Este trabajo ha sido posible gracias al apoyo financiero de la Fondation pour la Recherche Médicale (FRM), la Beca Postdoctoral Milton Safenowitz Investigación de la ELA (ALS Association), NIH NINDS subvenciones NS05462 y NS034382 y ANR subvención HyperMND.

1. Paso uno Medicación preanestésica: 10-15 min antes de la inducción de la anestesia, inyectar atropina (0,20 mg / kg) y methylprenidsolone (0,05 mg) por vía subcutánea para prevenir la salivación y edema, respectivamente. 2. Paso dos La inducción de la anestesia: inyectar pentobarbital sódico (70 mg / kg) o una mezcla de ketamina / xilazina (100 mg / kg y 10 mg / kg, respectivamente) por vía intraperitoneal. Vamos a pasar por debajo de la del ratón hasta que no reflejo del pellizco dedo del pie se puede conseguir. Si la anestesia parece demasiado clara, completar con 1/4 de la dosis. 3. Paso tres * Nota: esta cirugía es un procedimiento terminal. Cuando un plano quirúrgico de anestesia ha sido alcanzado, transferir el ratón sobre una manta caliente levantado en posición prona. <ol><li> Cubra el hocico del ratón con una máscara de la entrega de O 2 puro con un caudal de alrededor de 100 ml / min. </li><li&gt; Uso de una cortadora de cabello, afeitarse la región dorsal de la nuca del cuello hasta la base de la cola. </li><li> Afeitado también la extremidad posterior derecha. </li><li> Mueva el ratón a la posición supina, pero tenga cuidado de dejar la mascarilla de oxígeno en su lugar. </li></ol> 4. Paso cuatro Asegure el ratón en el lugar con suturas enrolladas alrededor de los miembros y asegurados en las cuatro esquinas de la superficie de trabajo. 5. Paso cinco Insertar una sonda de temperatura para controlar la temperatura del núcleo del ratón. Ajuste manta de calefacción / potencia de la lámpara para mantener la temperatura de núcleo entre 36 ° C y 38 ° C. 6. La traqueotomía y ventilación artificial <ol><li> Con unas tijeras romas, haga un corte a través de la tráquea y tire de la piel de distancia en ambos lados. </li><li> Utilizando pinzas romas, desgarrar la glándula salival para exponer dos músculos delgados (esternohioideo) que cubren la tráquea. </li><li> Con unas pinzas romas, separe el musc dosles a lo largo de su separación medial para revelar la tráquea. </li><li> El uso de un Dumont 7 pinzas, deslizar dos longitudes de sutura de seda 4.0 en la tráquea. </li><li> Hacer un corte transversal en la tráquea entre dos anillos cartilaginosos pero tener cuidado de no completamente sección de la tráquea. </li><li> Insertar el tubo traqueal por la tráquea, a continuación, asegurar en ambos lados de la abertura de inserción atando las suturas del mismo. El tubo traqueal está conectado a un ventilador ratón (SAR-830/AP, CWE Inc.) y un capnógrafo (μcapstar, CWE Inc.). El ventilador está conectado, a través de una bolsa de cumplimiento, a una fuente de puro O 2. Ajuste de los parámetros del ventilador (frecuencia respiratoria 100-150 lpm, final de la espiración volumen 170-310 l), de modo que el ratón no está luchando contra la ventilación artificial y la pCO final de la espiración 2 es estable entre un 4 y 5%. </li></ol> 7. La colocación de los catéteres intravenosos <ol><li> Utilizando técnicas de disección roma, exponer la yugularvena en un lado del cuello. En este nivel, la vena yugular se divide en dos troncos principales, las venas faciales anterior y posterior. </li><li> Realice los siguientes pasos dos veces, uno para cada uno de estos troncos: <ol><li> Usando Dumont 4 o 5 fórceps, separe con cuidado la vena de su tejido circundante conjuntivo. </li><li> Usando Dumont 7 pinzas, deslizar dos longitud de 6,0 puntos de seda debajo de la vena. separar la medida de lo posible a lo largo de la longitud de la vena. </li><li> Colocar una grapa de vaso pequeño en el lado proximal de la vena (el lado más cercano al corazón), y atar el lado distal de la vena. </li><li> Con unas tijeras iris muy finas, hacer una pequeña incisión transversal en la vena, no teniendo mucho cuidado a la sección de la vena por completo. </li><li> Insertar un catéter 1FR precargada (premicath, Vygon) en la abertura, hasta la grapa de vaso. </li><li> Sosteniendo la vena y el catéter juntos en una Dumont 4 pinzas, retire con cuidado el clip buque, a continuación, empuje el catéter unos pocos mormilímetros correos en la vena. </li><li> Fijar el catéter mediante la vinculación con las dos suturas en ambos lados de la inserción. </li></ol></li><li> Uno de los catéteres está conectado a una jeringa o una bomba de jeringa para inyectar dosis complementarias de anestésico siempre que sea necesario (generalmente cada min 10-30). La dosis IV es o bien 6 mg / kg de pentobarbital sódico o 1250 40 g / kg / min de ketamina / xilazina 12. El otro catéter se conecta a una bomba de jeringa para una infusión intravenosa lenta (50 l / h) de una solución de glucosa al 4% que contenía NaHCO 3 (1%) y plasmion (14%). </li></ol> 8. Cierre la piel del cuello con la aguja y la sutura, y devolver el ratón a la posición prona 9. La disección de los músculos de las extremidades posteriores y nervios <ol><li> Con unas tijeras, hacer una incisión desde la parte superior del muslo en el tendón de Aquiles. Separar la piel de los músculos subyacentes, teniendo cuidado de no dañar los vasos sanguíneos. Cauterizar según sea necesario. </li><li&gt; Identificar el borde anterior del bíceps femoral, que aparece como una línea blanca corriendo por el muslo. Usando tijeras, abra cuidadosamente a lo largo de la línea de la rodilla hasta el final a hueso de la cadera. Separe los músculos. El nervio ciático es ahora visible en el bíceps femoral. </li><li> Diseccionar cuidadosamente el bíceps femoral. Cauterize / ligadura según sea necesario para prevenir el sangrado. El bíceps femoral puede ser totalmente eliminado o simplemente recostado para exponer el nervio ciático y el tríceps sural músculos. </li><li> Diseccionar el nervio sural. </li><li> Con 8/0 hilos de seda, atar la parte distal del nervio peroneo común y corte distal al nudo. Disección del nervio hasta el final para arriba, tan cerca de la cadera como sea posible. </li><li> Identifique el nervio tibial entre los nervios peroneo común y Sural. Entre las diferentes ramas del nervio tibial, identificar las ramas que inervan el tríceps sural de las ramas que van más profunda (en adelante el nervio tibial). </li><li> Con 8/0 sedahilo, unir todas las ramas del nervio tibial, manteniendo intactas las ramas que inervan el tríceps sural. Cortar el nervio tibial distal a la del nudo y diseccionar el nervio hasta la medida de lo posible. </li><li> Cubrir toda la extremidad posterior con una gasa embebida con solución salina para prevenir la desecación mientras que proceder con el paso siguiente. </li></ol> 10. Laminectomía <ol><li> Con unas tijeras, hacer una incisión a lo largo de la columna vertebral. Se separa la piel de los músculos subyacentes. </li><li> Hacer dos incisiones a cada lado de las vértebras a separar los músculos subcutáneos. A continuación, cortar cada tendón de los músculos que se conectan en el lado de las vértebras a cada lado. </li><li> Utilizando pinzas romas y una cureta fina, retire el resto del músculo en el lado dorsal de las vértebras alrededor de las apófisis espinosas para identificar claramente cada vértebra individual. </li><li> Identificar vértebras T13 y L1. T13 es la última vértebra tener costillas adjunto. La columna vertebral wenfermo puede inmovilizar con las abrazaderas Cunningham Médula Espinal vertebrales (Stoelting Co.). Coloque las abrazaderas de la columna vertebral en cada lado de T13 y L1. Tenga cuidado de no comprimir la médula espinal, pero poner un poco de tensión en el eje longitudinal. Compruebe que la columna vertebral se asegura bien presionando hacia abajo suavemente con fórceps. </li><li> Usando pinzas gubias finas, eliminar los procesos espinales entonces las láminas sobre T13 y L1, exponiendo así la médula espinal. </li><li> Cubra la médula espinal expuesta con pequeños trozos de algodón o spongel absorbido con solución salina para evitar la desecación. </li><li> Colocar una costumbre baño hecho de plástico en la parte superior de la espalda, que rodea la médula espinal. Fije la bañera en su lugar con 4/0 hilos de seda. Asegúrese de que el baño es a prueba de agua, sellando con Kwik-Cast sellador (WPI). </li><li> Una vez que el Kwik-Cast se ha secado, retire el algodón que cubre la médula espinal, y llenar la bañera con aceite mineral. </li><li> Usando Dumont 5 pinzas y tijeras iris muy finos, tire suavemente de la pareja durar rodea la médula espinal, y lo abre en la medida de lo posible en ambas direcciones. Doble la duramadre en cada lado de la médula espinal. </li><li> Bajar la plataforma elevada en la que el ratón está mintiendo a fin de mantenerlo suspendido por la abrazadera vertebral. </li><li> Use otra abrazadera vertebral para sujetar una apófisis espinosa de la región sacra para apoyar la región de la espalda del animal. </li><li> Una abrazadera tercero se utiliza para inmovilizar la extremidad posterior derecha y el tobillo, doblada en un ángulo de 90 grados a la rodilla. </li></ol> 11. Aquiles Tendón Disección <ol><li> Con la extremidad doblada a 90 ° en la rodilla y en el tobillo, quitar la gasa que cubre el área de miembros posteriores, y diseccionar el tendón de Aquiles libre de los tejidos circundantes. Diseccionar tanto como sea posible el tríceps sural del tejido circundante también. </li><li> Cortar el tendón del músculo plantar de cierre en el calcáneo, entonces cortar de nuevo para eliminar completamente el tendón. </li><li> Usando una aguja roscada, coloque 6/0 seda hilo a través de the tendón de Aquiles y hacer un nudo triple alrededor del tendón. </li><li> Coloque el transductor de fuerza cerca del nudo, corte la parte distal del tendón, y adjuntarlo al transductor de fuerza con el hilo de seda. </li><li> Coloque dos trozos de alambre de acero inoxidable debajo de la fascia de los músculos tríceps sural. Estos cables están conectados a un amplificador extracelular de CA para la grabación de EMG. </li><li> Coloque el nervio peroneo común y el nervio tibial en dos electrodos bipolares gancho. </li><li> Coloque el nervio tríceps sural en el cátodo de un electrodo de gancho, con el ánodo de tocar un músculo cercano. </li><li> Conecte todos los electrodos de estimulación a una unidad de aislamiento. </li><li> Colocar un electrodo de bola en la superficie dorsal de la médula espinal, conectado a un amplificador extracelular de CA a registrar los potenciales de médula dorso. Colocar un electrodo de referencia de Ag / AgCl en contacto con un músculo de la espalda. </li></ol> 12. Paso Doce Estimular el nervio tríceps suralusando un 50 microsegundos pulso cuadrado de intensidad creciente a una frecuencia baja (&lt;1 Hz), hasta que la amplitud de contracción máxima se observa. Lentamente mover el transductor de fuerza para estirar el músculo, mientras que el control de la amplitud de la respuesta de contracción hasta que la amplitud de contracción alcanza un máximo. 13. Registros intracelulares de las neuronas motoras Desde este punto en adelante, las técnicas electrofisiológicas estándar se utilizan para preparar un electrodo intracelular, una neurona penetrar en la médula espinal y lo identifican como un motoneuron. <ol><li> Tire de una micropipeta de vidrio a una punta de m ~ 1, utilizando un extractor de pipeta (P-97 Micropipeta Puller, Instrumentos Sutter). Llenar el electrodo con una solución de KCl 3M (resistencia del electrodo de 10-20 mW). </li><li> Uso de un microposicionador, conducir la micropipeta, conectado a un amplificador intracelular (Axoclamp 2B, Axon Instruments) en la médula espinal. Monitorear los potenciales de campo locales producidas por la estimulación de correoada para localizar el nervio motor de la piscina del tríceps sural. </li><li> Cuidadosamente acercarse motoneuronas putativo mientras se monitoriza la resistencia del microelectrodo. Cuando se empuja contra una membrana, la resistencia aumenta. Penetración en algún momento puede ser facilitado mediante el &quot;buzz&quot; la función del amplificador intracelular. </li></ol> 14. La eutanasia Procedimiento Al final del experimento, los animales se sacrificaron mediante una sobredosis de pentobarbital (210 mg / kg IV), seguido de decapitación.

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No hay conflictos de interés declarado.

La preparación se describe aquí es el primero que permite, en el ratón adulto, la grabación simultánea intracelular de una motoneuron lumbar y la medición de la fuerza producida por las fibras musculares inervadas por su axón. Debido al pequeño tamaño del animal, las habilidades quirúrgicas requeridas para esta preparación puede ser difícil de adquirir. Sin embargo, una vez que esas habilidades se dominan, toda la cirugía se puede realizar en tres horas, y los animales pueden so...

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> Nombre del reactivo </td><td> Empresa </td><td> Número de catálogo </td><td> Comentarios (opcional) </td></tr><tr><td> Atropina sulfato </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> Sodio pentobarbital </td><td> Sanofi-Aventis </td><td> Pentobarbital </td><td></td></tr><tr><td> La ketamina </td><td></td><td></td><td></td></tr><tr><td> Xilazina </td><td></td><td></td><td></td></tr><tr><td> Glucosa </td><td></td><td></td><td></td></tr><tr><td> Expansor plasmático </td><td> Roger Bellon </td><td> Plasmagel </td><td></td></tr><tr><td> Tijeras de punta roma </td><td> FST </td><td> 14079-10 </td><td></td></tr><tr><td> Blunt tijeras finas </td><td> FST </td><td> 15025-10 </td><td></td></tr><tr><td> Tijeras Vannas Primavera </td><td> FST </td><td> 15002-08 </td><td></td></tr><tr><td> Fine fórceps dentado </td><td> FST </td><td> 11370-32 </td><td></td></tr><tr><td> Fine fórceps dentado </td><td> FST </td><td> 11370-31 </td><td></td></tr><tr><td> Cunningham espinal Adaptador </td><td> Stoelting Co. </td><td></td><td></td></tr><tr><td> Kwik-Cast sellador </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> Capnografo </td><td> CWE Inc </td><td> μcapstar </td><td></td></tr><tr><td> Calefacción manta </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> Pipetear extractor </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

La motoneurona espinal ha sido durante mucho tiempo un buen sistema modelo para el estudio de la función neural, ya que es una neurona del sistema nervioso central con las propiedades únicas de (1) con objetivos fácilmente identificables (las fibras musculares) y por lo tanto tiene una función muy conocida (para controlar la contracción muscular), (2) ser el objetivo convergente de muchas redes espinales y descendente, de ahí el nombre de &quot;vía final común&quot;, y (3) que tiene una gran soma que hace posible penetrar en ellos con afilados electrodos intracelulares . Además, cuando se estudió in vivo, es posible registrar simultáneamente la actividad eléctrica de las neuronas motoras y la fuerza desarrollada por sus objetivos musculares. Realización de grabaciones intracelulares de las neuronas motoras en vivo por lo tanto poner el experimentador en la posición única de ser capaz de estudiar, al mismo tiempo, todos los compartimentos de la &quot;unidad de motor&quot; (el nombre dado a la motoneurona, su axón, ylas fibras musculares que inerva 1): los insumos que inciden en la motoneurona, las propiedades electrofisiológicas de las neuronas motoras, y el impacto de estas propiedades de la función fisiológica de las neuronas motoras, es decir, la fuerza producida por su unidad de motor. Sin embargo, este enfoque es muy difícil porque la preparación no puede ser paralizada y por lo tanto la estabilidad mecánica para el registro intracelular se reduce. Así, este tipo de experimentos sólo se ha logrado en los gatos y en ratas. Sin embargo, el estudio de los sistemas motores espinales podría dar un salto formidable si era posible llevar a cabo experimentos similares en ratones normales y genéticamente modificados. Por razones técnicas, el estudio de las redes espinal en ratones se ha limitado principalmente a recién nacidos en preparaciones in vitro, donde las neuronas motoras espinales y las redes son inmaduros, las motoneuronas son separados de sus objetivos, y cuando se estudió en rodajas, el motoneurons se separan de la mayoría de sus entradas. Hasta hace poco, sólo algunos grupos habían logrado realizar grabaciones intracelulares de las neuronas motoras en vivo 2-4, incluyendo nuestro equipo, que publicó una nueva preparación que nos ha permitido obtener grabaciones muy estables de las motoneuronas in vivo en ratones adultos 5,6. Sin embargo, estos registros se obtuvieron en animales paralizados, es decir, sin la posibilidad de grabar la salida de la fuerza de estas motoneuronas. A continuación les presentamos una extensión de esta preparación original en el que hemos sido capaces de obtener grabaciones simultáneas de las propiedades electrofisiológicas de las neuronas motoras y de la fuerza desarrollada por su unidad de motor. Este es un logro importante, ya que nos permite identificar los diferentes tipos de neuronas motoras en base a su perfil de la fuerza, y revelando así su función. Junto con los modelos genéticos perturbar circuitos espinal segmentaria 7-9, o reproducting humano enferme10,11 sí, esperamos que esta técnica es una herramienta esencial para el estudio del sistema motor espinal.

La Figura 1 muestra cómo identificar un motoneuron del grupo tríceps sural después de la penetración. En la intensidad de estimulación mínima, sólo una EPSP monosináptico se puede observar (figura 1A). A mayor intensidad, la EPSP podría ser lo suficientemente grande como para provocar una &quot;ortodrómica&quot; punta (Figura 1B). En la intensidad de estimulación incluso superior, un todo-o-nada antidrómica pico aparece, con una latencia más corta que la EP...

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 nuevo método permite la grabación simultánea intracelular de una motoneuron adulto solo ratón y la medición de la fuerza producida por sus fibras musculares. La investigación combinada de las propiedades eléctricas y mecánicas de las unidades motoras en los animales normales y genéticamente modificadas es un gran avance para el estudio del sistema neuromuscular.

Grabación simultánea intracelular de una motoneurona lumbar y la Fuerza producida por su unidad motora en el ratón adulto<em&gt; En 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 vistas.  Northwestern University Feinberg School of Medicine. Este nuevo método permite la grabación simultánea intracelular de una motoneuron adulto solo ratón y la medición de la fuerza producida por sus fibras musculares. La investigación combinada de las propiedades eléctricas y mecánicas de las unidades motoras en los animales normales y genéticamente modificadas es un gran avance para el estudio del sistema neuromuscular.

Video: 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.

La transfección de células de ratón del ganglio de la retina por In vivo La electroporación

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

Investigación

Neurociencias

A Functional Motor Unit in the Culture Dish: Co-culture of Spinal Cord Explants and Muscle Cells

Human primary muscle cells cultured aneurally in monolayer rarely contract spontaneously because, in the absence of a nerve component, cell differentiation is limited and motor neuron stimulation is missing1. These limitations hamper the in vitro study of many neuromuscular diseases in cultured muscle cells. Importantly, the experimental constraints of monolayered, cultured muscle cells can be overcome by functional innervation of myofibers with spinal cord explants in co-cultures.
Here, we show the different steps required to achieve an efficient, proper innervation of human primary muscle cells, leading to complete differentiation and fiber contraction according to the method developed by Askanas2. To do so, muscle cells are co-cultured with spinal cord explants of rat embryos at ED 13.5, with the dorsal root ganglia still attached to the spinal cord slices. After a few days, the muscle fibers start to contract and eventually become cross-striated through innervation by functional neurites projecting from the spinal cord explants that connecting to the muscle cells. This structure can be maintained for many months, simply by regular exchange of the culture medium.
The applications of this invaluable tool are numerous, as it represents a functional model for multidisciplinary analyses of human muscle development and innervation. In fact, a complete de novo neuromuscular junction installation occurs in a culture dish, allowing an easy measurement of many parameters at each step, in a fundamental and physiological context.
Just to cite a few examples, genomic and/or proteomic studies can be performed directly on the co-cultures. Furthermore, pre- and post-synaptic effects can be specifically and separately assessed at the neuromuscular junction, because both components come from different species, rat and human, respectively. The nerve-muscle co-culture can also be performed with human muscle cells isolated from patients suffering from muscle or neuromuscular diseases3, and thus can be used as a screening tool for candidate drugs. Finally, no special equipment but a regular BSL2 facility is needed to reproduce a functional motor unit in a culture dish. This method thus is valuable for both the muscle as well as the neuromuscular research communities for physiological and mechanistic studies of neuromuscular function, in a normal and disease context.

Cultured muscle cells are an inadequate model to recapitulate innervated muscle in vivo. A functional motor unit can be reproduced in vitro by innervation of differentiated human primary muscle cells using rat embryo spinal cord explants. This article describes how co-cultures of spinal cord explants and muscle cells are established.

Una unidad motora funcional en la placa de cultivo: Co-cultivo de explantes de la médula espinal y las células musculares

Human primary muscle cells cultured  aneurally in monolayer rarely contract spontaneously because, in the absence of  a nerve component, cell ...

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.

Marcaje fluorescente retrógrada Permite dirigida extracelular sola unidad de grabación de neuronas identificadas<em&gt; En 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.

Time-lapse de imágenes de Migración neuroblasto en rodajas agudas del cerebro anterior ratón 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.

Varias unidades Métodos de grabación para caracterizar la actividad neuronal en el Locust (<em&gt; Schistocerca americana</em&gt;) Circuitos olfativas

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

A Guide to In vivo Single-unit Recording from Optogenetically Identified Cortical Inhibitory Interneurons

A major challenge in neurophysiology has been to characterize the response properties and function of the numerous inhibitory cell types in the cerebral cortex. We here share our strategy for obtaining stable, well-isolated single-unit recordings from identified inhibitory interneurons in the anesthetized mouse cortex using a method developed by Lima and colleagues1. Recordings are performed in mice expressing Channelrhodopsin-2 (ChR2) in specific neuronal subpopulations. Members of the population are identified by their response to a brief flash of blue light. This technique &#8211; termed &#8220;PINP&#8221;, or Photostimulation-assisted Identification of Neuronal Populations &#8211; can be implemented with standard extracellular recording equipment. It can serve as an inexpensive and accessible alternative to calcium imaging or visually-guided patching, for the purpose of targeting extracellular recordings to genetically-identified cells. Here we provide a set of guidelines for optimizing the method in everyday practice. We refined our strategy specifically for targeting parvalbumin-positive (PV+) cells, but have found that it works for other interneuron types as well, such as somatostatin-expressing (SOM+) and calretinin-expressing (CR+) interneurons.

A Guide to In vivo Single-unit Recording from Optogenetically Identified Cortical Inhibitory Interneurons

Here we describe our strategy for obtaining stable, well-isolated single-unit recordings from identified inhibitory interneurons in the anesthetized mouse cortex. Neurons expressing ChR2 are identified by their response to blue light. The method uses standard extracellular recording equipment, and serves as an inexpensive alternative to calcium imaging or visually-guided patching.

Una guía para<em&gt; En vivo</em&gt; Single-unidad de grabación de Optogenetically Identificado cortical inhibitorio interneuronas

A major challenge in neurophysiology has been to characterize the response properties and function of the numerous inhibitory cell types in the cerebral ...

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.

Simultáneo<em&gt; Ex vivo</em&gt; Pruebas funcionales de dos retinas por<em&gt; In vivo</em&gt; Sistema Electrorretinograma

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

Simultaneous Two-photon In Vivo Imaging of Synaptic Inputs and Postsynaptic Targets in the Mouse Retrosplenial Cortex

This video shows the craniotomy procedure that allows chronic imaging of neurons in the mouse retrosplenial cortex (RSC) using in vivo two-photon microscopy in Thy1-GFP transgenic mouse line. This approach creates a possibility to investigate the correlation of behavioural manipulations with changes in neuronal morphology in vivo.
The cranial window implantation procedure was considered to be limited only to the easily accessible cortex regions such as the barrel field. Our approach allows visualization of neurons in the highly vascularized RSC. RSC is an important element of the brain circuit responsible for spatial memory, previously deemed to be problematic for in vivo two-photon imaging.
The cranial window implantation over the RSC is combined with an injection of mCherry-expressing recombinant adeno-associated virus (rAAVmCherry) into the dorsal hippocampus. The expressed mCherry spreads out to axonal projections from the hippocampus to RSC, enabling the visualization of changes in both presynaptic axonal boutons and postsynaptic dendritic spines in the cortex. 
This technique allows long-term monitoring of experience-dependent structural plasticity in RSC.

Simultaneous Two-photon In Vivo Imaging of Synaptic Inputs and Postsynaptic Targets in the Mouse Retrosplenial Cortex

This video shows the craniotomy procedure that allows chronic imaging of neurons in mouse retrosplenial cortex using in vivo two photon microscopy in Thy1-GFP transgenic line. This approach is combined with injection of mCherry-expressing adeno-associated virus into dorsal hippocampus. These techniques allow long-term monitoring of experience-dependent structural plasticity in RSC.

Simultánea de dos fotones<em&gt; En Vivo</em&gt; Imágenes de entradas sinápticas y Objetivos postsinápticos en la corteza del ratón retroesplenial

This video shows the craniotomy procedure that allows chronic imaging of neurons in the mouse retrosplenial cortex (RSC) using in vivo two-photon ...

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 electrofisiológico para registrar las respuestas de voltaje intracelular<em&gt; Drosophila</em&gt; Los fotorreceptores y interneuronas a la luz Estímulos<em&gt; En Vivo</em

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

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.