The overall goal of the following experiment is to quantify the replenishment and mobilization of the specific synaptic vesicle pools in central nerve terminals using live fluorescence imaging techniques. This is achieved by monitoring two rounds of synaptic vesicle recycling, using FM dyes in the same nerve terminals to provide valid internal controls. As a second step, the readily releasable pool and the reserve pool are sequentially depleted using specific unloading protocols, which allow the quantification of relative pool sizes.
Next, change either loading or unloading conditions in order to study the effect of the chosen variation on either synaptic vesicle, pool replenishment, or mobilization. Results show that FM d loading of different synaptic physical pools is reproducible based on the quantification and analysis of the evoked dye unloading. The main advantage of this technique over existing methods, like the fluent family of genetic reporters, is that this technique tracks the replenishment of pools specifically by synaptic vesicles, which have undergone endocytosis.
It also allows accurate quantification. Since this protocol is performed twice in the same nerve terminals providing an internal control. This method can help answer key questions in the synaptic, CLE, endocytosis and exocytosis field, such as how manipulation in specific endocytosis modes can affect the replenishment of both the total recycling pool and the synaptic CLE pools within it, such as the readily reusable pool and the reserve pool.
Basic experimental setup should consist of an inverted epi fluorescence microscope, a cooled CCD camera, a fluorescent light source, a gravity perfusion apparatus, an imaging chamber with parallel platinum electrodes and electrical stimulator and a computer. Perform the experiments in the dark or under red light conditions with minimum fluorescent illumination of the sample to avoid FM dye bleaching. Begin this procedure by using seven day old spra dolly rat pups to prepare the primary cultures of cerebellar granule neurons.
After the cultures have been prepared, plate the neurons in the center of a poly D lysine coated 25 millimeter cover slip in a droplet, not exceeding seven millimeters in diameter after one hour, add two milliliters of culture media to the neurons after seven days. Place a single cover slip in the saline solution at room temperature for 10 minutes to allow stabilization in the new medium. After that, remove the cover slip, dry its underside, and the area surrounding the attached cells with a small piece of paper towel or absorbent paper.
Subsequently, use the silicone grease to glue the cover slip to the bottom of the imaging chamber. Cells should be between the two parallel wires. Use sufficient silicone grease to completely seal the chamber without encroaching into the center of the bath chamber.
Next, gently fill the bath chamber with about 260 microliters saline solution. Then fill the input and output tubing with the same solution. Seal the chamber by gluing a clean cover slip with silicone grease to the top.
After that, remove any air bubbles trapped in the chamber with the input and output tubing. Then immobilize the imaging chamber in a stainless steel platform. Check for leaks by gently profusing saline solution through the input tubing.
The next step is to mount the assembled chamber on the stage of an inverted microscope. Prime the inlet of the gravity perfusion system with saline solution and connect the chamber to the inlet. Then attach the connecting wires of the chamber to the electrical stimulator.
Perfuse the neurons with 1.5 milliliters of diluted fm dye. Subsequently stimulate the neurons using the attached stimulator to evoke dye uptake After stimulation, perfuse the neurons with fresh saline solution for two minutes to wash away excess FM dye. Then keep the neurons in the saline solution for eight minutes.
In the meantime, locate the axonal networks where the individual FM D loaded nerve terminals are visible. Using fluorescein wavelengths. Avoid the areas with clusters of cells.
Then refocus image immediately before image acquisition since a slight drift may have occurred during the rest period. Begin time-lapse image acquisition at the rate of one frame every four seconds. After acquiring five to 10 baseline images evoke exocytosis of the readily releasable pool by delivering a 30 hertz stimulation for two seconds.
After acquiring another 10 images, evoke synaptic vesicle, exocytosis of the reserve pool using three stimulations of 40 hertz for 10 seconds with 40 seconds interval between stimulations, acquire another five to 10 images. Then pause the image acquisition. Allow the neurons to recover for at least 20 minutes, and repeat this S one phase protocol again in S two phase for the control experiment.
For data analysis, convert the images to a stack. Using an image J built-in function, then adjust the brightness and contrast of the stack to maximize the dynamic range if significant horizontal drift has occurred. During the experiment, run stack reg and turbo reg plugins on image J to align image stack.
Then runtime series analyzer plugin. Next, define the regions of interest over at least 90 nerve terminals. It is helpful to toggle between the images before and after die unloading to reveal active nerve terminals.
An ideal region of interest size is one that is slightly bigger than a typical nerve terminal. Then obtain the total fluorescence intensity of each region of interest over time. After that, export to Microsoft Excel.
Shown here is a flow chart of a control experiment where cerebellar granule neurons were loaded with 10 micromolar FM 1 43 using 80 hertz 10 seconds stimulation. Here is an image showing nerve terminals loaded with FM 1 43. Here is the same image showing 90 numbered regions of interest selected for analysis after identifying active nerve terminals, and these are the images of the nerve terminals at selected time points.
These images are presented in pseudo color to illustrate the changes in fluorescence as the different synaptic vesicle pools were depleted. Finally, here shown the normalized fluorescent units of these 90 nerve terminals. In S one and S two phases, the fluorescence drop of each sequential stimulation can be measured and compared between S one and S two phases.
In this control experiment, both S one and S two resulted in the same sizes of the readily releasable and the reserve pools. After watching this video, you should have a good understanding of how to quantify the replenishment and mobilization of specific synaptic vesicle pools in the central nerve terminals using life fluorescence imaging techniques. While attempting this procedure, it is important to remember to confirm cell viability and responsiveness by performing the S one control experiments before varying the S two experimental conditions.
