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).

Ética Declaración: Todos los procedimientos que involucran sujetos animales siguen las directrices del Comité de Cuidado de Animales y Ética (ACEC) en la Universidad de Nueva Gales del Sur (el número ACEC aprobado es de 14 / 94A). 1. Preparación de la cuerda del ratón transparente espinal <ol><li> Preparación de hielo solución de hidrogel frío <ol><li> Preparación de solución de paraformaldehído al 16% (PFA) <ol><li> Añadir 16 g de polvo en 70 ml de paraformaldehído-calentado previamente en agua destilada (50-55 ° C) y se agita en un agitador magnético se calienta hasta que se disuelva paraformaldehído. Nota: No permita que la solución se caliente más de 55 ° C y tenga en cuenta que paraformaldehído es tóxico. </li><li> Transferir la solución de paraformaldehído a un cilindro y añadir agua destilada hasta 100 ml. Se enfría la solución de paraformaldehído usando un 4 ° C refrigerador. </li></ol></li><li> Disolver 125 mg iniciador VA-044 en 26,25 ml de agua destilada y se enfría la solución en un 4° C nevera. </li><li> solución fresca 5 ml de acrilamida (40%), 1,25 ml de solución de bis (2%) (PRECAUCIÓN: Bis y soluciones de acrilamida son tóxicos, puede causar defectos genéticos o cáncer), 5 ml 10x PBS, solución PFA 12,5 ml 16%, y el solución de iniciador VA-044 en hielo. </li><li> Mezclar las soluciones anteriores en hielo. Nota: el volumen total es de 50 ml. </li></ol></li><li> Perfusión y la Colección de la cuerda del ratón espinal <ol><li> Anestesiar un ratón con una inyección intraperitoneal de ketamina (80 mg / kg) y xyzaline (5 mg / kg) diluido en solución salina normal al 0,9%. La dosificación para el ratón puede ser inferior a la de las ratas. </li><li> En una superficie de plástico adecuado en una campana extractora, fijar las extremidades del ratón fuera del cuerpo (cinta adhesiva es eficaz) una vez que el ratón no responde a una pizca de su piel del pie. </li><li> Cortar la piel del ratón sobre el pecho y luego el pecho hueso para exponer el corazón con una tijera. </li><li> Usando una bomba peristáltica, con una aguja de 25 G unida a the extremo del tubo, inserte la aguja en el ventrículo izquierdo del corazón, snip la aurícula derecha para crear un punto de salida para la sangre y luego comenzar el lavado con solución salina normal 40 ml de 0,9% a razón de aproximadamente 10 ml / min. </li><li> Cuando se haya completado, perfundir el ratón con 35 ml de solución de hidrogel de hielo frío. Nota: la perfusión a una velocidad de 10 ml / min para evitar la hinchazón del tejido nervioso. </li><li> Incisión en la piel sobre la parte posterior con una cuchilla y luego retire los músculos al lado del vertebrado. Después de cortar los arcos vertebrales en un lado y dándoles la vuelta hacia el otro lado, tomar la médula espinal de la columna vertebral con una multa de tijera. </li></ol></li><li> Borrado de la cuerda del ratón espinal <ol><li> Coloque la médula espinal en 15 ml de solución de hidrogel durante la noche a 4 ° C. </li><li> Eliminar 10 ml de la solución de hidrogel y verter el resto de la solución con los segmentos de la médula espinal a un tubo de 5 ml. Llenar el tubo 5 ml con la solución de hidrogel hasta que esté completamente lleno. </li><li> Estirar un pequeño trozo de Parafilm sobre la parte superior del tubo de 5 ml y luego envolver Parafilm alrededor del cuello del tubo. Nota: Asegúrese de que los contactos parafilm la solución de hidrogel y no haya burbujas entre la solución de hidrogel y la parafina. </li><li> Poner el tubo de 5 ml en un 37 &#160; ° C horno durante la noche. Observe la solución de hidrogel hasta que se convierte en un gel. </li><li> Tome el tubo de 5 ml del horno y quitar el gel del tubo con una llave (tal como una punta de pipeta 1 ml pala). Quitar el gel a partir del tejido de la médula espinal por pegar el tejido grueso para el gel y luego tomar el papel grueso de distancia (el gel se adhiere al tejido y, por tanto, será eliminado por el tejido). </li><li> Después de quitar el gel a partir del tejido de la médula espinal, lavar la médula espinal 4 veces con 1 x PBS (pH 7,4) durante 24 horas en un agitador. </li><li> Preparar la solución de limpieza CUBIC disolviendo 3,85 g de urea y 3,85 g de N, N, N &#39;, N&#39;-tetrakis (2-hidroxipropil) ethylenediamine en 5,38 ml de agua destilada. Nota: Se utiliza un agitador caliente. Añadir 2,31 g de polietilenglicol mono-p-isooctilfenil éter / Triton X-100 a la solución una vez que es claro y se enfría a temperatura ambiente. Nota: 10 g de estos tres productos químicos son de un ratón. </li><li> Cortar la médula espinal de ratón coronal en 2-3 mm de largo segmentos con una hoja de afeitar y ponerlas en 5 ml de la solución de aclaramiento CÚBICO anteriormente. Colocar en un agitador en el horno a 37 &#160; ° C durante 3 días. </li><li> Tres días más tarde, cambiar la solución de intercambio de información para una nueva. Nota: la solución anterior se prepara antes de su uso. </li><li> Dos o tres días más tarde después de actualizar la solución de aclaramiento cúbico, comprobar la transparencia del tejido contra un papel con tipo de letra 8 letras. Si es transparente, las letras se pueden ver a través del tejido despejado. Eliminar la solución de compensación y añadir 4 ml de PBST (0,1% Triton-X100) para lavar el tejido 4 veces al día (cada 6 horas). </li></ol></li></ol><ol><li> Incubar los segmentos de la médula espinal en la solución de anticuerpo primario (anti-serotonina, criado en conejo, diluido 1: 100 en PBST) durante 3 días en un agitador en un horno de 37 ° C. </li><li> Eliminar la solución de anticuerpo y añadir 4 ml de PBST (0,1% Triton-X100) para lavar los segmentos de la médula espinal 4 veces al día (cada 6 horas) en un horno a 37 ° C. </li><li> Quitar la solución PBST y agregar la solución de anticuerpo secundario (conjugado de cabra 594 IgG anti-conejo, diluido 1: 100 en PBST) incubar el tejido durante 3 días en un agitador en un horno de 37 ° C. </li><li> Retirar la solución de anticuerpo secundario y añadir 4 ml de PBST para lavar los segmentos de la médula espinal 4 veces al día (cada 6 horas) en un agitador en un horno de 37 ° C. Al día siguiente, el tejido está listo para la imagen. </li></ol> 3. Imaging <ol><li> Eliminar la solución de PBST y añadir 4 ml de 85% de glicerol para hacer que el índice de refracción del tejido uniforme. </li><li&gt; Comprobar la claridad del tejido. Una vez que es evidente en aproximadamente 1 hora, poner el tejido sobre un gran tobogán, vidrio delgado que se ajusta el bastidor de soporte del microscopio de fluorescencia de múltiples fotos. </li><li> Ponga unas gotas de 85% de glicerol al lado del tejido de la médula espinal y el cubreobjetos con un vaso cubreobjetos 22 x 50 mm 2. Nota: La orientación de la médula espinal decide cómo la imagen se parece. </li><li> Ponga el portaobjetos de vidrio en el bastidor de soporte del microscopio de fluorescencia de múltiples fotones y mover el tejido en la vía de la luz. </li><li> Elija el helio-neón línea de láser 594 nm y ajustar la intensidad del láser al nivel óptimo marcando el brillo de la señal positiva en la imagen en vivo. </li><li> Seleccionar el área de escaneo del tejido de la médula espinal utilizando el objetivo 20X (en agua, NA 0,7) y se preparan para crear una pila Z, y la profundidad de cada paso se establece en 3 m. </li><li> Analiza el tejido desde la parte superior a la parte inferior de la pila Z bajo el objec 20Xtivo y luego objetivo 63X (en aceite, NA 1,4) (tamaño del paso fue de 1 m), respectivamente. Reconstruir videos en 3D utilizando un software de 3D ​​28. </li></ol>

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The authors have nothing to disclose.

El protocolo descrito se muestra cómo las fibras de imagen serotoninérgicos en la médula espinal de ratón con la claridad combinado y técnicas cúbico. Se introduce un proceso de compensación más rápido en comparación con el protocolo de compensación pasiva desarrollada por Cheung et al. 14 y Tomer et al. 15 y permite que el tejido de la médula espinal a estar bien apoyado por el hidrogel durante la limpieza. Un paso importante durante la fi...

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

Las fibras largas de descender a la médula espinal son esenciales para la locomoción, la percepción del dolor, y otros comportamientos. El patrón de terminación de fibras en la médula espinal de la mayoría de estos sistemas de fibra no se han investigado a fondo en cualquier especie. fibras serotoninérgicos, que se proyectan a la médula espinal, se han estudiado en ratas y zarigüeyas en secciones histológicas y su significado funcional se ha deducido en base a su patrón de terminación de fibras en la médula espinal. Con el desarrollo de la claridad y técnicas cúbico, es posible investigar este sistema de fibras y su distribución en la médula espinal, que es probable que revelar características hasta ahora desconocidas de las vías supraespinales serotoninérgicos. A continuación, ofrecemos un protocolo detallado para obtener imágenes de las fibras serotoninérgicos en la médula espinal de ratón usando la claridad combinado y técnicas cúbicos. El método consiste en la perfusión de un ratón con una solución de hidrogel y la clarificación del tejido con una combinación de la limpieza de los reactivos. el tejido de la médula espinal fue despejada en menos de dos semanas, y la posterior tinción de inmunofluorescencia contra la serotonina se completó en menos de diez días. Con un microscopio de fluorescencia multi-fotón, el tejido fue escaneada y una imagen 3D se reconstruyó usando software Osirix.

Esta sección muestra los resultados de la tinción de anticuerpos de la serotonina en la médula espinal de ratón transparente usando una combinación de la claridad y protocolos cúbicos. Se demuestra que las fibras serotoninérgicos están presentes en todas las láminas de la médula espinal con un predominio en la parte ventral del asta ventral (Figura 1, también ver video 1). El tejido de control no tuvo fibras positivos (resultado no se muestra)...

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.

Las fibras serotoninérgicos proyección de imagen en la médula espinal de ratón El uso de la claridad / Técnica CÚBICO

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 vistas.  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.

Imágenes in vivo de la médula espinal de ratón Utilizando microscopía de dos fotones

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 ...

Investigación

Neurociencias

Imaging Cleared Intact Biological Systems at a Cellular Level by 3DISCO

Tissue clearing and subsequent imaging of transparent organs is a powerful method to analyze fluorescently labeled cells and molecules in 3D, in intact organs. Unlike traditional histological methods, where the tissue of interest is sectioned for fluorescent imaging, 3D imaging of cleared tissue allows examination of labeled cells and molecules in the entire specimen. To this end, optically opaque tissues should be rendered transparent by matching the refractory indices throughout the tissue. Subsequently, the tissue can be imaged at once using laser-scanning microscopes to obtain a complete high-resolution 3D image of the specimen. A growing list of tissue clearing protocols including 3DISCO, CLARITY, Sca/e, ClearT2, and SeeDB provide new ways for researchers to image their tissue of interest as a whole. Among them, 3DISCO is a highly reproducible and straightforward method, which can clear different types of tissues and can be utilized with various microscopy techniques. This protocol describes this straightforward procedure and presents its various applications. It also discusses the limitations and possible difficulties and how to overcome them.

To obtain high-resolution images of fluorescently labeled cells within large tissues, ideally, the biological samples should be imaged without sectioning. 3DISCO is a straightforward tissue clearing procedure based on sequential incubation with organic solvents. Upon clearing, the organs become transparent allowing an end-to-end laser scan of the specimen.

Borrado de imágenes Sistemas Biológicos intactos a nivel celular por 3DISCO

Tissue clearing and subsequent imaging of transparent organs is a powerful method to analyze fluorescently labeled cells and molecules in 3D, in intact ...

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.

Un procedimiento para el implante Cámara Espinal para Longitudinal<em&gt; En Vivo</em&gt; Imágenes de la médula espinal de ratón

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.

De dos fotones de imágenes de Dinámica Celular en la médula espinal de ratón

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.

Clearing óptico del sistema nervioso central del ratón Utilizando CLARITY pasiva

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.

Inmunostaining De Retinas De Montaje Completo Con El Método De Claro De Tejido CLARITY

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.

Obtención de imágenes y cuantificación de dendritas neuronales intactas a través de CLARITY Tissue Clearing

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.

Un método de limpieza de tejidos para imágenes neuronales de escalas mesoscópicas a microscópicas

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.

Establecimiento de un modelo de lesión de la médula espinal por contusión en ratones basado en una técnica mínimamente invasiva

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

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.

<meta charset="utf8"></head><body>Imágenes del intestino con "CLARIDAD"

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.

Imágenes del intestino con "CLARIDAD"

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