Name: optical clearing of the mouse central nervous system using passive clarity
Uploaded: 2016-06-30T13:00:00.000+00:00
Duration: 10 min 28 s
Description: Watch this Scientific Journal Video about Optical Clearing of the Mouse Central Nervous System Using Passive CLARITY at JoVE.com

This work was generously supported by the National Institutes of Health/National Institute of Neurological Disorders and Stroke (NIH/NINDS) grant 1R01NS086981 and the Conrad N. Hilton Foundation. We thank Dr. Laurent Bentolila and Dr. Matthew Schibler for their invaluable assistance with confocal microscopy. We thank those that have contributed to the CLARITY forum (http://forum.claritytechniques.org). We especially thank Dr. Karl Deisseroth for opening up his lab to teach this fascinating technique. The authors are grateful for the generous support from the Brain Mapping Medical Research Organization, Brain Mapping Support Foundation, Pierson-Lovelace Foundation, The Ahmanson Foundation, Capital Group Companies Charitable Foundation, William M. and Linda R. Dietel Philanthropic Fund, and Northstar Fund. Research reported in this publication was also partially supported by the National Center for Research Resources and by the Office of the Director of the National Institutes of Health under award numbers C06RR012169, C06RR015431, and S10OD011939. The content is solely the responsibility of the authors and does not necessarily represent the official views of the National Institutes of Health.

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).
<ol>
	<li>Sacrifice the animal in euthanasia chamber with ~1 ml of isoflurane until breathing ceases (~2 - 3 min).</li>
	<li>Use a peristaltic pump set to ~5 ml/min to intracardially perfuse the mouse with perfusion solution (Table 1) at 4 &#176;C until the blood is cleared from the tissues (5 - 10 min).</li>
	<li>Change the perfusate to fixative solution (Table 1) at 4 &#176;C until the neck and tail have significantly stiffened (5 - 10 min).</li>
	<li>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.
	<ol>
		<li>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.</li>
	</ol>
	</li>
	<li>Hemisect the brain by placing it in a plastic mouse brain matrix and cutting along the midline with a matrix blade.</li>
	<li>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.</li>
	<li>Incubate the tube containing the sample in hydrogel at 4 &#176;C with gentle shaking (~30 rpm) for 7 days.</li>
</ol>
2. Polymerization
<ol>
	<li>Discard ~25&#160;ml of hydrogel, preserving the sample and ~25&#160;ml of hydrogel in the centrifuge tube.</li>
	<li>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.</li>
	<li>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.</li>
	<li>Polymerize the hydrogel by placing the tube with sample in a 37 &#176;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.</li>
	<li>Use a spatula to remove the polymerized sample from the centrifuge tube, then cut away the excess hydrogel from the sample.</li>
	<li>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.</li>
</ol>
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).
<ol>
	<li>Transfer polymerized sample to a clean 50 ml centrifuge tube and add 45 ml of clearing buffer (Table 1). Incubate at 45 &#176;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.</li>
	<li>After the initial 7 days, incubate the sample at 40 &#176;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.</li>
	<li>Once tissue is cleared, transfer sample to a new 50 ml centrifuge tube and wash 4 times in PBST (Table 1) at 40 &#176;C with gentle shaking (~30 rpm) over 24 hr to remove residual SDS.</li>
</ol>
4. Molecular Staining
Caution: 4&#39;,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).
<ol>
	<li>Transfer the sample to a clean 15 ml centrifuge tube and fill with 1 &#181;g/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.</li>
	<li>Wash 3 times in PBST at RT for at least 6 hr/wash.</li>
</ol>
5. Refractive Index Matching
Caution: 2,2&#8242;-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).
<ol>
	<li>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 &#176;C with gentle shaking (~30 rpm) for 24 hr.</li>
	<li>Exchange 30% TDE with 63% (v/v) TDE in 1x PBS, pH 7.5. Incubate at 40 &#176;C with gentle shaking for 24 hr. 
	Note: The sample will shrink back to approximately original size.</li>
	<li>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.</li>
	<li>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.</li>
	<li>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.</li>
	<li>Place ~100 &#181;l 63% TDE onto the glass dish and spread around to cover the glass surface within the circle of reusable adhesive.</li>
	<li>Using a spatula, carefully place the sample in the middle of the circle, taking care not to form bubbles.</li>
	<li>Pipette another 100 &#181;l of 63% TDE directly onto the sample, then immediately place a 40 mm circular cover glass over the reusable adhesive.</li>
	<li>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.</li>
	<li>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.</li>
	<li>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.</li>
</ol>
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.
<ol>
	<li>Place the imaging chamber with the sample inside on the stage of a laser scanning confocal microscope.</li>
	<li>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.
	<ol>
		<li>Return to the acquire tab at the top. In the large box &#34;Beam Path Settings&#34; go to the load/save setting box and select the drop down menu to select an appropriate wavelength channel to image YFP (527 nm).</li>
	</ol>
	</li>
	<li>Select appropriate objectives for imaging by locating the red hyperlink labeled &#34;objective: current object&#34; under the channel settings box. Clicking on the tab opens all available objectives and a selection will automatically rotate the turret.
	<ol>
		<li>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.</li>
	</ol>
	</li>
	<li>On the left column of drop down menus, open the &#34;XY: resolution&#34; window to set acquisition matrix, called &#34;format&#34;, 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).</li>
	<li>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.</li>
	<li>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.</li>
	<li>Select &#34;tile scan&#34; 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&#39;s optical section resolution limit. 
	Note: An objective&#39;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 &#181;m.</li>
	<li>Start the acquisition. 
	Note: This may take multiple hours.</li>
	<li>Collect higher resolution images using higher magnification objectives from specific regions of interest.</li>
</ol>

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The authors have nothing to disclose.

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.
The relative strengths and weaknesses of optical clearing techniques have been discussed previously in the literature13,17,22,, but are most clearly laid out in 23. Passive CLARITY is well suited for experiments where large numbers of samples are being cleared simultaneously and reproducibility is of great importance. The use of a hydrogel to provide additional support to the tissue facilitates multiple probing of the tissue. In this report, a small molecular weight dye, DAPI, was demonstrated, but multiple rounds of antibody probing are also possible with this approach.
The approach described above deviates from our previously published protocol15 in a couple of significant ways. Firstly, in the current approach, the specimens were not intracardially perfused with hydrogel. This was due to the fact that the hydrogel can prematurely polymerize in the tubing and needles of the perfusion apparatus, leading to incomplete perfusions and inconsistent clearing. Incubating paraformaldehyde fixed tissues in hydrogel for 7 days is consistent with the original CLARITY protocol14, though a 1 day incubation is used for 1 - 3 mm thick tissue sections in the PAssive CLARITY Technique (PACT)17. Secondly, this approach performs lipid removal at a slightly higher temperature that previously reported15-17. Higher temperature accelerates lipid removal, but must be balanced against the possibility of a catastrophic loss of tissue integrity (melting). By increasing the incubation temperature by 8 &#176;C (to 45 &#176;C) for 7 days and then by 3 &#176;C (to 40 &#176;C) for the remainder of the lipid removal process, tissue clearing is accelerated, but without the concurrent risk of the loss of tissue integrity. This increase in temperature is consistent with temperatures reported during Electrophoretic Tissue Clearing (ETC) in the original CLARITY protocol (50 &#176;C)14. Furthermore, no decrease in the intensity of transgenically expressed fluorophores nor decrease in DAPI staining has been observed.
Deoxygenation is an important step in the creation of the hydrogel. The hydrogel may polymerize in the presence of oxygen dissolved in the hydrogel solution, but in our hands the rate and completeness of polymerization was much more variable without deoxygenation. If the hydrogel does not polymerize, the most common reason is the presence of oxygen in the hydrogel.
The optical clearing process is remarkably easy to implement, however, the microscopy that follows may be quite complicated. Of critical importance are the objectives used to image the tissue. The working distance of an objective defines the depth that it can image into a tissue sample. An objective with a working distance of 2 mm can only image 2 mm into a sample, therefore the working distance of an objective must be greater than the thickness of the sample in order to image it completely. Similarly, the numerical aperture of an objective places a limit the size of features that it can resolve both in-plane and the thickness of optical sections that it can meaningfully acquire. Selective Plane Illumination Microscopy (SPIM) and Light Sheet Microscopy (LSM) can overcome the limitation on optical section thickness by the nature of the illumination, but these microscopes are not widely available. High numerical aperture, large working distance objectives are rare and expensive, but are necessary for high quality imaging from large optically cleared sections.
Once the images have been acquired, they are often very large, ranging from several hundred MB for a low resolution (both in-plane and the z-direction) image to a few TB for a high resolution image collected on an LSM. The computers used for image analysis require not only a great deal of storage, but large amounts of random access memory (RAM) for image processing. Software packages suitable of the analysis of large optically cleared image volumes include Amira, Imaris, and ImageJ14-16.
Among the limitations of this approach, passive CLARITY requires more time to clear tissues than other optical clearing approaches. Decreasing the volume of the tissue to be cleared by trimming it or sectioning it into smaller pieces can accelerate lipid-removal. Also, decreasing the concentration of paraformaldehyde in the hydrogel will in turn decrease crosslinking and increase the hydrogel pore size, leading to more rapid clearing17, though this may lead to greater tissue expansion, individual experimenters may find that it is worth the decrease in clearing time. Further, combined with TDE as a clearing agent, the amount of overall tissue expansion may be limited.
Though this report describes the use of small molecular weight dyes, antibody probing is a very natural next step. The presence of the hydrogel not only provides physical support to the cleared tissues, but also serves as an impediment for the penetration of antibodies deep into the tissue. Decreasing the acrylamide concentration or the paraformaldehyde concentration will increase the pore size in the hydrogel, facilitating (and thus accelerating) antibody penetration17, but this has a deleterious effect on tissue expansion. Nevertheless, antibody probing of blocks of tissue can be accomplished in a matter of days and whole brain in a couple of weeks using the acrylamide and paraformaldehyde concentrations described in the original report14.
Optical clearing techniques allow one to look deeply into the brain, furthering the understanding of its structure and function. These new tools will enable researchers to visualize the brain at cellular and molecular resolutions, increasing our comprehension of this most complex of organs.

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 tissue1-3. 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 &#181;m in laser scanning confocal microscopy and to less than 800 &#181;m using two-photon excitation microscopy4-8.
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 times15-17. 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 developed17-19. 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.

<table><tbody><tr><td>10x phosphate buffered saline (PBS)</td><td>Fisher</td><td>BP399-1</td><td>buffers</td></tr><tr><td>32% paraformaldehyde (PFA)</td><td>Electron Microscopy Sciences</td><td>15714-5</td><td>perfusion and hydrogel</td></tr><tr><td>2,2&#039;-thiodiethanol (TDE)</td><td>Sigma-Aldrich</td><td>166782-500G</td><td>refractive index matching solution</td></tr><tr><td>40% acrylamide</td><td>Bio-Rad</td><td>161-0140</td><td>hydrogel</td></tr><tr><td>2% bis-acrylamide</td><td>Bio-Rad</td><td>161-0142</td><td>hydrogel</td></tr><tr><td>VA-044 initiator</td><td>Wako Pure Chemical Industries, Ltd.</td><td>VA044</td><td>hydrogel</td></tr><tr><td>Boric acid</td><td>Sigma-Aldrich</td><td>B7901</td><td>clearing buffer</td></tr><tr><td>Sodium dodecyl sulfate (SDS)</td><td>Fisher</td><td>BP166-5</td><td>clearing buffer</td></tr><tr><td>Sodium hydroxide</td><td>Fisher</td><td>55255-1</td><td>buffers</td></tr><tr><td>Sodium azide</td><td>Sigma-Aldrich</td><td>52002-100G</td><td>preservative</td></tr><tr><td>Triton X-100</td><td>Sigma-Aldrich</td><td>X100-500G</td><td>buffers</td></tr><tr><td>Heparin</td><td>Sigma-Aldrich</td><td>H3393-50KU</td><td>perfusion</td></tr><tr><td>Sodium nitrate</td><td>Fisher</td><td>BP360-500</td><td>perfusion</td></tr><tr><td>DAPI</td><td>Molecular Probes</td><td>D1306</td><td>nuclear stain</td></tr><tr><td>99.9% isoflurane</td><td>Phoenix</td><td>57319-559-06</td><td>anesthetic</td></tr><tr><td>Hydrocholoric acid</td><td>Fisher</td><td>A1445-500</td><td>buffers</td></tr><tr><td>Glass bottom dish</td><td>Willco</td><td>HBSB-5040</td><td>Willco dishes</td></tr><tr><td>Reusable adhesive</td><td>Bostick</td><td>371351</td><td>Blu-Tack</td></tr><tr><td>Silicon elastomer</td><td>World Precision Instruments</td><td>KWIK-CAST</td><td>Kwik-Cast</td></tr><tr><td>Lint-free wipe</td><td>Kimberly-Clark</td><td>34120</td><td>KimWipe</td></tr><tr><td>Mouse brain matrix</td><td>Roboz</td><td>SA-2175</td><td>sectioning tissue</td></tr><tr><td>Matrix blades</td><td>Roboz</td><td>RS-9887</td><td>sectioning tissue</td></tr><tr><td>Peristaltic pump</td><td>Cole-Parmer</td><td>77122-24</td><td>pefusion</td></tr><tr><td>Laser scanning confocal microscope</td><td>Leica</td><td>SP5</td><td>microscopy</td></tr><tr><td>Imaging software</td><td>Leica</td><td>LAS AF</td><td>microscopy</td></tr></tbody></table>

optical clearing of the mouse central nervous system using passive clarity

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&#8242;-thiodiethanol (TDE), a previously described refractive index matching solution.

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 tease apart the complex pattern of neural connectivity in the brain. THY1-YFP transgenic mice have been used extensively in the optical clearing literature because these mice express YFP in a subset of projection neurons that reside in the cerebral cortex and hippocampus, as well as other structures in the brain (Figure 2a &#38; b). Furthermore, YFP+ cortical layer V neurons project through the corticospinal tract in the spinal cord (Figure 2c &#38; d), making them very useful in evaluating the effect of axonal insult and/or injury in the spinal cord on cortical neurons15. Through the use of confocal microscopy and optical sectioning of optically cleared tissues one can easily evaluate differences in neuronal density across large anatomical structures.
In addition to THY1-YFP transgenic mice, any number of transgenic mice can be imaged. PLP-EGFP transgenic mice express enhanced green fluorescent protein (EGFP) in mature oligodendrocytes expressing proteolipid protein (PLP) throughout the brain and spinal cord (Figure 3a). PV-TdTomato transgenic mice express TdTomato in a subset of parvalbumin (PV) expressing interneurons in the cerebellum, striatum and hippocampus (Figure 3b). Even in mice that do not express fluorescent transgenes, small molecular weight fluorescent dyes, such as DAPI, can readily diffuse into hydrogel embedded tissues and permit the evaluation of cellular nuclei in the brain (Figure 4).
<img alt="Figure 1" src="/files/ftp_upload/54025/54025fig1.jpg" /> 
Figure 1: Different Stages of Brain Clearing. Photograph of THY1-YFP+ mouse brain that has been removed from the skull (a). Photograph of hydrogel-embedded THY1-YFP+ mouse brain hemisphere (b). Photograph of THY1-YFP+ mouse brain hemisphere after lipid removal (c). Scale bars are 5 mm in each panel. <a href="https://www-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/54025/54025fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" src="/files/ftp_upload/54025/54025fig2.jpg" /> 
Figure 2: THY1-YFP Expression in the Brain and Spinal Cord. THY1-YFP+ neurons in the cerebral cortex and hippocampus imaged at 5X magnification and reconstructed in 3D (a). A maximum intensity projection of a 10X image stack of the cerebral cortex demonstrating the dendritic arborization of cortical layer V pyramidal neurons (b). THY1-YFP expression in an intact cervical and thoracic spinal cord imaged at 5X magnification and reconstructed in 3D (c). A maximum intensity projection of a 10X image stack of the spinal cord demonstrating individual axons (d). Scale bars are 1 mm in (a), 100 &#181;m in (b), 2 mm in (c), and 100 &#181;m in (d). <a href="https://www-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/54025/54025fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" src="/files/ftp_upload/54025/54025fig3.jpg" /> 
Figure 3: PLP-EGFP and PV-TdTomato Expression in the Brain. PLP-EGFP expression in a 2 mm thick coronal brain section imaged at 5X magnification (a). This image is approximately 1 mm from the surface of the tissue section. PV-TdTomato expression in the cerebellum of an intact cerebral hemisphere imaged at 10X magnification (b). This image is approximately 0.5 mm from the midsagittal plane. Scale bars are 1 mm in (a) and 250 &#181;m in (b). <a href="https://www-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/54025/54025fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" src="/files/ftp_upload/54025/54025fig4.jpg" /> 
Figure 4: DAPI Staining in the Brain. DAPI expression in the cerebellum of an intact cerebral hemisphere imaged at 5X magnification. The image is approximately 1 mm from the midsagittal plane. The inset shows the level of detail visible at this imaging depth at 10X magnification. Scale bars are 250 &#181;m in the main panel and 50 &#181;m in the inset. <a href="https://www-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/54025/54025fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Buffer</td>
			<td>Chemical</td>
			<td>Final concentration</td>
		</tr>
		<tr>
			<td>Perfusion solution&#160;</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td></td>
			<td>10x phosphate buffered saline</td>
			<td>1x</td>
		</tr>
		<tr>
			<td></td>
			<td>Sodium nitrate</td>
			<td>0.5% w/v</td>
		</tr>
		<tr>
			<td></td>
			<td>Heparin</td>
			<td>10 U/ml</td>
		</tr>
		<tr>
			<td>Fixative solution&#160;</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td></td>
			<td>10x phosphate buffered saline</td>
			<td>1x</td>
		</tr>
		<tr>
			<td></td>
			<td>Paraformaldehyde</td>
			<td>4% v/v</td>
		</tr>
		<tr>
			<td>Hydrogel solution&#160;</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td></td>
			<td>10x phosphate buffered saline</td>
			<td>1x</td>
		</tr>
		<tr>
			<td></td>
			<td>Paraformaldehyde</td>
			<td>4% v/v</td>
		</tr>
		<tr>
			<td></td>
			<td>Acrylamide</td>
			<td>4% v/v</td>
		</tr>
		<tr>
			<td></td>
			<td>Bis-acrylamide</td>
			<td>0.05% v/v</td>
		</tr>
		<tr>
			<td></td>
			<td>VA-044 Initiator</td>
			<td>0.25% w/v</td>
		</tr>
		<tr>
			<td>Clearing buffer&#160;</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td></td>
			<td>Boric acid</td>
			<td>200 mM</td>
		</tr>
		<tr>
			<td></td>
			<td>Sodium dodecyl sulfate</td>
			<td>4% w/v</td>
		</tr>
		<tr>
			<td>PBST&#160;</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td></td>
			<td>10x phosphate buffered saline</td>
			<td>1x</td>
		</tr>
		<tr>
			<td></td>
			<td>Triton X-100</td>
			<td>0.1% v/v</td>
		</tr>
		<tr>
			<td></td>
			<td>Sodium azide</td>
			<td>0.1% w/v</td>
		</tr>
	</tbody>
</table>
Table 1: List of Buffers and Solutions.

Watch this Scientific Journal Video about Optical Clearing of the Mouse Central Nervous System Using Passive CLARITY at JoVE.com

Optical Clearing of the Mouse Central Nervous System Using Passive CLARITY

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.

Traditionally, tissue visualization has required that the tissue of interest be serially sectioned and imaged, subjecting each tissue section to unique ...

optical-clearing-mouse-central-nervous-system-using-passive

Universidad San Sebastian

Research

JoVE Journal

Neuroscience

13.4K Views.  University of California, Los Angeles. 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.

Video: Optical Clearing of the Mouse Central Nervous System Using Passive CLARITY

Multiphoton Microscopy of Cleared Mouse Brain Expressing YFP

Multiphoton microscopy of intrinsic fluorescence and second harmonic generation (SHG) of whole mouse organs is made possible by optically clearing the organ before imaging.1,2 However, for organs that contain fluorescent proteins such as GFP and YFP, optical clearing protocols that use methanol dehydration and clear using benzyl alcohol:benzyl benzoate (BABB) while unprotected from light3 do not preserve the fluorescent signal. The protocol presented here is a novel way in which to perform whole organ optical clearing on mouse brain while preserving the fluorescence signal of YFP expressed in neurons. Altering the optical clearing protocol such that the organ is dehydrated using an ethanol graded series has been found to reduce the damage to the fluorescent proteins and preserve their fluorescent signal for multiphoton imaging.4 Using an optimized method of optical clearing with ethanol-based dehydration and clearing by BABB while shielded from light, we show high-resolution multiphoton images of yellow fluorescent protein (YFP) expression in the neurons of a mouse brain more than 2 mm beneath the tissue surface.

Multiphoton microscopy of whole mouse organs is possible by optically clearing the organ before imaging, but not all protocols preserve the fluorescent signal of fluorescent proteins. Using an optical clearing method with ethanol-based dehydration and benzyl alcohol:benzyl benzoate clearing, we show high-resolution multiphoton images of whole mouse brain expressing YFP.

Multiphoton microscopy of intrinsic fluorescence and second harmonic generation (SHG) of whole mouse organs is made possible by optically clearing the ...

Micron-scale Resolution Optical Tomography of Entire Mouse Brains with Confocal Light Sheet Microscopy

Understanding the architecture of mammalian brain at single-cell resolution is one of the key issues of neuroscience. However, mapping neuronal soma and projections throughout the whole brain is still challenging for imaging and data management technologies. Indeed, macroscopic volumes need to be reconstructed with high resolution and contrast in a reasonable time, producing datasets in the TeraByte range. We recently demonstrated an optical method (confocal light sheet microscopy, CLSM) capable of obtaining micron-scale reconstruction of entire mouse brains labeled with enhanced green fluorescent protein (EGFP). Combining light sheet illumination and confocal detection, CLSM allows deep imaging inside macroscopic cleared specimens with high contrast and speed. Here we describe the complete experimental pipeline to obtain comprehensive and human-readable images of entire mouse brains labeled with fluorescent proteins. The clearing and the mounting procedures are described, together with the steps to perform an optical tomography on its whole volume by acquiring many parallel adjacent stacks. We showed the usage of open-source custom-made software tools enabling stitching of the multiple stacks and multi-resolution data navigation. Finally, we illustrated some example of brain maps: the cerebellum from an L7-GFP transgenic mouse, in which all Purkinje cells are selectively labeled, and the whole brain from a thy1-GFP-M mouse, characterized by a random sparse neuronal labeling.

In this article we describe the full experimental procedure to reconstruct, with high resolution, the fine brain anatomy of fluorescently labeled mouse brains. The described protocol includes sample preparation and clearing, specimen mounting for imaging, data post-processing and multi-scale visualization.

Understanding the architecture of mammalian  brain at single-cell resolution is one of the key issues of neuroscience.  However, mapping neuronal soma and ...

Imaging Serotonergic Fibers in the Mouse Spinal Cord Using the CLARITY/CUBIC Technique

Long descending fibers to the spinal cord are essential for locomotion, pain perception, and other behaviors. The fiber termination pattern in the spinal cord of the majority of these fiber systems have not been thoroughly investigated in any species. Serotonergic fibers, which project to the spinal cord, have been studied in rats and opossums on histological sections and their functional significance has been deduced based on their fiber termination pattern in the spinal cord. With the development of CLARITY and CUBIC techniques, it is possible to investigate this fiber system and its distribution in the spinal cord, which is likely to reveal previously unknown features of serotonergic supraspinal pathways. Here, we provide a detailed protocol for imaging the serotonergic fibers in the mouse spinal cord using the combined CLARITY and CUBIC techniques. The method involves perfusion of a mouse with a hydrogel solution and clarification of the tissue with a combination of clearing reagents. Spinal cord tissue was cleared in just under two weeks, and the subsequent immunofluorescent staining against serotonin was completed in less than ten days. With a multi-photon fluorescent microscope, the tissue was scanned and a 3D image was reconstructed using Osirix software.

Supraspinal projections are important for pain perception and other behaviors, and serotonergic fibers are one of these fiber systems. The present study focused on the application of the combined CLARITY/CUBIC protocol to the mouse spinal cord in order to investigate the termination of these serotonergic fibers.

Long descending fibers to the spinal cord are essential for locomotion, pain perception, and other behaviors. The fiber termination pattern in the spinal ...

Induction of Paralysis and Visual System Injury in Mice by T Cells Specific for Neuromyelitis Optica Autoantigen Aquaporin-4

While it is recognized that aquaporin-4 (AQP4)-specific T cells and antibodies participate in the pathogenesis of neuromyelitis optica (NMO), a human central nervous system (CNS) autoimmune demyelinating disease, creation of an AQP4-targeted model with both clinical and histologic manifestations of CNS autoimmunity has proven challenging. Immunization of wild-type (WT) mice with AQP4 peptides elicited T cell proliferation, although those T cells could not transfer disease to na&#239;ve recipient mice. Recently, two novel AQP4 T cell epitopes, peptide (p) 135-153 and p201-220, were identified when studying immune responses to AQP4 in AQP4-deficient (AQP4-/-) mice, suggesting T cell reactivity to these epitopes is normally controlled by thymic negative selection. AQP4-/- Th17 polarized T cells primed to either p135-153 or p201-220 induced paralysis in recipient WT mice, that was associated with predominantly leptomeningeal inflammation of the spinal cord and optic nerves. Inflammation surrounding optic nerves and involvement of the inner retinal layers (IRL) were manifested by changes in serial optical coherence tomography (OCT). Here, we illustrate the approaches used to create this new in vivo model of AQP4-targeted CNS autoimmunity (ATCA), which can now be employed to study mechanisms that permit development of pathogenic AQP4-specific T cells and how they may cooperate with B cells in NMO pathogenesis.

Here, we present a protocol to induce paralysis and opticospinal inflammation by transfer of aquaporin-4 (AQP4)-specific T cells from AQP4-/- mice into WT mice. In addition, we demonstrate how to use serial optical coherence tomography to monitor visual system dysfunction.

While it is recognized that aquaporin-4 (AQP4)-specific T cells and antibodies participate in the pathogenesis of neuromyelitis optica (NMO), a human ...

Immunology and Infection

Novel Passive Clearing Methods for the Rapid Production of Optical Transparency in Whole CNS Tissue

Since the development of CLARITY, a bioelectrochemical clearing technique that allows for three-dimensional phenotype mapping within transparent tissues, a multitude of novel clearing methodologies including CUBIC (clear, unobstructed brain imaging cocktails and computational analysis), SWITCH (system-wide control of interaction time and kinetics of chemicals), MAP (magnified analysis of the proteome), and PACT (passive clarity technique), have been established to further expand the existing toolkit for the microscopic analysis of biological tissues. The present study aims to improve upon and optimize the original PACT procedure for an array of intact rodent tissues, including the whole central nervous system (CNS), kidneys, spleen, and whole mouse embryos. Termed psPACT (process-separate PACT) and mPACT (modified PACT), these novel techniques provide highly efficacious means of mapping cell circuitry and visualizing subcellular structures in intact normal and pathological tissues. In the following protocol, we provide a detailed, step-by-step outline on how to achieve maximal tissue clearance with minimal invasion of their structural integrity via psPACT and mPACT.

Here, we present two novel methodologies, psPACT and mPACT, for achieving maximal optical transparency and subsequent microscopic analysis of tissue vasculature in the intact rodent whole CNS.

Since the development of CLARITY, a bioelectrochemical clearing technique that allows for three-dimensional phenotype mapping within transparent tissues, ...

Immunostaining of Whole-Mount Retinas with the CLARITY Tissue Clearing Method

The tissue hydrogel delipidation method (CLARITY), originally developed by the Deisseroth laboratory, has been modified and widely used for immunostaining and imaging of thick brain samples. However, this advanced technology has not yet been used for whole-mount retinas. Although the retina is partially transparent, its thickness of approximately 200 &#181;m (in mice) still limits the penetration of antibodies into the deep tissue as well as reducing light penetration for high-resolution imaging. Here, we adapted the CLARITY method for whole-mount mouse retinas by polymerizing them with an acrylamide monomer to form a nanoporous hydrogel and then clearing them in sodium dodecyl sulfate to minimize protein loss and avoid tissue damage. CLARITY-processed retinas were immunostained with antibodies for retinal neurons, glial cells, and synaptic proteins, mounted in a refractive index matching solution, and imaged. Our data demonstrate that CLARITY&#160;can improve the quality of standard immunohistochemical staining and imaging for retinal neurons and glial cells in whole-mount preparation. For instance, 3D resolution of fine axon-like and dendritic structures of dopaminergic amacrine cells were much improved by CLARITY. Compared to non-processed whole-mount retinas, CLARITY can reveal immunostaining for synaptic proteins such as postsynaptic density protein 95. Our results show that CLARITY renders the retina more optically transparent after the removal of lipids and preserves fine structures of retinal neurons and their proteins, which can be routinely used for obtaining high-resolution imaging of retinal neurons and their subcellular structures in whole-mount preparation.

Here we present a protocol to adapt the CLARITY method of the brain tissues for whole-mount retinas to improve the quality of standard immunohistochemical staining and high-resolution imaging of retinal neurons and their subcellular structures.

The tissue hydrogel delipidation method (CLARITY), originally developed by the Deisseroth laboratory, has been modified and widely used for immunostaining ...

Imaging and Quantification of Intact Neuronal Dendrites via CLARITY Tissue Clearing

Brain activity, the electrochemical signals passed between neurons, is determined by the connectivity patterns of neuronal networks, and from the morphology of processes and substructures within these neurons. As such, much of what is known about brain function has arisen alongside developments in imaging technologies that allow further insight into how neurons are organized and connected in the brain. Improvements in tissue clearing have allowed for high-resolution imaging of thick brain slices, facilitating morphological reconstruction and analyses of neuronal substructures, such as dendritic arbors and spines. In tandem, advances in image processing software provide methods of quickly analyzing large imaging datasets. This work presents a relatively rapid method of processing, visualizing, and analyzing thick slices of labeled neural tissue at high-resolution using CLARITY tissue clearing, confocal microscopy, and image analysis. This protocol will facilitate efforts toward understanding the connectivity patterns and neuronal morphologies that characterize healthy brains, and the changes in these characteristics that arise in diseased brain states.

Neuronal dendritic morphology often underlies function. Indeed, many disease processes that affect the development of neurons manifest with a morphological phenotype. This protocol describes a simple and powerful method for analyzing intact dendritic arbors and their associated spines.

Brain activity, the electrochemical signals passed between neurons, is determined by the connectivity patterns of neuronal networks, and from the ...

A Tissue Clearing Method for Neuronal Imaging from Mesoscopic to Microscopic Scales

A detailed protocol is provided here to visualize neuronal structures from mesoscopic to microscopic levels in brain tissues. Neuronal structures ranging from neural circuits to subcellular neuronal structures are visualized in mouse brain slices optically cleared with ScaleSF. This clearing method is a modified version of ScaleS and is a hydrophilic tissue clearing method for tissue slices that achieves potent clearing capability as well as a high-level of preservation of fluorescence signals and structural integrity. A customizable three dimensional (3D)-printed imaging chamber is designed for reliable mounting of cleared brain tissues. Mouse brains injected with an adeno-associated virus vector carrying enhanced green fluorescent protein gene were fixed with 4% paraformaldehyde and cut into slices of 1-mm thickness with a vibrating tissue slicer. The brain slices were cleared by following the clearing protocol, which include sequential incubations in three solutions, namely, ScaleS0 solution, phosphate buffer saline (&#8211;), and ScaleS4 solution, for a total of 10.5&#8211;14.5 h. The cleared brain slices were mounted on the imaging chamber and embedded in 1.5% agarose gel dissolved in ScaleS4D25(0) solution. The 3D image acquisition of the slices was carried out using a confocal laser scanning microscope equipped with a multi-immersion objective lens of a long working distance. Beginning with mesoscopic neuronal imaging, we succeeded in visualizing fine subcellular neuronal structures, such as dendritic spines and axonal boutons, in the optically cleared brain slices. This protocol would facilitate understanding of neuronal structures from circuit to subcellular component scales.

The protocol provides a detailed method of neuronal imaging in brain slice using a tissue clearing method, ScaleSF. The protocol includes brain tissue preparation, tissue clarification, handling of cleared slices and confocal laser scanning microscopy imaging of neuronal structures from mesoscopic to microscopic levels.

A detailed protocol is provided here to visualize neuronal structures from mesoscopic to microscopic levels in brain tissues. Neuronal structures ranging ...

Three-dimensional Reconstruction of the Vascular Architecture of the Passive CLARITY-cleared Mouse Ovary

The ovary is the main organ of the female reproductive system and is essential for the production of female gametes and for controlling the endocrine system, but the complex structural relationships and three-dimensional (3D) vasculature architectures of the ovary are not well described. In order to visualize the 3D connections and architecture of blood vessels in the intact ovary, the first important step is to make the ovary optically clear. In order to avoid tissue shrinkage, we used the hydrogel fixation-based passive CLARITY (Clear Lipid-exchanged Acrylamide-hybridized Rigid Imaging/ Immunostaining/In situ-hybridization-compatible Tissue Hydrogel) protocol method to clear an intact ovary. Immunostaining, advanced multiphoton confocal microscopy, and 3D image-reconstructions were then used for the visualization of ovarian vessels and follicular capillaries. Using this approach, we showed a significant positive correlation (P &#60;0.01) between the length of the follicular capillaries and volume of the follicular wall.

Here we present an adaptation of the passive CLARITY and 3D reconstruction method for visualization of the ovarian vasculature and follicular capillaries in intact mouse ovaries.

The ovary is the main organ of the female reproductive system and is essential for the production of female gametes and for controlling the endocrine ...

Biology

Imaging the Gut with "CLARITY"

CLARITY (Clear Lipid-exchanged Acrylamide-hybridized Rigid Imaging compatible Tissue hYdrogel) has recently evolved as a valuable technique involving acrylamide embedding to delipidate tissue (without sectioning) and to preserve the 3-D tissue structure for immunostaining. The technique is highly relevant in imaging the dynamic gut environment where different cell types interact during homeostasis and disease states. This method optimized for the mouse gut is described here, which helps to trace cell types like epithelia, enteroendocrine, neurons, glia, and the neuronal projections into the epithelia or enteroendocrine cells that mediate microbial sensing or nutrient chemo sensing respectively. The gut tissue (1-1.5 cm) is fixed in 4% paraformaldehyde (PFA) in phosphate buffered saline (PBS) at 4 &#176;C overnight on day 1. On day 2, PFA is discarded, and the tissue is washed thrice with PBS. The tissue is hydrogel embedded to preserve its integrity by incubation in 4% hydrogel (acrylamide) solution in PBS (diluted from 30% ProtoGel) overnight at 4 &#176;C. On day 3, the tissue-hydrogel solution is incubated at 37&#160;&#176;C for 1 h to allow hydrogel polymerization. Tissue is then washed thrice gently with PBS to remove excess hydrogel. The subsequent step of delipidation (clearing) involves tissue incubation in sodium dodecyl sulfate (8% SDS in PBS) at 37 &#176;C for 2 days (days 4 &#38; 5) on a shaker at room temperature (RT). On day 6, the cleared tissue is thoroughly washed with PBS to remove SDS. Tissue can be immunostained by incubation in primary antibodies (diluted in 0.5% normal donkey serum in PBS containing 0.3% Triton X-100), overnight at 4&#176;C, and subsequent incubation in appropriate secondary Alexa Fluor antibodies for 1.5 h at RT, and nuclear staining with DAPI (1: 10000). The tissue is transferred onto a clean glass slide and mounted using VectaShield for confocal imaging.

We describe the protocol for passive CLARITY (PACT) staining of mouse intestine to enable visualization of subepithelial tissues, including neurons, glia and enteroendocrine cells (EEC) without tissue sectioning. The protocol involves hydrogel embedding of formaldehyde-fixed tissue, and subsequent delipidation using an anionic detergent to "clear" the tissue.

CLARITY (Clear Lipid-exchanged Acrylamide-hybridized Rigid Imaging compatible Tissue hYdrogel) has recently evolved as a valuable technique involving ...