Este trabajo fue financiado por los Institutos Nacionales de Salud (K22NS050137 a JCB).

1. De cultivo celular <ol><li> Las neuronas corticales son aisladas y cultivadas con técnicas descritas anteriormente y se cultivaron en placas de cultivo con fondo de cristal (MatTek Corporation, Ashland, MA), recubierta con poli-D-lisina y laminina 1. </li></ol> 2. Preparación de las soluciones madre de los fluorescentes TMRM sondas y H 2 DCF-DA <ol><li> Preparar una solución de 10 mM de TMRM disolviendo 5,0 mg de TMRM en 1 ml de dimetilsulfóxido anhidro. Vortex durante 1 min. Entonces, hacer alícuotas y almacenarlos a -20 ° C, protegido de la luz, y su uso dentro de un mes. </li><li> A continuación, preparar una solución de 10 mM de H 2 DCF-DA mediante la disolución de 4,87 mg de H 2 DCF-DA en 1 ml de DMSO anhidro. Del mismo modo, vortex durante 1 min. Entonces, hacer alícuotas y almacenarlos a -20 ° C, protegido de la luz, y su uso dentro de una semana. </li></ol> 3. Cargando corticales de rata neuronas con TMRM y H 2 DCF-DA TMRM es un potenciómetro, penetran en las células indicador fluorescente que se acumula en el interior de alta carga negativa de las mitocondrias. Es importante la utilización de las bajas concentraciones (10-50 nm de rango) de TMRM para evitar la auto-extinción de TMRM mitocondrial. Luego, la señal de fluorescencia de TMRM pueden ser directamente relacionados con la co-ΔΨm través de la membrana mitocondrial interna. Una pérdida de ΔΨm causas TMRM de fuga de la mitocondria que resulta en una pérdida de intensidad de fluorescencia. H 2 DCF-DA se penetra en las células de la sonda convierte en DCF-DA por las esterasas intracelulares, y sus resultados en la oxidación de DCF fluorescente. La concentración final de H 2 DCF-DA oscila entre 2.10 M y que debe ser probado empíricamente en neuronas derivadas de las diferentes regiones del cerebro ya que las altas concentraciones de carga podría dar lugar a la saturación de fluorescencia de DCF, incluso en ausencia de H 2 O 2. La presencia de oxidantes endógenos o exógenos (por ejemplo, el óxido nítrico peróxido de hidrógeno) se incrementará la intensidad de fluorescencia DFC. A continuación, ofrecemos un protocolo para la carga de las neuronas corticales de rata con TMRM y H 2 DCF-DA. <ol><li> Para cargar las neuronas corticales de rata con TMRM, en primer lugar, lavar los cultivos de neuronas 3 veces con tampón de Tyrode (superposición de texto: TB: 145 mM NaCl, 5 mM KCl, 10 mM de glucosa, 1,5 mM CaCl 2, 1 mM MgCl 2, y 10 mM HEPES, ajustar el pH a 7,4 con NaOH). A continuación, preparar 20 nM de TMRM diluyendo el stock de 10 mM TMRM 1 / 1000 veces la tuberculosis y después agregue 2 l de TMRM diluidas por 1 ml de la tuberculosis. Incubar las neuronas con TMRM durante 45 minutos en la oscuridad a temperatura ambiente. Después de 45 minutos, coloque la placa de cultivo en la platina del microscopio y empezar de imágenes. </li><li> Para cargar las neuronas corticales de rata con H 2 DCF-DA, lavar los cultivos de neuronas 3 veces con la tuberculosis. A continuación, preparar 2 M de H 2 DCF-DA mediante la dilución de los 10 mm de H 2 DCF-DA stock 1 / 10 veces en la tuberculosis y después agregue 2 l de diluirse H 2 DCF-DA por 1 ml de TB. A continuación, se incuban las neuronas con H 2 DCF-DA durante 45 minutos en oscuridad a temperatura ambiente. Después de 45 minutos, lavar las neuronas 4 veces con TB para eliminar el exceso de indicador fluorescente antes de obtener las imágenes. </li></ol> 4. Imágenes en directo de las neuronas se incubaron con TMRM para determinar ΔΨm <ol><li> Para realizar las imágenes en directo de las neuronas se incubaron con TMRM, la microscopía de escaneo láser confocal (superposición de texto: LSM 510, Carl Zeiss, Inc.), con la aplicación de vivir de series de tiempo del programa, se utiliza. Aplicar de baja resolución y atenuado la potencia del láser (superposición de texto: baja resolución: 256 x 256, la potencia del láser: 1%) para reducir al mínimo el tiempo necesario para obtener imágenes y para evitar photobleaching. </li><li> . A continuación, ajustar el enfoque de las neuronas montado cargado con TMRM con la luz reflejada. Examine la fluorescencia TMRM por la iluminación a 514 nm y la detección a 570 nm. Ajustar la ganancia de la detección de una cámara debajo del nivel de saturación. </li><li> Una vez que todos los parámetros que incluyen la resolución, la potencia del láser, el aumento de la detección de una cámara, y el intervalo de tiempo transcurrido para obtener las imágenes se establecen, no cambian estos valores entre los experimentos. A continuación, cambie el campo. Comenzar a recoger las imágenes. </li><li> Para probar los cambios en ΔΨm, estímulos, tales como una M de FCCP o 2 mg / ml de oligomicina, se puede aplicar, lo cual permitirá despolarización o hiperpolarización del potencial de membrana mitocondrial, respectivamente. Estos cambios se reflejarán en una disminución en la intensidad de fluorescencia TMRM en comparación con la intensidad de fluorescencia de referencia en el caso del FCCP, o un aumento en la intensidad de fluorescencia TMRM en el caso de oligomicina. </li></ol> 5. Imágenes en vivo de neuronas incubadas con H 2 DCF-DA para determinar ROS <ol><li> Para realizar las imágenes en directo de las neuronas incubadas con H 2 DCF-DA, en primer lugar, montar la placa de cultivo en el escenario de un microscopio. Ajustar el enfoque de las células que nosción refleja la luz. Examine DCF de fluorescencia de excitación a 488 nm y emisión a 515 nm. </li><li> A continuación, ajustar la potencia del láser a un 5-7%, la ganancia del detector, y resolución de 256 x 256. No cambie estos ajustes entre los experimentos. A continuación, establezca la frecuencia para la obtención de imágenes en directo mediante el programa de series de tiempo. </li><li> Seleccione un nuevo campo y comenzar a adquirir las imágenes. Para detectar cambios en los niveles de ROS, tratar las células con 100 a 200 M de H 2 O 2. Esto se refleja en un aumento en la intensidad de fluorescencia de DCF en comparación con el nivel de referencia. </li></ol> 6. Análisis de los datos <ol><li> Use la región de interés (Superposición de texto: ROI) de herramientas del programa de LSM para seleccionar las áreas. A continuación, medir la intensidad de ROS TMRM o fluorescencia. Seleccione el ROI de las regiones mitocondriales o regiones de interés desde el cuerpo celular en las células de todo imágenes para medir la intensidad de fluorescencia de TMRM o ROS, respectivamente. </li><li> Calcular la intensidad media de fluorescencia de todas las regiones de interés de cada célula de TMRM o de los cuerpos de célula entera para todas las células de imágenes de ROS para cada momento. Seleccione las regiones próximas a las células para calcular la intensidad de fluorescencia de fondo. Tomar varias mediciones y calcular la intensidad media de fondo. </li><li> Restar la intensidad de fluorescencia de fondo promedio de intensidad de fluorescencia media de rendimiento de la inversión en cada celda para cada momento el uso de Microsoft Excel. Después de restar la intensidad de fondo, normalizar la intensidad de fluorescencia de DCF o TMRM a la fluorescencia basal mediante esta fórmula (superposición de texto: Df = FF o / F o x 100, donde F = intensidad de la fluorescencia en cualquier punto del tiempo, para = línea de base de fluorescencia). A continuación, utilice el programa Sigma Plot para generar el gráfico que muestra los cambios en la intensidad de la fluorescencia con el tiempo. </li></ol> 7. Resultados representante La figura 1A muestra una imagen de fluorescencia de las neuronas corticales de rata incubados con TMRM. Además del FCCP, un desacoplador mitocondrial, conduce a la despolarización mitocondrial y una pérdida de intensidad de fluorescencia TMRM (Fig. 1B). El nivel básico TMRM fluorescencia se mantiene estable antes de la adición de FCCP (los primeros 350 segundos;. Fig. 1C). El análisis cuantitativo de TMRM cambios de fluorescencia en el tiempo muestra una disminución significativa de la fluorescencia TMRM tras la adición de FCCP (Fig. 1C). Figura 1D muestra la imagen de fluorescencia de las neuronas corticales de rata cargado con DCF. Además de H 2 O 2 da como resultado una mayor intensidad de fluorescencia de DCF en los cuerpos celulares (Fig. 1E). La línea de base DCF nivel de fluorescencia no se ha modificado (los primeros 120 segundos) antes de la aplicación de H 2 O 2. Lapso de tiempo-las medidas de fluorescencia de DCF muestran sus niveles estables, lo que aumenta después de H 2 O 2 tratamiento (Fig. 1F). <img src="/files/ftp_upload/2704/2704fig1.jpg" alt="Figura 1"/> Figura 1. Evaluación del potencial de membrana mitocondrial y los niveles de ROS en las neuronas corticales de rata viva. (A) Imagen Representante de fluorescencia de las neuronas corticales cargado con TMRM. Después de escanear la línea de base TMRM fluorescencia, las neuronas fueron tratadas con el FCCP protonophore (1 M). A la derecha hay una barra de intensidad de la fluorescencia pseudocolor TMRM con un máximo de brillantes que representan a amarillo y negro y la intensidad mínima, respectivamente. La pérdida de fluorescencia TMRM de las regiones mitocondrial indica el colapso de ΔΨm a FCCP tratamiento (grupo B). La representación cuantitativa de los cambios en la intensidad de fluorescencia TMRM en diferentes momentos antes y después del tratamiento FCCP se muestra en el panel C. (D) Imagen de fluorescencia de las neuronas corticales de rata cargado con H2DCF-DA. Después de determinar la base de DCF de fluorescencia, las células fueron tratadas con 200 mM H 2 O 2, y el cambio en la fluorescencia de DCF se evaluó. Un aumento en la fluorescencia de DCF refleja el aumento en los niveles de ROS en H 2 O 2 tratamiento (E). El análisis cuantitativo de los cambios en la fluorescencia de DCF, antes y después de H 2 O 2 tratamiento, se muestra en el panel de F. barra de escala = 10 m Video.7.1 - labmedia 2704_Joshi.avi En vivo de imágenes de células en las neuronas corticales TMRM antes y después de la adición FCCP con objetivo de 40X. La intensidad de pseudocolor muestra un máximo (de color amarillo brillante, antes de añadir FCCP) y la disminución (de color rojo, después de la adición FCCP) la intensidad de fluorescencia después de la adición TMRM FCCP. <a href="http://www.jove.com/files/ftp_upload/2704/supplemental_video1.avi">Haga clic aquí para ver el video</a> Vídeo. 7.5 - labmedia 2704_Joshi.avi En vivo de imágenes de células de DCF en las neuronas corticales antes y después de H 2 O 2 además con objetivo de 40X. La línea de base DCF fluorescencia de color verde claro en los cuerpos celulares y H2O2 Además aumenta la DCF intensidad de la fluorescencia de color verde brillante. <a href="http://www.jove.com/files/ftp_upload/2704/supplemental_video2.avi">Haga clic aquí para ver el video</a>

<ol>
	<li>Hilgenberg, L. G., Smith, M. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Preparation+of+dissociated+mouse+cortical+neuron+cultures.">Preparation of dissociated mouse cortical neuron cultures.</a> J. Vis. Exp. , (2007).</li><li>Scaduto, R. C., Grotyohann, L. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Measurement+of+mitochondrial+membrane+potential+using+fluorescent+rhodamine+derivatives.">Measurement of mitochondrial membrane potential using fluorescent rhodamine derivatives.</a> Biophys. J. 76, 469-477 (1999).</li><li>Ward, M. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+amyloid+precursor+protein+intracellular+domain+(AICD)+disrupts+actin+dynamics+and+mitochondrial+bioenergetics.">The amyloid precursor protein intracellular domain (AICD) disrupts actin dynamics and mitochondrial bioenergetics.</a> J. Neurochem. 113, 275-284 (2010).</li><li>Gunasekar, P. G., Kanthasamy, A. G., Borowitz, J. L., Isom, G. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=NMDA+receptor+activation+produces+concurrent+generation+of+nitric+oxide+and+reactive+oxygen+species:+implication+for+cell+death.">NMDA receptor activation produces concurrent generation of nitric oxide and reactive oxygen species: implication for cell death.</a> J. Neurochem. 65, 2016-2021 (1995).</li></ol>

Hemos presentado un procedimiento paso a paso describe cómo determinar ΔΨm y ROS en ratas las neuronas corticales con el fluorescente TMRM indicadores y H 2 DCF-DA, respectivamente. Para otros tipos de células, es importante para determinar empíricamente la concentración final y el tiempo de carga de TMRM o H 2 DCF-DA. En general, la gama de concentraciones de 20 a 200 nM TMRM, y el tiempo de incubación de células con TMRM varía de 20 a 60 min. La concentración final de H 2 DCF-DA varía desde 2 hasta 10 M, y la incubación de las células en una solución de carga que contiene este indicador varía entre 30-45 minutos. Es importante para optimizar la potencia del láser y la velocidad de escaneo de tomar las imágenes para evitar tanto la foto-toxicidad para las células y cambios en la intensidad de fluorescencia (por ejemplo, el parpadeo de TMRM fluorescencia) en ausencia de cualquier estímulo. Los ajustes ópticos optimizados debería resultar en una señal de fluorescencia que no es mayor o menor saturación (umbral) en ausencia de estímulo. Las condiciones óptimas para recoger las imágenes de un campo seleccionado en una potencia del láser en particular y una velocidad de escaneado se logra cuando no hay cambios en la intensidad de la fluorescencia de la sonda en la ausencia de cualquier estímulo durante 10-15 minutos de imágenes en vivo. Otras sondas de fluorescencia para determinar ΔΨm incluyen rodamina 123 y tetra metil éster etílico rodamina (TMRE). Sin embargo, se encontró que inhibe los procesos respiratorios en dos mitocondrias aisladas. Es importante destacar que TMRM no tiene ningún efecto sobre la respiración mitocondrial en bajas concentraciones y tiene dos bajas y fototoxicidad photobleaching 3 en comparación con otras sondas. H 2 DCF-DA es un buen indicador de ROS como es bien retenido en las células y reconoce varias especies oxidantes, como peróxidos, óxidos de super, y el óxido nítrico 4.

<table border="1"><tbody><tr><th> Nombre del reactivo </th><th> Empresa </th><th> Número de catálogo </th></tr><tr><td> Con fondo de cristal cultura plato </td><td> MatTek Corporación </td><td> P35G-1.5-14-C </td></tr><tr><td> NbActive4 </td><td> BrainBits </td><td> NbActive4 </td></tr><tr><td> TMRM </td><td> Invitrogen </td><td> T668 </td></tr><tr><td> H 2 DCF-DA </td><td> Invitrogen </td><td> C400 </td></tr><tr><td> NaCl </td><td> Sigma </td><td> S6191 </td></tr><tr><td> KCl </td><td> Sigma </td><td> P3911 </td></tr><tr><td> CaCl2 • 2H 2 O </td><td> Sigma </td><td> C3306 </td></tr><tr><td> MgCl2 • 6H 2 O </td><td> Sigma </td><td> M2670 </td></tr><tr><td> D-glucosa </td><td> Sigma </td><td> G6152 </td></tr><tr><td> HEPES </td><td> Invitrogen </td><td> 15630 </td></tr></tbody></table>

determination of mitochondrial membrane potential and reactive oxygen species in live rat cortical neurons

Potencial de membrana mitocondrial (ΔΨm) es fundamental para mantener la función fisiológica de la cadena respiratoria para generar ATP. Una pérdida significativa de ΔΨm hace agotadas las células de la energía con la posterior muerte. Las especies reactivas del oxígeno (ROS) son importantes moléculas de señalización, pero su acumulación en condiciones patológicas conduce a estrés oxidativo. Las dos principales fuentes de ROS en las células son las toxinas ambientales y el proceso de fosforilación oxidativa. Disfunción mitocondrial y de estrés oxidativo han sido implicados en la fisiopatología de muchas enfermedades, por lo tanto, la capacidad de determinar ΔΨm y ROS pueden proporcionar pistas importantes sobre el estado fisiológico de la célula y la función de la mitocondria. Varias sondas fluorescentes (Rodamina 123, TMRM, TMRE, JC-1) se puede utilizar para determinar Δψm en una variedad de tipos celulares, y muchos indicadores de fluorescencia (dihydroethidium, dihidrorodamina 123, H 2 DCF-DA) se puede utilizar para determinar ROS . Casi todas las sondas de fluorescencia que se utilizan para evaluar ΔΨm o ROS son de una sola longitud de onda de los indicadores, que aumentan o disminuyen su intensidad de fluorescencia proporcional a un estímulo que aumenta o disminuye los niveles de ΔΨm o ROS. Por lo tanto, es imprescindible para medir la intensidad de la fluorescencia de las sondas a nivel basal y después de la aplicación de un estímulo específico. Esto permite determinar el porcentaje de cambio en la intensidad de fluorescencia entre el nivel de referencia y un estímulo. Este cambio en la intensidad de la fluorescencia refleja el cambio en los niveles relativos de ΔΨm o ROS. En este video, que muestra cómo aplicar el indicador de fluorescencia, TMRM, en ratas las neuronas corticales para determinar el porcentaje de cambio en la intensidad de fluorescencia TMRM entre el nivel basal y después de aplicar FCCP, un desacoplador mitocondrial. Los niveles más bajos de TMRM fluorescencia que resulta del tratamiento FCCP reflejan la despolarización del potencial de membrana mitocondrial. También se muestra cómo aplicar la fluorescencia de la sonda H 2 DCF-DA para evaluar el nivel de ROS en las neuronas corticales, en primer lugar al inicio del estudio y después de la aplicación de H 2 O 2. Este protocolo (con modificaciones menores) también se puede utilizar para determinar los cambios en ΔΨm y ROS en diferentes tipos celulares y en neuronas aisladas de otras regiones del cerebro.

Watch this Scientific Journal Video about Determination of Mitochondrial Membrane Potential and Reactive Oxygen Species in Live Rat Cortical Neurons at JoVE.com

Determination of Mitochondrial Membrane Potential and Reactive Oxygen Species in Live Rat Cortical Neurons

Se demuestra la aplicación del indicador de fluorescencia, TMRM, en las neuronas corticales para determinar los cambios relativos en la intensidad de fluorescencia TMRM antes y después de la aplicación de un estímulo específico. También muestran la aplicación de la fluorescencia de la sonda H<sub&gt; 2</sub&gt; DCF-DA para evaluar el nivel relativo de especies reactivas del oxígeno en las neuronas corticales.

Determinación de Especies de la membrana mitocondrial de oxígeno potencial y reactiva en las neuronas corticales de rata en vivo

Mitochondrial membrane potential (&Delta;&Psi;m) is critical for maintaining the physiological function of the respiratory chain to generate ATP.  A ...

determination-mitochondrial-membrane-potential-reactive-oxygen

Research

JoVE Journal

Neuroscience

64.7K vistas.  Loyola University Chicago. Se demuestra la aplicación del indicador de fluorescencia, TMRM, en las neuronas corticales para determinar los cambios relativos en la intensidad de fluorescencia TMRM antes y después de la aplicación de un estímulo específico. También muestran la aplicación de la fluorescencia de la sonda H 2 DCF-DA para evaluar el nivel relativo de especies reactivas del oxígeno en las neuronas corticales.

Video: Determination of Mitochondrial Membrane Potential and Reactive Oxygen Species in Live Rat Cortical Neurons

Neuromodulation and Mitochondrial Transport: Live Imaging in Hippocampal Neurons over Long Durations

To understand the relationship between mitochondrial transport and neuronal function, it is critical to observe mitochondrial behavior in live cultured neurons for extended durations1-3. This is now possible through the use of vital dyes and fluorescent proteins with which cytoskeletal components, organelles, and other structures in living cells can be labeled and then visualized via dynamic fluorescence microscopy. For example, in embryonic chicken sympathetic neurons, mitochondrial movement was characterized using the vital dye rhodamine 1234. In another study, mitochondria were visualized in rat forebrain neurons by transfection of mitochondrially targeted eYFP5. However, imaging of primary neurons over minutes, hours, or even days presents a number of issues. Foremost among these are: 1) maintenance of culture conditions such as temperature, humidity, and pH during long imaging sessions; 2) a strong, stable fluorescent signal to assure both the quality of acquired images and accurate measurement of signal intensity during image analysis; and 3) limiting exposure times during image acquisition to minimize photobleaching and avoid phototoxicity.
Here, we describe a protocol that permits the observation, visualization, and analysis of mitochondrial movement in cultured hippocampal neurons with high temporal resolution and under optimal life support conditions. We have constructed an affordable stage-top incubator that provides good temperature regulation and atmospheric gas flow, and also limits the degree of media evaporation, assuring stable pH and osmolarity. This incubator is connected, via inlet and outlet hoses, to a standard tissue culture incubator, which provides constant humidity levels and an atmosphere of 5-10% CO2/air. This design offers a cost-effective alternative to significantly more expensive microscope incubators that don't necessarily assure the viability of cells over many hours or even days. To visualize mitochondria, we infect cells with a lentivirus encoding a red fluorescent protein that is targeted to the mitochondrion. This assures a strong and persistent signal, which, in conjunction with the use of a stable xenon light source, allows us to limit exposure times during image acquisition and all but precludes photobleaching and phototoxicity. Two injection ports on the top of the stage-top incubator allow the acute administration of neurotransmitters and other reagents intended to modulate mitochondrial movement. In sum, lentivirus-mediated expression of an organelle-targeted red fluorescent protein and the combination of our stage-top incubator, a conventional inverted fluorescence microscope, CCD camera, and xenon light source allow us to acquire time-lapse images of mitochondrial transport in living neurons over longer durations than those possible in studies deploying conventional vital dyes and off-the-shelf life support systems.

We describe a protocol that allows imaging of mitochondria in living neurons via fluorescence microscopy over long durations. Imaging over extended periods is accomplished through lentivirus-mediated expression of a mitochondrially targeted fluorescent protein and use of an inexpensive stage-top incubator that was designed and built in our laboratory.

La neuromodulación y mitocondrial de transporte: Imágenes en vivo en las neuronas del hipocampo durante largos períodos de tiempo

To understand the relationship between mitochondrial transport and neuronal function, it is critical to observe mitochondrial behavior in live cultured ...

Investigación

Neurociencias

Quantitative Analysis of Synaptic Vesicle Pool Replenishment in Cultured Cerebellar Granule Neurons using FM Dyes

After neurotransmitter release in central nerve terminals, SVs are rapidly retrieved by endocytosis. Retrieved SVs are then refilled with neurotransmitter and rejoin the recycling pool, defined as SVs that are available for exocytosis1,2. The recycling pool can generally be subdivided into two distinct pools - the readily releasable pool (RRP) and the reserve pool (RP). As their names imply, the RRP consists of SVs that are immediately available for fusion while RP SVs are released only during intense stimulation1,2. It is important to have a reliable assay that reports the differential replenishment of these SV pools in order to understand 1) how SVs traffic after different modes of endocytosis (such as clathrin-dependent endocytosis and activity-dependent bulk endocytosis) and 2) the mechanisms controlling the mobilisation of both the RRP and RP in response to different stimuli.
FM dyes are routinely employed to quantitatively report SV turnover in central nerve terminals3-8. They have a hydrophobic hydrocarbon tail that allows reversible partitioning in the lipid bilayer, and a hydrophilic head group that blocks passage across membranes. The dyes have little fluorescence in aqueous solution, but their quantum yield increases dramatically when partitioned in membrane9. Thus FM dyes are ideal fluorescent probes for tracking actively recycling SVs. The standard protocol for use of FM dye is as follows. First they are applied to neurons and are taken up during endocytosis (Figure 1). After non-internalised dye is washed away from the plasma membrane, recycled SVs redistribute within the recycling pool. These SVs are then depleted using unloading stimuli (Figure 1). Since FM dye labelling of SVs is quantal10, the resulting fluorescence drop is proportional to the amount of vesicles released. Thus, the recycling and fusion of SVs generated from the previous round of endocytosis can be reliably quantified.
Here, we present a protocol that has been modified to obtain two additional elements of information. Firstly, sequential unloading stimuli are used to differentially unload the RRP and the RP, to allow quantification of the replenishment of specific SV pools. Secondly, each nerve terminal undergoes the protocol twice. Thus, the response of the same nerve terminal at S1 can be compared against the presence of a test substance at phase S2 (Figure 2), providing an internal control. This is important, since the extent of SV recycling across different nerve terminals is highly variable11.
Any adherent primary neuronal cultures may be used for this protocol, however the plating density, solutions and stimulation conditions are optimised for cerebellar granule neurons (CGNs)12,13.

A live fluorescence imaging technique to quantify the replenishment and mobilisation of specific synaptic vesicle (SV) pools in central nerve terminals is described. Two rounds of SV recycling are monitored in the same nerve terminals providing an internal control.

Análisis cuantitativo de Reposición Synaptic piscina vesículas en cultivos de neuronas granulares del cerebelo con tintes FM

After neurotransmitter release in central nerve terminals, SVs are rapidly retrieved by endocytosis.  Retrieved SVs are then refilled with ...

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 ...

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 ...

Membrane Potential Dye Imaging of Ventromedial Hypothalamus Neurons From Adult Mice to Study Glucose Sensing

Studies of neuronal activity are often performed using neurons from rodents less than 2 months of age due to the technical difficulties associated with increasing connective tissue and decreased neuronal viability that occur with age. Here, we describe a methodology for the dissociation of healthy hypothalamic neurons from adult-aged mice. The ability to study neurons from adult-aged mice allows the use of disease models that manifest at a later age and might be more developmentally accurate for certain studies. Fluorescence imaging of dissociated neurons can be used to study the activity of a population of neurons, as opposed to using electrophysiology to study a single neuron. This is particularly useful when studying a heterogeneous neuronal population in which the desired neuronal type is rare such as for hypothalamic glucose sensing neurons. We utilized membrane potential dye imaging of adult ventromedial hypothalamic neurons to study their responses to changes in extracellular glucose. Glucose sensing neurons are believed to play a role in central regulation of energy balance. The ability to study glucose sensing in adult rodents is particularly useful since the predominance of diseases related to dysfunctional energy balance (e.g.&#160;obesity) increase with age.

The activity of single neurons from adult-aged mice can be studied by dissociating neurons from specific brain regions and using fluorescent membrane potential dye imaging. By testing responses to changes in glucose, this technique can be used to study the glucose sensitivity of adult ventromedial hypothalamic neurons.

Potencial de Membrana Tinte Imaging de ventromedial del hipotálamo neuronas de ratones adultos para el Estudio de la glucosa Sensing

Studies of neuronal activity are often performed using neurons from rodents less than 2 months of age due to the technical difficulties associated with ...

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 ...

In Vivo Two-photon Imaging of Cortical Neurons in Neonatal Mice

Two-photon imaging is a powerful tool for the in vivo analysis of neuronal circuits in the mammalian brain. However, a limited number of in vivo imaging methods exist for examining the brain tissue of live newborn mammals. Herein we summarize a protocol for imaging individual cortical neurons in living neonatal mice. This protocol includes the following two methodologies: (1) the Supernova system for sparse and bright labeling of cortical neurons in the developing brain, and (2) a surgical procedure for the fragile neonatal skull. This protocol allows the observation of temporal changes of individual cortical neurites during neonatal stages with a high signal-to-noise ratio. Labeled cell-specific gene silencing and knockout can also be achieved by combining the Supernova with RNA interference and CRISPR/Cas9 gene editing systems. This protocol can, thus, be used for analyzing the developmental dynamics of cortical neurons, molecular mechanisms that control the neuronal dynamics, and changes in neuronal dynamics in disease models.

In Vivo Two-photon Imaging of Cortical Neurons in Neonatal Mice

We present an in vivo two-photon imaging protocol for imaging the cerebral cortex of neonatal mice. This method is suitable for analyzing the developmental dynamics of cortical neurons, the molecular mechanisms that control the neuronal dynamics, and the changes in neuronal dynamics in disease models.

En Vivo Proyección de imagen de dos fotones de neuronas corticales en ratones neonatales

Two-photon imaging is a powerful tool for the in vivo analysis of neuronal circuits in the mammalian brain. However, a limited number of in vivo imaging ...

Stimulus-specific Cortical Visual Evoked Potential Morphological Patterns

This paper presents a methodology for the recording and analysis of cortical visual evoked potentials (CVEPs) in response to various visual stimuli using 128-channel high-density electroencephalography (EEG). The specific aim of the described stimuli and analyses is to examine whether it is feasible to replicate previously reported CVEP morphological patterns elicited by an apparent motion stimulus, designed to simultaneously stimulate both ventral and dorsal central visual networks, using object and motion stimuli designed to separately stimulate ventral and dorsal visual cortical networks.&#160; Four visual paradigms are presented: 1. Randomized visual objects with consistent temporal presentation. 2. Randomized visual objects with inconsistent temporal presentation (or jitter).&#160; 3. Visual motion via a radial field of coherent central dot motion without jitter.&#160; 4. Visual motion via a radial field of coherent central dot motion with jitter.&#160; These four paradigms are presented in a pseudo-randomized order for each participant.&#160; Jitter is introduced in order to view how possible anticipatory-related effects may affect the morphology of the object-onset and motion-onset CVEP response.&#160; EEG data analyses are described in detail, including steps of data exportation from and importation to signal processing platforms, bad channel identification&#160;and removal, artifact rejection, averaging, and categorization of average CVEP morphological pattern type based upon latency ranges of component peaks. Representative data show that the methodological approach is indeed sensitive in eliciting differential object-onset and motion-onset CVEP morphological patterns and may, therefore, be useful in addressing the larger research aim. Given the high temporal resolution of EEG and the possible application of high-density EEG in source localization analyses, this protocol is ideal for the investigation of distinct CVEP morphological patterns and the underlying neural mechanisms which generate these differential responses.&#160; &#160; &#160;&#160;

In this paper, we present a protocol to investigate differential cortical visual evoked potential morphological patterns through stimulation of ventral and dorsal networks using high-density EEG. Visual object and motion stimulus paradigms, with and without temporal jitter, are described. Visual evoked potential morphological analyses are also outlined.&#160;&#160;

Patrones morfológicos potenciales evocados visuales corticales específicos del estímulo

This paper presents a methodology for the recording and analysis of cortical visual evoked potentials (CVEPs) in response to various visual stimuli using ...

Generation of Neurospheres from Mixed Primary Hippocampal and Cortical Neurons Isolated from E14-E16 Sprague Dawley Rat Embryo

Primary neuron culture is an essential technique in the field of neuroscience. To gain deeper mechanistic insights into the brain, it is essential to have a robust in vitro model that can be exploited for various neurobiology studies. Though primary neuron cultures (i.e., long-term hippocampal cultures) have provided scientists with models, it does not yet represent the complexity of brain network completely. In the wake of these limitations, a new model has emerged using neurospheres, which bears a closer resemblance to the brain tissue. The present protocol describes the plating of high and low densities of mixed cortical and hippocampal neurons isolated from the embryo of embryonic day 14-16 Sprague Dawley rats. This allows for the generation of neurospheres and long-term primary neuron culture as two independent platforms to conduct further studies. This process is extremely simple and cost-effective, as it minimizes several steps and reagents previously deemed essential for neuron culture. This is a robust protocol with minimal requirements that can be performed with achievable results and further used for a diversity of studies related to neuroscience.

Presented here is a protocol for the spontaneous generation of neurospheres enriched in neural progenitor cells from high density plated neurons. During the same experiment, when neurons are plated at a lower density, the protocol also results in prolonged primary rat neuron cultures.

Generación de neuroesferas a partir de neuronas de hipocampo primario mixto y cortical esinas aisladas del e14-E16 Sprague Dawley Rat Embryo

Primary neuron culture is an essential technique in the field of neuroscience. To gain deeper mechanistic insights into the brain, it is essential to have ...

Determination of Mitochondrial Respiration and Glycolysis in Ex Vivo Retinal Tissue Samples

Mitochondrial respiration is a critical energy-generating pathway in all cells, especially retinal photoreceptors that possess a highly active metabolism. In addition, photoreceptors also exhibit high aerobic glycolysis like cancer cells. Precise measurements of these metabolic activities can provide valuable insights into cellular homeostasis under physiological conditions and in disease states. High throughput microplate-based assays have been developed to measure mitochondrial respiration and various metabolic activities in live cells. However, a vast majority of these are developed for cultured cells and have not been optimized for intact tissue samples and for application ex vivo. Described here is a detailed step-by-step protocol, using microplate-based fluorescence technology, to directly measure oxygen consumption rate (OCR) as an indicator of mitochondrial respiration, as well as extracellular acidification rate (ECAR) as an indicator of glycolysis, in intact ex vivo retinal tissue. This method has been used to successfully assess metabolic activities in adult mouse retina and demonstrate its application in investigating cellular mechanisms of aging and disease.

Determination of Mitochondrial Respiration and Glycolysis in Ex Vivo Retinal Tissue Samples

Described here is a detailed protocol for performing mitochondrial stress assay and glycolytic rate assay in ex vivo retinal tissue samples using a commercial bioanalyzer.

Determinación de la respiración mitocondrial y la glucólisis en muestras de tejido retiniano ex vivo

Mitochondrial respiration is a critical energy-generating pathway in all cells, especially retinal photoreceptors that possess a highly active metabolism. ...