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The overall goal of this new experimental protocol is to demonstrate the accessibility of hair follicles in the mouse ear, for investigating the function of fully differentiated mechanosensory nerve terminals. This method can help answer key questions in the mechanosensory neuroscience field, such as the role of synaptic-like vesicles in mechanical sensation. The two main advantages of this technique are that is is quick, and it gives easy optical access to innumerable living fully-differentiated mechanosensory nerve terminals.

Generally, individuals new to this method may struggle with getting skin separation and adequately clearing cartilage without damaging follicles. As well as showing the basic technique, here we will also show the example of an experiment, looking at the calcium dependence of a vital part of the nerve terminus function. Fill a 50 millimeter silicone-lined dish with Liley's solution, or other physiological saline.

Refresh the solution every 15 minutes, or use a constant perfusion. After decapitating a young adult mouse, using scissors, remove the external ears near the base, just above the dense hair line, and transfer the pinnae in the dish. Now, pin an ear, concave side down, along the margins, using fine insect pins.

Next, carefully peel away the exposed posterior skin using blunt dissection. Using Number 3 forceps, grip the central posterior skin, and with another pair of forceps, pierce the gap between the skin and the cartilage at the base of the ear. Gently work the forceps from side to side within the gap, gradually separating the anterior and posterior skin layers.

Systematically move across the central cartilage to the thin cartilage-free margins. This will take practice. After removing the posterior skin, pin it, dermal side up, next to the anterior skin.

Once the cartilage is removed on both skin preparations, remove the layer of foamy fibroelastic cartilage that covers the base of the hair follicles. Gently peel, pluck, and rub this tissue off. This will also take practice to master.

Removing too little obstructs the dye and its visualization, while removing too much damages the nerve terminals. Using a young mouse minimizes the adhesion between the anterior and posterior skin layers and the difficulty of removing the underlying cartilage. Both of these will increase the likelihood of seeing successful staining.

Each skin preparation may be cut in half from the apex to the base, to maximize the number of replicants and minimize animals used. This section describes how to stain tissues with FM1-43, but it should work with most other styryl pyridinium dyes of this class with suitable adjustment to concentrations. Perform this procedure in minimal ambient-light levels.

Before beginning, make enough fresh 10 micromolar FM1-43 in the appropriate-gassed physiological saline for 3 milliliters per preparation. Put each preparation in a separate silicone-lined 35 millimeter dish, with 2 milliliters of the appropriate saline. To compare the ability of calcium and barium to sustain dye uptake, incubate in separate dishes from this point.

Put some in normal Liley's 2 millimolar calcium and others in Liley's where barium substitutes for calcium. Next, place the open dish preparations on a slightly-submerged platform in a 30 degree Celsius water bath. The water depth should be sufficient to warm the tissues without spilling into the dish.

Into each dish, pin a fine gas-line onto the silicone base to gas the preparations at about 10 bubbles per second. Also, warm up the FM1-43 solutions and 100 milliliters of dye-free saline in tightly-stoppered bottles. Let everything equilibrate to 30 degree Celsius, and after 30 minutes, replace the dye-free saline on the tissue using the appropriate warmed solution.

Incubate for a further 30 minutes. After the incubation, pour away the dye-free saline and quickly and carefully blot away any liquid remaining around the preparation. Be careful not to touch the exposed dermal surfaces.

Return the preparations to the water bath, keeping the gassing lines in place throughout. Now add 2 to 3 milliliters of calcium-or barium-containing warm dye solution. After 40 minutes, return the tissues to room temperature for the remainder of the procedure, to inhibit the re-release of internalized dye.

Now, remove the non-internalized dye in three stages:First, pour away the labeling solution and quickly rinse the tissues three times with room-temperature saline in rapid succession, and then let them incubate for 30 minutes to departition most of the remaining dye from the surface membranes. Lastly, remove the persistent dye stuck to the external leaflet of the exposed membranes by chelating the preparation with a sulfonated b-cyclodextrin derivative, made up in calcium or barium saline as appropriate. Thus, all remaining dye will be within the tissues.

After chelating, return the preparations to the appropriate fresh dye-free saline and dry the outside of the dishes thoroughly with tissue paper. Then, image them as soon as possible. Continue operating under minimal ambient light.

Before imaging, using a stereo microscope, remove surface debris that would auto-fluorescence and contaminate the images. Then, position the dish under an upright epifluorescence microscope with a standard fluorescein filter set. Next, attenuate the excitation light intensity using neutral-density filters until the preparation is illuminated just enough to comfortably locate the nerve terminals.

Full illumination will kill the nerve terminals rather quickly. Once located, capture images of the labeled lanceolate terminals using a 10x dry objective, or a 20x water immersion objective. Minimize the light intensity and exposure time by using a camera integration time of 1/2 to 1 second.

For each preparation, image about 20 lanceolate endings, starting at the margin furthest from the cut edge of the base of the pinna skin, and working along this margin, imaging each follicle in turn. Work in slightly-overlapping fields parallel to the cut edge without imaging the same follicle twice or imaging obviously damaged follicles. After completing the marginal strip, move one field-width towards the base of the pinna and repeat the process in the reverse direction.

Analysis is described in the text protocol. After imaging the first dish, showing follicles labeled in calcium-containing saline, image the follicles in barium-containing saline. Continue in this manner.

Ensure the imaging of the remaining control and experimental tissues are time-matched, as after labeling, dye will subsequently be released again by the exocytosis arm of constitutive SLV recycling. In a typical pinna preparation, hair follicles are easily seen under transillumination without fluorescence, illustrating the wafer-thin nature of the preparation and the relative ease of accessibility it affords to these mechanosensory terminals. Under epifluorescence, the labeled lanceolate terminals surrounding each hair follicle are clearly seen and show the typically robust spontaneous FM1-43 uptake.

The punctate pattern reflects terminals being observed in an optical plane orthogonal to the skin, along the length of the terminal. One of the many applications of this method is in the study of exocytosis kinetics. As expected, if dye uptake is due to vesicle recycling, in the top panels, a labeled terminal is seen de-staining spontaneously.

After applying a powerful stimulant of exocytosis, alpha-latrotoxin, the natural de-staining process is accelerated, as seen in the bottom two panels. Another application, as observed in the video, is to examine the calcium dependence of endocytosis. Distinct dye uptake was seen for lanceolate terminals in both calcium-and barium-containing salines.

On quantifying the fluorescence intensity of the terminal seen labeling during the video, no significant difference was found in the dye uptake. This indicates endocytosis is supported equally well by barium and calcium. Further repetitions of the procedure on other ear preparations will test this conclusion.

Once mastered, this entire protocol and imaging can be completed in two-and-a-half hours. When attempting this procedure, it is important to remember to use small mice, to dissect quickly, and to be maximally dark-adapted to minimize the light intensity required. We adapted the idea for this method from one originally developed by a close colleague and now-retired professor, Clarke Slater, and his MSc student Michael Caine.

When we told them of the possible role for synaptic-like vesicles in mechanosensation in the muscle spindle, but of the need for a better way of visualizing than with FM1-43. This procedure can also be used in combination with other methods, such as pharmacology and electrophysiology. Then we can study how synaptic-like vesicle recycling is regulated, and how this relates to responsiveness of the mechanosensory endings.