Nos gustaría dar las gracias a los Profs. Huibert Mansvelder y Bert Sakmann de amplio apoyo, el Dr. Marcel Oberlaender para un debate fructífero y proporcionar rastreo neuronal, y Brendan Lodder para la asistencia técnica. Los datos fueron adquiridos utilizando el ntrode VI para LabVIEW, generosamente proporcionado por R. Bruno (Columbia Univ., NY, EE.UU.). Esta investigación fue apoyada por la Sociedad Max Planck y el Centro Bernstein para la Neurociencia Computacional, Tubinga (financiado por el Ministerio Federal Alemán de Educación e Investigación (BMBF; FKZ: 01GQ1002)) (RTN), Centro de Investigación Cognitiva y Neurogenomics (CNCR) , Campus de Neurociencias Amsterdam (NCA), la financiación de CPJdK (NWO-ALW # 822.02.013 y ENC-red # p3-c3) y la Universidad VU de Amsterdam.

1. Preparación del animal Todos los procedimientos experimentales se llevan a cabo de conformidad con la ley holandesa y después de una evaluación por un comité de ética local de la Universidad VU de Amsterdam, Países Bajos. <ol><li> Anestesiar a un rata Wistar (P25-P45, ♂ / ♀) con isoflurano (2-3% en oxígeno) y, posteriormente, con uretano (20% en 0,9% de NaCl, 1.6 hasta 1.7 g / kg) por inyección intraperitoneal. Evaluar la profundidad de la anestesia mediante la supervisión de la retirada pellizco, los reflejos palpebrales, y los movimientos de vibrisas. </li><li> Administrar una dosis adicional de uretano (10% de la dosis inicial) en caso de que el animal no llega a suficiente profundidad de la anestesia o cuando se observan espasmos vibrisas durante el curso del experimento. </li><li> Colocar el animal anestesiado en el marco estereotáxico equipado con una almohadilla eléctrica, inserte la sonda de temperatura rectal y mantener la temperatura corporal del animal a 37,5 ± 0,5 ° C por una almohadilla térmica. </li><li&gt; posición de la cabeza del animal en el marco estereotáxico; cortar el pelo en la parte superior de la cabeza con unas tijeras para hacer que la piel visible. </li><li> Inyectar 100 l 1% de lidocaína (en 0,9% de NaCl) por vía subcutánea para la anestesia local en el sitio de operación y esperar 3-5 minutos. Retire la piel que cubre la superficie del cráneo por el corte en la dirección-caudal a rostral y limpiar el tejido restante. </li><li> Limpiar el cráneo extensamente con 0,9% de NaCl y hisopos absorbentes. </li><li> Determinar las coordenadas de la zona de destino en el hemisferio izquierdo (para la corteza somatosensorial primaria (S1): 2,5 mm posterior y 5,5 mm lateral al bregma). Marque la ubicación en el cráneo con un rotulador quirúrgico de la piel. </li><li> Raspar el hueso suavemente por un taladro dental de la región de interés hasta que el hueso se vuelve transparente y los vasos sanguíneos son claramente visibles. </li><li> Hacer un 0,5 mm x 0,5 mm craneotomía mediante la reducción de una ventana en el cráneo adelgazada con un escalpelo (# 11), evitando daños a la duramadrey los vasos sanguíneos. Opcional: marca los bordes de la craneotomía utilizando un rotulador quirúrgico de la piel para mejorar la visibilidad. </li><li> Aplicar una capa delgada de pegamento en el cráneo seco y luego cemento dental para construir un baño (es decir, cámara de registro) que encierra la craneotomía. </li><li> Recorte los bigotes en el lado contralateral a exactamente 5 mm desde el folículo bigote. Opcional: poner de relieve los bigotes cortados con rímel negro. </li></ol> 2. Juxtasomal Grabaciones y Biocitina Etiquetado <ol><li> Hacer pipetas de parche de vidrio de borosilicato con diámetro de la punta dentro de ~ 1 m para obtener electrodos con 3-5 MΩ de resistencia. Nota: La morfología de la pipeta ideal para la grabación juxtasomal es una reducción gradual esbelto, un ángulo de cono y un ~ 1 m de la punta (Figura 1). </li><li> Dependiendo de la profundidad de grabación (con respecto a la superficie de la pía madre), las dimensiones de la forma cónica del electrodo de registro se deben ajustar. Es de importancia crítica es que tque se estrechan diámetro debe ser inferior a 75 micras, con punto de entrada en el cerebro para evitar daños mecánicos o estrés en el área de grabación. Esto indica que para las grabaciones corticales superficiales, se requiere una conicidad de 300-500 micras con un diámetro exterior máximo de 75 micras (Figura 1A), mientras que las grabaciones de relativamente profundas capas corticales requieren una conicidad más largo de 1.500-1.800 m, de nuevo con máxima externa diámetro de 75 micras (a 1.800 m de la punta del electrodo). </li><li> Cargue el parche pipeta con el timbre de la rata normal (NRR), complementado con un 2% biocitina (véase la Tabla 1 para las soluciones y reactivos) y montar el parche pipeta en una etapa de la cabeza fija a un micromanipulador. </li><li> Ajuste el ángulo del soporte de electrodo a 34 ° con respecto al plano sagital para dirigirse específicamente a la columna de la D2 de S1 de rata. </li><li> Conecte la etapa de la cabeza a un amplificador en modo puente (es decir, la pinza de corriente). </li><li> Coloque el parchepipetear en estrecha proximidad a la craneotomía. Llene la bañera con 0,9% de NaCl y determinar la resistencia del electrodo que debe estar entre 3-5 MΩ, evaluada mediante la aplicación de un pulso cuadrado con inyección de corriente positiva (1 nA, 200 msec on / off). </li><li> Establecer sobrepresión de 100 a 150 mbar y avanzar en la pipeta de parche en 1 m pasos mientras se aplica corriente positiva en forma de pulsos cuadrados (1 nA, 200 msec on / off). Monitorear el cambio de resistencia robusta al establecer contacto con la duramadre. En este punto, establecer las coordenadas del micromanipulador para &quot;cero&quot; para permitir mediciones precisas de la profundidad. </li><li> Avanzar en 1 m en modo paso hasta que el parche pipeta penetra la duramadre se indica por un descenso brusco de la resistencia del electrodo. Quite la presión de mantenimiento de la pipeta. </li><li> Búsqueda de unidades individuales mientras se avanza en 1 m pasos. Vigilar la resistencia del electrodo de forma continua mediante la aplicación de 200 ms on / off pulsos. Un aumento de la resistencia del electrodo typicaliado indica la proximidad de una sola neurona. Haga avanzar el electrodo hasta que el potencial de acción positiva (AP) formas de onda de ~ 2 mV se registran (Figuras 2A y 2B). </li><li> Opcional: grabar el patrón Rematar espontánea de la neurona y determinar bigote-evocado clavar por, por ejemplo, desviando caudalmente barbas individuales para 200 mseg en un ángulo de 3,3 ° utilizando un dispositivo piezoeléctrico 18. </li><li> Haga avanzar el electrodo hasta que la resistencia es 25-35 MΩ y picos tienen amplitudes de 3-8 mV para obtener condiciones óptimas de llenado juxtasomal. Inicie el llenado juxtasomal aplicando pulsos cuadrados de corriente positiva (1 nA, 200 ms on / off). Poco a poco y gradualmente aumentar la corriente por pasos de 0,1 nA mientras que el control de la forma de onda de AP y la frecuencia (Figuras 2A y 2B). </li><li> Supervisar la apertura de la membrana como un claro aumento en la frecuencia de AP durante la fase en el pulso de bloque (Figura 2C </strong&gt;). La forma de onda de pico durante el llenado muestra un aumento de la anchura y la reducida (Figuras 2C y 2D) post-hiperpolarización. Parámetros adicionales incluyen el aumento de ruido o una pequeña (1-5 mV) cambio negativo de CC. </li><li> Aumenta o disminuye la corriente (1 nA) durante el llenado para mantener la infusión biocitina estable. Reducir o detener los impulsos de corriente en aumento repentino de la frecuencia AP para evitar la toxicidad por el exceso de afluencia de iones extracelulares, tales como iones de sodio. Nota: cada neurona responde de manera diferente a la inyección de corriente y juxtasomal parámetros de etiquetado deben ser ajustados en función de las condiciones de grabación individuales. </li><li> Seguir de cerca la señal después de detener la inyección de corriente. La forma de onda pico después de una sesión de relleno se suele ampliarse y muestra una fuerte reducción después de hiperpolarización. Esperar a que la recuperación de la neurona, que es evidente cuando la forma de onda de pico vuelve a sus propiedades originales (es decir, presencia denormal, la figura post-hiperpolarización 2). </li><li> Repita biocitina llenar sesiones después de la recuperación completa de la neurona para aumentar la carga biocitina para mejorar la calidad de la tinción. </li><li> Retraer la pipeta de parche en pasos de 1 m hasta que la amplitud pico disminuye para reducir cualquier tensión mecánica a la célula. Espere por lo menos 1 hora para la biocitina para difundir de forma intracelular, mientras que de cerca el seguimiento de la temperatura corporal del animal y de la respiración. </li></ol> 3. Perfusión del animal y extracción del cerebro <ol><li> Preparar la configuración de la perfusión, enjuague y precargar el tubo con 0,9% de NaCl. </li><li> Coloque el animal sobre una bandeja quirúrgica y asegurar con cinta de etiquetado estándar. Asegurar suficiente profundidad de la anestesia; pizca pie y los reflejos de los párpados deben estar ausentes. </li><li> Hacer una incisión medial a lateral (5-6 cm) a través de la pared abdominal justo debajo de la caja torácica y proceder en la dirección posterior-anterior para exponer el esternón.Tire del esternón por delante y separar cuidadosamente el hígado desde el diafragma. </li><li> Hacer una pequeña incisión en el diafragma, cortar a través de las costillas inferiores y continuar la incisión a lo largo de toda la longitud de la cavidad abdominal para exponer el corazón. </li><li> Retire el pericardio. </li><li> Inserte la aguja en el ventrículo izquierdo y hacer una incisión en la aurícula derecha. Perfundir con 0,9% de NaCl (~ 8 ml / min) hasta la decoloración del hígado es completa. </li><li> Cambie la infusión de 4% de paraformaldehído (PFA), hasta que la rigidez de la pata delantera y la mandíbula inferior es aparente. </li><li> Decapita a la rata con un par de tijeras </li><li> Quitar los músculos del cuello que quedan y exponer el cráneo completo. </li><li> Coloque las tijeras en el tronco cerebral en la parte dorsal y cortar el hueso con cuidado a lo largo de la sutura sagital, el mantenimiento de la posición dorsal. </li><li> Retirar los huesos de ambos lados de la sutura sagital para exponer el cerebro usando fórceps. Retire con cuidado la duramadre para evitar damage. </li><li> Inserte con cuidado una espátula roma para la parte ventral del cerebro y quitar el cerebro suavemente. </li><li> Posterior a arreglar toda la noche a la mañana en el cerebro PFA a 4 ° C. Cambie el cerebro a tampón fosfato 0,05 M (PB) y almacenar a 4 ° C. </li></ol> 4. Cortando el cerebro en las secciones tangenciales <ol><li> Tome el cerebro fuera de la M PB 0.05 y lo puso en un papel de filtro frente anterior. Utilice una hoja de afeitar afilada para cortar el cerebelo a lo largo del plano coronal y separar los hemisferios cortando a lo largo mediados el plano sagital. </li><li> Aplique pegamento en la plataforma de montaje y montar el hemisferio izquierdo en su plano sagital con anterior frente a la derecha. Asegure la plataforma de montaje en un ángulo de 45 ° en un vibratome y sumergir el cerebro en 0,05 M PB. </li><li> Asegure una hoja de afeitar en la vibrátomo y asegurarse de que el primer contacto con la superficie del cerebro se encuentra en el centro del plano anterior-posterior del hemisferio. Cortar 24 100 msecciones y recogerlas en una placa de 24 pocillos que contenía 0,05 M PB. </li></ol> 5. Procedimientos histológicos <ol><li> Realizar protocolos histológicos para la tinción citocromo oxidasa 24 y el método de avidina-biotina-peroxidasa de acuerdo con los métodos descritos previamente 25. Opcional: visualice biocitina utilizando conjugados de avidina fluorescente / estreptavidina-Alexa. Esto permite, además, de doble tinción con técnicas de rastreo retrógrado o anterógrado. </li><li> Secciones de lavado de 5 x 5 min con 0,05 M PB y preparar el tetrahidrocloruro 3,-3&#39;-diaminobencidina (DAB) solución para la tinción de la citocromo oxidasa de visualizar barriles en la capa 4 (L4) de S1-que contiene (véase la Tabla 2 para las soluciones y reactivos). Incubar las secciones 6-12 de la pia en la solución precalentado durante 30 a 45 min a 37 ° C. </li><li> Enjuague secciones con 0,05 M PB de 6 x 5 min y la amortiguación actividad peroxidasa endógena por incubación de todas las secciones en el 3% de H 2 </sub&gt; O 2 en PB 0,05 M durante 20 min a temperatura ambiente (RT). </li><li> Enjuague secciones con 0,05 M PB de 5 x 10 min. Incubar las secciones en ABC-solución (véase la Tabla 3 para las soluciones y reactivos) la noche a 4 ° C. </li><li> Enjuague secciones con 0,05 M PB de 5 x 10 min y preparar la solución de DAB para visualizar la neurona biocitina lleno (véase la Tabla 4 para las soluciones y reactivos). Incubar las secciones en solución filtrada durante 45-60 min a TA. </li><li> Enjuague secciones con 0,05 M PB de 5 x 10 min. Mount secciones sobre portaobjetos de microscopio y cubreobjetos con Mowiol (véase la Tabla 5 para las soluciones y reactivos). </li><li> Determinar la calidad del etiquetado mediante microscopía de luz. </li></ol>

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P., Huber, D., Svoboda, K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Neural+activity+in+barrel+cortex+underlying+vibrissa-based+object+localization+in+mice.">Neural activity in barrel cortex underlying vibrissa-based object localization in mice.</a> Neuron. 67 (6), 1048-1061 (2010).</li><li>Veinante, P., Deschenes, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Single-cell+study+of+motor+cortex+projections+to+the+barrel+field+in+rats.">Single-cell study of motor cortex projections to the barrel field in rats.</a> J. Comp. Neurol. 464 (1), 98-103 (2003).</li><li>Boudewijns, Z. S., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Layer-specific+high-frequency+action+potential+spiking+in+the+prefrontal+cortex+of+awake+rats.">Layer-specific high-frequency action potential spiking in the prefrontal cortex of awake rats.</a> Front. Cell. Neurosci. 7, 99 (2013).</li><li>Oberlaender, M., Bruno, R. M., Sakmann, B., Broser, P. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Transmitted+light+brightfield+mosaic+microscopy+for+three-dimensional+tracing+of+single+neuron+morphology.">Transmitted light brightfield mosaic microscopy for three-dimensional tracing of single neuron morphology.</a> J. Biomed. Opt. 12 (6), 064029 (2007).</li><li>Boudewijns, Z. S., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Semi-automated+three-dimensional+reconstructions+of+individual+neurons+reveal+cell+type-specific+circuits+in+cortex.">Semi-automated three-dimensional reconstructions of individual neurons reveal cell type-specific circuits in cortex.</a> Commun. Integr. Biol. 4 (4), 486-488 (2011).</li><li>Bruno, R. M., Hahn, T. T., Wallace, D. J., de Kock, C. P., Sakmann, B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Sensory+experience+alters+specific+branches+of+individual+corticocortical+axons+during+development.">Sensory experience alters specific branches of individual corticocortical axons during development.</a> J. Neurosci. 29 (10), 3172-3181 (2009).</li><li>Schubert, D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Observing+without+disturbing:+how+different+cortical+neuron+classes+represent+tactile+stimuli.">Observing without disturbing: how different cortical neuron classes represent tactile stimuli.</a> J. Physiol. 581 (1), 5 (2007).</li><li>Neumann, E., Kakorin, S., Toensing, K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Fundamentals+of+electroporative+delivery+of+drugs+and+genes).">Fundamentals of electroporative delivery of drugs and genes).</a> Bioelectrochem. Bioenerg. 48 (1), 3-16 (1999).</li><li>Haas, K., Sin, W. C., Javaherian, A., Li, Z., Cline, H. T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Single-cell+electroporation+for+gene+transfer+in+vivo.">Single-cell electroporation for gene transfer in vivo.</a> Neuron. 29 (3), 583-591 (2001).</li></ol>

Los authros tienen nada que revelar.

El método juxtasomal permite la grabación en vivo de acción potencial enriquecidas a partir de unidades individuales a través de las condiciones de comportamiento (anestesiados, despierto cabeza fija o moviéndose libremente) con la opción de biocitina-etiquetar la neurona registrada en post hoc clasificación de tipos de células y / o reconstrucción 3D. La principal ventaja es obtener parámetros fisiológicos en la configuración extracelular (de este modo no invasivo), sin embargo, ser capaz ...

Redes neuronales consisten en múltiples tipos de células, que se caracterizan por propiedades morfológicas y fisiológicas altamente específicos 1-7. Como consecuencia de ello, los tipos de células individuales realizan tareas especializadas dentro de la red (véase, por ejemplo Gentet et al. 8 y Burgalossi et al. 9). Sólo estamos empezando a entender las funciones específicas del tipo de células a través de redes neuronales y queda aún mucho por descubrir. Para este fin, muchos laboratorios están aplicando enfoques experimentales que permiten el análisis de las propiedades morfológicas de la misma población neuronal de la que se han obtenido parámetros fisiológicos 1,10-15. Aquí, se demuestra la técnica de etiquetado juxtasomal 16,17 que implica registros electrofisiológicos usando pipetas de parche convencionales en la configuración extracelular (de este modo no invasivo) en combinación con la electroporación de la neurona registrada con biocitina. Laimportante ventaja de este enfoque es que la naturaleza no invasiva asegura que el potencial de acción Rematar de las neuronas individuales se registra sin alterar (por ejemplo, diálisis) el contenido intracelular de la célula. Seguido por electroporación, el enfoque juxtasomal proporciona la opción de identificación post hoc celular y la reconstrucción de enlace de función (fisiología) a la estructura (morfología). Típicamente, la reconstrucción morfológica consiste en la reconstrucción de la morfología dendrítica y axonal que se puede extender a la cuantificación de la columna vertebral y / o Bouton densidades o incluso la reconstrucción de la morfología neuronal en resolución nanométrica utilizando microscopía electrónica. La técnica de grabación juxtasomal se puede utilizar para las grabaciones en vivo de varios tipos de células a través de las capas corticales o en áreas subcorticales en una gama de especies, aunque la mayoría de los estudios han aplicado la técnica en pequeños roedores tales como ratones o ratas. Nuestra investigación se centra en el registro y etiquetado de las neuronasde rata corteza somatosensorial primaria (S1) e implica la identificación visual de las neuronas registradas 18, reconstrucciones dendríticas en combinación con un registro preciso en un marco de referencia estandarizada para ingeniería inversa redes corticales 4,19 y reconstrucción detallada de la arquitectura axonal caracterizar específica del tipo celular local y proyección a largo plazo dirigido a 20. En comparación con las técnicas de grabación en vivo alternativas (intracelulares o de células enteras), grabaciones juxtasomal son relativamente estables y por lo tanto pueden aplicarse a distintos estados de comportamiento incluyendo anestesiado 21,22, sedados 14, Pista de los animales-fijo 23, o incluso se mueven libremente despiertos 9 . Aquí, mostramos etiquetado juxtasomal en S1 de una rata anestesiada con uretano, aunque hacemos hincapié en la aplicabilidad general de esta técnica para muchas preparaciones de elección.

<table border="0" cellpadding="0" cellspacing="0" width="784">
	<tbody>
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			<td>National Instruments, Austin, TX, USA</td>
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			<td height="21" style="height:21px;width:224px;">(Labview acq. software)</td>
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			<td height="21" style="height:21px;width:224px;">HEPES</td>
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			<td>H3375</td>
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			<td height="21" style="height:21px;width:224px;">NaCl</td>
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			<td height="21" style="height:21px;width:224px;">KCl</td>
			<td style="width:275px;">Sigma Aldrich</td>
			<td style="width:119px;">60130</td>
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			<td height="21" style="height:21px;width:224px;">CaCl</td>
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			<td height="21" style="height:21px;width:224px;">MgCl2</td>
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</table>

juxtasomal biocytin labeling to study the structure-function relationship of individual cortical neurons

La corteza cerebral se caracteriza por múltiples capas y muchos tipos de células diferentes que juntos como una red son responsables de muchas funciones cognitivas superiores, incluyendo la toma de decisiones, el comportamiento sensorial guiada o la memoria. Para entender cómo estas redes neuronales complejas que de tales tareas, un paso crucial es determinar la función (o la actividad eléctrica) de tipos de células individuales dentro de la red, de preferencia cuando el animal está realizando una tarea cognitiva relevante. Además, es igualmente importante para determinar la estructura anatómica de la red y la arquitectura morfológica de las neuronas individuales para permitir la ingeniería inversa de la red cortical. Avances técnicos disponibles en la actualidad permiten el registro de la actividad celular en despierto, comportándose animales con la opción valiosa de post hoc identificar las neuronas registradas. Aquí, nos demuestran la técnica de etiquetado biocitina juxtasomal, que implica la acción de grabación de potenciómetroal clavar en la configuración extracelular (o floja-patch) utilizando pipetas de parche convencionales. La configuración de la grabación juxtasomal es relativamente estable y aplicable a través de condiciones de comportamiento, incluyendo anestesiado, sedado, despierto cabeza fija, e incluso en el animal se mueve libremente. Por lo tanto, este método permite la vinculación de tipo celular específico acción potencial durante Rematar comportamiento de los animales a la reconstrucción de las neuronas individuales y en última instancia, la totalidad de microcircuito cortical. En este manuscrito de vídeo, se muestra cómo las neuronas individuales en la configuración juxtasomal se pueden marcar con biocitina en ratas anestesiadas con uretano para la identificación post hoc y la reconstrucción morfológica.

El conocimiento detallado de la estructura 3D de las neuronas individuales es crucial para elucidar los principios organizativos de las redes neuronales. Nuestro método consiste en una tubería para lograr etiquetado biocitina alta calidad a partir de una preparación in vivo, lo que permite la clasificación de puestos neuronal hoc y la reconstrucción detallada de la arquitectura dendrítica y axonal de las neuronas individuales en alta resolución. Dependiendo de la calidad de etiquetado j...

Watch this Scientific Journal Video about Juxtasomal Biocytin Labeling to Study the Structure-function Relationship of Individual Cortical Neurons at JoVE.com

Juxtasomal Biocytin Labeling to Study the Structure-function Relationship of Individual Cortical Neurons

Para entender la estructura de las redes neuronales, la caracterización morfológica y funcional de las neuronas individuales es una necesidad. Aquí, nos demuestran etiquetado juxtasomal biocitina, la cual permite que los registros electrofisiológicos en la configuración extracelular, aún mantiene la capacidad de etiquetar de forma intracelular de la neurona para la reconstrucción post hoc de dendríticas y la arquitectura axonal.

Juxtasomal Biocitina Etiquetado para estudiar la estructura-función de la relación de un individuo neuronas corticales

The cerebral cortex is characterized by multiple layers and many distinct cell-types that together as a network are responsible for many higher cognitive ...

juxtasomal-biocytin-labeling-to-study-structure-function-relationship

Research

JoVE Journal

Neuroscience

13.5K vistas.  VU University Amsterdam.  Para entender la estructura de las redes neuronales, la caracterización morfológica y funcional de las neuronas individuales es una necesidad. Aquí, nos demuestran etiquetado juxtasomal biocitina, la cual permite que los registros electrofisiológicos en la configuración extracelular, aún mantiene la capacidad de etiquetar de forma intracelular de la neurona para la reconstrucción post hoc de dendríticas y la arquitectura axonal. 

Video: Juxtasomal Biocytin Labeling to Study the Structure-function Relationship of Individual Cortical Neurons

Laser Capture Microdissection of Drosophila Peripheral Neurons

The dendritic arborization (da) neurons of the Drosophila peripheral nervous system (PNS) provide an excellent model system in which to investigate the molecular mechanisms underlying class-specific dendrite morphogenesis1,2. To facilitate molecular analyses of class-specific da neuron development, it is vital to obtain these cells in a pure population. Although a range of different cell, and tissue-specific RNA isolation techniques exist for Drosophila cells, including magnetic bead based cell purification3,4, Fluorescent Activated Cell Sorting (FACS)5-8, and RNA binding protein based strategies9, none of these methods can be readily utilized for isolating single or multiple class-specific Drosophila da neurons with a high degree of spatial precision. Laser Capture Microdissection (LCM) has emerged as an extremely powerful tool that can be used to isolate specific cell types from tissue sections with a high degree of spatial resolution and accuracy. RNA obtained from isolated cells can then be used for analyses including qRT-PCR and microarray expression profiling within a given cell type10-16. To date, LCM has not been widely applied in the analysis of Drosophila tissues and cells17,18, including da neurons at the third instar larval stage of development. 
Here we present our optimized protocol for isolation of Drosophila da neurons using the infrared (IR) class of LCM. This method allows for the capture of single, class-specific or multiple da neurons with high specificity and spatial resolution. Age-matched third instar larvae expressing a UAS-mCD8::GFP19 transgene under the control of either the class IV da neuron specific ppk-GAL420 driver or the pan-da neuron specific 21-7-GAL421 driver were used for these experiments. RNA obtained from the isolated da neurons is of very high quality and can be directly used for downstream applications, including qRT-PCR or microarray analyses. Furthermore, this LCM protocol can be readily adapted to capture other Drosophila cell types a various stages of development dependent upon the cell type specific, GAL4-driven expression pattern of GFP.

Laser Capture Microdissection of Drosophila Peripheral Neurons

In this video-article we present a method for isolating single or multiple Drosophila da neurons from third instar larvae using the infrared capture (IR) class of Laser Capture Microdissection (LCM). RNA obtained from the isolated neurons can be readily used for downstream applications including qRT-PCR or microarray analyses.

Láser microdisección de captura de Drosophila Neuronas periféricas

The dendritic arborization (da) neurons of the Drosophila peripheral nervous system (PNS) provide an excellent model system in which to investigate the ...

Investigación

Neurociencias

Bilaminar Co-culture of Primary Rat Cortical Neurons and Glia

This video will guide you through the process of culturing rat cortical neurons in the presence of a glial feeder layer, a system known as a bilaminar or co-culture model. This system is suitable for a variety of experimental needs requiring either a glass or plastic growth substrate and can also be used for culture of other types of neurons.
Rat cortical neurons obtained from the late embryonic stage (E17) are plated on glass coverslips or tissue culture dishes facing a feeder layer of glia grown on dishes or plastic coverslips (known as Thermanox), respectively. The choice between the two configurations depends on the specific experimental technique used, which may require, or not, that neurons are grown on glass (e.g. calcium imaging versus Western blot). The glial feeder layer, an astroglia-enriched secondary culture of mixed glia, is separately prepared from the cortices of newborn rat pups (P2-4) prior to the neuronal dissection.
A major advantage of this culture system as compared to a culture of neurons only is the support of neuronal growth, survival, and differentiation provided by trophic factors secreted from the glial feeder layer, which more accurately resembles the brain environment in vivo. Furthermore, the co-culture can be used to study neuronal-glial interactions1.
At the same time, glia contamination in the neuronal layer is prevented by different means (low density culture, addition of mitotic inhibitors, lack of serum and use of optimized culture medium) leading to a virtually pure neuronal layer, comparable to other established methods1-3. Neurons can be easily separated from the glial layer at any time during culture and used for different experimental applications ranging from electrophysiology4, cellular and molecular biology5-8, biochemistry5, imaging and microscopy4,6,7,9,10. The primary neurons extend axons and dendrites to form functional synapses11, a process which is not observed in neuronal cell lines, although some cell lines do extend processes.
A detailed protocol of culturing rat hippocampal neurons using this co-culture system has been described previously4,12,13. Here we detail a modified protocol suited for cortical neurons. As approximately 20x106 cells are recovered from each rat embryo, this method is particularly useful for experiments requiring large numbers of neurons (but not concerned about a highly homogenous neuronal population). The preparation of neurons and glia needs to be planned in a time-specific manner. We will provide the step-by-step protocol for culturing rat cortical neurons as well as culturing glial cells to support the neurons.

Here we provide a protocol for culturing rat cortical neurons in the presence of a glial feeder layer. The cultured neurons establish polarity and create synapses, and can be separated from the glia for use in various applications, such as electrophysiology, calcium imaging, cell survival assays, immunocytochemistry, and RNA/DNA/protein isolation.

Bilaminar Co-cultivo de neuronas de rata primaria cortical y la glía

This video will guide you through the process of culturing rat cortical neurons in the presence of a glial feeder layer, a system known as a bilaminar or ...

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

Here we present a technique to label the trajectories of small groups of DRG neurons into the embryonic spinal cord by diffusive staining using the lipophilic tracer 1,1'-dioctadecyl-3,3,3',3'-tetramethylindocarbocyanine perchlorate (DiI)1. The comparison of axonal pathways of wild-type with those of mouse lines in which genes are mutated allows testing for a functional role of candidate proteins in the control of axonal branching which is an essential mechanism in the wiring of the nervous system. Axonal branching enables an individual neuron to connect with multiple targets, thereby providing the physical basis for the parallel processing of information. Ramifications at intermediate target regions of axonal growth may be distinguished from terminal arborization. Furthermore, different modes of axonal branch formation may be classified depending on whether branching results from the activities of the growth cone (splitting or delayed branching) or from the budding of collaterals from the axon shaft in a process called interstitial branching2 (Fig. 1).
The central projections of neurons from the DRG offer a useful experimental system to study both types of axonal branching: when their afferent axons reach the dorsal root entry zone (DREZ) of the spinal cord between embryonic days 10 to 13 (E10 - E13) they display a stereotyped pattern of T- or Y-shaped bifurcation. The two resulting daughter axons then proceed in rostral or caudal directions, respectively, at the dorsolateral margin of the cord and only after a waiting period collaterals sprout from these stem axons to penetrate the gray matter (interstitial branching) and project to relay neurons in specific laminae of the spinal cord where they further arborize (terminal branching)3. DiI tracings have revealed growth cones at the dorsal root entry zone of the spinal cord that appeared to be in the process of splitting suggesting that bifurcation is caused by splitting of the growth cone itself4 (Fig. 2), however, other options have been discussed as well5.
This video demonstrates first how to dissect the spinal cord of E12.5 mice leaving the DRG attached. Following fixation of the specimen tiny amounts of DiI are applied to DRG using glass needles pulled from capillary tubes. After an incubation step, the labeled spinal cord is mounted as an inverted open-book preparation to analyze individual axons using fluorescence microscopy.

The stereotyped projections of sensory afferents into the rodent spinal cord offer an easily accessible experimental system to study axonal branching through the tracing of single axons.

DII-etiquetado de las neuronas DRG para el Estudio de ramificación axonal en una preparación Todo el montaje del cable del ratón embrionario espinal

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

In Vivo Two-photon Imaging Of Experience-dependent Molecular Changes In Cortical Neurons

The brain's ability to change in response to experience is essential for healthy brain function, and abnormalities in this process contribute to a variety of brain disorders1,2. To better understand the mechanisms by which brain circuits react to an animal's experience requires the ability to monitor the experience-dependent molecular changes in a given set of neurons, over a prolonged period of time, in the live animal. While experience and associated neural activity is known to trigger gene expression changes in neurons1,2, most of the methods to detect such changes do not allow repeated observation of the same neurons over multiple days or do not have sufficient resolution to observe individual neurons3,4. Here, we describe a method that combines in vivo two-photon microscopy with a genetically encoded fluorescent reporter to track experience-dependent gene expression changes in individual cortical neurons over the course of day-to-day experience. 
One of the well-established experience-dependent genes is Activity-regulated cytoskeletal associated protein (Arc)5,6. The transcription of Arc is rapidly and highly induced by intensified neuronal activity3, and its protein product regulates the endocytosis of glutamate receptors and long-term synaptic plasticity7. The expression of Arc has been widely used as a molecular marker to map neuronal circuits involved in specific behaviors3. In most of those studies, Arc expression was detected by in situ hybridization or immunohistochemistry in fixed brain sections. Although those methods revealed that the expression of Arc was localized to a subset of excitatory neurons after behavioral experience, how the cellular patterns of Arc expression might change with multiple episodes of repeated or distinctive experiences over days was not investigated. 
In vivo two-photon microscopy offers a powerful way to examine experience-dependent cellular changes in the living brain8,9. To enable the examination of Arc expression in live neurons by two-photon microscopy, we previously generated a knock-in mouse line in which a GFP reporter is placed under the control of the endogenous Arc promoter10. This protocol describes the surgical preparations and imaging procedures for tracking experience-dependent Arc-GFP expression patterns in neuronal ensembles in the live animal. In this method, chronic cranial windows were first implanted in Arc-GFP mice over the cortical regions of interest. Those animals were then repeatedly imaged by two-photon microscopy after desired behavioral paradigms over the course of several days. This method may be generally applicable to animals carrying other fluorescent reporters of experience-dependent molecular changes4.

In Vivo Two-photon Imaging Of Experience-dependent Molecular Changes In Cortical Neurons

Experience-dependent molecular changes in neurons are essential for the brain's ability to adapt in response to behavioral challenges. An in vivo two-photon imaging method is described here that allows the tracking of such molecular changes in individual cortical neurons through genetically encoded reporters.

In vivo de dos fotones de imágenes de dependientes Experience-Los cambios moleculares en las neuronas corticales

The brain's ability to change in response to experience is essential for healthy brain function, and abnormalities in this process contribute to a variety ...

A Method for High Fidelity Optogenetic Control of Individual Pyramidal Neurons In vivo

Optogenetic methods have emerged as a powerful tool for elucidating neural circuit activity underlying a diverse set of behaviors across a broad range of species. Optogenetic tools of microbial origin consist of light-sensitive membrane proteins that are able to activate (e.g., channelrhodopsin-2, ChR2) or silence (e.g., halorhodopsin, NpHR) neural activity ingenetically-defined cell types over behaviorally-relevant timescales. We first demonstrate a simple approach for adeno-associated virus-mediated delivery of ChR2 and NpHR transgenes to the dorsal subiculum and prelimbic region of the prefrontal cortex in rat. Because ChR2 and NpHR are genetically targetable, we describe the use of this technology to control the electrical activity of specific populations of neurons (i.e., pyramidal neurons) embedded in heterogeneous tissue with high temporal precision. We describe herein the hardware, custom software user interface, and procedures that allow for simultaneous light delivery and electrical recording from transduced pyramidal neurons in an anesthetized in vivo preparation. These light-responsive tools provide the opportunity for identifying the causal contributions of different cell types to information processing and behavior.

A Method for High Fidelity Optogenetic Control of Individual Pyramidal Neurons In vivo

The recent development of neuroscience tools that combine genetics and optics, termed "optogenetics", enables control over neural circuit activity with an unprecedented level of spatial and temporal resolution. Here we provide a protocol for integrating in vivo recording with optogenetic manipulation of genetically-defined subsets of prefrontal cortical and subicular pyramidal neurons.

Un método para la alta fidelidad Optogenética control de un individuo neuronas piramidales<em&gt; En vivo</em

Optogenetic methods  have emerged as a powerful tool for elucidating neural circuit activity  underlying a diverse set of behaviors across a broad range ...

Genetic Manipulation of Cerebellar Granule Neurons In Vitro and In Vivo to Study Neuronal Morphology and Migration

Developmental events in the brain including neuronal morphogenesis and migration are highly orchestrated processes. In vitro and in vivo analyses allow for an in-depth characterization to identify pathways involved in these events. Cerebellar granule neurons (CGNs) that are derived from the developing cerebellum are an ideal model system that allows for morphological analyses. Here, we describe a method of how to genetically manipulate CGNs and how to study axono- and dendritogenesis of individual neurons. With this method the effects of RNA interference, overexpression or small molecules can be compared to control neurons. In addition, the rodent cerebellar cortex is an easily accessible in vivo system owing to its predominant postnatal development. We also present an in vivo electroporation technique to genetically manipulate the developing cerebella and describe subsequent cerebellar analyses to assess neuronal morphology and migration.

Genetic Manipulation of Cerebellar Granule Neurons In Vitro and In Vivo to Study Neuronal Morphology and Migration

Neuronal morphogenesis and migration are crucial events underlying proper brain development. Here, we describe methods to genetically manipulate cultured cerebellar granule neurons and the developing cerebellum for the assessment of morphology and migratory characteristics of neurons.

La manipulación genética de las neuronas cerebelosa gránulo<em&gt; In Vitro</em&gt; Y<em&gt; En Vivo</em&gt; Estudiar Neuronal Morfología y Migración

Developmental events in the brain including neuronal morphogenesis and migration are highly orchestrated processes. In vitro and in vivo analyses allow ...

Differential Labeling of Cell-surface and Internalized Proteins after Antibody Feeding of Live Cultured Neurons

In order to demonstrate the cell-surface localization of a putative transmembrane receptor in cultured neurons, we labeled the protein on the surface of live neurons with a specific primary antibody raised against an extracellular portion of the protein. Given that receptors are trafficked to and from the surface, if cells are permeabilized after fixation then both cell-surface and internal protein will be detected by the same labeled secondary antibody. Here, we adapted a method used to study protein trafficking (&#8220;antibody feeding&#8221;) to differentially label protein that had been internalized by endocytosis during the antibody incubation step and protein that either remained on the cell surface or was trafficked to the surface during this period. The ability to distinguish these two pools of protein was made possible through the incorporation of an overnight blocking step with highly-concentrated unlabeled secondary antibody after an initial incubation of unpermeabilized neurons with a fluorescently-labeled secondary antibody. After the blocking step, permeabilization of the neurons allowed detection of the internalized pool with a fluorescent secondary antibody labeled with a different fluorophore. Using this technique we were able to obtain important information about the subcellular location of this putative receptor, revealing that it was, indeed, trafficked to the cell-surface in neurons. This technique is broadly applicable to a range of cell types and cell-surface proteins, providing a suitable antibody to an extracellular epitope is available.

We describe a method to label protein on the surface of living neurons using a specific polyclonal antibody to extracellular epitopes. Protein bound by the antibody on the cell surface and subsequently internalized via endocytosis can be distinguished from protein remaining on, or trafficked to, the surface during the incubation.

Etiquetado diferencial de la superficie celular e internalizado Proteínas después Antibody alimentación de vivo cultivadas neuronas

In order to demonstrate the cell-surface localization of a putative transmembrane receptor in cultured neurons, we labeled the protein on the surface of ...

Laser Capture Microdissection - A Demonstration of the Isolation of Individual Dopamine Neurons and the Entire Ventral Tegmental Area

Laser capture microdissection (LCM) is used to isolate a concentrated population of individual cells or precise anatomical regions of tissue from tissue sections on a microscope slide. When combined with immunohistochemistry, LCM can be used to isolate individual cells types based on a specific protein marker. Here, the LCM technique is described for collecting a specific population of dopamine neurons directly labeled with&#160;tyrosine hydroxylase immunohistochemistry and for isolation of the dopamine neuron containing region of the ventral tegmental area using indirect tyrosine hydroxylase immunohistochemistry on a section adjacent to those used for LCM. An infrared (IR) capture laser is used to both dissect individual neurons as well as the ventral tegmental area off glass slides and onto an LCM cap for analysis. Complete dehydration of the tissue with 100% ethanol and xylene is critical. The combination of the IR capture laser and the ultraviolet (UV) cutting laser is used to isolate individual dopamine neurons or the ventral tegmental area when using PEN membrane slides. A PEN membrane slide has significant advantages over a glass slide as it offers better consistency in capturing and collecting cells, is faster collecting large pieces of tissue, is less reliant on dehydration and results in complete removal of the tissue from the slide. Although removal of large areas of tissue from a glass slide is feasible, it is considerably more time consuming and frequently leaves some residual tissue behind. Data shown here demonstrate that RNA of sufficient quantity and quality can be obtained using these procedures for quantitative PCR measurements. Although RNA and DNA are the most commonly isolated molecules from tissue and cells collected with LCM, isolation and measurement of microRNA, protein and epigenetic changes in DNA can also benefit from the enhanced anatomical and cellular resolution obtained using LCM.

The isolation of individual dopamine neurons or the ventral tegmental area with direct or indirect immunohistochemistry is demonstrated using laser capture microdissection. Parameters for isolation of tissue from a glass slide using an infrared laser and from membrane slides using the combination of an infrared and ultraviolet laser are discussed.

Microdisección de captura por láser - Una demostración del aislamiento individual de dopamina neuronas y el Área ventral tegmental completo

Laser capture microdissection (LCM) is used to isolate a concentrated population of individual cells or precise anatomical regions of tissue from tissue ...

Immunohistochemistry and Multiple Labeling with Antibodies from the Same Host Species to Study Adult Hippocampal Neurogenesis

Adult neurogenesis is a highly regulated, multi-stage process in which new neurons are generated from an activated neural stem cell via increasingly committed intermediate progenitor subtypes. Each of these subtypes expresses a set of specific molecular markers that, together with specific morphological criteria, can be used for their identification. Typically, immunofluorescent techniques are applied involving subtype-specific antibodies in combination with exo- or endogenous proliferation markers. We herein describe immunolabeling methods for the detection and quantification of all stages of adult hippocampal neurogenesis. These comprise the application of thymidine analogs, transcardial perfusion, tissue processing, heat-induced epitope retrieval, ABC immunohistochemistry, multiple indirect immunofluorescence, confocal microscopy and cell quantification. Furthermore we present&#160;a sequential multiple immunofluorescence protocol which circumvents problems usually arising from the need of using primary antibodies raised in the same host species. It allows an accurate identification of all hippocampal progenitor subtypes together with a proliferation marker within a single section. These techniques are a powerful tool to study the regulation of different progenitor subtypes in parallel, their involvement in brain pathologies and their role in specific brain functions.

This video article illustrates a comprehensive protocol to detect and quantify all stages of adult hippocampal neurogenesis within the same tissue section. We elaborated a method to overcome the limitations of indirect multiple immunofluorescence that arise when suitable antibodies from different host species are unavailable.

La inmunohistoquímica y Etiquetado múltiple con anticuerpos de la misma especie anfitrión para adultos estudio del hipocampo neurogénesis

Adult neurogenesis is a highly regulated, multi-stage process in which new neurons are generated from an activated neural stem cell via increasingly ...

Quantification of Filamentous Actin (F-actin) Puncta in Rat Cortical Neurons

Filamentous actin protein (F-actin) plays a major role in spinogenesis, synaptic plasticity, and synaptic stability. Changes in dendritic F-actin rich structures suggest alterations in synaptic integrity and connectivity. Here we provide a detailed protocol for culturing primary rat cortical neurons, Phalloidin staining for F-actin puncta, and subsequent quantification techniques. First, the frontal cortex of E18 rat embryos are dissociated into low-density cell culture, then the neurons grown in vitro for at least 12-14 days. Following experimental treatment, the cortical neurons are stained with AlexaFluor 488 Phalloidin (to label the dendritic F-actin puncta) and microtubule-associated protein 2 (MAP2; to validate the neuronal cells and dendritic integrity). Finally, specialized software is used to analyze and quantify randomly selected neuronal dendrites. F-actin rich structures are identified on second order dendritic branches (length range 25-75 &#181;m) with continuous MAP2 immunofluorescence. The protocol presented here will be a useful method for investigating changes in dendritic synapse structures subsequent to experimental treatments.

Quantification of Filamentous Actin F-actin Puncta in Rat Cortical Neurons

Filamentous actin (F-actin) plays an important role in spinogenesis, synaptic plasticity, and synaptic stability. Quantification of F-actin puncta is therefore a useful tool to study the integrity of synaptic structures. This protocol describes the procedures of quantifying F-actin puncta labeled with Phalloidin in low-density primary cortical neuronal cultures.

La cuantificación de la actina filamentosa (F-actina) puntos lagrimales en las neuronas corticales de rata

Filamentous actin protein (F-actin) plays a major role in spinogenesis, synaptic plasticity, and synaptic stability. Changes in dendritic F-actin rich ...