each pH value from 6.96 to 7.11. Use the transfection method described in 2.2 - 2.11 and a validated plasmid expressing green fluorescent protein (GFP). Determine the number of transfected cells per coverslip for every transfection buffer pH value to assess which buffer works the best.
- Aliquot the buffer with the highest transfection efficiency into 2 mL microcentrifuge tubes Freeze and store the tubes at -20 °C.
- Pre-warm the reduced serum medium, cell culture medium, and distilled water to 37 °C in the water bath.
- Prepare the transfection mix in a 1.5 mL microcentrifuge tube. Work under the laminar flow hood to ensure sterile working conditions.
- Mix 7.5 µL of 2 M calcium chloride with 4 µg of each endotoxin-free DNA (Synaptophysin-mOrange and mGFP/GFP-Rogdi). Add water to reach a total volume of 60 µL in a 1.5 mL microcentrifuge tube.
- Add 60 µL of transfection buffer to the mix. To obtain the best results, add the transfection buffer dropwise while shaking the DNA-mix gently on the vortex.
- Incubate at room temperature (RT) for 20 minutes. Avoid shaking the incubation tube during the incubation time by placing the tube next to the laminar flow hood.
- Under the laminar flow hood, remove the cell culture medium ("conditioned medium") from the wells using a 1000 µL pipet and store it in a separate container in the incubator.
- Add 500 µL of reduced serum medium to each well. Incubate the cells at 37 °C and 5% CO2 until the 20-minute incubation period (step 2.3.3) is over.
- Add 30 µL of transfection mix to each well by pipetting several drops. Discard the residue at the bottom of the tube.
- After all the wells have been supplied with transfection mix, gently shake the 24-well plate to ensure the distribution of the transfection mix in the medium.
- Incubate the wells for 60 minutes at 37 °C and 5% CO2.
- Remove and discard the transfection mix and wash it three times with cell culture medium. Add 1 mL of cell culture medium to each well and incubate them for 30 seconds at RT. Remove 750 µL of medium and add the same amount of fresh medium. Repeat this three times.
NOTE: The washing step is critical. Keep the time that each well has no medium at a minimum (i.e., remove and replace well-by-well) and add the washing medium gently.
- Remove and discard the cell culture medium and add 450 µL of the conditioned medium well-by-well.
- Let the neurons mature in the incubator at 37 °C and 5% CO2 to DIV 10.
3. Stimulation and Syt1 Uptake
NOTE: The following protocol applies the uptake to 3 wells. For depolarization of any other number of wells, adjust the amounts accordingly.
- Prepare 50 mL of 10x depolarization buffer (640 mM NaCl, 700 mM KCl, 10 mM magnesium chloride, MgCl2, 20 mM calcium chloride, CaCl2, 300 mM glucose, 200 mM HEPES, pH 7.4) in an Erlenmeyer flask.
NOTE: Depolarization buffer can be kept at 4 °C for several weeks. If a non-depolarizing solution is also used, prepare a 10x Tyrode's solution consisting of 1290 mM NaCl, 50 mM KCl, 10 mM MgCl2, 20 mM CaCl2, 300 mM Glucose, 200 mM HEPES pH 7.4 in order to compare stimulation-induced recycling with spontaneous recycling. After dilution to 1x, add 1 µM of Tetrodotoxin before use to block action potential generation.
- Dissolve 1.87 g of NaCl, 2.61 g of KCl, 0.1 g of MgCl2-6H2O, 0.15 g of CaCl2-2H2O and 3.0 g of glucose-1 H2O and 2.38 g of HEPES in 50 mL of distilled water in an Erlenmeyer flask. Adjust the pH with NaOH and sterile-filter the solution. Dilute the buffer 1:10 in distilled water to achieve a 1x concentration.
- Prepare 4% paraformaldehyde (PFA) in 1x phosphate-buffered saline (PBS) (pH 7.4) for fixation after stimulation.
- For 500 mL of 4% PFA in 1x PBS, dissolve 20 g of paraformaldehyde in 450 mL of distilled H2O.
NOTE: Heating the solution may speed up dissolving, but do not heat the solution over 70 °C, as the PFA may disintegrate.
CAUTION: PFA is toxic, potentially carcinogenic, and teratogenic. Wear gloves when working with PFA, work under the fume hood, and avoid ingestion.
- Let the solution cool to RT and add 50 mL of 10x PBS stock solution. Adjust the pH to 7.4 with NaOH/HCl(hydrochloric acid) using a pH meter.
- Pre-warm 600 µL of 1x depolarization buffer and 10 mL of cell culture medium to 37 °C in the water bath.
- Add 1 µL of mouse anti-Syt1 antibody (clone 604.2) to the 1x depolarization buffer and vortex for 10 seconds.
- Remove and discard the cell culture medium from the cells. Add 200 µL of the depolarization-antibody mix to each well and incubate for 5 minutes at 37 °C and 5% CO2 in the incubator.
- Remove and discard the depolarization-antibody mix and wash it three times with cell culture medium. Add 1 mL of cell culture medium to each well and incubate for 30 seconds at RT. Remove 750 µL of medium and add the same amount of fresh medium. Repeat three times.
- Remove and discard the cell culture medium and add 300 µL of 4% PFA in 1x PBS. Incubate for 20 minutes at 4 °C.
- Wash three times for 5 minutes each with 1x PBS.
NOTE: The protocol can be paused here.
4. Immunocytochemistry
- Prepare 50 mL of blocking buffer.
NOTE: Blocking buffer can be kept at -20 °C for several months.
- Dissolve 2.5 g of sucrose and 1 g of bovine serum albumin (BSA) in 5 mL of 10x PBS stock solution. Add 1.5 mL of 10% detergent stock solution. Stir the solution until all the components have properly dissolved and add distilled H2O until reaching a final volume of 50 mL. Aliquot the solution and freeze the aliquots for storage.
- Dilute the secondary, fluorophore-coupled antibody (directed against the primary Syt1-antibody species) in 200 µL of blocking buffer in each well at a dilution of 1:1000.
- Remove and discard the 1x PBS from each well containing a coverslip.
- Add 200 µL of blocking buffer-antibody mix to each well and incubate for 60 minutes at RT.
CAUTION: Because the secondary antibodies are light-sensitive, all steps moving forward must be performed in the dark.
- After incubation, wash the cells three times for 5 minutes with 1 mL of 1x PBS.
- Embed the coverslips on microscope slides with the embedding medium.
- Add a 7 µL drop of embedding medium onto the microscope slide. Remove the coverslip from the 24-well plate by lifting it with a syringe and grabbing it with forceps.
CAUTION: Cells on the surface of the coverslip are easily damaged, so forceps must be handled with care.
- Dip the coverslip into distilled water to remove the PBS and dry it carefully by touching one edge to soft tissue.
- Flip the coverslip onto the embedding medium droplet, so that the surface carrying the cells faces the microscope slide, thereby embedding the cells into the embedding medium.
- Leave the slides to dry under the hood for 1 - 2 h (cover them to avoid light exposure) and store them in a microscope slide box at 4 °C.
NOTE: The protocol can be paused here.