Optical Clearing of Biological Tissue with ScaleA2
Atsushi Miyawaki (sakurai-h at brain dot riken dot jp)
Brain Science Institute, RIKEN, Japan
DOI
http://dx.doi.org/10.13070/mm.en.1.188
Date
last modified : 2014-06-09; original version : 2011-09-27
Cite as
MATER METHODS 2011;1:188
Abstract

A step by step protocol for optical clearing of biological tissue with ScaleA2 by ScaleA2 inventor Dr. Atsushi Miyawaki's lab

Editor's note: since the publication of Atsushi Miyawaki's ScaleA2 in 2011, at least three different clearing methods have been published. Clarity in the article "Structural and molecular interrogation of intact biological systems" in 2013 with a detailed protocol available; ClearT "ClearT a detergent- and solvent-free clearing method for neuronal and non-neuronal tissue." in 2013 [1] ; SeeDB "SeeDB: a simple and morphology-preserving optical clearing agent for neuronal circuit reconstruction" in 2013, with a detailed, updated protocol [2].

This step-by-step protocol for optical clearing of biological tissue with ScaleA2 [3] is provided by ScaleA2 inventor - Dr. Atsushi Miyawaki lab.

Solution preparation

PDF version.

Stock solution

10% (wt/vol) Triton X-100 solution
  1. Dissolve 10 g of Triton X-100 (e.g., Nacalai Tesque, Code 355-01 or similar grade) in 80 ml of Milli-Q water by stirring.
  2. Add Milli-Q water to make 100 ml and stir until well mixed.
  3. Store at 4 ºC.
ScaleA2 solution (1 liter)
  1. Dissolve 240.24 g of urea crystals (e.g., Wako Chemical, Code 217-615 or similar grade) in 800 ml of Milli-Q water by stirring.
  2. Add 10 ml of 10% (wt/vol) Triton X-100 solution.
  3. Add 100 g of glycerol (e.g., Sigma, Code 191612 or similar grade).
  4. Mix well by stirring.
  5. Add Milli-Q water to make 1,000 ml and stir until well mixed.
  6. Store at room temperature.
ScaleB4 solution (1 liter)
  • 8 M urea
  • 0.1% (wt/vol) Triton X-100
  1. Dissolve 480.48 g of urea crystals in 800 ml of Milli-Q water by stirring.
  2. Add 10 ml of 10% (wt/vol) Triton X-100 solution.
  3. Add Milli-Q water to make 1,000 ml and stir until well mixed.
  4. Store at room temperature.
Remarks
  • Allow each solution to stand after preparation for at least 1 day prior to use.
  • Sterilization is not required.
  • Addition of preservative is not required.
The following is a typical protocol for mouse brain.

PDF version.

  1. Fix a mouse via transcardial perfusion with 4% PFA ( paraformaldehyde) / PBS (w/v) (pH 7.5–8.0) at room temperature (RT).
    Acid fixatives (pH < 7.0) may quench FPs irreversibly. On the other hand, use of alkaline fixatives (pH > 8.0) may results in damage to samples later at Step 8.
  2. Remove the brain and postfix it with 4% PFA/PBS (pH 7.5–8.0) at 4 ºC for 8–12 hrs.
    It may be hard to fix the brain of older mice (> 3 weeks old) across the pia mater. For older mice, it is recommended to split the brain into two pieces. For example, unless commissural connections are examined, cut the brain at mid-plane into two cerebral hemispheres. The following procedure assumes that a cerebral hemisphere is processed as a sample to be cleared. Alternatively, if you prefer to keep the entire brain intact, making a few incisions will facilitate fixation throughout the whole brain.
  3. After washing with PBS, incubate the sample in 20% sucrose /PBS (w/v) (pH 7.4–7.8) at 4 ºC (or RT) for 1–2 days.
  4. Embed the sample in OCT compound (Sakura) and freeze with liquid N2.
  5. Thaw the sample in PBS (25 ml/ 0.5 g tissue) at RT for 20 min with gentle shaking.
  6. Rinse the sample in PBS (25 ml/ 0.5 g tissue) at RT for 20 min with gentle shaking.
  7. Fix the sample again with 4% PFA/PBS (pH 7.5–8.0) for 30 min at RT.
    It is highly recommended that sample fluorescence is checked at every step prior to Scaling (Step 8). It is somewhat common that the fluorescence can easily be lost during the fixation process. For example, incomplete fixation may result in wash out of soluble forms of fluorescent proteins (FPs).
  8. Transfer the sample into ScaleA2 solution (20 ml/ 0.5 g tissue) in a see-through vial.
    Since ScaleA2 is free of salt, PBS-derived salt remaining in the sample is gradually washed out. In the process of tissue clearing, salts cause white precipitates and thus should be avoided.
    It is important to use a container where you can easily assess sample transparency.
  9. Incubate the sample in ScaleA2 at 4 ºC (or RT) for 2–14 days or longer with gentle shaking. Exchange ScaleA2 if necessary.
    Clearing larger or harder (from older animals) samples requires longer incubation times. Check the transparency of the sample intermittently. Scale makes samples soft and fragile, like jelly, so care should be taken not to damage or destroy the sample.
    It may be hard to clear the brain of old mice (> 3 weeks old) across the pia mater. It is thus recommended the brain be split into two pieces or introduced with a few slits (see Step 2).
  10. Observe the sample under an upright microscope. Fresh ScaleA2 solution is used as the immersion medium.
    If the sample needs to be stabilized over an extended time period for observation, please see below for “the procedure for immobilizing cleared sample.”
  11. Store the cleared sample in fresh ScaleA2 solution at 4 ºC (or RT). Fluorescence is not lost over time during long-term storage. There is no need to add preservatives such as sodium azide.
Immobilizing scaled samples

PDF version. PDF version also has a schematic representation.

Melted agarose-water solution:

0.35% (w/v) agarose dissolved in Milli-Q water (Melt in a microwave oven and cool down to 37–40°C).

Steps
  1. Transfer a Scaled sample from Scale solution onto a plastic dish measuring 60–100 mm diameter. Air-dry for 10 min at RT. To dry the surface of the sample lightly, wick away excess Scale solution carefully with filter paper.
    Scaled samples are difficult to cut due to their softness. If the sample needs to be trimmed for observation, follow the procedures in Steps 2–6. If not, go to Step 7.
  2. Fill the dish with a melted agarose-water solution to embed the sample completely. Avoid introducing air bubbles into the solution.
  3. Allow the agarose to get to harden at RT and air-dry its surface.
  4. Trim the sample embedded in the agarose gel with a scalpel.
  5. Remove the agarose gel from the trimmed sample carefully.
  6. Place the (trimmed) Scaled sample onto a plastic dish measuring 60–100 mm diameter for observation using an upright confocal or two-photon excitation microscope. Position the sample with the observation part on top.
  7. Pour a melted agarose-water solution gently over the top of the sample. Let the viscous solution run radially down the sides. After hardening, the mountain-shaped agarose gel covers most of the sample deeply, but the summit should be covered with only a thin film of agarose gel. In this setup, the observation part should be accessible to an objective lens.
  8. Air-dry the surface of the entire agarose gel for approximately 30 min at RT. Then immobilize the gel edge onto the surface of the plastic dish with fast-drying adhesive (Aron Alpha or Krazy Glue).
  9. Pour ScaleA2 solution into the dish until the mountain is submerged. Gently shake the plastic dish using an orbital shaker for 3 hours at RT. The ScaleA2 solution is being diluted with the remaining water. Repeat this step with new ScaleA2 solution.
  10. When ScaleA2 is fully substituted as an immersion medium, the sample is ready for observation.
References
  1. Kuwajima T, Sitko A, Bhansali P, Jurgens C, Guido W, Mason C. ClearT: a detergent- and solvent-free clearing method for neuronal and non-neuronal tissue. Development. 2013;140:1364-8 pubmed publisher
  2. Ke M, Fujimoto S, Imai T. SeeDB: a simple and morphology-preserving optical clearing agent for neuronal circuit reconstruction. Nat Neurosci. 2013;16:1154-61 pubmed publisher
  3. Hama H, Kurokawa H, Kawano H, Ando R, Shimogori T, Noda H, et al. Scale: a chemical approach for fluorescence imaging and reconstruction of transparent mouse brain. Nat Neurosci. 2011;14:1481-8 pubmed publisher
ISSN : 2329-5139