Wir sind dankbar, dass das Department of Physiology and Membrane Biology, University of California Davis, die uns mit Einrichtungen und ein ausgezeichnetes Umfeld für die Studien an Ionenkanälen.

1. Vorbereitung der Ruhe menschlichen T-Lymphozyten <ol><li> Mit RosetteSep Menschliche T-Zell-Anreicherung Cocktail und RosetteSep Density Medium, reinigen T-Zellen aus menschlichen Blutproben nach den Anweisungen des Herstellers. Die daraus resultierende Zellpopulation sollte 95% CD3 + ruhende T-Zellen. Wir reinigen menschlichen T-Lymphozyten aus dem peripheren Blutproben von gesunden Freiwilligen in Übereinstimmung mit dem Protokoll von der UC Davis Internal Review Board genehmigt gesammelt. </li><li> Nach der Isolierung resuspendieren ruhende T-Zellen bei einer Dichte von 0,5 x 10 6 Zellen / ml in Zellkulturmedium RPMI-1640 Medium mit Glutamin und HEPES, ergänzt mit 10% fötalem Kälberserum, 2% Glutamax-I, 1% RPMI 1640 Vitamin-Lösung, 1% RPMI 1640 Aminosäuren-Lösung, 1% Natrium-Pyruvat und 0,03% β-Mercaptoethanol. </li><li> Legen Sie Zellsuspension in Zellkulturflaschen und pflegen bei 37 ° C in einem befeuchteten Inkubator mit 5% CO 2 bis zu 24 h vor dem Experiment. </li></ol> 2. Vorbereitung der Aufnahme Kammer <ol><li> Die folgenden sind notwendig, um die Aufnahme Kammern vorzubereiten: a) Silizium-O-Ringe, b) um, 25-mm Deckgläser mit 70% Ethanol und destilliertem H 2 O, C gereinigt) gemischt Sylgard184, zubereitet nach den Anweisungen des Herstellers, und d) zwei 60 x 15 mm Petrischalen. </li><li> Legen ~ 0,5 ml gemischt Sylgard184 in ein 60x15-mm-Petrischale. </li><li> Tauchen Sie den unteren Teil der O-Ring in die Sylgard184 mit einer Pinzette. </li><li> Entfernen Sie überschüssiges Sylgard184 vom Rand der O-Ring, indem Sie den O-Ring auf einer sauberen Fläche, wie eine weitere Petrischale. </li><li> Platzieren Sie den Ring auf das Deckglas und drücken Sie ihn nach unten. </li><li> Cure den Sylgard184 durch Erhitzen der Kammern bei 85 ° C für 40-60 min oder bis die Sylgard184 polymerisiert wird. </li><li> Bewahren Sie die Aufnahme Kammern in einem staubfreien Behälter. </li></ol> 3. Erstellung von Patch-Pipetten <ol><li> Am Tag des Experiments, ziehen Pipetten mit a ~ 2-um Spitzendurchmesser mit Glaskapillaren und einer Pipette Abzieher. Wir verwenden Borosilikat Schlauch mit Filament (OD: 1,5 mm und ID: 1,10 mm). Shop-Pipetten in einem staubfreien Ort. </li><li> Coat Pipettenspitzen mit Sylgard184 oder HIPEC R6101 Semiconductor Schutzschicht, nach einem Standard-Verfahren 9. </li><li> Gently Feuer-polish die Pipetten vor dem Experiment. </li></ol> 4. Patch-Clamp-Setup Vorbereitung <ol><li> Konfigurieren und speichern Sie die Makros für Gigaseal Bildung in der Software-Steuerung Ihres Patch-Clamp-Verstärker. Wir verwenden ein EPC-10-Verstärker von Windows XP und Pulse-Software für die Datenerfassung unterstützt. Wir setzen die &quot;set-up&quot;, &quot;on-cell&quot; und &quot;whole-cell&quot;-Makros nach dem EPC-10 Bedienungsanleitung und verwenden Sie sie für alle Experimente. </li><li> Bereiten Bad-Lösungen und-Pipette Lösungen in Tabelle 1 aufgeführten vor dem Experiment und lagern Sie sie bei 4 ° C für bis zu 1 Woche. </li></ol><table border="1"><tbody><tr><td> Chemicals </td><td colspan="3"> Bath Solutions (mM) </td><td> Pipette 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> 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-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> Glucose </td><td> 10 </td><td> 10 </td><td> 10 </td><td> - </td></tr></tbody></table> Tabelle 1. Lösungen für whole-cell CARC aktuelle Aufnahmen. Alle Chemikalien wurden von Sigma-Aldrich, St. Louis, MO gekauft. <ol start="3"><li> Bestimmen Sie Flüssigkeitsverbindung Potentiale (LJ) zwischen der Patch-Pipette Lösung und jede Badlösung 10. Speichern Sie die LJ-Wert an der entsprechenden Stelle mit Ihrem Erwerb Software. Zum Beispiel in der Pulse-Programm legen Sie die LJ-Wert in &quot;LJ&quot; Kontrolle im &quot;Amplifier&quot;-Fenster. </li><li> Konfigurieren und speichern Sie die Stimulationsprotokolle, wie in Abbildung 1 gezeigt, an der entsprechenden Stelle Ihrer Software zur Erfassung, bevor die Experimente. In der Pulse-Programm setzen wir die Übernahme-Protokoll in der &quot;Pulse Generation&quot;-Fenster. <img src="/files/ftp_upload/2346/2364fig1.jpg" alt="Abbildung 1" /> Abbildung 1. . Stimulationsprotokoll Spannungsrampe von -120 bis +100 mV von 50 ms Dauer wird alle 0,2 angewendet - 2 s (5 bis 0,5 Hz). Halten Potential (V h) zwischen Rampen ist bei +30 mV aufrechterhalten. </li></ol> 5. Montage Cells auf dem Mikroskoptisch <ol><li> Bestreichen Sie die Aufnahme Kammern mit Poly-L-Lysin für ~ 20 min und wäscht mit H 2 O und dann mit Hanks balanced salt solution (HBSS) vor dem Ausplattieren der Zellen. </li><li> Platte 500 ul Zellsuspension mit 0,2-0,5 x10 6 Zellen in die Kammer und inkubieren für 20 min bei 37 ° C. </li><li> Sichern Sie die Aufnahme Kammer mit der anhaftenden Zellen in eine Kammer Halter. Wir verwenden eine maßgeschneiderte Kammer Halter, wo die Basis unterstützt der Unterseite des Deckglases, während der obere Teil sanft drückt gegen die Oberseite des Deckglases. </li><li> Setzen Sie die Kammer Halter auf der Bühne des inversen Mikroskop. </li><li> Konzentrieren Sie sich auf die Zellen im Durchlicht. Wir verwenden ein 40X Ölimmersionsobjektiv. </li><li> Legen Sie die Ag / AgCl-Referenzelektrode in die Wanne. </li><li> Richten Sie die mehrgehäusigen, Schwerkraft-Perfusion System. Die Spitze des Zulaufrohr sollte in einem 45 ° Winkel zur Deckglas in der Nähe des Sichtfeldes platziert werden. </li><li> Positionieren Sie den Saugschlauch gegenüber der Spitze des Zulaufrohr. </li><li> Versorgen die Kammer mit Phosphat-gepufferter Salzlösung (PBS) oder HBSS abzuwaschen den Zellkulturmedien und lose anhaftenden Zellen. </li></ol> 6. CRAC-Channel Currents Recording <ol><li> Finden Sie eine Zelle, die fest mit dem Deckglas befestigt ist. Wir in der Regel wählen Sie einen durchschnittlich großen, runden Zellen mit einer glatten Außenfläche. </li><li> Versorgen die Aufnahme Kammer mit Ca 2 +-frei, 3 mM Mg 2 +-haltigen Badlösung # 1 (Tabelle 1) mit 0,5-1 pM Thapsigargin für 8-10 min auf die SERCA-Pumpe blockieren. Wir führen diesen Schritt, um das Geschäft vor Gigaseal Bildung führen. </li><li> Mit einer Eppendorf Microloader und eine P-20 Pipette, füllen Sie die Patch-Pipette mit der Pipette Lösung (Tabelle 1) und legen Sie die Patch-Pipette in die Pipettenhalter am Verstärker headstage. Die Ca 2 +-Chelator BAPTA ist in der Pipette Lösung enthalten, passiv zum Abbau der zu speichern und Ca 2 +-abhängigen Inaktivierung CRAC Kanal zu reduzieren. </li><li> Eine kleine Menge von positivem Druck (~ 3 Zoll H 2 O) in der Patch-Pipette vor dem Betreten des Bades. Wir verwenden eine maßgeschneiderte Druck-Saug-Gerät, das an Druckluft-und Vakuum-Leitungen zu positiven oder negativen Druck innerhalb einer Patch-Pipette zu schaffen. </li><li> Senken Sie die Patch-Pipette in das Bad unter visueller Kontrolle über das Mikroskop. In der &quot;Oszilloskop&quot;-Fenster, sollten Sie einen Strom durch die Pipette. Eine Schritt-für, wie Änderung der Stromamplitude sollte durch eine kleine Amplitude Spannungsimpuls (Test-Impuls) durch die voreingestellten Makro angewendet induziert werden. Zu diesem Zeitpunkt sollte es möglich sein, den Wert der Pipette Widerstand zu bestimmen. Der Widerstand der Pipetten ist in der Regel 5-6 MOhm. </li><li> Korrigieren Sie die &quot;Offset&quot; Potential zwischen der Pipette und der Referenz-Elektrode erzeugt. </li><li> Unter visueller Kontrolle über das Mikroskop bringen die Pipettenspitze näher an die Zelle, bis sie die Zellmembran berührt. </li><li> Bewerben Unterdruck (20-40 cm H 2 O) in der Patch-Pipette. </li><li> Überwachen Sie die Gigaseal Bildung in der &quot;Oszilloskop&quot;-Fenster. Sie sehen die Amplitude des Stromes, durch den Test Puls sinkt und der Wert der Pipette Widerstand steigt auf&gt; 5 GOhm induziert. Normalerweise Gigaseal Formen innerhalb einer few Sekunden, aber es kann 1-2 min, um eine stabile Gigaseal erreichen. </li><li> Kompensieren Sie die Pipette Kapazität (&quot;C-Fast&quot;). </li><li> Break the patch Membran durch zusätzliche Unterdruck mittels einer 20 ml-Spritze. </li><li> Stellen Sie die Haltepotential bis +30 mV. Eine positive Haltepotential hilft, Calcium-abhängigen Inaktivierung CRAC Kanal zu verhindern. </li><li> Ausgleich für die Membran Kapazität der Zelle (&quot;C-slow&quot;); speichern oder notieren Sie den Wert von C-slow. </li><li> Überprüfen Sie den Wert des Zugangs Widerstand &quot;R s&quot;. In unseren Experimenten ist die R s in der Regel 7-20 MOhm. </li><li> Starten Sie zu einer Reihe von Spannung Rampen mit einer Frequenz von 0,5 Hz (Abbildung 1) in Ca 2 +-freien 3 mM Mg 2 +-haltigen Badlösung Nr. 1 gelten. In dieser Lösung ist CRAC aktuellen vernachlässigbar klein wie bei den &quot;Leck&quot; aktuellen aufgrund der schlechten Durchlässigkeit von CRAC Kanäle für Mg 2 + verglichen. Speichern und verwenden Sie die aktuelle Spuren in Badlösung Nr. 1 in zukünftigen Analysen aufgezeichnet als &quot;Leck&quot; Ströme. Die absolute Amplitude der &quot;Leck&quot; Strom bei -100 mV sollte weniger als oder gleich 5 pA. Wenn die Zelle &quot;leaky&quot;, den Test beenden und neu zu beginnen mit einem neuen Patch-Pipette und eine neue Zelle. </li><li> Zur Aufnahme Ca 2 + Strom über CRAC Kanäle (I Ca-CRAC), ersetzen Ca 2 +-freien 3 mM Mg 2 +-haltigen Badlösung Nr. 1 mit 20 mM Ca 2 +-haltigen Badlösung # 2 (Tabelle 1) durch Einschalten der Perfusion Ventile. Dies ermöglicht eine Ca 2 + zu fließen über CRAC Kanäle und erzeugen eine nach innen gerichteten Strom. In der &quot;Oszilloskop&quot; Fenster, sehen Sie die Entwicklung von nach innen Behebung I Ca-CRAC (Abbildung 2). Die Stromamplitude bei negativen Spannungen wird weiterhin für ca. 1 min durch die Ca 2 +-abhängige Potenzierung der CRAC aktuellen erhöhen. </li><li> Erhöhen Sie die Frequenz der Stimulation mit Spannung Rampen bis 5 Hz, die notwendig sind, um die transiente Na + Strom über CRAC Kanäle aufnehmen wird. </li><li> Ersetzen Sie die 20 mM Ca 2 +-haltigen Badlösung # 2 mit Na +-haltigem zweiwertiges Kation-freie Lösung # 3 (Tabelle 1) durch Einschalten der Perfusion Ventile. In der &quot;Oszilloskop&quot; Fenster, sehen Sie eine vorübergehende Entwicklung größerer Amplitude innerlich Behebung Na + Strom über die CRAC-Kanäle (I Na-CRAC; Abbildung 2). </li><li> Man kann gelten 20 mM Ca 2 +-haltigen Badlösung Nr. 2 mit 1-5 uM La 3 + am Ende des Experiments, die &quot;Leck&quot; aktuellen Datensatz. Allerdings haben wir nicht finden, dieser Ansatz nützlich, da die &quot;Leck&quot; aktuelle Veränderungen im Laufe der Zeit und es kann größer oder kleiner werden am Ende des Experiments. Wir bevorzugen es, den Strom in der Ca 2 +-freien aufgezeichnet 3 mM Mg 2 +-haltigen Badlösung # 1 am Anfang des Experiments als &quot;Leck&quot; Strom. Nehmen </li><li> Speichern Sie die aufgezeichneten Strom Spuren für die weitere Analyse. </li></ol> 7. Data Analysis <ol><li> Abrufen der gespeicherten aktuellen Datensätze in die Analyse-Software. Wir nutzen die Pulse-Programm für die Analyse. </li><li> Messen Sie den Strom Amplituden am Anfang und am Ende des Spannungsrampen. Wir messen typischerweise die Stromamplituden bei -100 mV und +100 mV unter Verwendung von Paaren von Cursor in die &quot;Oszilloskop&quot; Fenster der Pulse-Programm. </li><li> Exportieren Sie die Werte der Stromamplituden in ein Grafikprogramm zur weiteren Analyse und grafische Darstellung. Wir verwenden Origin Wissenschaftliche Visualisierungs-und Analyse-Software, Version 7. </li></ol> 8. Repräsentative Ergebnisse <img src="/files/ftp_upload/2346/2346fig2.jpg" alt="Abbildung 2" /> Abbildung 2. CRAC Ströme in einem ruhenden humanen T-Zell. (A), Zeitlicher Verlauf der CRAC Ströme in whole-cell Voltage-Clamp-Konfiguration bei -100 mV (gefüllte Kreise) und +100 mV (offene Kreise) aufgezeichnet. Vor Gigaseal Bildung, wurde die Zelle für ~ 10 min in Ca 2 +-freien vorinkubiert 3 mM Mg 2 +-haltigen Badlösung Nr. 1 mit 0,5 uM Thapsigargin. Nach Einbruch, wurden Badlösungen nacheinander wie folgt angewandt: Ca 2 +-freien 3 mM Mg 2 +-haltigen Badlösung # 1 (0 Ca + 3 Mg), gefolgt von 20 mM, gefolgt Ca 2 +-haltigen Badlösung # 2 (20 Ca), durch zweiwertige Kationen-freie Badlösung # 3 (DVF), gefolgt von Badlösung # 2 (20 Ca) gefolgt. Die Zelle wurde mit einer Reihe von Spannungsrampen wie in Abbildung 1 gezeigt, stimuliert. Die Häufigkeit der Rampen wurde 5 Hz in Badlösung # 3 (DVF) und 0,5 Hz in allen anderen Lösungen. Beachten Sie die langsame Entwicklung von I Ca-CRAC nach der Applikation von 20 mM Ca 2 +-haltigen Badlösung Nr. 2 und die schnelle transiente Entwicklung von I Na-CRAC in DVF Badlösung Nr. 3. (B), Vertreter aktuellen Spuren während Spannungsrampen in Ca 2 +-freien aufgezeichnet 3 mM Mg 2 +-haltigen Badlösung # 1 (&quot;Leck&quot;), 20 mM Ca 2 +-haltigen Badlösung # 2 (20 Ca) und Badlösung # 3 (DVF). (C, D), Strom-Spannungs-Beziehungen von I Ca-CRAC (C) und ich Na-CRAC (D) durch Subtraktion &quot;Leck&quot; Strom aus der Strömung während des Spannungs-Rampen in 20 mM aufgezeichnet Ca 2 +-haltigen Bad erhalten Lösung # 2 (20 Ca) und Badlösung # 3 (DVF) in Panel dargestellt (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>

Die elektrophysiologische Untersuchung von CRAC Ströme in Ruhe menschlichen T-Zellen ist eine anspruchsvolle Aufgabe, da die endogene CRAC Stromamplitude in diesen Zellen ist wegen der geringen Größe der Zellen (die ruhende humane T-Zell-Durchmesser liegt im Bereich von 5-8 um). Hier präsentieren wir eine Schritt-für-Schritt-Verfahren zuverlässig zu erfassen CRAC Ströme in Ruhe menschlichen T-Lymphozyten aus dem peripheren Blut mononukleären Zellen isoliert. Diese Technologie ermöglicht es uns, die Physiologie ...

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		<th>Material Name</th>
		<th>Type</th>
		<th>Company</th>
		<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>
<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

In T-Lymphozyten, die Erschöpfung des Ca 2 + aus den intrazellulären Ca 2 + zu speichern führt zur Aktivierung von plasmalemmal Ca 2 +-Kanäle, genannt Calcium Release-Activated Calcium (CRAC) Kanäle. CRAC-Kanäle spielen eine wichtige Rolle bei der Regulation der T-Zell-Proliferation und Genexpression. Abnormal CRAC-Kanal-Funktion in T-Zellen hat schwere kombinierte Immundefizienz und Autoimmunerkrankungen 1, 2 in Verbindung gebracht worden. Studieren CRAC-Kanal-Funktion in menschlichen T-Zellen können gemeinsam neue molekulare Regulationsmechanismen normalen Immunantwort und lösen die Ursachen verwandter Krankheiten beim Menschen. Elektrophysiologischen Ableitungen von Membran-Ströme liefern die genaue Beurteilung der funktionellen Kanal Eigenschaften und ihre Regulation. Elektrophysiologische Beurteilung der CRAC Kanal Ströme in Jurkat T-Zellen, einer menschlichen Leukämie-T-Zell-Linie, zuerst durchgeführt wurde mehr als 20 Jahren 3, jedoch bleibt CRAC Strommessungen in normalen menschlichen T-Zellen eine anspruchsvolle Aufgabe. Die Schwierigkeiten bei der Aufnahme CRAC Kanal Ströme in normalen T-Zellen werden durch die Tatsache, dass Blut gewonnene T-Lymphozyten viel kleiner als Jurkat T-Zellen und sind daher die endogene whole-cell CRAC Ströme sehr gering in der Amplitude zusammengesetzt sind. Hier geben wir eine Schritt-für-Schritt-Verfahren, das wir regelmäßig verwenden, um die Ca 2 + Rekord oder Na +-Ströme über CRAC Kanäle in Ruhe menschlichen T-Zellen aus dem peripheren Blut von gesunden Probanden isoliert. Die hier beschriebene Methode wurde von den Verfahren für die Aufnahme der CRAC Ströme in Jurkat T-Zellen und aktivierten humanen T-Zellen 4-8 verwendet hat.

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

Wir bieten Ihnen eine Schritt-für-Schritt-Protokoll für whole-cell Patch-Clamp-Aufnahme von Calcium Release-Activated Calcium (CRAC) Ströme in peripheren mononukleären Zellen gewonnenen humanen T-Lymphozyten.

Whole-Cell Aufnahme von Calcium Release-Activated Calcium (CRAC) Ströme in menschlichen T-Lymphozyten

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 Aufrufe.  University of California, Davis. Wir bieten Ihnen eine Schritt-für-Schritt-Protokoll für whole-cell Patch-Clamp-Aufnahme von Calcium Release-Activated Calcium (CRAC) Ströme in peripheren mononukleären Zellen gewonnenen humanen T-Lymphozyten.

Serie: 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.

Loading Drosophila Nerve Terminals mit Calcium-Indikatoren

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 ...

Forschung

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 Calcium Imaging in Layer 2 / 3 der Mäuse

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.

Ganzzellableitungen of Light evozierten exzitatorischen synaptischen Ströme in der Netzhaut Scheiben

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.

Messen in der Nähe Plasmamembran und Global intrazellulären Calcium-Dynamics in Astrozyten

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.

Überwachung dynamischen Veränderungen in der mitochondrialen Kalzium während der Apoptose mit einem genetisch kodierten Kalzium-Sensor

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.

Mess-Fast-Flux Calcium in Kardiomyozyten

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.

Bewertung von Calcium in Sparks Intact Skelettmuskelfasern

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.

Cytosolischen Calcium Messungen in Nierenepithelzellen mittels Durchflusszytometrie

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.

In Echtzeit In Vivo Aufnahme von Arabidopsis Kalziumsignale während der Fütterung mit einer fluoreszierenden Biosensor Insekt

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.

Bildgebende Calciumdynamik in Subpopulationen von Maus-Pankreas-Islet-Zellen

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