Visualization of Thalamocortical Axon Branching and Synapse Formation in Organotypic Cocultures

All experiments were performed according to the guidelines established by the animal welfare committees of Osaka University and the Japan Neuroscience Society.

1. Organotypic cocultures of the thalamus and cerebral cortex

Note: For the detailed procedure, refer to the original publications^7,8,13. All procedures should be performed under sterile conditions. Sprague-Dawley (SD) rats are used for neuronal cultures.

1. Preparations of reagents
   Note: Volumes of the following reagents are for one rat brain.
   1. Prepare 10x modified N2 Supplement¹³: Insulin (50 μg/mL), progesterone (200 nM), hydrocortisone (200 nM), sodium selenite (300 nM), transferrin (1 mg/mL), putrescine (1 mM), glucose (60 mg/mL)
   2. Prepare serum-containing culture medium (10 mL): 85% DMEM/F12, 10% 10x modified N2 supplement, 5% fetal bovine serum (FBS)
   3. Prepare serum-free culture medium (20 mL): 90% DMEM/F12, 10% 10x modified N2 supplement
   4. Prepare rat tail collagen solution (0.5 mL). In brief, take tender fibers from a rat tail under sterile conditions and incubate in acetic acid solution overnight. After centrifugation, collect the supernatant (a few mg/mL) and store in a refrigerator¹³.
2. Preparation of culture dishes
   1. Place a membrane insert into a 35 mm Petri dish.
   2. Add 40 – 50 μL of rat tail collagen solution to the membrane, and spread the collagen solution around the center.
   3. Air dry the membrane completely in a clean bench.
   4. Put 1.5 – 2 mL of the serum-containing culture medium into the dish, so that the membrane is soaked with the medium.
   5. Place the culture dish in an incubator (37 °C, humidified 95% air, and 5% CO₂) before plating.
3. Preparation of cortical slices
   1. Sterilize all surgical instruments with 70% ethanol for 10 min or longer.
   2. Dissect the whole brain quickly from a postnatal day (P) 2 rat under ice-cold anesthesia (immersed in ice-chips for a few minutes).
   3. Place the brain into a 100 mm Petri dish containing 100 mL of cold Hanks’ balanced salt solution.
   4. Remove the pia matter from the brain with fine forceps, and cut cortical slices (300 – 400 μm thickness) from the visual and somatosensory cortex with microsurgical scissors (for detail, see Figure 1).
      Note: The thickness of cortical slices is roughly estimated by the scale of grids under a binocular microscope.
      Note: Alternatively, cortical slices can be made with a vibratome under sterile conditions.
   5. Transfer a few cortical slices onto the membrane insert with a disposable plastic pipette.
   6. Adjust the positions of the slices near the center of the culture insert with a fire-polished Pasteur pipette (for details, see^8,13) and remove excess medium from the insert.
      Note: Adjust the level of the medium slightly below the surface of the slices so that the slices can receive a sufficient supply of both the gas and medium. This is crucial for cell viability.
   7. Maintain the slices in the serum-containing culture medium at 37 °C in an environment of humidified 95% air and 5% CO₂, and culture alone for one day before a thalamic block is taken and placed next to it.
4. Preparation of thalamic blocks
   1. Sterilize all surgical instruments with 70% ethanol for 10 min or longer.
   2. Anesthetize a pregnant rat (embryonic day 15) by intraperitoneal injection of pentobarbital (50 mg/kg) containing anesthetic.
   3. Dissect each embryo from the uterine horns, and place the embryos into a 100 mm Petri dish containing 100 mL of cold Hanks’ balanced salt solution.
   4. Under a binocular microscope, remove the brain from each embryo and cut thalamic blocks which mainly contain the ventrobasal complex and lateral geniculate nucleus (approximately 0.5 mm x 3)⁷ using microsurgical scissors (see Figure 1).
5. Culture and maintain organotypic cocultures
   1. Using a Pasteur pipette, place the thalamic block next to the ventricular surface of the cortical slice which has been placed on the membrane filter.
   2. Maintain the cocultures in the serum-containing culture medium at 37 °C in an environment of humidified 95% air and 5% CO₂.
   3. Exchange half of the medium with the serum-free culture medium after two days in culture. Thereafter, exchange the medium every 2 – 3 days.
      Note: The level of the medium should be checked after the medium exchange, as it is important for cell survival.

2. Electroporation

All procedures should be performed under sterile conditions.

Note: Perform electroporation one day after the preparation of the thalamic blocks to prevent detachment of the blocks.

1. Plasmids
   1. Make a mixture of plasmids as follows: pCAGGS-DsRed 2 μg/μL; pCAG-SYP-EGFP 3.5 μg/μL.
      Note: Purify the both plasmids using an endotoxin-free Maxiprep kit and dissolve in Hanks’ balanced salt solution.
2. Electroporation equipment setup
   The electrical instruments such as a stimulator and an isolator should be connected as shown in Figure 2.
   1. Connect the stimulator to the biphasic isolator.
   2. Connect an electrode (a silver wire 0.2 mm diameter) to the negative terminal of the isolator.
   3. Connect an electrode (a silver wire 1 mm diameter) in series to a 100 Ω resistor and the positive terminal of the isolator.
   4. Connect the amplifier to the oscilloscope to monitor whether electrical currents are passed across the resistor by measuring the voltage with the amplifier.
3. Preparation of glass pipettes
   Note: Two types of glass pipettes are used for ejection of the plasmid solution and application of electrical pulses.
   1. Make the pipettes from glass capillaries with an electrode puller.
   2. Break the tip for the ejection pipette (the tip diameter should be 20 – 50 μm).
   3. Grind and polish the tip of the electrode pipette with sandpaper followed by fire-polishing (the inner diameter should be 50 – 200 μm).
      Note: The electrode tip should be flat and smooth, as it directly touches the tissue.
4. Procedure of electroporation
   1. Connect the ejection and electrode pipettes to the manipulators.
   2. Insert a silver wire (0.2 mm diameter) into the electrode pipette.
   3. Attach the ejection pipette to a 1 mL syringe with a plastic tube.
   4. Take up >5 μL of plasmid solution into the ejection pipette by syringe suction.
   5. Put the coculture on the electroporation stage, and insert the 1 mm diameter electrode into the culture medium.
   6. Place the ejection pipette on the thalamic explant.
   7. Push the syringe manually after the tip touches the surface of the thalamic block.
      Note: About 0.5 μL of the plasmid solution is usually used per one thalamic block.
   8. Place the electrode pipette on the thalamic block immediately after retraction of the ejection pipette.
   9. Deliver electrical pulses (50 – 400 μA, 3 to 5 trains of 200 square pulses of 1 ms duration at 200 Hz). Monitor the amplitude and the number of trains on the oscilloscope.
      Note: The amplitude of electrical currents depends on the inner diameter of an electrode pipette¹³.
   10. Retract the electrode pipette and return the dish to the incubator.
5. Microscopic observation
   1. Put the culture dish on a microscopic stage of an upright confocal microscope equipped with two filter sets for EGFP (excitation, 488 nm; emission, 515 nm) and DsRed (excitation, 514 nm; emission, 565 nm).
   2. Take images of 10 – 25 optical sections at 1 – 7 µm-step sizes with a 20X long-working distance objective lens. Select individually distinguishable labeled axons which express both DsRed and SYP-EGFP.
      Note: The observed area is approximately 0.7 x 0.7 mm (corresponding to 1024 x 1024 pixels, that is, 0.68 μm/pixel).
   3. Capture images every 24 h for daily imaging. Complete within 10 min at room temperature, and return the cultures to the incubator. For short-interval time lapse imaging, keep the culture in a culture chamber, which maintains the environment of humidified 95% air and 5% CO₂ at 37 °C.