Imaging Serotonergic Fibers in the Mouse Spinal Cord Using the CLARITY/CUBIC Technique

The overall goal of this CLARITY/CUBIC experiment is to reveal the 3D features of serotonergic fibers in the mouse spinal cord, which are not observable by traditional histology. This method can help answer key questions in the neuroanatomy field, such as how are serotonergic fibers distributed throughout the spinal cord? The main advantages of this technique are that it allows serotonergic fibers to be observed in situ, along with identification of vectors that are not observable by traditional histology.

Although this method can provide insight into the organization of the central nervous system, it can also be applied to other systems, such as the cardiovascular system. We first sought to use this method when we reached limitations using a traditional tracing method to analyze serotonergic fibers in the mouse spinal cord. Begin by fixing the limbs of an anesthetized mouse to a plastic dissection stage in a fume hood.

Confirm a lack of response to toe pinch and use a pair of scissors to open the skin over the chest. Next, open the chest to expose the heart and insert a 25 gauge needle attached to a peristaltic pump tubing into the left ventricle. Snip the right atrium to create an exit point for the blood.

Then, wash the heart with 40 milliliters of 9 percent saline at approximately 10 milliliters per minute. When all of the blood has been flushed, perfuse the animal with 35 milliliters of freshly prepared, ice cold hydrogel at the same perfusion rate. Then, incise the skin over the back of the mouse and dissect the muscles nest to the vertebrae.

After cutting the vertebral arches on one side and flipping them to the other, excise the spinal cord from the vertebral column with fine scissors. Immediately after harvesting, place the spinal cord in 15 milliliters of freshly prepared hydrogel solution overnight at four degrees Celsius. The next day, remove 10 milliliters of the hydrogel and pour the rest of the solution and spinal cord into a five milliliter tube.

Add fresh hydrogel solution until the tube is full and then stretch a small piece of parafilm over the top of the tube, taking care that the parafilm contacts the hyrdogel and that there are no bubbles between the solution and the film. Removing air from the tube is the most difficult step. To make sure there are no bubbles, take care to fill the tube slowly with the hydrogel solution and then carefully place the parafilm across the top of the tube, without spilling the gel.

Now, wrap a second piece of parafilm around the neck of the tube and incubate the spinal cord in a 37 degree Celsius oven overnight. When the hydrogel solution becomes a gel, use a spanner to remove the gel and neural tissue from the tube. To remove the hydrogel from the spinal cord, briefly press a course tissue against the gel.

The gel will stick to the tissue as it is removed. When the spinal cord is clean, place it in PBS for four, six hour washes on a shaker. After 24 hours, use a razor blade to cut the spinal cord coronally into two to three millimeter long segments and transfer the tissue pieces into five milliliters of freshly prepared CUBIC clearing solution.

Shake the pieces in the 37 degree Celsius oven for 72 hours and then replace the three day old clearing solution with freshly prepared clearing solution. Two to three days later, check the transparency of the tissue against the paper with font size 8 letters. When the letters can be observed through the cleared tissue, replace the CUBIC clearing solution with four milliliters of fresh PBST for four, six hour washes.

To stain the spinal cord segments for fluorescent imaging, incubate the tissue in the appropriate primary antibody solution for three days on a shaker in the 37 degree Celsius oven. At the end of the primary antibody staining, immerse the tissue in four milliliters of PBST for four, six hour washes in the 37 degree Celsius oven. Then, replace the PBST with the appropriate secondary antibody solution for three more days on the shaker in the 37 degree Celsius oven.

On the fourth day, wash away the unbound secondary antibody with PBST, as just demonstrated. To image the segments, replace the last PBST wash with four milliliters of 85 percent glycerol to make the refractive index of the tissue even. After about an hour, check the transparency of the tissue.

When the tissue is clear, transfer the spinal cord segments on to a large, thin glass slide. Next, place a few drops of 85 percent glycerol next to the spinal cord tissue and cover the tissue with a 22 by 50 millimeter glass cover slip. Place the slide into the holding frame of the microscope and move the tissue into the light path.

Then, select the Helium Neon laser and check the brightness of the positive signal in the live image to adjust the intensity of the laser. With the laser at the optimal intensity, use a 20x objective to select the scanning area of the spinal cord tissue. Then, set the instrument to create a z-stack with the depth of each stack at three microns.

Finally, scan the tissue from the top to the bottom of the z-stack, first, under the 20x objective, followed by a z-stack scan under the 63x objective. 3D software can then be used to reconstruct the appropriate 3D videos. Here, the presence of serotonergic fibers in all of the laminae of the spinal cord, with a predominance in the ventral portion of the ventral horn, can be observed.

In the ventral horn, densely packed serotonergic fibers are present in the ventral medial area, extending toward the lateral portion of the ventral horn, with some of the immunopositive fibers also extending from the ventral horn, toward the dorsal horn, or central canal. Immunopositive fibers are present in all of the laminae of the dorsal horn, but there is a small gap between the positive fibers in laminae two and four, particularly, in the lateral part of the dorsal horn. In a horizontal section, densely packed sertonergic fibers in ventral horn along the longitudinal axis of the spinal cord, can be observed issuing branches regularly along the axis perpendicular to the fiber bundle.

These branches further diverge along their paths, to the lateral portion of the ventral horn. Compared with these branches, those extending toward the midline are more irregular and smaller. Under the 63x objective, the serotonergic fibers in the dorsal horn, are observed to travel along the axis of the spinal cord and to terminate at various points along the path of the tract with thick fibers, sporadically distributed among the thin fibers.

Once mastered, this technique can be completed in less than three weeks, if it is performed properly. Following this procedure, other methods, like double labeling with choline acetyltransferase antibody, can be performed to answer additional questions about whether serotonergic fibers terminate motor neurons. After its development, this technique paved the way for researchers in the field of biomedical imaging to explore the complex spatial relationships between various cell types in animal specimens or human tissues.

After watching this video, you should have a good understanding of how to clarify the mouse spinal cord tissue for 3D imaging after immunofluorescent staining.