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 and Tomer et al.15, is to keep all the solutions cool on ice, which prevents the polymerization of the chemicals. The other critical step is to perfuse the mouse slowly with the hydrogel solution in order to avoid swelling of the nervous tissue. For the hydrogel solution to become a gel, degassing is required. As an alternative to this, we used parafilm to cover the top of the tube and left no air bubbles between the hydrogel solution and the parafilm. This worked very well and the hydrogel solution became a gel after a few hours. Although some bubbles were found in the gel, they did not interfere with the following experiments, which was demonstrated by the absence of bubbles in the spinal cord tissue. Clearing is the most critical step, and requires a combination of reagents. The clearing solution, containing amino-alcohol, removes the lipid of the spinal cord tissue much faster than the SDS/boric acid based clearing solution used in the original CLARITY protocol16.
The advantage of CLARITY and CUBIC over traditional immunohistochemical techniques is that the entire tissue block can be imaged without sectioning the tissue, as a result, the continuity of the serotonergic fibers is retained, and it is possible to follow the same fiber for a long distance within the z-stack. With this advantage, collateralization can be confidently identified with the 3D images, and novel conclusions can therefore be made about the nature of serotonergic pathways. The protocol described is a convenient tool for investigation of fiber systems, as well as nuclei by simply labeling the tissue with specific antibodies. This technique is also an ideal choice for interrogating different systems in the same mouse brain by using double or triple labeling. In our laboratory setting, a multi-photon fluorescent microscope was available with a working distance of approximately 300 &#181;m. This limits our image to a maximum of 600 &#181;m if both sides of the tissue are scanned. However, a light sheet fluorescent microscope, can image the entire mouse brain and spinal cord through their full width, length and depth. The other advantage of this technique is that the antibodies can be eluted and then another different antibody or antibodies applied again to the same tissue. This reduces animal usage considerably as well as the costs associated with preparing new brain tissue for each experiment. For the same reason, this technique can be applied to other tissues. There have already been studies conducted on the lungs, kidneys, heart, intestine, and tumor tissue26,27.
There are also limitations to this technique, for example, the raphe nuclei in the mouse hindbrain contains many types of neurons, including GABAergic neurons. Although CLARITY and CUBIC will allow for the study of the distribution of the diverse types of raphespinal fibers in a single mouse brain and spinal cord. Those neurons, which are not identified specifically by their neurotransmitters or protein markers, can not be identified in this setting. An alternative is to label the specific neurons or fiber systems with nucleic acid probes such as in in situ hybridization. This is, on the other hand, limited by the labeling system and the available filters for fluorescent imaging. Imaging all the raphespinal fibers therefore remains a challenge. If these techniques were combined with an intracranial injection of an anterograde tracer (such as phaseolus vulgaris leucoagglutinin - PHA-L), it might be possible to see serotonergic fibers, GABAergic fibers, and others.

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 border="0" cellpadding="0" cellspacing="0" width="2081">
	<tbody>
		<tr height="22">
			<td height="22" style="height:22px;width:495px;">Photoinitiator VA044</td>
			<td style="width:495px;">Wako</td>
			<td style="width:284px;">va-044/225-02111</td>
			<td style="width:807px;">http://www.wako-chem.co.jp/specialty/waterazo/VA-044.htm</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">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 height="21">
			<td height="21" style="height:21px;">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=5e7a4f31-879c-4d63-ba0b-82556a0ccf1d</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">paraformaldehyde</td>
			<td>Sigma</td>
			<td align="right">158127</td>
			<td>http://www.sigmaaldrich.com/catalog/product/sial/158127?lang=en&#38;region=AU</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">urea</td>
			<td>Merck Millipore</td>
			<td align="right">66612</td>
			<td>http://www.merckmillipore.com/AU/en/product/Urea---CAS-57-13-6---Calbiochem,EMD_BIO-66612</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">N,N,N&#8217;,N&#8217;-tetrakis (2-hydroxypropyl) ethylenediamine</td>
			<td>Merck Millipore</td>
			<td align="right">821940</td>
			<td>http://www.merckmillipore.com/AU/en/product/Ethylenediamine-N,N,N&#39;,N&#39;-tetra-2-propanol,MDA_CHEM-821940</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Triton-X 100</td>
			<td>Merck Millipore</td>
			<td align="right">648462</td>
			<td>http://www.merckmillipore.com/AU/en/product/TRITON&#174;-X-100-Detergent---CAS-9002-93-1---Calbiochem,EMD_BIO-648462</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">sucrose</td>
			<td>Sigma</td>
			<td>S0389</td>
			<td>http://www.sigmaaldrich.com/catalog/product/sigma/s0389?lang=en&#38;region=AU</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">2,2&#8217;,2&#8217;&#8217;- nitrilotriethanol</td>
			<td>Merck Millipore</td>
			<td align="right">137002</td>
			<td>http://www.merckmillipore.com/AU/en/product/Triethanolamine-(Trolamine),MDA_CHEM-137022</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">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 height="21">
			<td height="21" style="height:21px;">goat anti rabbit IgG (H+L) Secondary Antibody, Alexa Fluor&#174; 594 conjugate</td>
			<td>Life Technologies</td>
			<td>&#160;A-11012</td>
			<td>https://www.lifetechnologies.com/order/genome-database/antibody/Rabbit-IgG-H-L-Secondary-Antibody-Polyclonal/A-11012</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">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 present in the ventromedial portion of the ventral horn and they extend towards the lateral portion of the ventral horn (Figure 2, also see Video 2). Some immunopositive fibers also extend from the ventral horn towards the dorsal horn or the central canal. Immunopositive fibers are present in all laminae of the dorsal horn but there is a small gap between positive fibers in lamina 2 and 4, especially in the lateral part of the dorsal horn. The majority of serotonergic fibers in the dorsal horn are of small diameter and only a small number of them are of large diameter (Video 1). In a horizontal section, densely packed serotonergic fibers in the ventral horn were observed to travel along the longitudinal axis of the spinal cord and form a large fiber bundle. The most interesting finding is that this large fiber bundle issues branches regularly along the longitudinal axis, which are perpendicular to the fiber bundle. These branches further collateralize along their paths to the lateral portion of the ventral horn. Compared with these branches, the ones extending towards the midline are irregular and smaller (Figure 2). Under the 63X objective, the serotonergic fibers in the dorsal horn were observed to travel along the axis of the spinal cord and terminate at various points along the path of the tract. The thick fibers are sporadically distributed among the thin fibers (Figure 3, also see Video 3).
<img alt="Figure 1" src="/files/ftp_upload/53673/53673fig1.jpg" /> 
Figure 1: Serotonergic fibers in a coronal section of the lumbar cord at 200X magnification. Left is the ventral horn, right is the dorsal horn, dorsal is the medial part of the spinal cord, and ventral is the lateral part of the spinal cord. The scale bar is 100 &#181;m. Serotonergic fibers are present in all laminae, especially in the ventromedial part of the ventral horn where the fiber density is high. These fibers extend towards both the lateral part of the ventral horn and the central canal or dorsal horn. The density of fibers in other laminae is low. The majority of serotonergic fibers are thin. Only a small number of fibers are thick and they are seen in both the dorsal horn and the ventral horn (<a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/53673/Video1.mp4">also see Video 1 (Right click to download)</a>).
<img alt="Figure 2" src="/files/ftp_upload/53673/53673fig2.jpg" /> 
Figure 2: Serotonergic fibers in the ventral portion of the ventral horn at 200X magnification. Left and right are the lateral sides of the spinal cord, dorsal is the rostral part of the spinal cord, and ventral is the caudal part of the spinal cord. The scale bar is 100 &#181;m. Serotonergic fibers are shown travelling along the longitudinal axis of the spinal cord and packed in the medial portion of the ventral horn where they form a thick fiber bundle. Out of this bundle, some branches are extended at regular intervals towards the lateral portion of the ventral horn, whereas others that extend towards the midline are irregular and shorter. These fibers terminate at many points along the path of the tract (<a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/53673/Video2.mp4">also see Video 2 (Right click to download)</a>).
<img alt="Figure 3" src="/files/ftp_upload/53673/53673fig3.jpg" /> 
Figure 3: Serotonergic fibers in the dorsal horn under at 630X magnification. Left is the lateral part of the spinal cord, right is the medial part of the spinal cord, dorsal and ventral refer to the dorsal and ventral part of the dorsal horn, respectively. The scale bar is 7 &#181;m. Serotonergic fibers are travelling along the longitudinal axis of the spinal cord and terminate along the path of the tract. These fibers are mainly of small diameter with a small number of thick fibers intermingled. These fibers rarely overlap with each other (<a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/53673/Video3.mp4">also see Video 3 (Right click to download)</a>).

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

Nanyang Technological University

Research

JoVE Journal

Neuroscience

10.4K 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

Spinal Cord Electrophysiology II: Extracellular Suction Electrode Fabrication

Development of neural circuitries and locomotion can be studied using neonatal rodent spinal cord central pattern generator (CPG) behavior. We demonstrate a method to fabricate suction electrodes that are used to examine CPG activity, or fictive locomotion, in dissected rodent spinal cords. The rodent spinal cords are placed in artificial cerebrospinal fluid and the ventral roots are drawn into the suction electrode. The electrode is constructed by modifying a commercially available suction electrode. A heavier silver wire is used instead of the standard wire given by the commercially available electrode. The glass tip on the commercial electrode is replaced with a plastic tip for increased durability. We prepare hand drawn electrodes and electrodes made from specific sizes of tubing, allowing consistency and reproducibility. Data is collected using an amplifier and neurogram acquisition software. Recordings are performed on an air table within a Faraday cage to prevent mechanical and electrical interference, respectively.

A demonstration of the fabrication and use of an extracellular suction electrode used to measure electrophysiological recordings of neonatal rodent spinal cords in vitro

Development of neural circuitries and locomotion can be studied using neonatal rodent spinal cord central pattern generator (CPG) behavior. We demonstrate ...

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 ...

Spinal Cord Transection in the Larval Zebrafish

Mammals fail in sensory and motor recovery following spinal cord injury due to lack of axonal regrowth below the level of injury as well as an inability to reinitiate spinal neurogenesis. However, some anamniotes including the zebrafish Danio rerio exhibit both sensory and functional recovery even after complete transection of the spinal cord. The adult zebrafish is an established model organism for studying regeneration following spinal cord injury, with sensory and motor recovery by 6&#160;weeks post-injury. To take advantage of in vivo analysis of the regenerative process available in the transparent larval zebrafish as well as genetic tools not accessible in the adult, we use the larval zebrafish to study regeneration after spinal cord transection. Here we demonstrate a method for reproducibly and verifiably transecting the larval spinal cord. After transection, our data shows sensory recovery beginning at 2&#160;days post-injury (dpi), with the C-bend movement detectable by 3 dpi and resumption of free swimming by 5 dpi. Thus we propose the larval zebrafish as a companion tool to the adult zebrafish for the study of recovery after spinal cord injury.

After spinal transection, adult zebrafish have functional recovery by six weeks post-injury. To take advantage of larval transparency and faster recovery, we present a method for transecting the larval spinal cord. After transection, we observe sensory recovery beginning at 2&#160;days post-injury, and C-bend movement by 3 days post-injury.

Mammals fail in sensory and motor recovery following spinal cord injury due to lack of axonal regrowth below the level of injury as well as an inability ...

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 ...

Diffusion Imaging in the Rat Cervical Spinal Cord

Magnetic resonance imaging (MRI) is the state of the art approach for assessing the status of the spinal cord noninvasively, and can be used as a diagnostic and prognostic tool in cases of disease or injury. Diffusion weighted imaging (DWI), is sensitive to the thermal motion of water molecules and allows for inferences of tissue microstructure. This report describes a protocol to acquire and analyze DWI of the rat cervical spinal cord on a small-bore animal system. It demonstrates an imaging setup for the live anesthetized animal and recommends a DWI acquisition protocol for high-quality imaging, which includes stabilization of the cord and control of respiratory motion. Measurements with diffusion weighting along different directions and magnitudes (b-values) are used. Finally, several mathematical models of the resulting signal are used to derive maps of the diffusion processes within the spinal cord tissue that provide insight into the normal cord and can be used to monitor injury or disease processes noninvasively.

The goal of this protocol is to obtain high-quality diffusion weighted magnetic resonance imaging (DWI) of the rat spinal cord for noninvasive characterization of tissue microstructure. This protocol describes optimizations of the MRI sequence, radiofrequency coil, and analysis methods to enable DWI images free from artifacts.

Magnetic resonance imaging (MRI) is the state of the art approach for assessing the status of the spinal cord noninvasively, and can be used as a ...

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 ...

Imaging Neural Activity in the Primary Somatosensory Cortex Using Thy1-GCaMP6s Transgenic Mice

The mammalian brain exhibits marked symmetry across the sagittal plane. However, detailed description of neural dynamics in symmetric brain regions in adult mammalian animals remains elusive. In this study, we describe an experimental procedure for measuring calcium dynamics through dual optical windows above bilateral primary somatosensory corticies (S1) in Thy1-GCaMP6s transgenic mice using 2-photon (2P) microscopy. This method enables recordings and quantifications of neural activity in bilateral mouse brain regions one at a time in the same experiment for a prolonged period in vivo. Key aspects of this method, which can be completed within an hour, include minimally invasive surgery procedures for creating dual optical windows, and the use of 2P imaging. Although we only demonstrate the technique in the S1 area, the method can be applied to other regions of the living brain facilitating the elucidation of structural and functional complexities of brain neural networks.

Imaging Neural Activity in the Primary Somatosensory Cortex Using Thy1-GCaMP6s Transgenic Mice

We describe an experimental procedure for measuring neuronal activity through dual optical windows above bilateral primary somatosensory corticies (S1) in Thy1-GCaMP6s transgenic mice using 2-photon (2P) microscopy in vivo.

The mammalian brain exhibits marked symmetry across the sagittal plane. However, detailed description of neural dynamics in symmetric brain regions in ...

Laminectomy and Spinal Cord Window Implantation in the Mouse

This protocol describes a method for spinal cord laminectomy and glass window implantation for in vivo imaging of the mouse spinal cord. An integrated digital vaporizer is utilized to achieve a stable plane of anesthesia at a low-flow rate of isoflurane. A single vertebral spine is removed, and a commercially available cover-glass is overlaid on a thin agarose bed. A 3D-printed plastic backplate is then affixed to the adjacent vertebral spines using tissue adhesive and dental cement. A stabilization platform is used to reduce motion artifact from respiration and heartbeat. This rapid and clamp-free method is well-suited for acute multi-photon fluorescence microscopy. Representative data are included for an application of this technique to two-photon microscopy of the spinal cord vasculature in transgenic mice expressing eGFP:Claudin-5 &#8212; a tight junction protein.

This protocol describes implantation of a glass window onto the spinal cord of a mouse to facilitate visualization by intravital microscopy.

This protocol describes a method for spinal cord laminectomy and glass window implantation for in vivo imaging of the mouse spinal cord. An integrated ...

Diffusion Tensor Magnetic Resonance Imaging in Chronic Spinal Cord Compression

Chronic spinal cord compression is the most common cause of spinal cord impairment in patients with nontraumatic spinal cord damage. Conventional magnetic resonance imaging (MRI) plays an important role in both confirming the diagnosis and evaluating the degree of compression. However, the anatomical detail provided by conventional MRI is not sufficient to accurately estimate neuronal damage and/or assess the possibility of neuronal recovery in chronic spinal cord compression patients. In contrast, diffusion tensor imaging (DTI) can provide quantitative results according to the detection of water molecule diffusion in tissues. In the present study, we develop a methodological framework to illustrate the application of DTI in chronic spinal cord compression disease. DTI fractional anisotropy (FA), apparent diffusion coefficients (ADCs), and eigenvector values are useful for visualizing microstructural pathological changes in the spinal cord. Decreased FA and increases in ADCs and eigenvector values were observed in chronic spinal cord compression patients compared to healthy controls. DTI could help surgeons understand spinal cord injury severity and provide important information regarding prognosis and neural functional recovery. In conclusion, this protocol provides a sensitive, detailed, and noninvasive tool to evaluate spinal cord compression.

Here, we present a protocol for the application of diffusion tensor imaging parameters to evaluate spinal cord compression.

Chronic spinal cord compression is the most common cause of spinal cord impairment in patients with nontraumatic spinal cord damage. Conventional magnetic ...

Intra-Arterial Delivery of Neural Stem Cells to the Rat and Mouse Brain: Application to Cerebral Ischemia

Neural stem cell (NSC) therapy is an emerging innovative treatment for stroke, traumatic brain injury and neurodegenerative disorders. As compared to intracranial delivery, intra-arterial administration of NSCs is less invasive and produces a more diffuse distribution of NSCs within the brain parenchyma. Further, intra-arterial delivery allows the first-pass effect in the brain circulation, lessening the potential for trapping of cells in peripheral organs, such as liver and spleen, a complication associated with peripheral injections. Here, we detail the methodology, in both mice and rats, for delivery of NSCs through the common carotid artery (mouse) or external carotid artery (rat) to the ipsilateral hemisphere after an ischemic stroke. Using GFP-labeled NSCs, we illustrate the widespread distribution achieved throughout the rodent ipsilateral hemisphere at 1 d, 1 week and 4 weeks after postischemic delivery, with a higher density in or near the ischemic injury site. In addition to long-term survival, we show evidence of differentiation of GFP-labeled cells at 4 weeks. The intra-arterial delivery approach described here for NSCs can also be used for administration of therapeutic compounds, and thus has broad applicability to varied CNS injury and disease models across multiple species.

A method for delivering neural stem cells, adaptable for injecting solutions or suspensions, through the common carotid artery (mouse) or external carotid artery (rat) after ischemic stroke is reported. Injected cells are distributed broadly throughout the brain parenchyma and can be detected up to 30 d after delivery.

Neural stem cell (NSC) therapy is an emerging innovative treatment for stroke, traumatic brain injury and neurodegenerative disorders. As compared to ...