Nous sommes reconnaissants envers le Département de physiologie et biologie membranaire, Université de Californie à Davis pour nous fournir des installations et un excellent environnement pour les études des canaux ioniques.

1. Préparation de repos lymphocytes T humains <ol><li> Utiliser RosetteSep homme T Cocktail enrichissement cellulaire et de densité moyenne RosetteSep, purifier les cellules T à partir d&#39;échantillons de sang humain selon les instructions du fabricant. La population cellulaire obtenue doit contenir 95% cellules T CD3 + repos. Nous purifions lymphocytes T humains à partir d&#39;échantillons de sang périphérique prélevés sur des volontaires sains en conformité avec le protocole approuvé par le Conseil d&#39;examen interne UC Davis. </li><li> Après isolement, resuspendre lymphocytes T à une densité de 0,5 x 10 6 cellules / mL dans le milieu de culture cellulaire contenant RPMI-1640 avec de la glutamine et l&#39;HEPES, complété avec 10% sérum de veau foetal, de 2% glutamax-I, 1% du RPMI 1640 la vitamine solution, une solution de RPMI 1640% d&#39;acides aminés, le pyruvate de sodium à 1%, et 0,03% β-mercaptoéthanol. </li><li> Suspension cellulaire place dans des flacons de culture cellulaire et de maintenir à 37 ° C dans un incubateur humidifié avec 5% de CO 2 jusqu&#39;à 24 h avant l&#39;expérience. </li></ol> 2. Préparation de la chambre d&#39;enregistrement <ol><li> Les éléments suivants sont nécessaires pour préparer des chambres d&#39;enregistrement: a) de silicium joints toriques, b) rond, 25 mm lamelles nettoyé avec de l&#39;éthanol 70% et distillé H 2 O, c) mélangé Sylgard184, préparé selon les instructions du fabricant, et d) deux 60 x 15 mm boîte de Pétri. </li><li> Placez ~ 0,5 ml de Sylgard184 mélangé dans un plat 60x15 mm Petri. </li><li> Trempez la partie inférieure de l&#39;O-ring dans la pince Sylgard184 aide. </li><li> Retirer l&#39;excès de Sylgard184 le bord de la O-ring en plaçant le joint torique sur une surface propre, comme une autre boîte de Pétri. </li><li> Placez l&#39;anneau sur la lamelle et appuyez vers le bas. </li><li> Guérissez les Sylgard184 par le chauffage des chambres à 85 ° C pendant 40-60 min ou jusqu&#39;à ce que le Sylgard184 est polymérisé. </li><li> Boutique des chambres d&#39;enregistrement dans un conteneur sans poussière. </li></ol> 3. Préparation de pipettes Patch <ol><li> Le jour de l&#39;expérience, tirez les pipettes avec un diamètre de l&#39;extrémité ~ 2-um en utilisant capillaires en verre et un extracteur de pipette. Nous utilisons des tubes en borosilicate avec filament (OD: 1,5 mm et ID: 1,10 mm). Pipettes Stocker dans un endroit exempt de poussière. </li><li> Manteau pipette avec Sylgard184 ou revêtement CHIP R6101 Semiconductor protection, selon une procédure standard 9. </li><li> Doucement feu polir les pipettes avant l&#39;expérience. </li></ol> 4. Préparation de configuration patch clamp <ol><li> Configurer et sauvegarder les macros pour la formation gigaseal dans le logiciel de contrôle de votre amplificateur de patch-clamp. Nous utilisons un amplificateur EPC-10 pris en charge par Windows XP et le logiciel d&#39;impulsions pour l&#39;acquisition de données. Nous avons fixé le &quot;set-up&quot;, &quot;sur la cellule&quot;, et &quot;cellule entière&quot; macros selon le manuel d&#39;EPC-10 et les utiliser pour toutes les expériences. </li><li> Préparer des solutions de bain et des solutions pipette énumérés dans le tableau 1 avant l&#39;expérience et les stocker à 4 ° C pendant 1 semaine. </li></ol><table border="1"><tbody><tr><td> Produits chimiques </td><td colspan="3"> Bain Solutions (mM) </td><td> Solutions pipette (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> MgCl 2 </td><td> 3 </td><td> 1 </td><td> - </td><td> 5 </td></tr><tr><td> MgSO 4 </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-unespartate </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> Le glucose </td><td> 10 </td><td> 10 </td><td> 10 </td><td> - </td></tr></tbody></table> Tableau 1. Solutions pour la cellule entière CRAC enregistrements actuels. Tous les produits chimiques ont été achetés chez Sigma-Aldrich, St. Louis, MO. <ol start="3"><li> Déterminer les potentiels de jonction liquide (LJ) entre la solution de la pipette de patch et chaque solution 10 de bain. Enregistrer la valeur LJ à l&#39;endroit approprié à l&#39;aide de votre logiciel d&#39;acquisition. Par exemple, dans le programme d&#39;impulsion insérer la valeur LJ en «LJ» de contrôle dans la &quot;amplificateur&quot; fenêtre. </li><li> Configurer et sauvegarder les protocoles de stimulation, comme le montre la figure 1, à l&#39;endroit approprié de votre logiciel d&#39;acquisition, avant les expériences. Dans le programme d&#39;impulsion, nous avons mis le protocole d&#39;acquisition de la «génération d&#39;impulsion&quot; fenêtre. <img src="/files/ftp_upload/2346/2364fig1.jpg" alt="Figure 1" /> Figure 1. . Protocole de stimulation rampe de tension de -120 à 100 mV de 50 ms dans la durée est appliqué toutes les 0,2 - 2 s (5 - 0,5 Hz). Potentiel de maintien (V h) entre les rampes est maintenue à 30 mV. </li></ol> 5. Cellules de montage sur la platine du microscope <ol><li> Enduire les chambres d&#39;enregistrement avec la poly-L-lysine pour ~ 20 min et laver avec du H 2 O puis avec une solution saline équilibrée de Hank (HBSS) avant l&#39;étalement des cellules. </li><li> Plate 500 pl de suspension cellulaire contenant 0.2 au 0.5 x10 6 cellules dans la chambre et incuber pendant 20 min à 37 ° C. </li><li> Fixez la chambre d&#39;enregistrement avec les cellules attachées à un support de chambre. Nous utilisons un porte-chambre de mesure, où la base supporte le bas de la lamelle, tandis que la partie supérieure légèrement en appui sur le haut de la lamelle. </li><li> Placez le porte-chambre sur la scène du microscope inversé. </li><li> Focus sur les cellules en lumière transmise. Nous utilisons un objectif 40X d&#39;huile d&#39;immersion. </li><li> Placer l&#39;électrode de référence Ag / AgCl dans le bain. </li><li> Alignez les multibarrel, système de perfusion par gravité. La pointe du tube d&#39;introduction devrait être placé à un angle de 45 ° à l&#39;étroite lamelle au champ de vue. </li><li> Placez le tuyau d&#39;aspiration en face de l&#39;extrémité du tube entrées. </li><li> Perfuser la chambre avec un tampon phosphate salin (PBS) ou HBSS pour laver les milieux de culture cellulaire et les cellules lâchement. </li></ol> 6. CRAC canaliser les courants d&#39;enregistrement <ol><li> Trouver une cellule qui est fermement attaché à la lamelle. Nous choisissent généralement une taille moyenne, de cellules rondes avec une surface extérieure lisse. </li><li> Perfuser la chambre d&#39;enregistrement de Ca 2 + libre, 3 mM de Mg 2 +-solution du bain contenant # 1 (Tableau 1) contenant de 0,5 à 1 thapsigargine pM pour 8-10 min pour bloquer la pompe SERCA. Nous effectuons cette démarche pour épuiser le magasin avant la formation gigaseal. </li><li> En utilisant une microloader Eppendorf et une pipette P-20 de déplacement, remplir la pipette de patch avec la solution pipette (tableau 1) et insérez la pipette de patch dans le porte-pipette sur la headstage amplificateur. Le Ca 2 + chélateur BAPTA est inclus dans la solution passive pipette pour épuiser les stocker et de réduire de Ca 2 +-dépendante l&#39;inactivation du canal CRAC. </li><li> Appliquez une petite quantité de pression positive (~ 3 pouces de H 2 O) à l&#39;intérieur de la pipette de patch avant d&#39;entrer dans le bain. Nous utilisons une sur-mesure à la pression d&#39;aspiration périphérique connecté à l&#39;air comprimé et les lignes de vide pour créer une pression positive ou négative à l&#39;intérieur d&#39;une pipette de patch. </li><li> Basse de la pipette de patch dans le bain sous contrôle visuel via le microscope. Dans le &quot;Oscilloscope&quot; fenêtre, vous devriez voir un courant circulant dans la pipette. Un changement étape comme dans l&#39;amplitude du courant doit être induite par une impulsion de tension de faible amplitude (de test par impulsion) appliquée par la macro prédéfinie. À ce moment, vous devriez être en mesure de déterminer la valeur de la résistance pipette. La résistance de nos pipettes est généralement 5-6 MQ. </li><li> Corriger les «compenser» le potentiel généré entre la pipette et l&#39;électrode de référence. </li><li> Sous contrôle visuel par le biais du microscope, amener la pointe de la pipette rapprocher de la cellule jusqu&#39;à ce qu&#39;elle touche la membrane cellulaire. </li><li> Appliquer une pression négative (20-40 pouces H 2 O) à l&#39;intérieur de la pipette de patch. </li><li> Surveiller la formation gigaseal dans le &quot;oscilloscope&quot; fenêtre. Vous verrez l&#39;amplitude du courant, induite par l&#39;impulsion d&#39;essai, diminue et la valeur de résistance augmente pipette&gt; 5 GQ. Habituellement gigaseal formes au sein d&#39;une few secondes, mais cela peut prendre 1-2 min pour obtenir un gigaseal stable. </li><li> Compenser la capacitance pipette («C-Fast&quot;). </li><li> Pause de la membrane de patch en appliquant une pression négative supplémentaires en utilisant une seringue de 20 cc. </li><li> Réglez le potentiel de maintien à +30 mV. Un potentiel de détention positif aide à prévenir les calcium-dépendante l&#39;inactivation du canal CRAC. </li><li> Compenser la capacitance membranaire de la cellule («C-Lent»); enregistrer ou enregistrer la valeur de C-lent. </li><li> Vérifiez la valeur de la résistance d&#39;accès &quot;R s&quot;. Dans nos expériences, le s R est généralement de 7 à 20 MQ. </li><li> Commencez à appliquer une série de rampes de tension avec une fréquence de 0,5 Hz (figure 1) dans le Ca 2 + libre 3 mM Mg 2 +-solution du bain contenant # 1. Dans cette solution, le courant CRAC est négligeable par rapport à la &quot;fuite&quot; en cours en raison de la faible perméabilité des canaux CRAC pour Mg 2 +. Enregistrer et utiliser les traces actuelles enregistrées dans la solution de bain n ° 1 dans les analyses futures comme «fuite» des courants. L&#39;amplitude absolue de la &quot;fuite&quot; courant à 100 mV doit être inférieure ou égale à 5 pA. Si la cellule est &quot;fuite&quot;, arrêter l&#39;expérience et de recommencer à l&#39;aide d&#39;une pipette nouveau patch et une nouvelle cellule. </li><li> Pour enregistrer Ca 2 + via les canaux CRAC (I Ca-CRAC), remplacer Ca 2 + libre 3 mM Mg 2 +-solution du bain contenant n ° 1 avec 20 mM de 2 Ca solution de bain + contenant # 2 (Tableau 1) par commutation des vannes de perfusion. Cela permet de Ca 2 + au flux via les canaux CRAC et de produire un courant entrant. Dans le &quot;Oscilloscope&quot; fenêtre, vous verrez le développement de rectifier intérieurement I Ca-CRAC (figure 2). L&#39;amplitude du courant à des tensions négatives va continuer à augmenter pendant environ 1 min en raison de la Ca 2 +-dépendante de potentialisation de l&#39;actuelle CRAC. </li><li> Augmenter la fréquence de stimulation avec des rampes de tension à 5 Hz, ce qui est nécessaire pour enregistrer le transitoire courant Na + via les canaux CRAC. </li><li> Remplacer le 20 mM de Ca 2 +-n ° 2 contenant une solution de bain avec Na +-cations divalents contenant une solution sans # 3 (tableau 1) par commutation des vannes de perfusion. Dans le &quot;Oscilloscope&quot; fenêtre, vous verrez un développement transitoires de plus grande amplitude intérieurement rectifier courant Na + via les canaux CRAC (I Na-CRAC; figure 2). </li><li> On peut appliquer 20 mm 2 Ca solution de bain + contenant n ° 2 contenant de 1 à 5 uM La 3 + à la fin de l&#39;expérience d&#39;enregistrer la «fuite» actuel. Cependant, nous n&#39;avons pas trouvé cette approche utile, car la «fuite» des changements en cours dans le temps et il peut devenir plus ou moins à la fin de l&#39;expérience. Nous préférons prendre le courant enregistré dans le Ca 2 + libre 3 mM Mg 2 +-solution du bain contenant n ° 1 au début de l&#39;expérience comme une &quot;fuite&quot; en cours. </li><li> Enregistrer les traces enregistrées en cours d&#39;analyse. </li></ol> 7. Analyse des données <ol><li> Récupérer les enregistrements sauvés en cours dans le logiciel d&#39;analyse. Nous utilisons le programme d&#39;impulsion pour analyse. </li><li> Mesurer les amplitudes actuelles au début et à la fin de rampes de tension. Nous généralement mesurer les amplitudes de courant à -100 mV et 100 mV à l&#39;aide de paires de curseurs dans la &quot;Oscilloscope&quot; fenêtre du programme d&#39;impulsion. </li><li> Exporter les valeurs des amplitudes de courant dans un programme graphique pour approfondir l&#39;analyse et la présentation graphique. Nous utilisons des graphiques origine scientifique et logiciels d&#39;analyse, la version 7. </li></ol> 8. Les résultats représentatifs <img src="/files/ftp_upload/2346/2346fig2.jpg" alt="Figure 2" /> Figure 2. CRAC courants dans une cellule T humaine de repos. (A), évolution dans le temps des courants CRAC enregistrée dans la cellule entière voltage-clamp de configuration à -100 mV (cercles pleins) et 100 mV (cercles ouverts). Avant la formation gigaseal, la cellule a été pré-incubées pendant 10 min dans ~ Ca 2 + libre 3 mM Mg 2 +-solution du bain contenant n ° 1 contenant 0,5 thapsigargine uM. Après le rodage, les solutions de bain ont été appliqués séquentiellement comme suit: Ca 2 + libre 3 mM Mg 2 +-solution du bain contenant # 1 (0 + 3 Ca Mg), suivie par 20 mM de solution de Ca 2 + bain contenant # 2 (20 Ca), suivie par la solution de bain de cations divalents libres # 3 (DVF), suivie par solution du bain # 2 (20 Ca). La cellule a été stimulé par une série de rampes de tension comme le montre la figure 1. La fréquence de 5 Hz a été rampes en solution de bain # 3 (DVF) et 0,5 Hz dans tous les autres solutions. Notez le lent développement de I Ca-CRAC, après application de 20 mM de Ca 2 +-solution du bain contenant # 2 et le développement rapide aux transitoires du I Na-CRAC en solution de bain DVF # 3. (B), représentant les traces actuelles enregistrées au cours des rampes de tension en Ca 2 + libre 3 mM Mg 2 +-solution du bain contenant # 1 («Fuite»), 20 mM de Ca 2 + solution contenant de bain # 2 (20 Ca), et la solution de bain # 3 (DVF). (C, D), courant-tension des relations de I Ca-CRAC (C) et j&#39;ai Na-CRAC (D) obtenue en soustrayant &quot;fuite&quot; en cours à partir des courants enregistrés au cours des rampes de tension dans 20 mM de Ca 2 +-bain contenant Solution n ° 2 (20 Ca), et la solution de bain # 3 (DVF) montré dans le panneau (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>

L&#39;exploration électrophysiologique des courants de repos CRAC lymphocytes T humains est une tâche difficile parce que le CRAC endogènes d&#39;amplitude en cours dans ces cellules est faible en raison de la taille des cellules de petite taille (diamètre de l&#39;reposant humaine de cellules T est dans la gamme de 5-8 um). Ici, nous présentons une procédure étape par étape pour enregistrer de manière fiable les courants de repos CRAC lymphocytes T humains isolés à partir des cellules mononucléées du sang ...

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		<th>Material Name</th>
		<th>Type</th>
		<th>Company</th>
		<th>Catalogue Number</th>
		<th>Comment</th>
		</tr>
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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>
<tr>
<td>Sylgard 184 Silicon Elastomer Kit</td>
<td>&nbsp;</td>
<td>Dow Corning, Midland, MI</td>
<td>3097358-1004</td>
<td>&nbsp;</td>
<tr>
<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

Dans les lymphocytes T, l&#39;épuisement de Ca 2 + de la intracellulaire de Ca2 + magasin conduit à l&#39;activation des plasmalemmal Ca 2 + canaux, appelés calcium Release-Activated calcium (CRAC) des canaux. Canaux CRAC jouent un rôle important dans la régulation de la prolifération des cellules T et l&#39;expression des gènes. Anomalie de la fonction de canal CRAC dans les cellules T a été liée à une immunodéficience sévère combinée et auto-immunes 1, 2. Etudier la fonction du canal CRAC dans les lymphocytes T humains peuvent découvrir de nouveaux mécanismes moléculaires régulant les réponses immunitaires normales et de démêler les causes des maladies liées à l&#39;humain. Enregistrements électrophysiologiques des courants membranaires fournissent l&#39;évaluation la plus précise des propriétés fonctionnelles de canal et de leur régulation. L&#39;évaluation électrophysiologique des courants des canaux CRAC dans les cellules T Jurkat, une ligne de la leucémie humaine des cellules T, a d&#39;abord été effectué plus de 20 ans il ya trois, cependant, les mesures actuelles dans le CRAC lymphocytes T humains normaux reste une tâche difficile. Les difficultés dans l&#39;enregistrement des courants de canal CRAC dans les cellules T normales sont aggravés par le fait que les dérivés du sang les lymphocytes T sont beaucoup plus petits en taille que les cellules T Jurkat et, par conséquent, les courants endogènes CRAC cellules entières sont très faibles en amplitude. Ici, nous donnons une procédure étape par étape que nous utilisent couramment pour enregistrer le Ca 2 + ou Na + courants via les canaux CRAC au repos des cellules T isolées du sang périphérique de volontaires sains. La méthode décrite ici a été adoptée par les procédures utilisées pour l&#39;enregistrement des courants CRAC dans les cellules T Jurkat et activé des cellules T 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

Nous fournissons un protocole étape par étape pour la cellule entière d&#39;enregistrement de patch-clamp de Calcium Release-Activated calcium (CRAC) des courants dans les zones périphériques mononucléées du sang de cellules dérivées de lymphocytes T humains.

Cellules entières d&#39;enregistrement de Calcium Release-Activated calcium (CRAC) Courants dans les lymphocytes T humains

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 vues.  University of California, Davis. Nous fournissons un protocole étape par étape pour la cellule entière d'enregistrement de patch-clamp de Calcium Release-Activated calcium (CRAC) des courants dans les zones périphériques mononucléées du sang de cellules dérivées de lymphocytes T humains.

Vidéo: 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.

Terminaux de chargement nerveuse chez la drosophile avec des indicateurs de calcium

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 ...

Recherche

Biologie

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 .

In Vivo 2-Photon Imaging calcium dans la couche 2 / 3 de souris

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.

Cellules entières Recordings of Light évoqués courants synaptiques excitateurs dans la tranche la rétine

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.

Mesure Proche membrane plasmique et mondiale dynamique du calcium intracellulaire dans les 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, ...

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.

Surveillance des changements dynamiques dans les niveaux de calcium mitochondrial cours de l&#39;apoptose en utilisant un capteur de calcium codées génétiquement

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.

Mesurer les flux de calcium dans les cardiomyocytes rapide

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.

Évaluation de calcium Sparks dans intactes les fibres musculaires squelettiques

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.

Mesures de calcium cytosolique dans les cellules épithéliales rénales par cytométrie en flux

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.

Enregistrement en temps réel In Vivo des signaux calciques Arabidopsis pendant la tétée insectes à l’aide d’un biocapteur Fluorescent

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.

Imagerie de la dynamique du calcium dans les sous-populations de cellules d'islet pancréatique de souris

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