This work was supported by NIH R01 EY020533, WSU Startup Fund, and RPB grants.

Declaración de Ética: Los procedimientos que implican sujetos animales fueron aprobados por el Cuidado y Uso de Animales Comité Institucional (IACUC) de la Universidad Estatal de Wayne. 1. Preparación de la solución Experimental <ol><li> Prepare la solución de disección 1 día a 1 semana antes del experimento real. Use una solución tampón HEPES para la disección de la retina debido a su gran capacidad de amortiguación a temperaturas más bajas 16. Mezclar todos los productos químicos de la siguiente manera (en mM): 115 NaCl, 2,5 KCl, CaCl2 2,5, 1,0 MgCl2, 10 HEPES, 28 glucosa. Ajustar el pH a 7,4 con NaOH. Conservar la solución en el refrigerador hasta por 1 semana. </li><li> La solución de registro es medio de Ames, que es un líquido cefalorraquídeo artificial (LCRa) diseñado para preparaciones de la retina 17. Por el día del experimento, pesar polvo Ames &#39;(4,4 g por 500 ml de solución) en un tubo. Además, pesar NaHCO3 (0,95 g por 500 ml de solución) y mezclar con la Apolvo mes. </li><li> Prepare la solución de la pipeta intracelular para las grabaciones de patch clamp por el día del experimento. Mezclar todos los productos químicos de la siguiente manera (en mM): 111 K-gluconato, 1,0 CaCl2, 10 HEPES, 1,1 EGTA, 10 de NaCl, 1,0 MgCl 2, 5 ATP-Mg y 1,0 GTP-Na. Ajustar el pH a 7,2 con KOH. Se filtra la solución de la pipeta con un filtro de jeringa convencional. Haga ~ 500 ml de alícuotas y almacenarlos en un congelador o un congelador. </li></ol> 2. Preparación para el Día del Experimento <ol><li> La noche antes del experimento, coloca un ratón en una jaula (C57BL / 6J o antecedentes similares, 4-8 semanas de edad, macho) en un espacio oscuro de O / N adaptación a la oscuridad. </li><li> Preparación para la disección: <ol><li> Rellene disección solución (100 - 200 ml) en un vaso de vidrio, colóquelo sobre hielo, y la burbuja de oxígeno durante al menos 10 min. </li><li> Iniciar la oxigenación de la caja oscura para el almacenamiento de preparación de la retina. </li><li> Preparar cubreobjetos de plástico con rieles de engrase para rerebanada preparativos intes- y colocar cada uno de ellos en un plato de plástico de 35 mm. </li><li> Adjunte una nueva hoja de afeitar para un helicóptero (máquina de cortar el tejido). </li><li> Cortar una membrana de filtro en dos mitades. Esto es para el corte en lonchas de la retina. </li><li> Alinear las herramientas de disección y pipetas de transferencia cerca del microscopio de disección. El área de trabajo debe estar bien organizado para que cada herramienta se puede acceder fácilmente en la oscuridad. </li></ol></li><li> Preparación para las grabaciones de patch clamp: <ol><li> Disuelva el polvo de Ames y NaHCO3 en agua destilada (4,4 g Ames y NaHCO 3 0,95 g / 500 ml ddH 2 O). Medio de la burbuja el Ames &#39;con oxígeno mixta (95% O 2 y 5% de CO 2) durante al menos 30 min mientras se calienta a un nivel de temperatura de grabación (~ 35 ° C). Ajustar el pH a 7,4 con NaHCO 3 mientras que todavía burbujeante. </li><li> Descongelar la solución de la pipeta interna durante la disección. Después de que se haya descongelado, añadir 0,01% sulforodamina B para la tinción intracelular. </li></ol></li></ol> 3. Disección de retina <ol><li> En un cuarto oscuro, la eutanasia con el ratón utilizando dióxido de carbono y el neumotórax bilateral. Al cabo de 2 - 3 minutos de usar el dióxido de carbono, el ratón perder el conocimiento. </li><li> Cuando el ratón ya no responde a una pizca de cola, enuclear rápidamente los ojos y colocarlos en una solución de disección fría en un plato de plástico. Realice el neumotórax bilateral para asegurar la muerte del animal. Almacenar rápidamente el ojo en el plato en la caja oscura. Utilice un visor de infrarrojos para llevar a cabo el procedimiento en la oscuridad. NOTA: Como alternativa, isoflurano, la decapitación o dislocación cervical se pueden utilizar para la eutanasia. El método de la eutanasia debe seguir las directrices del Instituto Nacional de Salud (NIH) y / o el Comité de Cuidado y Uso de Animales institucional. </li><li> En una habitación oscura, ajuste el microscopio de disección y la iluminación infrarroja. Bajo el microscopio, transferir un ojo de su plato en un plato de plástico de 10 cm wITH la solución de disección enfriado. Continuamente burbuja de oxígeno a través de la tubería colocada en la parte inferior del plato de plástico. Ajustar el volumen de flujo por lo que no es demasiado baja, pero asegurarse de que no es lo suficientemente alta como para perturbar la disección. </li><li> Córnea y el cristalino eliminación: <ol><li> Hacer una incisión en la parte superior de la córnea con un cuchillo micro-quirúrgica. Sosteniendo el nervio óptico en el otro extremo del globo ocular evita que el ojo se mueva. Extienda el corte con un par de tijeras quirúrgicas. Recorta la córnea con algunos esclerótica alrededor. </li><li> Coge la lente con una pinza fina y tire lentamente el objetivo. Si el vítreo es demasiado pegajosa para quitar la lente, cortar el vítreo con un par de tijeras. </li><li> Una vez que se hace el ojo-taza, vierta suavemente solución de disección fría y oxigenada en el ocular con una pequeña pipeta de transferencia. NOTA: El globo ocular se desinfla rápidamente después se hace una incisión en la córnea. Es importante para mantener la forma del ojo como deformar el ojo durante este proprocedimiento daña la retina y causar desprendimiento de retina. </li></ol></li><li> La identificación de las partes dorsal y ventral de la retina: NOTA: Debido a la desigual distribución de la verde y la luz ultravioleta (UV) conos 18, la identificación de un área específica de la retina que es importante para las grabaciones de respuesta de luz. <ol><li> Identificar la línea que va a través del ocular de la retina que pasa cerca de la cabeza del nervio óptico. Esta línea pasa sobre todo a través de la parte ventral del nervio óptico; el lado incluyendo el nervio óptico es el lado dorsal, mientras que el otro es ventral 19. Ver este hito claramente bajo espectadores infrarrojos. </li><li> Dependiendo del propósito del experimento, hacer un corte en el lado innecesario. Esto será útil después de aislar la retina de la copa para el ojo. </li></ol></li><li> Eliminación Vítreo: <ol><li> Opcionalmente, se incuba el ojo-taza con hialuronidasa (0,5 mg / ml) durante 15 min. </li><li> Retire el vítreo con pinzas extra-finas. El humor vítreo es principalmenteunido a la parte inferior y el borde exterior del ojo-cup. Retire con cuidado sin tocar o meter la retina. Como el vítreo podría estar estrechamente unida a la ojera, retire con cuidado para evitar el desprendimiento de retina del ojo taza. Continúe tirando hasta que no sienta la tensión. NOTA: Este paso es especialmente importante para la disección de la retina del ratón; Sin embargo, no es un problema para las retinas de otras especies, tales como la rata o salamandra. </li></ol></li><li> Aislar la retina del ojo taza. Coge la esclerótica y pelar suavemente de la retina utilizando la parte trasera de fórceps. NOTA: Todo el montaje preparativos de la retina se puede hacer de la retina aislada. Hacer tres y cincuenta y siete cortes en los bordes y aplanar la retina, o cortar la retina en varios pedazos. </li><li> Recortar la retina y deseche el medio innecesario de la retina (paso 3.5). En el mismo plato de plástico, cortar la retina restante en dos piezas. Por cada pieza de la retina, cortar los bordes para hacer unalosa de la retina, y recortar los bordes plegables. </li><li> Transferir una losa de la retina en una placa de vidrio usando una pipeta de transferencia de gran tamaño (~ 2 ml). Aspirar el exceso de solución con una pequeña pipeta, usar un trozo de papel de filtro para aplanar la losa, y luego colocar rápidamente una pieza de la membrana de filtro en la parte superior del tejido. <ol><li> Utilizando una pequeña pipeta de transferencia, colocar una gota de solución en la disección de la membrana de filtro. Espere ~ 15 seg hasta que la solución se extiende sobre la membrana. Durante este tiempo, el tejido de la retina se adhieren a la membrana de filtro. </li><li> Verter solución de disección más fría debajo y alrededor del filtro. El tejido de la retina con la membrana de filtro flotará. Coge la membrana de filtro con una pinza y colocarla en la cámara de corte. Vierta la disección de solución sobre el tejido, y almacenar la preparación en la caja oscura. NOTA: El procedimiento descrito aquí es fundamental. Es importante para llevar a cabo estos pasos con rapidez para garantizar que los palos tejido de la retina a los filtros de membrana y no se seque. </li></ol></li><li> El uso de un interruptor, cortar el tejido de la retina en rodajas. Alrededor de 200 micras de espesor es factible para los estudios de patch clamp. </li><li> Coloque un cubreobjetos de plástico con rieles de grasa en la cámara de corte. Transferir una rebanada de la retina en la parte superior de los carriles de grasa en un cubreobjetos de plástico. Girar la rebanada 90 ° de modo que la sección transversal es visible. Presione la membrana de filtro hacia abajo sobre el cubreobjetos, a continuación, cubrir los lados del papel de filtro con grasa. <ol><li> Después de la rebanada se inmoviliza sobre un cubreobjetos de plástico, agarrar el cubreobjetos y la transfiere a un plato de plástico de 35 mm. Vierta la solución de disección fría en la preparación de la retina y sumergir la preparación. Almacenar cada plato en la caja oscura, que debe ser oxigenada continuamente. NOTA: Todo el procedimiento que se debe hacer con cuidado para que la membrana de filtro no se deforma. De lo contrario, la rebanada de la retina se separa fácilmente de la membrana de filtro. </li></ol></li></ol><ol><li> Grabación de preparación: <ol><li> Prime los tubos de perfusión con medio de Ames. Permitir todas las burbujas que atraviesan al llenar el tubo. </li><li> Haga pipetas de registro patch clamp con un extractor. Para llenar una pipeta con la solución de la pipeta, baño en la parte trasera de una pipeta de 1 min hasta que se rellena la punta de la pipeta. A continuación, llenar ~ 1/3 de la pipeta usando una carga micro-pipeta. Guarde cada pipeta en una caja pipeta húmedo. </li><li> Encienda el equipo para patch clamp grabaciones; incluyendo la computadora, un amplificador, cámara CCD, y el microscopio. </li></ol></li><li> Apague la luz de la habitación. Coloque una rebanada preparación de la retina en la cámara de platina del microscopio utilizando un visor de infrarrojos. Después de que se inmoviliza, comenzar la perfusión continua. Ajuste la temperatura de perfusión a 33 a 37 ° C. </li><li> Ver la superficie rebanada con la cámara CCD. Centrarse en la ubicación de destino donde el objetivo cetipos ll residen. Seleccione una celda de aspecto saludable para la grabación de patch clamp (es decir, debe tener la superficie de aspecto suave y una buena forma de la célula). </li><li> Coloque una pipeta de grabación en un soporte de pipetas. Avanzar en la pipeta para la preparación rebanada. Cuando se está cerca de la rebanada preparación (~ 2 mm por encima), encontrar la punta de la pipeta con el microscopio. Una vez que la punta de la pipeta es visible bajo el microscopio, mover la punta hacia abajo, hacia la célula diana. </li><li> Ajuste el amplificador. Ajustar la pipeta a 0 mV / 0 pA (puesta a cero) y comenzar un pulso eléctrico continuo de ~ 5 mV a ~ 10 Hz. Compruebe la resistencia de la pipeta; resistencia ideal está entre 5 y 10 MΩ para la mayoría de las neuronas de la retina. </li><li> Comience soplando desde la punta. Utilice una boquilla, o utilizar una jeringa, para aplicar presión positiva cuando la pipeta se sumerge en la solución del baño. Continuamente soplar la solución interna hasta que la punta está en la superficie de la célula diana. <ol><li> Cuando la presión positiva hace un pequeño hoyuelo en el thsuperficie celular e, avanzar en la punta ligeramente y dejar de soplar. Compruebe la resistencia de la pipeta, que debería aumentar. Si está aumentando continuamente, déjelo y vigilar la resistencia hasta que llegue&gt; 1 GΩ (gigaseal). Si la resistencia no aumenta de forma espontánea, aplicar suavemente presión negativa hasta que se convierte en un gigaseal. </li></ol></li><li> Después de lograr la gigaseal, cambie el potencial de mantenimiento de -70 mV. A continuación, aplicar intermitentemente presión negativa para romper la membrana dentro de la punta de la pipeta. Cuando se realiza la configuración de célula completa, la resistencia de la pipeta puede estar entre 500 MΩ y 1 GΩ, y la corriente capacitiva se ve. A veces las corrientes postsinápticas espontáneas se pueden observar. </li><li> Anote la relación IV -80-40 mV. Los diferentes tipos de canales dependientes de voltaje se activan dependiendo del tipo de célula. </li><li> Evocó luz corrientes o tensiones sinápticas Record. </li></ol>

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The authors have nothing to disclose.

Good recordings can only be achieved with good retinal preparations and well-designed patch clamp setups. Although all the steps described above are important, the discussion highlights some critical steps both for the dissection and recordings. 
For dissection, two things are especially important: cooling and oxygenation. After enucleating the eye, quickly remove the front part of the eye in a dissecting chamber with oxygen-bubbled, cooled dissecting solution, and pour cold solution into the eye-cup. When the retinal tissue is in the dissecting chamber, cold solution should be poured on the tissue every 10 min. The tissue should be sliced quickly without pulling or poking the retinal tissue during the dissection, which could easily damage its structure. Using good dissecting tools (sharp forceps and scissors) is also crucial. A chopper is equally as effective as a vibratome because it allows for speedy maneuvering. Extra-fine forceps should be used to remove vitreous matter as efficiently as possible because residual vitreous damages the retinal structure when slicing the tissue.
Beginners tend to damage the retina when removing the lens. The retina can detach from the eye-cup, or can even be accidentally removed with the lens. To avoid this issue, the sclera can be cut around the Ora Serrata which is located close to the extra-ocular muscle attachment point. An eye-cup with a wide opening makes it easier to remove the lens. Also, it is important not to deform the eye when removing the front portion. Another critical point is fixing the retinal slice onto a coverslip. This can be facilitated by practicing rotating the filter paper without distorting it to prevent the retinal slice from coming off during the experiment. 
A sturdy, well-built rig is crucial for patch clamp recordings. It must be free of electric noise, vibration, and pipette drifting. In addition, light response can be recorded in dark conditions to prevent desensitization. Our cage is made with a metal frame and aluminum panels with black plastic drapes that act as an electric shield while also completely blocking light from entering the cage; computer monitor illumination will not affect the retinal preparation. Use a remote controller for the pipette manipulator and close the cage while setting up the patch clamp configuration, which also avoid illuminating the preparation.
Although we have been tried to simplify the procedures involved, patch clamp study is still a difficult experiment that requires extensive knowledge and training. An automated patch clamp has been developed by some groups20; this system is applicable to dissociated cells and might be an efficient technique for pharmacological screenings. 
In summary, patch clamp recording in dark-adapted retinal slice preparations is a useful technique to investigate visual information processing. Compared to other methods, patch clamp offers better temporal and spatial resolution. Additional technical innovations will make the patch clamp method more user-friendly in the near future.

The retina is one of the unique parts of the nervous system. As an accessible part of the brain, its synaptic architecture has been well characterized. In addition, the functions of this neural network can be examined with a physiological stimulus: light. If the retinal tissue is isolated in a dark room with appropriate procedures, neurons in the tissue will respond to light. This preparation has been used to study visual signal processing and elucidate various synaptic mechanisms and neural network functions, as well as disease mechanisms. 
Light responses in retinal neurons have been recorded for decades. Early studies used sharp electrodes to make intracellular recordings from mudpuppy retinal neurons1. In the 1980s, the patch clamp technique was invented2, and soon became a popular method among vision researchers3,4. Single cell recordings from lower vertebrates, including mudpuppy and fish retinal neurons, were popular methods that contributed to the elucidation of visual signal processing mechanisms5,6. 
After genetic mutation techniques were developed, the mouse retina became a more popular model for vision researchers7-9. The mammalian retina is more attractive than that of lower vertebrates because it is evolutionarily closer to the human retina, and there is an opportunity to use disease models. However, mouse retinal cells are small and fragile10, and making retinal preparations and conducting patch clamp recordings in a dark room is challenging. As technology has improved, diverse approaches have become available to study visual signaling mechanisms such as imaging studies11 and the electroretinogram (ERG)12. Nevertheless, single cell recording with the patch clamp method is still important because it is highly temporally and spatially sensitive compared to other methods. Therefore, we have continuously conducted patch clamp recordings and improved our methods to investigate visual signal processing in mouse retinal slice preparations13-15.
In this video tutorial, the protocols are presented with important tips. Good recordings can only be achieved with good preparation. Practicing animal dissection and building a sturdy patch clamp rig will enable most researchers to achieve successful recordings.

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		<tr height="42">
			<td height="42" style="height:42px;width:216px;">Name of Material/ Equipment</td>
			<td style="width:145px;">Company</td>
			<td style="width:203px;">Catalog Number</td>
			<td style="width:323px;">Comments/Description</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">mice (28-60 days old, male)</td>
			<td style="width:145px;">Jackson laboratory</td>
			<td style="width:203px;">C57BL/6J strain</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Ames&#39; medium powder</td>
			<td style="width:145px;">Sigma</td>
			<td style="width:203px;">A1420</td>
			<td>excellent</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Stereo microscope</td>
			<td style="width:145px;">Nikon</td>
			<td style="width:203px;">SMZ745</td>
			<td>excellent</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">dissecting tool_forceps</td>
			<td style="width:145px;">Dumont</td>
			<td style="width:203px;">#4, #5, #55</td>
			<td>excellent</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">dissecting tool_scissors</td>
			<td style="width:145px;">Roboz</td>
			<td style="width:203px;">RS-5605</td>
			<td>excellent</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">dissecting tool_surgery knife</td>
			<td style="width:145px;">Surgistar</td>
			<td style="width:203px;">7514</td>
			<td>excellent</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">razor blade (for chopper)</td>
			<td style="width:145px;">EMS</td>
			<td style="width:203px;">71970</td>
			<td>excellent</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">chopper</td>
			<td style="width:145px;">handmade</td>
			<td style="width:203px;"></td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">infrared viewer</td>
			<td style="width:145px;">Night Owl Optics</td>
			<td style="width:203px;">NOBG1</td>
			<td>It shows bright view.&#160; Focusing small objects is an issue.</td>
		</tr>
		<tr height="27">
			<td height="27" style="height:27px;width:216px;">infrared pocket scopes</td>
			<td style="width:145px;">B.E. Meyers</td>
			<td style="width:203px;">OWL Gen 3 NV pocketscope</td>
			<td>excellent view</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">puller</td>
			<td style="width:145px;">Sutter</td>
			<td style="width:203px;">P-1000</td>
			<td>excellent.&#160; Make consistent size pipettes.</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">dark box</td>
			<td style="width:145px;">Pelican</td>
			<td style="width:203px;">dark box</td>
			<td>excellent</td>
		</tr>
		<tr height="67">
			<td height="67" style="height:67px;width:216px;">patch clamp system</td>
			<td style="width:145px;">Scientifica</td>
			<td style="width:203px;">slice scope 2000</td>
			<td style="width:323px;">Excellent setup.&#160; Most key components are included in one package.&#160; Micromanipulators are excellent.</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">amplifier</td>
			<td style="width:145px;">Molecular Devices</td>
			<td style="width:203px;">multiclamp 700B</td>
			<td>Excellent and easy control.</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">acquiring software</td>
			<td style="width:145px;">Molecular Devices</td>
			<td style="width:203px;">pClamp software</td>
			<td>Excellent and easy control.</td>
		</tr>
		<tr height="28">
			<td height="28" style="height:28px;width:216px;">light source (LED)</td>
			<td style="width:145px;">Cool LED</td>
			<td style="width:203px;">pE-2 4 channel system</td>
			<td>Excellent</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">CCD camera</td>
			<td style="width:145px;">Q-imaging</td>
			<td style="width:203px;">Retiga 2000</td>
			<td>Excellent</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Faraday cage</td>
			<td style="width:145px;">handmade</td>
			<td style="width:203px;"></td>
			<td></td>
		</tr>
	</tbody>
</table>

recording light-evoked postsynaptic responses in neurons in dark-adapted, mouse retinal slice preparations using patch clamp techniques

La retina es la puerta de entrada al sistema visual. Para entender los mecanismos de procesamiento de señales visuales, investigamos las funciones de redes neuronales de la retina. Neuronas de la retina en la red forman parte de numerosos subtipos. Más de 10 subtipos de células bipolares, células ganglionares, células amacrinas y han sido identificados por los estudios morfológicos. Múltiples subtipos de neuronas de la retina se cree que codificar distintas características de señalización visual, tales como el movimiento y el color, y formar múltiples vías neurales. Sin embargo, los papeles funcionales de cada neurona en el procesamiento de señal visual no se entienden completamente. El método de patch clamp es útil para abordar esta cuestión fundamental. Aquí, un protocolo para registrar las respuestas sinápticas luz evocadas en las neuronas de la retina de ratón utilizando grabaciones de patch clamp en condiciones adaptadas a la oscuridad se proporciona. Los ojos del ratón son adaptado a la oscuridad O / N, y el tramo preparativos de la retina son disecados en un cuarto oscuro utilizando iluminación infrarroja y los espectadores. La luz infrarroja noactivar fotorreceptores de ratón y así preserva su capacidad de respuesta luz. Patch clamp se utiliza para registrar las respuestas de luz evocadas en las neuronas de la retina. Se inyecta un tinte fluorescente durante las grabaciones para caracterizar los subtipos morfológicos neuronales. Este procedimiento nos permite determinar las funciones fisiológicas de cada neurona en la retina del ratón.

Una rodaja de preparación representativo se muestra en la Figura 1. La preparación rebanada está en una posición recta, mostrando fotorreceptores a las células ganglionares en una superficie plana, y no hay desprendimiento del papel de filtro. Si se inclina una rebanada, sólo una parte de la preparación está en foco, lo que hace difícil identificar una célula apropiada para la sujeción del parche. Para las grabaciones, es importante seleccionar una buena soma, que generalmente tiene una superficie brillante, una forma incluso redonda, y no hay placas oscuros visibles. Después se hizo la configuración de célula completa, la rodaja se expone a la luz de fondo y luego a un paso de la luz (Figura 2). Potenciales postsinápticos excitatorios-Light evocado (L-EPSPS) fueron evocados en respuesta a una luz paso (sincronización se muestra en amarillo). La amplitud de pico y el tiempo de decaimiento variar dependiendo del tipo de célula y subtipo. Una de las células ganglionares genera potenciales de acción en respuesta a los estímulos luminosos (Figura 2 D &amp; E). El cell tipo y su subtipo morfológico fueron revelados por medio del etiquetado sulforrodamina después de las grabaciones fisiológicas (Figura 2, derecha). <img alt="Figura 1" src="/files/ftp_upload/52422/52422fig1highres.jpg" width="650" /> Figura 1. preparativos trozo de retina. (A) Una rodaja de preparación de la retina se ve con un objetivo de 10X. Una rodaja de preparación de la retina (parte superior) fue unido a un trozo de papel de filtro (la parte inferior). (B) Una compilación de cuatro imágenes que muestra una rebanada preparación de la retina se ve con un objetivo 60X. Cada capa de células se observa claramente (ONL: capa exterior nuclear, OPL: capa plexiforme externa, INL: la capa nuclear interna, IPL: capa plexiforme interna, GCL: capa de células ganglionares) <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/52422/52422fig1highres.jpg" target="_blank">Haga clic aquí para ver una versión más grande de esta figura.</a> Figura 2. potenciales postsinápticos excitatorios-Light evocado (L-EPSPS) de las neuronas de la retina. Paso a la luz evocaba L-EPSPS (izquierda). La intensidad de la luz fue 30 - 60% de contraste Weber. Antecedentes nivel adaptación a la luz fue de 4 x 10 4 fotones / micras 2 / seg. Sulforodamina B se inyectó durante patch clamp grabación para visualizar las neuronas registradas (derecha). Una pipeta de grabación estaba todavía unido a la soma (A) L-EPSP y tinción sulforrodamina de un cono de célula bipolar ON. (B) OFF célula bipolar cono. (C) de células amacrinas. (D) en las células ganglionares. (E) OFF de células ganglionares. La barra de escala indica 2 o 5 mV como se ha señalado. Estimulación de la luz se aplicó durante 1 seg. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/52422/52422fig2highres.jpg" target="_blank">Por favor, haga clic aquí</a>para ver una versión más grande de esta figura.

Watch this Scientific Journal Video about Recording Light-evoked Postsynaptic Responses in Neurons in Dark-adapted, Mouse Retinal Slice Preparations Using Patch Clamp Techniques at JoVE.com

Recording Light-evoked Postsynaptic Responses in Neurons in Dark-adapted, Mouse Retinal Slice Preparations Using Patch Clamp Techniques

We will demonstrate how to prepare retinal slices from the mouse eye and record light responses in retinal neurons. The entire procedure is conducted in dark-adapted conditions.

Grabación Luz evocó-Respuestas postsinápticos en las neuronas en adaptadas oscuro, Preparaciones ratón retina Slice Uso de técnicas Patch Clamp

The retina is the gateway to the visual system. To understand visual signal processing mechanisms, we investigate retinal neural network functions. ...

recording-light-evoked-postsynaptic-responses-neurons-dark-adapted

Nanyang Technological University

Research

JoVE Journal

Neuroscience

8.4K vistas.  Wayne State University School of Medicine. We will demonstrate how to prepare retinal slices from the mouse eye and record light responses in retinal neurons. The entire procedure is conducted in dark-adapted conditions.

Video: Recording Light-evoked Postsynaptic Responses in Neurons in Dark-adapted, Mouse Retinal Slice Preparations Using Patch Clamp Techniques

Analyzing Responses of Mouse Olfactory Sensory Neurons Using the Air-phase Electroolfactogram Recording

Animals depend on olfaction for many critical behaviors, such as finding food sources, avoiding predators, and identifying conspecifics for mating and other social interactions. The electroolfactogram (EOG) recording is an informative, easy to conduct, and reliable method to assay olfactory function at the level of the olfactory epithelium. Since the 1956 description of the EOG by Ottoson in frogs1, the EOG recording has been applied in many vertebrates including salamanders, rabbits, rats, mice, and humans (reviewed by Scott and Scott-Johnson, 2002, ref. 2). The recent advances in genetic modification in mice have rekindled interest in recording the EOG for physiological characterization of olfactory function in knock-out and knock-in mice. EOG recordings have been successfully applied to demonstrate the central role of olfactory signal transduction components3-8, and more recently to characterize the contribution of certain regulatory mechanisms to OSN responses9-12.
Odorant detection occurs at the surface of the olfactory epithelium on the cilia of OSNs, where a signal transduction cascade leads to opening of ion channels, generating a current that flows into the cilia and depolarizes the membrane13. The EOG is the negative potential recorded extracellularly at the surface of the olfactory epithelium upon odorant stimulation, resulting from a summation of the potential changes caused by individual responsive OSNs in the recording field2. Comparison of the amplitude and kinetics of the EOG thus provide valuable information about how genetic modification and other experimental manipulations influence the molecular signaling underlying the OSN response to odor. 
Here we describe an air-phase EOG recording on a preparation of mouse olfactory turbinates. Briefly, after sacrificing the mouse, the olfactory turbinates are exposed by bisecting the head along the midline and removing the septum. The turbinate preparation is then placed in the recording setup, and a recording electrode is placed at the surface of the olfactory epithelium on one of the medial turbinates. A reference electrode is electrically connected to the tissue through a buffer solution. A continuous stream of humidified air is blown over the surface of the epithelium to keep it moist. The vapor of odorant solutions is puffed into the stream of humidified air to stimulate the epithelium. Responses are recorded and digitized for further analysis.

The electroolfactogram (EOG) recording is an informative, easy-to-conduct, and reliable way of assessing olfactory function at the level of the olfactory epithelium. This protocol describes a recording setup, mouse tissue preparation, data collection, and basic data analysis.

Las respuestas de los análisis de las neuronas sensoriales olfativas ratón Utilizando la grabación Electroolfactogram Aire fase

Animals depend on olfaction for many critical behaviors, such as finding food sources, avoiding predators, and identifying conspecifics for mating and ...

Investigación

Neurociencias

Patch Clamp Recordings from Mouse Retinal Neurons in a Dark-adapted Slice Preparation

Our visual experience is initiated when the visual pigment in our retinal photoreceptors absorbs photons of light energy and initiates a cascade of intracellular events that lead to closure of cyclic-nucleotide-gated channels in the cell membrane. The resulting change in membrane potential leads in turn to reductions in the amount of neurotransmitter release from both rod and cone synaptic terminals. To measure how the light-evoked change in photoreceptor membrane potential leads to downstream activity in the retina, scientists have made electrophysiological recordings from retinal slice preparations for decades1,2. In the past these slices have been cut manually with a razor blade on retinal tissue that is attached to filter paper; in recent years another method of slicing has been developed whereby retinal tissue is embedded in low gelling temperature agar and sliced in cool solution with a vibrating microtome3,4. This preparation produces retinal slices with less surface damage and very robust light-evoked responses. Here we document how this procedure can be done under infrared light to avoid bleaching the visual pigment.

Here we describe a procedure for generating dark-adapted slices of the mouse retina for electrophysiological recordings.

Grabaciones de patch clamp de neuronas de ratón de retina en una rebanada preparación adaptada a la oscuridad

Our visual experience is initiated when the visual pigment in our retinal photoreceptors absorbs photons of light energy and initiates a cascade of ...

Transfection of Mouse Retinal Ganglion Cells by in vivo Electroporation

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

Transfection of Mouse Retinal Ganglion Cells by in vivo Electroporation

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

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

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

Patch-clamp Capacitance Measurements and Ca2+ Imaging at Single Nerve Terminals in Retinal Slices

Visual stimuli are detected and conveyed over a wide dynamic range of light intensities and frequency changes by specialized neurons in the vertebrate retina. Two classes of retinal neurons, photoreceptors and bipolar cells, accomplish this by using ribbon-type active zones, which enable sustained and high-throughput neurotransmitter release over long time periods. ON-type mixed bipolar cell (Mb) terminals in the goldfish retina, which depolarize to light stimuli and receive mixed rod and cone photoreceptor input, are suitable for the study of ribbon-type synapses both due to their large size (~10-12 &mu;m diameter) and to their numerous lateral and reciprocal synaptic connections with amacrine cell dendrites. Direct access to Mb bipolar cell terminals in goldfish retinal slices with the patch-clamp technique allows the measurement of presynaptic Ca2+ currents, membrane capacitance changes, and reciprocal synaptic feedback inhibition mediated by GABAA and GABAC receptors expressed on the terminals. Presynaptic membrane capacitance measurements of exocytosis allow one to study the short-term plasticity of excitatory neurotransmitter release 14,15. In addition, short-term and long-term plasticity of inhibitory neurotransmitter release from amacrine cells can also be investigated by recordings of reciprocal feedback inhibition arriving at the Mb terminal 21. Over short periods of time (e.g. ~10 s), GABAergic reciprocal feedback inhibition from amacrine cells undergoes paired-pulse depression via GABA vesicle pool depletion 11. The synaptic dynamics of retinal microcircuits in the inner plexiform layer of the retina can thus be directly studied.
The brain-slice technique was introduced more than 40 years ago but is still very useful for the investigation of the electrical properties of neurons, both at the single cell soma, single dendrite or axon, and microcircuit synaptic level 19. Tissues that are too small to be glued directly onto the slicing chamber are often first embedded in agar (or placed onto a filter paper) and then sliced 20, 23, 18, 9. In this video, we employ the pre-embedding agar technique using goldfish retina. Some of the giant bipolar cell terminals in our slices of goldfish retina are axotomized (axon-cut) during the slicing procedure. This allows us to isolate single presynaptic nerve terminal inputs, because recording from axotomized terminals excludes the signals from the soma-dendritic compartment. Alternatively, one can also record from intact Mb bipolar cells, by recording from terminals attached to axons that have not been cut during the slicing procedure. Overall, use of this experimental protocol will aid in studies of retinal synaptic physiology, microcircuit functional analysis, and synaptic transmission at ribbon synapses.

Patch-clamp Capacitance Measurements and Ca2+ Imaging at Single Nerve Terminals in Retinal Slices

Here we describe a protocol for the preparation of agar-embedded retinal slices that are suitable for electrophysiology and Ca2+ imaging. This method allows one to study ribbon-type synapses in retinal microcircuits using direct patch-clamp recordings of single presynaptic nerve terminals.

Patch-clamp mediciones de capacitancia y Ca 2 + Imágenes en las terminaciones nerviosas individuales en rebanadas de retina

Visual stimuli are detected and conveyed over a wide dynamic range of light intensities and frequency changes by specialized neurons in the vertebrate ...

Imaging Neuronal Responses in Slice Preparations of Vomeronasal Organ Expressing a Genetically Encoded Calcium Sensor

The vomeronasal organ (VNO) detects chemosensory signals that carry information about the social, sexual and reproductive status of the individuals within the same species 1,2. These intraspecies signals, the pheromones, as well as signals from some predators 3, activate the vomeronasal sensory neurons (VSNs) with high levels of specificity and sensitivity 4. At least three distinct families of G-protein coupled receptors, V1R, V2R and FPR 5-14, are expressed in VNO neurons to mediate the detection of the chemosensory cues. To understand how pheromone information is encoded by the VNO, it is critical to analyze the response profiles of individual VSNs to various stimuli and identify the specific receptors that mediate these responses.
The neuroepithelia of VNO are enclosed in a pair of vomer bones. The semi-blind tubular structure of VNO has one open end (the vomeronasal duct) connecting to the nasal cavity. VSNs extend their dendrites to the lumen part of the VNO, where the pheromone cues are in contact with the receptors expressed at the dendritic knobs. The cell bodies of the VSNs form pseudo-stratified layers with V1R and V2R expressed in the apical and basal layers respectively 6-8. Several techniques have been utilized to monitor responses of VSNs to sensory stimuli 4,12,15-19. Among these techniques, acute slice preparation offers several advantages. First, compared to dissociated VSNs 3,17, slice preparations maintain the neurons in their native morphology and the dendrites of the cells stay relatively intact. Second, the cell bodies of the VSNs are easily accessible in coronal slice of the VNO to allow electrophysiology studies and imaging experiments as compared to whole epithelium and whole-mount preparations 12,20. Third, this method can be combined with molecular cloning techniques to allow receptor identification.
Sensory stimulation elicits strong Ca2+ influx in VSNs that is indicative of receptor activation 4,21. We thus develop transgenic mice that express G-CaMP2 in the olfactory sensory neurons, including the VSNs 15,22. The sensitivity and the genetic nature of the probe greatly facilitate Ca2+ imaging experiments. This method has eliminated the dye loading process used in previous studies 4,21. We also employ a ligand delivery system that enables application of various stimuli to the VNO slices. The combination of the two techniques allows us to monitor multiple neurons simultaneously in response to large numbers of stimuli. Finally, we have established a semi-automated analysis pipeline to assist image processing.

The vomeronasal organ (VNO) detects intraspecies chemical signals that convey social and reproductive information. We have performed Ca2+ imaging experiments using transgenic mice expressing G-CaMP2 in VNO tissue. This approach allows us to analyze the complicated response patterns of the vomeronasal neurons to large numbers of pheromone stimuli.

Las respuestas de imagen neuronal en los preparativos de la rebanada del órgano vomeronasal Expresando un sensor de calcio genéticamente codificados

The vomeronasal organ (VNO) detects chemosensory signals that carry information about the social, sexual and reproductive status of the individuals within ...

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

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

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

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

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

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

Perforated Patch-clamp Recording of Mouse Olfactory Sensory Neurons in Intact Neuroepithelium: Functional Analysis of Neurons Expressing an Identified Odorant Receptor

Analyzing the physiological responses of olfactory sensory neurons (OSN) when stimulated with specific ligands is critical to understand the basis of olfactory-driven behaviors and their modulation. These coding properties depend heavily on the initial interaction between odor molecules and the olfactory receptor (OR) expressed in the OSNs. The identity, specificity and ligand spectrum of the expressed OR are critical. The probability to find the ligand of the OR expressed in an OSN chosen randomly within the epithelium is very low. To address this challenge, this protocol uses genetically tagged mice expressing the fluorescent protein GFP under the control of the promoter of defined ORs. OSNs are located in a tight and organized epithelium lining the nasal cavity, with neighboring cells influencing their maturation and function. Here we describe a method to isolate an intact olfactory epithelium and record through patch-clamp recordings the properties of OSNs expressing defined odorant receptors. The protocol allows one to characterize OSN membrane properties while keeping the influence of the neighboring tissue. Analysis of patch-clamp results yields&#160;a precise quantification of ligand/OR interactions, transduction pathways and pharmacology, OSNs&#39; coding properties and their modulation at the membrane level.&#160;

Analyzing the physiological properties of olfactory sensory neurons still faces technical limitations. Here we record them through perforated patch-clamp in an intact preparation of the olfactory epithelium in gene-targeted mice. This technique allows the characterization of membrane properties and responses to specific ligands of neurons expressing defined olfactory receptors.

Perforado grabación de patch-clamp de ratón neuronas sensoriales olfativas en neuroepitelio Intacto: Análisis funcional de las neuronas que expresan un odorante del receptor Identificado

Analyzing the physiological responses of olfactory sensory neurons (OSN) when stimulated with specific ligands is critical to understand the basis of ...

Whole-cell Patch-clamp Recordings in Brain Slices

Whole-cell patch-clamp recording is an electrophysiological technique that allows the study of the&#160;electrical properties of a substantial part of the neuron. In this configuration, the micropipette is in tight contact with the cell membrane, which prevents current leakage and thereby provides more accurate ionic current measurements than the previously used intracellular sharp electrode recording method. Classically, whole-cell recording can be performed on neurons in various types of preparations, including cell culture models, dissociated neurons, neurons in brain slices, and in intact anesthetized or awake animals. In summary, this technique has immensely contributed to the understanding of passive and active biophysical properties of excitable cells. A major advantage of this technique is that it provides information on how specific manipulations (e.g., pharmacological, experimenter-induced plasticity) may alter specific neuronal functions or channels in real-time. Additionally, significant opening of the plasma membrane allows the internal pipette solution to freely diffuse into the cytoplasm, providing means for introducing drugs, e.g., agonists or antagonists of specific intracellular proteins, and manipulating these targets without altering their functions in neighboring cells. This article will focus on whole-cell recording performed on neurons in brain slices,&#160;a preparation that has the advantage of recording neurons in relatively well preserved brain circuits, i.e., in a physiologically relevant context. In particular,&#160;when combined with appropriate pharmacology, this technique is a powerful tool allowing identification of specific neuroadaptations that occurred following any type of experiences, such as learning, exposure to drugs of abuse, and stress. In summary, whole-cell patch-clamp recordings in brain slices provide means to measure in ex vivo preparation long-lasting changes in neuronal functions that have developed in intact awake animals.

This protocol describes basic procedural steps for performing whole-cell patch-clamp recordings. This technique allows the study of&#160;the electrical behavior of neurons, and when performed in brain slices, allows the assessment of various neuronal functions from neurons that are still integrated in relatively well preserved brain circuits.

Las grabaciones de células enteras Patch-clamp en rodajas de cerebro

Whole-cell patch-clamp recording is an electrophysiological technique that allows the study of the&#160;electrical properties of a substantial part of the ...

Patch Clamp Recording of Starburst Amacrine Cells in a Flat-mount Preparation of Deafferentated Mouse Retina

The mammalian retina is a layered tissue composed of multiple neuronal types. To understand how visual signals are processed within its intricate synaptic network, electrophysiological recordings are frequently used to study connections among individual neurons. We have optimized a flat-mount preparation for patch clamp recording of genetically marked neurons in both GCL (ganglion cell layer) and INL (inner nuclear layer) of mouse retinas. Recording INL neurons in flat-mounts is favored over slices because both vertical and lateral connections are preserved in the former configuration, allowing retinal circuits with large lateral components to be studied. We have used this procedure to compare responses of mirror-partnered neurons in retinas such as the cholinergic starburst amacrine cells (SACs).

This protocol demonstrates how to perform whole-cell patch clamp recording on retinal neurons from a flat-mount preparation.

Patch Clamp grabación de células amacrinas Starburst en un plano de montaje Preparación de Deafferentated retina del ratón

The mammalian retina is a layered tissue composed of multiple neuronal types. To understand how visual signals are processed within its intricate synaptic ...

Slice Patch Clamp Technique for Analyzing Learning-Induced Plasticity

The slice patch clamp technique is a powerful tool for investigating learning-induced neural plasticity in specific brain regions. To analyze motor-learning induced plasticity, we trained rats using an accelerated rotor rod task. Rats performed the task 10 times at 30-s intervals for 1 or 2 days. Performance was significantly improved on the training days compared to the first trial. We then prepared acute brain slices of the primary motor cortex (M1) in untrained and trained rats. Current-clamp analysis showed dynamic changes in resting membrane potential, spike threshold, afterhyperpolarization, and membrane resistance in layer II/III pyramidal neurons. Current injection induced many more spikes in 2-day trained rats than in untrained controls.
To analyze contextual-learning induced plasticity, we trained rats using an inhibitory avoidance (IA) task. After experiencing foot-shock in the dark side of a box, the rats learned to avoid it, staying in the lighted side. We prepared acute hippocampal slices from untrained, IA-trained, unpaired, and walk-through rats. Voltage-clamp analysis was used to sequentially record miniature excitatory and inhibitory postsynaptic currents (mEPSCs and mIPSCs) from the same CA1 neuron. We found different mean mEPSC and mIPSC amplitudes in each CA1 neuron, suggesting that each neuron had different postsynaptic strengths at its excitatory and inhibitory synapses. Moreover, compared with untrained controls, IA-trained rats had higher mEPSC and mIPSC amplitudes, with broad diversity. These results suggested that contextual learning creates postsynaptic diversity in both excitatory and inhibitory synapses at each CA1 neuron.
AMPA or GABAA receptors seemed to mediate the postsynaptic currents, since bath treatment with CNQX or bicuculline blocked the mEPSC or mIPSC events, respectively. This technique can be used to study different types of learning in other regions, such as the sensory cortex and amygdala.

The slice patch clamp technique is an effective method for analyzing learning-induced changes in the intrinsic properties and plasticity of excitatory or inhibitory synapses.