Este trabalho foi possível graças ao apoio financeiro da Fondation pour la Recherche Médicale (FRM), o Milton bolsa de pós-doutorado Safenowitz para a pesquisa de ALS (ALS Association), Grants NIH NINDS NS05462 e NS034382, e Grant ANR HyperMND.

1. Um passo Medicação pré-anestésica: 10-15 min antes da indução de anestesia, injetar atropina (0,20 mg / kg) e methylprenidsolone (0,05 mg) subcutaneamente para evitar salivação e edema, respectivamente. 2. Passo dois Indução da anestesia: injectar pentobarbital sódico (70 mg / kg) ou de uma mistura de cetamina / xilazina (100 mg / kg e 10 mg / kg, respectivamente), intraperitonealmente. Deixe o rato ir até sob nenhum reflexo pitada do dedo do pé pode ser obtida. Se a anestesia parece muito leve, completar com 1/4 da dose. 3. Terceiro Passo * Nota: esta cirurgia é um procedimento terminal. Quando um plano cirúrgico de anestesia foi alcançada, transferir o rato sobre um cobertor aquecido levantada numa posição inclinada. <ol><li> Cobrir o focinho do rato com uma máscara de entrega O 2 puro com um caudal de cerca de 100 ml / min. </li><li&gt; Usando uma máquina de cortar cabelo, fazer a barba na região dorsal da nuca do pescoço até a base da cauda. </li><li> Barbear também o membro posterior direito. </li><li> Virar o mouse para a posição supina, mas tome cuidado para deixar a máscara de oxigênio no local. </li></ol> 4. Passo Quatro Fixar o rato no lugar com suturas looped em torno das pernas e fixado nos quatro cantos da superfície de trabalho. 5. Passo Cinco Insira uma sonda de temperatura para monitorar a temperatura do núcleo do rato. Ajuste cobertor de aquecimento / potência da lâmpada para manter a temperatura do núcleo entre 36 ° C e 38 ° C. 6. Traqueostomia e ventilação artificial <ol><li> Com uma tesoura sem corte, faça um corte sobre a traquéia e puxe a pele longe de ambos os lados. </li><li> Usando fórceps rombas, rasgar da glândula salivar de modo a expor os dois músculos finos (esterno), cobrindo a traqueia. </li><li> Utilizando uma pinça sem corte, separar o musc doisles ao longo da sua separação medial para revelar a traqueia. </li><li> Usando uma Dumont 7 fórceps, deslize dois comprimento de 4,0 fio de seda sob a traquéia. </li><li> Faça um corte transversal na traquéia entre dois anéis cartilaginosos, mas tome cuidado para não completamente secção da traquéia. </li><li> Insira o tubo traqueal para baixo da traquéia, em seguida, fixe-o em ambos os lados da abertura de inserção, amarrando as suturas sobre ele. O tubo traqueal é ligada a um ventilador de rato (SAR-830/AP, CWE Inc.) e um capnógrafo (μcapstar, CWE Inc.). O ventilador está ligado, através de um saco de cumprimento, com uma fonte de O 2 puro. Ajustar os parâmetros do ventilador (respiração taxa de 100-150 bpm, end-tidal volume de 170-310 ul) para que o mouse não está lutando contra a ventilação artificial e da pCO final de corrente 2 é estável entre 4 e 5%. </li></ol> 7. Colocação das linhas intravenosas <ol><li> Usando técnicas de raspagem, expor a jugularveia de um lado do pescoço. A este nível, a veia jugular divide-se em dois troncos principais, as veias faciais anterior e posterior. </li><li> Siga os seguintes passos duas vezes, um conjunto para cada um desses troncos: <ol><li> Usando Dumont 4 ou 5 fórceps, separe cuidadosamente a veia do seu tecido circundante conjuntivo. </li><li> Usando Dumont 7 fórceps, deslize dois comprimento de 6,0 fio de seda sob a veia. separá-los tanto quanto possível ao longo do comprimento da veia. </li><li> Colocar um clipe navio pequeno no lado proximal da veia (o lado mais próximo do coração), e amarrar o lado distal da veia. </li><li> Com uma tesoura muito finas íris, fazer uma pequena incisão na veia, tomando muito cuidado para não seção da veia completamente. </li><li> Inserir um cateter pré-cheia 1FR (premicath, Vygon) na abertura, até o grampo do vaso. </li><li> Segurando a veia eo cateter juntos em um 4 Dumont pinça, remova cuidadosamente o clipe de navio, em seguida, empurre o cateter um mor poucosmilímetros de e para a veia. </li><li> Fixar o cateter, amarrando-o com as duas suturas em ambos os lados da inserção. </li></ol></li><li> Um dos cateteres está ligada a uma seringa ou uma bomba de seringa para injectar doses suplementares de anestésico, quando necessário (normalmente a cada min 10-30). A dose IV é ou 6 mg / kg de pentobarbital de sódio ou 1250 40 ug / kg / min de cetamina / xilazina 12. O outro cateter está ligado a uma bomba seringa para infusão intravenosa lenta (50 uL / hr) de uma solução de glucose a 4% contendo NaHCO3 (1%) e plasmion (14%). </li></ol> 8. Feche a pele do pescoço com agulha e sutura, e retornar o mouse para a posição prona 9. Dissecção do músculo do membro-Hind e Nervos <ol><li> Com uma tesoura, fazer uma incisão da parte superior da coxa ao tendão de Aquiles. Separar a pele dos músculos subjacentes, tendo o cuidado para não danificar vasos sanguíneos. Cauterizar, conforme necessário. </li><li&gt; Identifique a borda anterior do músculo bíceps femoral, que aparece como uma linha branca correndo pela coxa. Com uma tesoura, abra cuidadosamente ao longo da linha do joelho todo o caminho até o osso do quadril. Separa-se os músculos. O nervo ciático é agora visível sob o bíceps femoral. </li><li> Dissecar cuidadosamente os bíceps femoral. Cauterizar / ligadura como necessário para evitar sangramento. O bíceps femoral pode ser totalmente removidos ou simplesmente reclinada para expor o nervo ciático e os tríceps sural músculos. </li><li> Dissecar o nervo sural. </li><li> Usando 8/0 fio de seda, amarre-se a parte distal do nervo fibular comum e cortar distal para o nó. Dissecar o nervo todo o caminho até, o mais próximo do hip possível. </li><li> Identificar o nervo tibial entre os nervos fibular comum e sural. Entre os diferentes ramos do nervo tibial, identificar os ramos que inervam o tríceps sural dos ramos que vão mais fundo (doravante referida como do nervo tibial). </li><li> Usando 8/0 sedafio, unir todos os ramos do nervo tibial, mantendo intactas as filiais da inervação do tríceps sural. Corte o nervo tibial distal ao nó e dissecar o nervo-se, tanto quanto possível. </li><li> Cobrir a totalidade dos membros posteriores com uma gaze embebida com uma solução salina para evitar a dessecação enquanto prosseguir com o passo seguinte. </li></ol> 10. Laminectomia <ol><li> Com uma tesoura, fazer uma incisão ao longo da espinha dorsal. Separou-se a pele contra os músculos subjacentes. </li><li> Fazer duas incisões em cada lado das vértebras para separar os músculos subcutâneos. Em seguida, retira cada tendão dos músculos que se fixam no lado das vértebras de cada lado. </li><li> Utilizando uma pinça sem corte e uma cureta bem, remover o restante do músculo no lado dorsal da vértebra ao redor das apófises espinhosas de identificar claramente cada vértebra individual. </li><li> Identificar vértebras T13 e L1. T13 é a última vértebra ter nervuras em anexo. A coluna vertebral wdoente ser imobilizado com as Cunningham grampos da Medula Espinal vertebrais (Stoelting Co.). Colocar os grampos em cada lado da coluna vertebral de T13 e L1. Tenha cuidado para não comprimir a medula espinhal, mas colocar um pouco de tensão no eixo longitudinal. Verifique se a coluna vertebral é bem protegido, pressionando delicadamente com uma pinça. </li><li> Usando fórceps finos, remover as apófises espinhosas, em seguida, as lâminas durante T13 e L1, expondo assim a medula espinhal. </li><li> Cubra a medula espinal exposta com pequenos pedaços de algodão ou spongel embebidas com soro fisiológico para evitar a dessecação. </li><li> Colocar num banho de personalizado feito de plástico no topo da parte de trás, que envolve a medula espinal. Garantir o banho no local usando 4/0 fio de seda. Certifique-se que o banho é à prova d&#39;água, selando-o com Kwik-Cast selante (WPI). </li><li> Uma vez que o Kwik-Cast ter secado, remova o algodão que cobre a medula espinhal, e encher o banho de óleo mineral. </li><li> Usando Dumont 5 pinças e tesouras muito finas íris, gentilmente puxar o companheiro durar envolve a medula espinal, e abri-lo tanto quanto possível, em ambos os sentidos. Dobrar a duração de cada lado da coluna vertebral. </li><li> Baixar a plataforma elevada na qual o rato é deitada de modo a mantê-lo em suspensão pelo grampo vertebral. </li><li> Use outro grampo vertebral para prender um processo espinhoso da região sacral para apoiar a região posterior do animal. </li><li> Uma braçadeira de terceiro é usado para imobilizar o membro posterior direito e tornozelo, flexionado em um ângulo de 90 ° para o joelho. </li></ol> 11. Aquiles Tendão Dissecção <ol><li> Com o membro dobrado em 90 ° no joelho e no tornozelo, remover a gaze cobrindo a área dos membros posteriores, e dissecar o tendão de Aquiles livre de tecido circundante. Dissecar tanto quanto possível sural Triceps a partir de tecido circundante também. </li><li> Cortar o tendão do músculo plantar perto de calcâneo, depois cortá-lo novamente para remover o tendão completamente. </li><li> Usando uma agulha enfiada, anexar fio 6/0 através de seda ªe tendão de Aquiles e faça um nó triplo ao redor do tendão. </li><li> Coloque o transdutor de força perto do nó, corte a parte distal do tendão, e anexá-lo ao transdutor de força usando o fio de seda. </li><li> Insira dois comprimentos de fio de aço inoxidável sob a fáscia do músculo tríceps sural. Estes fios são ligados a um amplificador de AC para o registo extracelular EMG. </li><li> Coloque o nervo fibular comum e nervo tibial em dois eletrodos bipolares de gancho. </li><li> Coloque o nervo sural Triceps sobre o cátodo de um eléctrodo de gancho, com o ânodo tocar um músculo nas proximidades. </li><li> Conectar todos os eléctrodos de estimulação para uma unidade de isolamento. </li><li> Coloque um eléctrodo de bola sobre a superfície dorsal da espinal-medula, ligado a um amplificador AC extracelular para gravar os potenciais cordão dorso. Coloque um eletrodo de referência Ag / AgCl em contato com um músculo de volta. </li></ol> 12. Doze Passos Estimular o nervo tríceps suralutilizando um 50 mseg pulso quadrado de intensidade crescente a uma frequência baixa (&lt;1 Hz), até que a amplitude máxima é observada contracção. Mova lentamente o transdutor de força para esticar o músculo enquanto monitorando a amplitude da resposta de contracção até que a amplitude twitch atinge um máximo. 13. Gravações intracelulares de motoneurônios A partir deste ponto, técnicas padrão de electrofisiológicos são usados ​​para preparar um eléctrodo intracelular, um neurónio penetrar na medula espinhal e identificá-lo como um motoneurónio. <ol><li> Puxe uma micropipeta de vidro a uma ponta mM ~ 1 usando um extrator pipeta (P-97 Micropipeta Puller, Instrumentos de Sutter). Encher o eléctrodo com uma solução 3 M KCl (resistência do eléctrodo de 10-20 MQ). </li><li> Usando um micropositioner, conduzir a micropipeta, ligado a um amplificador intracelular (Axoclamp 2B, Axon Instruments) para a medula espinhal. Monitorar os potenciais de campo locais induzidas pela estimulação de e-ach nervo para localizar o pool do motor do tríceps sural. </li><li> Cuidadosamente aproximar motoneurónios putativas ao monitorar a resistência do microeléctrodo. Ao empurrar contra uma membrana, a resistência aumenta. Penetração algures pode ser facilitada usando o &quot;buzz&quot; função do amplificador intracelular. </li></ol> 14. Procedimento de eutanásia No final da experiência, o animal é sacrificado com uma overdose de pentobarbital (210 mg / kg IV), seguido por decapitação.

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A preparação descrita aqui é o primeiro que permite, no rato adulto, a gravação simultânea intracelular de um motoneurônio lombar e a medição da força produzida pelas fibras musculares inervadas por seu axónio. Devido ao pequeno tamanho do animal, as técnicas cirúrgicas necessárias para esta preparação pode ser um desafio para adquirir. No entanto, uma vez que essas capacidades são dominados, a cirurgia inteira pode ser realizada em três horas, e os animais podem sobreviver...

A correction was made to <a href="http://www.jove.com/video/4312">Simultaneous Intracellular Recording of a Lumbar Motoneuron and the Force Produced by its Motor Unit in the Adult Mouse In vivo</a>. There was an error in the name of one author, Marin Manuel. The author's name has been corrected to:
Marin Manuel 
 
instead of: 
 
Manuel Marin

A correction to author list has been made for article <a href="http://www.jove.com/video/4312">Simultaneous Intracellular Recording of a Lumbar Motoneuron and the Force Produced by its Motor Unit in the Adult Mouse In vivo</a>.

Erratum: Simultaneous Intracellular Recording of a Lumbar Motoneuron and the Force Produced by its Motor Unit in the Adult Mouse In Vivo

<table border="1"><tbody><tr><td> Nome do reagente </td><td> Companhia </td><td> Número de catálogo </td><td> Comentários (opcional) </td></tr><tr><td> Sulfato de atropina </td><td> Aguettant </td><td></td><td></td></tr><tr><td> Methylprenidsolone </td><td> Pfizer </td><td> Solu-Medrol </td><td></td></tr><tr><td> Pentobarbitona de sódio </td><td> Sanofi-Aventis </td><td> Pentobarbital </td><td></td></tr><tr><td> Cetamina </td><td></td><td></td><td></td></tr><tr><td> Xilazina </td><td></td><td></td><td></td></tr><tr><td> Glicose </td><td></td><td></td><td></td></tr><tr><td> Plasma expansor </td><td> Roger Bellon </td><td> Plasmagel </td><td></td></tr><tr><td> Tesouras cegas </td><td> FST </td><td> 14079-10 </td><td></td></tr><tr><td> Blunt tesoura fina </td><td> FST </td><td> 15025-10 </td><td></td></tr><tr><td> Vannas Primavera Tesoura </td><td> FST </td><td> 15002-08 </td><td></td></tr><tr><td> Bem fórceps serrilhada </td><td> FST </td><td> 11370-32 </td><td></td></tr><tr><td> Bem fórceps serrilhada </td><td> FST </td><td> 11370-31 </td><td></td></tr><tr><td> Cunningham Adaptador Spinal </td><td> Stoelting Co. </td><td></td><td></td></tr><tr><td> Kwik-Cast selante </td><td> WPI </td><td> # KWIK-CAST </td><td></td></tr><tr><td> Ventilador </td><td> CWE Inc </td><td> SAR-830/AP </td><td></td></tr><tr><td> Capnógrafo </td><td> CWE Inc </td><td> μcapstar </td><td></td></tr><tr><td> Cobertor </td><td> Harvard Apparatus </td><td> 507221F </td><td></td></tr><tr><td> Amplificador intracelular </td><td> Axon Instruments </td><td> Axoclamp 2B </td><td></td></tr><tr><td> Pipeta extrator </td><td> Sutter Instruments </td><td> P-97 </td><td></td></tr><tr><td> KCl </td><td> Sigma-Aldrich </td><td> P9333-500G </td><td></td></tr></tbody></table>

simultaneous intracellular recording of a lumbar motoneuron and the force produced by its motor unit in the adult mouse in vivo

O motoneurônio espinhal tem sido um sistema modelo bom para estudar a função neural, porque é um neurónio do sistema nervoso central, com as propriedades únicas de (1) tendo alvos prontamente identificáveis ​​(fibras musculares) e, portanto, tem uma função bem conhecido (para controlar a contracção muscular), (2) ser o alvo convergente de muitas redes e descendente da coluna vertebral, daí o nome de &quot;via comum final&quot;, e (3), com uma soma grande que faz com que seja possível a penetração los com afiadas eletrodos intracelulares . Além disso, quando estudados in vivo, é possível gravar, simultaneamente, a actividade eléctrica dos motoneurónios e da força desenvolvida pelo músculo seus alvos. Realizando gravações intracelulares de motoneurónios in vivo, por conseguinte, colocar o experimentalista, na posição única de ser capaz de estudar, ao mesmo tempo, todos os compartimentos da &quot;unidade motora&quot; (o nome dado à motoneurónio, do seu axónio, eas fibras musculares que inerva 1): Os insumos que incidem sobre o motoneurônio, as propriedades eletrofisiológicas do motoneurônio, eo impacto destas propriedades em função fisiológica dos neurônios motores, ou seja, a força produzida por seu motor. No entanto, esta abordagem é muito difícil, porque a preparação não pode ser paralisado e, assim, a estabilidade mecânica para a gravação intracelular é reduzida. Assim, este tipo de experiências só foi alcançado em gatos e em ratos. No entanto, o estudo dos sistemas motores espinais poderia fazer um salto formidável se que era possível realizar experiências semelhantes em ratinhos normais e geneticamente modificados. Por razões técnicas, o estudo das redes espinhal em ratos, principalmente se limitado a neonatal em preparações in vitro, onde os neurônios motores e as redes espinhal são imaturos, os neurônios motores são separados de suas metas, e quando estudado em fatias, o MOtoneurons são separadas a maior parte das suas entradas. Até recentemente, apenas alguns grupos conseguiram realizar gravações intracelulares de motoneurônios in vivo 2-4, incluindo a nossa equipe, que publicou uma nova preparação que nos permitiu obter gravações muito estáveis ​​de motoneurônios in vivo em camundongos adultos 5,6. No entanto, estas gravações foram obtidos em animais paralisados, isto é, sem a possibilidade de gravar a saída força destes neurónios motores. Aqui apresentamos uma extensão da preparação original em que fomos capazes de obter gravações simultâneas das propriedades eletrofisiológicas das motoneurônios e da força desenvolvida por seu motor. Esta é uma conquista importante, pois nos permite identificar os diferentes tipos de neurônios motores com base no seu perfil de força, e revelando assim a sua função. Juntamente com modelos genéticos perturbar circuito segmentar vertebral 7-9, ou reproduzindo doen humanaSE 10,11, esperamos que esta técnica para ser uma ferramenta essencial para o estudo do sistema motor espinhal.

A Figura 1 mostra como identificar um motoneurônio do grupo tríceps sural após a penetração. Em intensidade de estimulação baixo, apenas uma EPSP monossináptico pode ser observado (Figura 1A). Com maior intensidade, o EPSP pode ser suficientemente grande para provocar uma &quot;ortodrómica&quot; pico (Figura 1B). Em intensidade de estimulação ainda maior, um tudo-ou-nada antidrômica espiga aparecer, com uma latência mais curta do que o EPSP monossináptico ...

Watch this Scientific Journal Video about Simultaneous Intracellular Recording of a Lumbar Motoneuron and the Force Produced by its Motor Unit in the Adult Mouse In vivo at JoVE.com

Simultaneous Intracellular Recording of a Lumbar Motoneuron and the Force Produced by its Motor Unit in the Adult Mouse In vivo

Este método novo permite a gravação simultânea intracelular de um motoneurônio único adulto rato e a medição da força produzida por suas fibras musculares. A investigação combinado das propriedades eléctricas e mecânicas das unidades motoras em animais normais e geneticamente modificadas é um avanço para o estudo do sistema neuromuscular.

Gravação Intracelular simultânea de um motoneurônio lombar e a força produzida pelo seu motor no Rato Adulto<em&gt; Em vivo</em

The  spinal motoneuron has long been a good model system for studying neural  function because it is a neuron of the central nervous system with the ...

simultaneous-intracellular-recording-lumbar-motoneuron-force-produced

Research

JoVE Journal

Neuroscience

Simultaneous Intracellular Recording of a Lumbar Motoneuron and the Force Produced by its Motor Unit in the Adult Mouse In vivo

14.6K visualizações.  Northwestern University Feinberg School of Medicine. Este método novo permite a gravação simultânea intracelular de um motoneurônio único adulto rato e a medição da força produzida por suas fibras musculares. A investigação combinado das propriedades eléctricas e mecânicas das unidades motoras em animais normais e geneticamente modificadas é um avanço para o estudo do sistema neuromuscular.

Vídeo: Simultaneous Intracellular Recording of a Lumbar Motoneuron and the Force Produced by its Motor Unit in the Adult Mouse In vivo

Transfection of Mouse Retinal Ganglion Cells by in vivo Electroporation

The targeting and refinement of RGC projections to the midbrain is a popular and powerful model system for studying how precise patterns of neural connectivity form during development. In mice, retinofugal projections are arranged in a topographic manner and form eye-specific layers in the Lateral Geniculate Nucleus (dLGN) of the thalamus and the Superior Colliculus (SC). The development of these precise patterns of retinofugal projections has typically been studied by labeling populations of RGCs with fluorescent dyes and tracers, such as horseradish peroxidase1-4. However, these methods are too coarse to provide insight into developmental changes in individual RGC axonal arbor morphology that are the basis of retinotopic map formation. They also do not allow for the genetic manipulation of RGCs.
Recently, electroporation has become an effective method for providing precise spatial and temporal control for delivery of charged molecules into the retina5-11. Current retinal electroporation protocols do not allow for genetic manipulation and tracing of retinofugal projections of a single or small cluster of RGCs in postnatal mice. It has been argued that postnatal in vivo electroporation is not a viable method for transfecting RGCs since the labeling efficiency is extremely low and hence requires targeting at embryonic ages when RGC progenitors are undergoing differentiation and proliferation6. 
In this video we describe an in vivo electroporation protocol for targeted delivery of genes, shRNA, and fluorescent dextrans to murine RGCs postnatally. This technique provides a cost effective, fast and relatively easy platform for efficient screening of candidate genes involved in several aspects of neural development including axon retraction, branching, lamination, regeneration and synapse formation at various stages of circuit development. In summary we describe here a valuable tool which will provide further insights into the molecular mechanisms underlying sensory map development.

Transfection of Mouse Retinal Ganglion Cells by in vivo Electroporation

We demonstrate an in vivo electroporation protocol for transfecting single or small clusters of retinal ganglion cells (RGCs) and other retinal cell types in postnatal mice over a wide range of ages. The ability to label and genetically manipulate postnatal RGCs in vivo is a powerful tool for developmental studies.

Transfecção de células de camundongo por Ganglionares da Retina In vivo Eletroporação

The targeting and refinement of RGC projections to the midbrain is a popular and powerful model system for studying how precise patterns of neural ...

Neurociência

Retrograde Fluorescent Labeling Allows for Targeted Extracellular Single-unit Recording from Identified Neurons In vivo

The overall goal of this method is to record single-unit responses from an identified population of neurons. In vivo electrophysiological recordings from individual neurons are critical for understanding how neural circuits function under natural conditions. Traditionally, these recordings have been performed 'blind', meaning the identity of the recorded cell is unknown at the start of the recording. Cellular identity can be subsequently determined via intracellular1, juxtacellular2 or loose-patch3 iontophoresis of dye, but these recordings cannot be pre-targeted to specific neurons in regions with functionally heterogeneous cell types. Fluorescent proteins can be expressed in a cell-type specific manner permitting visually-guided single-cell electrophysiology4-6. However, there are many model systems for which these genetic tools are not available. Even in genetically accessible model systems, the desired promoter may be unknown or genetically homogenous neurons may have varying projection patterns. Similarly, viral vectors have been used to label specific subgroups of projection neurons7, but use of this method is limited by toxicity and lack of trans-synaptic specificity. Thus, additional techniques that offer specific pre-visualization to record from identified single neurons in vivo are needed. Pre-visualization of the target neuron is particularly useful for challenging recording conditions, for which classical single-cell recordings are often prohibitively difficult8-11. The novel technique described in this paper uses retrograde transport of a fluorescent dye applied using tungsten needles to rapidly and selectively label a specific subset of cells within a particular brain region based on their unique axonal projections, thereby providing a visual cue to obtain targeted electrophysiological recordings from identified neurons in an intact circuit within a vertebrate CNS.
The most significant novel advancement of our method is the use of fluorescent labeling to target specific cell types in a non-genetically accessible model system. Weakly electric fish are an excellent model system for studying neural circuits in awake, behaving animals12. We utilized this technique to study sensory processing by "small cells" in the anterior exterolateral nucleus (ELa) of weakly electric mormyrid fish. "Small cells" are hypothesized to be time comparator neurons important for detecting submillisecond differences in the arrival times of presynaptic spikes13. However, anatomical features such as dense myelin, engulfing synapses, and small cell bodies have made it extremely difficult to record from these cells using traditional methods11, 14. Here we demonstrate that our novel method selectively labels these cells in 28% of preparations, allowing for reliable, robust recordings and characterization of responses to electrosensory stimulation.

Retrograde Fluorescent Labeling Allows for Targeted Extracellular Single-unit Recording from Identified Neurons In vivo

Retrograde transport of fluorescent dye labels a sub-population of neurons based on anatomical projection. Labeled axons can be visually targeted in vivo, permitting extracellular recording from identified axons. This technique facilitates recording when neurons cannot be labeled through genetic manipulation or are difficult to isolate using 'blind' in vivo approaches.

Labeling fluorescentes retrógrada Permite alvejado Extracelular Gravação Single-unidade dos neurônios identificados<em&gt; In vivo</em

The overall goal of this method is to record single-unit responses from an identified population of neurons. In vivo electrophysiological recordings from ...

Time-lapse Imaging of Neuroblast Migration in Acute Slices of the Adult Mouse Forebrain

There is a substantial body of evidence indicating that new functional neurons are constitutively generated from an endogenous pool of neural stem cells in restricted areas of the adult mammalian brain. Newborn neuroblasts from the subventricular zone (SVZ) migrate along the rostral migratory stream (RMS) to their final destination in the olfactory bulb (OB)1. In the RMS, neuroblasts migrate tangentially in chains ensheathed by astrocytic processes2,3 using blood vessels as a structural support and a source of molecular factors required for migration4,5. In the OB, neuroblasts detach from the chains and migrate radially into the different bulbar layers where they differentiate into interneurons and integrate into the existing network1, 6.
In this manuscript we describe the procedure for monitoring cell migration in acute slices of the rodent brain. The use of acute slices allows the assessment of cell migration in the microenvironment that closely resembling to in vivo conditions and in brain regions that are difficult to access for in vivo imaging. In addition, it avoids long culturing condition as in the case of organotypic and cell cultures that may eventually alter the migration properties of the cells. Neuronal precursors in acute slices can be visualized using DIC optics or fluorescent proteins. Viral labeling of neuronal precursors in the SVZ, grafting neuroblasts from reporter mice into the SVZ of wild-type mice, and using transgenic mice that express fluorescent protein in neuroblasts are all suitable methods for visualizing neuroblasts and following their migration. The later method, however, does not allow individual cells to be tracked for long periods of time because of the high density of labeled cells. We used a wide-field fluorescent upright microscope equipped with a CCD camera to achieve a relatively rapid acquisition interval (one image every 15 or 30 sec) to reliably identify the stationary and migratory phases. A precise identification of the duration of the stationary and migratory phases is crucial for the unambiguous interpretation of results. We also performed multiple z-step acquisitions to monitor neuroblasts migration in 3D. Wide-field fluorescent imaging has been used extensively to visualize neuronal migration7-10. Here, we describe detailed protocol for labeling neuroblasts, performing real-time video-imaging of neuroblast migration in acute slices of the adult mouse forebrain, and analyzing cell migration. While the described protocol exemplified the migration of neuroblasts in the adult RMS, it can also be used to follow cell migration in embryonic and early postnatal brains.

We describe a protocol for real-time videoimaging of neuronal migration in the mouse forebrain. The migration of virally-labeled or grafted neuronal precursors was recorded in acute live slices using wide-field fluorescent imaging with a relatively rapid acquisition interval to study the different phases of cell migration, including the durations of the stationary and migration phases and the speed of migration.

Lapso de tempo de imagem de migração neuroblastos em fatias agudas do cérebro anterior Rato Adulto

There is a substantial body of evidence indicating that new functional neurons are constitutively generated from an endogenous pool of neural stem cells ...

Multi-unit Recording Methods to Characterize Neural Activity in the Locust (Schistocerca Americana) Olfactory Circuits

Detection and interpretation of olfactory cues are critical for the survival of many organisms. Remarkably, species across phyla have strikingly similar olfactory systems suggesting that the biological approach to chemical sensing has been optimized over evolutionary time1. In the insect olfactory system, odorants are transduced by olfactory receptor neurons (ORN) in the antenna, which convert chemical stimuli into trains of action potentials. Sensory input from the ORNs is then relayed to the antennal lobe (AL; a structure analogous to the vertebrate olfactory bulb). In the AL, neural representations for odors take the form of spatiotemporal firing patterns distributed across ensembles of principal neurons (PNs; also referred to as projection neurons)2,3. The AL output is subsequently processed by Kenyon cells (KCs) in the downstream mushroom body (MB), a structure associated with olfactory memory and learning4,5. Here, we present electrophysiological recording techniques to monitor odor-evoked neural responses in these olfactory circuits.
First, we present a single sensillum recording method to study odor-evoked responses at the level of populations of ORNs6,7. We discuss the use of saline filled sharpened glass pipettes as electrodes to extracellularly monitor ORN responses. Next, we present a method to extracellularly monitor PN responses using a commercial 16-channel electrode3. A similar approach using a custom-made 8-channel twisted wire tetrode is demonstrated for Kenyon cell recordings8. We provide details of our experimental setup and present representative recording traces for each of these techniques.

Multi-unit Recording Methods to Characterize Neural Activity in the Locust Schistocerca Americana Olfactory Circuits

We demonstrate variations of the extracellular multi-unit recording technique to characterize odor-evoked responses in the first three stages of the invertebrate olfactory pathway. These techniques can easily be adapted to examine ensemble activity in other neural systems as well.

Multi-unidade Métodos de gravação para caracterizar a atividade neural no Locust (<em&gt; Schistocerca Americana</em&gt;) Circuitos olfatórias

Detection and interpretation of olfactory cues are critical for the survival  of many organisms. Remarkably, species across phyla have strikingly similar ...

Simultaneous ex vivo Functional Testing of Two Retinas by in vivo Electroretinogram System

An In vivo electroretinogram (ERG) signal is composed of several overlapping components originating from different retinal cell types, as well as noise from extra-retinal sources. Ex vivo ERG provides an efficient method to dissect the function of retinal cells directly from an intact isolated retina of animals or donor eyes. In addition, ex vivo ERG can be used to test the efficacy and safety of potential therapeutic agents on retina tissue from animals or humans. We show here how commercially available in vivo ERG systems can be used to conduct ex vivo ERG recordings from isolated mouse retinas. We combine the light stimulation, electronic and heating units of a standard in vivo system with custom-designed specimen holder, gravity-controlled perfusion system and electromagnetic noise shielding to record low-noise ex vivo ERG signals simultaneously from two retinas with the acquisition software included in commercial in vivo systems. Further, we demonstrate how to use this method in combination with pharmacological treatments that remove specific ERG components in order to dissect the function of certain retinal cell types.

Simultaneous ex vivo Functional Testing of Two Retinas by in vivo Electroretinogram System

Ex vivo ERG can be used to record electrical activity of retinal cells directly from isolated intact retinas of animals or humans. We demonstrate here how common in vivo ERG systems can be adapted for ex vivo ERG recordings in order to dissect the electrical activity of retinal cells.

Simultâneo<em&gt; Ex vivo</em&gt; Teste Funcional de Dois retinas por<em&gt; In vivo</em&gt; Sistema Eletrorretinograma

An In vivo electroretinogram (ERG) signal is composed of several overlapping components originating from different retinal cell types, as well as noise ...

Electrophysiological Method for Recording Intracellular Voltage Responses of Drosophila Photoreceptors and Interneurons to Light Stimuli In Vivo

Voltage responses of insect photoreceptors and visual interneurons can be accurately recorded with conventional sharp microelectrodes. The method described here enables the investigator to measure long-lasting (from minutes to hours) high-quality intracellular responses from single Drosophila R1-R6 photoreceptors and Large Monopolar Cells (LMCs) to light stimuli. Because the recording system has low noise, it can be used to study variability among individual cells in the fly eye, and how their outputs reflect the physical properties of the visual environment. We outline all key steps in performing this technique. The basic steps in constructing an appropriate electrophysiology set-up for recording, such as design and selection of the experimental equipment are described. We also explain how to prepare for recording by making appropriate (sharp) recording and (blunt) reference electrodes. Details are given on how to fix an intact fly in a bespoke fly-holder, prepare a small window in its eye and insert a recording electrode through this hole with minimal damage. We explain how to localize the center of a cell's receptive field, dark- or light-adapt the studied cell, and to record its voltage responses to dynamic light stimuli. Finally, we describe the criteria for stable normal recordings, show characteristic high-quality voltage responses of individual cells to different light stimuli, and briefly define how to quantify their signaling performance. Many aspects of the method are technically challenging and require practice and patience to master. But once learned and optimized for the investigator's experimental objectives, it grants outstanding in vivo neurophysiological data.

Electrophysiological Method for Recording Intracellular Voltage Responses of Drosophila Photoreceptors and Interneurons to Light Stimuli In Vivo

Sharp microelectrodes enable accurate electrophysiological characterization of photoreceptor and visual interneuron output in living Drosophila. Here we show how to use this method to record high-quality voltage responses of individual cells to controlled light stimulation. This method is ideal for studying neural information processing in insect compound eyes.

Método eletrofisiológico para respostas gravação intracelular de tensão<em&gt; Drosophila</em&gt; Fotorreceptores e Interneurônios à luz Estímulos<em&gt; In Vivo</em

Voltage responses of insect photoreceptors and visual interneurons can be accurately recorded with conventional sharp microelectrodes. The method ...

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 ...

In Vivo Single-Molecule Tracking at the Drosophila Presynaptic Motor Nerve Terminal

An increasing number of super-resolution microscopy techniques are helping to uncover the mechanisms that govern the nanoscale cellular world. Single-molecule imaging is gaining momentum as it provides exceptional access to the visualization of individual molecules in living cells. Here, we describe a technique that we developed to perform single-particle tracking photo-activated localization microscopy (sptPALM) in Drosophila larvae. Synaptic communication relies on key presynaptic proteins that act by docking, priming, and promoting the fusion of neurotransmitter-containing vesicles with the plasma membrane. A range of protein-protein and protein-lipid interactions tightly regulates these processes and the presynaptic proteins therefore exhibit changes in mobility associated with each of these key events. Investigating how mobility of these proteins correlates with their physiological function in an intact live animal is essential to understanding their precise mechanism of action. Extracting protein mobility with high resolution in vivo requires overcoming limitations such as optical transparency, accessibility, and penetration depth. We describe how photoconvertible fluorescent proteins tagged to the presynaptic protein Syntaxin-1A can be visualized via slight oblique illumination and tracked at the motor nerve terminal or along the motor neuron axon of the third instar Drosophila larva.

In Vivo Single-Molecule Tracking at the Drosophila Presynaptic Motor Nerve Terminal

Here we illustrate how single molecule photo-activated localization microscopy can be carried out on the motor nerve terminal of a live Drosophila melanogaster third instar larva.

Na Vivo Único-molécula de rastreamento no Terminal do nervo Motor pré-sináptica da drosófila

An increasing number of super-resolution microscopy techniques are helping to uncover the mechanisms that govern the nanoscale cellular world. ...

Ablation of a Neuronal Population Using a Two-photon Laser and Its Assessment Using Calcium Imaging and Behavioral Recording in Zebrafish Larvae

To identify the role of a subpopulation of neurons in behavior, it is essential to test the consequences of blocking its activity in living animals. Laser ablation of neurons is an effective method for this purpose when neurons are selectively labeled with fluorescent probes. In the present study, protocols for laser ablating a subpopulation of neurons using a two-photon microscope and testing of its functional and behavioral consequences are described. In this study, prey capture behavior in zebrafish larvae is used as a study model. The pretecto-hypothalamic circuit is known to underlie this visually-driven prey catching behavior. Zebrafish pretectum were laser-ablated, and neuronal activity in the inferior lobe of the hypothalamus (ILH; the target of the pretectal projection) was examined. Prey capture behavior after pretectal ablation was also tested.

Here, we present a protocol to ablate a genetically labeled subpopulation of neurons by a two-photon laser from Zebrafish larvae.

Ablação de uma população Neuronal, usando um Laser de dois fotões e sua avaliação usando imagens de cálcio e gravação comportamental em larvas de Zebrafish

To identify the role of a subpopulation of neurons in behavior, it is essential to test the consequences of blocking its activity in living animals. Laser ...

Investigating Mammalian Axon Regeneration: In Vivo Electroporation of Adult Mouse Dorsal Root Ganglion

Electroporation is an essential non-viral gene transfection approach to introduce DNA plasmids or small RNA molecules into cells. A sensory neuron in the dorsal root ganglion (DRGs) extends a single axon with two branches. One branch goes to the peripheral nerve (peripheral branch), and the other branch enters the spinal cord through the dorsal root (central branch). After the neural injury, the peripheral branch regenerates robustly whereas the central branch does not regenerate. Due to the high regenerative capacity, sensory axon regeneration has been widely used as a model system to study mammalian axon regeneration in both the peripheral nervous system (PNS) and the central nervous system (CNS). Here, we describe a previously established approach protocol to manipulate gene expression in mature sensory neurons in vivo via electroporation. Based on transfection with plasmids or small RNA oligos (siRNAs or microRNAs), the approach allows for both loss- and gain-of-function experiments to study the roles of genes-of-interests or microRNAs in regulation of axon regeneration in vivo. In addition, the manipulation of gene expression in vivo can be controlled both spatially and temporally within a relatively short time course. This model system provides a unique tool to investigate the molecular mechanisms by which mammalian axon regeneration is regulated in vivo.

Investigating Mammalian Axon Regeneration: In Vivo Electroporation of Adult Mouse Dorsal Root Ganglion

Electroporation is an effective approach to deliver genes of interests into cells. By applying this approach in vivo on the neurons of adult mouse dorsal root ganglion (DRG), we describe a model to study axon regeneration in vivo.

Investigação regeneração do axônio mamíferos: In Vivo Electroporation do gânglio de raiz Dorsal do rato adulto

Electroporation is an essential non-viral gene transfection approach to introduce DNA plasmids or small RNA molecules into cells. A sensory neuron in the ...