We are thankful to the Department of Physiology and Membrane Biology, University of California Davis for providing us with facilities and an excellent environment for the studies of ion channels.

1. Preparation of Resting Human T Lymphocytes
<ol>
 <li>Using RosetteSep Human T Cell Enrichment Cocktail and RosetteSep Density Medium, purify T cells from human blood samples according to the manufacturer's instructions. The resulting cell population should contain 95% CD3+ resting T cells. We purify human T lymphocytes from peripheral blood samples collected from healthy volunteers in accordance with the protocol approved by the UC Davis Internal Review Board.</li>
 <li>After isolation, resuspend resting T cells at a density of 0.5 x 106 cells/ mL in cell culture medium containing RPMI-1640 medium with glutamine and HEPES, supplemented with 10% fetal calf serum, 2% GlutaMAX-I, 1% RPMI 1640 vitamin solution, 1% RPMI 1640 amino acids solution, 1% sodium pyruvate, and 0.03% &beta;-mercaptoethanol.</li>
 <li>Place cell suspension into cell culture flasks and maintain at 37&deg;C in a humidified incubator with 5% CO2 for up to 24 h prior to the experiment.</li>
</ol>
2. Preparation of the Recording Chamber
<ol>
 <li>The following are needed to prepare recording chambers: a) silicon O-rings, b) round, 25-mm coverslips cleaned with 70% ethanol and distilled H2O, c) mixed Sylgard184, prepared according to the manufacturer's instructions, and d) two 60 x 15 mm Petri dishes.</li>
 <li>Place ~0.5 mL of mixed Sylgard184 into a 60x15-mm Petri dish. </li>
 <li>Dip the bottom part of the O-ring into the Sylgard184 using forceps.</li>
 <li>Remove excess Sylgard184 from the rim of the O-ring by placing the O-ring on a clean surface, such as another Petri dish. </li>
 <li>Place the ring onto the coverslip and press it down.</li>
 <li>Cure the Sylgard184 by heating the chambers at 85&deg; C for 40-60 min or until the Sylgard184 is polymerized.</li>
 <li>Store the recording chambers in a dust-free container.</li>
</ol>
3. Preparation of Patch Pipettes
<ol>
 <li>On the day of the experiment, pull pipettes with a ~2-&mu;m tip diameter using glass capillaries and a pipette puller. We use borosilicate tubing with filament (O.D.: 1.5 mm and I.D.: 1.10 mm). Store pipettes in a dust-free location.</li>
 <li>Coat pipette tips with Sylgard184 or HIPEC R6101 Semiconductor Protective Coating, according to a standard procedure 9.</li>
 <li>Gently fire-polish the pipettes before the experiment. </li>
</ol>
4. Patch Clamp Setup Preparation
<ol>
 <li>Configure and save the macros for gigaseal formation in the software controlling your patch-clamp amplifier. We use an EPC-10 amplifier supported by Windows XP and Pulse software for the data acquisition. We set the "set-up", "on-cell", and "whole-cell" macros according to the EPC-10 manual and use them for all experiments.</li>
 <li>Prepare bath solutions and pipette solutions listed in Table 1 prior to the experiment and store them at 4&deg; C for up to 1 week.</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>&nbsp;</td>
 <td># 1</td>
 <td># 2</td>
 <td># 3</td>
 <td>&nbsp;</td>
 </tr>
 </tbody>
 <tbody>
 <tr>
 <td>CH3SO3Na</td>
 <td>130</td>
 <td>110</td>
 <td>125</td>
 <td>-</td>
 </tr>
 <tr>
 <td>NaCl</td>
 <td>2</td>
 <td>4</td>
 <td>5</td>
 <td>-</td>
 </tr>
 <tr>
 <td>Ca(OH)2</td>
 <td>-</td>
 <td>20</td>
 <td>-</td>
 <td>-</td>
 </tr>
 <tr>
 <td>MgCl2</td>
 <td>3</td>
 <td>1</td>
 <td>-</td>
 <td>5</td>
 </tr>
 <tr>
 <td>MgSO4</td>
 <td>-</td>
 <td>-</td>
 <td>-</td>
 <td>2</td>
 </tr>
 <tr>
 <td>HEPES</td>
 <td>10</td>
 <td>10</td>
 <td>10</td>
 <td>15</td>
 </tr>
 <tr>
 <td>HEDTA</td>
 <td>-</td>
 <td>&nbsp;</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>
Table 1. Solutions for whole-cell CARC current recordings.
All chemicals were purchased from Sigma-Aldrich, St. Louis, MO.
<ol start="3">
 <li>Determine liquid junction potentials (LJ) between the patch pipette solution and each bath solution 10. Save the LJ value in the appropriate place using your acquisition software. For example, in the Pulse program insert the LJ value into "LJ" control in the "Amplifier" window.</li>
 <li>Configure and save the stimulation protocols, as shown in Figure 1, in the appropriate place of your acquisition software before the experiments. In the Pulse program, we set the acquisition protocol in the "Pulse Generation" window. <img src="/files/ftp_upload/2346/2364fig1.jpg" alt="Figure 1" /> Figure 1. Stimulation protocol. Voltage ramp from -120 to +100 mV of 50 ms in duration is applied every 0.2 - 2 s (5 - 0.5 Hz). Holding potential (Vh) between ramps is maintained at +30 mV.</li>
</ol>
5. Mounting Cells on the Microscope Stage
<ol>
 <li>Coat the recording chambers with poly-L-lysine for ~20 min and wash with H2O and then with Hank's 	balanced salt solution (HBSS) before plating the cells.</li>
<li>Plate 	500 &mu;L of cell suspension containing 0.2-0.5 x106 cells into the chamber and incubate for 20 min at 37&deg; C.</li>
<li>Secure 	the recording chamber with the attached cells in a chamber holder. 	We use a custom-made chamber holder, where the base supports the 	bottom of the coverslip, while the top part gently presses against 	the top of the coverslip. </li>
<li>Place 	the chamber holder on the stage of the inverted microscope.</li>
<li>Focus on the cells in transmitted light. We use a 40X oil immersion objective.</li>
<li>Place 	the Ag/AgCl reference electrode into the bath.</li>
<li>Align 	the multibarrel, gravity-driven perfusion system. The tip of the 	inflow tube should be placed at a 45&deg; angle to the coverslip close to the field of view.</li>
<li>Position 	the suction tube opposite to the tip of the inflow tube.</li>
<li>Perfuse 	the chamber with phosphate-buffered saline (PBS) or HBSS to wash 	away the cell culture media and loosely attached cells. </li>
</ol>
6. CRAC Channel Currents Recording
<ol>
 <li>Find a cell that is firmly attached to the coverslip. We typically select an average-sized, round cell with a smooth outer surface. </li>
 <li>Perfuse the recording chamber with Ca2+-free, 3 mM Mg2+-containing bath solution #1 (Table 1) containing 0.5-1 &mu;M thapsigargin for 8-10 min to block the SERCA pump. We perform this step to deplete the store prior to gigaseal formation. </li>
 <li>Using an Eppendorf microloader and a P-20 displacement pipette, fill the patch pipette with the pipette solution (Table 1) and insert the patch pipette into the pipette holder on the amplifier headstage. The Ca2+ chelator BAPTA is included in the pipette solution to passively deplete the store and to reduce Ca2+-dependent CRAC channel inactivation.</li>
 <li>Apply a small amount of positive pressure (~3 inches of H2O) inside the patch pipette before entering the bath. We use a custom-made pressure-suction device connected to pressurized air and vacuum lines to create positive or negative pressure inside a patch pipette. </li>
 <li>Lower the patch pipette into the bath under visual control via the microscope. In the "Oscilloscope" window, you should see a current flowing through the pipette. A step-like change in the current amplitude should be induced by a small amplitude voltage pulse (test pulse) applied by the preset macro. At this time, you should be able to determine the value of pipette resistance. The resistance of our pipettes is typically 5-6 M&#8486;.</li>
 <li>Correct the "offset" potential generated between the pipette and the reference electrode.</li>
 <li>Under visual control via the microscope, bring the pipette tip closer to the cell until it touches the cell membrane.</li>
 <li>Apply negative pressure (20-40 inches of H2O) inside the patch pipette. </li>
 <li>Monitor the gigaseal formation in the "oscilloscope" window. You will see the amplitude of the current, induced by the test pulse, decreases and the value of pipette resistance increases to &gt;5 G&Omega;. Usually gigaseal forms within a few seconds, but it may take 1-2 min to achieve a stable gigaseal.</li>
 <li>Compensate for the pipette capacitance ("C-Fast"). </li>
 <li>Break the patch membrane by applying additional negative pressure using a 20 cc syringe.</li>
 <li>Set the holding potential to +30 mV. A positive holding potential helps to prevent calcium-dependent CRAC channel inactivation.</li>
 <li>Compensate for the membrane capacitance of the cell ("C-slow"); save or record the value of C-slow. </li>
 <li>Check the value of the access resistance "Rs". In our experiments, the Rs is typically 7-20 M&Omega;.</li>
 <li>Start to apply a series of voltage ramps with a frequency of 0.5 Hz (Figure 1) in Ca2+-free 3 mM Mg2+-containing bath solution #1. In this solution, CRAC current is negligibly small as compared with the "leak" current due to the poor permeability of CRAC channels for Mg2+. Save and use the current traces recorded in bath solution #1 in future analyses as "leak" currents. The absolute amplitude of the "leak" current at -100 mV should be less than or equal to 5 pA. If the cell is "leaky", stop the experiment and start over using a new patch pipette and a new cell.</li>
 <li>To record Ca2+ current via CRAC channels (ICa-CRAC), replace Ca2+-free 3 mM Mg2+-containing bath solution #1 with 20 mM Ca2+-containing bath solution #2 (Table 1) by switching the perfusion valves. This allows for Ca2+ to flow via CRAC channels and produce an inward current. In the "Oscilloscope" window, you will see development of inwardly rectifying ICa-CRAC (Figure 2). The current amplitude at negative voltages will continue to increase for about 1 min due to the Ca2+-dependent potentiation of the CRAC current.</li>
 <li>Increase the frequency of stimulation with voltage ramps to 5 Hz, which is necessary to record the transient Na+ current via CRAC channels.</li>
 <li>Replace the 20 mM Ca2+-containing bath solution #2 with Na+-containing divalent cation-free solution #3 (Table 1) by switching the perfusion valves. In the "Oscilloscope" window, you will see a transient development of larger amplitude inwardly rectifying Na+ current via the CRAC channels (INa-CRAC; Figure 2).</li>
 <li>One may apply 20 mM Ca2+-containing bath solution #2 containing 1-5 &mu;M La3+ at the end of the experiment to record the "leak" current. However, we did not find this approach useful because the "leak" current changes over time and it may become larger or smaller at the end of the experiment. We prefer to take the current recorded in the Ca2+-free 3 mM Mg2+-containing bath solution # 1 at the beginning of the experiment as a "leak" current.</li>
 <li>Save the recorded current traces for further analysis.</li>
</ol>
7. Data Analysis
<ol>
 <li>Retrieve the saved current records into analysis software. We use the Pulse program for analysis.</li>
 <li>Measure the current amplitudes at the beginning and at the end of voltage ramps. We typically measure the current amplitudes at -100 mV and at +100 mV using pairs of cursors in the "Oscilloscope" window of the Pulse program. </li>
 <li>Export the values of current amplitudes into a graphic program for further analysis and graphical presentation. We use Origin Scientific Graphing and Analysis Software, version 7.</li>
</ol>
8. Representative Results
<img src="/files/ftp_upload/2346/2346fig2.jpg" alt="Figure 2" /> Figure 2. CRAC currents in a resting human T cell. (A), Time course of CRAC currents recorded in whole-cell voltage-clamp configuration at -100 mV (filled circles) and +100 mV (open circles). Prior to gigaseal formation, the cell was preincubated for ~10 min in Ca2+-free 3 mM Mg2+-containing bath solution #1 containing 0.5 &mu;M thapsigargin. After break-in, bath solutions were sequentially applied as follows: Ca2+-free 3 mM Mg2+-containing bath solution #1 (0 Ca + 3 Mg), followed by 20 mM Ca2+-containing bath solution #2 (20 Ca), followed by divalent cation-free bath solution #3 (DVF), followed by bath solution #2 (20 Ca). The cell was stimulated with a series of voltage ramps as shown in Figure 1. The frequency of ramps was 5 Hz in bath solution #3 (DVF) and 0.5 Hz in all other solutions. Note the slow development of ICa-CRAC after application of 20 mM Ca2+-containing bath solution #2 and the fast transient development of INa-CRAC in DVF bath solution #3. (B), Representative current traces recorded during voltage ramps in Ca2+-free 3 mM Mg2+-containing bath solution #1 ("leak"), 20 mM Ca2+-containing bath solution #2 (20 Ca), and bath solution #3 (DVF). (C, D), Current-voltage relationships of ICa-CRAC (C) and INa-CRAC (D) obtained by subtracting "leak" current from the currents recorded during voltage-ramps in 20 mM Ca2+-containing bath solution #2 (20 Ca), and bath solution #3 (DVF) shown in panel (B).

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

No conflicts of interest declared.

The electrophysiological investigation of CRAC currents in resting human T cells is a challenging task because the endogenous CRAC current amplitude in these cells is small due to the small cell size (the resting human T cell diameter is in the range of 5-8 &mu;m). Here, we present a step-by-step procedure to reliably record CRAC currents in resting human T lymphocytes isolated from peripheral blood mononuclear cells. This technology allows us to investigate the physiology and functional expression of CRAC channels in re...

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

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

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

We provide a step-by-step protocol for whole-cell patch clamp recording of Calcium Release-Activated Calcium (CRAC) currents in peripheral blood mononuclear cell-derived human T lymphocytes.

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

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

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

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16.0K Views.  University of California, Davis. We provide a step-by-step protocol for whole-cell patch clamp recording of Calcium Release-Activated Calcium (CRAC) currents in peripheral blood mononuclear cell-derived human T lymphocytes.

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.

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 ...

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 .

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.

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 ...

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.

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.

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.

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

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.

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.

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

Forward Genetic Screen Using Transgenic Calcium Reporter Aequorin to Identify Novel Targets in Calcium Signaling

Forward genetic screens have been important tools in the unbiased identification of genetic components involved in several biological pathways. The basis of the screen is to generate a mutant population that can be screened with a phenotype of interest. EMS (ethyl methane sulfonate) is a commonly used alkylating agent for inducing random mutation in a classical forward genetic screen to identify multiple genes involved in any given process. Cytosolic calcium (Ca2+) elevation is a key early signaling pathway that is activated upon stress perception. However the identity of receptors, channels, pumps and transporters of Ca2+ is still elusive in many study systems. Aequorin is a cellular calcium reporter protein isolated from Aequorea victoria and stably expressed in Arabidopsis. Exploiting this, we designed a forward genetic screen in which we EMS-mutagenized the aequorin transgenic. The seeds from the mutant plants were collected (M1) and screening for the phenotype of interest was carried out in the segregating (M2) population. Using a 96-well high-throughput Ca2+ measurement protocol, several novel mutants can be identified that have a varying calcium response and are measured in real time. The mutants with the phenotype of interest are rescued and propagated till a homozygous mutant plant population is obtained. This protocol provides a method for forward genetic screens in Ca2+ reporter background and identify novel Ca2+ regulated targets.

A forward genetic screen based on Ca2+ elevation as a read-out leads to identification of genetic components involved in calcium dependent signaling pathways in plants.

Forward genetic screens have been important tools in the unbiased identification of genetic components involved in several biological pathways. The basis ...

Real-Time Measurements of Calcium and Contractility Parameters in hiPSC-CMs

Real-Time Measurements of Calcium and Contractility Parameters in Human Induced Pluripotent Stem Cell-Derived Cardiomyocytes

Human induced pluripotent stem cell-derived cardiomyocytes (hiPSC-CMs) represent a powerful tool for studying mutation-mediated changes in cardiomyocyte function and defining the effects of stressors and drug interventions. In this study, it is demonstrated that this optics-based system is a powerful tool to assess the functional parameters of hiPSC-CMs in 2D. By using this platform, it is possible to perform paired measurements in a well-preserved temperature environment on different plate layouts. Moreover, this system provides researchers with instant data analysis.
This paper describes a method for measuring the contractility of unmodified hiPSC-CMs. Contraction kinetics are measured at 37 &#176;C based on pixel correlation changes relative to a reference frame taken at relaxation at a 250 Hz sampling frequency. Additionally, simultaneous measurements of intracellular calcium transients can be acquired by loading the cell with a calcium-sensitive fluorophore, such as Fura-2. Using a hyperswitch, ratiometric calcium measurements can be performed on a 50 &#181;m diameter illumination spot, corresponding to the area of the contractility measurements.

Author Spotlight: Real-Time Measurements of Calcium and Contractility Parameters in Human Induced Pluripotent Stem Cell-Derived Cardiomyocytes  

Here, an established method is described to perform contractility and calcium measurements in human induced pluripotent stem cell-derived cardiomyocytes using an optics-based platform. This platform enables researchers to study the effect of mutations and the response to various stimuli in a rapid and reproducible manner.

Human induced pluripotent stem cell-derived cardiomyocytes (hiPSC-CMs) represent a powerful tool for studying mutation-mediated changes in cardiomyocyte ...