Os autores gostariam de agradecer ao Dr. Alistair Garratt (Max Delbrück Center, Berlin) para comentários úteis. Este trabalho foi financiado pelo centro de pesquisa colaborativa (SFB665) do Conselho de Pesquisa Alemã (DFG).

1. Procedimento de dissecção Nota: O uso experimental de camundongos devem seguir as orientações aprovadas oficialmente para o cuidado e uso de animais de laboratório. <ol><li> Antes da preparação, configure seu microscópio de dissecação e lay out dos instrumentos cirúrgicos necessários para a dissecção incluindo tesouras grandes e pequenas, grandes pinças dentadas, pinças curvas e quatro conjuntos de Dumont No.5 dissecar fórceps (dois dos quais no interior polido sugestões) ( para mais detalhes veja a tabela de reagentes e equipamentos específicos). Coloque uma folha de papel de filtro em uma de 100 mm placa de Petri Sylgard revestido. Despeje frio PBS nos poços de uma placa de 12 poços, um de 100 mm placa de Petri e um tubo de 12 ml e deixar no gelo. Pipetar 2 ml de tampão de fixação (paraformaldeído 4% em PBS, pH 7,4) em cada poço de uma placa de 12 segundo poço e colocar no gelo. </li><li> Depois de anestesiados, o sacrifício da barragem cronometrado grávida no E12.5 (detecção de plug vaginal foi designado como E0.5), coloque o mousecom o seu lado ventral para cima em três folhas de papel toalha e deixe a área abdominal com etanol 70%. </li><li> Abra a cavidade abdominopélvica, isolar os chifres bilateral uterina e transferi-los para um dos pratos de 100 mm com gelado PBS. </li><li> Sob controle microscópico, faça uma incisão longitudinal da parede uterina com pequenos, tesoura reta e cortar cada saco amniótico da placenta. </li><li> Descascar o saco amniótico de cada embrião e cortar o cordão umbilical. </li><li> Com uma tesoura pequena, decapitar os embriões e colocar a cabeça ou uma parte deles de lado para o isolamento de DNA genômico se genotipagem deve ser exigido. Posteriormente, transferir o torsi aos poços respectivos da placa de 12 bem com o frio PBS. </li><li> Molhar o papel de filtro no prato Sylgard com algumas gotas de PBS e posição de um tronco embrionárias com o lado dorsal até no papel. Para melhor estabilidade, acertar a sua cauda e extremidades do corpo. </li><li> Trabalharsob um microscópio de dissecação com uma ampliação de cerca de 16x, com cuidado prenda a pele do embrião acima da medula espinhal com dois pares de pinças com ponta fina e delicadamente desagregá-la. Começando no meio, vá primeiro para a cauda e, em seguida, retomar a partir do meio para o lado anterior. </li><li> Molhado o embrião de vez em quando com duas ou três gotas de PBS para impedir que a secagem. </li><li> Para evitar que DRG são teared fora da medula espinhal quando ela é removida do embrião destacar o DRG e da medula espinhal em torno da coluna vertebral cartilaginosa por movimentos deslizantes horizontal com a lâmina de uma pinça bem com interior polido dicas. A partir de meados do lado direito do embrião, trabalhar o seu caminho em direção à cauda e, em seguida, até o final anterior. Depois, repita o procedimento no outro lado. Tenha cuidado para não arrancar a DRG associados. </li><li> Para retirar completamente o cabo soltou da coluna vertebral do embrião, pegar seu cérvicoal peça com uma pinça fina e puxe-o em uma única peça para o seu fim caudal. </li><li> Submergir o cabo isolado da coluna vertebral com DRG anexado em um poço da placa de 12 bem preenchido com tampão de fixação e prosseguir com a preparação de medulas espinhais dos embriões restantes. </li><li> Fix medula espinhal pelo menos duas horas no gelo. </li></ol> 2. DII-rotulagem de neurônios DRG <ol><li> Para cada medula espinhal para ser rotulado, prepare duas agulhas de vidro com um puxador de micropipeta. Nós usamos um modelo de Sutter extrator P-97 eletrodo (configuração do programa: 1 HEAT = 625, VEL = 35 TEMPO = 175; 2 HEAT = 600, VEL = 25 TEMPO = 175; HEAT 3 = 630, = VEL... 30 TIME, = 150). </li><li> Sob controle microscópico molhe a ponta de cada agulha de vidro 3-5 vezes em uma solução de 5% (w / v) de DII em etanol. Evaporação do etanol cria uma fina camada de cristais de DII na agulha. </li><li> Trabalhando sob um microscópio de dissecação com uma ampliação de aproximadamente 40x, coloque uma medula espinhal com seus vlado entral up em um slide. Usando uma tesoura primavera, abrir a cabo no lado ventral em todo o seu comprimento cortando o floorplate para permitir a posterior montagem da preparação em um modo de livro aberto invertido (ver secção C). </li><li> Posição da medula espinhal com o lado dorsal até em um slide. Em seguida, manualmente abordagem o cabo com a ponta coberta de DII da agulha de vidro sob visualização microscópica e cuidadosamente furar DRG cada segundo de um lado da medula espinhal. Quase nenhum vestígio de DII deve ser visível no DRG perfurado para garantir a rotulagem de apenas um pequeno número de neurônios. Dê uma segunda agulha para o outro lado do cabo. Visando apenas a cada segundo DRG evita uma sobreposição de rotulados projeções axonal da DRG vizinhas que podem complicar a diferenciação de axônios individuais. </li><li> Retorno da medula espinhal para o buffer de fixação e incubar no escuro por pelo menos seis horas à temperatura ambiente ou durante a noite a 4 ° C para dar tempo para o corante para diffuse ao longo do axônio dentro da membrana plasmática. Permitir, por um período de incubação mais longo (até dois dias a 4 ° C) se a análise de crescimento garantia para a medula espinhal é desejada. </li></ol> 3. Análise microscópica de montagem e <ol><li> Seguintes corante difusão posição, um único cabo spinal com o lado dorsal para baixo em uma lamela em uma gota de PBS. Tome cuidado para que o DRG anexados são corretamente orientados lateralmente e não se misturam. Aspirar o excesso de fluido e achatar o cabo em um modo de livro aberto invertido com a pinça. </li><li> Monte a preparação sobre uma lâmina de microscópio usando PBS. </li><li> Para evitar o aumento indesejado de análise de fluorescência de fundo coloração microscópica de rotulados projeções axonal deve ser realizada no dia da montagem. Até então, manter os slides no escuro a 4 ° C. </li></ol> 4. Resultados representativos: A medula espinhal do rato recebe pr aferentesojections de 8 pares de colo do útero, 13 pares de torácicos, 5 pares de pares lombar e sacral DRG 4 de um total de 60 gânglios espinhais. Depois de algum treino uma medula espinhal com a maioria dos DRG ainda ligado pode ser isolado de um embrião em menos de cinco minutos. Este procedimento é adequado para o isolamento de medulas espinhais com DRG em anexo a partir de embriões E13.5 E11.5 para mouse. No entanto, melhores resultados são obtidos a partir E12.5. Resultados exemplar do processo de rotulagem descritos aqui são mostrados na Figura 3. A rotulagem dos DRG em toda a extensão da medula espinhal podem então ser usados ​​para quantificar o comportamento de ramificação axonal em diferentes níveis vertebrais (Fig. 4). <img alt="Figura 1" src="/files/ftp_upload/3667/3667fig1.jpg" /> Figura 1. Esquema representando os dois principais modos de ramificação axonal. (A) Desvios pelas atividades do cone de crescimento que poderia levar a uma bifurcação - como indicado aqui - assim como arbors terminal complexos ou(B) formação de colaterais no eixo axônio (intersticial ramificação). O último modo é o tipo dominante de ramificação de projetar axônios corticais e tálamo-cortical 6,7. <img alt="Figura 2" src="/files/ftp_upload/3667/3667fig2.jpg" /> Figura 2 projeções aferentes dos neurônios DRG para a medula espinhal embrionárias mostrar os dois tipos de ramificação axonal:. Axônios primeira filial na DREZ pela bifurcação (1) e da forma filha ramos colaterais resultantes após um período de espera por intersticial ramificação (2). <img alt="Figura 3" src="/files/ftp_upload/3667/3667fig3.jpg" /> Figura 3. Visualização de trajetórias individuais axonal de neurônios DRG embrionárias mouse. (A) da medula espinhal com DRG anexado preparados a partir de um embrião de rato E12.5. (Barra Escala, 1 mm.) (BD) vistas dorsal da DII marcado DRG de camundongos selvagens com ampliação crescente são mostrados. Em B a cada DRG segundo é labeled por DII. Imagens de fluorescência são invertidos, caudal está à esquerda e em C e D, lateral está no fundo. Em C um pequeno número de axônios é rotulado e em maior ampliação da presença de T-like ramos podem ser identificados no DREZ da medula espinhal. (Escala de barras, 250 mM (B), 100 mm (C) e 50 mm (D).) <img alt="Figura 4" src="/files/ftp_upload/3667/3667fig4.jpg" /> Figura 4. Quantificação dos ramos em forma de T, única gira rostral ou caudal no tipo selvagem e C-peptídeo natriurético do tipo (CNP) camundongos deficientes em E13.5. O número de axônios única contados são dadas entre parênteses para os níveis de tronco diferentes para cada genótipo. C-ou-R voltas - o crescimento somente na direção caudal ou rostral, respectivamente. <img alt="Figura 5" src="/files/ftp_upload/3667/3667fig5.jpg" /> Figura 5. A GMPc desencadeia sinalização bifurcação axônio sensorial na DREZ da medula espinhal. (A) Esquema dea via de sinalização composta cGMP do ligante CNP, o receptor de guanilato ciclase Npr2 ea serina / treonina quinase cGKIα em neurônios DRG embrionárias. Npr2 gera cGMP de GTP com a estimulação pelo CNP. (B) Dii rastreamento de axônios única de neurônios DRG em camundongos wild-type e CNP-deficiente. (Barra Escala, 25 mm.)

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Padrões estereotipados de projeção composto por dois tipos de formação ramo axonal juntamente com facilidade de preparação em combinação com o uso de tecido fixo para DII rotulagem faz o cabo embrionárias espinhal com DRG anexado um modelo favorável ao estudo axonal ramificação. A aplicação de pequenas quantidades de DII usando agulhas de vidro revestido permite - em contraste com a rotulagem grosso do DRG - a visualização de pequenos grupos de neurônios DRG e, assim, a análise de axônios individuais...

<table border="1"><tbody><tr><td> Nome do reagente </td><td> Companhia </td><td> Número de catálogo </td><td> Comments (opcional) </td></tr><tr><td> Estereomicroscópio Stemi RDC </td><td> Zeiss </td><td></td><td></td></tr><tr><td> Solução tampão fosfato (PBS) </td><td> Biochrom AG </td><td> L182-50 </td><td></td></tr><tr><td> Paraformaldeído </td><td> Merck </td><td> 8.18715.1000 </td><td></td></tr><tr><td> Tesoura cirúrgica padrão </td><td> Multa Ferramentas Ciência </td><td> 14001-13 </td><td></td></tr><tr><td> Dentada forceps padrão </td><td> Multa Ferramentas Ciência </td><td> 11021-14 </td><td></td></tr><tr><td> Extra fino tesoura de íris </td><td> Multa Ferramentas Ciência </td><td> 14088-10 </td><td></td></tr><tr><td> Fórceps curvos </td><td> Multa Ferramentas Ciência </td><td> 11003-13 </td><td></td></tr><tr><td> Dumont No.5 multa forceps dicas </td><td> Multa Ferramentas Ciência </td><td> 11254-20 </td><td></td></tr><tr><td> Dumont No.5 acabamento espelhado forceps </td><td> Multa Ferramentas Ciência </td><td> 11252-23 </td><td></td></tr><tr><td> Vannas-Tübingen Primavera tesoura </td><td> Multa Ferramentas Ciência </td><td> 15008-08 </td><td></td></tr><tr><td> Papel-filtro </td><td> Fisher Scientific </td><td> FB59041 </td><td></td></tr><tr><td> Sylgard 184 </td><td> Precisão Instrumentos mundo </td><td> SYLG184 </td><td></td></tr><tr><td> 100-mm placas de Petri </td><td> Greiner </td><td> 663102 </td><td></td></tr><tr><td> 12 ml de tubos de polipropileno </td><td> Carl Roth GmbH </td><td> ECO3.1 &lt;/ Td&gt; <td></td></tr><tr><td> 12 placa de cultura bem </td><td> Becton Dickinson </td><td> 35-3043 </td><td></td></tr><tr><td> Etanol </td><td> Merck </td><td> 1.00983.2500 </td><td></td></tr><tr><td> Flaming / Brown micropipeta extrator P-97 </td><td> Sutter Instrument Co. </td><td></td><td></td></tr><tr><td> Vidro borosilicato capilares </td><td> Harvard Apparatus </td><td> 30-0066 </td><td></td></tr><tr><td> DII (1,1 &#39;-Dioctadecyl-3, 3,3&#39;, 3&#39;-tetrametil - indocarbocyanine perclorato) </td><td> Sigma-Aldrich </td><td> 468495 </td><td></td></tr><tr><td> Microscópio slides SuperFrost Plus </td><td> Carl Roth GmbH </td><td> H867.1 </td><td></td></tr><tr><td> Tampa de vidro desliza </td><td> Carl Roth GmbH </td><td> 1870,2 </td><td></td&gt; </tr></tbody></table>

dii-labeling of drg neurons to study axonal branching in a whole mount preparation of mouse embryonic spinal cord

Apresentamos aqui uma técnica para identificar as trajetórias de pequenos grupos de neurônios DRG para a medula espinhal de embriões através da coloração difusivo usando o traçador lipofílico 1,1 &#39;-dioctadecyl-3, 3,3&#39;, 3&#39;-tetramethylindocarbocyanine perclorato (DII) 1 . A comparação das vias axonal de tipo selvagem com os de linhas de rato em que os genes são mutados permite testes para um papel funcional de proteínas candidato no controle de ramificação axonal que é um mecanismo essencial na fiação do sistema nervoso. Ramificação axonal permite que um neurônio individual para se conectar com múltiplos alvos, fornecendo assim a base física para o processamento paralelo de informações. Ramificações em regiões objectivo intermédio de crescimento axonal pode ser diferenciado de arborização terminal. Além disso, diferentes modos de formação ramo axonal pode ser classificada dependendo de resultados de ramificação das atividades do cone de crescimento (divisão ou bra adiadanching) ou a partir do brotamento de colaterais a partir do eixo do axônio em um processo chamado de ramificação intersticial 2 (Fig. 1). As projeções centrais de neurônios da DRG oferecer um sistema útil experimental para estudar os dois tipos de ramificação axonal: quando seus axônios aferentes alcançar a zona de entrada da raiz dorsal (DREZ) da medula espinhal entre os dias embrionários 10-13 (E10 - E13) que exibir um padrão estereotipado de T ou em forma de Y bifurcação. Os dois axônios filha resultando então prosseguir em direções rostral ou caudal, respectivamente, na margem dorsolateral da medula e só após um período de espera colaterais brotam esses axônios-tronco para penetrar na massa cinzenta (intersticial ramificação) e do projeto para os neurônios de revezamento em específico lâminas da medula espinhal, onde eles ainda arborize (terminal de ramificação) 3. Traçados DII revelaram cones de crescimento na zona da raiz dorsal entrada da medula espinhal que pareciam estar no processo de dividir o que sugere que bifurcação é causada pela divisão do cone de crescimento em si 4 (Fig. 2), no entanto, outras opções foram discutidas e 5. Este vídeo demonstra como primeiro dissecar a medula espinhal de ratos E12.5 deixando o DRG em anexo. Fixação dos montantes seguintes diminuto exemplar da DII são aplicadas a DRG usando agulhas de vidro retirado tubos capilares. Após uma etapa de incubação, o cordão espinhal é rotulada montado como uma preparação de livro aberto invertido para analisar axônios individuais usando microscopia de fluorescência.

Watch this Scientific Journal Video about DiI-Labeling of DRG Neurons to Study Axonal Branching in a Whole Mount Preparation of Mouse Embryonic Spinal Cord at JoVE.com

DiI-Labeling of DRG Neurons to Study Axonal Branching in a Whole Mount Preparation of Mouse Embryonic Spinal Cord

As projeções estereotipadas de aferentes sensoriais para a medula espinhal de roedores oferecem um sistema de fácil acesso experimental para o estudo de ramificação axonal através do rastreamento de axônios individuais.

DII-rotulagem de neurônios DRG para estudar em uma ramificação axonal Preparação Monte Whole of Mouse Embryonic Spinal Cord

Here we  present a technique to label the trajectories of small groups of DRG neurons  into the embryonic spinal cord by diffusive staining using the ...

dii-labeling-drg-neurons-to-study-axonal-branching-whole-mount

Research

JoVE Journal

Neuroscience

24.8K visualizações.  Max Delbrück Center for Molecular Medicine. As projeções estereotipadas de aferentes sensoriais para a medula espinhal de roedores oferecem um sistema de fácil acesso experimental para o estudo de ramificação axonal através do rastreamento de axônios individuais.

Vídeo: DiI-Labeling of DRG Neurons to Study Axonal Branching in a Whole Mount Preparation of Mouse Embryonic Spinal Cord

Dissection and Culture of Commissural Neurons from Embryonic Spinal Cord

Commissural neurons have been widely used to investigate the mechanisms underlying axon guidance during embryonic spinal cord development. The cell bodies of these neurons are located in the dorsal spinal cord and their axons follow stereotyped trajectories during embryonic development. Commissural axons initially project ventrally towards the floorplate. After crossing the midline, these axons turn anteriorly and project towards the brain. Each of these steps is regulated by the action of several guidance cues. Cultures highly enriched in commissural neurons are ideally suited for many experiments addressing the mechanisms of axon pathfinding, including turning assays, immunochemistry and biochemistry. Here, we describe a method to dissect and culture commissural neurons from E13 rat dorsal spinal cord. First, the spinal cord is isolated and dorsal strips are dissected out. The dorsal tissue is then dissociated into a cell suspension by trypsinization and mechanical disruption. Neurons are plated onto poly-L-lysine-coated glass coverslips or tissue-culture dishes. After 30 hours in vitro, most neurons have extended an axon. The purity of the culture (Yam et al. 2009), typically over 90%, can be assessed by immunolabeling with the commissural neuron markers DCC, LH2 and TAG1 (Helms and Johnson, 1998). This neuronal preparation is a useful tool for in vitro studies of the cellular and molecular mechanisms of commissural axon growth and guidance during spinal cord development.

This video demonstrates a method to dissect and culture commissural neurons from E13 rat dorsal spinal cord. Dissociated commissural neurons are useful to study the cellular and molecular mechanisms of axon growth and guidance.

Dissecção e Cultura de Neurônios da Medula Espinhal comissural embrionárias

Commissural neurons have been widely used to investigate the mechanisms underlying axon guidance during embryonic spinal cord development. The cell bodies ...

Neurociência

Visualization of the Embryonic Nervous System in Whole-mount Drosophila Embryos

The Drosophila embryo is an attractive model system for investigating the cellular and molecular basis of neuronal development. Here we describe the procedure for the visualization of Drosophila embryonic nervous system using antibodies to neuronal proteins. Since the entire embryonic peripheral nervous and central nervous systems are well characterized at the level of individual cells (Dambly-Chaudi&egrave;re et al., 1986; Bodmer et al., 1987; Bodmer et al., 1989), any aberrations to these systems can be easily identified using antibodies to different neuronal proteins. The developing embryos are collected at certain times to ensure that the embryos are in the proper developmental stages for visualization. After collection, the outer layers of the embryo, the chorion membrane and the vitelline envelope that surrounds the embryo, are removed before fixation. Embryos are then incubated with neuronal antibodies and visualized using fluorescently labeled secondary antibodies. Embryos at stages 12-17 are visualized to access the embryonic nervous system. At stage 12 the CNS germ band starts shortening and by stage 15 the definitive pattern of the commissure has been achieved. By stage 17 the CNS contracts and the PNS is fully developed (Campos-Ortega et al. 1985). Thus changes in the pattern of the PNS and CNS can be easily observed during these developmental stages.

Visualization of the Embryonic Nervous System in Whole-mount Drosophila Embryos

We describe the procedure to prepare staged Drosophila embryos for the visualization of the embryonic nervous system during embryogenesis.

Visualização do Sistema Nervoso embrionárias no Whole-mount Drosophila Os embriões

The Drosophila embryo is an attractive model system for investigating the cellular and molecular basis of neuronal development. Here we describe the ...

Whole Mount Preparation of the Adult Drosophila Ventral Nerve Cord for Giant Fiber Dye Injection

To analyze the axonal and dendritic morphology of neurons, it is essential to obtain accurate labeling of neuronal structures. Preparing well labeled samples with little to no tissue damage enables us to analyze cell morphology and to compare individual samples to each other, hence allowing the identification of mutant anomalies.
In the demonstrated dissection method the nervous system remains mostly inside the adult fly. Through a dorsal incision, the abdomen and thorax are opened and most of the internal organs are removed. Only the dorsal side of the ventral nerve cord (VNC) and the cervical connective 
(CvC) containing the big axons of the giant fibers (GFs)1 are exposed, while the brain containing the GF cell body and dendrites remains2 in the 
intact head. In this preparation most nerves of the VNC should remain attached to their muscles. 
Following the dissection, the intracellular filling of the giant fiber (GF) with a fluorescent dye is demonstrated. In the CvC the GF axons are located at the dorsal surface and thus can be easily visualized under a microscope with differential interference contrast (DIC) optics. This allows the injection of the GF axons with dye at this site to label the entire GF including the axons and their terminals in the VNC. This method results in reliable and strong staining of the GFs allowing the neurons to be imaged immediately after filling with an epifluorescent microscope. Alternatively, the fluorescent signal can be enhanced using standard immunohistochemistry procedures3 suitable for high resolution confocal microscopy.

Whole Mount Preparation of the Adult Drosophila Ventral Nerve Cord for Giant Fiber Dye Injection

An in vivo dissection of the adult Drosophila ventral nerve cord (VNC) is demonstrated. This particular dissection method causes little damage to the VNC allowing the subsequent labeling of the giant fiber neurons with fluorescent dye for high resolution imaging.

Preparação Monte todo o Adulto Drosophila Cordão nervoso ventral para injeção gigante de fibra Dye

To analyze the axonal and dendritic morphology of neurons, it is essential to obtain accurate labeling of neuronal structures. Preparing well labeled ...

An Ex Vivo Laser-induced Spinal Cord Injury Model to Assess Mechanisms of Axonal Degeneration in Real-time

Injured CNS axons fail to regenerate and often retract away from the injury site. Axons spared from the initial injury may later undergo secondary axonal degeneration. Lack of growth cone formation, regeneration, and loss of additional myelinated axonal projections within the spinal cord greatly limits neurological recovery following injury. To assess how central myelinated axons of the spinal cord respond to injury, we developed an ex vivo living spinal cord model utilizing transgenic mice that express yellow fluorescent protein in axons and a focal and highly reproducible laser-induced spinal cord injury to document the fate of axons and myelin (lipophilic fluorescent dye Nile Red) over time using two-photon excitation time-lapse microscopy. Dynamic processes such as acute axonal injury, axonal retraction, and myelin degeneration are best studied in real-time. However, the non-focal nature of contusion-based injuries and movement artifacts encountered during in vivo spinal cord imaging make differentiating primary and secondary axonal injury responses using high resolution microscopy challenging. The ex vivo spinal cord model described here mimics several aspects of clinically relevant contusion/compression-induced axonal pathologies including axonal swelling, spheroid formation, axonal transection, and peri-axonal swelling providing a useful model to study these dynamic processes in real-time. Major advantages of this model are excellent spatiotemporal resolution that allows differentiation between the primary insult that directly injures axons and secondary injury mechanisms; controlled infusion of reagents directly to the perfusate bathing the cord; precise alterations of the environmental milieu (e.g., calcium, sodium ions, known contributors to axonal injury, but near impossible to manipulate in vivo); and murine models also offer an advantage as they provide an opportunity to visualize and manipulate genetically identified cell populations and subcellular structures. Here, we describe how to isolate and image the living spinal cord from mice to capture dynamics of acute axonal injury.

An Ex Vivo Laser-induced Spinal Cord Injury Model to Assess Mechanisms of Axonal Degeneration in Real-time

We present a protocol utilizing two-photon excitation time-lapse microscopy to simultaneously visualize the dynamics of axon and myelin injuries in real time. This proposed protocol permits studies of both intrinsic and extrinsic factors which can influence central myelinated axon fate after injury and contribute to permanent clinical disability.

Um<em&gt; Ex Vivo</em&gt; Induzida por laser Lesão Medular modelo para avaliar mecanismos de axonal Degeneration em tempo real

Injured CNS axons fail to regenerate and often retract away from the injury site. Axons spared from the initial injury may later undergo secondary axonal ...

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 ...

Preparation of Primary Mixed Glial Cultures from Adult Mouse Spinal Cord Tissue

It has been well-accepted that spinal cord glial responses contribute significantly to the development of neuropathic pain. Tremendous information regarding glial activities at the cellular and molecular levels has been obtained through in vitro cell culture systems. The in vitro systems utilized, mainly include primary glia derived from neonatal brain cortical tissue and immortalized cell lines. However, these systems may not reflect the characteristics of spinal cord glial cells in vivo. In order to further investigate the roles of spinal cord glial cells in the development of peripheral nerve injury-induced neuropathic pain using a culture system that better reflects the in vivo condition, our laboratory has developed a method to establish primary spinal cord mixed glial cultures from adult mice. Briefly, spinal cords are collected from adult mice and processed through papain digestion followed by myelin removal with a density-gradient medium. Single cell suspensions are cultured in complete Dulbecco&#39;s modified Eagle media (cDMEM) supplemented with 2-mercaptoethanol (2-ME) at 35.9 oC. These culture conditions were optimized specifically for the growth of mixed glial cells. Under these conditions, cells are ready to be used for experimentation between 12 - 14 d&#160;(cells are usually in log phase during this time) after the establishment of the culture (D 0) and can be kept in culture conditions up to D 21. This culture system can be used to investigate the responses of spinal cord glial cells upon stimulation with various substances and agents. Besides neuropathic pain, this system can be used to study glial responses in other diseases that involve pathological changes of spinal cord glial cells.

The development of neuropathic pain involves pathological changes of spinal cord glial cells. A reliable glial culture system derived from adult spinal cord tissue and designed for studying these cells in vitro is lacking. Therefore, we show here how&#160;to establish primary mixed glial cultures from adult mouse spinal cord tissue.

Preparação de culturas mista glial primários de rato adulto Spinal Cord Tissue

It has been well-accepted that spinal cord glial responses contribute significantly to the development of neuropathic pain. Tremendous information ...

A Simple One-step Dissection Protocol for Whole-mount Preparation of Adult Drosophila Brains

There is an increasing interest in using Drosophila to model human brain degenerative diseases, map neuronal circuitries in adult brains, and study the molecular and cellular basis of higher brain functions. A whole-mount preparation of adult brains with well-preserved morphology is critical for such whole brain-based studies, but can be technically challenging and time-consuming. This protocol describes an easy-to-learn, one-step dissection approach of an adult fly head in less than 10 s, while keeping the intact brain attached to the rest of the body to facilitate subsequent processing steps. The procedure helps remove most of the eye and tracheal tissues normally associated with the brain that can interfere with the later imaging step, and also places less demand on the quality of the dissecting forceps. Additionally, we describe a simple method that allows convenient flipping of the mounted brain samples on a coverslip, which is important for imaging both sides of the brains with similar signal intensity and quality. As an example of the protocol, we present an analysis of dopaminergic (DA) neurons in adult brains of WT (w1118) flies. The high efficacy of the dissection method makes it particularly useful for large-scale adult brain-based studies in Drosophila.

A Simple One-step Dissection Protocol for Whole-mount Preparation of Adult Drosophila Brains

The adult Drosophila brain is a valuable system for studying neuronal circuitry, higher brain functions, and complex disorders. An efficient method to dissect whole brain tissue from the small fly head will facilitate brain-based studies. Here we describe a simple, one-step dissection protocol of adult brains with well-preserved morphology.

Um de uma etapa simples protocolo de dissecação para Whole mount-Preparação do Adulto<em&gt; Drosophila</em&gt; Brains

There is an increasing interest in using Drosophila to model human brain degenerative diseases, map neuronal circuitries in adult brains, and study the ...

Zebrafish In Situ Spinal Cord Preparation for Electrophysiological Recordings from Spinal Sensory and Motor Neurons

Zebrafish, first introduced as a developmental model, have gained popularity in many other fields. The ease of rearing large numbers of rapidly developing organisms, combined with the embryonic optical clarity, served as initial compelling attributes of this model. Over the past two decades, the success of this model has been further propelled by its amenability to large-scale mutagenesis screens and by the ease of transgenesis. More recently, gene-editing approaches have extended the power of the model.
For neurodevelopmental studies, the zebrafish embryo and larva provide a model to which multiple methods can be applied. Here, we focus on methods that allow the study of an essential property of neurons, electrical excitability. Our preparation for the electrophysiological study of zebrafish spinal neurons involves the use of veterinarian suture glue to secure the preparation to a recording chamber. Alternative methods for recording from zebrafish embryos and larvae involve the attachment of the preparation to the chamber using a fine tungsten pin1,2,3,4,5. A tungsten pin is most often used to mount the preparation in a lateral orientation, although it has been used to mount larvae dorsal-side up4. The suture glue has been used to mount embryos and larvae in both orientations. Using the glue, a minimal dissection can be performed, allowing access to spinal neurons without the use of an enzymatic treatment, thereby avoiding any resultant damage. However, for larvae, it is necessary to apply a brief enzyme treatment to remove the muscle tissue surrounding the spinal cord. The methods described here have been used to study the intrinsic electrical properties of motor neurons, interneurons, and sensory neurons at several developmental stages6,7,8,9.

Zebrafish In Situ Spinal Cord Preparation for Electrophysiological Recordings from Spinal Sensory and Motor Neurons

This manuscript describes methods for electrophysiological recordings from spinal neurons of zebrafish embryos and larvae. The preparation maintains neurons in situ and often involves minimum dissection. These methods allow for the electrophysiological study of a variety of spinal neurons, from the initial electrical excitability acquisition through the early larval stages.

zebrafish<em&gt; In Situ</em&gt; Spinal Cord Preparação para gravações eletrofisiológicas da Spinal Sensorial e Motor Neurons

Zebrafish, first introduced as a developmental model, have gained popularity in many other fields. The ease of rearing large numbers of rapidly developing ...

Visualization of the Axonal Projection Pattern of Embryonic Motor Neurons in Drosophila

The establishment of functional neuromuscular circuits relies on precise connections between developing motor axons and target muscles. Motor neurons extend growth cones to navigate along specific pathways by responding to a large number of axon guidance cues that emanate from the surrounding extracellular environment. Growth cone target recognition also plays a critical role in neuromuscular specificity. This work presents a standard immunohistochemistry protocol to visualize motor neuron projections of late stage-16 Drosophila melanogaster embryos. This protocol includes a few key steps, including a genotyping procedure, to sort the desired mutant embryos; an immunostaining procedure, to tag embryos with fasciclin II (FasII) antibody; and a dissection procedure, to generate filleted preparations from fixed embryos. Motor axon projections and muscle patterns in the periphery are much better visualized in flat preparations of filleted embryos than in whole-mount embryos. Therefore, the filleted preparation of fixed embryos stained with FasII antibody provides a powerful tool to characterize the genes required for motor axon pathfinding and target recognition, and it can also be applied to both loss-of-function and gain-of-function genetic screens.

Visualization of the Axonal Projection Pattern of Embryonic Motor Neurons in Drosophila

This work details a standard immunohistochemistry method to visualize motor neuron projections of late stage-16 Drosophila melanogaster embryos. The filleted preparation of fixed embryos stained with FasII antibody provides a powerful tool to characterize the genes required for motor axon pathfinding and target recognition during neural development.

Visualização do Padrão de Projeção Axonal de Neurônios de Motor Embrionários em<em&gt; Drosophila</em

The establishment of functional neuromuscular circuits relies on precise connections between developing motor axons and target muscles. Motor neurons ...

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.

Implante de janela de laminectomia e medula espinhal no 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 ...