Optical Clearing of the Mouse Central Nervous System Using Passive CLARITY

Summary

Optical clearing techniques are revolutionizing the way tissues are visualized. In this report we describe modifications of the original Clear Lipid-exchanged Acrylamide-hybridized Rigid Imaging-compatible Tissue-hYdrogel (CLARITY) protocol that yields more consistent and less expensive results.

Abstract

Traditionally, tissue visualization has required that the tissue of interest be serially sectioned and imaged, subjecting each tissue section to unique non-linear deformations, dramatically hampering one's ability to evaluate cellular morphology, distribution and connectivity in the central nervous system (CNS). However, optical clearing techniques are changing the way tissues are visualized. These approaches permit one to probe deeply into intact organ preparations, providing tremendous insight into the structural organization of tissues in health and disease. Techniques such as Clear Lipid-exchanged Acrylamide-hybridized Rigid Imaging-compatible Tissue-hYdrogel (CLARITY) achieve this goal by providing a matrix that binds important biomolecules while permitting light-scattering lipids to freely diffuse out. Lipid removal, followed by refractive index matching, renders the tissue transparent and readily imaged in 3 dimensions (3D). Nevertheless, the electrophoretic tissue clearing (ETC) used in the original CLARITY protocol can be challenging to implement successfully and the use of a proprietary refraction index matching solution makes it expensive to use the technique routinely. This report demonstrates the implementation of a simple and inexpensive optical clearing protocol that combines passive CLARITY for improved tissue integrity and 2,2′-thiodiethanol (TDE), a previously described refractive index matching solution.

Introduction

The ability to image complete neuroanatomical structures is immensely valuable for understanding the brain in health and disease. Traditionally, 3D imaging has required tissue sectioning to provide the axial resolution and to visualize deep anatomical structures. This approach can produce high-resolution data sets, but requires sophisticated image reconstruction techniques and is very labor intensive. As a result, it has been limited to the imaging of small volumes of tissue^1-3. Optical sectioning, on the other hand, is well suited for the creation of high-resolution 3D images of fluorescently labeled tissues. Since optical sectioning is inherently three dimensional, it does not require extensive computation to produce a 3D image volume. However, light scattering and tissue opacity limit the depth at which tissues can be optically sectioned. The depth of imaging is limited to about 150 µm in laser scanning confocal microscopy and to less than 800 µm using two-photon excitation microscopy^4-8.

In order to overcome these limitations, several optical clearing techniques have been recently developed and then further refined to permit the deep microscopic imaging of intact tissues. The use of benzyl alcohol and benzyl benzoate (BABB) to render fixed tissues transparent was among one of the earliest approaches⁹. However, this approach was limited by the quenching of fluorescence in the samples and incomplete clearing of highly myelinated structures¹⁰. Refinements of this technique, such as 3D Imaging of Solvent Cleared Organs (3DISCO), have led to very rapid and complete tissue clearing, but still suffer from rapid loss of fluorescent signals, especially yellow fluorescent protein (YFP)¹⁰. Water-based clearing solutions, such as Sca/e¹¹ and SeeDB¹², preserve fluorescent signals, but do not completely clear highly myelinated tissues. Clear, Unobstructed Brain Imaging Cocktails and Computational analysis (CUBIC) is a promising new optical clearing technology that appears to overcome the limitations of previously developed water-based clearing solutions¹³. In contrast with other optical clearing techniques, Clear Lipid-exchanged Acrylamide-hybridized Rigid Imaging-compatible Tissue-hYdrogel (CLARITY)¹⁴ embeds the brain in a porous matrix that provides structural integrity to proteins, nucleic acids and small molecules, while leaving lipids unbound. The lipids are then removed electrophoretically, culminating in an optically cleared brain that can be easily visualized and equally easily probed using commonly available techniques.

Electrophoretic tissue clearing (ETC) is used to remove lipids from hydrogel embedded tissues in CLARITY, however, ETC can be difficult to implement consistently and tissues subject to electrophoresis can exhibit tissue distortion, browning, loss of fluorescence and antibody reactivity. Passive lipid clearing avoids these limitations, but requires increased clearing times^15-17. Passive clearing also permits the clearing of large numbers of samples in parallel, since it is not limited by the number of ETC apparatuses available. Decreasing the concentration of paraformaldehyde and excluding bis-acrylamide from the hydrogel have led to great increases in the speed of clearing at the expense of greater tissue expansion¹⁶. Less expensive alternatives to the refractive index matching solution used in the original CLARITY technique have been developed^17-19. 2,2'-thiodiethanol (TDE) is an inexpensive and rapid brain clearing agent that reverses some of the tissue expansion that occurs during lipid removal¹⁸. This report incorporates passive clearing of hydrogel embedded tissue and the use of TDE as a refractive index matching solution to the original CLARITY technique to produce a highly reproducible and inexpensive protocol for the optical clearing of the mouse central nervous system.

Protocol

All experiments were performed in accordance with Institutional Animal Care and Use Committee (IACUC) guidelines. THY1-YFP, PLP-EGFP and PV-TdTomato mice were used for these experiments, but any mice expressing fluorescent proteins can be successfully cleared and imaged.

1. Tissue Preparation

Caution: Paraformaldehyde (PFA) and acrylamide are toxic and irritants. Perform experiments utilizing these reagents in a fume hood and with appropriate personal protective equipment (lab coat, gloves, and eye protection).

1. Sacrifice the animal in euthanasia chamber with ~1 ml of isoflurane until breathing ceases (~2 - 3 min).
2. Use a peristaltic pump set to ~5 ml/min to intracardially perfuse the mouse with perfusion solution (Table 1) at 4 °C until the blood is cleared from the tissues (5 - 10 min).
3. Change the perfusate to fixative solution (Table 1) at 4 °C until the neck and tail have significantly stiffened (5 - 10 min).
4. End the perfusion and decapitate the animal. Carefully dissect out the brain from the skull by first removing the connective tissue surrounding the skull and then removing bone from the ventral surface of the skull and lifting the brain out (Figure 1a), as described in ²⁰.
   Note: Give special care to the removal of bone surrounding the flocculonodular lobes of the cerebellum and the olfactory bulbs.
   1. Dissect the spinal cord from the vertebral column by gently inserting the tip of a fine, sharp scissor between the spinal cord and the vertebral column and cutting the individual vertebral column segments away, as described in ²¹.
5. Hemisect the brain by placing it in a plastic mouse brain matrix and cutting along the midline with a matrix blade.
6. Add each hemisphere and spinal cord to separate 50 ml centrifuge tubes with ~40 ml of hydrogel solution (Table 1). From this point forward cover the tubes with aluminum foil to protect the fluorophores during all subsequent steps.
7. Incubate the tube containing the sample in hydrogel at 4 °C with gentle shaking (~30 rpm) for 7 days.

2. Polymerization

1. Discard ~25 ml of hydrogel, preserving the sample and ~25 ml of hydrogel in the centrifuge tube.
2. Deoxygenate the sample by removing the cap and placing the centrifuge tube in desiccator jar and connecting the jar to a vacuum. Apply the vacuum for at least 10 min to permit dissolved gases to come out of solution and form bubbles. Gently shake to dislodge the bubbles.
3. Replace the vacuum in the desiccator jar with 100% nitrogen gas over 2 - 3 min. Once the pressure equalizes (once the top of the desiccator jar can be opened), quickly cap the tube to prevent the reintroduction of oxygen.
4. Polymerize the hydrogel by placing the tube with sample in a 37 °C water bath for 3 hr until polymerization is complete. Polymerized hydrogel will take on a gel-like consistency (Figure 1b). Note: A small amount of unpolymerized hydrogel at the top is acceptable.
5. Use a spatula to remove the polymerized sample from the centrifuge tube, then cut away the excess hydrogel from the sample.
6. Remove the remaining hydrogel by placing the sample on a lint free wipe and gently rubbing and rolling the tissue on it, leaving behind only a thin layer of polymerized hydrogel on the sample.

3. Lipid Removal

Note: Lipid-cleared tissue is fragile, handle with care. Use a cell strainer when draining the tube to avoid losing sample. Also, the sample will swell throughout the clearing process, however, swelling can be reversed during the refractive index matching process.

Caution: Sodium dodecyl sulfate (SDS) and boric acid are irritants. Perform experiments utilizing them in a fume hood and with appropriate personal protective equipment (lab coat, gloves and eye protection).

1. Transfer polymerized sample to a clean 50 ml centrifuge tube and add 45 ml of clearing buffer (Table 1). Incubate at 45 °C with gentle shaking (~30 rpm). Change the buffer daily for an initial 7 days by pouring off the used clearing buffer into chemical waste and adding ~45 ml of fresh clearing buffer. Keep the centrifuge tubes containing the samples covered in aluminum foil to protect the fluorophores.
2. After the initial 7 days, incubate the sample at 40 °C and exchange clearing buffer every 2 - 3 days. Continue clearing until all lipids are solubilized and tissue reaches transparency (Figure 1c). This will be 4 - 6 weeks for a mouse brain hemisphere and 2 - 4 weeks for a spinal cord.
3. Once tissue is cleared, transfer sample to a new 50 ml centrifuge tube and wash 4 times in PBST (Table 1) at 40 °C with gentle shaking (~30 rpm) over 24 hr to remove residual SDS.

4. Molecular Staining

Caution: 4',6-Diamidino-2-Phenylindole Dihydrochloride (DAPI) is an irritant and any experiments utilizing it should be performed in a fume hood and with appropriate personal protective equipment (lab coat, gloves, and eye protection).

1. Transfer the sample to a clean 15 ml centrifuge tube and fill with 1 µg/ml DAPI in 1x PBST, pH 7.5. Incubate at room temperature (RT) for 24 hr. Keep the centrifuge tubes containing the samples covered in aluminum foil to protect the fluorophores.
2. Wash 3 times in PBST at RT for at least 6 hr/wash.

5. Refractive Index Matching

Caution: 2,2′-Thiodiethanol (TDE) is an irritant and any experiments utilizing it should be performed in a fume hood and with appropriate personal protective equipment (lab coat, gloves, and eye protection).

1. Transfer the sample to a clean 15 ml centrifuge tube and fill with 30% (v/v) TDE in 1x PBS, pH 7.5. Incubate at 40 °C with gentle shaking (~30 rpm) for 24 hr.
2. Exchange 30% TDE with 63% (v/v) TDE in 1x PBS, pH 7.5. Incubate at 40 °C with gentle shaking for 24 hr.
   Note: The sample will shrink back to approximately original size.
3. On a flat, clean surface roll a piece of reusable adhesive into a cylinder with uniform diameter just slightly greater than the thickness of the sample.
4. Lay the reusable adhesive cylinder in an open-ended circle (~25 mm inner diameter with a small opening at the top) on a 40 mm glass bottom dish. If corrections need to be made, remove the cylinder from the glass and cut with a razor blade.
5. Use a p1000 pipette tip to create a seal between the reusable adhesive and glass by rolling it around the outer rim of the cylinder.
6. Place ~100 µl 63% TDE onto the glass dish and spread around to cover the glass surface within the circle of reusable adhesive.
7. Using a spatula, carefully place the sample in the middle of the circle, taking care not to form bubbles.
8. Pipette another 100 µl of 63% TDE directly onto the sample, then immediately place a 40 mm circular cover glass over the reusable adhesive.
9. Using the index, middle, and ring finger of both hands, carefully press around the perimeter of the circle until the cover glass has made contact with the sample.
10. Slowly bring the glass bottom dish to an upright position resting against a surface, then add 63% TDE with a pipette until the solution reaches the small opening at the top. Use a lint free wipe to dry off the pipette tip so that the solution does not drip on the glass. To avoid bubbles in the chamber, never empty the pipette all the way.
11. To the small opening, add silicon elastomer to enclose the circle and create a closed chamber. Wait 5 min for it to dry before imaging.

6. Microscopy

Caution: The lasers used in confocal, single plane illumination and light sheet microscopy are very powerful and can cause blindness. Any experiments utilizing lasers should be performed after extensive training and with great caution and appropriate eye protection.

1. Place the imaging chamber with the sample inside on the stage of a laser scanning confocal microscope.
2. Open the imaging software (see Table of Materials). To turn on the argon excitation laser, click the configuration tab at the top of the program then the laser icon. Check the box next to argon to power the laser and move the slide scale to 30%. Observe the pointer slowly warm up to the selected position.
   1. Return to the acquire tab at the top. In the large box "Beam Path Settings" go to the load/save setting box and select the drop down menu to select an appropriate wavelength channel to image YFP (527 nm).
3. Select appropriate objectives for imaging by locating the red hyperlink labeled "objective: current object" under the channel settings box. Clicking on the tab opens all available objectives and a selection will automatically rotate the turret.
   1. Use objectives (e.g., 10X) with a large working distance (greater than the thickness of the tissue to be imaged, e.g., 11 mm) and a large numerical aperture (e.g., 0.3) to maximize in-plane image resolution and minimize optical section thickness.
4. On the left column of drop down menus, open the "XY: resolution" window to set acquisition matrix, called "format", to 1,024 x 1,024 or a value appropriate for the lateral resolution limit of the objective. Set the zoom factor as low as possible (e.g., 1.7).
5. In the same window, set 8 line average and 1 frame average parameters by dropping down the individually labeled menus accordingly. Use large values for a high signal-to-noise ratio and small values for rapid acquisition.
   Note: 8 line averages and 1 frame average will acquire high-quality images of a mouse brain hemisphere in approximately 4 hr.
6. Locate the sample using the stage controls. Once a representative field is visible, set the gain and offset.
   Note: The gain and offset will vary substantially based on the tissue type, fluorophores, and/or anatomical feature being imaged. For YFP, set values from 760 to 780 for the gain and from -1.0 to -1.5 for the offset.
7. Select "tile scan" and select the boundaries of the region of interest. Set the upper and lower limits of the z-stack to be acquired. Set the thickness of the optical section based on the objective's optical section resolution limit.
   Note: An objective's optical section resolution limit is defined primarily by its numerical aperture. Most microscope control software will automatically calculate a value appropriate for the objective being used. For a 10X objective with a numerical aperture of 0.3 that would be approximately 11 µm.
8. Start the acquisition.
   Note: This may take multiple hours.
9. Collect higher resolution images using higher magnification objectives from specific regions of interest.

Results

Visualizing the structural organization of the brain is critical for understanding how morphology and connectivity affect brain function in health and disease. Optical clearing techniques make it possible to image cell populations in 3D in intact tissues, allowing us to study morphology and connectivity cohesively.

Mice that express fluorescent proteins driven by promoters specific for sub-populations of cells permit one to teas...

Discussion

Passive clearing of hydrogel embedded tissues (passive CLARITY) is a simple and inexpensive method for clearing large pieces of tissue. This approach does not require dedicated equipment and can easily be performed in a temperature-controlled shaker. Over the span of a few weeks, even large, highly myelinated tissues, such as an entire brain or spinal cord, will become transparent and suitable for microscopy. Even though this report has focused on the clearing of CNS tissues, passive CLARITY can be applied to any tissue....

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
10x phosphate buffered saline (PBS)	Fisher	BP399-1	buffers
32% paraformaldehyde (PFA)	Electron Microscopy Sciences	15714-5	perfusion and hydrogel
2,2'-thiodiethanol (TDE)	Sigma-Aldrich	166782-500G	refractive index matching solution
40% acrylamide	Bio-Rad	161-0140	hydrogel
2% bis-acrylamide	Bio-Rad	161-0142	hydrogel
VA-044 initiator	Wako Pure Chemical Industries, Ltd.	VA044	hydrogel
Boric acid	Sigma-Aldrich	B7901	clearing buffer
Sodium dodecyl sulfate (SDS)	Fisher	BP166-5	clearing buffer
Sodium hydroxide	Fisher	55255-1	buffers
Sodium azide	Sigma-Aldrich	52002-100G	preservative
Triton X-100	Sigma-Aldrich	X100-500G	buffers
Heparin	Sigma-Aldrich	H3393-50KU	perfusion
Sodium nitrate	Fisher	BP360-500	perfusion
DAPI	Molecular Probes	D1306	nuclear stain
99.9% isoflurane	Phoenix	57319-559-06	anesthetic
Hydrocholoric acid	Fisher	A1445-500	buffers
Glass bottom dish	Willco	HBSB-5040	Willco dishes
Reusable adhesive	Bostick	371351	Blu-Tack
Silicon elastomer	World Precision Instruments	KWIK-CAST	Kwik-Cast
Lint-free wipe	Kimberly-Clark	34120	KimWipe
Mouse brain matrix	Roboz	SA-2175	sectioning tissue
Matrix blades	Roboz	RS-9887	sectioning tissue
Peristaltic pump	Cole-Parmer	77122-24	pefusion
Laser scanning confocal microscope	Leica	SP5	microscopy
Imaging software	Leica	LAS AF	microscopy