We thank Dr. Noah Weisleder for reading and editing of this manuscript. This work was supported by Research Grants UMDNJ Foundation 62-09 to ZP, American Heart Association SDG2630086 to XZ, 0535555N to MB, and National Institutes of Health RC2AR058962-01 to MB.

1. Intracellular Ca2+ measurement for individual cells
<ol>
	<li>KYSE-150, a human esophageal squamous cell carcinoma (ECSS) cell line is cultured in 5% CO2 atmosphere at 37&deg;C in mixed RPMI 1640/Ham's F-12 medium (1:1) containing 5% fetal bovine serum.</li>
	<li>KYSE-150 cells are transfected with the plasmids either containing shRNA specifically against Orai1 or a scrambled sequence. The plasmids also included a gene encoding red fluorescent protein (RFP) as a reporter, which is driven by a separate promoter.</li>
	<li>The cells are grown in glass-bottom dishes (No. 1.5 cover glass, MatTek, MA) for 48 hours.</li>
	<li>Remove the medium and rinse the cells with a balanced salt solution (BSS) (140 mM NaCl, 2.8 mM KCl, 2 mM CaCl2, 2 mM MgCl2, 10 mM HEPES, pH 7.2).</li>
	<li>Add 1 ml BSS solution containing 2 &mu;M Fura-2 acetoxymethyl ester (Molecular Probes) into the dish and wrap the whole dish with foil to protect from light.</li>
	<li>Incubate the cells for 40 min at 37&deg;C and then leave them for an additional 15 min period at room temperature to allow ester hydrolysis to be completed. Wash the cells with BSS twice.</li>
	<li>Mount the dish on the stage of Nikon TE200 inverted microscope, which is connected to PTI spectrofluorometer, with excitation wavelengths at 350 and 390 nm and emission at 510 nm. The exact excitation wavelengths to be used can slightly vary depending on the optics of each individual system. The two wavelengths with best ratio dynamic are determined by excitation spectrum scanning of Fura-2 salt in solutions containing Ca2+ ranging from 0 to 39.8 &mu;M (or higher concentration at which Fura-2 fluorescence is saturated).</li>
	<li>Set up a gravity perfusion system with 4 different solutions in BSS: Ca2+ (2 mM), EGTA (0.5 mM), EGTA-TG (thapsigargin, 5 &mu;M), Ca2+-2-APB (a SOCE inhibitor, 75 &mu;M). Computer-controlled automated perfusion systems can also be used.</li>
	<li>Select the cells expressing RFP in visualization mode.</li>
	<li>Position the opening tip of the perfusion system at 10x magnification until the tip is right above the region of interest at a 45 degree angle.</li>
	<li>Simultaneously record the fluorescence signals F350nm and F390nm. Ratio (F350nm/F390nm) is displayed in the third window. Change the perfusion solutions in the following order: Ca2+; EGTA; Ca2+; EGTA-TG; Ca2+; Ca2+-2-APB.</li>
</ol>
The above protocol can be modified for measurement of intracellular Ca2+ in cell suspension system.
<ol start="12">
	<li>Culture KYSE-150 cells in a T25 flask with the same culture condition as above.</li>
	<li>When the cells reach 90% confluence, remove the medium and rinse the cells with a BSS solution.</li>
	<li>Add 2.5 ml BSS solution containing 2 &mu;M Fura-2 AM and incubate the cells for 40 min at 37&deg;C followed by deesterification.</li>
	<li>Remove the BSS, add 2.5 ml trypsin into the flask and incubate at 37&deg;C until the cells detach. Then add 2.5 ml culture medium to stop the trypsinization and harvest the cells by centrifugation at 800 rpm for 5 min.</li>
	<li>Wash the cells with culture medium one more time by centrifugation and re-suspend the cells' pellet into BSS solution (without addition of 2 mM CaCl2, containing &mu;M range Ca2+) and count the cells number using a hematometer.</li>
	<li>Add ~106 cells into a quartz cuvette (10 mm, Starna Cells) and fill the volume up to 2 ml with BSS.</li>
	<li>Place the cuvette (stirring with a magnetic bar) into the spectrofluorometer.</li>
	<li>Simultaneously record the fluorescence signals F340nm and F380nm with emission wavelength at 510 nm. The running protocol is: BSS, addition of 0.5 &mu;l EGTA (0.25 M stock), addition of 1 &mu;l thapsigargin (TG, 10 mM stock), addition of 4 &mu;l Ca2+(1 M stock).</li>
</ol>
2. Mn quenching assay in cultured cells
<ol>
	<li>Perform excitation wavelength scanning of Fura-2 in solutions containing various Ca2+ concentration ranging from 0 to 39.8 &mu;M to determine the Ca concentration independent wavelength as the isosbestic point. Usually, the isosbestic point is at or around 360 nm.</li>
	<li>KYSE-150 cells are cultured in glass-bottom dishes and loaded with Fura-2 AM.</li>
	<li>Set up the perfusion system with 5 different BSS solutions: Ca2+ (2 mM), EGTA/TG (0.5 mM and 5 &mu;M, respectively), Mn2+ (0.5 mM, without addition of EGTA or Ca2+, containing free Ca2+ in &mu;M range), Mn2+/2-APB (75 &mu;M), EGTA/Triton X-100 (0.1%).</li>
	<li>Mount the dish on the stage of the microscope and select the "Excitation Ratio" mode with excitation wavelengths at 360 nm and 390 nm and emission at 510 nm.</li>
	<li>Simultaneously record the fluorescence signals F360nm and F390nm with Ratio (F360nm/F390nm) displayed in the third window. The running protocol is: Ca2+ for 1 min to stabilize the fluorescence, Mn2+ for 30 sec, EGTA/TG for 10 min, Mn2+ for 2 min, EGTA-Triton X-100 (0.1%) for 1 min to obtain the background reading. Mn2+/2-APB solution can be used instead of Mn2+ to examine the abolished fluorescence quenching, which indicates that the Mn2+ entry is through SOCE pathway.</li>
</ol>
3. Mn quenching assay in muscle cells
<ol>
	<li>C2C12, a mouse myogenic cell line, is cultured on glass-bottom dishes in 5% CO2 atmosphere at 37&deg;C in DMEM medium containing 10% fetal bovine serum, 10% horse serum.</li>
	<li>After the cells reach confluence, the culture medium is changed to differentiation medium (remove fetal bovine serum and reduce horse serum to 2.5%). The C2C12 myoblasts will differentiate into myotubes after 4 days.</li>
	<li>Load C2C12 myotubes with a final concentration of 5 &mu;M Fura-2 AM as described in 1.4-1.6.</li>
	<li>Add a final concentration of 20 &mu;M N-benzyl-p-toluene sulphonamide (BTS) to inhibit muscle contractions and the motion artifacts caused by them and allow 15 min before initiating the experiment.</li>
	<li>Mount the dish on the stage of the microscope connected to the PTI device and load the following BSS solutions into the perfusion system: 2 Ca2+ (2 mM), 0 Ca2+, Mn2+ (0.5 mM); Mn2+/TG (0.5 mM, 20 &mu;M, respectively).</li>
	<li>Set up the perfusion system as described above.</li>
	<li>Simultaneously record the fluorescence signals F360nm and F390nm with the Ratio (F360nm/F390nm) displayed in the third window. The running protocol is: 2 Ca2+ for 1 min, 0 Ca2+ for 1 min to flush away Ca2+, Mn2+ for 0.5 min, Mn2+/TG for 10 min, and Mn2+/EGTA-Triton X-100 (0.1%).</li>
</ol>
4. Spatially and temporally resolved SOCE in skinned muscle fibers
<ol>
	<li>Prepare the following solutions. 
	Isotonic Tyrode buffer: 140 mM NaCl, 5 mM KCl, 10 mM HEPES, 2 mM MgCl2, 2.5 mM CaCl2, pH 7.2, 290 mosm 
	Culture medium: DMEM with 2% horse serum and 1% penicillin and streptomycin 
	Wash solution #1: 109.6 mM K-glutamate, 2 mM EGTA-KOH, 6.7 mM MgCl2, 2 mM ATP, 6 mM creatine phosphate (CP), 20 mM N,N-bis(2-hydroxyethyl)-2-aminoethanesulfonic acid (BES)-KOH, pH 7.0 
	Wash solution #2: 140 mM K-glutamate, 5 mM EGTA-KOH, 6.5 mM MgCl2, 2 mM ATP, 6 mM CP, 20 mM BES-KOH, pH 7.0 
	Skinning solution: 140 mM K-glutamate, 6.5 mM MgCl2, 2 mM ATP, 6 mM CP, 20 mM BES-KOH, pH 7.0 
	TT-loading solution: 90.6 mM K-glutamate, 18 mM Na-glutamate, 0.55 mM CaCl2, 2 mM EGTA-KOH, 6.7 mM MgCl2, 5.4 mM ATP, 15 mM CP, 0.0025 mg/ml creatine kinase (CK), 20 mM BES-KOH, 5 &mu;M carbonyl cyanide p-trifluoromethoxyphenylhydrazone (FCCP), pH 7.0, pCa 7.0 
	SR loading solution: 107.8 mM K-glutamate, 0.98 mM CaCl2, 2 mM EGTA-KOH, 6.6 mM MgCl2, 5.4 mM ATP, 15 mM CP, 0.0025 mg/ml CK, 20 mM BES-KOH, 5 &mu;M FCCP, pH 7.0, pCa 6.6 
	SR depleting solution: 100 mM K-glutamate, 40 mM Na-glutamate, 10 mM EGTA-KOH, 10 mM 1,2-bis(o-aminophenoxy)ethane-N,N,N',N'-tetraacetic acid (BAPTA), 0.35 mM MgCl2, 0.5 mM ATP, 1 mM CP, 20 mM BES-KOH, 5 &mu;M FCCP, pH 7.0. 25 mM caffeine/20 &mu;M thapsigargin (TG) are added before experiments.</li>
	<li>To dissect the Extensor digitorum longus (EDL) muscle, first lay down the mice and arrange the leg at lateral position, then remove the skin from the ankle area up to the knee, cut the superficial muscle layers of tissue to expose the EDL, and sever both the upper and lower tendons to free the intact EDL muscle; it is important to keep tendons as long as possible. The EDL is transferred to a Tyrode solution containing 0 Ca2+ and 0.1 mM EGTA to prevent any contractions.</li>
	<li>Under a stereomicroscope, use two tweezers to hold lower insertion tendons and split the EDL muscle into two bundles. Repeat the process to obtain 4 and then 8 bundles. It is critical that tendons remain for each of the 8 bundles at the end of the process. If they are lost from some bundles, discard them.</li>
	<li>Under a stereomicroscope, each EDL bundle strip is griped at both tendons and carefully stretched until 3~4 separated single muscle fibers are left intact with tendon on both sides.</li>
	<li>Put 1 drop of Ca2+ free tyrode buffer in the middle of the glass-bottom dish, lay down the EDL strips straight on the dish, quickly remove as much as possible of the solution, then tape down both tendons using water resistant scotch tape, and make sure the tape is tight. Wash the fiber first with the 0 Ca2+ solution and then with a 2.5 mM Ca2+ solution 2 times. If the fiber hyper-contracts and damage is noticed, discard the fiber.</li>
	<li>If needed, the fiber can be cultured for up to 96 hours, allowing for genetic manipulations. Wash the fiber 3 times with complete culture medium and add 2 ml medium into the dish, place in 5% CO2 and 37&deg;C incubator. Before each experiment, examine muscle fibers and discard those without clear striation or with signs of contamination, or with signs of damage such as contracture of the sarcolemmal membrane. Good fibers respond to KCl, electrical stimulation, and caffeine stimulation.</li>
	<li>Single muscle fibers are washed 3 times with the wash solution # 1 and bathed in intracellular-like skinning solution with 500 &mu;M Rhod-5N and 0.5 mM Ca2+ for 5 min.</li>
	<li>Using a Tungsten needle attached to a pin holder, mechanically skin the fiber under a stereomicroscope, allowing transverse tubules (TT) to automatically reseal and to trap the Rhod-5N conjugated with Ca2+ in the TT, wash the skinned fibers twice with washing solution #2. The mechanical skinning process is optimal when less than 25% of the cell width is removed during the skinning process.</li>
	<li>To load the SR, skinned fibers are incubated with the skinning solution containing 20 &mu;M Fluo-5N AM for 1 h at room temperature, followed by extensive washes using washing solution #2, and incubate for an additional 30 min to allow complete de-esterification of the dye.</li>
	<li>After 30 min, skinned fibers are visualized under 60X objective of the confocal microscope. Fluorescence images are acquired at wavelengths of the corresponding dyes (excitation at 543 nm and emission longer than 570 nm for Rhod-5N; excitation at 488 nm and emission at 530-560 nm for Fluo-5N). Regions of interest (ROIs) are selected for each dye loaded areas.</li>
	<li>The experimental protocol is as following: TT-loading solution for 120 s, SR-loading solution for 120 s, SR-depletion solution for 400-750 s to deplete the SR Ca2+ store. During this process, changes in Rhod-5N intensity (TT Ca2+) and Fluo-5N intensity (SR Ca2+) are recorded, creating a time course determination of relative fluorescence changes in the TT and SR. Mean values of fluorescence intensity from multiple fibers are further analyzed. The maximal loading intensity at the end of TT/SR-loading is normalized to 100%.</li>
</ol>
5. Representative Results
We examined SOCE activity in KYSE-150 cells using intracellular Ca2+ measurement (Fig 2.). Using RFP as reporter, we could select the individual cells transfected with plasmid containing specific shRNA against orai1, a gene encoding the SOCE channel. Compared with cells transfected with plasmids containing scramble shRNA (black trace), the knocking down of Orai1 protein results in decreased SOCE activity (red trace).
SOCE in KYSE-150 cells was also confirmed with the Mn2+ quenching assay (Fig 3.). The excitation wavelength of 360 nm reports the isosbestic point of Fura-2, where the fluorescence is independent of the Ca2+ concentration. After TG completely depleted ER Ca2+ stores, the perfusion of Mn2+ resulted in a significant fluorescence decrease. The overall SOCE activity can be measured by the decrease rate of Fura-2 fluorescence intensity with a steeper slope indicating a more active SOCE, while a shallower slope meaning a less active SOCE. The slope of fluorescence decrease in 2-APB treated cells appeared to be much shallower, which indicated that SOCE activity was blocked by this compound. The Mn2+ quenching rates were determined from the record of the initial 10 seconds, where the quenching was still in linear range without saturation.
A similar Mn2+ quenching assay was performed in the muscle (Fig 4.). In this case Mn2+ was applied together with TG. While TG was depleting the SR Ca2+ stores, the fluorescence quenching slope gradually changed until it reached its maximal rate. The sigmoidal curve indicates the graded activation of SOCE under these experimental conditions.
Healthy intact single muscle fibers in culture showed clear and uniform striation, no signs of contaminations, and no signs of contraction-induced damage. These fibers were able to contract in response to electrical stimulation or depolarizing solutions. Under confocal imaging, skinned fiber trapping of Rhod-5N dye in the TT compartment showed the characteristic mammalian doublet pattern (visualized in the red channel). After loading of the SR with Fluo-5N AM, the typical punctuated SR pattern was visible (in green channel). Upon perfusion of the skinned fiber with TT/SR loading solution, fluorescence intensity of both Rhod-5N and Fluo-5N increased; upon perfusion with SR depletion solution, fluorescence levels in the SR compartment and the TT compartment started to decrease, indicating tight coupling between SR Ca2+ store depletion and SOCE activation, respectively (Fig 5.).
<img src="/files/ftp_upload/3415/3415fig1.jpg" alt="Figure 1" /> 
Figure 1. Activation of store-operated Ca2+ entry (SOCE). Orai1, a channel pore forming unit, is located on the plasma membrane (PM) and STIM1, a Ca2+ sensor, is located on the endoplasmic or sarcoplasmic reticulum (ER/SR) membranes. When ER/SR Ca2+ stores are reduced either due to blocking of ER/SR Ca2+ pump (SERCA) or Ca2+ release through IP3 receptor or ryanodine receptor, STIM1 is activated. Activated STIM1 molecules form patches and induce the aggregation of Orai1, which further leads the activation of SOCE.
<img src="/files/ftp_upload/3415/3415fig2.jpg" alt="Figure 2" /> 
Figure 2. Measurement of intracellular Ca2+ in KYSE-150 cells. Exchange of extracellular solution from 0 Ca2+ (0.5 mM EGTA) to 2 mM Ca2+ did not induce any change in intracellular Ca2+. 5 &mu;M TG in 0 Ca2+ bath solution induced passive ER Ca2+ release. After ER Ca2+ stores were depleted (&gt;10 min), addition of extracellular Ca2+ (2 mM) activated a sustained intracellular Ca2+ elevation through SOCE, which can be blocked by SOCE inhibitors, e.g. skf-96365 and 2-APB (data not show). Cells transfected with plasmids containing shRNA specifically against orai1 (red) demonstrated significantly reduced SOCE than cells transfected with scramble sequence (black). 
<img src="/files/ftp_upload/3415/3415fig3.jpg" alt="Figure 3" /> 
Figure 3. Mn2+ quenching assay of SOCE in KYSE-150 cells. The quenching of Fura-2 fluorescence by Mn2+ (0.5 mM) was measured at the Ca2+-independent excitation wavelength of Fura-2 (360 nm). Cells were treated with 5 &mu;M TG for 10 min to completely deplete ER Ca2+ stores. The decay slope of Fura-2 fluorescence upon Mn2+ addition (dash lines, within the first 10 sec) represented the activation of SOCE, which was expressed as percent decrease in fluorescence per unit time (the initial value as 100%). The maximally quenched fluorescence signal was established at the end of the experiment by lysing the cells with 0.1% Triton X-100 (as 0%). Cells treated with 2-APB (75 &mu;M) demonstrated a much shallower slope, suggesting SOCE is blocked by this compound in KYSE-150 cells. 
<img src="/files/ftp_upload/3415/3415fig4.jpg" alt="Figure 4" /> 
Figure 4. Graded activation of SOCE in muscle cells revealed by Mn2+ quenching assay. Activation of SOCE could be registered while TG was depleting SR Ca2+ stores. Simultaneous application of Mn2+ (0.5mM) and TG (20 &mu;M) induced a graded- and sigmoidal decrease of intracellular Fura-2 fluorescence (cyan dash line to show the fastest quenching point), which was distinct from initial Mn2+ quenching rate (blue dash line). Mn2+ quenching slope remained almost the same as basal level in cells treated 2-APB (75 &mu;M), suggesting that SOCE was inhibited.
<img src="/files/ftp_upload/3415/3415fig5.jpg" alt="Figure 5" /> 
Figure 5. Spatially and temporally resolved SOCE in skinned muscle fiber. (A) Rhod-5N salt was loaded into TT and Fluo-5N AM was loaded into SR. (B) After applying Ca2+ depletion solution, the fluorescence in both TT and SR compartments decreased. The loss of Fluo-5N fluorescence indicated rapidly released SR Ca2+ content and the loss of Rhod-5N fluorescence suggested activation of SOCE.

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No conflicts of interest declared.

Although the excitation wavelengths for Ca2+-binding and Ca2+ free Fura-2 are 340 nm and 380 nm, respectively, the best ratio dynamic range of Fura-2 for Ca2+ concentration measurement may occur at other wavelengths in a particular microscope system. Such shifts in wavelength are usually due to changes in the optical path with the addition of various optical components. In this study, the excitation wavelengths for Fura-2 were determined as 350 nm and 390 nm by performing spectral analysis of Fura-2 fluorescence.
The intracellular Ca2+ level in the cytosol results from a balance of several sources, including intracellular Ca2+ release, extracellular Ca2+ entry, as well as Ca2+ exclusion mechanism at the ER and plasma membrane. To isolate the unidirectional SOC-mediated Ca2+ influx from intracellular Ca2+ release and Ca2+ extrusion, the Mn2+ quenching assay can be used. Mn2+ is known to be able to permeate into cells via SOCE but is impervious to surface membrane extrusion or ER uptake by Ca2+ pumps. Therefore, fluorescence quenching represents a measurement of unidirectional Mn2+ flux into cells that estimates the degree of activation of SOCE. Mn2+ quenching assay is performed at the isosbestic point, at such wavelength Fura-2 fluorescence intensity is independent of Ca2+ concentration. The isosbestic point for each system should be determined by spectrum scanning. Alternatively, Mn2+ quenching can be recorded by adjusted fluorescence at any two wavelengths in such a way that the final value is independent of Ca2+ concentration13.
To gain the spatial and temporal information of activity of SOCE in skeletal muscles, we employed a dual-dye method, which allows simultaneous measurement of SOCE activity and SR Ca2+ content in skinned adult mammalian EDL muscle fibers. The correlation between changes in SR Ca2+ content and SOCE activity can be plotted to indicate the threshold and sensitivity of SOCE activation to depletion of the SR Ca2+ storage. The TT-loading solution is optimized to additionally load the T-tubules with Ca2+ and for priming of the T-tubules and the SR-loading solution is designed to promote maximal SR Ca2+ loading and to additionally load the T-tubules with ~500 &mu;M Ca2+. FCCP is included in the TT/SR-loading solution and the SR-depleting solution to eliminate the effects from the mitochondria.

Table of specific reagents and equipment:
<table border="1">
<tbody>
 <tr>
 <td>Name 			of the reagent</td>
 <td>Company</td>
 <td>Catalogue 			number</td>
 <td>Comments </td>
 </tr>
 <tr>
 <td>RPMI 			1640</td>
 <td>Mediatech</td>
 <td>10-040-CV</td>
 <td>&nbsp;
 </td>
 </tr>
 <tr>
 <td>Ham's 			F-12</td>
 <td>Mediatech</td>
 <td>10-080-CV</td>
 <td>&nbsp;
 </td>
 </tr>
 <tr>
 <td>DMEM</td>
 <td>Mediatech</td>
 <td>10-013-CV</td>
 <td>&nbsp;
 </td>
 </tr>
 <tr>
 <td>Glass 			Bottom Dish</td>
 <td>MatTek</td>
 <td>P35G-1.5-14-C</td>
 <td>&nbsp;
 </td>
 </tr>
 <tr>
 <td>Fura-2 			AM</td>
 <td>Invitrogen 			(Molecular Probes)</td>
 <td>F1221</td>
 <td>-20&deg;C, 			seal tight, protected from light, dissolved into DMSO</td>
 </tr>
 <tr>
 <td>Fluo-5N 			AM</td>
 <td>Invitrogen 			(Molecular Probes)</td>
 <td>F14204</td>
 <td>-20&deg;C, 			seal tight, protected from light</td>
 </tr>
 <tr>
 <td>Rhod-5N</td>
 <td>Invitrogen 			(Molecular Probes)</td>
 <td>R14207</td>
 <td>-20&deg;C, 			seal tight, protected from light</td>
 </tr>
 <tr>
 <td>Quartz 			Cuvette</td>
 <td>Starna 			Cells</td>
 <td>3-Q-10</td>
 <td>&nbsp;
 </td>
 </tr>
 <tr>
 <td>thapsigargin</td>
 <td>Tocris</td>
 <td>1138</td>
 <td>&nbsp;
 </td>
 </tr>
 <tr>
 <td>2-Aminoethoxydiphenylborane 			(2-APB)</td>
 <td>Tocris</td>
 <td>1224</td>
 <td>&nbsp;
 </td>
 </tr>
 <tr>
 <td>N-benzyl-p-toluene 			sulphonamide (BTS)</td>
 <td>Sigma-Aldrich</td>
 <td>435600</td>
 <td>&nbsp;
 </td>
 </tr>
 <tr>
 <td>Collagenase 			Type I</td>
 <td>Sigma</td>
 <td>C0130</td>
 <td>&nbsp;
 </td>
 </tr>
 </tbody>
</table>
Equipment:
<ol>
<li> A spectrofluorometer (Photon Technology International, NJ): xenon arc lamp, computer-controlled high-speed random access monochromator, cuvette-based emission monochromator, Nikon TE-200 inverted fluorescence microscope, S Fluor 40x/1.30 oil objective, PMT detectors.</li>
<li> Laser Scanning Confocal Microscope (BioRad Radiance2100): argon laser 453/477/488/514, HeNe laser 543nm, Nikon TE2000 inverted microscope, 60X objective (N.A. 1.4, oil).</li>
<li> A stereomicroscope with a magnification power &gt; 100 (Zeiss Stemi SV11 Apo).</li>
</ol>

fluorescence-based measurement of store-operated calcium entry in live cells: from cultured cancer cell to skeletal muscle fiber

Store operated Ca2+ entry (SOCE), earlier termed capacitative Ca2+ entry, is a tightly regulated mechanism for influx of extracellular Ca2+ into cells to replenish depleted endoplasmic reticulum (ER) or sarcoplasmic reticulum (SR) Ca2+ stores1,2. Since Ca2+ is a ubiquitous second messenger, it is not surprising to see that SOCE plays important roles in a variety of cellular processes, including proliferation, apoptosis, gene transcription and motility. Due to its wide occurrence in nearly all cell types, including epithelial cells and skeletal muscles, this pathway has received great interest3,4. However, the heterogeneity of SOCE characteristics in different cell types and the physiological function are still not clear5-7.
The functional channel properties of SOCE can be revealed by patch-clamp studies, whereas a large body of knowledge about this pathway has been gained by fluorescence-based intracellular Ca2+ measurements because of its convenience and feasibility for high-throughput screening. The objective of this report is to summarize a few fluorescence-based methods to measure the activation of SOCE in monolayer cells, suspended cells and muscle fibers5,8-10. The most commonly used of these fluorescence methods is to directly monitor the dynamics of intracellular Ca2+ using the ratio of F340nm and F380nm (510 nm for emission wavelength) of the ratiometric Ca2+ indicator Fura-2. To isolate the activity of unidirectional SOCE from intracellular Ca2+ release and Ca2+ extrusion, a Mn2+ quenching assay is frequently used. Mn2+ is known to be able to permeate into cells via SOCE while it is impervious to the surface membrane extrusion processes or to ER uptake by Ca2+ pumps due to its very high affinity with Fura-2. As a result, the quenching of Fura-2 fluorescence induced by the entry of extracellular Mn2+ into the cells represents a measurement of activity of SOCE9. Ratiometric measurement and the Mn+2 quenching assays can be performed on a cuvette-based spectrofluorometer in a cell population mode or in a microscope-based system to visualize single cells. The advantage of single cell measurements is that individual cells subjected to gene manipulations can be selected using GFP or RFP reporters, allowing studies in genetically modified or mutated cells. The spatiotemporal characteristics of SOCE in structurally specialized skeletal muscle can be achieved in skinned muscle fibers by simultaneously monitoring the fluorescence of two low affinity Ca2+ indicators targeted to specific compartments of the muscle fiber, such as Fluo-5N in the SR and Rhod-5N in the transverse tubules9,11,12.

Watch this Scientific Journal Video about Fluorescence-based Measurement of Store-operated Calcium Entry in Live Cells: from Cultured Cancer Cell to Skeletal Muscle Fiber at JoVE.com

Fluorescence-based Measurement of Store-operated Calcium Entry in Live Cells: from Cultured Cancer Cell to Skeletal Muscle Fiber

The extent of store-operated Ca2+ entry (SOCE) can be monitored using fluorescent Ca2+ indicators. Mn2+ quenching of such indicators assays SOCE in cultured cells and skeletal muscle fibers. A technique allowing spatial and temporal resolution of SOCE by confocal imaging of mechanically skinned muscle fibers is also described.

Store operated Ca2+ entry (SOCE), earlier termed capacitative Ca2+ entry, is a tightly regulated mechanism for influx of extracellular Ca2+ into cells to ...

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21.0K Views.  Robert Wood Johnson Medical School. The extent of store-operated Ca2+ entry (SOCE) can be monitored using fluorescent Ca2+ indicators. Mn2+ quenching of such indicators assays SOCE in cultured cells and skeletal muscle fibers. A technique allowing spatial and temporal resolution of SOCE by confocal imaging of mechanically skinned muscle fibers is also described.

Video: Fluorescence-based Measurement of Store-operated Calcium Entry in Live Cells: from Cultured Cancer Cell to Skeletal Muscle Fiber

Mitochondrial Isolation from Skeletal Muscle

Mitochondria are organelles controlling the life and death of the cell. They participate in key metabolic reactions, synthesize most of the ATP, and regulate a number of signaling cascades2,3. Past and current researchers have isolated mitochondria from rat and mice tissues such as liver, brain and heart4,5. In recent years, many researchers have focused on studying mitochondrial function from skeletal muscles.
Here, we describe a method that we have used successfully for the isolation of mitochondria from skeletal muscles 6. Our procedure requires that all buffers and reagents are made fresh and need about 250-500 mg of skeletal muscle. We studied mitochondria isolated from rat and mouse gastrocnemius and diaphragm, and rat extraocular muscles. Mitochondrial protein concentration is measured with the Bradford assay. It is important that mitochondrial samples be kept ice-cold during preparation and that functional studies be performed within a relatively short time (~1 hr). Mitochondrial respiration is measured using polarography with a Clark-type electrode (Oxygraph system) at 37&deg;C7. Calibration of the oxygen electrode is a key step in this protocol and it must be performed daily. Isolated mitochondria (150 &mu;g) are added to 0.5 ml of experimental buffer (EB). State 2 respiration starts with addition of glutamate (5mM) and malate (2.5 mM). Then, adenosine diphosphate (ADP) (150 &mu;M) is added to start state 3. Oligomycin (1 &mu;M), an ATPase synthase blocker, is used to estimate state 4. Lastly, carbonyl cyanide p-[trifluoromethoxy]-phenyl-hydrazone (FCCP, 0.2 &mu;M) is added to measurestate 5, or uncoupled respiration 6. The respiratory control ratio (RCR), the ratio of state 3 to state 4, is calculated after each experiment. An RCR &ge;4 is considered as evidence of a viable mitochondria preparation.
In summary, we present a method for the isolation of viable mitochondria from skeletal muscles that can be used in biochemical (e.g., enzyme activity, immunodetection, proteomics) and functional studies (mitochondrial respiration).

This protocol describes a procedure to study the respiration of mitochondria isolated from skeletal muscles. This method was adapted from Scorrano et al. (2007). The mitochondrial isolation procedure requires about 2 hours. The mitochondrial respiration can be completed in about 1 hour.

Mitochondria are organelles controlling the life and death of the cell. They participate in key metabolic reactions, synthesize most of the ATP, and ...

Purification of Progenitors from Skeletal Muscle

Skeletal muscle contains multiple progenitor populations of distinct embryonic origins and developmental potential. Myogenic progenitors, usually residing in a "satellite cell position" between the myofiber plasma membrane and the laminin-rich basement membrane that ensheaths it, are self-renewing cells that are solely committed to the myogenic lineage1,2. We have recently described a second class of vessel associated progenitors that can generate myofibroblasts and white adipocytes, which responds to damage by efficiently entering proliferation and provides trophic support to myogenic cells during tissue regeneration3,4. One of the most trusted assays to determine the developmental and regenerative potential of a given cell population relies on their isolation and transplantation5-7. To this end we have optimized protocols for their purification by flow cytometry from enzymatically dissociated muscle, which we will outline in this article. The populations obtained using this method will contain either myogenic or fibro/adipogenic colony forming cells: no other cell types are capable of expanding in vitro or surviving in vivo delivery. However, when these populations are used immediately after the sort for molecular analysis (e.g qRT-PCR) one must keep in mind that the freshly sorted subsets may contain other contaminant cells that lack the ability of forming colonies or engrafting recipients.

Method for the enzymatic dissociation, surface labeling and purification by flow cytometry of fibro/adipogenic and myogenic progenitors from murine skeletal muscle.

Skeletal muscle contains multiple progenitor populations of distinct embryonic origins and developmental potential. Myogenic progenitors, usually residing ...

Isolation and Culture of Individual Myofibers and their Satellite Cells from Adult Skeletal Muscle

Muscle regeneration in the adult is performed by resident stem cells called satellite cells. Satellite cells are defined by their position between the basal lamina and the sarcolemma of each myofiber. Current knowledge of their behavior heavily relies on the use of the single myofiber isolation protocol. In 1985, Bischoff described a protocol to isolate single live fibers from the Flexor Digitorum Brevis (FDB) of adult rats with the goal to create an in vitro system in which the physical association between the myofiber and its stem cells is preserved 1. In 1995, Rosenblattmodified the Bischoff protocol such that myofibers are singly picked and handled separately after collagenase digestion instead of being isolated by gravity sedimentation 2, 3. The Rosenblatt or Bischoff protocol has since been adapted to different muscles, age or conditions 3-6. The single myofiber isolation technique is an indispensable tool due its unique advantages. First, in the single myofiber protocol, satellite cells are maintained beneath the basal lamina. This is a unique feature of the protocol as other techniques such as Fluorescence Activated Cell Sorting require chemical and mechanical tissue dissociation 7. Although the myofiber culture system cannot substitute for in vivo studies, it does offer an excellent platform to address relevant biological properties of muscle stem cells. Single myofibers can be cultured in standard plating conditions or in floating conditions. Satellite cells on floating myofibers are subjected to virtually no other influence than the myofiber environment. Substrate stiffness and coating have been shown to influence satellite cells' ability to regenerate muscles 8, 9 so being able to control each of these factors independently allows discrimination between niche-dependent and -independent responses. Different concentrations of serum have also been shown to have an effect on the transition from quiescence to activation. To preserve the quiescence state of its associated satellite cells, fibers should be kept in low serum medium 1-3. This is particularly useful when studying genes involved in the quiescence state. In serum rich medium, satellite cells quickly activate, proliferate, migrate and differentiate, thus mimicking the in vivo regenerative process 1-3. The system can be used to perform a variety of assays such as the testing of chemical inhibitors; ectopic expression of genes by virus delivery; oligonucleotide based gene knock-down or live imaging. This video article describes the protocol currently used in our laboratory to isolate single myofibers from the Extensor Digitorum Longus (EDL) muscle of adult mice (6-8 weeks old).

Isolation and culture of myofibers is the gold standard in vitro system to study the transition of satellite cells through quiescence, activation and differentiation. Importantly, the single myofiber culture system preserves the myofiber/stem cell association, which is an essential component of the muscle stem cell niche.

Muscle  regeneration in the adult is performed by resident stem cells called satellite  cells. Satellite cells are defined by  their position between the ...

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 ...

Isolation of Blood-vessel-derived Multipotent Precursors from Human Skeletal Muscle

Since the discovery of mesenchymal stem/stromal cells (MSCs), the native identity and localization&#160;of MSCs have been obscured by their retrospective isolation in culture. Recently, using fluorescence-activated cell sorting (FACS), we and other researchers prospectively identified and purified three subpopulations of multipotent precursor cells associated with the vasculature of human skeletal muscle. These three cell populations: myogenic endothelial cells (MECs), pericytes (PCs), and adventitial cells (ACs), are localized respectively to the three structural layers of blood vessels: intima, media, and adventitia. All of these human blood-vessel-derived stem cell (hBVSC) populations not only express classic MSC markers but also possess mesodermal developmental potentials similar to typical MSCs. Previously, MECs, PCs, and ACs have been isolated through distinct protocols and subsequently characterized in separate studies. The current isolation protocol, through modifications to the isolation process and adjustments in the selective cell surface markers, allows us to simultaneously purify all three hBVSC subpopulations by FACS from a single human muscle biopsy. This new method will not only streamline the isolation of multiple BVSC subpopulations but also facilitate future clinical applications of hBVSCs for distinct therapeutic purposes.

Blood vessels within human skeletal muscle harbor several multi-lineage precursor populations that are ideal for regenerative applications. This isolation method allows simultaneous purification of three multipotent precursor cell populations respectively from three structural layers of blood vessels: myogenic endothelial cells from intima, pericytes from media, and adventitial cells from adventitia.

Since the discovery of mesenchymal stem/stromal cells (MSCs), the native identity and localization&#160;of MSCs have been obscured by their retrospective ...

Measurement of Calcium Fluctuations Within the Sarcoplasmic Reticulum of Cultured Smooth Muscle Cells Using FRET-based Confocal Imaging

Maintenance of steady-state calcium (Ca2+) levels in the sarcoplasmic reticulum (SR) of vascular smooth muscle cells (VSMCs) is vital to their overall health. A significant portion of intracellular Ca2+ content is found within the SR stores in VSMCs. As the only intracellular organelle with a close association to the surrounding extracellular space through plasma membrane-SR junctions, the SR can be considered to constitute the first line of response to any irregularity in Ca2+ transients, or stress experienced by the cell. Among its many functions, one of the most important is its role in the transmission of Ca2+-regulated signals throughout the cell to induce further cell-wide reactions downstream. The more common use of cytoplasmic Ca2+ indicators in this regard is overall insufficient for research into the highly dynamic changes to the intraluminal SR Ca2+ store that have yet to be fully characterized. Here, we provide a detailed protocol for the direct and clear measurement of luminal SR Ca2+. This tool is useful for investigation into the nuanced changes in SR Ca2+ that have significant subsequent effects on the normal function and health of the cell. Fluctuations in SR Ca2+ content specifically can provide us with a significant amount of information pertaining to cellular responses to disease or stress conditions experienced by the cell. In this method, a modified version of a SR-targeted Ca2+ indicator, D1SR, is used to detect Ca2+ fluctuations in response to the introduction of agents to cultured rat aortic smooth muscle cells (SMCs). Following incubation with the D1SR indicator, confocal fluorescence microscopy and fluorescence resonance energy transfer (FRET)-based imaging are used to directly observe changes to intraluminal SR Ca2+ levels under control conditions and with the addition of agonist agents that function to induce intracellular Ca2+ movement.

Currently, most available calcium indicators are used to quantify cytoplasmic calcium transients as indirect measures of calcium released from the sarcoplasmic reticulum in cultured smooth muscle cells. This protocol describes the use of a specific FRET-based indicator that allows direct measurement of calcium signals within the sarcoplasmic reticulum lumen.

Maintenance of steady-state calcium (Ca2+) levels in the sarcoplasmic reticulum (SR) of vascular smooth muscle cells (VSMCs) is vital to their overall ...

Simultaneous Measurement of Mitochondrial Calcium and Mitochondrial Membrane Potential in Live Cells by Fluorescent Microscopy

Apart from their essential role in generating ATP, mitochondria also act as local calcium (Ca2+) buffers to tightly regulate intracellular Ca2+ concentration. To do this, mitochondria utilize the electrochemical potential across their inner membrane (&#916;&#936;m) to sequester Ca2+. The influx of Ca2+ into the mitochondria stimulates three rate-limiting dehydrogenases of the citric acid cycle, increasing electron transfer through the oxidative phosphorylation (OXPHOS) complexes. This stimulation maintains &#916;&#936;m, which is temporarily dissipated as the positive calcium ions cross the mitochondrial inner membrane into the mitochondrial matrix.
We describe here a method for simultaneously measuring mitochondria Ca2+ uptake and &#916;&#936;m in live cells using confocal microscopy. By permeabilizing the cells, mitochondrial Ca2+ can be measured using the fluorescent Ca2+ indicator Fluo-4, AM, with measurement of &#916;&#936;m using the fluorescent dye tetramethylrhodamine, methyl ester, perchlorate (TMRM). The benefit of this system is that there is very little spectral overlap between the fluorescent dyes, allowing accurate measurement of mitochondrial Ca2+ and &#916;&#936;m simultaneously. Using the sequential addition of Ca2+ aliquots, mitochondrial Ca2+ uptake can be monitored, and the concentration at which Ca2+ induces mitochondrial membrane permeability transition and the loss of &#916;&#936;m determined.

Mitochondria can utilize the electrochemical potential across their inner membrane (&#916;&#936;m) to sequester calcium (Ca2+), allowing them to shape cytosolic Ca2+ signaling within the cell. We describe a method for simultaneously measuring mitochondria Ca2+ uptake and &#916;&#936;m in live cells using fluorescent dyes and confocal microscopy.

Apart from their essential role in generating ATP, mitochondria also act as local calcium (Ca2+) buffers to tightly regulate intracellular Ca2+ ...

Live-cell Imaging of Fungal Cells to Investigate Modes of Entry and Subcellular Localization of Antifungal Plant Defensins

Small cysteine-rich defensins are one of the largest groups of host defense peptides present in all plants. Many plant defensins exhibit potent in vitro antifungal activity against a broad-spectrum of fungal pathogens and therefore have the potential to be used as antifungal agents in transgenic crops. In order to harness the full potential of plant defensins for diseases control, it is crucial to elucidate their mechanisms of action (MOA). With the advent of advanced microscopy techniques, live-cell imaging has become a powerful tool for understanding the dynamics of the antifungal MOA of plant defensins. Here, a confocal microscopy based live-cell imaging method is described using two fluorescently labeled plant defensins (MtDef4 and MtDef5) in combination with vital fluorescent dyes. This technique enables real-time visualization and analysis of the dynamic events of MtDef4 and MtDef5 internalization into fungal cells. Importantly, this assay generates a wealth of information including internalization kinetics, mode of entry and subcellular localization of these peptides. Along with other cell biological tools, these methods have provided critical insights into the dynamics and complexity of the MOA of these peptides. These tools can also be used to compare the MOA of these peptides against different fungi.

Plant defensins play an important role in plant defense against pathogens. For effective use of these antifungal peptides as antifungal agents, understanding their modes of action (MOA) is critical. Here, a live-cell imaging method is described to study critical aspects of the MOA of these peptides.

Small cysteine-rich defensins are one of the largest groups of host defense peptides present in all plants. Many plant defensins exhibit potent in vitro ...

Isolation of High Quality Murine Atrial and Ventricular Myocytes for Simultaneous Measurements of Ca2+ Transients and L-Type Calcium Current

Mouse models play a crucial role in arrhythmia research and allow studying key mechanisms of arrhythmogenesis including altered ion channel function and calcium handling. For this purpose, atrial or ventricular cardiomyocytes of high quality are necessary to perform patch-clamp measurements or to explore calcium handling abnormalities. However, the limited yield of high-quality cardiomyocytes obtained by current isolation protocols does not allow both measurements in the same mouse. This article describes a method to isolate high-quality murine atrial and ventricular myocytes via retrograde enzyme-based Langendorff perfusion, for subsequent simultaneous measurements of calcium transients and L-type calcium current from one animal. Mouse hearts are obtained, and the aorta is rapidly cannulated to remove blood. Hearts are then initially perfused with a calcium-free solution (37 &#176;C) to dissociate the tissue at the level of intercalated discs and afterwards with an enzyme solution containing little calcium to disrupt extracellular matrix (37 &#176;C). The digested heart is subsequently dissected into atria and ventricles. Tissue samples are chopped into small pieces and dissolved by carefully pipetting up and down. The enzymatic digestion is stopped, and cells are stepwise reintroduced to physiologic calcium concentrations. After loading with a fluorescent Ca2+-indicator, isolated cardiomyocytes are prepared for simultaneous measurement of calcium currents and transients. Additionally, isolation pitfalls are discussed and patch-clamp protocols and representative traces of L-type calcium currents with simultaneous calcium transient measurements in atrial and ventricular murine myocytes isolated as described above are provided.

Isolation of High Quality Murine Atrial and Ventricular Myocytes for Simultaneous Measurements of Ca2+ Transients and L-Type Calcium Current

Mouse models allow studying key mechanisms of arrhythmogenesis. For this purpose, high quality cardiomyocytes are necessary to perform patch-clamp measurements. Here, a method to isolate murine atrial and ventricular myocytes via retrograde enzyme-based Langendorff perfusion, which allows simultaneous measurements of calcium-transients and L-type calcium current, is described.

Mouse models play a crucial role in arrhythmia research and allow studying key mechanisms of arrhythmogenesis including altered ion channel function and ...

Quantification of Mitochondrial Membrane Potential and Superoxide Levels

Fluorescence-Based Quantification of Mitochondrial Membrane Potential and Superoxide Levels Using Live Imaging in HeLa Cells

Mitochondria are dynamic organelles critical for metabolic homeostasis by controlling energy production via ATP synthesis. To support cellular metabolism, various mitochondrial quality control mechanisms cooperate to maintain a healthy mitochondrial network. One such pathway is mitophagy, where PTEN-induced kinase 1 (PINK1) and Parkin phospho-ubiquitination of damaged mitochondria facilitate autophagosome sequestration and subsequent removal from the cell via lysosome fusion. Mitophagy is important for cellular homeostasis, and mutations in Parkin are linked to Parkinson's disease (PD). Due to these findings, there has been a significant emphasis on investigating mitochondrial damage and turnover to understand the molecular mechanisms and dynamics of mitochondrial quality control. Here, live-cell imaging was used to visualize the mitochondrial network of HeLa cells, to quantify the mitochondrial membrane potential and superoxide levels following treatment with carbonyl cyanide m-chlorophenyl hydrazone (CCCP), a mitochondrial uncoupling agent. In addition, a PD-linked mutation of Parkin (ParkinT240R) that inhibits Parkin-dependent mitophagy was expressed to determine how mutant expression impacts the mitochondrial network compared to cells expressing wild-type Parkin. The protocol outlined here describes a simple workflow using fluorescence-based approaches to quantify mitochondrial membrane potential and superoxide levels effectively.

Author Spotlight: Fluorescence-Based Quantification of Mitochondrial Membrane Potential and Superoxide Levels Using Live Imaging in HeLa Cells  

This technique describes an effective workflow to visualize and quantitatively measure mitochondrial membrane potential and superoxide levels within HeLa cells using fluorescence-based live imaging.

Mitochondria are dynamic organelles critical for metabolic homeostasis by controlling energy production via ATP synthesis. To support cellular metabolism, ...