Immunolabeling and Counting Ribbon Synapses in Young Adult and Aged Gerbil Cochleae

This protocol enables the evaluation of the synaptic integrity in the cochlea of the gerbil of all ages. This is important for fundamental research into hearing problems, in particular, due to aging. The protocol is relatively fast and yields quantitative data from several locations along the cochlea.

Furthermore, it addresses a specific problem of aging tissue by using an autofluorescence quencher. Degeneration of synapses is the earliest sign of loss of function in the cochlea. Quantifying it is an important tool in research towards developing non-invasive diagnostic tools for synaptopathy.

The cochlear dissection is the most crucial part that needs practice, so it is better to start with non-experimental cochli. Begin by removing the bullae. Cut the bone with scissors or crack the bone with forceps and remove it with forceps to reach the cochlea.

Remove the malleus, incus, and semicircular canals. Carefully remove the bone covering the apex of the cochlea and scratch the cochlear bone with forceps to make small holes at the apex and in the basal turn or carefully poke holes in both locations. Immediately transfer the bullae into at least 50 milliliters of cold fixative.

To cut the cochli in half, position a piece of razor blade longer than the coiled length of the cochlea into a blade holder. Then, place the cochlea in a Petri dish under a stereo microscope. Cut away excess tissue with a razor blade or with fine scissors.

Hold the cochlea in place with fine forceps and cut in half along the modiolus. Prepare a 5%solution of the autofluorescence quencher by mixing it with 70%ethanol. Place the cochli in this solution and incubate it for one minute at room temperature on a shaker followed by washing thrice with one milliliter of PBS for five minutes.

For the dissection of the cochlea, collect two fine forceps, super-fine forceps, vannas, spring scissors, a blade holder, a razor blade, and fill polystyrol Petri dish and its lid with PBS. Break pieces from the razor blade to obtain a cutting surface of two to four millimeters. Cut the cochlea in half as shown in the section for tissue preparation.

Carefully fix the cochlear half with the forceps such that the cut edge is facing upwards. Use fine spring scissors to cut away the bone of the cochlea above the helicotrema covering the apex. Then, use scissors to isolate the apical turn by cutting through the modiolus and auditory nerve and scala vestibuli.

Make two cuts on each side of the cochlear bone above, or if the stria vascularis is not needed for further evaluation, directly through the stria vascularis of the middle turn. Use the razor blade to separate the cochlear pieces. Alternatively, cut between the Organ of Corti and the stria vascularis.

Leave the stria vascularis connected to the spiral ligament and remove the decalcified bone using forceps. Next, transfer the cochlear pieces into a Petri dish lid filled with PBS. Remove excess tissue on the pillar and modiolar sides such that the whole mounts will later lie on the slide as flat as possible.

Large cochlear pieces, in particular, those originating from the basal turn, should be cut into two pieces. Carefully remove the tectorial membrane with super-fine forceps. Place the cochlear pieces onto a slide into a drop of mounting medium, ensuring that the Organ of Corti faces up.

Shift the cochlear piece into the sagittal plane and look for invagination of the spiral limbus in close proximity to the inner hair cells. This invagination of the spiral limbus can be seen more clearly in the middle and basal cochlear pieces. Transfer the entire cochlea and the entire stria vascularis onto the microscope slide by repeating these steps.

Cover slip the slide and add black nail polish around the edges for sealing. Dry in the dark and store them at four degree Celsius. Open a copy of the deconvolved stacks in ImageJ software loaded with BioVoxxel plugin.

Split the channels by clicking on Image, then Color, and then Split Channels. Merge them by clicking on Image, then Color, and then Merge Channels to assign different colors. Convert the image to an RGB stack by clicking on Image, then go to Color and click Stack to RGB.

Adjust the brightness and contrast by clicking on Image, Adjust, and then Brightness/Contrast. Use either auto or slider to ensure that the pre-and post-synaptic structures and the inner hair cell labels are distinct from the background. Label five inner hair cells with the text tool by clicking on the desired location within the stack.

Next, go to Analyze, then Tools, and select ROI Manager. Check the tick box labels to label the points with a number. Zoom in on the inner hair cell of interest.

Right click on the point tool to choose Multi-point mode. Scroll through the Z dimension and count the functional ribbon synapses, which are identified by a pre-synaptic ribbon in juxtaposition to a post-synaptic glutamate patch. After selecting all the structures of interest, click Add the ROI Manager, then click on the arbitrary name and choose Rename.

With the hand tool, adjust the image to fully display the next inner hair cell. Change from Multi-point tool to Point tool to avoid adding more puncta to the previously stored data. Click on a structure of interest within the next inner hair cell and change to Multi-point tool.

Continue adding the functional ribbon synapse in the ROI Manager until all the inner hair cells are evaluated. Click on a data set in the ROI Manager and then on Measure. Wait for a new window to appear that lists the measured data points and click on File and Save As to save this list as a spreadsheet.

Save the image by activating Show All in the ROI Manager. Click on Flatten to permanently add the point labels to the stack and click on Agree to close by saving the changes. Z stack projections from the cochlea of a young gerbil aged 10 months show that the myosin 7A-labeled inner hair cells outlined here by white dashed lines were in sharp contrast to the background.

The pre-synaptic ribbons were labeled with anti-CtBP2 two shown in green, while the post-synaptic glutamate patches were labeled with anti-GluA2 shown in red. The cochlea from an old gerbil aged 38 months was treated with the autofluorescence quencher and showed distinct inner hair cells with clearly visible pre-and post-synaptic structures. To test the stability of the immunolabeling, the same piece of cochlea from the young gerbil was imaged after three months and 29 1/2 months.

In spite of the reduced signal-to-noise ratio, the functional synapses were distinct and quantifiable. Even if a cut fails, mount the remaining piece of tissue to later estimate the full cochlear length most accurately. The data from this protocol are particularly valuable if preceded by, and then individually correlated with, electrophysiological or psychophysical hearing tests.