Estamos agradecidos con el Departamento de Fisiología y Biología de Membrana de la Universidad de California Davis para que nos proporciona las instalaciones y un excelente ambiente para los estudios de los canales iónicos.

1. Preparación de los linfocitos T en reposo Humanos <ol><li> Utilizando RosetteSep Humanos Cocktail de enriquecimiento de células T y de densidad media RosetteSep, purificar las células T de muestras de sangre humana de acuerdo a las instrucciones del fabricante. La población de células resultante debe contener un 95% CD3 + células T en reposo. Purificamos linfocitos T humanos de sangre periférica recogida de voluntarios sanos, de acuerdo con el protocolo aprobado por el Consejo de la Universidad de California Davis Revisión Interna. </li><li> Tras el aislamiento, volver a suspender las células T en reposo, con una densidad de 0,5 x 10 6 células / ml en medio de cultivo celular que contiene medio RPMI-1640 con glutamina y HEPES, complementado con un 10% de suero fetal bovino, 2% Glutamax-I, 1% de RPMI 1640 vitamina solución, una solución de RPMI 1640% aminoácidos, 1% de piruvato de sodio, y el 0,03% β-mercaptoetanol. </li><li> Lugar la suspensión de células en frascos de cultivo celular y mantener a 37 ° C en una incubadora humidificada con 5% de CO 2 durante un máximo de 24 horas antes del experimento. </li></ol> 2. Preparación de la Cámara de grabación <ol><li> Los siguientes son necesarios para preparar las cámaras de grabación: a) el silicio O-rings, b) redondo, 25 mm cubreobjetos limpian con 70% de etanol y H 2 O destilada, c) las mezclas Sylgard184, preparados de acuerdo con las instrucciones del fabricante, yd) dos de 60 x 15 mm Petri. </li><li> Lugar ~ 0,5 ml de la mezcla en un plato Sylgard184 60x15 mm-Petri. </li><li> Sumerja la parte inferior de la junta tórica en la pinza Sylgard184 utilizando. </li><li> Eliminar el exceso de Sylgard184 desde el borde del anillo mediante la colocación de la junta tórica en una superficie limpia, como la otra placa de Petri. </li><li> Coloque el anillo en el cubreobjetos y se presiona. </li><li> Curar la Sylgard184 por las cámaras de calefacción a 85 ° C durante 40-60 minutos o hasta que el Sylgard184 se polimeriza. </li><li> Tienda de las cámaras de grabación en un recipiente de polvo. </li></ol> 3. Preparación de pipetas de parches <ol><li> En el día del experimento, tire pipetas con un diámetro de la punta ~ 2 micras utilizando capilares de vidrio y un extractor de la pipeta. Usamos tubos de borosilicato con filamento (OD: 1.5 mm y ID: 1,10 mm). Pipetas guárdela en un lugar libre de polvo. </li><li> Escudo puntas de pipeta con Sylgard184 o HIPEC revestimiento Semiconductor R6101 de protección, de acuerdo con un procedimiento estándar de 9. </li><li> Suavemente el fuego pulido las pipetas antes del experimento. </li></ol> 4. Preparación de la instalación de patch clamp <ol><li> Configurar y guardar las macros para la formación de gigaseal en el software que controla el amplificador de patch-clamp. Utilizamos un amplificador EPC-10 compatible con Windows XP y el software de pulso para la adquisición de datos. Hemos creado el &quot;set-up&quot;, &quot;en las células&quot;, y &quot;conjunto de células&quot; macros de acuerdo con el manual de EPC-10 y los utilizan para todos los experimentos. </li><li> Preparar soluciones para baños y soluciones de la pipeta en la Tabla 1 antes del experimento y almacenarlas a 4 º C durante un máximo de 1 semana. </li></ol><table border="1"><tbody><tr><td> Productos químicos </td><td colspan="3"> Soluciones de Bath (mM) </td><td> Pipeta Solutions (mM) </td></tr><tr><td></td><td> # 1 </td><td> # 2 </td><td> # 3 </td><td></td></tr></tbody><tbody><tr><td> CH 3 SO 3 Na </td><td> 130 </td><td> 110 </td><td> 125 </td><td> - </td></tr><tr><td> NaCl </td><td> 2 </td><td> 4 </td><td> 5 </td><td> - </td></tr><tr><td> Ca (OH) 2 </td><td> - </td><td> 20 </td><td> - </td><td> - </td></tr><tr><td> MgCl2 </td><td> 3 </td><td> 1 </td><td> - </td><td> 5 </td></tr><tr><td> MgSO4 </td><td> - </td><td> - </td><td> - </td><td> 2 </td></tr><tr><td> HEPES </td><td> 10 </td><td> 10 </td><td> 10 </td><td> 15 </td></tr><tr><td> HEDTA </td><td> - </td><td></td><td> 10 </td><td> - </td></tr><tr><td> EDTA </td><td> - </td><td> - </td><td> 1 </td><td> - </td></tr><tr><td> CS-Aspartate </td><td> - </td><td> - </td><td> - </td><td> 125 </td></tr><tr><td> BAPTA </td><td> - </td><td> - </td><td> - </td><td> 12 </td></tr><tr><td> Glucosa </td><td> 10 </td><td> 10 </td><td> 10 </td><td> - </td></tr></tbody></table> Tabla 1. Soluciones para el conjunto de células grabaciones actuales CARC. Todos los productos químicos fueron adquiridos de Sigma-Aldrich, St. Louis, MO. <ol start="3"><li> Determinar los potenciales de unión líquida (LJ) entre la solución de parche pipeta y cada solución del baño 10. Guardar el valor LJ en el lugar adecuado con el software de adquisición. Por ejemplo, en el programa Pulso insertar el valor en LJ &quot;LJ&quot; control en el &quot;amplificador&quot; de la ventana. </li><li> Configurar y guardar los protocolos de estimulación, como se muestra en la Figura 1, en el lugar apropiado de su software de adquisición antes de los experimentos. En el programa Pulso, se establece el protocolo de adquisición en la &quot;Generación de pulso&quot; de la ventana. <img src="/files/ftp_upload/2346/2364fig1.jpg" alt="Figura 1" /> Figura 1. . Protocolo de estimulación Rampa de tensión -120 a 100 mV de 50 ms de duración se aplica cada 0,2 a 2 s (5 a 0,5 Hz). Potencial de mantenimiento (V h) entre las rampas se mantiene en 30 mV. </li></ol> 5. Las células de montaje de la platina del microscopio <ol><li> Abrigo de las cámaras de grabación con poli-L-lisina de aproximadamente 20 minutos y lavar con H 2 O y luego con solución salina balanceada de Hank (HBSS) de las placas, las células. </li><li> Placa de 500 l de suspensión celular que contiene 0,2-0,5 x 10 6 células en la cámara y se incuba durante 20 min a 37 ° C. </li><li> Asegure la cámara de registro con las células colocadas en un soporte de cámara. Utilizamos un soporte de cámara a medida, donde la base soporta la parte inferior del cubreobjetos, mientras que la parte superior presiona suavemente contra la parte superior del cubreobjetos. </li><li> Coloque el soporte de la cámara en la platina del microscopio invertido. </li><li> Centrarse en las células de la luz transmitida. Nosotros usamos un objetivo de inmersión 40X. </li><li> Coloque el electrodo de referencia Ag / AgCl en el baño. </li><li> Alinear la multibarrel, impulsado por la gravedad del sistema de perfusión. La punta del tubo de entrada debe ser colocado en un ángulo de 45 ° hasta el fin cubreobjetos para el campo de visión. </li><li> Coloque el tubo de aspiración frente a la punta del tubo de entrada. </li><li> Perfusión de la cámara con tampón fosfato salino (PBS) o HBSS para lavar los medios de cultivo celular y las células unen libremente. </li></ol> 6. CRAC Canal Corrientes grabación <ol><li> Encontrar una célula que se une firmemente a la cubreobjetos. Por lo general selecciona uno de tamaño medio, de células redondas, con una superficie exterior lisa. </li><li> Perfusión de la cámara de grabación con Ca 2 + libre, 3 mM Mg 2 + que contienen solución del baño # 1 (Tabla 1) que contiene 0.5-1 thapsigargin M durante 8-10 minutos para bloquear la bomba SERCA. Llevamos a cabo este paso al agotamiento de la tienda antes de la formación gigaseal. </li><li> El uso de un microloader Eppendorf y un P-20 pipeta de desplazamiento, llenar el parche pipeta con la solución de la pipeta (Tabla 1) e insertar el parche pipeta en el soporte de la pipeta en el headstage amplificador. El Ca 2 + quelante BAPTA está incluido en la solución de la pipeta a agotar pasivamente la tienda y para reducir el Ca 2 +-dependiente de inactivación del canal CRAC. </li><li> Aplique una pequeña cantidad de presión positiva (~ 3 cm de H 2 O) en el interior de la pipeta parche antes de entrar en el baño. Usamos una medida de presión y absorber dispositivo conectado a las líneas de aire a presión y vacío para crear presión positiva o negativa dentro de un parche pipeta. </li><li> Baje el parche pipeta en el baño bajo control visual a través del microscopio. En el &quot;osciloscopio&quot; de la ventana, debería ver una corriente que fluye a través de la pipeta. Un cambio de ritmo-como en la amplitud de la corriente debe ser inducida por un pulso de amplitud pequeña tensión (prueba de impulso) aplicado por el macro preestablecido. En este momento, usted debería ser capaz de determinar el valor de la resistencia a la pipeta. La resistencia de nuestras pipetas es típicamente 6.5 mW. </li><li> Corregir el &quot;offset&quot; potencial generada entre la pipeta y el electrodo de referencia. </li><li> Bajo control visual a través del microscopio, llevar la punta de la pipeta cerca de la celda hasta que toque la membrana celular. </li><li> Aplique una presión negativa (20-40 pulgadas de H 2 O) en el interior del parche pipeta. </li><li> Monitorear la formación gigaseal en el &quot;osciloscopio&quot; de la ventana. Usted verá la amplitud de la corriente inducida por el pulso de prueba, disminuye y aumenta el valor de la resistencia a la pipeta&gt; 5 GΩ. Por lo general, las formas gigaseal dentro de una few segundos, pero se puede tomar 1-2 minutos para lograr una gigaseal estable. </li><li> Compensar la capacitancia pipeta (&quot;C-Fast&quot;). </li><li> Ruptura de la membrana de parches mediante la aplicación de presión negativa adicional usando una jeringa de 20 cc. </li><li> Establecer el potencial de retención de 30 mV. Un potencial de retención positiva ayuda a evitar que el calcio dependientes de inactivación del canal CRAC. </li><li> Compensar la capacitancia de la membrana de la célula (&quot;C-lento&quot;), guardar o registrar el valor de C-lento. </li><li> Comprobar el valor de la resistencia de acceso &quot;I s&quot;. En nuestros experimentos, la s R es típicamente 7.20 mW. </li><li> Empezar a aplicar una serie de rampas de tensión con una frecuencia de 0.5 Hz (Figura 1) en Ca 2 + libre de 3 mM Mg 2 + que contienen solución del baño # 1. En esta solución, la corriente CRAC es insignificante en comparación con la &quot;fuga&quot; de corriente debido a la poca permeabilidad de los canales CRAC para Mg 2 +. Guardar y utilizar las huellas corriente registró en la solución de baño # 1 en los futuros análisis de &quot;filtrar&quot; las corrientes. La amplitud absoluta de la &quot;fuga&quot; de corriente a -100 mV debe ser menor o igual a 5 PA. Si la célula es &quot;permeable&quot;, detener la prueba y comenzar de nuevo con una pipeta nueva revisión y una nueva célula. </li><li> Para grabar Ca 2 + a través de los canales actuales CRAC (I Ca-CRAC), sustitución de Ca 2 + libre de 3 mM Mg 2 + que contienen solución del baño # 1 mM Ca con 20 + 2 que contienen solución del baño # 2 (Tabla 1) al cambiar las válvulas de la perfusión. Esto permite Ca 2 + a fluir a través de los canales CRAC y producir una corriente hacia el interior. En el &quot;osciloscopio&quot; de la ventana, podrás ver el desarrollo del interior rectificar I Ca-CRAC (Figura 2). La amplitud de la corriente con tensiones negativas seguirá aumentando durante 1 min, debido a la Ca 2 +-dependiente de la potenciación de la CRAC actual. </li><li> Aumentar la frecuencia de la estimulación con rampas de tensión a 5 Hz, lo que es necesario registrar el transitorio corriente de Na + a través de los canales CRAC. </li><li> Vuelva a colocar el 20 mM Ca 2 + que contienen solución del baño # 2 con Na + que contienen cationes divalentes sin solución n º 3 (Tabla 1) al cambiar las válvulas de la perfusión. En el &quot;osciloscopio&quot; de la ventana, verás un desarrollo transitorio de mayor amplitud interior rectificar corriente de Na + a través de los canales CRAC (I Na-CRAC, Figura 2). </li><li> Se puede aplicar 20 mM Ca 2 + que contienen solución del baño # 2, que contiene 5.1 M La 3 + en el final del experimento para registrar la &quot;fuga&quot; de corriente. Sin embargo, no encontramos este enfoque es útil porque la &quot;fuga&quot; los cambios actuales en el tiempo y puede llegar a ser más grande o más pequeña al final del experimento. Nosotros preferimos aprovechar la corriente registrado en el Ca 2 + libre de 3 mM Mg 2 + que contienen solución del baño # 1 en el inicio del experimento como una &quot;fuga&quot; de corriente. </li><li> Guarde las huellas registradas en curso para su posterior análisis. </li></ol> 7. Análisis de Datos <ol><li> Recuperar los registros guardados en el actual software de análisis. Utilizamos el programa de impulso para el análisis. </li><li> Medir las amplitudes de corriente al principio y al final de las rampas de tensión. Nos suelen medir las amplitudes de corriente a -100 mV y 100 mV en el uso de pares de cursores en el &quot;osciloscopio&quot; de la ventana del programa de impulso. </li><li> Exportar los valores de las amplitudes de corriente en un programa gráfico para su posterior análisis y presentación gráfica. Usamos gráfica origen científico y software de análisis, la versión 7. </li></ol> 8. Resultados representante <img src="/files/ftp_upload/2346/2346fig2.jpg" alt="Figura 2" /> Figura 2. CRAC corrientes en una de las células T humanas en reposo. (A), Evolución temporal de las corrientes CRAC registrado en células enteras voltaje-clamp configuración a -100 mV (círculos llenos) y 100 mV (círculos). Antes de la formación gigaseal, el celular se incuba durante unos 10 min en Ca 2 + libre de 3 mM Mg 2 + que contienen solución del baño # 1 que contiene 0,5 M thapsigargin. Después del rodaje, las soluciones de baño se aplican de forma secuencial de la siguiente manera: Ca 2 + libre de 3 mM Mg 2 + que contienen solución del baño # 1 (0 + 3 Ca Mg), seguido de 20 mM Ca 2 + que contienen solución del baño # 2 (20 Ca), seguido por la solución de baño de cationes divalentes-free # 3 (DVF), seguido de solución del baño # 2 (20 Ca). La célula fue estimulado con una serie de rampas de tensión como se muestra en la Figura 1. La frecuencia de las rampas de los 5 Hz en la solución del baño # 3 (DVF) y 0,5 Hz en todas las otras soluciones. Tenga en cuenta el lento desarrollo de la I Ca-CRAC después de la aplicación de 20 mM Ca 2 + que contienen solución del baño # 2 y el rápido desarrollo transitoria de I Na-CRAC en la solución de baño de DVF N º 3. (B), Representante de las huellas actuales registrados en las rampas de tensión en Ca 2 + libre de 3 mM Mg 2 + que contienen solución del baño # 1 (&quot;Escape&quot;), 20 mM Ca 2 + que contienen solución del baño # 2 (20 Ca), y la solución de baño # 3 (DVF). (C, D), relaciones corriente-voltaje de la I Ca-CRAC (C) y Na-CRAC (D) restando &quot;fugas&quot; en curso de las corrientes registradas durante las rampas de tensión en 20 mM Ca 2 + que contienen baño solución # 2 (20 Ca), y la solución de baño # 3 (DVF) que se muestran en el panel (B).

<ol>
	<li>Parekh, A. B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Store-operated+CRAC+channels:+function+in+health+and+disease.">Store-operated CRAC channels: function in health and disease.</a> Nat Rev Drug Discov. 9, 399-410 (2010).</li><li>Feske, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=CRAC+channelopathies.">CRAC channelopathies.</a> Pflugers Arch. , (2010).</li><li>Lewis, R. S., Cahalan, . <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Mitogen-induced+oscillations+of+cytosolic+Ca2++and+transmembrane+Ca2++current+in+human+leukemic+T+cells.">Mitogen-induced oscillations of cytosolic Ca2+ and transmembrane Ca2+ current in human leukemic T cells.</a> Cell Regul. 1, 99-112 (1989).</li><li>Zweifach, A., Lewis, R. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Rapid+inactivation+of+depletion-activated+calcium+current+(ICRAC)+due+to+local+calcium+feedback.">Rapid inactivation of depletion-activated calcium current (ICRAC) due to local calcium feedback.</a> J Gen Physiol. 105, 209-226 (1995).</li><li>Zweifach, A., Lewis, R. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Slow+calcium-dependent+inactivation+of+depletion-activated+calcium+current.+Store-dependent+and+-independent+mechanisms.">Slow calcium-dependent inactivation of depletion-activated calcium current. Store-dependent and -independent mechanisms.</a> J Biol Chem. 270, 14445-1451 (1995).</li><li>Zweifach, A., Lewis, R. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Calcium-dependent+potentiation+of+store-operated+calcium+channels+in+T+lymphocytes.">Calcium-dependent potentiation of store-operated calcium channels in T lymphocytes.</a> J Gen Physiol. 107, 597-610 (1996).</li><li>Prakriya, M., Lewis, R. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Separation+and+characterization+of+currents+through+store-operated+CRAC+channels+and+Mg2+-inhibited+cation+(MIC)+channels.">Separation and characterization of currents through store-operated CRAC channels and Mg2+-inhibited cation (MIC) channels.</a> J Gen Physiol. 119, 487-507 (2002).</li><li>Fomina, A. F., Fanger, C. M., Kozak, J. A., Cahalan, . <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Single+channel+properties+and+regulated+expression+of+Ca(2+)+release-activated+Ca(2+)+(CRAC)+channels+in+human+T+cells.">Single channel properties and regulated expression of Ca(2+) release-activated Ca(2+) (CRAC) channels in human T cells.</a> J Cell Biol. 150, 1435-1444 (2000).</li><li>Sakmann, B., Neher, E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Single-Channel Recording. , (1995).</li><li>Neher, E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Correction+for+liquid+junction+potentials+in+patch+clamp+experiments.">Correction for liquid junction potentials in patch clamp experiments.</a> Methods Enzymol. 207, 123-1231 (1992).</li></ol>

La investigación electrofisiológica de las corrientes de CRAC en reposo las células T humanas es una tarea difícil debido a la amplitud endógeno CRAC actual en estas células es pequeño debido al tamaño de células pequeñas (el diámetro de descanso de células T humanas está en el rango de 5.8 micras). Aquí se presenta un procedimiento paso a paso para registrar de forma fiable las corrientes CRAC en el descanso los linfocitos T humanos aislados a partir de células mononucleares de sangre periférica. Esta t...

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		<th>Type</th>
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		<th>Catalogue Number</th>
		<th>Comment</th>
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<tr>
<td>RosetteSep Human T Cell Enrichment Cocktail</td>
<td>&nbsp;</td>
<td>StemCell Technologies, Vancouver, BC, Canada</td>
<td>15061</td>
<td>&nbsp;</td>
<tr>
<td>RosetteSep Density Medium</td>
<td>&nbsp;</td>
<td>StemCell Technologies </td>
<td>15705</td>
<td>&nbsp;</td>
<tr>
<td>RPMI-in 1640 medium w/glutamine/HEPES</td>
<td>&nbsp;</td>
<td>Fisher, Waltham, MA</td>
<td>SH3025501</td>
<td>&nbsp;</td>
<tr>
<td>Fetal Calf Serum</td>
<td>&nbsp;</td>
<td>Omega Scientific, Tarzana, CA</td>
<td>FB-01</td>
<td>&nbsp;</td>
<tr>
<td>GlutaMAX-I (100X solution)</td>
<td>&nbsp;</td>
<td>Invitrogen, Carlsbad, CA</td>
<td>35050</td>
<td>&nbsp;</td>
<tr>
<td>RPMI 1640 vitamin solution (100X)</td>
<td>&nbsp;</td>
<td>Sigma-Aldrich</td>
<td>7256</td>
<td>&nbsp;</td>
<tr>
<td>1640 amino acids solution (50X)</td>
<td>&nbsp;</td>
<td>Sigma-Aldrich</td>
<td>R7131</td>
<td>&nbsp;</td>
<tr>
<td>Sodium pyruvate</td>
<td>&nbsp;</td>
<td>Sigma-Aldrich</td>
<td>S8636</td>
<td>&nbsp;</td>
<tr>
<td>&beta;-Mercaptoethanol</td>
<td>&nbsp;</td>
<td>Sigma-Aldrich</td>
<td>M7522</td>
<td>&nbsp;</td>
<tr>
<td>Inositol trisphosphate</td>
<td>&nbsp;</td>
<td>Sigma-Aldrich</td>
<td>19766</td>
<td>&nbsp;</td>
<tr>
<td>BAPTA</td>
<td>&nbsp;</td>
<td>Sigma-Aldrich</td>
<td>A4926</td>
<td>&nbsp;</td>
<tr>
<td>Poly-L-Lysine Hydrobromide</td>
<td>&nbsp;</td>
<td>Sigma-Aldrich</td>
<td>P2636</td>
<td>&nbsp;</td>
<tr>
<td>Lanthanum Chloride</td>
<td>&nbsp;</td>
<td>Sigma-Aldrich</td>
<td>262072</td>
<td>&nbsp;</td>
<tr>
<td>Thapsigargin</td>
<td>&nbsp;</td>
<td>Calbiochem</td>
<td>586005</td>
<td>&nbsp;</td>
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<td>Sylgard 184 Silicon Elastomer Kit</td>
<td>&nbsp;</td>
<td>Dow Corning, Midland, MI</td>
<td>3097358-1004</td>
<td>&nbsp;</td>
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<td>HIPEC R6101 Semiconductor Protective Coating</td>
<td>&nbsp;</td>
<td>Dow Corning, Midland, MI</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<tr>
<td>63-500 Series High-Performance Vibration Isolation Lab Table</td>
<td>&nbsp;</td>
<td>Technical Manufacturing, Peabody, MA</td>
<td>63-540</td>
<td>&nbsp;</td>
<tr>
<td>EPC 10 patch clamp amplifier with headstage</td>
<td>&nbsp;</td>
<td>HEKA Instruments, Bellmore, NY</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<tr>
<td>Micromanipulator </td>
<td>&nbsp;</td>
<td>Sutter Instrument, Novato, CA </td>
<td>MP-285</td>
<td>&nbsp;</td>
<tr>
<td>Olympus 1X71 Inverted microscope with 40x oil immersion objective</td>
<td>&nbsp;</td>
<td>Olympus America, Center Valley, PA</td>
<td>1X71</td>
<td>&nbsp;</td>
<tr>
<td>Windows Computer</td>
<td>&nbsp;</td>
<td>Dell </td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<tr>
<td>Pulse software</td>
<td>&nbsp;</td>
<td>HEKA Instruments</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<tr>
<td>Origin Scientific Graphing and Analysis Software</td>
<td>&nbsp;</td>
<td>OriginLab, Northampton, MA</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<tr>
<td>Patch pipette puller</td>
<td>&nbsp;</td>
<td>Sutter Instrument</td>
<td>P-97</td>
<td>&nbsp;</td>
<tr>
<td>Borosilicate glass with filament (O.D.: 1.5mm and I.D.: 1.10mm</td>
<td>&nbsp;</td>
<td>Sutter Instrument</td>
<td>BF150-110-7.5</td>
<td>&nbsp;</td>
<tr>
<td>Narashige's Microforge</td>
<td>&nbsp;</td>
<td>Tritech Research, Los Angeles, CA</td>
<td>MF-830</td>
<td>&nbsp;</td>
<tr>
<td>Silicon O-rings </td>
<td>&nbsp;</td>
<td>McMASTER-CARR, Santa Fe Springs, CA</td>
<td>111 S70</td>
<td>&nbsp;</td>
<tr>
<td>Coverslips 25 mm </td>
<td>&nbsp;</td>
<td>Fisher Scientific </td>
<td>12-545-102 25 mm 25CIR.-1</td>
<td>&nbsp;</td>		</tbody>
		</table>
		</div>

whole-cell recording of calcium release-activated calcium (crac) currents in human t lymphocytes

In T lymphocytes, depletion of Ca2+ from the intracellular Ca2+ store leads to activation of plasmalemmal Ca2+ channels, called Calcium Release-Activated Calcium (CRAC) channels. CRAC channels play important role in regulation of T cell proliferation and gene expression. Abnormal CRAC channel function in T cells has been linked to severe combined immunodeficiency and autoimmune diseases 1, 2 . Studying CRAC channel function in human T cells may uncover new molecular mechanisms regulating normal immune responses and unravel the causes of related human diseases. Electrophysiological recordings of membrane currents provide the most accurate assessment of functional channel properties and their regulation. Electrophysiological assessment of CRAC channel currents in Jurkat T cells, a human leukemia T cell line, was first performed more than 20 years ago 3, however, CRAC current measurements in normal human T cells remains a challenging task. The difficulties in recording CRAC channel currents in normal T cells are compounded by the fact that blood-derived T lymphocytes are much smaller in size than Jurkat T cells and, therefore, the endogenous whole-cell CRAC currents are very low in amplitude. Here, we give a step-by-step procedure that we routinely use to record the Ca2+ or Na+ currents via CRAC channels in resting human T cells isolated from the peripheral blood of healthy volunteers. The method described here was adopted from the procedures used for recording the CRAC currents in Jurkat T cells and activated human T cells 4-8.

Watch this Scientific Journal Video about Whole-Cell Recording of Calcium Release-Activated Calcium CRAC Currents in Human T Lymphocytes at JoVE.com

Whole-Cell Recording of Calcium Release-Activated Calcium CRAC Currents in Human T Lymphocytes

Ofrecemos un protocolo de paso a paso para la grabación de células enteras patch clamp de calcio activados por calcio de lanzamiento (CRAC) las corrientes en mononucleares de sangre periférica de células derivadas de los linfocitos T humanos.

De células enteras de grabación de calcio activados por calcio de lanzamiento (CRAC) Las corrientes en los linfocitos T humanos

In T lymphocytes, depletion of Ca2+ from the intracellular Ca2+ store leads to activation of plasmalemmal Ca2+ channels, called Calcium Release-Activated ...

whole-cell-recording-calcium-release-activated-calcium-crac-currents

Research

JoVE Journal

Biology

Whole-Cell Recording of Calcium Release-Activated Calcium (CRAC) Currents in Human T Lymphocytes

16.0K vistas.  University of California, Davis. Ofrecemos un protocolo de paso a paso para la grabación de células enteras patch clamp de calcio activados por calcio de lanzamiento (CRAC) las corrientes en mononucleares de sangre periférica de células derivadas de los linfocitos T humanos.

Video: Whole-Cell Recording of Calcium Release-Activated Calcium CRAC Currents in Human T Lymphocytes

Loading Drosophila Nerve Terminals with Calcium Indicators

Calcium plays many roles in the nervous system but none more impressive than as the trigger for neurotransmitter release, and none more profound than as the messenger essential for the synaptic plasticity that supports learning and memory. To further elucidate the molecular underpinnings of Ca2+-dependent synaptic mechanisms, a model system is required that is both genetically malleable and physiologically accessible. Drosophila melanogaster provides such a model. In this system, genetically-encoded fluorescent indicators are available to detect Ca2+ changes in nerve terminals. However, these indicators have limited sensitivity to Ca2+ and often show a non-linear response. Synthetic fluorescent indicators are better suited for measuring the rapid Ca2+ changes associated with nerve activity. Here we demonstrate a technique for loading dextran-conjugated synthetic Ca2+ indicators into live nerve terminals in Drosophila larvae. Particular emphasis is placed on those aspects of the protocol most critical to the technique's success, such as how to avoid static electricity discharges along the isolated nerves, maintaining the health of the preparation during extended loading periods, and ensuring axon survival by providing Ca2+ to promote sealing of severed axon endings. Low affinity dextran-conjugated Ca2+-indicators, such as fluo-4 and rhod, are available which show a high signal-to-noise ratio while minimally disrupting presynaptic Ca2+ dynamics. Dextran-conjugation helps prevent Ca2+ indicators being sequestered into organelles such as mitochondria. The loading technique can be applied equally to larvae, embryos and adults.

Calcium is a ubiquitous messenger in the nervous system, essential for triggering neurotransmitter release and changes in synaptic strength. Here we demonstrate a technique for loading Ca2+-indicators into Drosophila nerve terminals. We also demonstrate fabrication of the required apparatus and emphasize points critical for the technique's success.

Cargando Drosophila terminales nerviosas con indicadores de calcio

Calcium plays many roles in the nervous system but none more impressive than as the trigger for neurotransmitter release, and none more profound than as ...

Investigación

Biología

In Vivo 2-Photon Calcium Imaging in Layer 2/3 of Mice

To understand network dynamics of microcircuits in the neocortex, it is essential to simultaneously record the activity of a large number of neurons . In-vivo two-photon calcium imaging is the only method that allows one to record the activity of a dense neuronal population with single-cell resolution . The method consists in implanting a cranial imaging window, injecting a fluorescent calcium indicator dye that can be taken up by large numbers of neurons and finally recording the activity of neurons with time lapse calcium imaging using an in-vivo two photon microscope. Co-injection of astrocyte-specific dyes allows one to differentiate neurons from astrocytes. The technique can be performed in mice expressing fluorescent molecules in specific subpopulations of neurons to better understand the network interactions of different groups of cells.

To understand network dynamics of microcircuits in the neocortex, it is essential to simultaneously record the activity of a large number of neurons . In-vivo two-photon calcium imaging is the only method that allows one to record the activity of a dense neuronal population with single-cell resolution .

En Vivo 2-Photon imágenes de calcio en la capa 2 / 3 de los ratones

To understand network dynamics of microcircuits in the neocortex, it is essential to simultaneously record the activity of a large number of neurons .  ...

Whole-cell Recordings of Light Evoked Excitatory Synaptic Currents in the Retinal Slice

We use the whole-cell patch clamp technique to study the synaptic circuitry that underlies visual information processing in the retina. In this video, we will guide you through the process of performing whole-cell recordings of light evoked currents of individual cells in the retinal slice preparation. We use the aquatic tiger salamander as an animal model. We begin by describing the dissection of the eye and show how slices are mounted for electrophysiological recordings. Once the slice is placed in the recording chamber, we demonstrate how to perform whole-cell voltage clamp recordings. We then project visual stimuli onto the photoreceptors in the slice to elicit light-evoked current responses. During the recording we perfuse the slice with pharmacological agents, whereby an 8-channel perfusion system allows us to quickly switch between different agents.  The retinal slice preparation is widely used for patch clamp recordings in the retina, in particular to study amacrine or bipolar cells, which are not accessible in a whole-mount preparation.

This video shows the process of whole-cell voltage clamp recordings in the retinal slice of the aquatic tiger salamander. We demonstrate the preparation of the slice as well as how to perform patch clamp recordings during visual stimulation of the retina.

De células enteras grabaciones de la Luz evocados excitatorios de las corrientes sinápticas en la porción de retina

We use the whole-cell patch clamp technique to study the synaptic circuitry that underlies visual information processing in the retina. In this video, we ...

Measuring Near Plasma Membrane and Global Intracellular Calcium Dynamics in Astrocytes

The brain contains glial cells.  Astrocytes, a type of glial cell, have long been known to provide a passive supportive role to neurons. However, increasing evidence suggests that astrocytes may also actively participate in brain function through functional interactions with neurons.  However, many fundamental aspects of astrocyte biology remain controversial, unclear and/or experimentally unexplored. One important issue is the dynamics of intracellular calcium transients in astrocytes. This is relevant because calcium is well established as an important second messenger and because it has been proposed that astrocyte calcium elevations can trigger the release of transmitters from astrocytes. However, there has not been any detailed or satisfying description of near plasma membrane calcium signaling in astrocytes. Total internal reflection fluorescence (TIRF) microscopy is a powerful tool to analyze physiologically relevant signaling events within about 100 nm of the plasma membrane of live cells. Here, we use TIRF microscopy and describe how to monitor near plasma membrane and global intracellular calcium dynamics almost simultaneously. The further refinement and systematic application of this approach has the potential to inform about the precise details of astrocyte calcium signaling. A detailed understanding of astrocyte calcium dynamics may provide a basis to understand if, how, when and why astrocytes and neurons undergo calcium-dependent functional interactions.

We describe how to measure near membrane and global intracellular calcium dynamics in cultured astrocytes using total internal reflection and epifluorescence microscopy.

Cerca de la medición de la membrana plasmática y Global Dynamics calcio intracelular en los astrocitos

The brain contains glial cells.  Astrocytes, a type of glial cell, have long been known to provide a passive supportive role to neurons. However, ...

Monitoring Dynamic Changes In Mitochondrial Calcium Levels During Apoptosis Using A Genetically Encoded Calcium Sensor

Dynamic changes in intracellular calcium concentration in response to various stimuli regulates many cellular processes such as proliferation, differentiation, and apoptosis1. During apoptosis, calcium accumulation in mitochondria promotes the release of pro-apoptotic factors from the mitochondria into the cytosol2. It is therefore of interest to directly measure mitochondrial calcium in living cells in situ during apoptosis. High-resolution fluorescent imaging of cells loaded with dual-excitation ratiometric and non-ratiometric synthetic calcium indicator dyes has been proven to be a reliable and versatile tool to study various aspects of intracellular calcium signaling. Measuring cytosolic calcium fluxes using these techniques is relatively straightforward. However, measuring intramitochondrial calcium levels in intact cells using synthetic calcium indicators such as rhod-2 and rhod-FF is more challenging. Synthetic indicators targeted to mitochondria have blunted responses to repetitive increases in mitochondrial calcium, and disrupt mitochondrial morphology3. Additionally, synthetic indicators tend to leak out of mitochondria over several hours which makes them unsuitable for long-term experiments. Thus, genetically encoded calcium indicators based upon green fluorescent protein (GFP)4 or aequorin5 targeted to mitochondria have greatly facilitated measurement of mitochondrial calcium dynamics. Here, we describe a simple method for real-time measurement of mitochondrial calcium fluxes in response to different stimuli. The method is based on fluorescence microscopy of 'ratiometric-pericam' which is selectively targeted to mitochondria. Ratiometric pericam is a calcium indicator based on a fusion of circularly permuted yellow fluorescent protein and calmodulin4. Binding of calcium to ratiometric pericam causes a shift of its excitation peak from 415 nm to 494 nm, while the emission spectrum, which peaks around 515 nm, remains unchanged. Ratiometric pericam binds a single calcium ion with a dissociation constant in vitro of ~1.7 &mu;M4. These properties of ratiometric pericam allow the quantification of rapid and long-term changes in mitochondrial calcium concentration. Furthermore, we describe adaptation of this methodology to a standard wide-field calcium imaging microscope with commonly available filter sets. Using two distinct agonists, the purinergic agonist ATP and apoptosis-inducing drug staurosporine, we demonstrate that this method is appropriate for monitoring changes in mitochondrial calcium concentration with a temporal resolution of seconds to hours. Furthermore, we also demonstrate that ratiometric pericam is also useful for measuring mitochondrial fission/fragmentation during apoptosis. Thus, ratiometric pericam is particularly well suited for continuous long-term measurement of mitochondrial calcium dynamics during apoptosis.

This protocol describes a method for real-time measurement of mitochondrial calcium fluxes by fluorescent imaging. The method takes advantage of a circularly permutated YFP-based dual-excitation ratiometric calcium sensor (ratiometric pericam-mt) selectively expressed in mitochondria.

Seguimiento de los cambios dinámicos en niveles de calcio mitocondrial de la apoptosis mediante un sensor de calcio genéticamente codificados

Dynamic changes in intracellular calcium concentration in response to various stimuli regulates many cellular processes such as proliferation, ...

Measuring Fast Calcium Fluxes in Cardiomyocytes

Cardiomyocytes have multiple Ca2+ fluxes of varying duration that work together to optimize function 1,2. Changes in Ca2+ activity in response to extracellular agents is predominantly regulated by the phospholipase C&beta;- G&alpha;q pathway localized on the plasma membrane which is stimulated by agents such as acetylcholine 3,4. We have recently found that plasma membrane protein domains called caveolae5,6 can entrap activated G&alpha;q 7. This entrapment has the effect of stabilizing the activated state of G&alpha;q and resulting in prolonged Ca2+ signals in cardiomyocytes and other cell types8. We uncovered this surprising result by measuring dynamic calcium responses on a fast scale in living cardiomyocytes. Briefly, cells are loaded with a fluorescent Ca2+ indicator. In our studies, we used Ca2+ Green (Invitrogen, Inc.) which exhibits an increase in fluorescence emission intensity upon binding of calcium ions. The fluorescence intensity is then recorded for using a line-scan mode of a laser scanning confocal microscope. This method allows rapid acquisition of the time course of fluorescence intensity in pixels along a selected line, producing several hundreds of time traces on the microsecond time scale. These very fast traces are transferred into excel and then into Sigmaplot for analysis, and are compared to traces obtained for electronic noise, free dye, and other controls. To dissect Ca2+ responses of different flux rates, we performed a histogram analysis that binned pixel intensities with time. Binning allows us to group over 500 traces of scans and visualize the compiled results spatially and temporally on a single plot. Thus, the slow Ca2+ waves that are difficult to discern when the scans are overlaid due to different peak placement and noise, can be readily seen in the binned histograms. Very fast fluxes in the time scale of the measurement show a narrow distribution of intensities in the very short time bins whereas longer Ca2+ waves show binned data with a broad distribution over longer time bins. These different time distributions allow us to dissect the timing of Ca2+fluxes in the cells, and to determine their impact on various cellular events.

We present a method to isolate rapid (microsecond) calcium events from slower fluxes in living cells using laser scanning confocal microscopy. The method measures fluorescence intensity fluctuations of calcium indicators by recording line scans of several hundred pixels in a cell. Histogram analysis allows us to isolate the time scales of different calcium fluxes.

La medición de flujos rápidos de calcio en los cardiomiocitos

Cardiomyocytes have multiple Ca2+ fluxes of varying duration that work together to optimize function 1,2. Changes in Ca2+ activity in response to ...

Assessment of Calcium Sparks in Intact Skeletal Muscle Fibers

Maintaining homeostatic Ca2+ signaling is a fundamental physiological process in living cells. Ca2+ sparks are the elementary units of Ca2+ signaling in the striated muscle fibers that appear as highly localized Ca2+ release events mediated by ryanodine receptor (RyR) Ca2+ release channels on the sarcoplasmic reticulum (SR) membrane. Proper assessment of muscle Ca2+ sparks could provide information on the intracellular Ca2+&#160;handling properties of healthy and diseased striated muscles. Although Ca2+ sparks events are commonly seen in resting cardiomyocytes, they are rarely observed in resting skeletal muscle fibers; thus there is a need for methods to generate and analyze sparks in skeletal muscle fibers.
Detailed here is an experimental protocol for measuring Ca2+ sparks in isolated flexor digitorm brevis (FDB) muscle fibers using fluorescent Ca2+ indictors and laser scanning confocal microscopy. In this approach, isolated FDB fibers are exposed to transient hypoosmotic stress followed by a return to isotonic physiological solution. Under these conditions, a robust Ca2+ sparks response is detected adjacent to the sarcolemmal membrane in young healthy FDB muscle fibers. Altered Ca2+ sparks response is detected in dystrophic or aged skeletal muscle fibers. This approach has recently demonstrated that membrane-delimited signaling involving cross-talk between inositol (1,4,5)-triphosphate receptor (IP3R) and RyR contributes to Ca2+ spark activation in skeletal muscle. In summary, our studies using osmotic stress induced Ca2+ sparks showed that this intracellular response reflects a muscle signaling mechanism in physiology and aging/disease states, including mouse models of muscle dystrophy (mdx mice) or amyotrophic lateral sclerosis (ALS model).

Described here is a method to directly measure calcium sparks, the elementary units of Ca2+ release from sarcoplasmic reticulum&#160;in intact skeletal muscle fibers. This method utilizes osmotic-stress-mediated triggering of Ca2+ release from ryanodine receptor in isolated muscle fibers. The dynamics and homeostatic capacity of intracellular Ca2+ signaling can be employed to assess muscle function in health and disease.

Evaluación de calcio en Sparks intactas las fibras musculares esqueléticas

Maintaining homeostatic Ca2+ signaling is a fundamental physiological process in living cells. Ca2+ sparks are the elementary units of Ca2+ signaling in ...

Cytosolic Calcium Measurements in Renal Epithelial Cells by Flow Cytometry

A variety of cellular processes, both physiological and pathophysiological, require or are governed by calcium, including exocytosis, mitochondrial function, cell death, cell metabolism and cell migration to name but a few. Cytosolic calcium is normally maintained at low nanomolar concentrations; rather it is found in high micromolar to millimolar concentrations in the endoplasmic reticulum, mitochondrial matrix and the extracellular compartment. Upon stimulation, a transient increase in cytosolic calcium serves to signal downstream events. Detecting changes in cytosolic calcium is normally performed using a live cell imaging set up with calcium binding dyes that exhibit either an increase in fluorescence intensity or a shift in the emission wavelength upon calcium binding. However, a live cell imaging set up is not freely accessible to all researchers. Alternative detection methods have been optimized for immunological cells with flow cytometry and for non-immunological adherent cells with a fluorescence microplate reader. Here, we describe an optimized, simple method for detecting changes in epithelial cells with flow cytometry using a single wavelength calcium binding dye. Adherent renal proximal tubule epithelial cells, which are normally difficult to load with dyes, were loaded with a fluorescent cell permeable calcium binding dye in the presence of probenecid, brought into suspension and calcium signals were monitored before and after addition of thapsigargin, tunicamycin and ionomycin.

Calcium is involved in numerous physiological and pathophysiological signaling pathways. Live cell imaging requires specialized equipment and can be time consuming. A quick, simple method using a flow cytometer to determine relative changes in cytosolic calcium in adherent epithelial cells brought into suspension was optimized.

Las mediciones del calcio citosólico en las células epiteliales renales mediante citometría de flujo

A variety of cellular processes, both physiological and pathophysiological, require or are governed by calcium, including exocytosis, mitochondrial ...

Real-time In Vivo Recording of Arabidopsis Calcium Signals During Insect Feeding Using a Fluorescent Biosensor

Calcium ions are predicted to be key signaling entities during biotic interactions, with calcium signaling forming an established part of the plant defense response to microbial elicitors and to wounding caused by chewing insects, eliciting systemic calcium signals in plants. However, the role of calcium in vivo during biotic stress is still unclear. This protocol describes the use of a genetically-encoded calcium sensor to detect calcium signals in plants during feeding by a hemipteran pest. Hemipterans such as aphids pierce a small number of cells with specialized, elongated sucking mouthparts, making them the ideal tool to study calcium dynamics when a plant is faced with a biotic stress, which is distinct from a wounding response. In addition, fluorescent biosensors are revolutionizing the measurement of signaling molecules in vivo in both animals and plants. Expressing a GFP-based calcium biosensor, GCaMP3, in the model plant Arabidopsis thaliana allows for the real-time imaging of plant calcium dynamics during insect feeding, with a high spatial and temporal resolution. A repeatable and robust assay has been developed using the fluorescence microscopy of detached GCaMP3 leaves, allowing for the continuous measurement of cytosolic calcium dynamics before, during, and after insect feeding. This reveals a highly-localized rapid calcium elevation around the aphid feeding site that occurs within a few minutes. The protocol can be adapted to other biotic stresses, such as additional insect species, while the use of Arabidopsis thaliana allows for the rapid generation of mutants to facilitate the molecular analysis of the phenomenon.

Real-time In Vivo Recording of Arabidopsis Calcium Signals During Insect Feeding Using a Fluorescent Biosensor

This protocol outlines a simple method for analyzing calcium signals in plants generated by feeding hemipteran insects, such as aphids. Arabidopsis thaliana transformed with the GFP calcium biosensor GCaMP3 allow for the real-time in vivo imaging of calcium dynamics with a high temporal and spatial resolution.

En tiempo real en Vivo la grabación de señales de calcio Arabidopsis durante la alimentación de insectos utilizando un Biosensor fluorescente

Calcium ions are predicted to be key signaling entities during biotic interactions, with calcium signaling forming an established part of the plant ...

Imaging Calcium Dynamics in Subpopulations of Mouse Pancreatic Islet Cells

Pancreatic islet hormones regulate blood glucose homeostasis. Changes in blood glucose induce oscillations of cytosolic calcium in pancreatic islet cells that trigger secretion of three main hormones: insulin (from &#946;-cells), glucagon (&#945;-cells) and somatostatin (&#948;-cells). &#946;-Cells, which make up the majority of islet cells and are electrically coupled to each other, respond to the glucose stimulus as one single entity. The excitability of the minor subpopulations, &#945;-cells and &#948;-cells (making up around 20% (30%) and 4% (10%) of the total rodent1 (human2) islet cell numbers, respectively) is less predictable and is therefore of special interest.
Calcium sensors are delivered into the peripheral layer of cells within the isolated islet. The islet or a group of islets is then immobilized and imaged using a fluorescence microscope. The choice of the imaging mode is between higher throughput (wide-field) and better spatial resolution (confocal). Conventionally, laser scanning confocal microscopy is used for imaging tissue, as it provides the best separation of the signal between the neighboring cells. A wide-field system can be utilized too, if the contaminating signal from the dominating population of &#946;-cells is minimized.
Once calcium dynamics in response to specific stimuli have been recorded, data are expressed in numerical form as fluorescence intensity vs. time, normalized to the initial fluorescence and baseline-corrected, to remove the effects linked to bleaching of the fluorophore. Changes in the spike frequency or partial area under the curve (pAUC) are computed vs. time, to quantify the observed effects. pAUC is more sensitive and quite robust whereas spiking frequency provides more information on the mechanism of calcium increase.
Minor cell subpopulations can be identified using functional responses to marker compounds, such as adrenaline and ghrelin, that induce changes in cytosolic calcium in a specific populations of islet cells.

Here, we present a protocol for imaging and quantifying calcium dynamics in heterogeneous cell populations, such as pancreatic islet cells. Fluorescent reporters are delivered into the peripheral layer of cells within the islet, which is then immobilized and imaged, and per-cell analysis of the dynamics of fluorescence intensity is performed.

Imagen de la dinámica del calcio en subpoblaciones de células de islotes pancreáticos de ratón

Pancreatic islet hormones regulate blood glucose homeostasis. Changes in blood glucose induce oscillations of cytosolic calcium in pancreatic islet cells ...