Research in the laboratories is funded by the Fritz-Bender Foundation, Munich, Germany (to T.D.) and a University of Witten/Herdecke internal research grant (to W.-K.L). We would like to thank Prof Dr. Dr. Frank Th&#233;venod (Institute for Physiology, Pathophysiology &#38; Toxicology, University of Witten/Herdecke) for helpful suggestions.

1. Preparation of Reagents and Solutions
<ol>
	<li>Prepare a modified Hank&#8217;s balanced salt solution (HBSS) (in mM: 136.9 NaCl, 5.37 KCl, 0.34 Na2HPO4, 0.44 KH2PO4, 4.17 NaHCO3, pH 7.2, no phenol red). Store at 4 &#176;C.</li>
	<li>Make 1.5 M CaCl2 and 2 M 4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid (HEPES) (pH 7.2) stock solutions using distilled water.</li>
	<li>Prepare a 250 mM stock solution of probenecid. For the free acid form (often referred to as water insoluble), dissolve probenecid powder in 1 N NaOH, fill to three quarters of the final volume with distilled water and slowly titrate to pH 7.4 with 1 N HCl. Store at 4 &#176;C.</li>
	<li>Dissolve fluorescent membrane permeable calcium binding dye in anhydrous dimethyl sulfoxide (DMSO) to a stock concentration of 1 mM. Store at -20 &#176;C in the dark.</li>
	<li>Prior to each experiment, freshly prepare probenecid-containing HBSS flux (PHF) buffer by combining modified HBSS with (in final concentrations) 1.5 mM CaCl2, 10 mM HEPES, pH 7.2, 5% fetal bovine serum, 2 mM probenecid, and 5.55 mM glucose. Warm to 37 &#176;C.</li>
</ol>
2. Cell Culture
<ol>
	<li>Plate 2.5 x 105 kidney proximal tubule cells (WKPT-0293 Cl.2) per well of a 6 well plate or 35 mm dish in standard culture medium and grow for 2 days reaching a confluency of approximately 70%.</li>
</ol>
3. Calcium Dye Loading
<ol>
	<li>For each well or dish, mix 1 ml culture medium containing 5% fetal bovine serum, 4 &#181;M cell permeable calcium binding dye and 2 mM probenecid.</li>
	<li>Replace the culture medium with 1 ml loading dye mixture in each well and incubate at 37 &#176;C for 30 min in a humidified incubator with 5% CO2. Cover the plate with aluminium foil to prevent bleaching of the fluorescent indicator.</li>
</ol>
4. Preparation of Cells for Flow Cytometry
<ol>
	<li>Warm 0.05% trypsin-ethylenediaminetetraacetic acid (EDTA) solution to 37 &#176;C.</li>
	<li>Aspirate calcium loading dye solution from each well and add 1 ml trypsin along the wall of the well.</li>
	<li>Incubate at 37 &#176;C until all cells are detached.</li>
	<li>Transfer cells to a 1.5 ml tube and pellet by centrifuging in a microcentrifuge at 10,000 g for 1 min.</li>
	<li>Carefully aspirate the supernatant without disturbing the pellet.</li>
	<li>Wash cells by adding 1 ml PHF buffer and resuspend by pipetting up and down.</li>
	<li>Pellet cells by centrifuging at 10,000 g for 1 min.</li>
	<li>Repeat steps 4.6. and 4.7. a further two times.</li>
	<li>Resuspend cells in a final volume of 500 &#181;l PHF buffer and use the cells for experimentation within 1 hr.</li>
	<li>Optional: count cells to determine the cell concentration and use 5 x 104 &#8211; 1 x 105 cells per measurement.</li>
</ol>
5. Flow Cytometer Setup
<ol>
	<li>Start the flow cytometer, open the fluidics drawer and flip the vent valve toggle switch to increase the air pressure of the sheath fluid tank.</li>
	<li>&#8220;Prime&#8221; the instrument to remove potential residing air bubbles from within the inner tubes and the flow chamber.</li>
	<li>In the flow cytometry software, open two dot plot windows.
	<ol>
		<li>The first dot plot requires forward scattered light (FSC) for the X parameter and side scattered light (SSC) for the Y parameter to control for the quality of the cells within the sample through their size and their granularity, respectively.</li>
		<li>The second dot plot measures the fluorescence intensity of the calcium-bound dye. Select the appropriate fluorescence detection channel for the Y parameter using a logarithmic scale and time in seconds for the X parameter on a linear scale.</li>
	</ol>
	</li>
	<li>Set the flow rate to &#8216;high&#8217;.</li>
</ol>
6. Flow Cytometry Measurement
<ol>
	<li>Perform the measurements at RT. In a flow cytometry sample tube (11.5 x 75 mm), add 480 &#181;l PHF buffer.</li>
	<li>Add 20 &#181;l cell suspension and briefly vortex to mix.</li>
	<li>Move the tube support arm to the side and position the sample tube over the sample injection tube until a tight seal is formed. Return the tube support arm to its original centered position.</li>
	<li>Optimize measurement by adjusting the fluorescence channel such that the mean fluorescence intensity of the sample is at approximately 102 on the y-axis. Save and use for subsequent experiments.</li>
	<li>To start an experiment, repeat steps 6.1 to 6.3.</li>
	<li>Record baseline fluorescence for 50 sec.</li>
	<li>Pause measurement, remove the sample tube, add compound of interest, vortex briefly and return to the sample injection tube. This step should not take more than 15 sec.</li>
	<li>Resume measurement and continue for a total of 204.8 sec.</li>
	<li>Use a calcium ionophore, such as ionomycin (10 &#181;M), and a calcium chelator, such as ethylene glycol tetraacetic acid (EGTA), MnCl2 or CoCl2 (2 mM), for positive and negative controls, respectively.</li>
</ol>
7. Data Analysis
<ol>
	<li>Analyze data using dedicated flow cytometry software for offline analysis, such as WinMDI (Windows Multiple Document Interface for Flow Cytometry), a free software package developed by Joe Trotter at The Scripps Institute, Flow Cytometry Core Facility.</li>
	<li>Open a dot plot with SSC and FSC parameters.</li>
	<li>Select the region of cells for analysis using the &#8220;Regions&#8221; tool (right-click on the image) to exclude dead cells or cell debris found in the bottom left corner from the data analysis (Figure 1).</li>
	<li>Quadrant analysis (Figure 2)
	<ol>
		<li>Open a density plot with time on the x-axis and fluorescence intensity on the y-axis. Use all events.</li>
		<li>Use the same gating region as in 7.3.</li>
		<li>Quantify the data by using the &#8220;Quadrant&#8221; tool.</li>
		<li>Adjust the quadrant separators so that the vertical line is placed at the time of compound addition and the horizontal line is placed at the center of the baseline fluorescence in the controls. Ensure that the positioning of the quadrant separators is equal in all samples.</li>
		<li>The percentage of cells in each quadrant is displayed in each corner (Figure 2). Compare the upper right quadrant (UR) between samples.</li>
	</ol>
	</li>
	<li>Histogram analysis (Figure 3)
	<ol>
		<li>Open the control data in a histogram window with fluorescence intensity on the x-axis and events on the y-axis. Use all events.</li>
		<li>For a pictorial overview, add the test data to be compared to the control plot as overlay plots (go to File &#8594; Open File &#8594; Select file &#8594; Overlay).</li>
		<li>For quantitative analysis, single histogram windows must be used. Gate the cell population using the region R1 from step 7.3.</li>
		<li>In the non-treated control, use the marker function (&#8220;Markers&#8221;) to mark the area for analysis starting from the midpoint between the minimum and maximum fluorescence of the control peak and ending at the maximum fluorescence intensity. The percentage of cells in this area is calculated.</li>
		<li>Retrieve data from the statistics window. Repeat for remaining data files using the same marked area for analysis.</li>
	</ol>
	</li>
	<li>Fluorescence intensity analysis (Figure 4)
	<ol>
		<li>Open a dot plot with time on the x-axis and fluorescence intensity on the y-axis. Use all events.</li>
		<li>Enable &#8220;Kinetics Mode&#8221; and &#8220;Overlay Kinetics Line&#8221;. A blue line will appear in the dot plot representing the mean fluorescence intensity.</li>
		<li>Quantify the relative calcium influx by creating multiple rectangular regions. Each region should have a width of about &#8220;50&#8221;, which is equivalent to &#8220;10 sec&#8221;. Define up to 15 rectangular regions.</li>
		<li>Retrieve quantification from the statistics window and copy into a spreadsheet.</li>
		<li>Use the y-mean data for analysis. Calculate the mean of R2 to R6, which is equivalent to 20 to 50 sec. This is the baseline value.</li>
		<li>Choose an appropriate time interval for calcium influx analysis (depends on onset, duration and intensity).</li>
		<li>Calculate the calcium signals from each region within this time interval relative to the mean baseline value, that is, the fluorescence signal before compound addition, which is set to 100%. Determine the mean and standard deviation of the regions post compound addition; this is the mean calcium signal evoked by the compound.</li>
	</ol>
	</li>
</ol>

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The authors have nothing to disclose.

Calcium is a key event in multiple cellular processes acting as a second messenger signaling molecule and also as a means to communicate between the ER and mitochondria. The ability to measure changes in free cytosolic calcium concentrations is a useful technique that can apply to all areas of cell biology. There are various methods to detect cytosolic calcium signals. Classically, signals from a ratiometric calcium indicator, such as Fura-2, are monitored in live cells using a fluorescence imaging setup and absolute calcium concentrations can be derived from determining minimum and maximum fluorescence intensities. However, this technique requires specialized equipment, such as a fluorescence microscope, a live cell imaging set up and analysis software. Here, we describe a simple method for calcium detection in the cytosol of adherent epithelial cells using flow cytometry, which is accessible to most laboratories.
Epithelia are in possession of an array of transporters responsible for the transport of metabolites, xenobiotics and drugs. Of particular importance is the SLC22A superfamily, to which organic cation and organic anion transporters belong 22, and the ATP-binding cassette (ATP) superfamily, which counts the multidrug resistance P-glycoprotein ABCB1 and multidrug resistance proteins (MRPs) amongst its members 23. Initial experiments using the rat kidney proximal tubule cell line WKPT-0293 Cl.2 were performed as described in the paper by Gergely et al.16. The cells were brought into suspension and loaded with Fluo-3 AM. However, only a modest change was observed with 10 &#181;M ionomycin (38.14% versus 50.95% in control by quadrant analysis or 1.78 fold by multiple region analysis). We hypothesized that epithelial transporters were responsible for the poor loading of the cells with Fluo-3 AM. To circumvent this problem, probenecid and PSC833, inhibitors of organic anion transporters and the multidrug resistance P-glycoprotein, respectively, were applied simultaneously with the calcium binding dye to cell monolayers. Probenecid improved calcium binding dye loading, as indicated by an increase in baseline fluorescence intensity, whereas PSC833 had no effect, which is in contrast to a previous report in Chinese hamster ovary cells24. This difference probably lies in the levels of ABCB1 in the different cell lines; the WKPT-0293 Cl.2 cell line harbors a low endogenous level of ABCB1, which can be induced by toxic substances, such as toxic metals25,26. The use of probenecid to improve calcium dye loading can be extended to other cell types that exhibit probenecid-sensitive transport systems, such as cells of the nervous system, blood-brain barrier and the liver, where calcium signaling may play a role in intracellular signaling pathways.
The easy detection of calcium binding to Fluo-3 lies in its requirement of only one excitation wavelength (506 nm) and one emission wavelength (525 nm). However, the data generated with single wavelength calcium binding dyes, as a non-ratiometric dye, can only be used to quantify relative increases in cytosolic calcium over non-treated controls. To determine absolute calcium concentrations, a ratiometric dye is necessary because a spectral shift occurs after calcium binding that allows for correction of unequal dye loading and bleaching. The ratiometric dye Indo-1 has been used in detection of calcium mobilization in immunological cells by flow cytometry27,28 but so far, this has not been performed in epithelial cells. Using the two emission spectra of UV-excitable Indo-1 (400 nm when calcium bound and 475 nm when calcium free), absolute cytosolic calcium concentrations can be determined as described by Grynkiewicz et al.11.
Our example data employed pharmacological compounds that induce a permanent change in cytosolic calcium. Physiological calcium signals are of low intensity, display fast kinetics and are transient, thus the question remains as to whether the flow cytometry method is sensitive enough to detect physiological calcium signals. Experiments with ATP were performed with the WKPT-0293 Cl.2 cell line, which harbors purinergic receptors as demonstrated by increased calcium signal upon application of ATP (measured by live cell imaging). Using the flow cytometer, increased cytosolic calcium could be observed with 1 - 100 &#181;M ATP, but the rapid nature of the signal combined with the delay in resuming measurement after compound addition meant that only a portion of the signal could be recorded. Thus a major disadvantage of the flow cytometry method is its limited capacity to record rapid (&#60;10 sec) and transient calcium signals; rather the method is more suitable for stimuli that give rise to slower transient signals (&#62;10 sec) or persistent elevated calcium signals. These limitations could be circumvented by reducing stimulator concentration or by applying inhibitors for calcium reuptake mechanisms to retain the released calcium in the cytosol but caution must be taken to select an inhibitor concentration that does not increase cytosolic calcium per se.
The information acquired by live cell imaging and flow cytometry are quite different. With live cell imaging, the high resolution means that spatial changes in calcium can be measured by selecting intracellular regions of single cells for imaging. Furthermore, fluorescence signals are acquired nonstop every 1 - 2 sec therefore rapid calcium signals can be recorded. In contrast, flow cytometry is more suited for slower and long lasting calcium signals and measures calcium signals from a whole cell population. Only relative changes in calcium signals can be quantified with the flow cytometry method. But, the flow cytometry method has a number of advantages over conventional live cell imaging: 1) high throughput screening of compounds (stimulators and inhibitors of calcium signaling) can be performed; 2) specific imaging expertise is not required; 3) research technicians can perform the measurements; 4) clinical and non-clinical laboratories that do not have the means to perform high resolution imaging and ratiometric calcium measurements can investigate calcium signaling. Taken together, the flow cytometry method is useful if the investigator initially wishes to establish a role for calcium and could easily apply inhibitors to determine upstream and downstream signaling pathways, whereas live cell imaging would be required to characterize the calcium signal (e.g., kinetics, intensity, spatial changes) in further detail.
In summary, we describe a quick, simple and sensitive method for the detection of real-time relative changes in cytosolic calcium in difficult-to-load adherent epithelial kidney cells, which have been brought into suspension. For experimenters without access to a flow cytometry, this method can be easily applied to a spectrofluorimeter with the cell suspension in a cuvette.

Calcium is an important second messenger responsible for the transmission of cellular physiological and pathophysiological signals1. Cytosolic calcium concentrations are kept low through the activity of calcium pumps and calcium exchangers. The sarcoplasmic/endoplasmic reticulum calcium ATPase (SERCA) refills the endoplasmic reticulum (ER) calcium store as part of a &#8220;pump leak&#8221; system whereas the plasma membrane calcium ATPase (PMCA) extrudes calcium to the extracellular compartment, both in ATP-dependent manners2. Physiological messengers, such as hormones or neurotransmitters, transmit their signals through G-protein coupled receptors, activation of phospholipase C, which hydrolyses phosphatidylinositol 4,5-bisphosphate (PIP2) in the plasma membrane to generate diacylglycerol and inositol triphosphate (IP3)3. While diacylglycerol remains in the plasma membrane, IP3 diffuses into the cytosol and binds to IP3 receptors (IP3Rs), which are ligand activated calcium channels, found in the ER membrane and calcium is released from the ER luminal store culminating in an increase in cytosolic calcium concentrations4. An alternative route for calcium to reach the cytosol is through calcium channels and exchangers present in the plasma membrane. Crosstalk between these two compartments have been described: calcium induced calcium release (CICR) where extracellular calcium induces calcium release from ER stores5, and store operated calcium entry (SOCE) where emptying of the ER stores is sensed by STIM and causes opening of Orai calcium channels in the plasma membrane to promote refilling of the ER stores6.
Under pathophysiological conditions, the cellular calcium response is deregulated and cytosolic calcium increases in the absence of physiological stimulators1. The calcium response may be affected in a number of ways: prolonged calcium signal, delayed calcium removal from the cytosol, depletion of calcium stores or localized changes in calcium. Furthermore, the mitochondria take on a central role in buffering and releasing calcium7. Prolonged and/or supramaximal cytosolic calcium increase leads to cell death. In fact, calcium is more often than not involved in the cellular pathophysiological response and is a key event in a wide array of diseases, such as neurodegenerative disease, heart disease, cancer, bone disease and kidney disease, which are mainly associated with cell death and loss or alteration of organ or tissue function8-10. Moreover, perturbation of calcium signals has been linked to cellular adaptation and changes in cellular functions and responses.
Traditionally, calcium signals are measured with a negatively charged fluorescent calcium indicator that is loaded into cells in a cell permeable acetoxymethyl (AM) ester form. Once cleaved by cellular esterases, the fluorescent indicators remain within the intracellular compartment and increase in fluorescence intensity when calcium is bound. The most well known and used calcium indicator is the ratiometric Fura-2 developed by Roger Tsien and colleagues11. Calcium indicators with differing affinities for calcium allow different calcium pools to be monitored. Detection methods include live cell imaging, fluorescence microplate reader and flow cytometry. The relatively fast kinetics of calcium signals (usually within a few seconds to minutes) makes live cell imaging the best method for gaining the most information about characteristics of the calcium signal. Aside from the fast kinetics of calcium signals (within milliseconds), live cell imaging is a better method for studying cellular compartmentalization of calcium signaling within a single cell 12. Absolute calcium concentrations can be calculated by determining the minimum and maximum fluorescence of the calcium indicator by adding a calcium chelator and permeabilizing the cells, respectively, as described by Grynkiewicz et al.11.
The use of flow cytometry to measure calcium signals was first developed in the 1980s in immunological cells where the opportunity to measure multiple parameters, such as membrane integrity and separation of cell population, and the requirement of cells in suspension combined with the development of single wavelength calcium indicators made flow cytometry an ideal and convenientdetection method13-15. In adherent cells, live cell imaging provides the most information about calcium signaling, most importantly the kinetics, but requires a complicated setup comprising a fluorescence microscope, a perfusion system, maintenance of the cellular environment, such as temperature, and specialized microscope software for acquiring and analyzing data. Alternative methods such as a fluorescence microplate reader or pharmacological means through use of calcium chelators are incomparable in terms of gained information. Flow cytometry is becoming more widely used to monitor cytosolic calcium in adherent cells though, in the majority of studies, only end-point measurements are performed. Using the method initially developed by Gergely et al. in immunological B cells16, changes in cytosolic free calcium concentration by flow cytometry with real-time kinetics and monitoring the fluorescence intensity in individual cells from a sample population have been successfully measured in cancer epithelial cells and cancer stem cell hybrids17. This method was further adapted for use in adherent epithelial cells, which are difficult to load with calcium indicators 18.
Here, using an immortalized adherent epithelial cell line derived from the S1 segment of the rat kidney proximal tubule (WKPT-0293 Cl.2)19, we describe an optimized simplified method for measurement of changes in cytosolic calcium concentration by flow cytometry. Since many epithelial cells possess a number of organic transporters for anionic compounds, loading of cells with a calcium indicator can prove to be a challenge. To prevent the efflux of calcium indicators, probenecid, the prototypical inhibitor of organic anion transporters and originally developed to decrease renal excretion of penicillin20, is applied simultaneously in the incubation medium. The cells are maintained in monolayers for calcium indicator loading, brought into suspension and calcium signals can be detected immediately after compound addition.

<table><tbody><tr><td>Fluo-3, AM</td><td>Invitrogen</td><td>F-1241&amp;nbsp;</td><td>Cell permeable</td></tr><tr><td>Probenecid</td><td>Sigma-Aldrich</td><td>P-8761</td><td>Water insoluble, dissolve in 1 N NaOH</td></tr><tr><td>0.05% Trypsin-EDTA (1X)</td><td>Invitrogen</td><td>25300-062</td><td></td></tr><tr><td>Ionomycin</td><td>Sigma-Aldrich</td><td>I9657</td><td></td></tr><tr><td>Thapsigargin</td><td>Tocris Bioscience</td><td>1138</td><td></td></tr><tr><td>Tunicamycin</td><td>Sigma-Aldrich</td><td>T7765</td><td></td></tr><tr><td>FACSCalibur Flow Cytometer + CELLQuest software</td><td>Becton Dickinson</td><td></td><td></td></tr><tr><td>Windows Multiple Document Interface for Flow Cytometry (WinMDI)</td><td>Joe Trotter, The Scripps Institute</td><td></td><td>Please note that this is an older 16-bit application that reads FCS 2.0 compliant files but does not recognize FCS 3.0 digital data.</td></tr><tr><td>WKPT-0293 Cl.2 rat kidney proximal tubule cell line</td><td></td><td></td><td>Made available by Dr. Ulrich Hopfer (Department of Physiology &amp;amp; Biophysics, Case&lt;br/&gt; Western Reserve University, Cleveland, OH)</td></tr></tbody></table>

cytosolic calcium measurements in renal epithelial cells by flow cytometry

A variety of cellular processes, both physiological and pathophysiological, require or are governed by calcium, including exocytosis, mitochondrial function, cell death, cell metabolism and cell migration to name but a few. Cytosolic calcium is normally maintained at low nanomolar concentrations; rather it is found in high micromolar to millimolar concentrations in the endoplasmic reticulum, mitochondrial matrix and the extracellular compartment. Upon stimulation, a transient increase in cytosolic calcium serves to signal downstream events. Detecting changes in cytosolic calcium is normally performed using a live cell imaging set up with calcium binding dyes that exhibit either an increase in fluorescence intensity or a shift in the emission wavelength upon calcium binding. However, a live cell imaging set up is not freely accessible to all researchers. Alternative detection methods have been optimized for immunological cells with flow cytometry and for non-immunological adherent cells with a fluorescence microplate reader. Here, we describe an optimized, simple method for detecting changes in epithelial cells with flow cytometry using a single wavelength calcium binding dye. Adherent renal proximal tubule epithelial cells, which are normally difficult to load with dyes, were loaded with a fluorescent cell permeable calcium binding dye in the presence of probenecid, brought into suspension and calcium signals were monitored before and after addition of thapsigargin, tunicamycin and ionomycin.

To increase cytosolic calcium, two pharmacological compounds were applied to renal proximal tubule cells (WKPT-0293 Cl.2) loaded with cell permeable calcium binding dye, Fluo-3-AM. Nontreated control samples were loaded with calcium binding dye in the presence of probenecid and were mixed but without the addition of any compounds. Thapsigargin (TG) is a classical inhibitor of the SERCA pump leading to leakage out of the ER and resulting in increased cytosolic calcium. Tunicamycin (TUN) blocks N-glycosylation of proteins leading to activation of the unfolded protein response but also increases cytosolic calcium18,21. As a positive control, ionomycin (IONO), a calcium ionophore making the plasma membrane permeable for extracellular calcium, was used. For analysis, the cell population was gated by selecting a region in the SSC/FSC dot plot (R1 in Figure 1). A density plot for fluorescence intensity vs. time was created for each data file. By using the quadrant function, the dot plot was separated into four areas representing Fluo-3 fluorescence intensity before and after compound addition and above and below the baseline fluorescence. Quantification as percentage of gated cell population in each quadrant allows determination of increased cytosolic calcium concentration. In this example, 3 &#181;M TG causes an increase in the percentage of cells in the upper right quadrant indicating that a calcium signal has been induced. The percentage of cells increases from 42.8% in control to 51.3% (1.20 fold) in TG treated cells. Similarly, 6 &#181;M TUN augments the percentage of cells with increased calcium fluorescence intensity to 49.9% (1.17 fold). Application of 10 &#181;M IONO resulted in 65.5% (1.53 fold) of cells in the upper right quadrant (Figure 2). The histogram analysis resulted in 1.41 fold, 1.36 fold and 2.11 fold by TG, TUN and IONO, respectively (Figure 3) (for statistics, see 18). Finally, using the multiple region analysis mode, TG, TUN and IONO caused 1.55 fold, 1.29 fold and 4.54 fold increases in calcium signal over control, respectively (Figure 4).
The most sensitive analysis mode was the multiple region analysis. In contrast to the quadrant and histogram analysis modes, where the distribution of the cell population is quantified, the multiple region analysis mode quantifies the change in fluorescent signal from the total cell population. The largest discrepancy in the results from different analysis modes was for ionomycin that ranged from 1.53 fold increase over control from the quadrant analysis to 4.54 fold from the multiple region analysis. The compounds tested did not exhibit such variance. However, throughout all analysis modes, the extent of calcium signal evoked remained the same though the absolute values differed, i.e., IONO had the largest effect followed by TG and then TUN.
Application of TG, TUN or IONO results in permanent changes in cytosolic calcium. To determine whether physiological calcium signals, which are transient, can be measured using this method, ATP was applied to WKPT-0293 Cl.2 cells. ATP evoked a rapid and transient calcium signal, which could not be recorded in its entirety by the flow cytometry method. ATP concentrations at 20 - 100 &#181;M were difficult to detect therefore the concentration was reduced to &#8220;slow down&#8221; the calcium signals. As shown in Figure 5, a change in cytosolic calcium could be measured after addition of 5 &#181;M ATP. Still, the peak of fluorescence change was not recorded. Using the quadrant analysis (Figure 5B) and histogram analysis (Figure 5C) modes, no difference in calcium signals in ATP could be detected due to the transient nature of the calcium signal. However, with selected parts of the trace, the multiple rectangular region analysis mode recorded a 1.85 fold increase in calcium signal immediately after resumed measurement (Figure 5A).
<img alt="Figure 1" fo:content-width="5in" src="/files/ftp_upload/51857/51857fig1highres.jpg" width="500" /> 
Figure 1. SSC versus FSC dot plot. The integrity of the cell population is assessed using side scatter (SSC) and forward scatter (FSC). A region is selected (R1) for further analysis. Dead cells and cell debris, which have reduced light scattering, are excluded. A representative dot plot is shown.
<img alt="Figure 2" fo:content-width="5in" src="/files/ftp_upload/51857/51857fig2highres.jpg" width="500" /> 
Figure 2. Quadrant analysis. Baseline fluorescence of calcium binding dye-loaded WKPT-0293 Cl.2 cells was measured for 50 sec. Measurement was stopped, compound or diluent was added, vortexed to mix and measurement was resumed immediately for a total of 204.8 sec. Quantification was performed using quadrant analysis. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/51857/51857fig2highres.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" fo:content-width="5in" src="/files/ftp_upload/51857/51857fig3highres.jpg" width="500" /> 
Figure 3. Histogram analysis. Histogram plot of calcium binding dye-loaded WKPT-0293 Cl.2 cells treated with TG, TUN or IONO. Using the marker function, the percentage of gated cells with high calcium binding dye fluorescence intensity was quantified by positioning the starting point of the marker at the peak of the control curve and ending at the maximum fluorescence. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/51857/51857fig3highres.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" fo:content-width="5in" src="/files/ftp_upload/51857/51857fig4highres.jpg" width="500" /> 
Figure 4. Multiple region analysis. Baseline fluorescence of calcium binding dye-loaded WKPT-0293 Cl.2 cells was measured for 50 sec. Measurement was stopped, compound or diluent was added, vortexed to mix and measurement was resumed immediately for a total of 204.8 sec. Quantification was performed using multiple rectangular region analysis. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/51857/51857fig4highres.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" fo:content-width="5in" src="/files/ftp_upload/51857/51857fig5highres.jpg" width="500" /> 
Figure 5. Analysis of ATP-induced calcium signals. Baseline fluorescence of calcium binding dye-loaded WKPT-0293 Cl.2 cells was measured for 50 sec. Measurement was stopped, 5 &#181;M ATP was added, vortexed to mix and measurement was resumed immediately for a total of 204.8 sec. Data was quantified using multiple rectangular region analysis (A), quadrant analysis (B) or histogram analysis (C). For the multiple region analysis, only the transient calcium signal was analyzed. Representative data or analysis from n = 3 - 7 are shown. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/51857/51857fig5highres.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Cytosolic Calcium Measurements in Renal Epithelial Cells by Flow Cytometry at JoVE.com

Cytosolic Calcium Measurements in Renal Epithelial Cells by Flow Cytometry

Calcium is involved in numerous physiological and pathophysiological signaling pathways. Live cell imaging requires specialized equipment and can be time consuming. A quick, simple method using a flow cytometer to determine relative changes in cytosolic calcium in adherent epithelial cells brought into suspension was optimized.

A variety of cellular processes, both physiological and pathophysiological, require or are governed by calcium, including exocytosis, mitochondrial ...

cytosolic-calcium-measurements-renal-epithelial-cells-flow

Nanyang Technological University

Research

JoVE Journal

Biology

15.2K Views.  University of Witten/Herdecke.  Calcium is betrokken bij tal van fysiologische en pathofysiologische signaalwegen. Live cell imaging vereist gespecialiseerde apparatuur en kan tijdrovend zijn. Een snelle, eenvoudige werkwijze met een flow cytometer om relatieve veranderingen in cytosolische calcium in hechtende epitheelcellen gebracht suspensie bepalen geoptimaliseerd. 

Video: Cytosolic Calcium Measurements in Renal Epithelial Cells by Flow Cytometry

Measuring Calpain Activity in Fixed and Living Cells by Flow Cytometry

Calpains are ubiquitous intracellular, calcium-sensitive, neutral cysteine proteases 1. Calpains play crucial roles in many physiological processes, including signaling, cytoskeletal remodeling, regulation of gene expression, apoptosis and cell cycle progression 1. Calpains have been implicated in many pathologies including muscular dystrophies, cancer, diabetes, Alzheimer's disease and multiple sclerosis 1. Calpain regulation is complex and incompletely understood. mRNA and protein levels correlate poorly with activity, limiting the use of gene or protein expression techniques to measure calpain activity. This video protocol details a flow cytometric assay developed in our laboratory for measuring calpain activity in fixed and living cells. This method uses the fluorescent substrate BOC-LM-CMAC, which is cleaved specifically by calpain, to measure calpain activity. 2 In this video, calpain activity in fixed and living murine 32Dkit leukemia cells, alone or as part of a splenocyte population is measured using an LSRII (BD Bioscience). 32Dkit cells are shown to have elevated activity compared to normal splenocytes.

This article will detail the protocol for measuring calpain activity in fixed and living cells using flow cytometry.

Calpains are ubiquitous intracellular, calcium-sensitive, neutral cysteine proteases 1. Calpains play crucial roles in many physiological processes, ...

Immunology and Infection

Real-time Live-cell Flow Cytometry to Investigate Calcium Influx, Pore Formation, and Phagocytosis by P2X7 Receptors in Adult Neural Progenitor Cells

Live-cell flow cytometry is increasingly used among cell biologists to quantify biological processes in a living cell culture. This protocol describes a method whereby live-cell flow cytometry is extended upon to analyze the multiple functions of P2X7 receptor activation in real-time. Using a time module installed on a flow cytometer, live-cell functionality can be assessed and plotted over a given time period to explore the kinetics of calcium influx, transmembrane pore formation, and phagocytosis. This simple method is advantageous as all three canonical functions of the P2X7 receptor can be assessed using one machine, and the gathered data plotted over time provides information on the entire live-cell population rather than single-cell recordings often obtained using technically challenging patch-clamp methods. Calcium influx experiments use a calcium indicator dye, while P2X7 pore formation assays rely on ethidium bromide being allowed to pass through the transmembrane pore formed upon high agonist concentrations. Yellow-green (YG) latex beads are utilized to measure phagocytosis. Specific agonists and antagonists are applied to investigate the effects of P2X7 receptor activity. Individually, these methods can be modified to provide quantitative data on any number of calcium channels and purinergic and scavenger receptors. Taken together, they highlight how the use of real-time live-cell flow cytometry is a rapid, cost-effective, reproducible, and quantifiable method to investigate P2X7 receptor function.

Providing single-cell sensitivity, real-time flow cytometry is uniquely suited to quantify multimodal receptor functions of live cultures. Using adult neural progenitor cells, the P2X7 receptor function was assessed via calcium influx detected by calcium indicator dye, transmembrane pore formation by ethidium bromide uptake, and phagocytosis using fluorescent latex beads.

Live-cell flow cytometry is increasingly used among cell biologists to quantify biological processes in a living cell culture. This protocol describes a ...

Developmental Biology

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.

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 ...

Analysis of T-cell Receptor-Induced Calcium Influx in Primary Murine T-cells by Full Spectrum Flow Cytometry

Calcium influx in response to T-cell receptor stimulation is a common measure of T-cell signaling. Several calcium indicator dyes have been developed to assess calcium signaling by band-pass flow cytometry. This protocol is designed to measure calcium responses in primary murine T-cells using full spectrum flow cytometry. Total splenocytes are labeled with the ratiometric calcium indicator dye Indo-1, along with a panel of fluorochrome-conjugated antibodies to cell surface molecules. Leveraging the capabilities of full spectrum flow cytometry provides a platform for utilizing a wide array of cell surface stains in combination with Indo-1. Cells are then analyzed in real-time at 37 &#176;C before and after the addition of an anti-CD3 antibody to stimulate the T-cell receptor. After unmixing the spectral signals, the ratio of calcium-bound to calcium-free Indo-1 is calculated and can be visualized over time for each gated population of splenocytes. This technique can allow for the simultaneous analysis of calcium responses in multiple cell populations.

Calcium influx, a measure of T-cell signaling, is an effective way to analyze responses to T-cell receptor stimulation. This protocol for multiplexing Indo-1 with panels of antibodies directed at cell surface molecules takes advantage of the highly flexible capabilities of full spectrum flow cytometry.

Calcium influx in response to T-cell receptor stimulation is a common measure of T-cell signaling. Several calcium indicator dyes have been developed to ...

Single-channel Analysis and Calcium Imaging in the Podocytes of the Freshly Isolated Glomeruli

Podocytes (renal glomerular epithelial cells) are known to regulate glomerular permeability and maintain glomerular structure; a key role for these cells in the pathogenesis of various renal diseases has been established since podocyte injury leads to proteinuria and foot process effacement. It was previously reported that various endogenous agents may cause a dramatic overload in intracellular Ca2+ concentration in podocytes, presumably leading to albuminuria, and this likely occurs via calcium-conducting ion channels. Therefore, it appeared important to study calcium handling in the podocytes both under normal conditions and in various pathological states. However, available experimental approaches have remained somewhat limited to cultured and transfected cells. Although they represent a good basic model for such studies, they are essentially extracted from the native environment of the glomerulus. Here we describe the methodology of studying podocytes as a part of the freshly isolated whole glomerulus. This preparation retains the functional potential of the podocytes, which are still attached to the capillaries; therefore, podocytes remain in the environment that conserves the major parts of the glomeruli filtration apparatus. The present manuscript elaborates on two experimental approaches that allow 1) real-time detection of calcium concentration changes with the help of ratiometric confocal fluorescence microscopy, and 2) the recording of the single ion channels activity in the podocytes of the freshly isolated glomeruli. These methodologies utilize the advantages of the native environment of the glomerulus that enable researchers to resolve acute changes in the intracellular calcium handling in response to applications of various agents, measure basal concentration of calcium within the cells (for instance, to evaluate disease progression), and assess and manipulate calcium conductance at the level of single ion channels.

Changes in the intracellular calcium levels in the podocytes are one of the most important means to control the filtration function of glomeruli. Here we explain a high-throughput approach that allows detection of real-time calcium handling and single ion channels activity in the podocytes of the freshly isolated glomeruli.

Podocytes (renal glomerular epithelial cells) are known to regulate glomerular permeability and maintain glomerular structure; a key role for these cells ...

Implementing Patch Clamp and Live Fluorescence Microscopy to Monitor Functional Properties of Freshly Isolated PKD Epithelium

Cyst initiation and expansion during polycystic kidney disease is a complex process characterized by abnormalities in tubular cell proliferation, luminal fluid accumulation and extracellular matrix formation. Activity of ion channels and intracellular calcium signaling are key physiologic parameters which determine functions of tubular epithelium. We developed a method suitable for real-time observation of ion channels activity with patch-clamp technique and registration of intracellular Ca2+ level in epithelial monolayers freshly isolated from renal cysts. PCK rats, a genetic model of autosomal recessive polycystic kidney disease (ARPKD), were used here for ex vivo analysis of ion channels and calcium flux. Described here is a detailed step-by-step procedure designed to isolate cystic monolayers and non-dilated tubules from PCK or normal Sprague Dawley (SD) rats, and monitor single channel activity and intracellular Ca2+ dynamics. This method does not require enzymatic processing and allows analysis in a native setting of freshly isolated epithelial monolayer. Moreover, this technique is very sensitive to intracellular calcium changes and generates high resolution images for precise measurements. Finally, isolated cystic epithelium can be further used for staining with antibodies or dyes, preparation of primary cultures and purification for various biochemical assays.

Ion channels expressed in renal tubular epithelium play a significant role in the pathology of polycystic kidney disease. Here we describe experimental protocols used to perform patch-clamp analysis and intracellular calcium level measurements in cystic epithelium freshly isolated from rodent kidneys.

Cyst initiation and expansion during polycystic kidney disease is a complex process characterized by abnormalities in tubular cell proliferation, luminal ...

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 ...

Determination of the Relative Cell Surface and Total Expression of Recombinant Ion Channels Using Flow Cytometry

Inherited or de novo mutations in cation-selective channels may lead to sudden cardiac death. Alteration in the plasma membrane trafficking of these multi-spanning transmembrane proteins, with or without change in channel gating, is often postulated to contribute significantly in this process. It has thus become critical to develop a method to quantify the change of the relative cell surface expression of cardiac ion channels on a large scale. Herein, a detailed protocol is provided to determine the relative total and cell surface expression of cardiac L-type calcium channels CaV1.2 and membrane-associated subunits in tsA-201 cells using two-color fluorescent cytometry assays. Compared with other microscopy-based or immunoblotting-based qualitative methods, flow cytometry experiments are fast, reproducible, and large-volume assays that deliver quantifiable end-points on large samples of live cells (ranging from 104 to 106 cells) with similar cellular characteristics in a single flow. Constructs were designed to constitutively express mCherry at the intracellular C-terminus (thus allowing a rapid assessment of the total protein expression) and express an extracellular-facing hemagglutinin (HA) epitope to estimate the cell surface expression of membrane proteins using an anti-HA fluorescence conjugated antibody. To avoid false negative, experiments were also conducted in permeabilized cells to confirm the accessibility and proper expression of the HA epitope. The detailed procedure provides: (1) design of tagged DNA (deoxyribonucleic acid) constructs, (2) lipid-mediated transfection of constructs in tsA-201 cells, (3) culture, harvest, and staining of non-permeabilized and permeabilized cells, and (4) acquisition and analysis of fluorescent signals. Additionally, the basic principles of flow cytometry are explained and the experimental design, including the choice of fluorophores, titration of the HA antibody and control experiments, is thoroughly discussed. This specific approach offers objective relative quantification of the total and cell surface expression of ion channels that can be extended to study ion pumps and plasma membrane transporters.

Inherited cardiac arrhythmias are often caused by mutations that alter the surface delivery of one or more ion channels. Here, we adapt flow cytometry assays to provide a quantification of the relative total and cell surface protein expression of recombinant ion channels expressed in tsA-201 cells.

Inherited or de novo mutations in cation-selective channels may lead to sudden cardiac death. Alteration in the plasma membrane trafficking of these ...

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.

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.

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

Single Cell Calcium Imaging for Channel Activation Studies

Single-Cell Calcium Imaging for Studying the Activation of Calcium Ion Channels

Single cell Ca2+ imaging is essential for the study of Ca2+ channels activated by various stimulations like temperature, voltage, native compound and chemicals et al. It primarily relies on microscopy imaging technology and the related Ca2+ indicator Fura-2/AM (AM is the abbreviation for Acetoxymethyl ester). Inside the cells, Fura-2/AM is hydrolyzed by esterases into Fura-2, which can reversibly bind with free cytoplasmic Ca2+. The maximum excitation wavelength shifts from 380nm to 340nm (when saturated with Ca2+) upon binding. The emitted fluorescence intensity is quantitatively related to the concentration of bound Ca2+. By measuring the 340/380 ratio, the Ca2+ concentration in the cytoplasm can be determined, eliminating errors caused by variations in the loading efficiency of the fluorescent probe among different samples. This technology allows for real-time, quantitative, and simultaneous monitoring of Ca2+ changes in multiple cells. The results are stored in &#8220;.XLSX&#8221; format for subsequent analysis, which is fast and generates intuitive change curves, greatly improving the detection efficiency. From different experimental perspectives, this article lists the use of this technology to detect Ca2+ signals in cells with endogenous or overexpressed channel proteins. Meantime, different methods for activating cells were also showed and compared. The aim is to provide readers with a clearer understanding of the usage and applications of single cell Ca2+ imaging.

This article presents a method for real-time, quantitative monitoring of calcium ion (Ca2+) concentrations in cells using single-cell Ca2+ imaging with the Fura-2/AM dye. This technique enables efficient dye loading and accurate calculation of Ca2+ levels through fluorescence intensity ratios, making it a simple and rapid approach for research applications.

Single cell Ca2+ imaging is essential for the study of Ca2+ channels activated by various stimulations like temperature, voltage, native compound and ...

Cytosolic Calcium Measurements in Renal Epithelial Cells by Flow Cytometry

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