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).

Declaração de Ética: Todos os procedimentos envolvendo indivíduos animais siga as orientações do Comitê de Cuidados com Animais e Ética (ACEC) da Universidade de New South Wales (o número ACEC aprovado é de 14 / 94A). 1. Preparação da Transparente cabo do rato Spinal <ol><li> Preparação de Ice Cold Hidrogel Solution <ol><li> Preparação de solução de paraformaldeído a 16% (PFA) <ol><li> Adicionar 16 g de paraformaldeído em pó em 70 ml de pré-aquecida de água destilada (50-55 ° C) e agita-se num agitador magnético até aquecida paraformaldeído é dissolvido. Nota: Não permitir que a solução para aquecer acima de 55 ° C e estar ciente de que paraformaldeído é tóxico. </li><li> Transferir a solução de paraformaldeído a um cilindro e adicionar água destilada até 100 ml. Arrefece-se a solução de paraformaldeído a 4 utilizando um refrigerador ° C. </li></ol></li><li> Dissolve-se 125 mg de VA-044 iniciador em 26.25 ml de água destilada e arrefece-se a solução em um 4° C frigorífico. </li><li> Arrefecer 5 ml de solução de acrilamida (40%), 1,25 ml de solução de bis (2%) (CUIDADO: Bis e soluções de acrilamida são tóxicos, podem causar defeitos genéticos ou cancro), 5 ml de PBS 10X, ml de 16% solução de 12,5 PFA, eo solução de iniciador VA-044 em gelo. </li><li> Misturar as soluções acima em gelo. Nota: O volume total é 50 ml. </li></ol></li><li> Perfusão e Colecção do Cordão do rato Spinal <ol><li> Anestesiar um rato com uma injecção intraperitoneal de cetamina (80 mg / kg) e xyzaline (5 mg / kg) diluído em 0,9% de solução salina normal. A dosagem para rato pode ser menor do que para os ratos. </li><li> Em uma superfície de plástico adequado em um exaustor, corrigir os membros do mouse para longe do corpo (fita adesiva é eficaz) uma vez que o mouse não responder a uma pitada a sua pele do pé. </li><li> Cortar a pele do rato sobre o peito e depois do peito osso para expor o coração com uma tesoura. </li><li> Usando uma bomba peristáltica, com uma agulha de 25 G ligada a the das extremidades do tubo, inserir a agulha no ventrículo esquerdo do coração, cortar o átrio direito para criar um ponto de saída para o sangue e, em seguida, iniciar a lavagem com 40 ml de solução salina normal a 0,9% de uma taxa de aproximadamente 10 ml / min. </li><li> Quando completo, perfundir o mouse com 35 solução de hidrogel frio ml de gelo. Nota: perfundir a uma velocidade de 10 ml / min para evitar inchaço do tecido nervoso. </li><li> Faça uma incisão na pele sobre a parte de trás com uma lâmina e em seguida, remover os músculos ao lado da vertebrados. Após o corte dos arcos vertebrais de um lado e lançando-os para o outro lado, levar a medula espinal da coluna vertebral com uma tesoura fina. </li></ol></li><li> Limpando o cabo do rato Spinal <ol><li> Coloque a espinal medula em 15 ml de solução de hidrogel durante a noite a 4 ° C. </li><li> Retirar 10 mL da solução de hidrogel e despeje o resto da solução com os segmentos da medula espinhal de um tubo de 5 ml. Encher o tubo de 5 ml com a solução de hidrogel até que esteja completamente cheia. </li><li> Estende um pequeno pedaço de parafilme sobre a parte superior do tubo de 5 ml e, em seguida, enrolar em torno do pescoço parafilme do tubo. Nota: Certifique-se os contactos parafilme a solução de hidrogel e não há bolhas entre a solução de hidrogel e o parafilme. </li><li> Colocar o tubo de 5 ml de uma 37 &#160; ° C forno durante a noite. Observe a solução hidrogel até que se torna um gel. </li><li> Aqui o tubo de 5 ml de sair do forno e remover o gel a partir do tubo com uma chave de fendas (por exemplo, uma ponta de pipeta de 1 ml pá). Remover o gel do tecido da medula espinhal furando o tecido grosseiro ao gel e em seguida, tomar o tecido grosseiro de distância (o gel vai ficar com o tecido e, portanto, serão removidos pelo tecido). </li><li> Após a remoção do gel a partir do tecido da medula espinal, lava-se a medula espinal 4 vezes com 1x de PBS (pH 7,4) durante 24 horas num agitador. </li><li> Preparar a solução de compensação CUBIC por dissolução de 3,85 g de ureia e 3,85 g de N, N, N &#39;, N&#39;-tetraquis (2-hidroxipropil) ethylenediamine em 5,38 ml de água destilada. Nota: Um agitador quente é usado. Adicionar 2,31 g de polietileno-glicol mono-p-isooctylphenyl éter / Triton X-100 à solução, uma vez que é clara e arrefece até à temperatura ambiente. Nota: 10 g destes três produtos químicos são para um rato. </li><li> Cortar a medula espinhal de rato coronariamente em longos segmentos de 2-3 mm com uma lâmina de barbear e colocá-los em 5 ml da solução de limpeza CUBIC acima. Colocar num agitador no forno a 37 &#160; ° C durante 3 dias. </li><li> Três dias mais tarde, mudar a solução de limpeza para uma fresca. Nota: a solução acima é preparada antes da utilização. </li><li> Dois a três dias mais tarde depois de atualizar a solução de compensação cúbicos, verificar a transparência do tecido contra um papel com tipo de letra 8 letras. Se é transparente, as letras podem ser vistas através do tecido limpo. Remover a solução de compensação e adicionar 4 ml de PBST (0,1% de Triton-X100) para lavar o tecido 4 vezes por dia (a cada 6 horas). </li></ol></li></ol><ol><li> Incubar os segmentos da medula espinhal na solução de anticorpo primário (anti-serotonina, produzidos em coelho, diluído a 1: 100 em PBST) durante 3 dias num agitador numa estufa de 37 ° C. </li><li> Remover a solução de anticorpo e adicionar 4 ml de PBST (0,1% de Triton-X100) para lavar os segmentos da medula espinhal 4 vezes por dia (a cada 6 horas) num forno de 37 ° C. </li><li> Remover a solução de PBST, e adicionar a solução de anticorpo secundário (594 anti-IgG de coelho de cabra conjugada, diluídos 1: 100 em PBST) a incubar o tecido durante 3 dias num agitador numa estufa de 37 ° C. </li><li> Remover a solução de anticorpo secundário e adicionar 4 ml de PBST para lavar os segmentos da medula espinhal 4 vezes por dia (a cada 6 horas) num agitador num forno de 37 ° C. No dia seguinte, o tecido é preparado para imagiologia. </li></ol> 3. Imagem <ol><li> Remover a solução de PBST e adicionar 4 ml de 85% de glicerol para tornar o índice de refracção do tecido mesmo. </li><li&gt; Verifique a clareza do tecido. Uma vez que é evidente em aproximadamente 1 hora, colocar o tecido para uma grande lâmina de vidro, fino que se encaixa o quadro realização da multi-foto fluorescente microscópio. </li><li> Coloque algumas gotas de 85% de glicerol ao lado do tecido da medula espinhal e que lamela de vidro com uma lamela de 22 x 50 mm 2. Nota: A orientação da medula espinhal decide como a imagem se parece. </li><li> Colocar a lâmina de vidro sobre a armação de retenção do microscópio fluorescente multi-fotão e mover o tecido para a via de luz. </li><li> Escolha do Hélio Néon linha de laser 594 nm e ajustar a intensidade do laser para o nível óptimo, verificando o brilho do sinal positivo na imagem real. </li><li> Seleccione a área de leitura do tecido da medula espinal utilizando a objectiva de 20x (em água, NA 0,7) e preparam-se para criar um z-pilha, e a profundidade de cada degrau definir a 3 um. </li><li> Digitalizar o tecido a partir do topo para o fundo da pilha-Z sob o objec 20Xtiva e, em seguida, objectiva 63X (em óleo, NA 1.4) (tamanho do passo foi de 1? M), respectivamente. Reconstruir vídeos em 3D usando um software 3D 28. </li></ol>

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The authors have nothing to disclose.

O protocolo descrito mostra como fibras imagem serotoninérgicos na medula espinhal de rato com a clareza combinadas e técnicas cúbico. Ele apresenta um processo de limpeza mais rápido em comparação com o protocolo de compensação passiva desenvolvido por Cheung et al. 14 e Tomer et ai. 15 e permite que o tecido da medula espinal a ser bem suportado pelo hidrogel durante a limpeza. Um passo importante durante a fixação da medula espinhal de rato, como relatado por Cheung et al. 14 e Tomer et ai. 15, é a de manter todas as soluções arrefecer em gelo, o que impede a polimerização dos produtos químicos. O outro passo crucial é o rato para perfundir-se lentamente com a solução de hidrogel, a fim de evitar o inchaço do tecido nervoso. Para a solução de hidrogel a tornar-se um gel, de desgaseificação é necessária. Como uma alternativa a isso, utilizou-se parafilme a cobrir a parte superior do tubo e deixaram as bolhas de ar entre as empresolução Rogel eo parafilme. Este sistema funcionou muito bem e a solução tornou-se um gel de hidrogel depois de algumas horas. Embora algumas bolhas foram encontrados no gel, que não interferiu com as experiências seguintes, o que foi demonstrado pela ausência de bolhas no tecido da medula espinal. Clearing é o passo mais crítico, e requer uma combinação de reagentes. A solução de limpeza, que contém amino-álcool, remove o lípido do tecido da medula espinhal muito mais rápido do que o / solução de compensação de SDS bórico base de ácido utilizado no protocolo original de 16 Clareza. A vantagem de clareza e cúbicas em relação às técnicas de imuno-histoquímica tradicionais é que a totalidade do bloco de tecido podem ser visualizados sem a secção do tecido, como um resultado, a continuidade das fibras serotoninérgicos é retida, e é possível seguir a mesma fibra por uma longa distância dentro do Z-pilha. Com esta vantagem, a colateralização pode ser confiantemente identificados com as imagens 3D, umND novos conclusões podem, portanto, ser feita acerca da natureza das vias serotoninérgicos. O protocolo descrito é uma ferramenta conveniente para investigação de sistemas de fibras, bem como os núcleos por simplesmente rotular o tecido com anticorpos específicos. Esta técnica é também uma escolha ideal para interrogar sistemas diferentes no mesmo cérebro de camundongo usando marcação dupla ou tripla. No nosso ambiente de laboratório, um multi-fotão microscópio fluorescente estava disponível com uma distância de trabalho de aproximadamente 300 um. Isto limita a imagem para um máximo de 600 um, se ambos os lados do tecido são digitalizados. No entanto, uma folha de microscópio de fluorescência de luz, pode imagem de todo o cérebro do rato e da medula espinhal através da sua largura, comprimento e profundidade. A outra vantagem desta técnica é que os anticorpos podem ser eluída e, em seguida, um outro anticorpo ou anticorpos diferentes aplicada novamente para o mesmo tecido. Isso reduz o uso de animais consideravelmente, bem como os custos associados com a preparação de um novo tecido cerebralpara cada experimento. Pela mesma razão, esta técnica pode ser aplicada a outros tecidos. Houve já estudos realizados sobre os pulmões, rins, coração, intestino e tecido tumoral 26,27. Também existem limitações para esta técnica, por exemplo, os núcleos da rafe na parte posterior do cérebro do rato contém muitos tipos de neurônios, incluindo neurônios GABAérgicos. Embora Clareza e cúbicas permitirá o estudo da distribuição dos diversos tipos de fibras raphespinal num cérebro do rato e da medula espinhal. Esses neurônios, que não são identificados especificamente pelos seus neurotransmissores ou marcadores de proteínas, não pode ser identificado neste cenário. Uma alternativa é a rotular os neurónios específicos ou sistemas de fibras com sondas de ácidos nucleicos, tais como na hibridação in situ. Este é, por outro lado, limitada por o sistema de rotulagem e os filtros disponíveis para imagiologia fluorescente. Imagiologia todas as fibras raphespinal, portanto, permanece um desafio.Se estas técnicas foram combinadas com uma injecção intracraniana de um traçador anterógrada (tais como Phaseolus vulgaris leucoagglutinin - PHA-L), pode ser possível ver as fibras, fibras GABAérgicas serotonérgicos, e outros.

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

fibras descendentes longos para a medula espinhal são essenciais para a locomoção, a percepção da dor, e outros comportamentos. O padrão de terminação da fibra na medula espinal da maioria destes sistemas de fibra não foram cuidadosamente investigados em qualquer espécie. fibras serotoninérgicos, que se projectam para a espinal-medula, foram estudados em ratos e em secções histológicas marsupiais e o seu significado funcional foi deduzida com base no seu padrão de terminação da fibra na medula espinal. Com o desenvolvimento de clareza e técnicas cúbicos, é possível investigar este sistema de fibra e a sua distribuição na medula espinal, a qual é susceptível de revelar características anteriormente desconhecidos de vias supraespinhais serotoninérgicos. Aqui, nós fornecemos um protocolo detalhado para geração de imagens das fibras serotoninérgicos na medula espinhal de rato usando o CLAREZA combinados e técnicas cúbicos. O método envolve a perfusão de um rato com uma solução de hidrogel e clarificação do tecido com uma combinção de limpar reagentes. tecido da medula espinhal foi liberado em pouco menos de duas semanas, e a coloração de imunofluorescência subsequente contra a serotonina foi concluída em menos de dez dias. Com um microscópio de fluorescência multi-fotão, o tecido foi digitalizado e uma imagem 3D foi reconstruído usando software Osirix.

Esta secção mostra os resultados de coloração de anticorpos de serotonina na medula espinal do rato transparente usando uma combinação de a clareza e protocolos cúbico. Mostramos que as fibras serotoninérgicas estão presentes em todas as lâminas da medula espinhal com predominância na porção ventral do corno ventral (Figura 1, também vemos Video 1). O tecido de controlo não têm fibras positivas (resultado não foi apresentado). No corno ventral, fibras serotoninérgicas densamente estão presentes na porção ventromedial do corno anterior e se estendem para a parte lateral do corno anterior (Figura 2, também veja vídeo 2). Algumas fibras imunopositivas também se estendem a partir do corno anterior para o corno dorsal ou para o canal central. imunopositivas fibras estão presentes em todas as lâminas do corno dorsal, mas há uma pequena diferença entre as fibras positivas na lâmina 2 e 4, especialmente nas lateraisparte do corno dorsal. A maioria das fibras serotonérgicos no corno dorsal são de diâmetro reduzido e apenas um pequeno número delas são de grande diâmetro (de Vídeo 1). Em uma secção horizontal, fibras serotoninérgicos densamente empacotados no corno ventral foram observadas a viajar ao longo do eixo longitudinal da medula espinhal e formam uma grande feixe de fibras. A descoberta mais interessante é que este grande feixe de fibras de ramificações emite regularmente ao longo do eixo longitudinal, os quais são perpendiculares ao feixe de fibras. Estes ramos hipotecar ainda mais ao longo de seus caminhos para a parte lateral do corno anterior. Em comparação com estes ramos, os que se prolongam no sentido da linha média são irregulares e menores (Figura 2). No âmbito do objectivo 63X, as fibras serotonérgicos no corno dorsal foram observados a viajar ao longo do eixo da medula espinhal e terminam em vários pontos ao longo do caminho do trato. As fibras grossas são esporadicamente distribuídos entre os f fina<html> IBERS (Figura 3, também ver vídeo 3). <img alt="figura 1" src="/files/ftp_upload/53673/53673fig1.jpg" /> Figura 1:. Fibras de serotonina num corte coronal da medula lombar no 200X Esquerda é o corno anterior, certo é o corno dorsal, dorsal é a parte medial da medula espinhal, e ventral é a parte lateral da medula espinhal. A barra de escala é de 100 mm. fibras serotoninérgicos estão presentes em todas as lâminas, especialmente na parte ventromedial do corno anterior, onde a densidade das fibras é alta. Estas fibras prolongar-se para tanto a parte lateral do corno anterior e o canal ou dorsal chifre central. A densidade de fibras na outra lâminas é baixo. A maioria das fibras serotoninérgicas são finas. Apenas um pequeno número de fibras são grossas e eles são vistos tanto no corno dorsal e corno anterior (&gt; ver também Video 1 (Clique direito a download)). <img alt="Figura 2" src="/files/ftp_upload/53673/53673fig2.jpg" /> Figura 2:. Fibras serotonérgica a porção ventral do corno ventral a uma ampliação de 200X esquerda e direita estão os lados laterais da medula espinal, dorsal, é a porção rostral da medula espinal, e ventral é a parte caudal da medula espinhal. A barra de escala é de 100 mm. fibras serotoninérgicos são mostrados viajando ao longo do eixo longitudinal da medula espinhal e embalado na porção medial do corno ventral onde formam um feixe de fibras de espessura. Fora deste pacote, alguns ramos são estendidos em intervalos regulares para a parte lateral do corno anterior, enquanto outros que se estendem em direção à linha média são irregulares e mais curtos. Essas fibras terminam em muitos pontos ao longo do caminho do trato ( <a href="https://www-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/53673/Video2.mp4">ver também Video 2</a>(Clique direito a download)). <img alt="Figura 3" src="/files/ftp_upload/53673/53673fig3.jpg" /> Figura 3: fibras Serotoninérgicos no corno dorsal sob a 630X ampliação Esquerda é a parte lateral da medula espinhal, certo é a parte medial da medula espinhal, dorsal e ventral referem-se à dorsal e parte ventral do corno dorsal, respectivamente. . A barra de escala é de 7 mm. fibras serotoninérgicos viaje ao longo do eixo longitudinal da medula espinhal e terminam ao longo do caminho do trato. Estas fibras são, principalmente, de pequeno diâmetro, com um pequeno número de fibras espessas de permeio. Estas fibras raramente se sobrepõem uns com os outros ( <a href="https://www-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/53673/Video3.mp4">ver também Video 3 (Clique direito a 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.

Imagiologia Fibras Serotoninérgicos na medula espinhal mouse usando o CLARIDADE / Técnica CUBIC

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

Universidad San Sebastian

Research

JoVE Journal

Neuroscience

10.4K visualizações.  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.

Vídeo: 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.

Em Imaging vivo do Cordão do rato Spinal Usando Two-photon Microscopia

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 ...

Neurociência

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.

Um Processo de Implantação de uma Câmara Spinal para Longitudinal<em&gt; In Vivo</em&gt; Criação de Imagens do Cordão do rato Spinal

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.

Dois fótons de imagem da Cellular Dynamics na Medula Espinhal Rato

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 óptica do Sistema Nervoso Central Rato Usando a claridade passiva

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.

Imunostaining de Retinas de Monte Inteiro com o Método de Limpeza tecidual 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.

Imagem e quantificação de dendritos neuronais intactos via clareza de tecido

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.

Um método de limpeza de tecidos para imagens neuronais 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.

Estabelecendo um Modelo de Lesão da Medula Espinhal por Contusão de Camundongo Baseado em uma Técnica Minimamente Invasiva

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.

Estabelecimento do Modelo de Síndrome do Cordão Central em Camundongos C57BL/6J

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 ...