Feel free to ask questions, and we’ll add them to the list.
What tissues can I use iDISCO with?
We validated iDISCO in mouse embryos up to E16.5, as well as E18.5 for specific cut regions. In adult mouse tissues, we validated the method for brain, spinal cord, intestine, skin, kidney, stomach, heart, skeletal muscle, spleen, liver.
Why does the protocol use Heparin?
The use of heparin is inspired by cell binding assays, where it is use to reduce the background staining. The heparin binds to cell-surface glycoproteins and prevent them from sticking to the ligands and antibodies.
How long are the washes?
We wash the primary and secondary antibodies out for 24h, and usually change the wash solutions more frequently over the first hour, then we change the solution a couple of times over the rest of the day (approximately every 2 hours). Exact timing of washes is not critical.
What are the convenient stop points?
The sample can be stored in methanol 100%. The time in blocking buffer can be extended a couple of days to be more convenient with your schedule. We have also found leaving the sample washing out of antibody an extra day has no deleterious effects.
Can I do multiple rounds of staining?
We do not know of a method to reverse the clearing.
The antibody did not diffuse to the center of the sample.
If the sample is completely dark at the center, the antibody is depleted. Use a higher concentration of antibody. If there is a gradient of staining from the periphery but the center is still stained, use slightly higher concentrations and increase the incubation time.
It is important to consider the density of antigens in the tissu when designing the staining experiment. The diffusion of the IgG in the adult tissues after methanol fixation is not limited by the ability of the IgG to randomly diffuse, but is limited by the interaction with antigens. Therefore, if the antigens are distributed as a dense mesh, they will form a “net” that will capture the IgG and prevent their diffusion. Examples of IgG that will diffuse very poorly in adult brains are Neurofilament and GFAP.
There is a strong surface background / ring-like background staining.
This seems to happen because the primary antibody is too concentrated. Reduce the concentration.
The samples turned opaque!
There are only two reasons the clearing could fail:
- You used a THF without BHT. The protocol reference contains BHT in the THF, which is essential to the clearing without oxydation.
- You had too much air in the vial of DBE or THF. For THF, fill the tube up to more than half at least. For DBE, fill the tube to the top, and limit the shaking to 1h.
THF solution appears cloudy.
This will happen on a rare occasion if you introduced to much air while mixing water and THF.
The samples have an amber color.
If the colouring is light, this is normal and will not prevent the imaging. If the amber color is too pronounced, the sample was kept for too long in THF, or it got oxidised because too much air was present in the tube.
The samples are hard after the clearing.
This is normal and also makes samples easier to handle!
The sample shrinks during the clearing.
This is normal because of the dehydration, but you can limit the amount of shrinkage by reducing the time in THF 100%. The total time spent in THF 100% is not critical to the clearing. If you are working with embryos or early post-natal brains, you can also limit the amount of time in DCM as well.
How long can I keep the samples after clearing?
We have not found the upper limit yet, we have samples over a year old that still look good as new. As long as the sample is protected from light and the DBE tube is filled to the top, the sample and staining signal should last many months.