Name: imaging serotonergic fibers in the mouse spinal cord using the clarity/cubic technique
Uploaded: 2016-02-26T14:00:00.000+00:00
Duration: 9 min 54 s
Description: Watch this Scientific Journal Video about Imaging Serotonergic Fibers in the Mouse Spinal Cord Using the CLARITY/CUBIC Technique at JoVE.com

This work was supported by the Australian Research Council Centre of Excellence for Integrative Brain Function (ARC Centre Grant CE140100007), an NHMRC project grant (#1086643). Prof. George Paxinos is supported by a Senior Principal Research Fellow NHMRC grant (#1043626).

Ethics Statement: All procedures involving animal subjects follow the guidelines of the Animal Care and Ethics Committee (ACEC) at The University of New South Wales (the approved ACEC number is 14/94A).
1. Preparation of the Transparent Mouse Spinal Cord
<ol>
	<li>Preparation of Ice Cold Hydrogel Solution
	<ol>
		<li>Preparation of 16% paraformaldehyde solution (PFA)
		<ol>
			<li>Add 16 g paraformaldehyde powder into 70 ml pre-warmed distilled water (50-55 &#176;C) and stir on a heated magnetic stirrer until paraformaldehyde is dissolved. Note: Do not allow the solution to heat over 55&#160;&#176;C and be aware that paraformaldehyde is toxic.</li>
			<li>Transfer the paraformaldehyde solution to a cylinder and add distilled water to 100 ml. Cool the paraformaldehyde solution using a 4&#160;&#176;C refrigerator.</li>
		</ol>
		</li>
		<li>Dissolve 125 mg VA-044 initiator in 26.25 ml distilled water and cool the solution in a 4&#160;&#176;C fridge.</li>
		<li>Cool 5 ml acrylamide solution (40%), 1.25 ml Bis solution (2%) (CAUTION: Bis and acrylamide solutions are toxic, may cause genetic defects or cancer), 5 ml 10x PBS, 12.5 ml 16% PFA solution, and the VA-044 initiator solution on ice.</li>
		<li>Mix the above solutions on ice. 
		Note: the total volume is 50 ml.</li>
	</ol>
	</li>
	<li>Perfusion and Collection of the Mouse Spinal Cord
	<ol>
		<li>Anaesthetize a mouse with an intraperitoneal injection of ketamine (80 mg/kg) and xyzaline (5 mg/kg) diluted in 0.9% normal saline.&#160;&#160;The dosage for mouse can be lower than that for rats.</li>
		<li>On a suitable plastic surface in a fume hood, fix the limbs of the mouse away from the body (sticky tape is effective) once the mouse does not respond to a pinch to its skin of the foot.</li>
		<li>Cut the mouse skin over the chest and then the bone chest to expose the heart with a scissor.</li>
		<li>Using a peristaltic pump, with a 25 G needle attached to the end of the tubing, insert the needle into the left ventricle of the heart, snip the right atrium to create an exit point for the blood and then start washing with 40 ml 0.9% normal saline a rate of approximately 10 ml/min.</li>
		<li>When complete, perfuse the mouse with 35 ml ice cold hydrogel solution. 
		Note: perfuse at a speed of 10 ml/min to avoid swelling of the nervous tissue.</li>
		<li>Incise the skin over the back with a blade and then remove the muscles next to the vertebrate. After cutting the vertebral arches on one side and flipping them to the other side, take the spinal cord out of the vertebral column with a fine scissor.</li>
	</ol>
	</li>
	<li>Clearing the Mouse Spinal Cord
	<ol>
		<li>Place the spinal cord in 15 ml of hydrogel solution overnight at 4 &#176;C.</li>
		<li>Remove 10 ml of the hydrogel solution and pour the rest of the solution with the spinal cord segments to a 5 ml tube. Fill the 5 ml tube with the hydrogel solution until it is completely full.</li>
		<li>Stretch a small piece of parafilm over the top of the 5 ml tube and then wrap parafilm around the neck of the tube. Note: Ensure the parafilm contacts the hydrogel solution and there are no bubbles between the hydrogel solution and the parafilm.</li>
		<li>Put the 5 ml tube in a 37&#160;&#176;C oven overnight. Observe the hydrogel solution till it becomes a gel.</li>
		<li>Take the 5 ml tube out of the oven and remove the gel from the tube with a spanner (such as a shoveled 1 ml pipette tip). Remove the gel from the spinal cord tissue by sticking the coarse tissue to the gel and then take the coarse tissue away (the gel will stick to the tissue and therefore will be removed by the tissue).</li>
		<li>After removing the gel from the spinal cord tissue, wash the spinal cord 4 times with 1x PBS (pH 7.4) for 24 hr on a shaker.</li>
		<li>Prepare the CUBIC clearing solution by dissolving 3.85 g urea and 3.85 g N,N,N&#39;,N&#39;-tetrakis (2-hydroxypropyl) ethylenediamine in 5.38 ml distilled water. 
		Note: A hot stirrer is used. Add 2.31 g polyethylene glycol mono-p-isooctylphenyl ether/Triton X-100 to the solution once it is clear and cools down to room temperature. 
		Note: 10 g of these three chemicals are for one mouse.</li>
		<li>Cut the mouse spinal cord coronally into 2-3 mm long segments with a razor blade and put them into 5 ml of the above CUBIC clearing solution. Place on a shaker in the oven at 37&#160;&#176;C for 3 days.</li>
		<li>Three days later, change the clearing solution to a fresh one. Note: the above solution is prepared prior to use.</li>
		<li>Two to three days later after refreshing the CUBIC clearing solution, check the transparency of the tissue against a paper with font 8 letters. If it is transparent, the letters can be seen through the cleared tissue. Remove the clearing solution and add 4 ml of PBST (0.1% Triton-X100) to wash the tissue 4 times a day (every 6 hours).</li>
	</ol>
	</li>
</ol>
2. Immunofluorescence Staining
<ol>
	<li>Incubate the spinal cord segments in the primary antibody solution (anti-serotonin, raised in rabbit, diluted 1:100 in PBST) for 3 days on a shaker in a 37&#160;&#176;C oven.</li>
	<li>Remove the antibody solution and add 4 ml of PBST (0.1% Triton-X100) to wash the spinal cord segments 4 times a day (every 6 hr) in a 37&#160;&#176;C oven.</li>
	<li>Remove the PBST solution and add the secondary antibody solution (594 conjugated goat anti-rabbit IgG, diluted 1:100 in PBST) to incubate the tissue for 3 days on a shaker in a 37&#160;&#176;C oven.</li>
	<li>Remove the secondary antibody solution and add 4 ml of PBST to wash the spinal cord segments 4 times a day (every 6 hours) on a shaker in a 37&#160;&#176;C oven. The next day the tissue is ready for imaging.</li>
</ol>
3. Imaging
<ol>
	<li>Remove the PBST solution and add 4 ml of 85% glycerol to make the refractive index of the tissue even.</li>
	<li>Check the clarity of the tissue. Once it is clear in approximately 1 hr, put the tissue onto a large, thin glass slide which fits the holding frame of the multi-photo fluorescent microscope.</li>
	<li>Put a few drops of 85% glycerol next to the spinal cord tissue and coverslip it with a 22 x 50 mm2 coverslip glass. 
	Note: The orientation of the spinal cord decides how the image looks like.</li>
	<li>Put the glass slide onto the holding frame of the multi-photon fluorescent microscope and move the tissue into the light pathway.</li>
	<li>Choose the Helium Neon laser line 594 nm and adjust the intensity of the laser to the optimal level by checking the brightness of the positive signal in the live image.</li>
	<li>Select the scanning area of the spinal cord tissue using the 20X objective (in water, NA 0.7) and prepare to create a z-stack, and the depth of each step set to 3 &#181;m.</li>
	<li>Scan the tissue from the top to the bottom of the z-stack under the 20X objective and then 63X objective (in oil, NA 1.4) (step size was 1 &#181;m), respectively. Reconstruct 3D videos using a 3D software28.</li>
</ol>

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The authors have nothing to disclose.

The protocol described shows how to image serotonergic fibers in the mouse spinal cord with the combined CLARITY and CUBIC techniques. It introduces a faster clearing process compared to the passive clearing protocol developed by Cheung et al.14 and Tomer et al.15 and allows the spinal cord tissue to be well supported by the hydrogel during clearing.
An important step during fixation of the mouse spinal cord, as reported by Cheung et al.14 ...

Supraspinal projections are responsible for the modulation of diverse behaviors such as pain perception. One of the projections carrying nociceptive information contains serotoninergic fibers, which originate from the hindbrain raphe and adjacent reticular nuclei1,2. Physiological and pharmacological studies have demonstrated an increased release of serotonin in the dorsal horn of the spinal cord after electrical stimulation of the raphe nuclei in the hindbrain3-5. In the rat and opossum, serotonergic raphespinal fibers have dense terminals, not only in the dorsal horn6-8, but also in the intermediate zone7,9,10, the ventral horn7,11, and even lamina 1012,13. There are no similar studies in the mouse. The present study aimed to map the termination pattern of serotonergic fibers arising from the hindbrain raphe nuclei and their adjacent reticular nuclei in the mouse spinal cord using the recently published CLARITY14 method and its modification - CUBIC15. 
Conventional fluorescence or peroxidase immunohistochemistry of the spinal cord clearly shows the distribution of serotonergic fibers in the gray matter of the spinal cord in 30-40 &#181;m thick cross-sections. However, this approach does not show the continuity of the serotonergic fiber tracts in the white matter and their collaterals in the gray matter. Although the 3D reconstruction of histological sections has advanced our knowledge of fiber tracts, it remains a challenge for histologists and anatomists to follow a single tract due to small distortions in the tissue caused by cutting. To circumvent this obstacle a number of researchers have developed various protocols for making the whole tissue structure transparent, and collecting an image of unaltered tissue in a single video file17-21. So far, the clear, lipid-exchanged, acrylamide-hybridized rigid, imaging/ immunostaining compatible, tissue hydrogel (CLARITY) technique, developed by Deisseroth's group14,15, as well as CUBIC, developed by Susaki et al16 are the most successful. Since the publication of the protocols, many researchers have started using these techniques to investigate various aspects of biological tissues, including, not only the brain22-25, but also the heart, kidneys, intestine, and the lungs26,27. 
By fixing the mouse spinal cord with the hydrogel solution (CLARITY) and clearing with the CUBIC reagents (which is a much faster method than that described by the original CLARITY protocol14,15), a spinal cord tissue block of 2-3 mm long was cleared within two weeks and immunofluorescence staining for serotonin completed in eight days. With just a combination of chemical agents, conventional immunohistochemistry can be used to create an image of individual fiber tracts in a 3D video file in approximately one month.

<table><tbody><tr><td>Photoinitiator VA044</td><td>Wako</td><td>va-044/225-02111</td><td>http://www.wako-chem.co.jp/specialty/waterazo/VA-044.htm</td></tr><tr><td>40% acrylamide solution</td><td>Bio Rad</td><td>161-0140</td><td>http://www.bio-rad.com/en-au/sku/161-0140-40-acrylamide-solution</td></tr><tr><td>2% Bis Solution</td><td>Bio Rad</td><td>161-0142</td><td>http://www.bio-rad.com/en-au/sku/161-0142-2-bis-solution?parentCategoryGUID=5e7a4f31-&lt;br/&gt;879c-4d63-ba0b-82556a0ccf1d</td></tr><tr><td>paraformaldehyde</td><td>Sigma</td><td>158127</td><td>http://www.sigmaaldrich.com/catalog/product/sial/158127?lang=en&amp;amp;region=AU</td></tr><tr><td>urea</td><td>Merck Millipore</td><td>66612</td><td>http://www.merckmillipore.com/AU/en/product/Urea---CAS-57-13-6---Calbiochem,EMD_BIO-66612</td></tr><tr><td>N,N,N&amp;rsquo;,N&amp;rsquo;-tetrakis (2-hydroxypropyl) ethylenediamine</td><td>Merck Millipore</td><td>821940</td><td>http://www.merckmillipore.com/AU/en/product/Ethylenediamine-N,N,N&#039;,N&#039;-tetra-2-propanol,MDA_CHEM-821940</td></tr><tr><td>Triton-X 100</td><td>Merck Millipore</td><td>648462</td><td>http://www.merckmillipore.com/AU/en/product/TRITON&amp;reg;-X-100-Detergent---CAS-9002-93-1---Calbiochem,EMD_BIO-648462</td></tr><tr><td>sucrose</td><td>Sigma</td><td>S0389</td><td>http://www.sigmaaldrich.com/catalog/product/sigma/s0389?lang=en&amp;amp;region=AU</td></tr><tr><td>serotonin antibody</td><td>Merck Millipore</td><td>AB938</td><td>http://www.merckmillipore.com/AU/en/product/Anti-Serotonin-Antibody,MM_NF-AB938</td></tr><tr><td>goat anti rabbit IgG (H+L) Secondary Antibody, Alexa Fluor&amp;reg; 594 conjugate</td><td>Life Technologies</td><td>&amp;nbsp;A-11012</td><td>https://www.lifetechnologies.com/order/genome-database/antibody/Rabbit-IgG-H-L-Secondary-Antibody-Polyclonal/A-11012</td></tr><tr><td>multi-photon microscope</td><td>Leica</td><td>Leica TCS SP5 MP STED</td><td>http://www.leica-microsystems.com/products/confocal-microscopes/details/product/leica-tcs-sp5-mp/</td></tr></tbody></table>

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.

This section shows results from serotonin antibody staining in the transparent mouse spinal cord using a combination of the CLARITY and CUBIC protocols. We show that serotonergic fibers are present in all laminae of the spinal cord with a predominance in the ventral portion of the ventral horn (Figure 1, also see Video 1). The control tissue did not have positive fibers (result was not shown). In the ventral horn, densely packed serotonergic fibers are pr...

Watch this Scientific Journal Video about Imaging Serotonergic Fibers in the Mouse Spinal Cord Using the CLARITY/CUBIC Technique at JoVE.com

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

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 ...

imaging-serotonergic-fibers-mouse-spinal-cord-using-claritycubic

Research

JoVE Journal

Neuroscience

10.5K Views.  Neuroscience Research Australia. 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.

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

In vivo Imaging of the Mouse Spinal Cord Using Two-photon Microscopy

In vivo imaging using two-photon microscopy 1 in mice that have been genetically engineered to express fluorescent proteins in specific cell types 2-3 has significantly broadened our knowledge of physiological and pathological processes in numerous tissues in vivo 4-7. In studies of the central nervous system (CNS), there has been a broad application of in vivo imaging in the brain, which has produced a plethora of novel and often unexpected findings about the behavior of cells such as neurons, astrocytes, microglia, under physiological or pathological conditions 8-17. However, mostly technical complications have limited the implementation of in vivo imaging in studies of the living mouse spinal cord. In particular, the anatomical proximity of the spinal cord to the lungs and heart generates significant movement artifact that makes imaging the living spinal cord a challenging task.
We developed a novel method that overcomes the inherent limitations of spinal cord imaging by stabilizing the spinal column, reducing respiratory-induced movements and thereby facilitating the use of two-photon microscopy to image the mouse spinal cord in vivo. This is achieved by combining a customized spinal stabilization device with a method of deep anesthesia, resulting in a significant reduction of respiratory-induced movements. This video protocol shows how to expose a small area of the living spinal cord that can be maintained under stable physiological conditions over extended periods of time by keeping tissue injury and bleeding to a minimum. Representative raw images acquired in vivo detail in high resolution the close relationship between microglia and the vasculature. A timelapse sequence shows the dynamic behavior of microglial processes in the living mouse spinal cord. Moreover, a continuous scan of the same z-frame demonstrates the outstanding stability that this method can achieve to generate stacks of images and/or timelapse movies that do not require image alignment post-acquisition. Finally, we show how this method can be used to revisit and reimage the same area of the spinal cord at later timepoints, allowing for longitudinal studies of ongoing physiological or pathological processes in vivo.

In vivo Imaging of the Mouse Spinal Cord Using Two-photon Microscopy

A minimally invasive protocol to stabilize the mouse spinal column and perform repetitive in vivo spinal cord imaging using two-photon microscopy is described. This method combines a spinal stabilization device and an anesthetic regimen to minimize respiratory-induced movements and produce raw imaging data that require no alignment or other post-processing.

In vivo imaging using two-photon microscopy 1 in mice that have been genetically engineered to express fluorescent proteins in specific cell types 2-3 has ...

A Procedure for Implanting a Spinal Chamber for Longitudinal In Vivo Imaging of the Mouse Spinal Cord

Studies in the mammalian neocortex have enabled unprecedented resolution of cortical structure, activity, and response to neurodegenerative insults by repeated, time-lapse in vivo imaging in live rodents. These studies were made possible by straightforward surgical procedures, which enabled optical access for a prolonged period of time without repeat surgical procedures. In contrast, analogous studies of the spinal cord have been previously limited to only a few imaging sessions, each of which required an invasive surgery. As previously described, we have developed a spinal chamber that enables continuous optical access for upwards of 8 weeks, preserves mechanical stability of the spinal column, is easily stabilized externally during imaging, and requires only a single surgery. Here, the design of the spinal chamber with its associated surgical implements is reviewed and the surgical procedure is demonstrated in detail. Briefly, this video will demonstrate the preparation of the surgical area and mouse for surgery, exposure of the spinal vertebra and appropriate tissue debridement, the delivery of the implant and vertebral clamping, the completion of the chamber, the removal of the delivery system, sealing of the skin, and finally, post-operative care. The procedure for chronic in vivo imaging using nonlinear microscopy will also be demonstrated. Finally, outcomes, limitations, typical variability, and a guide for troubleshooting are discussed.

A Procedure for Implanting a Spinal Chamber for Longitudinal In Vivo Imaging of the Mouse Spinal Cord

In this video, we describe a procedure for implanting a chronic optical imaging chamber over the dorsal spinal cord of a live mouse. The chamber, surgical procedure, and chronic imaging are reviewed and demonstrated.

Studies in the mammalian neocortex have enabled unprecedented resolution of cortical structure, activity, and response to neurodegenerative insults by ...

Two-photon Imaging of Cellular Dynamics in the Mouse Spinal Cord

Two-photon (2P) microscopy is utilized to reveal cellular dynamics and interactions deep within living, intact tissues. Here, we present a method for live-cell imaging in the murine spinal cord. This technique is uniquely suited to analyze neural precursor cell (NPC) dynamics following transplantation into spinal cords undergoing neuroinflammatory demyelinating disorders. NPCs migrate to sites of axonal damage, proliferate, differentiate into oligodendrocytes, and participate in direct remyelination. NPCs are thereby a promising therapeutic treatment to ameliorate chronic demyelinating diseases. Because transplanted NPCs migrate to the damaged areas on the ventral side of the spinal cord, traditional intravital 2P imaging is impossible, and only information on static interactions was previously available using histochemical staining approaches. Although this method was generated to image transplanted NPCs in the ventral spinal cord, it can be applied to numerous studies of transplanted and endogenous cells throughout the entire spinal cord. In this article, we demonstrate the preparation and imaging of a spinal cord with enhanced yellow fluorescent protein-expressing axons and enhanced green fluorescent protein-expressing transplanted NPCs.

A new ex vivo preparation for imaging the mouse spinal cord. This protocol allows for two-photon imaging of live cellular interactions throughout the spinal cord.

Two-photon (2P) microscopy is utilized to reveal cellular dynamics and interactions deep within living, intact tissues. Here, we present a method for ...

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.

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 ...

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 ...

Establishing a Mouse Contusion Spinal Cord Injury Model Based on a Minimally Invasive Technique

Using minimally invasive methods to model spinal cord injury (SCI) can minimize behavioral and histological differences between experimental animals, thereby improving the reproducibility of the experiments.
These methods need two requirements to be fulfilled: clarity of the surgical anatomical pathway and simplicity and convenience of the laboratory device. Crucially for the operator, a clear anatomical pathway provides minimally invasive exposure, which avoids additional damage to the experimental animal during the surgical procedures and allows the animal to maintain a consistent and stable anatomical morphology during the experiment.
In this study, the use of a novel integrated platform called the SCI coaxial platform for spinal cord injury in small animals to expose the T9 level spinal cord in a minimally invasive way and stabilize and immobilize the vertebra of mice using a vertebral stabilizer is researched, and, finally, a coaxial gravity impactor is used to contuse the spinal cord of mice to approach different degrees of T9 spinal cord injury. Finally, histological results are provided as a reference for the readers.

Minimally invasive techniques and a simple laboratory device improve the reproducibility of the spinal cord injury model by reducing operative damage to the experimental animals and allowing anatomical morphology maintenance. The method is worthwhile because the reliable results and reproducible procedure facilitate investigations of the mechanisms of disease reparation.

Using minimally invasive methods to model spinal cord injury (SCI) can minimize behavioral and histological differences between experimental animals, ...

Establishment of Central Cord Syndrome Model in C57BL/6J Mouse

Animal models of central cord syndrome (CCS) could substantially benefit preclinical research. Identifiable anatomical pathways can give minimally invasive exposure approaches and reduce extra injury to experimental animals during operation, enabling the maintenance of consistent and stable anatomical morphology during experiments to minimize behavioral and histological differences between individuals to improve the reproducibility of experiments. In this study, the C6 level spinal cord was exposed using a spinal cord injury coaxial platform (SCICP) and combination with a minimally invasive technique. With the assistance of a vertebral stabilizator, we fixed the vertebrae and compressed the spinal cords of C57BL/6J mice with 5 g/mm2 and 10 g/mm2 weights with SCICP to induce different degrees of C6 spinal cord injury. In line with the previous description of CCS, the results reveal that the lesion in this model is concentrated in the gray matter around the central cord, enabling further research into CCS. Finally, histological results are provided as a reference for the readers.

The present protocol simulating central cord syndrome (CCS) in mice has improved repeatability and minimized operation damage to the experimental animals, avoiding disrupting the anatomical structure excessively. The strategy in this study is advantageous because it allows for research into injury mechanisms by producing consistent results.

Animal models of central cord syndrome (CCS) could substantially benefit preclinical research. Identifiable anatomical pathways can give minimally ...

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 ...

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

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Rozdziały w tym wideo

In vivo Imaging of the Mouse Spinal Cord Using Two-photon Microscopy

A Procedure for Implanting a Spinal Chamber for Longitudinal In Vivo Imaging of the Mouse Spinal Cord

Two-photon Imaging of Cellular Dynamics in the Mouse Spinal Cord

Optical Clearing of the Mouse Central Nervous System Using Passive CLARITY

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

Imaging and Quantification of Intact Neuronal Dendrites via CLARITY Tissue Clearing

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

Establishing a Mouse Contusion Spinal Cord Injury Model Based on a Minimally Invasive Technique

Establishment of Central Cord Syndrome Model in C57BL/6J Mouse

Imaging the Gut with "CLARITY"