This work was funded by NIH GM62999, NIH EB007268, NIH AG15820, B&amp;B Seed Grant, and a NIH Supplemental Grant to FR, BB fellowship to CM, CDT fellowship to RG.

1. Slide Preparation
<ol>
	<li>Place 22 mm x 40 mm glass microscope slides in 6 cm cell culture dishes, 1 slide per dish.</li>
	<li>Expose each side of each slide as well as the 6 cm dish to ultraviolet (UV) light for 15-20 min to sterilize in a sterile hood.</li>
	<li>Cover the dishes with slides inside with parafilm and store at 4 &#176;C until ready to use.</li>
</ol>
2. Preparation of Media, Buffers, Solutions, and Reagents
<ol>
	<li>Prepare 1 L of Hank&#39;s balanced salt solution (HBSS) in deionized water and add 10 mM HEPES, 5 mM NaHCO3, and 1 mM EGTA. Adjust to pH 7.2-7.3 with NaOH. Filter buffer with a 0.22 &#181;m filter into an autoclaved bottle.</li>
	<li>Prepare 900 mL of high glucose Dulbecco&#39;s Modified Eagle&#39;s Medium (DMEM) h in deionized water. Add 44 mM NaHCO3 and adjust the pH to 7.2-7.3 with HCl. Filter media with a 0.22 &#181;m filter into an autoclaved bottle. Add 100 mL of fetal bovine serum (FBS) to the media to make the FBS concentration 10%. Store at 4 &#176;C.</li>
	<li>Prepare 1 L of intracellular buffer (125 mM KCl, 25 mM NaCl, 10 mM HEPES, 0.2 mM CaCl2, 0.5 mM EGTA, 0.2 mM MgCl2) in deionized water, and adjust the pH to 7.25. The final [Ca2+] will be ~100 nM. Before use, add 0.5 mM ATP. Store at room temperature.</li>
	<li>Prepare 1 L of Ringer&#39;s buffer (121 mM NaCl, 2.4 mM K2HPO4, 0.4 mM KH2PO4, 10 mM HEPES, and 1.2 mM MgCl2) in deionized water, and adjust the pH to 7.2. Store at room temperature. To use, aliquot 50 mL into a 50 mL tube and add 1.8 mM Ca2+ and 10 mM glucose.</li>
	<li>Prepare 1 L of KCl rinse solution (125 mM KCl, 25 mM NaCl, 10 mM HEPES, 0.2 mM MgCl2) in deionized water, and adjust the pH to 7.25.</li>
	<li>Dissolve 5 mg of ionomycin in 1 mL of dimethyl sulfoxide (DMSO). Aliquot 30 &#181;L into microtubes and store at -20 &#176;C.</li>
	<li>Prepare the cyclopiazonic acid (CPA) stock by dissolving CPA into DMSO. Make sure the final percent of DMSO is &#8804; 1% for the CPA added to the cells to prevent apoptosis. Prepare 20 mM 4-chloro-m-cresol (4-cmc) by dissolving into filtered H2O and letting it dissolve overnight by shaking at 37 &#176;C. Prepare the ATP stock by dissolving in filtered H2O to 100 mM.</li>
	<li>To determine the in situ Kd, prepare 15 mL each of 1 mM EGTA, 0.2 mM, 0.6 mM, 2 mM, 5 mM, 10 mM, 20 mM, 50 mM, and 100 mM Ca2+ in KCl buffer using a 100 mM EGTA stock and a 1 M CaCl2 stock dissolved in deionized water.</li>
</ol>
3. Cell Culture
<ol>
	<li>Culture C2C12 myoblast and HEK293 cells according to ATCC protocols for each using 10 cm cell culture dishes in a sterile UV hood. 
	NOTE: C2C12 cells should not be allowed to grow higher than ~70% confluence since the myoblasts will begin to differentiate into myotubes. Both HEK293 and C2C12 cells grow readily and should not be allowed to grow past 100% confluency to maintain cell integrity. Additionally, cells should not be passaged more than 10 times before using them for an experiment to preserve cell health.</li>
</ol>
4. Transfection of HEK293 Cells
<ol>
	<li>Seed HEK293 cells onto sterilized 22 mm x 40 mm glass microscope slides in 6 cm dishes so they are ~70% confluent the day of transfection.</li>
	<li>The next day, follow the manufacturer&#39;s protocol for the transfection reagent to transfect cells using 2 &#181;g of CatchER+ cDNA and a 1:3 (weight/volume) DNA:transfection reagent ratio in the reduced serum media (e.g. Opti-MEM).</li>
	<li>Empty the DMEM media from the dish, and add 3 mL of reduced serum media. Add DNA:transfection reagent mixture to the dish and incubate for 4-6 h at 37 &#176;C.</li>
	<li>After incubation, wash the cells with 5-6 mL of HBSS. Discard the HBSS from the dish and replace with 3 mL of fresh DMEM and incubate them at 37 &#176;C for 48 h to allow expression of the GECI.</li>
	<li>Transfection of C2C12 myoblast cells
	<ol>
		<li>For C2C12 myoblast cells, trypsinize cells by adding enough trypsin to cover the bottom of dish (1-2 mL). Place the dish in a 37 &#176;C incubator for 2-6 min to allow the trypsin to loosen the cells from the bottom of the dish.</li>
		<li>Once the incubation is complete, remove the trypsin and aspirate the cells in 8 mL of DMEM. Pipette the cells thoroughly with the DMEM to evenly disperse and re-suspend te cells and seed them onto sterile coverslips in 6 cm cell culture dishes containing 3 mL of DMEM so that the final confluence is 60%.</li>
		<li>Transfect the cells the same day as outlined in step 4.2-4.3. Incubate for 24 h at 37 &#176;C.</li>
		<li>Repeat step 4.4.</li>
	</ol>
	</li>
</ol>
5. Preparation of Slide and Fluorescence Microscope
<ol>
	<li>Turn on microscope and light source, then open the Simple PCI program. Allow the microscope to warm up for 15-20 min or until the charge-coupled device (CCD) camera cools to -65 &#176;C. Prepare the chamber and slide with cells while waiting.</li>
	<li>Cover the bottom of the low profile open diamond bath imaging chamber with a thin layer of sealant, using a cotton swab.</li>
	<li>Take the slide containing transfected cells out of the incubator and gently rinse three times with 1 mL of Ringer&#39;s buffer with 10 mM glucose and 1.8 mM Ca2+ preheated to 37 &#176;C.</li>
	<li>Leave 1 mL of Ringer&#39;s buffer in the dish, to prevent cells from drying, and use forceps to take the slide out of the dish. Allow excess solution to absorb onto a laboratory tissue by touching the edge of the slide to a laboratory tissue. Then place onto the side of the chamber containing the grease with cells facing up towards grease and secure chamber onto the stage mount using a screwdriver.</li>
	<li>Add 1 mL of Ringer&#39;s buffer to the chamber to prevent cells from drying while mounting the chamber onto the stage and completing the rest of the microscope setup. Once the vacuum suction hose is attached to the chamber, the final volume the chamber holds is ~450 &#181;L.</li>
	<li>Switch the microscope objective to the 40X oil immersion objective.</li>
	<li>Use lens paper and lens cleaning solution to clean and dry the objective. Do not rub the objective. Dab gently until the surface is clean.</li>
	<li>Add one drop of the immersion oil to the objective.</li>
	<li>Place the mount onto the microscope stage and add the vacuum tip to the chamber. Once the vacuum suction hose is attached to the chamber and turned on, the final volume the chamber holds is ~450 &#181;L. This final volume is level with the sides of the chamber. It is imperative for the chamber solution to be level, during the experiment, to prevent the solution from spilling down into the objective due to overfilling.</li>
	<li>Raise the objective and use the coarse adjustment until the oil on the objective touches the bottom of the slide.</li>
	<li>Using the bright field mode, adjust the gain to 175 and the exposure time to 0.03 s to focus the cells.</li>
	<li>Once cells are focused, switch to fluorescence mode using 488 nm excitation. Change the exposure time to 0.07 s. Locate a field of view that has enough cells that are healthy with adequate fluorescence to image.</li>
	<li>Once a field of view has been chosen, take pictures in fluorescence and bright field mode.</li>
	<li>Circle a region of interest (ROI) in the cells or circle the whole cell to record the intensity from the chosen area.</li>
</ol>
6. Imaging Drug-induced Ca2+ Transients and In Situ Kd Calibration
<ol>
	<li>Start intensity measurement with frame rate set at one every 5 s.</li>
	<li>Allow the baseline intensity to stabilize for 20-30 frames (100-150 s). If needed, decrease the frames/s during this step to prevent photobleaching.</li>
	<li>Calculate the amount of 4-cmc needed from a 20 mM stock based on the final chamber volume of ~450 &#181;L to make a final concentration of 200 &#181;M. Make dilutions of the stock as necessary.</li>
	<li>Carefully take 60 &#181;L of Ringer&#39;s buffer from the chamber and place in a microcentrifuge tube. Dilute the calculated amount of 4-cmc with this solution and add back to the chamber to evoke the intended response from the cells being imaged.</li>
	<li>Add the calculated amount of 4-cmc to the microcentrifuge tube and mix by pipetting.</li>
	<li>Gently add the solution over the field of view while simultaneously adding the event marker in the intensity measurement.</li>
	<li>Allow time for the drug to bind to the receptor and subsequent signal decrease, then wash with 3-6 mL of Ringer&#39;s buffer with 1.8 mM Ca2+ and 10 mM glucose while simultaneously adding the event marker. Since the chamber volume is 450 &#181;L and is connected to a vacuum, adding 3-6 mL of buffer ensures that the former solution containing the drug has been washed away and replaced with the newly added solution.</li>
	<li>Repeat 6.1-6.7 for 100 &#181;M ATP and 15 &#181;M CPA, using a new slide of transfected cells for each assay.</li>
	<li>Once the measurement is complete, end the data collection in the intensity measurement window in the Simple PCI program.</li>
	<li>Take fluorescence and brightfield pictures of the cells.</li>
	<li>Open the data file for the experiment. Here, re-circle the cells or regions of interest to re-calculate the intensity values, if needed.</li>
	<li>To determine the in situ Kd in C2C12 myoblast cells, follow the protocol as outlined in section 4.2 and section 5 and put 1 mL of 0 mM Ca2+ Ringer&#39;s buffer in the chamber.
	<ol>
		<li>Permeabilize the cells with 0.002-0.005% saponin in intracellular buffer for 15-30 s to empty the cells of any CatchER+ that may be in the cytosol. Wash the cells with 3-6 mL of 1 mL EGTA in KCl buffer with 10 &#181;M ionomycin. Allow the intensity to decrease until a plateau is reached.</li>
		<li>Rinse with 3-6 mL of KCl buffer with 10 &#181;M ionomycin and allow the intensity to stabilize, then add each Ca2+ solution detailed in step 2.7. Allow the intensity to reach a plateau between each addition.</li>
	</ol>
	</li>
	<li>To calibrate the sensor for basal [Ca2+] quantification, follow step 6.12. Use the EGTA and the 100 mM Ca2+ buffer from step 2.7 to get the minimum and maximum fluorescence intensity.</li>
</ol>
7. Data Processing
<ol>
	<li>Download the spreadsheet file of the intensity measurement data.</li>
	<li>Normalize the data for each cell to the baseline intensity (F/F0). Plot the normalized data versus time in any graphing program.</li>
	<li>To calculate the % change, subtract the normalized peak value from 1 and multiply by 100. Calculate the average and standard deviation if multiple cells were imaged.</li>
	<li>To calculate the signal to noise ratio, SNR, open up the image file saved from the experiment in an image processing software, such as ImageJ, and then measure the signal, transfected cells, and noise, background. Calculate the SNR using the equation SNR = signal/noise.</li>
	<li>To calculate the in situ Kd from the collected fluorescence imaging data, average the intensity data, comprising the plateaus, after the addition of each Ca2+ concentration and EGTA (0 mM Ca2+). Calculate the fractional saturation (relative intensity) using &#952; = (F &#8211; Fmin)/(Fmax- Fmin), where &#952; is the relative intensity, F is the fluorescence at any point, Fmin is the minimum fluorescence intensity, and Fmax is the maximum fluorescence intensity, to see the change in the intensity of the probe with the addition of Ca2+ compared to the minimum intensity. Plot the relative intensity data against the [Ca2+] in any graphing and curve fitting program. Use the 1:1 binding equation &#952; = [MT]/(Kd + [MT]) translated for any graphing and curve fitting program to calculate the Kd. MT is the total metal concentration.</li>
	<li>To quantify the basal calcium from the calibration outlined in 6.13, average the intensity data, comprising the plateaus, after the addition of 1 mM EGTA (Fmin) and 100 mM Ca2+ (Fmax). Use the calibration equation [Ca2+] = Kd([F &#8211; Fmin)/(Fmax &#8211; F)] to calculate the basal calcium.</li>
</ol>

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J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Light+microscopy+techniques+for+live+cell+imaging.">Light microscopy techniques for live cell imaging.</a> Science. 300 (5616), 82-86 (2003).</li><li>Bootman, M. D., Rietdorf, K., Collins, T., Walker, S., Sanderson, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Ca2+-sensitive+fluorescent+dyes+and+intracellular+Ca2++imaging.">Ca2+-sensitive fluorescent dyes and intracellular Ca2+ imaging.</a> Cold Spring Harb. 2013 (2), 83-99 (2013).</li></ol>

The authors have nothing to disclose.

Live single-cell imaging of fluorescent probes, such as CatchER+, is an effective technique to analyze intricate ER/SR Ca2+ signaling processes in each cell in response to receptor agonists or antagonists. This technique is also useful for imaging using multiple wavelengths concurrently, such as needed for Fura-2 or to image CatchER+ and Rhod-2 together to monitor both ER and cytosolic calcium changes, respectively. There are several critical steps in this protocol; cell transfection can ...

The spatio-temporal attributes of intracellular calcium (Ca2+) transients activate various biological functions1. These Ca2+ signaling events are triggered extracellularly through different stimuli and controlled intracellularly by the major Ca2+ storage organelle and by numerous Ca2+ pumps, channels, and Ca2+ binding proteins. Ca2+ transients can be significantly altered as a result of defects with signal modulation, leading to different diseases2. Because of the speed and intricacy of the Ca2+ signaling system, with the endo- (ER) and sarcoplasmic reticulum (SR) at the center, genetically-encoded Ca2+ probes that have been optimized for mammalian expression with fast kinetics are needed to observe global and local Ca2+ changes in different cells3.
The ER and the SR, its counterpart in muscle cells, are the major intracellular Ca2+ storage organelles and act as Ca2+ sinks that help to amplify the Ca2+ signal4. The ER/SR is an integral part in Ca2+ signaling with dual roles as a transmitter and receiver of signals5. The ryanodine receptor (RyR) and the inositol 1,4,5-triphosphate receptor (IP3R) are Ca2+ release receptors located on the membranes of the ER/ SR that are regulated by Ca2+ 6. Other agents directly or indirectly stimulate the function of these receptors. 4-chloro-m-cresol (4-cmc) is a potent agonist of the RyR, having a 10 fold higher sensitivity than caffeine for inducing SR Ca2+ release where both are regularly employed to study RyR-mediated Ca2+ release in healthy and diseased cells7. ATP increases IP3-mediated Ca2+ release through the IP3R8. ATP binds to the purinergic receptor P2YR, a G-protein coupled receptor (GPCR), triggering the production of IP3 that binds to the IP3R to release Ca2+ from the ER9,10. The sarco-endoplasmic reticulum calcium ATPase (SERCA) pump is a P-type ATPase pump, also located on the ER/SR membrane that reduces cytosolic Ca2+ and refills the ER/SR by actively pumping the ion into the ER/SR lumen11. Specific inhibitors of the SERCA pump include thapsigargin, from Thapsia garganica, and cyclopiazonic acid (CPA), from Aspergillus and Penicillium. CPA has a low affinity for the pump and reversibly blocks the Ca2+ access point12. Thapsigargin, on the other hand, irreversibly binds to the Ca2+ free pump at residue F256 in the M3 helix with nanomolar affinity11. Analyzing and quantifying the changes involved in Ca2+ stimulated events has been and remains a challenge. Since the ER/SR is the major subcellular Ca2+ containing compartment with a central function in the propagation of the Ca2+ signal, much work has been focused on understanding ER/SR Ca2+ signaling5.
The creation of synthetic Ca2+ dyes helped to advance the field and practice of Ca2+ imaging. Although dyes, such as Mag-Fura-2, have been widely used to measure compartmentalized Ca2+ in different cells,13,14,15 they have limitations such as uneven dye loading, photobleaching, and the inability to be targeted to specific organelles. The discovery of the green fluorescent protein (GFP) and the advancement of fluorescent protein-based Ca2+ probes has propelled the field of Ca2+ imaging forward16. Some of the existing GECIs are F&#246;rster resonance energy transfer (FRET) pairs involving yellow fluorescent protein (YFP), cyan fluorescent protein (CFP), calmodulin and the M13 binding peptide17,18. Troponin C-based GECIs are also available as FRET pairs of CFP and Citrine and as single fluorophore probes19,20,21. Others, such as GCaMP2 and R-GECO are single fluorophore sensors involving calmodulin22,23. To overcome the limitations of narrow tuning of Kd&#39;s and cooperative binding associated with multiple Ca2+ binding sites found in their Ca2+ binding domains24, a novel class of calcium sensors was created by designing a Ca2+ binding site on the surface of the beta barrel in a chromophore sensitive location of enhanced green fluorescent protein (EGFP)25,26. This highly touted sensor, called CatchER, has a Kd of ~0.18 mM, a kon near the diffusion limit, and a koff of 700 s-1. CatchER has been used to monitor receptor-mediated ER/SR calcium release in different mammalian cell lines such as HeLa, HEK293, and C2C1225. Because of its fast kinetics, CatchER was used in flexor digitorum brevis (FDB) muscle fibers of young and old Friend Virus B NIH Jackson (FVB) mice to reveal that more Ca2+ remains in the SR after 2 s of depolarization in the FDB fibers of old mice compared to that of young mice27. To overcome its low fluorescence at 37 &#176;C, which hinders its applications in calcium imaging of mammalian cells, we have developed an improved version of CatchER called CatchER+. CatchER+ exhibits enhanced fluorescence at 37 &#176;C for better application in mammalian cells. Additional mutations were incorporated into CatchER to improve the thermostability and fluorescence at 37 &#176;C28,29, to create CatchER+. CatchER+ exhibits a six-fold increase in its signal to noise ratio (SNR) over CatchER30.
Here, the protocols for the culture and transfection of HEK293 and C2C12 cells with CatchER+ and its application for monitoring ER/SR receptor-mediated calcium transients are presented. Representative results are shown for CatchER+ expressed in HEK293 cells treated with 4-cmc, CPA and ATP. We also provide a protocol for determining the in situ Kd of CatchER+ in C2C12 myoblast cells and quantification of basal [Ca2+].

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			<td>4-Chloro-3-methylphenol (4-CmC)</td>
			<td>Sigma-Aldrich</td>
			<td>C55402</td>
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			<td>515DCXR dichroic mirror</td>
			<td>Chroma Technology Corp.</td>
			<td>NC338059</td>
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			<td>Adenosine 5&#8242;-triphosphate disodium salt hydrate</td>
			<td>Sigma-Aldrich</td>
			<td>A26209</td>
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			<td>Calcium chloride dihydrate</td>
			<td>EMD Millipore</td>
			<td>102382</td>
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		<tr>
			<td>Corning tissue-culture treated culture dishes (100 mm)</td>
			<td>Sigma-Aldrich</td>
			<td>CLS430167</td>
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		<tr>
			<td>Corning tissue-culture treated culture dishes (60 mm)</td>
			<td>Sigma-Aldrich</td>
			<td>CLS430166</td>
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			<td>Cyclopiazonic Acid (CPA)</td>
			<td>EMD Millipore</td>
			<td>239805</td>
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			<td>D(+)-Glucose</td>
			<td>ACROS Organics</td>
			<td>41095-0010</td>
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			<td>Dow Corning 111 Valve Lubricant &#38; Sealant</td>
			<td>Warner Instruments</td>
			<td>64-0275</td>
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		<tr>
			<td>Dulbecco&#8217;s Modified Eagle&#8217;s Medium (DMEM)</td>
			<td>Sigma-Aldrich</td>
			<td>D7777</td>
		</tr>
		<tr>
			<td>Ethylenebis(oxyethylenenitrilo) 
			tetraacetic Acid (EGTA)</td>
			<td>ACROS Organics</td>
			<td>409911000&#160;</td>
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		<tr>
			<td>Fetal Bovine Serum (FBS)</td>
			<td>ThermoFisher</td>
			<td>26140087</td>
		</tr>
		<tr>
			<td>Fisherbrand Cover Glasses 22x40 mm</td>
			<td>Fisher Scientific</td>
			<td>&#160;12-544B</td>
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		<tr>
			<td>Hanks&#8217; Balanced Salts (HBSS)</td>
			<td>Sigma-Aldrich</td>
			<td>H4891</td>
		</tr>
		<tr>
			<td>HEPES, Free Acid, Molecular Biology Grade</td>
			<td>EMD Millipore</td>
			<td>391340</td>
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		<tr>
			<td>Immersion Oil without autofluorescence</td>
			<td>Leica</td>
			<td>11513859</td>
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		<tr>
			<td>Ionomycin, Free Acid</td>
			<td>Fisher Scientific</td>
			<td>50-230-5804</td>
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		<tr>
			<td>Leica DM6100B inverted microscope with a cooled EM-CCD camera</td>
			<td>Hamamatsu</td>
			<td>C9100-13</td>
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		<tr>
			<td>Lipofectamine 2000 Transfection Reagent</td>
			<td>ThermoFisher</td>
			<td>11668019</td>
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			<td>Lipofectamine 3000 Transfection Reagent</td>
			<td>ThermoFisher</td>
			<td>L3000015</td>
		</tr>
		<tr>
			<td>Low Profile Open Diamond Bath Imaging Chamber</td>
			<td>Warner Instruments</td>
			<td>RC-26GLP</td>
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			<td>Magnesium Chloride Hexahydrate</td>
			<td>Fisher Scientific</td>
			<td>M33-500</td>
		</tr>
		<tr>
			<td>Opti-MEM</td>
			<td>ThermoFisher</td>
			<td>51985034</td>
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		<tr>
			<td>Potassium Chloride</td>
			<td>EMD Millipore</td>
			<td>PX1405</td>
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		<tr>
			<td>Potassium Phosphate, Dibasic</td>
			<td>EMD Millipore</td>
			<td>PX1570</td>
		</tr>
		<tr>
			<td>Potassium Phosphate, Monobasic</td>
			<td>EMD Millipore</td>
			<td>PX1565</td>
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		<tr>
			<td>Saponin</td>
			<td>Sigma-Aldrich</td>
			<td>47036</td>
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		<tr>
			<td>SimplePCI Image Analysis Software</td>
			<td>Hamamatsu</td>
			<td>N/A</td>
		</tr>
		<tr>
			<td>Sodium Bicarbonate</td>
			<td>Fisher Scientific</td>
			<td>S233-3</td>
		</tr>
		<tr>
			<td>Sodium Chloride</td>
			<td>Fisher Scientific</td>
			<td>S271-500</td>
		</tr>
		<tr>
			<td>Sterivex-GV 0.22 &#181;m filter</td>
			<td>EMD Millipore</td>
			<td>SVGVB1010</td>
		</tr>
		<tr>
			<td>Till Polychrome V Xenon lamp</td>
			<td>Till Photonics</td>
			<td>N/A</td>
		</tr>
		<tr>
			<td>Trypsin (2.5%), no phenol red (10X)</td>
			<td>ThermoFisher</td>
			<td>15090046</td>
		</tr>
	</tbody>
</table>

monitoring er/sr calcium release with the targeted ca2+ sensor catcher+

Intracellular calcium (Ca2+) transients evoked by extracellular stimuli initiate a multitude of biological processes in living organisms. At the center of intracellular calcium release are the major intracellular calcium storage organelles, the endoplasmic reticulum (ER) and the more specialized sarcoplasmic reticulum (SR) in muscle cells. The dynamic release of calcium from these organelles is mediated by the ryanodine receptor (RyR) and the inositol 1,4,5-triphosphate receptor (IP3R) with refilling occurring through the sarco/endoplasmic reticulum calcium ATPase (SERCA) pump. A genetically encoded calcium sensor (GECI) called CatchER was created to monitor the rapid calcium release from the ER/SR. Here, the detailed protocols for the transfection and expression of the improved, ER/SR-targeted GECI CatchER+ in HEK293 and C2C12 cells and its application in monitoring IP3R, RyR, and SERCA pump-mediated calcium transients in HEK293 cells using fluorescence microscopy is outlined. The receptor agonist or inhibitor of choice is dispersed in the chamber solution and the intensity changes are recorded in real time. With this method, a decrease in ER calcium is seen with RyR activation with 4-chloro-m-cresol (4-cmc), the indirect activation of IP3R with adenosine triphosphate (ATP), and inhibition of the SERCA pump with cyclopiazonic acid (CPA). We also discuss protocols for determining the in situ Kd and quantifying basal [Ca2+] in C2C12 cells. In summary, these protocols, used in conjunction with CatchER+, can elicit receptor mediated calcium release from the ER with future application in studying ER/SR calcium related pathologies.

This section will illustrate the results that were achieved using the previously described methods using the optimized ER/SR-targeted GECI CatchER+ to monitor changes in ER/SR Ca2+ through different receptor mediated pathways.
Figure 1 illustrates ER emptying through the RyR stimulated with 200 &#181;M 4-cmc. 4-cmc is an agonist of the RyR. Addition of the drug induces a decrea...

Watch this Scientific Journal Video about Monitoring ER/SR Calcium Release with the Targeted Ca2+ Sensor CatchER+ at JoVE.com

Monitoring ER/SR Calcium Release with the Targeted Ca2+ Sensor CatchER+

We present protocols for the application of our targeted genetically-encoded calcium indicator (GECI) CatchER+ for monitoring rapid calcium transients in the endoplasmic/sarcoplasmic reticulum (ER/SR) of HEK293 and skeletal muscle C2C12 cells using real-time fluorescence microscopy. A protocol for the in situ Kd measurement and calibration is also discussed.

Monitoring ER/SR Calcium Release with the Targeted Ca2+ Sensor CatchER+

Intracellular calcium (Ca2+) transients evoked by extracellular stimuli initiate a multitude of biological processes in living organisms. At the center of ...

monitoring-ersr-calcium-release-with-the-targeted-ca2-sensor-catcher

Research

JoVE Journal

Biology

14.8K Views.  Georgia State University. We present protocols for the application of our targeted genetically-encoded calcium indicator (GECI) CatchER+ for monitoring rapid calcium transients in the endoplasmic/sarcoplasmic reticulum (ER/SR) of HEK293 and skeletal muscle C2C12 cells using real-time fluorescence microscopy. A protocol for the in situ Kd measurement and calibration is also discussed.

Video: Monitoring ER/SR Calcium Release with the Targeted Ca2+ Sensor CatchER+

Monitoring Actin Disassembly with Time-lapse Microscopy

Okay, so what I'd like to do today is to show you how to construct a flow chamber, a fusion chamber for imaging on an upright microscope. What then I then do is I create two spaces using film. So what I do is I take the para film, I cut it into small strips.Next I'm going to, this will basically form the channels for the, the walls, and I dry them with a tilted air. I put the cover slip on top of the para, And here's the finished. Then I heat it on a heat block to melt the param and basically create a seal.So what this is, is the, the cover glass here, and then on top of the cover glass is two layers of phim as you see right here. And then on top of this, again, there's a, it's a 22 by two two mil meter cover slip. This one right here, which I pressed against the param to fall the seal.Okay, so the liquid goes in in one direction here and I can suck it out with a filter paper here in this direction. And I'll show you how to do that in a bit. So what I like to do is I like to put it in a humidified chamber, which is very simple.Take two kind of elevated strips and have a filter of paper underneath. And to keep it noisy as I, I put the cover, cover slip on the flow chamber on. Now the way you introduce a solution to the chamber is that you just put the pipet tip at the entrance of the chamber and you just split.Now what I do is I incubate the chamber with the lid closed for 10 minutes. What I usually do to suck the liquid out from the other side is to use it, get a filter paper and cut it into thin strips like this. And I take one of these thin strips and I simultaneously apply the solution into the entrance while putting the filter paper at the exit to suck the exchange, the solution.So you'll see the liquid flowing through the solution. And then I wait for five minutes for the mid close. Now what I'm ready to do is to actually put that the actin it and the actin, I actually have to polymerize, I polymerize inside the flow chamber and it's just the same procedures before.So because like this, and then now that the action is polymerized, I wash out all the un excess un polymerized action using a large volume of just buffer. So here I have about a hundred microliters and it's a few reaction chamber volumes worth of buffer. Just, you know, when the filter paper becomes saturated, I have to change the direction.Okay?And so I'm ready to do imaging with this. So perfusion chamber with actin attached to it. Now I take the slide, mount it under the microscope, and since I'm imaging with an oil objective, I'm gonna put some immersion oil on the top surface of the cover, grass touch, such so assess solution or protein or anything on acting inside.So what I do is I can actually exchange the solution side while I'm doing the time lapse imaging. As such.

Monitoring Actin Demontaż z poklatkowy Mikroskopia

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 ...

Functional Imaging with Reinforcement, Eyetracking, and Physiological Monitoring

We use functional brain imaging (fMRI) to study neural circuits that underlie decision-making.  To understand how outcomes affect decision processes, simple perceptual tasks are combined with appetitive and aversive reinforcement.  However, the use of reinforcers such as juice and airpuffs can create challenges for fMRI.  Reinforcer delivery can cause head movement, which creates artifacts in the fMRI signal.  Reinforcement can also lead to changes in heart rate and respiration that are mediated by autonomic pathways.  Changes in heart rate and respiration can directly affect the fMRI (BOLD) signal in the brain and can be confounded with signal changes that are due to neural activity.  In this presentation, we demonstrate methods for administering reinforcers in a controlled manner, for stabilizing the head, and for measuring pulse and respiration.

This presentation demonstrates the use of fMRI to study neural circuits that underlie decision-making. Simple perceptual tasks are combined with appetitive and aversive reinforcements to investigate how outcomes affect decision processes.

We use functional brain imaging (fMRI) to study neural circuits that underlie decision-making.  To understand how outcomes affect decision processes, ...

Dopamine Release at Individual Presynaptic Terminals Visualized with FFNs

The nervous system transmits signals between neurons via neurotransmitter release during synaptic vesicle fusion. To observe neurotransmitter uptake and release from individual presynaptic terminals directly, we designed fluorescent false neurotransmitters as substrates for the synaptic vesicle monoamine transporter. Using these probes to image dopamine release in the striatum, we made several observations pertinent to synaptic plasticity. We found that the fraction of synaptic vesicles releasing neurotransmitter per stimulus was dependent on the stimulus frequency. A kinetically distinct "reserve" synaptic vesicle population was not observed under these experimental conditions. A frequency-dependent heterogeneity of presynaptic terminals was revealed that was dependent in part on D2 dopamine receptors, indicating a mechanism for frequency-dependent coding of presynaptic selection.

Hui Zhang and Niko G. Gubernator contributed equally to this work.

A new means to measure neurotransmission optically using fluorescent dopamine analogs.

The nervous system transmits signals between neurons via neurotransmitter release during synaptic vesicle fusion. To observe neurotransmitter uptake and ...

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.

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.

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

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, ...

Live Cell Calcium Imaging Combined with siRNA Mediated Gene Silencing Identifies Ca2+ Leak Channels in the ER Membrane and their Regulatory Mechanisms

In mammalian cells, the endoplasmic reticulum (ER) plays a key role in protein biogenesis as well as in calcium signalling1. The heterotrimeric Sec61 complex in the ER membrane provides an aqueous path for newly-synthesized polypeptides into the lumen of the ER. Recent work from various laboratories suggested that this heterotrimeric complex may also form transient Ca2+ leak channels2-8. The key observation for this notion was that release of nascent polypeptides from the ribosome and Sec61 complex by puromycin leads to transient release of Ca2+ from the ER. Furthermore, it had been observed in vitro that the ER luminal protein BiP is involved in preventing ion permeability at the level of the Sec61 complex9,10. We have established an experimental system that allows us to directly address the role of the Sec61 complex as potential Ca2+ leak channel and to characterize its putative regulatory mechanisms11-13. This system combines siRNA mediated gene silencing and live cell Ca2+ imaging13. Cells are treated with siRNAs that are directed against the coding and untranslated region (UTR), respectively, of the SEC61A1 gene or a negative control siRNA. In complementation analysis, the cells are co-transfected with an IRES-GFP vector that allows the siRNA-resistant expression of the wildtype SEC61A1 gene. Then the cells are loaded with the ratiometric Ca2+-indicator FURA-2 to monitor simultaneously changes in the cytosolic Ca2+ concentration in a number of cells via a fluorescence microscope. The continuous measurement of cytosolic Ca2+ also allows the evaluation of the impact of various agents, such as puromycin, small molecule inhibitors, and thapsigargin on Ca2+ leakage. This experimental system gives us the unique opportunities to i) evaluate the contribution of different ER membrane proteins to passive Ca2+ efflux from the ER in various cell types, ii) characterize the proteins and mechanisms that limit this passive Ca2+ efflux, and iii) study the effects of disease linked mutations in the relevant components.

Live Cell Calcium Imaging Combined with siRNA Mediated Gene Silencing Identifies Ca2+ Leak Channels in the ER Membrane and their Regulatory Mechanisms

The endoplasmic reticulum plays a key role in protein biogenesis and in calcium homeostasis. We have established an experimental system that allows us to address the role of Ca2+ leak channels and to characterize their putative regulatory mechanisms. This system involves siRNA mediated gene silencing and live cell Ca2+ imaging.

In mammalian cells, the endoplasmic reticulum (ER) plays a key role in protein biogenesis as well as in calcium signalling1. The heterotrimeric Sec61 ...

Long-term Lethal Toxicity Test with the Crustacean Artemia franciscana

Our research activities target the use of biological methods for the evaluation of environmental quality, with particular reference to saltwater/brackish water and sediment. The choice of biological indicators must be based on reliable scientific knowledge and, possibly, on the availability of standardized procedures. In this article, we present a standardized protocol that used the marine crustacean Artemia to evaluate the toxicity of chemicals and/or of marine environmental matrices. Scientists propose that the brine shrimp (Artemia) is a suitable candidate for the development of a standard bioassay for worldwide utilization. A number of papers have been published on the toxic effects of various chemicals and toxicants on brine shrimp (Artemia). The major advantage of this crustacean for toxicity studies is the overall availability of the dry cysts; these can be immediately used in testing and difficult cultivation is not demanded1,2. Cyst-based toxicity assays are cheap, continuously available, simple and reliable and are thus an important answer to routine needs of toxicity screening, for industrial monitoring requirements or for regulatory purposes3. The proposed method involves the mortality as an endpoint. The numbers of survivors were counted and percentage of deaths were calculated. Larvae were considered dead if they did not exhibit any internal or external movement during several seconds of observation4. This procedure was standardized testing a reference substance (Sodium Dodecyl Sulfate); some results are reported in this work. This article accompanies a video that describes the performance of procedural toxicity testing, showing all the steps related to the protocol.

Long-term Lethal Toxicity Test with the Crustacean Artemia franciscana

This study concerns the development and standardization of a valuable methodological protocol to determine long-term (14 days) lethal toxicity exerted by chemical substances, industrial wastewater or sewage and liquid environmental samples on the saltwater crustacean, Artemia franciscana.

Our research activities target the use of biological methods for the evaluation of environmental quality, with particular reference to saltwater/brackish ...

Direct Imaging of ER Calcium with Targeted-Esterase Induced Dye Loading (TED)

Visualization of calcium dynamics is important to understand the role of calcium in cell physiology. To examine calcium dynamics, synthetic fluorescent Ca2+ indictors have become popular. Here we demonstrate TED (= targeted-esterase induced dye loading), a method to improve the release of Ca2+ indicator dyes in the ER lumen of different cell types. To date, TED was used in cell lines, glial cells, and neurons in vitro. TED bases on efficient, recombinant targeting of a high carboxylesterase activity to the ER lumen using vector-constructs that express Carboxylesterases (CES). The latest TED vectors contain a core element of CES2 fused to a red fluorescent protein, thus enabling simultaneous two-color imaging. The dynamics of free calcium in the ER are imaged in one color, while the corresponding ER structure appears in red. At the beginning of the procedure, cells are transduced with a lentivirus. Subsequently, the infected cells are seeded on coverslips to finally enable live cell imaging. Then, living cells are incubated with the acetoxymethyl ester (AM-ester) form of low-affinity Ca2+ indicators, for instance Fluo5N-AM, Mag-Fluo4-AM, or Mag-Fura2-AM. The esterase activity in the ER cleaves off hydrophobic side chains from the AM form of the Ca2+ indicator and a hydrophilic fluorescent dye/Ca2+ complex is formed and trapped in the ER lumen. After dye loading, the cells are analyzed at an inverted confocal laser scanning microscope. Cells are continuously perfused with Ringer-like solutions and the ER calcium dynamics are directly visualized by time-lapse imaging. Calcium release from the ER is identified by a decrease in fluorescence intensity in regions of interest, whereas the refilling of the ER calcium store produces an increase in fluorescence intensity. Finally, the change in fluorescent intensity over time is determined by calculation of &Delta;F/F0.

Direct Imaging of ER Calcium with Targeted-Esterase Induced Dye Loading TED

Targeted-esterase induced dye loading (TED) supports the analysis of intracellular calcium store dynamics by fluorescence imaging. The method bases on targeting of a recombinant Carboxylesterase to the endoplasmic reticulum (ER), where it improves the local unmasking of synthetic low-affinity Ca2+ indicator dyes in the ER lumen.

Visualization of calcium dynamics is important to understand the role of calcium in cell physiology. To examine calcium dynamics, synthetic fluorescent ...

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 ...