Recording Light-evoked Postsynaptic Responses in Neurons in Dark-adapted, Mouse Retinal Slice Preparations Using Patch Clamp Techniques

The overall goal of this experiment is to obtain light response recordings from dark adapted retinal neurons. This is accomplished by first harvesting a mouse eye. The second step is to isolate the retina and cut it for slice preparations.

Next, the retinal preparation is mounted and viewed on a microscope to find a target cell for patch clamp recordings. After establishing a whole cell patch clamp configuration, a light stimulus is applied in the vicinity of the cell. Finally, as simultaneous electrophysiological response can be observed through acquisition software.

The main advantage of this technique of existing methods like KRC imaging, is that the recording sensitivity is high, and that recording cells can be morphologically identified. Generally, individuals new to this method will struggle because working in total darkness can be difficult and disorienting for those unaccustomed to doing so. To begin a range, the dissecting tools and transfer pipettes near the dissecting microscope.

The work area will be well organized so that each tool can be easily accessed in the dark. When ready, Dawn an infrared viewer to conduct the procedure in the dark. After euthanasia quickly in nucleate the eyes and place them in a dish of cold dissecting solution, kept in a dark box working under a microscope.

Transfer an eye into a dish with cool dissecting solution. Continuously bubbled with oxygen. To remove the cornea and lens, hold the optic nerve at the other end of the eyeball to keep it from moving and make an incision on the top of the cornea with a microsurgical knife.

Extend the cut using a pair of surgical scissors and remove the cornea, leaving some sclera surrounding it. Next, grab the lens with fine forceps and slowly pull it out. Once the eye cup is made, gently pour cold oxygenated dissecting solution into the eye cup with a small transfer pipette.

Work quickly to maintain the shape of the retina and use gentle pressure to identify the dorsal and ventral sides of the retina. Use infrared viewers to identify the line going across the retinal eye cup, passing near the optic nerve head. This line passes mostly across the ventral side of the optic nerve and the side, including the optic nerve, is the dorsal side.

Depending on the experiment, make a cut on either the ventral or dorsal side. This marks the unnecessary side and is a useful marker moving forward. Next, gently remove the vitreous with extra fine forceps without touching the retina.

The vitreous is mainly attached to the bottom and the outer edge of the eye cup. As the vitreous might be attached tightly to the eye cup, proceed gently to remove retinal detachment. Continue to pull until no tension is felt.

Once the retina is isolated, grab the sclera and gently peel off the retina. Using the backside of the forceps, trim the retina and discard the unnecessary half in the same plastic dish. Cut the remaining retina into two pieces for each piece.

Cut off the edges to make a retinal slab and trim the folding edges. Next, use a large transfer pipette to transfer a retinal slab onto a glass plate. Suck up the excess solution with a small pipette and use a piece of filter paper to flatten the slab.

Gently place a piece of the filter membrane on top of the tissue using a small transfer pipette. Place a drop of dissecting solution on the filter membrane. Wait about 15 seconds while the solution spreads and sticks the retinal tissue to the filter membrane.

Once stuck together, pour more cold dissecting solution under and around the filter until the membrane floats. Next, place the filter membrane into the slicing chamber and cover with dissecting solution. Working quickly to ensure that the membrane does not dry out, use the chopper to cut the retinal tissue into slices.

Place a plastic cover slip with grease rails into the slicing chamber. Transfer a retinal slice on top of the grease rails and rotate the slice 90 degrees so that the transverse section is visible. Press the filter membrane down onto the cover slip before covering the sides of the filter paper with grease.

After the slices immobilized on a plastic cover slip, transfer it to a 33 millimeter plastic dish. Submerge the preparation with coal dissecting solution and store each dish in a dark box that is continuously oxygenated. For patch clamp recordings.

Prime the profusion tubes with aims medium and allow all the bubbles to pass from the tubing. After making recording pipettes with a glass polar, backfill the tip with pipette solution. Next, use a micro pipette filler to fill about one third of the pipette once filled, store each pipette in a moist pipette box.

Next, turn on the equipment for patch plant recordings, including the computer amplifier CCD camera and microscope working in dark conditions. Place a retinal slice preparation onto the microscope stage with the aid of an infrared viewer. After it is immobilized, begin continuous perfusion.

Set the perfusion temperature to 33 to 37 degrees Celsius. View the slice surface with a CCD camera. Focus on the target location.

With the target cell types reside, select a healthy looking cell for a patch clat recording. Once a healthy cell is identified, place a recording pipette in a pipette holder and advance the pipette to the slice preparation. When it is close to the slice preparation, find the tip of the pipette with a microscope.

Once the pipette tip is visible under the microscope, move the tip down towards the target cell. Next, set the amplifier and adjust the pipette at pika amp. Five volts per Nana amp.

Start a continuous electric pulse of approximately five millivolts and approximately 10 hertz and check the pipette resistance. An ideal resistance is between five and 12 mega ohms. For most retinal neurons.

When ready, use a mouthpiece or syringe to blow out the internal solution until the tip of the pipette is on the surface of the target cell. When the positive pressure makes a small dimple on the cell surface, advance the tip slightly and stop blowing out. If the pipette resistance is continuously increasing, leave it and monitor the resistance until it reaches greater than one giga ohm.

If the resistance does not spontaneously increase, gently apply negative pressure until it becomes a giga seal. After the giga seal is achieved, change the holding potential to negative 70 millivolts 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 mega ohms and one giga ohm, and the capacitive current is observable.

Record the IV relationship from negative 80 to 40 millivolts. Different types of voltage gated channels will be activated depending on the cell type. Lastly, record light evoked synaptic currents or voltages.

This slice preparation viewed at 10 x shows both photoreceptors and ganglion cells with no detachment from the filter paper. When this preparation is viewed with a 60 x objective, each cell layer is clearly observed. Light evoked excitatory postsynaptic potentials from retinal neurons were evoked in response to a step light.

The peak amplitude and decay time vary depending on the cell type and subtype. The cell type and its morphological subtype were revealed by sulfur labeling after physiological recordings After its development. This technique paved the way for scientists in the field of retina research to study the functions of natural visual signaling, as well as the contributions of synaptic mechanisms.