Light-Evoked Postsynaptic Responses in Neurons of Mouse Retinal Slices

All procedures involving sample collection have been performed in accordance with the institute's IRB guidelines.

1. Preparation of Experimental Solution

1. Prepare the dissecting solution 1 day to 1 week before the actual experiment. Use a 4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid (HEPES) buffer solution for retinal dissection because of its strong buffering ability at lower temperatures16. Mix all chemicals as follows (in mM): 115 NaCl, 2.5 KCl, 2.5 CaCl₂, 1.0 MgCl₂, 10 HEPES, 28 glucose. Adjust pH to 7.4 with NaOH. Keep the solution in a refrigerator up to 1 week.
2. The recording solution is Ames' medium, which is an artificial cerebral spinal fluid (aCSF) designed for retinal preparations. By the day of the experiment, weigh out Ames' powder (4.4 g for 500 ml solution) in a tube. Also, weigh out NaHCO₃ (0.95 g for 500 ml solution) and mix with the Ames' powder.
3. Prepare the intracellular pipette solution for patch clamp recordings by the day of the experiment. Mix all chemicals as follows (in mM): 111 K-gluconate, 1.0 CaCl₂, 10 HEPES, 1.1 ethylene glycol tetraacetic acid (EGTA), 10 NaCl, 1.0 MgCl₂, 5 ATP-Mg, and 1.0 GTP-Na. Adjust pH to 7.2 with KOH. Filter the pipette solution with a conventional syringe filter. Make ~500 µl aliquots and store them in a freezer or a deep freezer.

2. Preparation for the Day of Experiment

1. The night before the experiment, place a mouse in a cage (C57BL/6J or similar background, 4 – 8 weeks old, male) in a dark space for overnight (O/N) dark adaptation.
2. Preparation for dissection:
   1. Fill dissecting solution (100 – 200 ml) in a glass beaker, place it on ice, and bubble with oxygen for at least 10 min.
   2. Start oxygenation of the dark box for retinal preparation storage.
   3. Prepare plastic coverslips with grease rails for retinal slice preparations and place each of them in a 35 mm plastic dish.
   4. Attach a new razor blade to a chopper (tissue slicer).
   5. Cut a filter membrane into two halves. This is for retinal slicing.
   6. Align the dissecting tools and transfer pipettes near the dissecting microscope. The work area must be well organized so that each tool can be easily accessed in the dark.
3. Preparation for patch clamp recordings:
   1. Dissolve Ames' powder and NaHCO₃ in distilled water (Ames 4.4 g and NaHCO₃ 0.95g/500 ml ddH₂O). Bubble the Ames' medium with mixed oxygen (95% O₂ and 5% CO₂) for at least 30 min while heating to a recording temperature level (~35 °C). Adjust the pH to 7.4 with NaHCO₃ while still bubbling.
   2. Thaw the internal pipette solution during the dissection. After it is thawed, add 0.01% sulforhodamine B for intracellular staining.

3. Patch Clamp Recordings from Retinal Slice Preparation

1. Recording preparation:
   1. Prime the perfusion tubes with Ames' medium. Allow all the bubbles to pass through when filling the tubing.
   2. Make patch clamp recording pipettes with a puller. To fill a pipette with the pipette solution, dip in the backside of a pipette for 1 min until the pipette tip is backfilled. Next, fill ~1/3 of the pipette using a micro-pipette filler. Store each pipette in a moist pipette box.
   3. Turn on the equipment for patch clamp recordings, including the computer, amplifier, charge-coupled device (CCD) camera, and microscope.
2. Shut off the room light. Place a retinal slice preparation onto the microscope stage chamber using an infrared viewer. After it is immobilized, begin continuous perfusion. Set the perfusion temperature at 33 to 37 °C.
3. View the slice surface with the CCD camera. Focus on the target location where the target cell types reside. Select a healthy-looking cell for patch clamp recording (i.e., it should have smooth-looking surface and a good cell shape).
4. Place a recording pipette in a pipette holder. Advance the pipette to the slice preparation. When it is close to the slice preparation (~2 mm above), find the tip of the pipette with the microscope. Once the pipette tip is visible under the microscope, move the tip down towards the target cell.
5. Set the amplifier. Adjust the pipette at 0 mV/0 pA (zeroing) and start a continuous electric pulse of ~5 mV at ~10 Hz. Check the pipette resistance; ideal resistance is between 5 and 10 MΩ for most retinal neurons.
6. Start blowing out from the tip. Use a mouthpiece, or use a syringe, to apply positive pressure when the pipette dips into the bath solution. Continuously blow out the internal solution until the tip is on the surface of the target cell.
   1. When the positive pressure makes a small dimple on the cell surface, advance the tip slightly and stop blowing out. Check the pipette resistance, which should increase. If it is continuously increasing, leave it and monitor the resistance until it reaches >1 GΩ (gigaseal). If the resistance does not spontaneously increase, gently apply negative pressure until it becomes a gigaseal.
7. After the gigaseal is achieved, change the holding potential to -70 mV. Then, intermittently apply negative pressure to rupture the membrane inside the pipette tip. When the whole-cell configuration is made, the pipette resistance can be between 500 MΩ and 1 GΩ, and the capacitive current is seen. Sometimes spontaneous postsynaptic currents can be observed.
8. Record the current–voltage (I-V) relationship from -80 to +40 mV. Different types of voltage-gated channels are activated depending on the cell type.
9. Record light-evoked synaptic currents or voltages.