The overall goal of this procedure is to isolate fluorescently labeled single cells from live whole mount retinas. This method can help answer key questions in neuroscience. Such as how many subtypes of a particular neuronal class exist and what are the genetic markers for each subtype?
The main advantage of this technique is that we forgo the need for retinal tissue dissociation, allowing the cells to survive in a healthier environment prior to isolation. The implications of this technique extend to any fluorescently labeled cell population and can thus be applied to other systems to understand cellular diversity and function in health and disease. Generally, individuals new to this method will struggle, because this technique relies on the ability of the researcher to patch-clamp a cell without compromising the viability of the cell.
Future demonstration of this method is critical as RNA isolation steps may be difficult to learn, because the small amount of RNA may easily degrade if not handled properly and quickly. To begin this procedure, poke the cornea with a needle and cut it away at the border of the cornea and sclera. Then, remove the lens using forceps.
Gently make a tear in the sclera and sever the optic nerve where the retina and sclera meet. Carefully finish removing the sclera from the retina. Next, remove the transparent vitreous.
Slice the retinas in half and store them in the oxygenated extracellular solution at room temperature until use. When ready to mount the tissue in the recording chamber, transfer a piece of retina to the enzyme solution, diluted in 500 microliters of oxygenated extracellular solution in a petri dish, and incubate it for two minutes at room temperature on a shaker. Subsequently, wash the retina in the oxygenated extracellular solution and transfer it to a glass bottom recording chamber with a plastic transfer pipette.
After that, use forceps to carefully flatten the tissue with the photoreceptor layer facing down. Remove excess fluid using a pipette and anchor the tissue using a platinum ring with nylon mesh. Then, fill the chamber with the oxygenated extracellular solution and mount it on to a microscope stage.
Perfuse the tissue with the oxygenated extracellular solution at two to four milliliters per minute. To prepare for this procedure, pulse some glass micropipettes for electrophysiological recordings using a micropipette puller. Observe the ganglion cell layer using IR-DIC optics.
Then, identify the GFP plus ganglion cells using epifluorescence at about 480 nanometers. Next, locate the pipette filled with intracellular solution in DIC. Apply slight positive pressure and zero any voltage offsets on the amplifier.
Subsequently, lower the glass micropipette on a GFP positive cell and apply test voltage command steps to monitor the seal resistance. The negative pressure should form a gigaohm seal between the pipette and the cell membrane. After forming a stable seal, rupture the membrane by applying brief pulses of negative pressure to gain whole cell access.
Wait one to two minutes for the dendrites of the cell to fill with fluorescent tracer. In this step, carefully extract the cell cytoplasmic content into the pipette by applying negative pressure with a ten milliliter syringe. In the meantime, monitor the extraction in DIC by visualizing the cell body decreasing in size.
After extracting the cytoplasmic contents, lift the pipette carefully off from the tissue and quickly remove the pipette from the solution. Next, quickly remove the pipette from the headstage holder and rinse the pipette tip briefly with DEPC-treated H2O. Then, connect the pipette to a one milliliter syringe via the tight-fitting tubing.
Immediately expel the cells into ten microliters of lysis buffer one in the 0.2 milliliter PCR tubes. Briefly centrifuge the tubes in a table-top mini centrifuge at 2000 times g for ten seconds. Following that immediately flash freeze the samples on dry ice for five minutes.
After freezing, store them at negative 80 degrees Celsius for up to two weeks for best results. To prepare for this procedure, set up a magnetic separator device by taping the top part of an inverted P20 or P200 tip holder to the 96 well magnetic stand. Then, prepare fresh 70%ethanol at approximately one milliliter per sample.
Remove the RNA magnetic beads from 4 degree C storage and thaw them at room temperature. Once the RNA beads are at room temperature, vortex them for 30 seconds to ensure that the solution is well mixed. After that, thaw the cells at room temperature for one minute.
Then, add five microliters of Rnase-free H2O to each sample and pipette up and down. Afterward, add 22 microliters of RNA beads to each tube and mix thoroughly. Incubate the samples at room temperature for five minutes to allow the RNA to interact and bind with the magnetic beads.
Following that, place the tubes on a magnetic separator device for eight minutes. Before proceeding, ensure that the supernatant is clear. Observe the beads from one palette and be sure not to detach it from the side of tube during pipetting.
Remove the supernatant from the samples and add 150 microliters of 70%ethanol. Then, remove the ethanol and repeat the wash two more times. Allow the samples to air dry for six minutes.
Check intermittently to see if more ethanol has been collected at the bottom of the tube and remove it accordingly. Remember it is critical to dry the beads appropriately. If the beads are not dry enough, ethanol may be carried through with the eluent and will decrease the total yields, while over dried beads can result in the loss of RNA.
While the samples are drying, prepare 10x reaction buffer by combining 19 microliters of lysis buffer two and one microliter of Rnase inhibitor. Briefly spin it down and keep it on ice. Once the samples are dry and the bead pellets no longer appear glossy, remove the tubes from the magnetic separator and add 9.5 microliters of Rnase-free H2O to rehydrate the samples.
Then, place the samples on ice and add one microliter of 10x reaction buffer to each sample. This image shows the ganglion cell layer of the retina, visualized using IR-DIC in the whole mount retina preparation. The same preparation was visualized in epifluorescence at about 480 nanometers to identify the GFP positive ganglion cells.
This image shows a GFP positive cell that was targeted for patch-clamp recording and filled with a fluorescent tracer. Confocal image of ipRGC dendrites imaged in the on and off sublamina of the inner plexiform layer allows for the classification of these cell types This bioanalyzer output shows the examples of libraries that have undergone reverse transcription, amplification, and purification. Lanes one to three show ideal DNA smears, while lane four represents a poorly processed sample.
The control lane should contain two clean peaks at the marker sizes of 35 bp and 10380 bp. Here is a detailed trace of one successfully prepared library with high-intensity around 2 kb. This trace also demonstrates a successful preparation, but with the smear centered around 500 bp.
This image shows the bioanalyzer output following tagmentation, amplification and purification of the samples. The detailed trace of a successfully tagmented sample can be seen here, corresponding to the sample in lane one. Incomplete tagmentation will result in a trace such as the one seen here, warranting retagmentation with a fresh dilution.
After watching this video you should have a good understanding of how to isolate RNA from the fluorescently labeled cell types in the intact retina. While attempting this procedure, it's important to remember that RNA is very sensitive to degradation and that having healthy retinal cells is the key. Once mastered, this technique can be done in two days if it is performed properly.
Following this procedure, other methods like qPCR, NC2 hybridization, or immunohistochemistry, can be performed in order to answer additional questions, like whether identified genes are specific to a given cellular subtype. This technique has paved the way for researchers in the field of neuroscience, trying to understand heterogeneity of neurons in various brain regions. Understanding this heterogeneity will allow for future studies of cellular function and connectivity in molecularly identified populations.
