Financiado por la NSF (0854551).

1. Pre-experimental Preparación <ol><li> Preparar las soluciones siguientes (véase el cuadro 1 para las composiciones): (a) 1 litro NFR (Ringer Frog Normal), (b) 100 ml de solución salina al 10%, (c) 100 ml CMF (Ca 2 + / Mg 2 + sin solución ), (d) 1 ml ITS (insulina-transferrina-selenio, Sigma I1884), (e) 100 ml de medio L-15 de la cultura, (f) 10 ml de HCG (gonadotropina coriónica humana Sigma CG-10), (g) 100 ml K +-solución interna, (h) 100 ml K +-interna solución de anfotericina B. La fuerza osmótica de la solución 1 se debe comprobar para asegurarse de que es aproximadamente 260 mOsM utilizando un osmómetro de presión de vapor (por ejemplo, modelo Wescor # 5100C) . </li><li> Filtrar esterilizar soluciones de (b) y (c). Soluciones (a) (b) y (c) se pueden almacenar hasta tres meses a 4 ° C. Preparar la solución de (d) mediante la adición de 1 ml de H2O desionizada al vial que contiene el polvo liofilizado a través de una aguja de calibre 25 y jeringa filtro de jeringa. Esta última solución cun ser almacenado a 4 º C durante 30 días. </li><li> Preparar la solución de (e) en una campana de flujo laminar mediante la combinación de todos los componentes en un vaso de precipitados o matraz. Esterilizar por filtración la solución final y transferir alícuotas de 10 ml en tubos de centrífuga estériles. Conservar a -20 ° C. </li><li> Preparar la HCG (solución (f)) mediante la adición de 10 mL de agua desionizada H 2 0 al vial que contiene el polvo liofilizado a través de una aguja de calibre 25 y jeringa filtro de jeringa. Esta solución final se puede almacenar a 4 ° C durante 30 días. </li><li> Fabricar al menos dos herramientas de microdisección por pegado un pasador Minutien (26002-10, herramientas Fine Science) al extremo de una pipeta Pasteur de vidrio con pegamento de cianoacrilato. Permitir que el extremo afilado de la espiga se extienda más allá del extremo de la pipeta de aproximadamente 0,5 cm. </li><li> Dos días antes de la preparación de cultivos, inducir la cría con el siguiente procedimiento. Identificar una pareja reproductora-ready de Xenopus mediante la observación de una cloaca prominente, de color rojizo en la pigmentación de la mujer y la oscuridad en el su planorface de las patas delanteras de la masculina. Uno a la vez, neta de cada rana y manténgalo lado ventral hacia abajo sobre un lavadero con la red de modo que no puede escapar. Inyectar 1 ml de la solución de (f) a través de la red y por vía subcutánea en uno de los sacos linfáticos dorsales. </li><li> Coloque la pareja reproductora juntos en un tanque de diez cubiertas de agua. Para asegurarse de que los animales no se pisotean los huevos recién puestos y fertilizados, instale un piso de malla con un tamaño de malla de aproximadamente ½ &quot;provistos de aproximadamente 1-2 cm por encima del fondo del tanque (Figura 1A). </li><li> Deja ranas perturbadas durante 12-48 horas hasta que los animales están en amplexus y los huevos fertilizados se observan en el piso debajo de la pantalla (Figura 1B). </li><li> Quite los animales del tanque, pero dejan los huevos en reposo durante al menos 24 horas más. Este par de ranas se puede volver a criados después de seis semanas. </li><li> Aflojar los embriones desde la parte inferior del tanque y transferirlos a cuatro o cinco culto mm 60x15ure platos que contienen 10% de solución salina (solución B). Clasificar los embriones por etapa según el esquema de Niewkoop y Faber (ref. 20). Embriones de interés serán aquellos en la etapa 22-24. Figura 2A muestra un embrión en etapa de aproximadamente 22, mientras que la Figura 2B muestra un embrión que es demasiado avanzado en el desarrollo de su utilidad (aproximadamente etapa 28). Es importante elegir embriones que son sanos: las que son lisas en apariencia con marrón claro y moteado blanco son ideales. Los embriones que tienen grandes manchas de color negro o blanco son generalmente insalubres e inutilizable. </li></ol> 2. Microdisección de embriones de Xenopus <ol><li> Dentro de una campana de flujo laminar, etiquetar y llenar hasta la mitad aproximadamente tres 60x15 mm platos estériles de cultivo con 10% de solución salina, y una de ellas con CMF. </li><li> Uso de una solución estéril, Pasteur de vidrio de pipeta de transferencia nueve y cincuenta y cinco etapa 22-24 embriones en una de las placas que contienen 10% de solución salina. Con la ayuda de un zoom estéreo diseccionarmicroscopio ing interior de la campana (0,6-5x con oculares 10 x), retire la capa gelatinosa y la membrana vitelina de cada embrión con dos pares de pinzas estériles º 5 (11251-30, herramientas Ciencias Artes). (Ver Figuras 3A y 3B.) </li><li> Lavar los embriones desnudos haciendo pasar ellos, uno a la vez, a través de los otros dos platos de 10% de solución salina y finalmente en el plato que contiene el CMF. Utilizar una pipeta estéril nueva para cada transferencia y minimizar el volumen de solución transferida de plato a plato. </li><li> A su vez, mantenga cada embrión suave, pero firmemente con un par de pinzas y, utilizando la herramienta de microdisección formado en el paso 1,5, extraer el tubo neural y miotomas asociados que están situados en el aspecto más dorsal del animal. Para ello, haciendo tres sectores, uno en cada extremo de la ubicación del tubo neural y la tercera sólo ventral a la misma (Figura 4A). Mover cada tubo neural diseccionado / miotoma a una parte limpia de la placa alejado de la yema GRANULES y otros desechos (Figura 4B). El eje dorso-ventral, y las localizaciones del tubo neural y miotomas se indica en la Figura 4. </li><li> Después de unos quince minutos en esta solución (CMF) pinzas de uso para levantar la piel pigmentada libre de los tejidos disecados y descarte. Después de un adicional de 30-60 min, las células se formará una &quot;pila de arena&quot;, como se disocian el uno del otro (Figura 4C). </li></ol> 3. Preparación del nervio-músculo Co-culturas <ol><li> Descongelar un tubo de 10 ml de medio de cultivo y añadir asépticamente 70 l ITS y 35 ng / ml de BDNF (Sigma B3795). </li><li> Etiqueta y llenar 35 cajas de cultivo estériles mm (FD35-100 World Precision Instruments) aproximadamente a mitad de camino con el medio de cultivo L-15 (solución de (e)), una para cada embrión disecado. </li><li> Se fabrica una pipeta chapado sujetando cada extremo de una pipeta Pasteur de vidrio mientras se mantiene la parte cónica sobre una llama. Tire de los extremos separados a 10 cm y se interrumpe el extremo para proporcionar una punta de aproximadamente 0,2 mm. </li><li> Use la pipeta para aspirar plating la &quot;pila de arena&quot; de un embrión minimizar la cantidad de solución dibujada. Expulsar las células en la parte inferior de una placa de cultivo. Placa de las células en varias líneas. Observe chapados en células no diferenciadas (Figura 5A y 5B). </li><li> Deja los platos de células establecidas inalteradas durante al menos quince minutos para que el tiempo las células se unan al plato. </li></ol> 4. Patch clamp nervio-músculo Sinapsis <ol><li> Después de 12-24 horas en cultivo, las células chapados asumir distintas características morfológicas: las células musculares se vuelven en forma de huso y las neuronas espinales ampliar los procesos largos (Figura 6). Contacto funcional sináptica entre las varices neuritas y células musculares se puede confirmar con parejas simultáneas registros electrofisiológicos. &quot;M&quot;, &quot;S&quot; y &quot;V&quot; se refieren respectivamente a la célula muscular, neuronalsoma y várices presináptica. </li><li> Preparar dos electrodos de parche para la grabación: una para la variz presináptica y uno para la célula muscular. </li><li> Para el electrodo presináptica, preparar una solución que contiene anfotericina como sigue: Añadir 100 l de DMSO a un tubo de microcentrífuga de 1,5 ml que contiene 5 mg de anfotericina B (Sigma A4888). Vortex esta solución durante 10 sec. A continuación, añadir 10 l de esta solución a un tubo de microcentrífuga que contiene 625 segundos K +-l interno solución de anfotericina B (solución de (h)). Vortex como anteriormente. </li><li> Llenar el electrodo pre-sináptica con la K que contiene anfotericina +-interna solución preparada en el paso 4,3 y el electrodo de post-sináptica con K +-interna solución (solución (g)). </li><li> Vuelva a colocar los medios de comunicación en la placa de cultivo con NFR y lo traslada a un microscopio invertido equipado con óptica de contraste de fase. </li><li> Coloque los dos electrodos justo por encima de sus respectivos objetivos. Obtener la cooperación de células enterasnfiguration con el electrodo de la célula muscular antes de la configuración de parche perforado con el electrodo de varicosidad. </li><li> Para confirmar la funcionalidad de la sinapsis, despolarizar la variz con un amplificador de patch clamp (por ejemplo Axopatch 200B, Molecular Devices) bajo control de software (por ejemplo, por pCLAMP 9, Molecular Devices) y observar las corrientes simultáneas presinápticos y postsinápticos. Figura 7 muestra las corrientes visto en respuesta a la despolarización pasos de voltaje a que la variz presináptica en incrementos de 10 mV desde -30 mV a +40 mV. Las corrientes hacia el interior se observan en la célula presináptica se realizan por Na + y Ca 2 + y las corrientes hacia el exterior de K +. Corrientes registradas en la célula postsináptica son respuestas a neurotransmisores liberados de la variz y se realizan principalmente por Na + a través de canales de receptores de acetilcolina nicotínicos. </li></ol>

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No hay conflictos de interés declarado.

Los pasos clave en el éxito de co-cultivo de las neuronas motoras y las células musculares son el uso de etapas apropiadamente embriones producidos a partir de la reproducción inducida de ranas Xenopus, y la cuidadosa disección aséptica de las neuronas espinales y mioblastos indiferenciados. Los huevos fertilizados se debe dejar reposar hasta que alcancen aproximadamente etapa 10 más o menos como en movimiento se detiene antes a menudo su desarrollo. Los embriones sanos se identifican por un aspecto suave y un color marrón y coloración blanco moteado. Etapa 22-24 embriones son las más útiles porque este es el punto durante el desarrollo justo después del cierre del tubo neural y antes de que los miocitos se han diferenciado significativamente. Además, las células de los embriones no mayores de disociar bien en CMF. Se debe tener cuidado al retirar la membrana vitelina, ya que se adhiere fuertemente al embrión. La membrana debe ser desgarrado utilizando pinzas cortantes para que el embrión emerge intacto. Otra importante precaution es cuidadosamente colocar las células en la parte inferior de la placa de cultivo (en lugar de dejar que establecerse allí). Este método es preferido ya que esto aumenta la probabilidad de que las células se adhieran a la placa de cultivo. Neurotransmisión funcional entre el nervio y el músculo se puede determinar con la grabación sólo postsináptica y, a menudo se puede determinar mediante la observación de la contracción muscular espontánea después de inervación. Un-inervados células musculares no crisparse en la cultura. Grabación Postsináptica solo es útil para grabar las corrientes de placa terminal en miniatura o potenciales, pero las mediciones de liberación provocada requiere estimulación presináptica, y la correlación de las corrientes de pre y postsinápticos requiere doble patch-clamp. Además del método de grabación de dos a dos parche descrito aquí, esta preparación ofrece la oportunidad de introducir una pipeta tercero en el soma neuronal 5 Esto permite la generación de un potencial de acción THAt se puede propagar a la variz presináptica y conducir a la liberación de neurotransmisores. Además, los agentes putativos que se cree que mediar o modular la transmisión sináptica puede ser introducido en cualquiera de los lados de la sinapsis a través de la pipeta célula muscular o a través de la difusión desde una pipeta tercero colocado en el soma.

<table border="1"><tbody><tr><td> Equipos / Suministros </td><td> Vendedor </td><td> Catálogo / Número de modelo </td><td></td></tr><tr><td> La insulina-transferrina-selenio </td><td> Sigma </td><td> I1884 </td><td></td></tr><tr><td> Gonadotropina coriónica humana </td><td> Sigma </td><td> CG-10 </td><td></td></tr><tr><td> Osmómetro de presión de vapor </td><td> Wescor </td><td> 5100C </td><td></td></tr><tr><td> Minutien Pines </td><td> Herramientas Artes Ciencias </td><td> 26002-10 </td><td></td></tr><tr><td> La anfotericina B </td><td> Sigma </td><td> A4888 </td><td></td></tr><tr><td> Fórceps </td><td> Herramientas Artes Ciencias </td><td> 11251-30 </td><td></td></tr><tr><td></td><td></td><td></td><td><table border="1"><tbody><tr><td> Solución Nombre </td><td> Receta </td></tr><tr><td> (A) NFR (Ringer rana normal (en mM) </td><td> 116 NaCl, 2 KCl, 1,8 CaCl 2, 5 Na +-HEPES, pH 7,35. </td></tr><tr><td> (B) solución salina 10% </td><td> 10 ml 90 ml NFR desionizada H 2 O </td></tr><tr><td> (C) CMF (Ca 2 + / Mg 2 + sin solución) </td><td> 125 NaCl, 2 KCl, 1,2 de EDTA, 5 Na +-HEPES, pH 7,35. </td></tr><tr><td> (E) Media L-15 Cultura </td><td> 50 ml de L-15 medios de comunicación (Sigma L1518) + 50 ml NFR. </td></tr><tr><td> (G) K +-solución interna (en mM) </td><td> 116 KCl, NaCl 1, 1 MgCl 2, 10 EGTA, 5 HEPES, pH 7,3 </td></tr><tr><td> (H) K +-interna solución de anfotericina B (en mM) </td><td> 52 K 2 SO 4, 38 KCl, 1 EGTA, 5 HEPES, pH 7,3 </td></tr></tbody></table></td></tr></tbody></table> Table 1.

simultaneous pre- and post-synaptic electrophysiological recording from xenopus nerve-muscle co-cultures

Mucha información sobre el acoplamiento de presinápticas corrientes iónicas con la liberación de neurotransmisor se ha obtenido a partir de preparaciones de invertebrados, especialmente el calamar gigante sinapsis 1. Sin embargo, excepto para la preparación descrito aquí, existen pocos preparativos vertebrados en el que es posible realizar mediciones simultáneas de la liberación de neurotransmisores presinápticos y corrientes iónicas. Motoneuronas embrionarias Xenopus y células musculares se pueden cultivar juntas en un medio de cultivo simple a temperatura ambiente, sino que se forman sinapsis funcionales dentro de doce hasta veinticuatro horas, y puede ser usado para estudiar nervio y el desarrollo de las células musculares y las interacciones sinápticas durante varios días (hasta crecimiento excesivo ocurre). Algunas de las ventajas de estos co-cultivos preparados a base de otros vertebrados incluyen la simplicidad de la preparación, la capacidad de mantener las culturas y el trabajo a temperatura ambiente, y la fácil accesibilidad de las sinapsis formadas 2-4. La preparación se ha utilizado ampliamente para estudiar las propiedades biofísicas de los canales de iones presináptica y la regulación de la liberación del transmisor 5-8. Además, la preparación se ha prestado a otros usos, incluyendo el estudio del crecimiento de neuritas y la sinaptogénesis 9-12, los mecanismos moleculares de la liberación de neurotransmisores 13-15, el papel de mensajeros difusibles en la neuromodulación 16,17, y en la plasticidad sináptica in vitro 18 - 19.

La figura 4B muestra la vista dorsal de un cordón espinal aislada / miotoma inmediatamente después de su extracción de una etapa 22 embriones de Xenopus. Figura 4C muestra que, después de la solución eliminación de la piel y la incubación en Ca 2 +-Mg 2 +-solución libre (CMF, ( c)), las células se disocian en una &quot;pila de arena&quot; y están listos para el revestimiento. Inmediatamente después de la siembra en medios de cultivo las células presentan poca variabilidad morfológica (Figura 5 B) sino que adquieren distintas formas después de veinticuatro horas de cultivo. Convertirse en células musculares en forma de huso mientras que las neuronas permanecen esférica mientras se extienden neuritas que varicosites elaborados en las sinapsis con las células musculares (Figura 6). Grabación simultánea emparejado parche de la varicosidad presináptica y postsináptica célula muscular (Figura 7) muestra las corrientes presinápticos entrada y de salida asociados con la liberación de neurotransmisores y resulta elnt excitatorios postsinápticos corrientes. <img src="/files/ftp_upload/50253/50253fig1.jpg" alt="Figura 1"/> Figura 1 A:.. Par Cría de ranas en balde apareamiento encima de la pantalla B: Ranas en amplexus con huevos fertilizados. <img src="/files/ftp_upload/50253/50253fig2.jpg" alt="Figura 2"/> Figura 2 A:. B &quot;adecuado&quot; etapa embrionaria 22:. &quot;Inadecuado&quot; etapa 28 embriones. La barra de escala representa 1 mm. <img src="/files/ftp_upload/50253/50253fig3.jpg" alt="Figura 3"/> Figura 3 A:. Stage 22 embriones antes de la extracción de la membrana vitelina y la capa de gelatina. La flecha apunta hacia el borde exterior de la capa gelatinosa. La membrana vitelina se adhiere estrechamente a la EMBRyo y es virtualmente invisible B:. embrión desnudo. La barra de escala representa 1 mm. <img src="/files/ftp_upload/50253/50253fig4.jpg" alt="Figura 4"/> Figura 4: Etapa 22 embriones con una línea discontinua indica a ser diseccionado parte B:.. Porción dorsal agudo aislado de embriones, &quot;d&quot; y &quot;v&quot; se refieren a las caras dorsal y ventral de los embriones, mientras que &quot;m&quot; y &quot;n&quot; indican la ubicación aproximada de las mytomes y el tubo neural C:. &quot;montón de arena&quot; de las células después de 60 minutos en CMF. La barra de escala representa 1 mm para A, y 0,5 mm para B y C. <img src="/files/ftp_upload/50253/50253fig5.jpg" alt="Figura 5"/> Figura 5 A:. Vista potencia 5x de células en cultivo inmediatamente después de placas. B: la misma cultura a 40x. Escalabarra representa 40 micras para A, y 5 micras para B. <img src="/files/ftp_upload/50253/50253fig6.jpg" alt="La figura 6"/> Figura 6. Unión neuromuscular en cultivo con identificación de soma neuronal (S), varicosidades presináptica (V), y células del músculo postsináptico (M). Barra de escala: 5 m. <img src="/files/ftp_upload/50253/50253fig7.jpg" alt="Figura 7" fo:content-width="5in" fo:src="/files/ftp_upload/50253/50253fig7highres.jpg"/> Figura 7. Presinápticos (parte superior) y postsinápticos (inferior) corrientes observadas en respuesta a la aplicación de voltaje de 10 mV incremental de pasos presentados a la variz presináptica de -30 mV a +40 mV. Pasos de voltaje se indican junto a algunas de las huellas presinápticos. El potencial de mantenimiento para ambas células fue -70 mV.

Watch this Scientific Journal Video about Simultaneous Pre- and Post-synaptic Electrophysiological Recording from Xenopus Nerve-muscle Co-cultures at JoVE.com

Simultaneous Pre- and Post-synaptic Electrophysiological Recording from Xenopus Nerve-muscle Co-cultures

Este video muestra los procedimientos utilizados para cultivar cultivos primarios de embrión<em&gt; Xenopus</em&gt; Células nerviosas y musculares y la utilidad de esta preparación para la fabricación de parches simultáneos grabaciones pre-y post sináptica-clamp.

Simultánea pre-y post-sináptica de registro electrofisiológico<em&gt; Xenopus</em&gt; Músculo Nervio-Co-culturas

Much information about the coupling of  presynaptic ionic currents with the release of neurotransmitter has been obtained  from invertebrate preparations, ...

simultaneous-pre-post-synaptic-electrophysiological-recording-from

Research

JoVE Journal

Neuroscience

Simultaneous Pre- and Post-synaptic Electrophysiological Recording from Xenopus Nerve-muscle Co-cultures

10.8K vistas.  David Geffen School of Medicine at UCLA. Este video muestra los procedimientos utilizados para cultivar cultivos primarios de embrión Xenopus Células nerviosas y musculares y la utilidad de esta preparación para la fabricación de parches simultáneos grabaciones pre-y post sináptica-clamp.

Video: Simultaneous Pre- and Post-synaptic Electrophysiological Recording from Xenopus Nerve-muscle Co-cultures

Derivation of Enriched Oligodendrocyte Cultures and Oligodendrocyte/Neuron Myelinating Co-cultures from Post-natal Murine Tissues

Identifying the molecular mechanisms underlying OL development is not only critical to furthering our knowledge of OL biology, but also has implications for understanding the pathogenesis of demyelinating diseases such as Multiple Sclerosis (MS). Cellular development is commonly studied with primary cell culture models. Primary cell culture facilitates the evaluation of a given cell type by providing a controlled environment, free of the extraneous variables that are present in vivo. While OL cultures derived from rats have provided a vast amount of insight into OL biology, similar efforts at establishing OL cultures from mice has been met with major obstacles. Developing methods to culture murine primary OLs is imperative in order to take advantage of the available transgenic mouse lines. 
Multiple methods for extraction of OPCs from rodent tissue have been described, ranging from neurosphere derivation, differential adhesion purification and immunopurification 1-3. While many methods offer success, most require extensive culture times and/or costly equipment/reagents. To circumvent this, purifying OPCs from murine tissue with an adaptation of the method originally described by McCarthy &amp; 
de Vellis 2 is preferred. This method involves physically separating OPCs from a mixed glial culture derived from neonatal rodent cortices. The result is a purified OPC population that can be differentiated into an OL-enriched culture. This approach is appealing due to its relatively short culture time and the unnecessary requirement for growth factors or immunopanning antibodies. 
While exploring the mechanisms of OL development in a purified culture is informative, it does not provide the most physiologically relevant environment for assessing myelin sheath formation. Co-culturing OLs with neurons would lend insight into the molecular underpinnings regulating OL-mediated myelination of axons. For many OL/neuron co-culture studies, dorsal root ganglion neurons (DRGNs) have proven to be the neuron type of choice. They are ideal for co-culture with OLs due to their ease of extraction, minimal amount of contaminating cells, and formation of dense neurite beds. While studies using rat/mouse myelinating xenocultures have been published 4-6, a method for the derivation of such OL/DRGN myelinating co-cultures from post-natal murine tissue has not been described. Here we present detailed methods on how to effectively produce such cultures, along with examples of expected results. These methods are useful for addressing questions relevant to OL development/myelinating function, and are useful tools in the field of neuroscience.

This article describes methods to derive enriched populations of murine oligodendrocyte precursor cells (OPCs) in primary culture, which differentiate to produce mature oligodendrocytes (OLs). In addition, this report describes techniques to produce murine myelinating co-cultures by seeding mouse OPCs onto a neurite bed of mouse dorsal root ganglion neurons (DRGNs).

Derivación de las Culturas enriquecido oligodendrocitos y neuronas oligodendrocitos / myelinating Co-culturas de Post-natal tejidos murinos

Identifying the molecular mechanisms underlying OL development is not only critical to furthering our knowledge of OL biology, but also has implications ...

Investigación

Neurociencias

Simultaneous Recording of Calcium Signals from Identified Neurons and Feeding Behavior of Drosophila melanogaster

To study neuronal networks in terms of their function in behavior, we must analyze how neurons operate when each behavioral pattern is generated. Thus, simultaneous recordings of neuronal activity and behavior are essential to correlate brain activity to behavior. For such behavioral analyses, the fruit fly, Drosophila melanogaster, allows us to incorporate genetically encoded calcium indicators such as GCaMP1, to monitor neuronal activity, and to use sophisticated genetic manipulations for optogenetic or thermogenetic techniques to specifically activate identified neurons2-5. Use of a thermogenetic technique has led us to find critical neurons for feeding behavior (Flood et al., under revision). As a main part of feeding behavior, a Drosophila adult extends its proboscis for feeding6 (proboscis extension response; PER), responding to a sweet stimulus from sensory cells on its proboscis or tarsi. Combining the protocol for PER7 with a calcium imaging technique8 using GCaMP3.01, 9, I have established an experimental system, where we can monitor activity of neurons in the feeding center &#x2013; the suboesophageal ganglion (SOG), simultaneously with behavioral observation of the proboscis. I have designed an apparatus ("Fly brain Live Imaging and Electrophysiology Stage": "FLIES") to accommodate a Drosophila adult, allowing its proboscis to freely move while its brain is exposed to the bath for Ca2+ imaging through a water immersion lens. The FLIES is also appropriate for many types of live experiments on fly brains such as electrophysiological recording or time lapse imaging of synaptic morphology. Because the results from live imaging can be directly correlated with the simultaneous PER behavior, this methodology can provide an excellent experimental system to study information processing of neuronal networks, and how this cellular activity is coupled to plastic processes and memory.

Simultaneous Recording of Calcium Signals from Identified Neurons and Feeding Behavior of Drosophila melanogaster

The fruit fly, Drosophila melanogaster, extends its proboscis for feeding, responding to a sugar stimulus from its proboscis or tarsus. I have combined observations of the proboscis extension response (PER) with a calcium imaging technique, allowing us to monitor the activity of neurons in the brain, simultaneously with behavioral observation.

La grabación simultánea de señales de calcio en neuronas identificadas y comportamiento de alimentación de Drosophila melanogaster</em

To study neuronal networks in terms of their function in behavior, we must analyze how neurons operate when each behavioral pattern is generated.  Thus, ...

Neural Explant Cultures from Xenopus laevis

The complex process of axon guidance is largely driven by the growth cone, which is the dynamic motile structure at the tip of the growing axon. During axon outgrowth, the growth cone must integrate multiple sources of guidance cue information to modulate its cytoskeleton in order to propel the growth cone forward and accurately navigate to find its specific targets1. How this integration occurs at the cytoskeletal level is still emerging, and examination of cytoskeletal protein and effector dynamics within the growth cone can allow the elucidation of these mechanisms. Xenopus laevis growth cones are large enough (10-30 microns in diameter) to perform high-resolution live imaging of cytoskeletal dynamics (e.g.2-4 ) and are easy to isolate and manipulate in a lab setting compared to other vertebrates. The frog is a classic model system for developmental neurobiology studies, and important early insights into growth cone microtubule dynamics were initially found using this system5-7 . In this method8, eggs are collected and fertilized in vitro, injected with RNA encoding fluorescently tagged cytoskeletal fusion proteins or other constructs to manipulate gene expression, and then allowed to develop to the neural tube stage. Neural tubes are isolated by dissection and then are cultured, and growth cones on outgrowing neurites are imaged. In this article, we describe how to perform this method, the goal of which is to culture Xenopus laevis growth cones for subsequent high-resolution image analysis. While we provide the example of +TIP fusion protein EB1-GFP, this method can be applied to any number of proteins to elucidate their behaviors within the growth cone.

Neural Explant Cultures from Xenopus laevis

Culturing neural explants from dissected Xenopus laevis embryos that express fluorescent fusion proteins allows for imaging of growth cone cytoskeletal dynamics.

Los cultivos neuronales de explantes de<em&gt; Xenopus laevis</em

The complex process of axon guidance is largely driven by the growth cone, which is the dynamic motile structure at the tip of the growing axon. During ...

Forebrain Electrophysiological Recording in Larval Zebrafish

Epilepsy affects nearly 3 million people in the United States and up to 50 million people worldwide. Defined as the occurrence of spontaneous unprovoked seizures, epilepsy can be acquired as a result of an insult to the brain or a genetic mutation. Efforts to model seizures in animals have primarily utilized acquired insults (convulsant drugs, stimulation or brain injury) and genetic manipulations (antisense knockdown, homologous recombination or transgenesis) in rodents. Zebrafish are a vertebrate model system1-3 that could provide a valuable alternative to rodent-based epilepsy research. Zebrafish are used extensively in the study of vertebrate genetics or development, exhibit a high degree of genetic similarity to mammals and express homologs for ~85% of known human single-gene epilepsy mutations. Because of their small size (4-6 mm in length), zebrafish larvae can be maintained in fluid volumes as low as 100 &mu;l during early development and arrayed in multi-well plates. Reagents can be added directly to the solution in which embryos develop, simplifying drug administration and enabling rapid in vivo screening of test compounds4. Synthetic oligonucleotides (morpholinos), mutagenesis, zinc finger nuclease and transgenic approaches can be used to rapidly generate gene knockdown or mutation in zebrafish5-7. These properties afford zebrafish studies an unprecedented statistical power analysis advantage over rodents in the study of neurological disorders such as epilepsy. Because the "gold standard" for epilepsy research is to monitor and analyze the abnormal electrical discharges that originate in a central brain structure (i.e., seizures), a method to efficiently record brain activity in larval zebrafish is described here. This method is an adaptation of conventional extracellular recording techniques and allows for stable long-term monitoring of brain activity in intact zebrafish larvae. Sample recordings are shown for acute seizures induced by bath application of convulsant drugs and spontaneous seizures recorded in a genetically modified fish.

A simple method to record extracellular field potentials in the larval zebrafish forebrain is described. The method provides a robust in vivo read-out of seizure-like activity. This technique can be used with genetically modified zebrafish larvae carrying epilepsy-related genes or seizures evoked by administration of convulsant drugs.

Prosencéfalo electrofisiológico de grabación en larvas de pez cebra

Epilepsy affects nearly 3 million people in the United States and up to 50 million people worldwide. Defined as the occurrence of spontaneous unprovoked ...

Simultaneous Electrophysiological Recording and Calcium Imaging of Suprachiasmatic Nucleus Neurons

Simultaneous electrophysiological and fluorescent imaging recording methods were used to study the role of changes of membrane potential or current in regulating the intracellular calcium concentration. Changing environmental conditions, such as the light-dark cycle, can modify neuronal and neural network activity and the expression of a family of circadian clock genes within the suprachiasmatic nucleus (SCN), the location of the master circadian clock in the mammalian brain. Excitatory synaptic transmission leads to an increase in the postsynaptic Ca2+ concentration that is believed to activate the signaling pathways that shifts the rhythmic expression of circadian clock genes. Hypothalamic slices containing the SCN were patch clamped using microelectrodes filled with an internal solution containing the calcium indicator bis-fura-2. After a seal was formed between the microelectrode and the SCN neuronal membrane, the membrane was ruptured using gentle suction and the calcium probe diffused into the neuron filling both the soma and dendrites. Quantitative ratiometric measurements of the intracellular calcium concentration were recorded simultaneously with membrane potential or current. Using these methods it is possible to study the role of changes of the intracellular calcium concentration produced by synaptic activity and action potential firing of individual neurons. In this presentation we demonstrate the methods to simultaneously record electrophysiological activity along with intracellular calcium from individual SCN neurons maintained in brain slices.

Procedures are described to perform simultaneous recordings of membrane potential or current and changes of intracellular calcium concentration. Suprachiasmatic nucleus neurons are filled with the calcium indicator bis-fura-2 using a patch clamp electrode in the whole cell patch clamp configuration.

Simultaneous electrophysiological and fluorescent imaging recording methods were used to study the role of changes of membrane potential or current in ...

Electrophysiological Recording in the Brain of Intact Adult Zebrafish

Previously, electrophysiological studies in adult zebrafish have been limited to slice preparations or to eye cup preparations and electrorentinogram recordings. This paper describes how an adult zebrafish can be immobilized, intubated, and used for in vivo electrophysiological experiments, allowing recording of neural activity. Immobilization of the adult requires a mechanism to deliver dissolved oxygen to the gills in lieu of buccal and opercular movement. With our technique, animals are immobilized and perfused with habitat water to fulfill this requirement. A craniotomy is performed under tricaine methanesulfonate (MS-222; tricaine) anesthesia to provide access to the brain. The primary electrode is then positioned within the craniotomy window to record extracellular brain activity. Through the use of a multitube perfusion system, a variety of pharmacological compounds can be administered to the adult fish and any alterations in the neural activity can be observed. The methodology not only allows for observations to be made regarding changes in neurological activity, but it also allows for comparisons to be made between larval and adult zebrafish. This gives researchers the ability to identify the alterations in neurological activity due to the introduction of various compounds at different life stages.

This paper describes how an adult zebrafish can be immobilized, intubated, and used for in vivo electrophysiological experiments to allow recordings and manipulation of neural activity in an intact animal.

Registro electrofisiológico en el cerebro del pez cebra adultos Intacto

Previously, electrophysiological studies in adult zebrafish have been limited to slice preparations or to eye cup preparations and electrorentinogram ...

A Method for Systematic Electrochemical and Electrophysiological Evaluation of Neural Recording Electrodes

New materials and designs for neural implants are typically tested separately, with a demonstration of performance but without reference to other implant characteristics.&#160;This precludes a rational selection of a particular implant as optimal for a particular application and the development of new materials based on the most critical performance parameters.&#160;This article develops a protocol for in vitro and in vivo testing of neural recording electrodes.&#160;Recommended parameters for electrochemical and electrophysiological testing are documented with the key steps and potential issues discussed.&#160;This method eliminates or reduces the impact of many systematic errors present in simpler in vivo testing paradigms, especially variations in electrode/neuron distance and between animal models.&#160;The result is a strong correlation between the critical in vitro and in vivo responses, such as impedance and signal-to-noise ratio.&#160;This protocol can easily be adapted to test other electrode materials and designs.&#160;The in vitro techniques can be expanded to any other nondestructive method to determine further important performance indicators.&#160;The principles used for the surgical approach in the auditory pathway can also be modified to other neural regions or tissue.

Different electrode coatings affect neural recording performance through changes to electrochemical, chemical and mechanical properties. Comparison of electrodes in vitro is relatively simple, however comparison of in vivo response is typically complicated by variations in electrode/neuron distance and between animals. This article provides a robust method to compare neural recording electrodes.

Un método para Sistemática Electroquímica y electrofisiológico Evaluación de Neuronales Grabación Electrodos

New materials and designs for neural implants are typically tested separately, with a demonstration of performance but without reference to other implant ...

Electrophysiological Recording From Drosophila Labellar Taste Sensilla

The peripheral taste response of insects can be powerfully investigated with electrophysiological techniques. The method described here allows the researcher to measure gustatory responses directly and quantitatively, reflecting the sensory input that the insect nervous system receives from taste stimuli in its environment. This protocol outlines all key steps in performing this technique. The critical steps in assembling an electrophysiology rig, such as selection of necessary equipment and a suitable environment for recording, are delineated. We also describe how to prepare for recording by making appropriate reference and recording electrodes, and tastant solutions. We describe in detail the method used for preparing the insect by insertion of a glass reference electrode into the fly in order to immobilize the proboscis. We show traces of the electrical impulses fired by taste neurons in response to a sugar and a bitter compound. Aspects of the protocol are technically challenging and we include an extensive description of some common technical challenges that may be encountered, such as lack of signal or excessive noise in the system, and potential solutions. The technique has limitations, such as the inability to deliver temporally complex stimuli, observe background firing immediately prior to stimulus delivery, or use water-insoluble taste compounds conveniently. Despite these limitations, this technique (including minor variations referenced in the protocol) is a standard, broadly accepted procedure for recording Drosophila neuronal responses to taste compounds.

Electrophysiological Recording From Drosophila Labellar Taste Sensilla

This protocol describes extracellular recording of the action potential responses fired by labellar taste neurons in Drosophila.

Electrofisiológico Grabación desde<em&gt; Drosophila</em&gt; Labellar Taste Sensilla

The peripheral taste response of insects can be powerfully investigated with electrophysiological techniques. The method described here allows the ...

Simultaneous Electrophysiological Recording and Micro-injections of Inhibitory Agents in the Rodent Brain

Here we describe a method for the construction of a single-use &#8220;injectrode&#8221; using commercially accessible and affordable parts. A probing system was developed that allows for the injection of a drug while recording electrophysiological signals from the affected neuronal population. This method provides a simple and economical alternative to commercial solutions. A glass pipette was modified by combining it with a hypodermic needle and a silver filament. The injectrode is attached to commercial microsyringe pump for drug delivery. This results in a technique that provides real-time pharmacodynamics feedback through multi-unit extracellular signals originating from the site of drug delivery. As a proof of concept, we recorded neuronal activity from the superior colliculus elicited by flashes of light in rats, concomitantly with delivery of drugs through the injectrode. The injectrode recording capacity permits the functional characterization of the injection site favoring precise control over the localization of drug delivery. Application of this method also extends far beyond what is demonstrated here, as the choice of chemical substance loaded into the injectrode is vast, including tracing markers for anatomic experiments.

Here, we craft a glass pipette with dual functions: inhibition of deep brain structures by microinjections of drugs and real-time monitoring of their effects through simultaneous electrophysiological recordings.

Grabación simultánea electrofisiológico y micro-inyecciones de agentes inhibitorios en el cerebro de roedores

Here we describe a method for the construction of a single-use &#8220;injectrode&#8221; using commercially accessible and affordable parts. A probing ...

Simultaneous Recording of Electroretinography and Visual Evoked Potentials in Anesthetized Rats

The electroretinogram (ERG) and visual evoked potential (VEP) are commonly used to assess the integrity of the visual pathway. The ERG measures the electrical responses of the retina to light stimulation, while the VEP measures the corresponding functional integrity of the visual pathways from the retina to the primary visual cortex following the same light event. The ERG waveform can be broken down into components that reflect responses from different retinal neuronal and glial cell classes. The early components of the VEP waveform represent the integrity of the optic nerve and higher cortical centers. These recordings can be conducted in isolation or together, depending on the application. The methodology described in this paper allows simultaneous assessment of retinal and cortical visual evoked electrophysiology from both eyes and both hemispheres. This is a useful way to more comprehensively assess retinal function and the upstream effects that changes in retinal function can have on visual evoked cortical function.

This protocol describes simultaneous measurement of electroretinogram and visual evoked potentials in anesthetized rats.

Grabación simultánea de Electrorretinografía y los potenciales evocados visuales en ratas anestesiadas

The electroretinogram (ERG) and visual evoked potential (VEP) are commonly used to assess the integrity of the visual pathway. The ERG measures the ...