The CLARITY staining protocol enables high-resolution, three-dimensional imaging of intact tissues by rendering them optically transparent and permeable to macromolecules. CLARITY is part of a broader family of optical clearing methods designed to improve imaging of biological tissues. This advanced tissue clearing method uses a hydrogel-based approach to preserve structural integrity while allowing deep antibody or probe penetration. CLARITY is compatible with fixed mouse brain samples and other fixed tissue types, making it suitable for a variety of research applications. Ideal for neuroscience and developmental biology, CLARITY is compatible with a wide range of tissues, including mouse, zebrafish, and post-mortem human brain samples. The protocol includes detailed steps for solution preparation, equipment setup, perfusion, and tissue processing, covering key tissue processing methods relevant for high-resolution imaging.

Introduction

CLARITY is a transformative technique in tissue imaging, developed to overcome the limitations of traditional sectioning and staining methods. Consistent tissue clearing is essential for achieving reliable and reproducible imaging results, as it preserves tissue morphology and ensures uniform transparency across samples. By embedding tissues in a hydrogel matrix and removing lipids through electrophoresis, CLARITY preserves fine anatomical structures while enabling deep molecular labeling, including antibody staining as a key application. This protocol is particularly valuable for researchers aiming to visualize complex neural networks or study disease pathology in whole organs, with a clearing process optimized for both structural and molecular interrogation. This CLARITY staining guide offers a step-by-step approach to preparing reagents, clearing tissues, and optimizing imaging conditions. It supports a wide range of applications, from basic research to translational studies, and is compatible with both fluorescent and non-fluorescent samples.

Background and principles

The CLARITY method is based on the principle of hydrogel-tissue hybridization, where acrylamide monomers are polymerized within fixed tissues to form a stable, porous mesh. Hydrogel-based tissue clearing techniques like CLARITY create hydrogel-based tissue constructs that preserve tissue physicochemical properties such as optical transparency, molecular permeability, and structural integrity. This mesh retains proteins and nucleic acids while allowing lipids to be removed using sodium dodecyl sulfate (SDS) and electrophoretic tissue clearing (ETC). The result is a transparent, structurally intact sample that can be labeled with antibodies or RNA probes. Developed by the Chung Lab, CLARITY revolutionized tissue imaging by enabling in situ analysis of large, intact samples. The protocol emphasizes temperature control, slow perfusion, and careful reagent handling to ensure optimal tissue preservation and imaging clarity.

Developed by the Chung Lab.

Materials required

Reagents

Beuthanasia-D

32% Paraformaldehyde (PFA)

40% ​Acrylamide solution

Azo-initiator

10X PBS

Ultrapure distilled water

Boric acid

Sodium dodecyl sulfate (SDS)

Lithium hydroxide monohydrate

N-methyl-D-glucamine

Diatrizoic acid

60% Iodixanol

Triton-X or NP-40

Sodium azide

1X PBS

Equipment

Transcardial perfusion of fixatives and hydrogel monomers

Dissection board (Styrofoam lid is fine)

20 mL syringes with luer lock ends

1 mL syringes

Winged infusion sets

Needles

Absorbent pads

50 mL Falcon tubes

Guillotine, for sacrificing larger animals

Surgical scissors

Hemostats

Forceps

Spatula

Hydrogel-tissue hybridization

Desiccator with 3-way stopcock

Vacuum pump

Compressed nitrogen tank

Compressed gas tank pressure regulator

Teflon tape

3/8” tubing

3/8” to 1/4" barbed tubing connector

ETC clearing system

Buffer Filter with Light-Blocking Blue Bowl

Platinum wire with 0.5 mm diameter

Bottle for Chamber fabrication

Nalgene Straight Side Jar – Poly, 32 oz

Single barbed tube fitting (7/16” hex for 1/4” tubing)

Tube to tube coupling for 3/32” to 1/16” tubing

3M Duo adhesive dispenser

3M Duo adhesive-mixing applicators

3M Duo adhesive cartridges

Sample holder

Bio-Rad HC PowerPac System

Banana to Large Alligator Test Lead Set

Clear 1/4" tubing

Clear 5/8” tubing

1/4" wye connector

4x Chemical resistant stopcock 1/4" to 1/4"

5/8" to 1/4" tubing connection

Elbow connection 1/4" male pipe to 1/4" barbed fitting

Elbow connection 1/4" barbed fitting

Rubber grounding plug

Magnetic water pump

Tissue

In principle, any tissue type from any animal of any age with or without fluorescence can be used. It has been demonstrated that CLARITY is compatible with the whole adult mouse brain, whole adult zebrafish brain, and extensively formalin-fixed post-mortem human brain section (without the perfusion step and further optimization in this case).

Tissues with strong fluorescent protein expression can undergo CLARITY processing described in this protocol and then be directly imaged; tissues without fluorescent proteins can be labelled with antibodies or RNA probes for subsequent imaging.

Stage 1 - Solution preparation

Steps

Keeping all reagents on ice, prepare a 10% stock solution of initiator solution by dissolving 1 g of azo-initiator in 10 mL UltraPure water.

• Then prepare the following solution:

Steps

Adjust the following solution to pH 8.5 using boric acid.

Steps

Make a solution consisting of 0.1% Triton-X (0.1% NP-40 can be used instead) and 0.1% Sodium Azide using 1X PBS.

Steps

This solution consists of 23.5% (w/v) N-methyl-D-glucamine, 29.4% (w/v) diatrizoic acid, and 32.4% (w/v) iodixanol in water.

• Use a stir bar (or shake if necessary) to fully dissolve the powders at each step.
• Ensure the solution is stored carefully to ensure no water is lost, as just a small amount of evaporation will result in precipitation.
• The optical clearing solution in the case would look as follows:

*To create this, add approximately 2.75 mL water to every 10 mL of 60% iodixanol solution.

• Add reagents in order

Stage 2 - Equipment setup

Steps

Mount the nitrogen tank with an appropriate tank bracket and attach the regulator to the tank outlet using Teflon tape if necessary to prevent leaking.

Run 3/8” tubing from the regulator outlet to the stopcock of the desiccator using a 3/8” to 1/4” barbed tube fitting.

Connect the vacuum pump to the desiccator by simply connecting the supplied tubing to the barbed fitting on the stopcock.

Steps

Create the measurement reservoir in a similar manner using a 32 oz Nalgene bottle and 1/4" barbed elbow connectors.

Apply epoxy to both the inside and outside parts of the connection and allow to dry overnight.

Create two more holes in the lid of the bottle, large enough for insertion of a pH probe and a thermometer for data acquisition.

• These holes should be left unsealed.

Create a heat exchange module by measuring out two pieces of around 2 ft of 1/4" tubing.

Connect these to the system in parallel using wye connectors and submerge in water.

Connect the water filter to the system using 1/4" male pipe thread to 1/4" tubing elbow connectors.

Tube all the components of the system together using 1/4" tubing, though 5/8” tubing will be needed for the pump connection.

• Use a reducing fitting to connect this larger tubing to the rest of the system.
• Critical step: the system should be connected in the following order (in the direction of flow): pump, filter, ETC chamber, measurement reservoir, and heat exchanger.

Stage 3 - Perfusion and tissue preparation

Steps

Make a fresh batch of hydrogel monomer solution, or thaw frozen stock solution at 4⁰C or on ice.

• After the solution is completely thawed and transparent (but still ice-cold), gently invert to mix.
• Ensure no precipitation or bubbles are seen in the solution.

Deeply anesthetize an animal with beuthanasia-D (0.5 mL per 1 kg of body weight intraperitoneally).

• Surgically open the chest cavity with a midline abdominal incision that bifurcates rostrally into a Y-shape.
• Punch a small hole in the right atrium and insert an injection needle into the left ventricle to allow perfusion.

Prepare two syringes filled with ice-cold PBS and hydrogel monomer solution, respectively, each with winged needle sets for each solution.

• In the case of mice, perfuse first with 20 mL of ice-cold PBS at a rate of less than 5 mL/min, carefully take the needle out and perfuse with 20 mL of the ice-cold hydrogel monomer solution. Rats require about 200 mL of each solution at the rate of 20 mL/min.
• Critical step: maintain a slow rate of perfusion. We found that injecting less than 5 mL per minute for both solutions in the case of mice yields better results. Use extreme caution not to introduce bubbles to the vasculature (especially when introducing needles), as this decreases the quality of perfusion.

Carefully harvest the organs of interest and place them immediately in a 50 mL conical tube containing 20 mL of the ice-cold hydrogel monomer solution for both post-fixation and even infiltration of monomers.

• Keep this on ice until it can be transferred to a 4⁰C refrigerator.

Incubate the sample for one day at 4⁰C to allow for further distribution of monomer and initiator molecules throughout the tissue.

Stage 4 - Hydrogel tissue embedding

Steps

After the tissues have been allowed to incubate in the hydrogel monomer solution for one day, move the samples to 10 mL of fresh hydrogel monomer solution.

Place the conical tubes in a desiccation chamber on a tube rack and unscrew the caps about halfway.

Connect nitrogen gas and a vacuum pump to the desiccator via the three-way stopcock.

• Open flow in all three directions and turn on the nitrogen gas.
• Allow the gas to flow for about five seconds.

Without turning off the nitrogen flow, turn on the vacuum pump and adjust the stopcock so that flow is only open to the desiccator and the vacuum pump.

• Allow the vacuum pump to run for at least ten minutes.

Very slowly turn the stopcock so that flow is only open to the nitrogen gas and the desiccator, then turn off the vacuum pump.

• Allow the desiccation chamber to fill with nitrogen gas.

Very quickly, lift the lid of the desiccator and tighten the caps of the conical tubes inside.

• The nitrogen gas can now be shut off.

Gently shake the samples in a 37°C warm room for two hours.

• This temperature will trigger radical initiation by the azo-initiator.

To remove unreacted PFA, wash the samples in 50 mL of clearing solution at 37°C for 24 hours, with gentle shaking.

• Do this a total of three times.

Stage 5 - Electrophoretic tissue-clearing

At this point, you should have already constructed an ETC system as detailed in the section equipment setup.

Steps

Add the sample to the ETC chamber and close the lid.

• Connect any remaining unconnected tubing.

Fill the system with clearing solution by first filling the measurement reservoir and placing it on a surface a few inches higher than the level of the heat exchanger and pump.

• This will allow the buffer to fill the tubing.
• Start the pump and add more clearing solution to the measurement reservoir as needed to fill the system.

Connect the electrodes to the lead cables and start the power supply.

• Use around 40 V.

Check the samples regularly to determine that the system is working properly and that clearing is progressing.

• The entire process should take several days.

Remove the cleared samples from the ETC system and wash them twice with for 24 hours each.

Place the sample in a volume of optical clearing solution that is sufficient to cover the tissue completely and allow it to incubate for two days.

• After the first day, move the sample to a container of fresh optical clearing solution.

To image the cleared sample, it must be mounted between a glass slide and a black Willco dish.

• Roll up a piece of Blu-Tack adhesive into cylinder shapes of a thickness slightly more than the thickness of your sample.
• Place them horizontally on the glass slide.
• Press down the edge of Blu-Tack to close up the gap between the Blu-Tack adhesive and the glass slide

Carefully place the sample in between the Blu-Tack pieces and add about 20 μL of optical clearing solution to the sample.

With the lipped side facing up, firmly press a Willco dish down onto the adhesive until it just comes into contact with the sample.

• Using a pipette, add more optical clearing solution to the gaps between adhesive until the imaging chamber is filled.

KWIK-SIL is an adhesive that cures rapidly – carefully add it to the gaps between the Blu-Tack to build a wall and seal in the sample.

Cover this construction with aluminum foil and store it away safely to cure.

• After about 20 minutes, the sample is ready for imaging.

Types of clearing methods

Tissue clearing methods have evolved to address the challenges of imaging intact biological systems by making tissues transparent while preserving their structural and molecular integrity. These methods are generally classified into three main categories: hydrophobic, hydrophilic, and hydrogel-based clearing methods.

Hydrophobic clearing methods, such as 3DISCO and iDISCO, utilize organic solvents to efficiently remove lipids from tissues, resulting in rapid clearing and high transparency. These approaches are particularly effective for large tissue samples and are often chosen for their speed. However, the use of harsh solvents can sometimes compromise the preservation of fluorescent proteins and delicate tissue structures.

Hydrophilic clearing methods, including CUBIC and Scale, employ water-soluble reagents to achieve tissue transparency. These methods are gentler on biological tissues and are well-suited for preserving endogenous fluorescent proteins, making them ideal for studies that rely on fluorescence imaging. Hydrophilic methods are also compatible with a wide range of tissue types, but may require longer clearing times compared to hydrophobic approaches.

Hydrogel-based clearing methods, such as CLARITY and SHIELD, have significantly advanced tissue clearing. These methods stabilize proteins and nucleic acids by embedding tissues in a hydrogel matrix while enabling complete lipid removal. This approach preserves the tissue’s physicochemical properties and allows for deep molecular interrogation, including multiplexed antibody labeling and RNA detection. Hydrogel-based methods are especially valuable for applications that require high-fidelity imaging and molecular retention, such as neuronal circuit reconstruction and whole-brain imaging.

The choice of tissue clearing method depends on the specific research goals, tissue type, and desired downstream applications. Whether prioritizing rapid lipid removal, preservation of fluorescent proteins, or compatibility with advanced imaging and labeling, researchers can select from a versatile palette of clearing methods to best suit their experimental needs.

Imaging techniques

The visualization of cleared tissues at high resolution relies on advanced imaging techniques that can capture the intricate details of intact organs and large tissue samples. Light sheet microscopy has emerged as a leading technology for imaging cleared tissues, offering rapid, volumetric imaging with minimal photobleaching. This technique is particularly well-suited for imaging whole mouse brains and other organ-scale tissues, enabling researchers to achieve single-cell resolution across large volumes.

Light sheet microscopy works by illuminating a thin plane of the sample with a sheet of light, allowing for fast acquisition of optical sections and three-dimensional reconstruction. This approach is ideal for high-resolution imaging of cleared brain tissue, facilitating the study of neuronal circuits, brain regions, and brain activity in intact tissues. The ability to image entire organs, such as the whole mouse brain, has transformed our understanding of complex biological systems.

Other imaging modalities, such as confocal and two-photon microscopy, are also compatible with cleared tissues and can provide detailed images at subcellular resolution. However, these techniques often require longer imaging times and may be more susceptible to photobleaching, especially when imaging large or thick samples.

Recent innovations, including lattice light-sheet microscopy and adaptive optics, have further expanded the capabilities of resolution imaging in cleared tissues. Lattice light-sheet microscopy enables rapid, high-resolution imaging of entire mouse brains, while adaptive optics corrects for optical aberrations, improving image quality and depth penetration.

By combining tissue clearing with state-of-the-art imaging techniques, researchers can achieve unprecedented insights into the structure and function of biological tissues, from the cellular to the organ level.

Comparison to other methods

Compared to traditional histology, CLARITY offers superior depth and resolution by eliminating the need for physical sectioning. Unlike solvent-based clearing methods such as iDISCO or BABB, which are commonly used for solvent-cleared organs and enable deep tissue imaging and long-term preservation, CLARITY preserves endogenous fluorescence and is less damaging to tissue morphology. While methods like CUBIC and Scale also achieve transparency, CLARITY’s hydrogel matrix provides better molecular retention and compatibility with electrophoretic clearing. However, CLARITY requires more specialized equipment and longer processing times. Although solvent-based methods are often preferred for whole body clearing and facilitate multi-modal brain imaging approaches by allowing comprehensive visualization and integration of different imaging techniques, CLARITY’s strength lies in its ability to combine structural preservation with deep molecular access, making it ideal for applications requiring high-fidelity 3D imaging of complex tissues.

Applications

CLARITY is widely used in neuroscience, developmental biology, and pathology for high-resolution imaging of intact tissues, and is especially applied to the nervous system, including the cerebral cortex. It enables detailed visualization of neuronal circuits, vascular networks, and cellular architecture in whole organs. Researchers use CLARITY for immunohistochemistry, in situ hybridization, and fluorescent protein imaging in mouse, zebrafish, and human tissues. CLARITY allows for the analysis of immediate early genes and early genes in the same mouse brain, enabling comprehensive activity mapping and neural circuit analysis. Stochastic electrotransport selectively enhances molecular labeling efficiency during tissue clearing, improving the penetration and targeting of specific molecules. It is particularly valuable for studying neurodegenerative diseases, brain development, and tumor microenvironments. The protocol supports both endogenous fluorescence and antibody-based labeling, making it versatile for a range of experimental designs. Its compatibility with confocal and light-sheet microscopy further enhances its utility in advanced imaging workflows.

Limitations

Despite its advantages, CLARITY has several limitations. The protocol is time-intensive, requiring multiple days for tissue preparation, clearing, and labeling. It also requires specialized equipment, such as vacuum pumps and electrophoretic chambers, which may not be readily available in all labs. Tissue shrinkage or expansion can occur if uneven monomer infiltration affects structural fidelity. Additionally, the method may not be suitable for very large or calcified tissues without further optimization. Fluorescent signal loss can happen if samples are not properly protected from light. These challenges necessitate careful planning and adherence to protocol details for successful outcomes.

Troubleshooting

Common issues in CLARITY staining include incomplete clearing, tissue damage, and poor antibody penetration. If tissues remain opaque, ensure SDS concentration and electrophoresis parameters are correct, and verify that the hydrogel polymerized uniformly. Bubbles during perfusion can disrupt tissue integrity; use slow, steady flow rates and avoid introducing air. Weak staining may result from insufficient incubation times or degraded antibodies—optimize concentrations and use fresh reagents. If fluorescence fades, protect samples from light and minimize exposure during processing. For uneven clearing, confirm that monomer infiltration was complete and that temperature was consistently maintained at 4°C during incubation. During troubleshooting, use microscopic observation to assess tissue integrity and staining quality.