The overall goal of this technique is to maintain neuronal tissue for prolonged periods without significant tissue degradation. Using this method, electrophysiological recordings and calcium imaging can be conducted more than 24 hours after tissue extraction. This method will greatly increase the viability of acutely extracted neuronal tissue and maintain its viability for greater than 24 hours setting a new gold standard for acute tissue incubation.
The main advantage of this technique is that the experimental time the tissue can be utilized is significantly increased. This reduces the number of animals needed and maximizes the amount of data that can be obtained from each experiment. The Braincubator is a automated system that maintains the tissue temperature and prevents bacterial proliferation in the extracellular fluid.
This provides a tightly-controlled environment for the tissue. Demonstrating this procedure will be Orsolya Kekesi, and Alba Bellot-Saez, PhD students from my lab. For brain slice preparation prepare 300-micrometer thick brain slices that contain the region of interest with a vibratome.
Then transfer the slices to a custom-built incubation system with carbogen flow, pH levels, and temperature controlled. Set the initial chamber temperature to 35 degrees Celsius for 15 to 30 minutes, and then slowly reduce it to 15 degrees Celsius. After that incubate the slices in the incubation system until electrophysiology recording or imaging.
For retinal whole mount preparation immediately enucleate the eyes from a euthanized animal. Make a small cut along the ora serrata and place the eyes and either Ames'media or aCSF with carbogen supply at room temperature. After that immediately remove the cornea, lens and vitreous by cutting along the ora serrata with small scissors and remove them with forceps.
Place the tissue in the incubation system at room temperature. To remove the inner limiting membrane transfer the eye cup containing the retina to a small glass jar containing papain, L-cysteine, EDTA and DNase at 37 degrees Celsius for 20 minutes. Apply carbogen to the solution through the lid, but do not bubble.
Stop the enzymatic digestion by placing the tissue in an ovomuccoid and BSA solution for 10 minutes in Earle's BSS. Then wash the tissue thoroughly with aCSF. Subsequently, transfer the tissue to the incubation system and reduce the temperature to about 15 to 16 degrees Celsius.
Following that, transfer the retinal tissue to the microscope. Isolate the retina from eye cup and cut it into four pieces with a razor blade. If the entire retina is required, make four small cuts in the periphery of the retina to allow it to lie flat.
This tissue can now be transferred to the microscope for electrophysiology or maintained in the Braincubator. The tissue is maintained in the main chamber containing the probes for pH and temperature measurements. A second chamber is isolated from the main chamber and is exposed to 1.1 watts UVC light in order to irradiate bacteria floating in the solution.
Use a peristaltic pump to circulate aCSF through the two chambers, and a Peltier thermoelectric cold plate to either cool or heat the main chamber. In this procedure, dissolve calcium dye in DMSO to make a one millimolar solution. Add 1%pluronic acid F-127 to the mixture to achieve a final volume of 50 microliters.
Then, sonicate it for 10 minutes. Prepare a glass-loading chamber with calcium dye diluted in 2.5 milliliters of aCSF to achieve a final concentration of 10 micromolar for brain slices, and 20 micromolar for retina. For retina and brain slices from young animals, incubate the sample in the bath for 45 minutes.
For adult animals, pipette 25 microliters of dye directly onto the brain slice and maintain it for 75 minutes to allow better penetration of the dye into the deep layers. To ensure adequate oxygenation of the submerged tissue during dye incubation, oxygenate it continuously with carbogen through the lid, but do not bubble. Following dye loading, wash the tissue with aCSF and transfer it to the incubation system.
Slowly reduce the temperature to about 15 to 16 degrees Celsius until use. For recording, place tissue in a submerged recording chamber under a microscope, and perfuse it with oxygenated aCSF at a flow rate of four to five milliliters per minute. Hold the tissue in by using a custom-made harp.
Next, prepare some recording pipettes using a micropipette puller to achieve a final resistance of five to six megaohms. Fill a pipette with three to four microliters of internal solution, and then place the solution on ice. Then visualize the cells using a CCD camera under IR-DIC.
Position the pipette on the cell membrane using a micromanipulator. Maintain positive pressure through a suction port on the pipette holder. Once the pipette is on the cell, apply gentle negative pressure to the pipette to achieve a gigaohm seal.
Then rupture the cell membrane with brief negative pressure. Subsequently, start whole cell current or voltage clamp recording. For ratiometric imaging of Fura-2 use an ultra high-speed wavelength switcher to provide excitation wavelengths of 340 nanometers and 380 nanometers.
Then capture the emitted light through an omission filter with a high sensitivity, high-speed digital camera. For a single excitation wavelength of Fluo-4, filter the excitation light through 460 to 490 nanometer bandpass filter, and emitted light through a 515 to 550 nanometer bandpass filter. After digestion, retinal ganglion cells and displaced amacrine cells can be clearly visualized under DIC illumination, and can be targeted for patch clamp recordings without any prior scraping of the inner limiting membrane.
Representative current clamp recordings from a retinal ganglion cell are shown here. Depolarization of the cell via the patch pipette caused dose-dependent action potential generation indicating the viability of the cells following papain treatment. Removal of the inner limiting membrane also allowed ubiquitous staining of the ganglion cell layer with Fura-2 AM.And the cells responded to 30 millimolar potassium chloride application with a large increase in intracellular calcium concentration as evidenced by an increase in 342 to 380 nanometer ratio relative to F0.After potassium chloride application, the calcium levels returned to baseline following stimulation.
And the cells could be subsequently stimulated to produce a similar amplitude response. Moreover, these responses were indistinguishable between retina recorded at shorter than four hours and longer than 24 hours post dissection. While attempting this procedure, it's important to remember that acute neuronal tissue is environmentally defenseless.
Therefore, tight regulation of aCSF through close monitoring of pH, temperature and bacterial levels is essential for maintaining the cellular activity and network integrity. This procedure can be used to maintain neuronal tissue for more than 24 hours without the need for aseptic techniques or the use of culture media containing growth factors or antibiotics that may affect physiology. We hope this method will set the gold standard for the ideal parameters to maximize tissue viability.
This reduces the variability in the health of the tissue itself and subsequently reduces experimental variability. Using the papain protocol to digest the inner limiting membrane of the retina makes it possible to achieve ubiquitous staining with calcium dyes. This protocol can also be used to obtain multisite intracellular recordings from neurons of the ganglion cell layer.
After watching this video, you should have a good understanding of how to maintain viable tissue for over 24 hours for imaging and electrophysiological experiments. Don't forget that working with ultraviolet light can be hazardous. Precautions such as protective eye wear should always be worn while performing this procedure.
