Siamo grati al Dipartimento di Fisiologia e membrana Biologia, Università di California Davis per averci fornito servizi e un ambiente eccellente per lo studio dei canali ionici.

1. Preparazione di riposo linfociti T umani <ol><li> Utilizzando Cocktail RosetteSep umani arricchimento cellule T e Medium Density RosetteSep, purificare le cellule T da campioni di sangue umano secondo le istruzioni del produttore. La popolazione cellulare risultante dovrebbe contenere 95% di cellule T CD3 + riposo. Noi purificare i linfociti T umani da campioni di sangue periferico raccolti da volontari sani in conformità al protocollo approvato dal Consiglio di UC Davis interno Review. </li><li> Dopo l&#39;isolamento, risospendere riposo le cellule T ad una densità di 0,5 x 10 6 cellule / ml nel terreno di coltura contenente RPMI-1640 medium con la glutamina e HEPES, integrati da siero di vitello fetale del 10%, 2% GlutaMAX-I, 1% RPMI 1640 vitamina soluzione, 1% RPMI 1640 aminoacidi soluzione, piruvato di sodio 1% e 0,03% β-mercaptoetanolo. </li><li> Luogo sospensione di cellule in fiasche di coltura cellulare e di mantenere a 37 ° C in un incubatore umidificato al 5% di CO 2 fino a 24 ore prima dell&#39;esperimento. </li></ol> 2. Preparazione della Camera di registrazione <ol><li> I seguenti sono necessari per preparare camere di registrazione: a) al silicio O-ring, b) rotondo, 25 mm coprioggetto pulito con etanolo al 70% e distillata H 2 O, c) miscelato Sylgard184, preparati secondo le istruzioni del produttore e d) due 60 x 15 mm Petri. </li><li> Luogo ~ 0,5 mL di misto Sylgard184 in un 60x15 mm piastra di Petri. </li><li> Immergere la parte inferiore della O-ring nel pinze Sylgard184 utilizzando. </li><li> Rimuovere l&#39;eccesso Sylgard184 dal bordo della O-ring mettendo l&#39;O-ring su una superficie pulita, come un altro piatto di Petri. </li><li> Posizionare l&#39;anello sul coprioggetto e premere verso il basso. </li><li> Curare la Sylgard184 dal riscaldamento delle camere a 85 ° C per 40-60 minuti o fino a quando il Sylgard184 è polimerizzato. </li><li> Memorizzare le camere di registrazione in un contenitore privo di polvere. </li></ol> 3. Preparazione del Pipette Patch <ol><li> Il giorno dell&#39;esperimento, tirare pipette con diametro punta ~ 2-micron utilizzando vetro capillari e un estrattore pipetta. Utilizziamo tubi in vetro borosilicato con filamento (OD: 1,5 mm e ID: 1,10 mm). Pipette conservare in un luogo privo di polvere. </li><li> Cappotto puntali con Sylgard184 o HIPEC rivestimento protettivo Semiconductor R6101, secondo una procedura standard 9. </li><li> Delicatamente fuoco lucidare le pipette prima dell&#39;esperimento. </li></ol> 4. Patch Morsetto Preparazione installazione <ol><li> Configurare e salvare le macro per la formazione gigaseal nel software di controllo del vostro patch-clamp amplificatore. Usiamo un EPC-10 amplificatore supportata da Windows XP e software Pulse per l&#39;acquisizione dei dati. Abbiamo impostato il &quot;set-up&quot;, &quot;a-cella&quot;, e &quot;a cellula intera&quot; macro secondo la EPC-10 manuale e utilizzarli per tutti gli esperimenti. </li><li> Preparare le soluzioni di bagno e soluzioni pipetta elencati nella tabella 1 prima l&#39;esperimento e conservarli a 4 ° C per un massimo di 1 settimana. </li></ol><table border="1"><tbody><tr><td> Prodotti chimici </td><td colspan="3"> Soluzioni per vasca da bagno (mM) </td><td> Soluzioni pipetta (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> Glucosio </td><td> 10 </td><td> 10 </td><td> 10 </td><td> - </td></tr></tbody></table> Tabella 1. Soluzioni per le registrazioni CARC intera-cella corrente. Tutti i prodotti chimici sono stati acquistati da Sigma-Aldrich, St. Louis, MO. <ol start="3"><li> Determinare i potenziali di giunzione liquida (LJ) tra la soluzione di pipetta di patch e ogni soluzione da bagno 10. Salvare il valore LJ nel luogo appropriato utilizzando il software di acquisizione. Per esempio, nel programma Pulse inserire il valore LJ in controllo &quot;LJ&quot; nella finestra &quot;amplificatore&quot;. </li><li> Configurare e salvare i protocolli di stimolazione, come mostrato nella Figura 1, nell&#39;apposito spazio del vostro software di acquisizione prima che gli esperimenti. Nel programma Pulse, abbiamo impostato il protocollo di acquisizione della &quot;Generazione Pulse&quot; finestra. <img src="/files/ftp_upload/2346/2364fig1.jpg" alt="Figura 1" /> Figura 1. . Protocollo di stimolazione rampa di tensione da -120 a +100 mV di 50 ms di durata viene applicata ogni 0,2 - 2 s (5 - 0,5 Hz). Potenziale Holding (V h) tra le rampe è mantenuta a 30 mV. </li></ol> 5. Le cellule di montaggio sullo stage microscopio <ol><li> Cappotto le camere di registrazione con poli-L-lisina per ~ 20 minuti e lavare con H 2 O e poi con soluzione salina bilanciata di Hank (HBSS) prima di placcatura le cellule. </li><li> Piatto 500 microlitri di sospensione cellulare contenente 0,2-0,5 x10 6 celle nella camera e incubare per 20 minuti a 37 ° C. </li><li> Fissare la camera di registrazione con le cellule attaccato in un supporto da camera. Noi usiamo una misura porta della camera, dove la base sostiene la parte inferiore del vetrino, mentre la parte superiore leggermente preme contro la parte superiore del coprioggetto. </li><li> Posizionare il supporto da camera sul palco del microscopio invertito. </li><li> Concentrarsi sulle cellule in luce trasmessa. Usiamo un obiettivo 40X immersione in olio. </li><li> Posizionare la Ag / AgCl elettrodo di riferimento nella vasca. </li><li> Allineare l&#39;, multibarrel gravità sistema guidato perfusione. La punta del tubo di flusso deve essere posizionato ad un angolo di 45 ° fino alla fine coprioggetto al campo di vista. </li><li> Posizionare il tubo di aspirazione di fronte alla punta del tubo di afflusso. </li><li> Profumato la camera con tampone fosfato salino (PBS) o HBSS per lavare via i media di coltura cellulare e le cellule debolmente collegate. </li></ol> 6. CRAC Canale Correnti di registrazione <ol><li> Trova una cella che è saldamente attaccato al vetrino. Noi di solito selezionare una media grandezza, cellule rotonde con una superficie esterna liscia. </li><li> Profumato della camera di registrazione con Ca 2 +-free, 3 mM Mg 2 + che contiene la soluzione bagno # 1 (tabella 1) contenente 0,5-1 thapsigargin mM per 8-10 minuti per bloccare la pompa SERCA. Eseguiamo questa operazione per esaurire il negozio prima della formazione gigaseal. </li><li> Utilizzando un microloader Eppendorf e un P-20 pipetta a spostamento, riempire la pipetta di patch con la soluzione di pipetta (Tabella 1) e inserire la pipetta di patch nel supporto pipetta sulla headstage amplificatore. Il Ca 2 + chelante Bapta è incluso nella soluzione pipetta ad esaurire passivamente il negozio e per ridurre Ca 2 +-dipendente inattivazione del canale CRAC. </li><li> Applicare una piccola quantità di pressione positiva (~ 3 pollici di H 2 O) dentro la pipetta di patch prima di entrare nel bagno. Noi usiamo una misura della pressione di aspirazione dispositivo collegato ad aria compressa e le linee di vuoto per creare una pressione positiva o negativa all&#39;interno di una pipetta di patch. </li><li> Abbassare la pipetta di patch nel bagno sotto controllo visivo tramite il microscopio. Nella finestra &quot;Oscilloscopio&quot;, dovreste vedere una corrente che fluisce attraverso la pipetta. Un passo simile cambiamento nella ampiezza della corrente deve essere indotto da un impulso di tensione piccola ampiezza (trigger) applicato dalla macro preimpostate. A questo punto, dovreste essere in grado di determinare il valore di resistenza pipetta. La resistenza delle nostre pipette è tipicamente 5-6 MW. </li><li> Correggere il potenziale &quot;offset&quot; generato tra la pipetta e l&#39;elettrodo di riferimento. </li><li> Sotto controllo visivo tramite il microscopio, portare la punta della pipetta più vicino alla cella fino a toccare la membrana cellulare. </li><li> Applicare una pressione negativa (20-40 cm di H 2 O) dentro la pipetta di patch. </li><li> Monitorare la formazione gigaseal nella finestra &quot;oscilloscopio&quot;. Vedrete l&#39;ampiezza della corrente, indotta dagli impulsi di test, diminuisce e aumenta il valore della resistenza pipetta&gt; 5 GΩ. Forme di solito gigaseal all&#39;interno di un fesecondo w, ma potrebbero essere necessari 1-2 minuti per ottenere un gigaseal stabile. </li><li> Compensare la capacità pipetta (&quot;C-Fast&quot;). </li><li> Rompere la membrana del cerotto da applicare un&#39;ulteriore pressione negativa utilizzando una siringa da 20 cc. </li><li> Impostare il potenziale tenendo a +30 mV. Un potenziale tenendo positivo aiuta a prevenire il calcio-dipendente inattivazione del canale CRAC. </li><li> Compensare la capacità della membrana della cellula (&quot;C-slow&quot;); salvare o registrare il valore di C-slow. </li><li> Controllare il valore della resistenza di accesso &quot;R s&quot;. Nei nostri esperimenti, la R s è tipicamente 7-20 MW. </li><li> Iniziare ad applicare una serie di rampe di tensione con una frequenza di 0,5 Hz (Figura 1) a Ca 2 +-gratis 3 mM Mg 2 + che contiene la soluzione da bagno # 1. In questa soluzione, la corrente CRAC è trascurabile rispetto alla &quot;fuga&quot; in corso a causa della scarsa permeabilità dei canali CRAC per Mg 2 +. Salvare e utilizzare le tracce registrate in soluzione attuale vasca n. 1 su future analisi come &quot;perdita&quot; delle correnti. L&#39;ampiezza assoluta della &quot;fuga&quot; in corso a -100 mV deve essere inferiore o uguale a 5 pA. Se la cella è &quot;permeabile&quot;, interrompere l&#39;esperimento e ricominciare con una pipetta nuova patch e una nuova cellula. </li><li> Per registrare Ca 2 + corrente attraverso canali CRAC (I Ca-CRAC), sostituire Ca 2 +-free 3 mM Mg 2 + che contiene la soluzione bagno # 1 con 20 mM di Ca 2 +-bagno che contiene la soluzione # 2 (tabella 1) cambiando le valvole di perfusione. Questo permette di Ca 2 + a fluire attraverso canali CRAC e producono una corrente verso l&#39;interno. Nella finestra &quot;Oscilloscopio&quot;, si vedrà lo sviluppo di dentro rettificare I Ca-CRAC (Figura 2). L&#39;ampiezza della corrente a tensioni negative continueranno ad aumentare per circa 1 minuto a causa del Ca 2 +-dipendente potenziamento della corrente CRAC. </li><li> Aumentare la frequenza di stimolazione con rampe di tensione a 5 Hz, che è necessario per registrare il transitorio di corrente Na + attraverso canali CRAC. </li><li> Sostituire il 20 mM Ca 2 +-bagno che contiene la soluzione # 2 con Na +-contenenti cationi bivalenti senza soluzione di # 3 (Tabella 1) cambiando le valvole di perfusione. Nella finestra &quot;Oscilloscopio&quot;, si vedrà uno sviluppo di maggiore ampiezza transitoria interiormente rettificazione Na + corrente attraverso i canali CRAC (I Na-CRAC; Figura 2). </li><li> Si può applicare 20 mM Ca 2 +-bagno che contiene la soluzione # 2, che contengono 1-5 mM La 3 +, alla fine dell&#39;esperimento per registrare la &quot;fuga&quot; di corrente. Tuttavia, non abbiamo trovato questo approccio utile perché la &quot;perdita&quot; cambiamenti in corso nel tempo e può diventare grandi o più piccoli alla fine dell&#39;esperimento. Noi preferiamo seguire la corrente registrato nel Ca 2 +-free 3 mM Mg 2 + che contiene la soluzione bagno # 1 all&#39;inizio dell&#39;esperimento come una &quot;fuga&quot; di corrente. </li><li> Salva le tracce registrate in corso per ulteriori analisi. </li></ol> 7. Analisi dei dati <ol><li> Recuperare il record salvati corrente in software di analisi. Usiamo il programma di impulsi per l&#39;analisi. </li><li> Misurare le ampiezze corrente all&#39;inizio e alla fine della rampa di tensione. Noi di solito misura le ampiezze di corrente a -100 mV e +100 mV con le coppie di cursori nella finestra &quot;Oscilloscopio&quot; del programma Pulse. </li><li> Esportare i valori di ampiezze di corrente in un programma di grafica per ulteriori analisi e presentazione grafica. Noi usiamo grafica scientifica Origin e software per l&#39;analisi, la versione 7. </li></ol> 8. Rappresentante Risultati <img src="/files/ftp_upload/2346/2346fig2.jpg" alt="Figura 2" /> Figura 2. CRAC correnti in una cella di riposo T umani. (A), corso Tempo di correnti CRAC registrati in whole-cell voltage-clamp configurazione a -100 mV (cerchi pieni) e +100 mV (cerchi aperti). Prima di formazione gigaseal, la cella era preincubate per circa 10 minuti a Ca 2 +-free 3 mM Mg 2 + che contiene la soluzione bagno # 1 thapsigargin contenenti 0,5 micron. Dopo il rodaggio, soluzioni bagno sono stati applicati in sequenza i seguenti: Ca 2 +-free 3 mM Mg 2 +, che contiene la soluzione bagno # 1 (0 Ca + 3 Mg), seguiti da 20 mM Ca 2 +-bagno che contiene la soluzione # 2 (20 Ca), seguita da cationi bivalenti soluzione senza bagno # 3 (DVF), seguita dalla soluzione del bagno # 2 (20 Ca). La cella è stata stimolata da una serie di rampe di tensione, come mostrato nella Figura 1. La frequenza delle rampe è stato di 5 Hz in soluzione bagno # 3 (DVF) e 0,5 Hz in tutte le altre soluzioni. Si noti il lento sviluppo di I Ca-CRAC dopo l&#39;applicazione di 20 mM Ca 2 +-bagno che contiene la soluzione # 2 e il rapido sviluppo transitoria di I Na-CRAC in soluzione bagno DVF # 3. (B), Rappresentante tracce registrate durante le rampe di attuale tensione a Ca 2 +-free 3 mM Mg 2 +, che contiene la soluzione bagno # 1 (&quot;Fuga&quot;), 20 mM Ca 2 +-bagno che contiene la soluzione # 2 (20 Ca), e la soluzione del bagno # 3 (DVF). (C, D), corrente-tensione I rapporti di Ca-CRAC (C) e io Na-CRAC (D) ottenuto sottraendo &quot;fuga&quot; di corrente dalle correnti registrati durante tensione rampe in 20 mM Ca 2 + contenenti bagno soluzione # 2 (20 Ca), e la soluzione del bagno # 3 (DVF) mostrato nel grafico (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>

Nessun conflitto di interessi dichiarati.

Le indagini elettrofisiologiche delle correnti CRAC a riposo cellule T è un compito impegnativo perché la endogeno ampiezza CRAC corrente in queste cellule è di piccole dimensioni a causa delle dimensioni piccole cellule (il riposo umano diametro delle cellule T è nel range di 5-8 micron). Qui vi presentiamo uno step-by-step procedura affidabile per registrare le correnti CRAC a riposo linfociti T umane isolate da cellule mononucleari del sangue periferico. Questa tecnologia ci permette di studiare la fisiologia e l&#39;espressione funzionale di canali CRAC a riposo le cellule T per capire meglio la natura del normale e patologico le risposte delle cellule immunitarie. Usando questo protocollo, si può misurare correnti CRAC a riposo cellule T umano così come in altre cellule del sistema immunitario, come i linfociti T attivati ​​umane, monociti e macrofagi.

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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

Nei linfociti T, l&#39;esaurimento di Ca 2 + dal intracellulare Ca 2 + negozio porta all&#39;attivazione di plasmalemmal Ca + 2 canali, chiamati Calcio Release-Activated di calcio (CRAC) canali. Canali CRAC un ruolo molto importante nella regolazione della proliferazione delle cellule T e l&#39;espressione genica. Anormale funzione del canale CRAC nelle cellule T è stato collegato a gravi malattie autoimmuni immunodeficienza combinata e 1, 2. Studiare la funzione CRAC canale in cellule T possono scoprire nuovi meccanismi molecolari che regolano le normali risposte immunitarie e svelare le cause di malattie umane correlate. Le registrazioni elettrofisiologiche delle correnti di membrana forniscono la valutazione più accurata delle proprietà del canale funzionale e la loro regolamentazione. Valutazione elettrofisiologica CRAC correnti di canale in cellule Jurkat T, un essere umano linea di leucemia a cellule T, è stata eseguita più di 20 anni fa 3, tuttavia, le misurazioni CRAC corrente in normali cellule T umano rimane un compito impegnativo. Le difficoltà di registrazione dei canali CRAC correnti in normali cellule T sono aggravate dal fatto che i derivati ​​dal sangue i linfociti T sono molto più piccoli nelle dimensioni di cellule Jurkat T e, di conseguenza, l&#39;intero endogeno delle cellule correnti CRAC sono molto bassi in ampiezza. Qui, diamo uno step-by-step procedura che abitualmente utilizzano per registrare il Ca 2 + o Na + correnti attraverso canali CRAC a riposo cellule T isolate dal sangue periferico di donatori sani. Il metodo qui descritto è stato adottato dalle procedure utilizzate per la registrazione delle correnti CRAC in cellule Jurkat T e cellule T attivate umane 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

Forniamo un passo-passo protocollo per la cellula intera registrazione patch clamp di calcio Release-Activated di calcio (CRAC) correnti in cellule mononucleate del sangue periferico derivati ​​linfociti T umani.

Whole-Cell Registrazione del calcio Release-Activated di calcio (CRAC) Correnti in Human Linfociti T

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

Nanyang Technological University

Research

JoVE Journal

Biology

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

16.0K Views.  University of California, Davis. Forniamo un passo-passo protocollo per la cellula intera registrazione patch clamp di calcio Release-Activated di calcio (CRAC) correnti in cellule mononucleate del sangue periferico derivati ​​linfociti T umani.

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

Loading Drosophila Nerve Terminals with Calcium Indicators

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

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

Caricamento terminazioni nervose Drosophila con gli indicatori di calcio

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

Ricerca

Biologia

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 di calcio a livello 2 / 3 dei topi

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.

A cellula intera Recordings of Light Evocati eccitatorio Correnti Synaptic nel Slice Retinal

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 ...

Calcium Imaging of Cortical Neurons using Fura-2 AM

Calcium imaging is a common technique that is useful for measuring calcium signals in cultured cells. Calcium imaging techniques take advantage of calcium indicator dyes, which are BAPTA-based organic molecules that change their spectral properties in response to the binding of Ca2+ ions. Calcium indicator dyes fall into two categories, ratio-metric dyes like Fura-2 and Indo-1 and single-wavelength dyes like Fluo-4. Ratio-metric dyes change either their excitation or their emission spectra in response to calcium, allowing the concentration of intracellular calcium to be determined from the ratio of fluorescence emission or excitation at distinct wavelengths.  The main advantage of using ratio-metric dyes over single wavelength probes is that the ratio signal is independent of the dye concentration, illumination intensity, and optical path length allowing the concentration of intracellular calcium to be determined independently of these artifacts. One of the most common calcium indicators is Fura-2, which has an emission peak at 505 nM and changes its excitation peak from 340 nm to 380 nm in response to calcium binding.  Here we describe the use of Fura-2 to measure intracellular calcium elevations in neurons and other excitable cells.

Calcium signals play a key role in many cellular processes including gene expression, survival and differentiation. Here we demonstrate how to perform calcium imaging using Fura-2 AM. Calcium imaging is a valuable tool to study the regulation of intracellular calcium in real time and its regulation of signaling cascades.

Imaging di calcio di neuroni corticali usando Fura-02:00

Calcium imaging is a common technique that is useful for measuring calcium signals in cultured cells. Calcium imaging techniques take advantage of calcium ...

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.

Misura Vicino membrana plasmatica e Global Dynamics calcio intracellulare negli astrociti

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.

Monitoraggio Cambiamenti dei livelli di calcio mitocondriale durante l&#39;apoptosi mediante un sensore di calcio geneticamente codificato

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.

Misurare flussi di calcio veloce in cardiomiociti

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.

Valutazione del calcio Sparks in intatte fibre muscolari scheletriche

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.

Misure calcio citosolico nelle cellule epiteliali renali mediante citometria di flusso

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.

Registrazione in tempo reale In Vivo di Arabidopsis segnali di calcio durante l'alimentazione dell'insetto utilizzando un biosensore fluorescente

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