Immunolabelling and clearing of intact spinal cord for visualization of lower urinary tract afferents

While immersed in phosphate buffered-saline (PBS), pH 7.2, trim nerve roots of fixed spinal cord to within approximately 2 mm of the spinal cord surface to facilitate the identification of segments later following imaging.

Wash samples in 1x Dulbecco’s PBS (DPBS)(6 x 15 mins).

Dehydrate samples in a series of methanol in DPBS dilutions while on rotation at 12 rpm:

1. 50% methanol in DPBS (1.5 h)
2. 80% methanol in DPBS (1.5 h)
3. 100% methanol (1.5 h)

Bleach samples overnight in 6% hydrogen peroxide in methanol at 4°C, protected from light.

Rehydrate samples in a series of methanol in DPBS dilutions while on rotation at 12 rpm:

1. 100% methanol (2 x 1.5 h)
2. 80% methanol in DPBS (1.5 h)
3. 50% methanol in DPBS (1.5 h)
4. DPBS (1.5 h)

Incubate samples in DPBS containing 0.2% gelatin, 0.5% Triton X-100 and 0.01% thimerosal (DPBSG-T) for 36 h while on rotation at 12 rpm

Incubate samples in primary antibody solution containing DPBSG-T with 0.1% saponin for 10 days at 37°C with agitation. Volume of solution need only be sufficient to cover the sample.

Wash spinal cords in 1x DPBS with 0.5% Triton X-100 (DPBST) (6 x 15 mins).

Incubate samples in secondary antibody solution containing DPBSG-T with 0.1% Saponin for 4 days at 37°C with agitation. Volume of solution need only be sufficient to cover the sample.

Wash spinal cords in DPBST (6 x 15 mins).

Dehydrate spinal cords in a series of methanol in DPBS dilutions while on rotation at 12 rpm:

1. 20% methanol in DPBS (1 h)
2. 40% methanol in DPBS (1 h)
3. 60% methanol in DPBS (1 h)
4. 80% methanol in DPBS (1h)
5. 100% methanol (2 x 1 h)

Incubate samples in a solution of 2/3 dichloromethane and 1/3 methanol overnight on rotation. Ensure that samples sink to the bottom of the vial at the end of this step, otherwise continue incubation in freshly made solution.

Incubate samples in 100% dichloromethane for 30 mins while on rotation at 12 rpm. Repeat this step until samples sink.

Incubate samples in dibenzyl ether until the samples have become clear. Ensure each vial is completely filled with dibenzyl ether to minimize sample oxidation as a result of large amounts of air in the vial. This process should not take longer than 2 h.

Store cleared samples in fresh dibenzyl ether. Keep away from light (wrap in foil and store in an opaque container).

Prior to visualization on a light sheet microscope, samples should be transferred into ethyl cinnamate, at least 3 h prior.

Remove sample from ethyl cinnamate and gently dry on a tissue. Affix sample to plastic mount using the minimum amount of super glue required. Tips:

• Avoid adhering the sample to the base via any region of the sample that is of interest; both the super glue and proximity to the plastic will reduce imaging quality in that area.
• Mount the sample perfectly in the middle; the light sheet microscope stage has limited mobility, particularly at higher magnifications.
• Orientate the sample such that the thinnest plane of the sample is perpendicular to the light sheet beams. The further light has to travel through a sample, the poorer the image quality.
• Orientate the sample so that the region of interest is facing as close to the lens as possible. In this protocol, the spinal cord was mounted perpendicular to the light sheet beams, with the dorsal horn facing the lens, and the ventral side of L5 being the point of adherence to the mounting platform.

In order to visualize the lower urinary tract afferents in the dorsal horn of the spinal cord, use the highest magnification lens available on the light sheet microscope.

For example, a 12x lens is adequate for afferent visualization.

Look for cholera toxin subunit B labelled projections in the lateral and medial boundaries of the dorsal horn. One should also find dense afferent innervation of the sacral preganglionic nucleus of L6 and S1 spinal cord .