Labeling

We extensively use the SWITCH strategy for large volume labeling. Make sure to read, read, and read the section above before you start doing any large volume labeling! Here we will provide two protocols: one for myelinated fiber labeling and one for antibody labeling. You can use these as a starting point for your labeling adventures.

Myelinated fiber labeling protocol

You will need the following reagents:

Make the following solutions EXACTLY:

• SWITCH-Off: 10 mM SDS in 1X PBS (dissolve some SDS into 1X PBS)
• SWITCH-On: 1X PBS, 0.2% Triton X-100 (aka PBST; dissolve some Triton X into 1X PBS. We recommend you make a 10% Triton X stock to let it dissolve faster)

We take advantage of the fact that DiD staining doesn't work at 10 mM SDS. So we let the DiD diffuse into the tissue with SDS (SWITCH-Off), then switch the buffer to one without SDS, to label the myelinated fibers (SWITCH-On).

Myelinated fibers can be readily visualized with the lipophilic DiD fluorescent molecule. The sample should be equilibrated in a solution of 10 mM SDS in PBS in order to distribute SDS molecules throughout the sample. The sample should then be placed in a volume of DiD-OFF solution just large enough to cover the sample and incubated at 37 ˚C with gentle shaking for 12 hrs to 7 days depending on the size of the sample (1 mm-thick section to whole mouse brain, respectively). The sample should then be moved to 40 mL of PBST and incubated at 37 ˚C for 12 hrs to 2 days (same deal). We have also observed that tomato lectin and nuclear stains such as DAPI or Syto16 can be used with this SWITCH approach.

Antibody labeling protocol

You will need the following:

You will also need some antibodies.

Again, make the following solutions:

• SWITCH-Off: 0.5 mM SDS in 1X PBS
• SWITCH-On: 1X PBS, 0.2% Triton X-100

You can add more SDS if you find that 0.5 mM SDS is not inhibiting your antibody well enough. You can add up to about 10 mM SDS, and then there's no additional benefits. This is because only the monomeric form of SDS inhibits binding, and the maximum amount of monomeric SDS is about 10 mM -- beyond that point (called the critical micelle concentration), the SDS will exist as a mixture of micelles and monomers (e.g. 200 mM SDS has 190 mM micelles and 10 mM monomers).

Onwards: the sample should be equilibrated in antibody-OFF solution in order to distribute SDS molecules throughout the sample. The sample should then be placed in a fresh volume of antibody-OFF solution just large enough to cover the sample, and then antibodies should be added in the desired proportions. The sample should then be incubated at 37 ˚C with gentle shaking for 12 hrs to 7 days depending on the size of the sample (1 mm-thick section to whole mouse brain). The sample should then be moved to 40 mL of PBST and incubated at 37 ˚C for 12 hrs to 2 days.

Delabeling

You can strip the antibodies for multiple round labeling by incubating the sample in the clearing solution at 70 ˚C for 2 hours to O/N depending on the size of the sample. Basically, think of it like you're clearing again, except you're clearing away the antibodies. You can then label again, following the labeling protocol. It's like a coloring book -- color one color, take a picture, erase, and color again.

Optical clearing

The tissue is transparent, but not transparent enough for imaging. Before you can image your labeled samples, you should optically clear the tissue by incubating it in a refractive index matching solution. We'll write another post that explains how this all works, but for now, you can follow the following protocol for our custom refractive index matching solution.

Our optical clearing solution requires:

Make a 23.5% (w/v) n-methyl-d-glucamine, 29.4% (w/v) diatrizoic acid, and 32.4% (w/v) iodixanol solution. Basically, add 40 g n-methyl-d-glucamine, 50 g diatrizoic acid, and 55 g iodixanol to 100 mL of water. Or, if you're using the 60% iodixanol solution from Sigma, add 40 g n-methyl-d-glucamine, 50 g diatrizoic acid, and 92 mL of 60% iodixanol to 63 mL of water.

Do not use heat when mixing the solution, as this will cause a color change. This solution should be stored carefully to ensure that no water is lost, as just a small amount of evaporation will result in precipitation. Teflon tape can be used to increase the security of the bottle’s seal, and parafilm can be used around the cap.

The sample should be washed in optical clearing solution at least 3x for 6 hrs each at 37 ˚C with gentle shaking. After the final wash, the sample should be clear enough to easily see through by eye. If the solution immediately surrounding the sample seems inhomogeneous, it suggests that the sample has not yet fully equilibrated with the solution.