Our visual experience begins when the visual pigment in our retinal photoreceptors absorbs photons of light energy, ultimately leading to the closure of cyclic nucleotide gated channels in the cell membrane. This suppresses the release of the neurotransmitter glutamate from both rods and cones. The released glutamate is sensed by downstream retinal cells to study the downstream signals.
The eyes are removed from a euthanized mouse. The S are isolated, embedded in an agar block and sliced using a vibrating microtome. Next, the retinal slices are placed under a microscope and stimulated with LED lighting.
While patch clamp recordings are taken to measure the synaptic currents or potentials generated. Generally individuals new to this method will struggle because of the added complexities of working in the dark. So being able to visualize the entire process will provide a feel that's otherwise difficult to describe.
Begin this procedure by coating the recording and reference wire electrodes that will be used with a layer of silver chloride. Dip the electrodes in one molar sodium chloride. Then using a 15 volt power source with a one hertz sine wave drive 15 volts between them for about 20 seconds.
Using a micro pipette puller, prepare patch pipettes from fire polished BO silica glass, choosing a pipette tip resistance based on the size of the target cells for cell bodies about 20 microns in diameter. Use a pipe PET with a tip resistance of five to 10 mega ohms for cell bodies. After about five microns in diameter, use a tip resistance of 12 to 15 mega ohms.
To prepare the ground reference electrodes, use a syringe to fill polyethylene tubing with a 3.5%agar solution prepared in one millimolar sodium chloride. Next, prepare the following solutions according to the instructions in the accompanying written protocol. External bath solution slicing solution, potassium aspartate pipette internal solution, and the agar backstop.
Store all solutions at four degrees Celsius until they are needed. On the day of the experiment, Thor around 1.5 milliliters of potassium aspartate internal solution. Then add diluted Fluor.
Pour about 50 milliliters of aims media containing sodium bicarbonate into the light tight storage container. Connect the plastic tubing to a gas tank containing 5%carbon dioxide and 95%oxygen to equilibrate the aims media with the gas, and then place the container into a 30 to 32 degrees Celsius water bath. Next, fill a bottle with 110 milliliters of aims, medium containing he peas and place it on ice.
Equate the medium by bubbling the solution with 100%oxygen gas. Place the remaining aimes medium containing sodium bicarbonate into a heated reservoir on top of the Faraday cage. Wrap paraform around the tubing and bottle top to trap gas in the space above the solution and equilibrate with 5%carbon dioxide, 95%oxygen gas.
Using a glass air dispersion tube, prepare 3%low gelling temperature aros by adding 0.75 grams to 25 milliliters of aims medium with heaps heat. The solution is about 115 degrees Celsius while stirring. When the solution becomes clear without bubbles, place the melted agar solution into a 42 degree Celsius water bath.
Set up the recording chamber on the inverted microscope. Slide a five centimeter agar bridge over the silver, silver chloride reference electrode and position it in the recording chamber. The pipette tip resistance can be measured by filling a pipette with internal solution, placing it on the pipette holder, putting the pipette holder in the amplifier head stage, and lowering the pipette into solution to preserve the visual pigment for physiological recordings, all procedures should be performed under infrared light using infrared goggles, but all procedures will be shown here in room light.
After the mouse has been euthanized, use curved scissors to quickly remove the eyes. Place the eyes on a piece of filter paper to prevent them from rolling. Then hold an eye in place with forceps and use a thin double-sided blade to make a slit in the cornea.
As soon as the eyeball is punctured. Transfer the eye into a 60 millimeter Petri dish filled with aims medium. Using the slit in the cornea as an entry point, use small scissors to cut away the remaining portions of the cornea.
Then remove the lens and vitreous humor, ensuring that the retina remains attached to the eye cup. Place it in a light tight container filled with aims. Medium equilibrated with 5%carbon dioxide in 95%oxygen at 32 degrees celsius.
Next hemis sec. One of the eye cups using a number 10 scalpel blade. Separate the retina from the retinal pigment epithelium.
Remove the edges of the retina to allow the tissue to life flat. Use a small glass pasta pipettes with a latex bulb to fill a 35 millimeter Petri dish with melted low geling temperature agar and drag the forceps through the agar to test its consistency. Once the agar begins to solidify, transfer the retina to the top of the agar.
Then without touching the retina, use a twisted piece of Kim wipe to absorb the excess solution around it. Place an additional two to three drops of agar directly on the retina and quickly place a plastic cylinder to form a wall around it while maintaining the retina near the center of the cylinder drip in additional melted agar, transfer the Petri dish to an ice water bath to cool. After 30 seconds, remove the tissue and use a plunger to extrude the agar block, pushing from the side farthest from the retina.
Use a one-sided razor blade to cut out a small block of agar containing the retina and super glue the block into place against the agar backstop on the specimen disc. Transfer the specimen disc to the vibrating microtome tree and pour ice cold aims heaps into the tree, allowing the block with the retina to be fully submerged. Cut retinal slices with a thickness of about 200 microns using the maximum blade vibrating rate with a forward blade speed set such that it takes four to five seconds to cut through the retina.Once.
Use a number 11 scalpel blade to cut each usable slice from the block one at a time. Place the slices containing the retina into recording chambers and hold them down. Using slice anchors the nylon threads crossing the slice anchor.
Hold down the agar surrounding the retina, leaving the retina unobstructed. For recordings. Transfer the recording chamber to the stage of the upright microscope.
Close the curtain and turn on the infrared light source to view the slice on an infrared sensitive camera. Under the microscope, identify a retinal slice with intact photoreceptors at the appropriate cutting angle. Some surface damage may be apparent due to the slicing process.
Using a pipette with a broken tip gently suck away the dead cell bodies To remove this thin layer of cells, once a healthy layer of live cells has been revealed, identify a cell body whose position is located in the strata of interest. Move the pipette tip such that it dimples the cell membrane and apply gentle, negative, or inward pressure to make a high resistant seal. Whole cell mode can be achieved by applying negative pressure to the electrode to break into the cell.
Next, stimulate the retinal tissue with light using LEDs to evoke responses. At the end of the recording, capture an image using the appropriate excitation light for the Fluor that dialyzed into the cell from the pipette internal solution. Here, light evoked potentials of a dark adapted retinal neuron to flashes of light of increasing strength are shown.
This figure is a fluorescence picture taken from a retinal slice where the cells have been filled with the Fluor Lucifer yellow evoked fluorescence shows the morphology of cells from which patch recordings were made. After watching this video, you should have a better understanding of some of the complexities of making patch clamp recordings from dark adapted retinal slices. The goal of this procedure is to measure light evoked responses under controlled illumination.
In order to do this, we have to first isolate the retina, embed it in auger, and then prepare retinal slices from which we can make single cell recordings.
