This work was supported by NIH grant R15ES030140 and NSF grant CBET1903568. Financial support from the Russ College of Engineering and Technology and the Department of Chemical and Biomolecular Engineering at Ohio University is also acknowledged.

All human blood samples used in the protocol described below were purchased as de-identified samples. No human subjects were directly involved or recruited for this study. The guidelines of the Declaration of Helsinki should be used when research involves human subjects.
1. Erythrocyte isolation from whole blood
<ol>
	<li>Add 500 &#181;L of whole blood in acid citrate dextrose (ACD) (stored at 4 &#176;C) to a microcentrifuge tube. 
	NOTE: Whole blood was purchased in ACD. According to the company, 1.5 mL of ACD is added to 7 mL of whole blood (8.5 mL total volume).</li>
	<li>Centrifuge the whole blood at 700 x g for 5 min at room temperature (RT) and remove the clear plasma and the thin buffy coat using a pipette to leave the red erythrocyte layer.</li>
	<li>Prepare 1 L of Ringer solution containing 125 mM NaCl, 5 mM KCl, 1 mM MgSO4, 32 mM HEPES, 5 mM glucose, and 1 mM CaCl2. Adjust the pH to 7.4 by adding 2 &#181;L drops of 1.0 M NaOH. To prepare glucose-free Ringer solution, follow the same protocol, but do not include glucose in the solution.</li>
	<li>Wash the erythrocytes 2x in Ringer solution by suspending the cell pellet in 1.5 mL of Ringer solution, centrifuging at 700 x g for 5 min at RT, and removing the supernatant.</li>
	<li>Make a 0.4% hematocrit by resuspending 40 &#181;L of the erythrocyte pellet in 9,960 &#181;L of glucose-free Ringer solution to reach a final volume of 10 mL. 
	NOTE: Hematocrit is a term used to refer to the volume fraction of erythrocytes in suspension. A 0.4% hematocrit is a suspension containing 0.4% erythrocytes.</li>
	<li>Incubate the cell suspension at 37 &#176;C for 7 days.</li>
</ol>
2. Treatment of erythrocytes with ionomycin and measurement of hemolysis
<ol>
	<li>Dissolve 1 mg of ionomycin calcium salt in 630 &#181;L of dimethyl sulfoxide (DMSO) to reach a final concentration of 2 mM. Aliquot and store at -20 &#176;C.</li>
	<li>Take 1 mL of the 0.4% hematocrit from step 1.5 and add 0.5 &#181;L of 2 mM ionomycin to reach a final concentration of 1 &#181;M. Incubate for 2 h at 37 &#176;C.
	<ol>
		<li>Use 1 mL of the hematocrit with no ionomycin treatment as a negative control.</li>
	</ol>
	</li>
	<li>Centrifuge the ionomycin-treated and untreated hematocrits at 700 x g for 5 min at RT, and remove their supernatants to leave the cell pellets at the bottom of the tubes.</li>
	<li>Wash the cells 3x with Ringer solution by suspending the cell pellets in 1.5 mL of Ringer solution, centrifuging at 700 x g for 5 min at RT and discarding the supernatants.</li>
	<li>To measure hemolysis, add 1 mL of the untreated 0.4% hematocrit from step 1.5 to a microcentrifuge tube and incubate for 2 h at 37 &#176;C as the negative control for hemolysis (0%).</li>
	<li>Add 1 mL of the untreated 0.4% hematocrit from step 1.5 to a microcentrifuge tube and centrifuge at 700 x g for 5 min at RT. Remove the supernatant and add 1 mL of distilled water to the cell pellet and incubate for 2 h at 37 &#176;C as the positive control for hemolysis (100%).</li>
	<li>Add 1 mL of the ionomycin-treated 0.4% hematocrit from step 2.2 to a microcentrifuge tube.</li>
	<li>Centrifuge the untreated cells, treated cells, and the cells in distilled water at 700 x g for 5 min at RT.</li>
	<li>Take 200 &#181;L of the supernatants and add to a 96-well plate.</li>
	<li>Measure the absorbance at 541 nm using a microplate reader.</li>
	<li>Calculate the hemolysis using Equation 132: 
	 
	%Hemolysis = (AT - A0)/(A100 - A0)*100 
	 
	where A0 is the absorbance of erythrocytes in Ringer solution, A100 is the absorbance of erythrocytes in water, and AT is the absorbance of treated erythrocytes by ionomycin.</li>
</ol>
3. Annexin-V binding assay
<ol>
	<li>Dilute 2 mL of the 5x annexin V binding buffer in 8 mL of phosphate-buffered saline (PBS) to obtain 1x binding buffer.</li>
	<li>Resuspend the ionomycin-treated and untreated cell pellets from step 2.4 in 1 mL of 1x binding buffer.</li>
	<li>Take 235 &#181;L of the cell suspensions in the binding buffer and add 15 &#181;L of Annexin V-Alexa Flour 488 conjugate.</li>
	<li>Incubate the cells at RT for 20 min in a dark place. Centrifuge at 700 x g for 5 min at RT and remove the supernatant.</li>
	<li>Wash the cells 2x with 1x binding buffer, by suspending the cell pellet in 1.5 mL of the binding buffer, centrifuging at 700 x g for 5 min at RT and removing the supernatant.</li>
	<li>Resuspend the cell pellets in 250 &#181;L of 1x binding buffer for flow cytometry measurements.</li>
</ol>
4. Flow cytometry
<ol>
	<li>Transfer 200 &#181;L of the annexin-V stained erythrocytes to 1 mL round bottom polystyrene tubes compatible with flow cytometry.</li>
	<li>Login to the flow cytometry software and click on the new experiment button. Click on the new tube button. Select the global sheet and choose the apply analysis to measure the fluorescence intensity with an excitation wavelength of 488 nm and an emission wavelength of 530 nm.</li>
	<li>Set number of cells to 20,000 to be collected for fluorescence-activated cell sorting (FACS) analysis.</li>
	<li>Select the desired tube and click on load button. Click on record button for forward scatter and side scatter measurements. Repeat for all samples.</li>
	<li>Right click on specimen button and click on apply batch analysis to generate the result file.</li>
	<li>Right click on specimen button and click on generate FSC files.</li>
	<li>Add the flow cytometry data (FSC files) into the workplace of flow cytometry software.</li>
	<li>Analyze the control data by selecting the cell population of interest and adding statistics for eryptosis value.
	<ol>
		<li>Double click on control and select histogram versus fluorescence intensity.</li>
		<li>Click on gate button to draw a gate on the histogram which indicates the percentage of eryptosis.</li>
	</ol>
	</li>
	<li>Apply the same statistics for all other experimental tubes to obtain the eryptosis values. Right click on control and select copy analysis to group.</li>
	<li>After properly gating all samples, transfer the analyzed data by dragging and dropping them into the layout editor.
	<ol>
		<li>Overlay the analyzed data with control in layout editor.</li>
		<li>Set the desired histograms and intensities by changing the x and y axis of the overlaid graphs.</li>
		<li>Export image files by clicking on export button and save the graphs in desired location.</li>
	</ol>
	</li>
</ol>
5. Confocal microscopy
<ol>
	<li>Transfer 5 &#181;L of annexin-V-stained cells on a microscope slide and cover it with a cover slip. Keep in a dark place to prevent photobleaching.</li>
	<li>Use Argon laser of the confocal fluorescence microscope to observe the cells excited at 488 nm with desired magnifications. 
	NOTE: A confocal microscope is not necessarily needed and any microscope with fluorescence capabilities can be used to obtain fluorescence images that demonstrate annexin-V binding.</li>
	<li>Obtain fluorescence images of the control (non-treated cells) and treated cells. 
	NOTE: Non-treated cells are expected to show very weak fluorescence signals, whereas treated cells are expected to show bright green fluorescence on their membranes.</li>
</ol>

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The authors have nothing to disclose.

The goal of this procedure is to provide optimal values for ionophore concentration, treatment time, and extracellular glucose concentration, which are important factors in ensuring successful induction of eryptosis. A critical step in the protocol is the depletion of extracellular glucose, which, despite its importance, has not been sufficiently emphasized in the literature. The sugar content in normal Ringer solution (5 mM) has an inhibitory effect on eryptosis. Glucose depletion in the extracellular environment induce...

Programmed cell death in erythrocytes, also known as eryptosis, is common in many clinical conditions and hematological disorders. Eryptosis is associated with cell shrinkage and the loss of phospholipid asymmetry in the cell plasma membrane1,2. Loss of asymmetry results in the translocation of phosphatidylserine (PS), a lipid normally localized in the inner leaflet3,4, to the cell outer leaflet, which signals to macrophages to phagocytose and remove defective erythrocytes5,6,7,8. At the end of the normal life span of erythrocytes, removal of eryptotic cells by macrophages ensures the balance of erythrocytes in circulation. However, in diseased conditions, such as sickle cell disease and thalassemia9,10,11, enhanced eryptosis may result in severe anemia2. Due to its importance in hematological diseases, there is significant interest in examining the factors inducing or inhibiting eryptosis and the molecular mechanisms underlying this process.
The plasma membrane of healthy erythrocytes is asymmetric, with different phospholipids localizing at the outer and inner leaflets. Membrane asymmetry is primarily regulated by the action of membrane enzymes. Aminophospholipid translocase facilitates the transport of aminophospholipids, PS and phosphatidylethanolamine (PE), by directing these lipids to the cell inner leaflet. On the other hand, floppase transports the choline containing phospholipids, phosphatidylcholine (PC) and sphingomyelin (SM), from the inner to the outer leaflet of the cell membrane12. However, unlike healthy cells, the membrane of eryptotic erythrocytes is scrambled. This is due to the action of a third enzyme, scramblase, which disrupts phospholipid asymmetry by facilitating the bidirectional transport of aminophospholipids13,14,15,16. Scramblase is activated by elevated intracellular levels of Ca2+. Therefore, calcium ionophores, which facilitate the transport of Ca2+ across the cell membrane12, are efficient inducers of eryptosis.
Ionomycin, a calcium ionophore, has been widely used to induce eryptosis in erythrocytes12,17,18,19,20,21,22,23,24,25,26. Ionomycin has both hydrophilic and hydrophobic groups, which are necessary to bind and capture Ca2+ ion, and transport it to the cytosolic space27,28,29. This leads to the activation of scramblase and translocation of PS to the outer leaflet, which can be easily detected using annexin-V, a cellular protein with a high affinity to PS12. Although triggering eryptosis by ionomycin is commonly reported, there is considerable method discrepancy in the literature (Table 1). The population of erythrocytes undergoing eryptosis depends on different factors such as ionophore concentration, treatment time with ionophore, and the sugar content of extracellular environment (glucose depletion activates cation channels and facilitates the entry of Ca2+ into the cytosolic space)30,31. However, there is little consistency in these factors in the literature, making it difficult to perform eryptosis reproducibly in vitro.
In this protocol, we present a step-by-step procedure to induce eryptosis in human erythrocytes. Factors affecting successful eryptosis including Ca2+ concentration, ionophore concentration, treatment time, and pre-incubation in glucose-depleted buffer are examined and optimal values are reported. This procedure demonstrates that pre-incubation of erythrocytes in a glucose-free buffer significantly increases the percentage of eryptosis compared to glucose-containing buffer. This protocol can be used in the laboratory to produce eryptotic erythrocytes for various applications.

<table><tbody><tr><td>96-well plate</td><td>Fisher Scientific</td><td>12-565-331</td><td></td></tr><tr><td>Annexin V Alexa Fluor 488 - apoptosis kit</td><td>Fisher Scientific</td><td>A10788</td><td>Store at 4 &amp;deg;C</td></tr><tr><td>BD FACSAria II flow cytometer</td><td>BD Biosciences</td><td>643177</td><td></td></tr><tr><td>CaCl&lt;sub&gt;2&lt;/sub&gt;</td><td>Fisher Scientific</td><td>C79-500</td><td></td></tr><tr><td>Centrifuge</td><td>Millipore Sigma</td><td>M7157</td><td>Model Eppendorf 5415C</td></tr><tr><td>Confocal fluorescence microscopy</td><td>Zeiss, LSM Tek Thornwood</td><td></td><td>Model LSM 510, Argon laser excited at 488 nm for taking images</td></tr><tr><td>Cover glasses circles</td><td>Fisher Scientific</td><td>12-545-100</td><td></td></tr><tr><td>Disposable round bottom flow cytometry tube</td><td>VWR</td><td>VWRU47729-566</td><td></td></tr><tr><td>DMSO</td><td>Sigma-Aldrich</td><td>472301-100ML</td><td></td></tr><tr><td>DPBS</td><td>VWR Life Science</td><td>SH30028.02</td><td></td></tr><tr><td>Glucose monohydrate</td><td>Sigma-Aldrich</td><td>Y0001745</td><td></td></tr><tr><td>HEPES Buffer (1 M)</td><td>Fisher Scientific</td><td>50-751-7290</td><td>Store at 4 &amp;deg;C</td></tr><tr><td>Ionomycin calcium salt</td><td>EMD Milipore Corp.</td><td>407952-1MG</td><td>Dissolve in DMSO to reach 2 mM. Store at -20 &amp;deg;C</td></tr><tr><td>KCl</td><td>Fisher Scientific</td><td>P330-500</td><td></td></tr><tr><td>MgSO&lt;sub&gt;4&lt;/sub&gt;</td><td>Fisher Scientific</td><td>M65-500</td><td></td></tr><tr><td>Microcentrifuge tube</td><td>Fisher Scientific</td><td>02-681-5</td><td></td></tr><tr><td>NaCl</td><td>Fisher Scientific</td><td>S271-500</td><td></td></tr><tr><td>Plain glass microscope slides</td><td>Fisher Scientific</td><td>12-544-4</td><td></td></tr><tr><td>Synergy HFM microplate reader</td><td>BioTek</td><td></td><td></td></tr><tr><td>Whole blood in ACD</td><td>Zen-Bio</td><td></td><td>Store at 4 &amp;deg;C and warm to 37 &amp;deg;C prior to use</td></tr></tbody></table>

induction of eryptosis in red blood cells using a calcium ionophore

Eryptosis, erythrocyte programmed cell death, occurs in a number of hematological diseases and during injury to erythrocytes. A hallmark of eryptotic cells is the loss of compositional asymmetry of the cell membrane, leading to the translocation of phosphatidylserine to the membrane outer leaflet. This process is triggered by increased intracellular concentration of Ca2+, which activates scramblase, an enzyme that facilitates bidirectional movement of phospholipids between membrane leaflets. Given the importance of eryptosis in various diseased conditions, there have been efforts to induce eryptosis in vitro. Such efforts have generally relied on the calcium ionophore, ionomycin, to enhance intracellular Ca2+ concentration and induce eryptosis. However, many discrepancies have been reported in the literature regarding the procedure for inducing eryptosis using ionomycin. Herein, we report a step-by-step protocol for ionomycin-induced eryptosis in human erythrocytes. We focus on important steps in the procedure including the ionophore concentration, incubation time, and glucose depletion, and provide representative result. This protocol can be used to reproducibly induce eryptosis in the laboratory.

Optimization of ionomycin concentration
While ionomycin is required to induce eryptosis, increased ionomycin concentrations can lead to hemolysis (i.e. lysis of erythrocytes and release of hemoglobin), which needs to be avoided. Treatment of erythrocytes with 1 &#181;M ionomycin in Ringer solution for 2 h is enough to induce eryptosis, as evidenced by successful labeling with annexin-V Alexa Flour 488 conjugate ...

Watch this Scientific Journal Video about Induction of Eryptosis in Red Blood Cells Using a Calcium Ionophore at JoVE.com

Induction of Eryptosis in Red Blood Cells Using a Calcium Ionophore

A protocol for the induction of eryptosis, programmed cell death in erythrocytes, using the calcium ionophore, ionomycin, is provided. Successful eryptosis is evaluated by monitoring the localization phosphatidylserine in the membrane outer leaflet. Factors affecting the success of the protocol have been examined and optimal conditions provided.

Eryptosis, erythrocyte programmed cell death, occurs in a number of hematological diseases and during injury to erythrocytes. A hallmark of eryptotic ...

induction-of-eryptosis-in-red-blood-cells-using-a-calcium-ionophore

Research

JoVE Journal

Biochemistry

8.7K Views.  Ohio University. Er zijn veel inspanningen geweest om eryptose te induceren met behulp van ionomycine. De exacte procedure voor dit proces wordt echter niet duidelijk beschreven in de literatuur. Wij bieden een stapsgewijze procedure om eryptose te induceren met behulp van ionomycine.Het belangrijkste voordeel van deze procedure is het op een betrouwbare en reproduceerbare manier induceren van eryptose bij menselijke erytrocyten. Om te beginnen, voeg 500 microliters koud vol bloed in zuurcitraat dextro...

Video: Induction of Eryptosis in Red Blood Cells Using a Calcium Ionophore

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

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

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

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

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

Biology

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

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

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

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

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

Cytosolic Calcium Measurements in Renal Epithelial Cells by Flow Cytometry

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

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

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

Functional Characterization of Endogenously Expressed Human RYR1 Variants

More than 700 variants in the RYR1 gene have been identified in patients with different neuromuscular disorders including malignant hyperthermia susceptibility, core myopathies and centronuclear myopathy. Because of the diverse phenotypes linked to RYR1 mutations it is fundamental to characterize their functional effects to classify variants carried by patients for future therapeutic interventions and identify non-pathogenic variants. Many laboratories have been interested in developing methods to functionally characterize RYR1 mutations expressed in patients&#39; cells. This approach has numerous advantages, including: mutations are endogenously expressed, RyR1 is not over-expressed, use of heterologous RyR1 expressing cells is avoided. However, since patients may present mutations in different genes aside RYR1, it is important to compare results from biological material from individuals harboring the same mutation, with different genetic backgrounds. The present manuscript describes methods developed to study the functional effects of endogenously expressed RYR1 variants in: (a) Epstein Barr virus immortalized human B-lymphocytes and (b) satellite cells derived from muscle biopsies and differentiated into myotubes. Changes in the intracellular calcium concentration triggered by the addition of a pharmacological RyR1 activators are then monitored. The selected cell type is loaded with a ratiometric fluorescent calcium indicator and intracellular [Ca2+] changes are monitored either at the single cell level by fluorescence microscopy or in cell populations using a spectrofluorometer. The resting [Ca2+], agonist dose response curves are then compared between cells from healthy controls and patients harboring RYR1 variants leading to insight into the functional effect of a given variant.

Here methods used to study the functional effect of RYR1 mutations endogenously expressed in Epstein Barr Virus immortalized human B-lymphocytes and muscle biopsy derived satellite cells differentiated into myotubes are described.

More than 700 variants in the RYR1 gene have been identified in patients with different neuromuscular disorders including malignant hyperthermia ...

Medicine

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 ...

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 ...

Neuroscience

Hemolysis Assay: An In Vitro Method to Measure Calcium Ionophore-Induced Lysis in Human Erythrocytes

Source: Bigdelou, P. et al., <a href="https://www.jove.com/v/60659">Induction of Eryptosis in Red Blood Cells Using a Calcium Ionophore.</a> J. Vis. Exp. (2020).
This video describes an in vitro assay to determine calcium-ionophore-induced hemolysis or lysis of red blood cells in a human sample. The assay measures the percentage of hemolysis by quantifying hemoglobin released from ruptured erythrocytes.

Source: Bigdelou, P. et al., Induction of Eryptosis in Red Blood Cells Using a Calcium Ionophore. J. Vis. Exp. (2020).
This video describes an in vitro ...

JoVE Encyclopedia of Experiments

Biological Techniques

Assay Techniques

Cell-based assays

Annexin V Binding Assay: A Fluorescence-Based Technique to Identify Apoptotic Erythrocytes via Phosphatidylserine Labeling

Source: Bigdelou, P., et al. <a href="https://www.jove.com/v/60659">Induction of Eryptosis in Red Blood Cells Using a Calcium Ionophore.</a> J. Vis. Exp. (2020)
In this video, we demonstrate the Annexin V binding assay to measure the degree of eryptosis, erythrocyte programmed cell death, using fluorescent Annexin V staining. Apoptotic erythrocytes demonstrate phosphatidylserine translocation from the inner to the outer leaflet of the cell membrane, causing the fluorophore-tagged Annexin V to bind to the translocated phosphatidylserine, thereby emitting a fluorescent signal.

Source: Bigdelou, P., et al. Induction of Eryptosis in Red Blood Cells Using a Calcium Ionophore. J. Vis. Exp. (2020)
In this video, we demonstrate the ...

Monitoring B Cell Activation with Antigenic Liposomes using a Calcium-Flux Assay

Source: Bednar, K. J.&#160;et al.,&#160;<a href="https://app.jove.com/v/58285">Antigenic Liposomes for Generation of Disease-specific Antibodies.</a>&#160;J. Vis. Exp.&#160;(2018)
This video showcases the assessment of B cell activation through the utilization of antigenic liposomes. B cells treated with calcium-sensitive Indo-1 dye are distinguished using flow cytometry, and their violet-to-blue fluorescence ratio is analyzed. As the cells encounter antigenic liposomes, it triggers B cell activation through calcium influx, progressively enhancing the violet-to-blue fluorescence signal ratio over time.

1. Conjugation of Protein Antigen to PEGylated Lipid

	Add 3 g of cross-linked dextran gel beads with a fractionation range of 1,500&#8211;30,000 Daltons ...

Immunology

Immunodiagnostics

Specialized Assays

An Imaging Technique to Study the Calcium Response in Bacteria-Infected Cells

Place the coverslip containing the Fluo-4 cells in an imaging or glass-bottom chamber. Use EM buffer to wash the sample three times to remove compounds potentially resulting from cell lysis. Then, add one milliliter of EM buffer. Place the chamber on an inverted fluorescence microscope stage, heated at 33 degrees Celsius. Select a microscopy field, and set up acquisition parameters, including exposure time and binning, if necessary, to optimize the Fluo-4 fluorescent signal.
To add bacteria to the sample, carefully remove 500 microliters of EM buffer from the chamber, and add 500 microliters of the bacterial suspension to obtain a final OD600 of 0.05. Immediately perform an acquisition, or wait 10 minutes to allow the bacteria to sediment onto the cells. At the end of the acquisition stream, acquire a phase contrast image of the selected field to visualize the bacteria contacting the cells and the membrane ruffles associated with bacterial invasion sites.
Repeat the acquisition procedure at intervals that are amenable to the duration of the image acquisitions, file savings, and selection of a new field to cover the whole process. At the end of the acquisition procedure, add to the sample, two micromolar of the calcium ionophore ionomycin to determine the maximal amplitude of the calcium signals. Acquire images every 3 to 5 seconds until the signal stabilizes, usually for less than 10 minutes.
Follow by adding the calcium chelator EGTA to a final concentration of 10 millimolar to determine the fluorescent signal in the absence of calcium. Acquire images every 3 to 5 seconds until signal stabilization, usually for less than 10 minutes. Carry out image analysis according to the text protocol.

Induction of Eryptosis in Red Blood Cells Using a Calcium Ionophore

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