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 induces cellular stress and activates protein kinase C (PKC), resulting in the activation of calcium and potassium channels. This results in an increase in the entry of Ca2+ in the cytosolic space30,31,34 and ultimately activates the scramblase16, which increases eryptosis. Activation of potassium channel also results in potassium chloride leakage out of the cell, which leads to erythrocyte shrinkage35.
The procedure outlined above needs to be performed with specific attention to hemolysis. It is important to use an optimized ionophore concentration, which is high enough to induce eryptosis, and low enough to prevent hemolysis. Similarly, incubating erythrocytes with ionomycin for a short period of time results in low eryptosis while very long incubation may lead to cell membrane disruption and hemolysis. It should also be noted that while the presented protocol is highly reliable when performed on the same erythrocyte sample, cells from different individuals respond differently to ionomycin and there might be inter-subject variability between different samples.
Particular attention should be paid to data analysis from flow cytometry. The percentage eryptosis obtained from the flow cytometer indicates the percentage of cell population with PS on their outer leaflet. However, cells with different intensities of annexin-V binding cannot be distinguished based on this number. Annexin-V binds to the PS exposed on cell surface, with a very high affinity and high specificity to PS36,37,38. However, as shown in the microscopy images in this report, different cells show differences in annexin-V binding intensity. The cells with low PS on their membranes have low fluorescence intensities, whereas higher PS occupancy on cell membrane results in higher fluorescence intensities.
The protocol presented in this paper can be modified by increasing the extracellular Ca2+ concentration. In this protocol, ionomycin was used to induce eryptosis in the presence of 1 mM CaCl2; higher Ca2+ concentrations might lead to enhanced intracellular calcium levels and may induce more eryptosis. In addition, different calcium ionophores, such as selectophore and calcimycin, might have different ability to enhance the intracellular concentration of Ca2+, compared to ionomycin, and could result in different eryptosis values. However, consistent eryptosis of erythrocytes can be achieved using ionomycin with the outlined protocol and can be used in the laboratory to examine the molecular mechanisms of eryptosis, mimic diseased conditions39,40 in vitro, and screen potential therapeutics that inhibit eryptosis, among other applications.

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 border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<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 &#176;C</td>
		</tr>
		<tr>
			<td>BD FACSAria II flow cytometer</td>
			<td>BD Biosciences</td>
			<td>643177</td>
			<td></td>
		</tr>
		<tr>
			<td>CaCl2</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 &#176;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 &#176;C</td>
		</tr>
		<tr>
			<td>KCl</td>
			<td>Fisher Scientific</td>
			<td>P330-500</td>
			<td></td>
		</tr>
		<tr>
			<td>MgSO4</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 &#176;C and warm to 37 &#176;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 and quantification by FACS analysis (Figure 1A). Higher concentrations of ionomycin (5 and 10 &#181;M) result in a slight increase in eryptosis (Figure 1A-D). However, such concentrations also enhance hemolysis (Figure 1E), which is not desired. In order to stay below 5% hemolysis, 1 &#181;M ionomycin should be used.
Treatment time with ionomycin
Incubation of erythrocytes with ionomycin in Ringer solution for as little as 30 min is enough to induce eryptosis (Figure 2A). Increased incubation time increases the level of eryptosis, as measured by the annexin V-binding assay, for up to 2 h (Figure 2B,C). However, further incubation time results in a slight decrease in the level of eryptosis (Figure 2D). Maximum eryptosis was obtained after 2 h of treatment with 1 &#181;M ionomycin, and for all other treatment times, lower eryptosis was obtained (Figure 2E). Representative flow cytometry histograms are presented in Figure 2A-D. In addition, average percentage eryptosis and hemolysis, for various treatment times with 1 &#181;M ionomycin, are presented in Figure 2E and Figure 2F, respectively. The higher value of hemolysis after 180 min explains the reduction in eryptosis after the same amount of incubation (Figure 2E) as less viable cells exist upon 180 min of treatment with ionomycin.
Moreover, cells were treated with low concentrations of ionomycin including 0, 0.25, 0.5, and 1 &#181;M for longer treatment times including 6 and 12 h, and eryptosis was measured (Figure 3). Cells treated with ionomycin concentrations of lower that 1 &#181;M for 6 and 12 h show lower eryptosis compared to the cells treated with 1 &#181;M ionomycin (Figure 3). Since decreasing the concentration and increasing the exposure time did not enhance eryptosis, 1 &#181;M was used to trigger eryptosis.
Eryptosis is dependent on incubation time and extracellular glucose concentration
Extracellular glucose concentration affects the outcome of the process. Higher eryptosis values are observed when erythrocytes are pre-incubated in glucose-free Ringer solution compared to glucose-containing Ringer solution prior to incubation with 1 &#181;M ionomycin for 2 h. The highest eryptosis values are obtained after 7 days of pre-incubation in both solutions. However, eryptosis is higher after pre-incubation in glucose-free Ringer solution compared to normal Ringer solution, which contains 5 mM glucose (see Figure 4A for representative plots and Figure 4B for comparison of global means). In addition, forward scatter histograms indicate the effect of glucose depletion on erythrocyte shrinkage (Figure 5A-D). Forward scatter is a measure for cell size based on the light refraction, and the level of light scattered is directly proportional to the size of cells33. The cells incubated in glucose-free Ringer solution show less forward scatter compared to the cells incubated in glucose-containing buffer (Figure 5E), indicating cell shrinkage in the glucose-free environment.
In addition to flow cytometry measurements, cells were observed under a confocal fluorescence microscope to confirm eryptosis. Erythrocytes with no treatment (Figure 6A) and with ionomycin treatment (Figure 6B) were labeled with annexin-V Alexa Flour 488 conjugate and observed under microscope. Treated cells showed a bright fluorescence signal (Figure 6B) due to the binding of annexin-V to PS in the outer leaflet. In contrast, cells with no treatment showed a very weak fluorescence signal (Figure 6A) indicating very low eryptosis. Further example images of eryptotic erythrocytes labeled with annexin-V with high fluorescence signal are shown in Figure 6C.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/60659/60659fig1.jpg" /> 
Figure 1: Representative graphs of the effect of various ionomycin concentrations on eryptosis and hemolysis. Flow cytometry histograms of erythrocytes treated with (A) 1 &#181;M, (B) 5 &#181;M, and (C) 10 &#181;M ionomycin (gray) at 37 &#176;C at 0.4% hematocrit in Ringer solution for 2 h. Black line indicates non-treated cells. Percentage of eryptosis is indicated in each figure. Phosphatidylserine exposure was measured using annexin-V binding. (D) Arithmetic means &#177; SD (n = 3) of the percentage eryptosis of cells treated with different concentrations of ionomycin after 2 h treatment, and (E) arithmetic means &#177; SD (n = 3) of hemolysis of erythrocytes by different concentrations of ionomycin under same conditions. <a href="https://www.jove.com/files/ftp_upload/60659/60659fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/60659/60659fig2a.jpg" /> 
Figure 2: Representative figures on the effect of various ionomycin treatment times on eryptosis. Flow cytometry histograms of erythrocytes treated with 1 &#181;M ionomycin (gray) at 37 &#176;C for (A) 30 min, (B) 60 min, (C) 120 min, and (D) 180 min at 0.4% hematocrit in Ringer solution. Black line indicates non-treated cells. Percentage of eryptosis is indicated in each figure. Phosphatidylserine exposure was measured through annexin-V binding. (E) Arithmetic means &#177; SD (n = 3) of percentage eryptosis of cells treated with 1 &#181;M ionomycin for different times. The highest eryptosis was obtained after 120 min treatment. (F) Arithmetic means &#177; SD (n = 3) of percentage hemolysis of cells treated with 1 &#181;M ionomycin for different times. For statistical analysis, one-way non-parametric ANOVA with Kruskal-Wallis test was performed, and eryptosis after 120 min treatment was significantly higher than control as indicated in panel E. * is for p &#60; 0.05. <a href="https://www.jove.com/files/ftp_upload/60659/60659fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/60659/60659fig3a.jpg" /> 
Figure 3: Effect of various ionomycin concentrations and treatment times on eryptosis. Arithmetic means &#177; SD (n = 3) of the percentage eryptosis of cells treated with different concentrations of ionomycin is shown after various treatment times. The cells were treated with low concentrations of ionomycin including 0, 0.25, 0.5, and 1 &#181;M for longer exposure (6 h and 12 h). Higher concentrations and longer treatments resulted in higher eryptosis values. <a href="https://www.jove.com/files/ftp_upload/60659/60659fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/60659/60659fig4a.jpg" /> 
Figure 4: Effect of energy depletion on eryptosis. (A) Flow cytometry histogram for erythrocytes treated with 1 &#181;M ionomycin (gray) at 37 &#176;C for 2 h at 0.4% hematocrit, after pre-incubation in glucose-free Ringer solution (top figures) and Ringer solution (bottom figures) from 1 to 7 days, reveals that energy depletion facilitates eryptosis. Black line indicates non-treated cells. Percentages of eryptosis are indicated in the graphs for each day. (B) Arithmetic means &#177; SD (n = 3) of the percentage eryptosis of erythrocytes treated with 1 &#181;M ionomycin at 37 &#176;C for 2 h at 0.4% hematocrit, after pre-incubation in Ringer solution (black bars) and glucose-free Ringer solution (white bars) from 1 to 7 days. <a href="https://www.jove.com/files/ftp_upload/60659/60659fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/60659/60659fig5a.jpg" /> 
Figure 5: Effect of energy depletion on cell size. Forward scatter histogram for erythrocytes treated with 1 &#181;M ionomycin at 37 &#176;C for 2 h at 0.4% hematocrit, after pre-incubation in glucose-free Ringer solution (gray) and Ringer solution (black line) for (A) 1 day, (B) 3 days, (C) 5 days, and (D) 7 days. The forward scatter histogram over time indicates erythrocyte shrinkage in glucose-free buffer. (E) Arithmetic means &#177; SD (n = 3) of forward scatter intensities of erythrocytes treated with 1 &#181;M ionomycin at 37 &#176;C for 2 h at 0.4% hematocrit, after pre-incubation in Ringer solution (black bars) and glucose-free Ringer solution (white bars) from 1 to 7 days. <a href="https://www.jove.com/files/ftp_upload/60659/60659fig5large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 6" class="xfigimg" src="/files/ftp_upload/60659/60659fig6a.jpg" /> 
Figure 6: Confocal fluorescence microscopy images of erythrocytes treated with (A) 0 &#181;M, (B) and (C) 1 &#181;M ionomycin at 37 &#176;C for 2 h at 0.4% hematocrit. 40x objective magnification was used for images in panels A and B, and 100x objective magnification was used to take images for panel C. PS in healthy erythrocytes is located on the inner leaflet of the cell membrane, therefore there is no fluorescence signal in panel A. In panels B and C erythrocytes have been induced for eryptosis and there is a bright fluorescence signal resulting from the binding of annexin-V to PS translocated to the outer leaflet of the cell membrane. <a href="https://www.jove.com/files/ftp_upload/60659/60659fig6large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Cell density /hematocrit</td>
			<td>Ionomycin concentration</td>
			<td>Buffer</td>
			<td>Pre-incubation</td>
			<td>Treatment time with ionomycin</td>
			<td>Detection method</td>
			<td>Reference</td>
		</tr>
		<tr>
			<td>1.65 x 108 cells/mL</td>
			<td>0.3 mM</td>
			<td>Buffer A*</td>
			<td>36 h in buffer A</td>
			<td>1 h</td>
			<td>Annexin V</td>
			<td>12</td>
		</tr>
		<tr>
			<td>0.40%</td>
			<td>1 mM</td>
			<td>Ringer solution</td>
			<td>48 h in Ringer</td>
			<td>1 h</td>
			<td>Annexin V</td>
			<td>17</td>
		</tr>
		<tr>
			<td>50%</td>
			<td>10 mM</td>
			<td>Buffer B**</td>
			<td>-</td>
			<td>3 h</td>
			<td>Merocyanine 540</td>
			<td>18</td>
		</tr>
		<tr>
			<td>0.40%</td>
			<td>1 mM</td>
			<td>Ringer solution</td>
			<td>48 h in Ringer</td>
			<td>1 h</td>
			<td>Annexin V</td>
			<td>19</td>
		</tr>
		<tr>
			<td>0.40%</td>
			<td>1 mM</td>
			<td>Ringer solution</td>
			<td>48 h in Ringer</td>
			<td>1 h</td>
			<td>Annexin V</td>
			<td>20</td>
		</tr>
		<tr>
			<td>2%</td>
			<td>1 mM</td>
			<td>Ringer solution</td>
			<td>-</td>
			<td>4 h</td>
			<td>Annexin V</td>
			<td>21</td>
		</tr>
		<tr>
			<td>0.40%</td>
			<td>1 mM</td>
			<td>Ringer solution</td>
			<td>-</td>
			<td>0.5 h</td>
			<td>Annexin V</td>
			<td>22</td>
		</tr>
		<tr>
			<td>10%</td>
			<td>1 mM</td>
			<td>Ringer solution</td>
			<td>-</td>
			<td>3 h</td>
			<td>Annexin V</td>
			<td>23</td>
		</tr>
		<tr>
			<td>0.40%</td>
			<td>10 mM</td>
			<td>Ringer solution</td>
			<td>-</td>
			<td>0.5 h</td>
			<td>Annexin V</td>
			<td>24</td>
		</tr>
		<tr>
			<td>0.40%</td>
			<td>1 mM</td>
			<td>Ringer solution</td>
			<td>48 h in Ringer</td>
			<td>0.5 h</td>
			<td>Annexin V</td>
			<td>25</td>
		</tr>
		<tr>
			<td>2 x 106 cells/mL</td>
			<td>1 mM</td>
			<td>HEPES-buffered saline (HBS)</td>
			<td>-</td>
			<td>0.5 h</td>
			<td>Annexin V</td>
			<td>26</td>
		</tr>
		<tr>
			<td colspan="7">*Buffer A: 10 mM HEPES, 150 mM NaCl, 5 mM KCl, 1 mM MgCl2&#183;6H2O, 10 mM glucose, and 1.8 mM CaCl2&#183;2H2O</td>
		</tr>
		<tr>
			<td colspan="7">**Buffer B: 5 mM Tris, 100 mM KC1, 60 mM NaCl, and 10 mM glucose</td>
		</tr>
	</tbody>
</table>
Table 1: Various protocols used in the literature to induce eryptosis using ionomycin.

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

Nanyang Technological University

Research

JoVE Journal

Biochemistry

8.6K Views.  Ohio University. 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.

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

Analyses of Mitochondrial Calcium Influx in Isolated Mitochondria and Cultured Cells

Ca2+ handling by mitochondria is a critical function regulating both physiological and pathophysiological processes in a broad spectrum of cells. The ability to accurately measure the influx and efflux of Ca2+ from mitochondria is important for determining the role of mitochondrial Ca2+ handling in these processes. In this report, we present two methods for the measurement of mitochondrial Ca2+ handling in both isolated mitochondria and cultured cells. We first detail a plate reader-based platform for measuring mitochondrial Ca2+ uptake using the Ca2+ sensitive dye calcium green-5N. The plate reader-based format circumvents the need for specialized equipment, and the calcium green-5N dye is ideally suited for measuring Ca2+ from isolated tissue mitochondria. For our application, we describe the measurement of mitochondrial Ca2+ uptake in mitochondria isolated from mouse heart tissue; however, this procedure can be applied to measure mitochondrial Ca2+ uptake in mitochondria isolated from other tissues such as liver, skeletal muscle, and brain. Secondly, we describe a confocal microscopy-based assay for measurement of mitochondrial Ca2+ in permeabilized cells using the Ca2+ sensitive dye Rhod-2/AM and imaging using 2-dimensional laser-scanning microscopy. This permeabilization protocol eliminates cytosolic dye contamination, allowing for specific recording of changes in mitochondrial Ca2+. Moreover, laser-scanning microscopy allows for high frame rates to capture rapid changes in mitochondrial Ca2+ in response to various drugs or reagents applied in the external solution. This protocol can be applied to measure mitochondrial Ca2+ uptake in many cell types including primary cells such as cardiac myocytes and neurons, and immortalized cell lines.

Here, we present two protocols for the measurement of mitochondrial Ca2+ influx in isolated mitochondria and cultured cells. For isolated mitochondria, we detail a plate reader-based Ca2+ import assay using the Ca2+ sensitive dye calcium green-5N. For cultured cells, we describe a confocal microscopy method using the Ca2+ dye Rhod-2/AM.

Ca2+ handling by mitochondria is a critical function regulating both physiological and pathophysiological processes in a broad spectrum of cells. The ...

Dissipative Microgravimetry to Study the Binding Dynamics of the Phospholipid Binding Protein Annexin A2 to Solid-supported Lipid Bilayers Using a Quartz Resonator

The dissipative quartz crystal microbalance technique is a simple and label-free approach to measure simultaneously the mass uptake and viscoelastic properties of the absorbed/immobilized mass on sensor surfaces, allowing the measurements of the interaction of proteins with solid-supported surfaces, such as lipid bilayers, in real-time and with a high sensitivity. Annexins are a highly conserved group of phospholipid-binding proteins that interact reversibly with the negatively charged headgroups via the coordination of calcium ions. Here, we describe a protocol that was employed to quantitatively analyze the binding of annexin A2 (AnxA2) to planar lipid bilayers prepared on the surface of a quartz sensor. This protocol is optimized to obtain robust and reproducible data and includes a detailed step-by-step description. The method can be applied to other membrane-binding proteins and bilayer compositions.

Here, we present an experimental protocol that can be employed to determine the binding affinities and mode of interaction of label-free phospholipid-binding protein annexin A2 with immobilized solid-supported bilayers (SLB) by simultaneously measuring the mass uptake and the viscoelastic properties of the protein annexin A2.

The dissipative quartz crystal microbalance technique is a simple and label-free approach to measure simultaneously the mass uptake and viscoelastic ...

In Vivo Calcium Imaging in C. elegans Body Wall Muscles

The model organism C. elegans provides an excellent system to perform in vivo calcium imaging. Its transparent body and genetic manipulability allow for the targeted expression of genetically encoded calcium sensors. This protocol outlines the use of these sensors for the in vivo imaging of calcium dynamics in targeted cells, specifically the body wall muscles of the worms. By utilizing the co-expression of presynaptic channelrhodopsin, stimulation of acetylcholine release from excitatory motor neurons can be induced using blue light pulses, resulting in muscle depolarization and reproducible changes in cytoplasmic calcium levels. Two worm immobilization techniques are discussed with varying levels of difficulty. Comparison of these techniques demonstrates that both approaches preserve the physiology of the neuromuscular junction and allow for the reproducible quantification of calcium transients. By pairing optogenetics and functional calcium imaging, changes in postsynaptic calcium handling and homeostasis can be evaluated in a variety of mutant backgrounds. Data presented validates both immobilization techniques and specifically examines the roles of the C. elegans sarco(endo)plasmic reticular calcium ATPase and the calcium-activated BK potassium channel in the body wall muscle calcium regulation.

In Vivo Calcium Imaging in C. elegans Body Wall Muscles

This method provides a way to couple optogenetics and genetically encoded calcium sensors to image baseline cytosolic calcium levels and changes in evoked calcium transients in the body wall muscles of the model organism C. elegans.

The model organism C. elegans provides an excellent system to perform in vivo calcium imaging. Its transparent body and genetic manipulability allow for ...

Translating Ribosome Affinity Purification (TRAP) for RNA Isolation from Endothelial Cells In Vivo

Many studies have been limited to using in vitro cellular assays and whole tissues or isolating of specific cell types from animals for in vitro analysis of transcriptome and gene expression by qPCR and RNA sequencing. Comprehensive transcriptome and gene expression analysis of specific cell types in complex tissues and organs will be critical to understand cellular and molecular mechanisms by which genes are regulated and their association with tissue homeostasis and organ functions. In this article, we demonstrate the methodology for isolation of ribosome-bound RNA directly in vivo in the vascular endothelia of animal lungs as an example. The specific materials and procedures for tissue processing and RNA purification will be described, including the assessment of RNA quality and yield as well as real time qPCR for arteriogenic gene assays. This approach, known as translating ribosome affinity purification (TRAP) technique, can be utilized for characterization of gene expression and transcriptome analysis of certain cell types directly in vivo in any specific type in complex tissues.

Translating Ribosome Affinity Purification TRAP for RNA Isolation from Endothelial Cells In Vivo

We present an approach to purify ribosome-bound mRNA from vascular endothelial cells (ECs) directly in mouse brain, lung and heart tissues via EC-specific genetic tag of enhanced green fluorescence protein (EGFP)in ribosomes in combination with RNA purification.

Many studies have been limited to using in vitro cellular assays and whole tissues or isolating of specific cell types from animals for in vitro analysis ...

SUMO-Binding Entities (SUBEs) as Tools for the Enrichment, Isolation, Identification, and Characterization of the SUMO Proteome in Liver Cancer

Post-translational modification is a key mechanism regulating protein homeostasis and function in eukaryotic cells. Among all ubiquitin-like proteins in liver cancer, the modification by SUMO (Small Ubiquitin MOdifier) has been given the most attention. Isolation of endogenous SUMOylated proteins in vivo is challenging due to the presence of active SUMO-specific proteases. Initial studies of SUMOylation in vivo were based on the molecular detection of specific SUMOylated proteins (e.g., by western blot). However, in many cases, antibodies, generally made with non-modified recombinant protein, did not immunoprecipitate SUMOylated forms of the protein of interest.&#160;Nickel chromatography has been the other approach to study SUMOylation by capturing histidine-tagged versions of SUMO molecules. This approach is mainly used in cells stably expressing or transiently transfected with His-SUMO molecules. To overcome these limitations, SUMO-binding entities (SUBEs) were developed to isolate endogenous SUMOylated proteins. Herein, we describe all the steps required for the enrichment, isolation, and identification of SUMOylated substrates from human hepatoma cells and hepatic tissues from a liver cancer mouse model by using SUBEs. Firstly, we describe methods involved in the preparation and lysis of the human hepatoma cells and liver tumor tissue samples. Then, a thorough explanation of the preparation of SUBEs and controls is detailed along with the protocol for the protein pull-down assays. Finally, some examples are provided regarding the options available for the identification and characterization of the SUMOylated proteome, namely the use of western-blot analysis for the detection of a specific SUMOylated substrate from liver tumors or the use of proteomics by mass spectrometry for high-throughput characterization of the SUMOylated proteome and interactome in hepatoma cells.

SUMO-Binding Entities SUBEs as Tools for the Enrichment, Isolation, Identification, and Characterization of the SUMO Proteome in Liver Cancer

Here, we present a protocol to enrich, isolate, identify, and characterize proteins modified by SUMO in vivo both from human hepatoma cells and liver tumors obtained from mouse models of hepatocellular carcinoma by using SUMO-binding entities (SUBEs).

Post-translational modification is a key mechanism regulating protein homeostasis and function in eukaryotic cells. Among all ubiquitin-like proteins in ...

Ultra-High-Speed Western Blot using Immunoreaction Enhancing Technology

A western blot (also known as an immunoblot) is a canonical method for biomedical research. It is commonly used to determine the relative size and abundance of specific proteins as well as post-translational protein modifications. This technique has a rich history and remains in widespread use due to its simplicity. However, the western blotting procedure famously takes hours, even days, to complete, with a critical bottleneck being the long incubation times that limit its throughput. These incubation steps are required due to the slow diffusion of antibodies from the bulk solution to the immobilized antigens on the membrane: the antibody concentration near the membrane is much lower than the bulk concentration. Here, we present an innovation that dramatically reduces these incubation intervals by improving antigen binding via cyclic draining and replenishing (CDR) of the antibody solution. We also utilized an immunoreaction enhancing technology to preserve the sensitivity of the assay. A combination of the CDR method with a commercial immunoreaction enhancing agent boosted the output signal and substantially reduced the antibody incubation time. The resulting ultra-high-speed western blot can be accomplished in 20 minutes without any loss in sensitivity. This method can be applied to western blots using both chemiluminescent and fluorescent detection. This simple protocol allows researchers to better explore the analysis of protein expression in many samples.

An ultra-high-speed western blotting technique is developed by improving the kinetics of antigen-antibody binding through cyclic draining and replenishing (CDR) technology in conjunction with an immunoreaction enhancing agent.

A western blot (also known as an immunoblot) is a canonical method for biomedical research. It is commonly used to determine the relative size and ...

Rapid Determination of Antibody-Antigen Affinity by Mass Photometry

Measurements of the specificity and affinity of antigen-antibody interactions are critically important for medical and research applications. In this protocol, we describe the implementation of a new single-molecule technique, mass photometry (MP), for this purpose. MP is a label- and immobilization-free technique that detects and quantifies molecular masses and populations of antibodies and antigen-antibody complexes on a single-molecule level. MP analyzes the antigen-antibody sample within minutes, allowing for the precise determination of the binding affinity and simultaneously providing information on the stoichiometry and the oligomeric state of the proteins. This is a simple and straightforward technique that requires only picomole quantities of protein and no expensive consumables. The same procedure can be used to study protein-protein binding for proteins with a molecular mass larger than 50 kDa. For multivalent protein interactions, the affinities of multiple binding sites can be obtained in a single measurement. However, the single-molecule mode of measurement and the lack of labeling imposes some experimental limitations. This method gives the best results when applied to measurements of sub-micromolar interaction affinities, antigens with a molecular mass of 20 kDa or larger, and relatively pure protein samples. We also describe the procedure for performing the required fitting and calculation steps using basic data analysis software.

We describe a single-molecule approach to antigen-antibody affinity measurements using mass photometry (MP). The MP-based protocol is fast, accurate, uses a very small amount of material, and does not require protein modification.

Measurements of the specificity and affinity of antigen-antibody interactions are critically important for medical and research applications. In this ...

Myosin-Specific Adaptations of In vitro Fluorescence Microscopy-Based Motility Assays

Myosin proteins bind and interact with filamentous actin (F-actin) and are found in organisms across the phylogenetic tree. Their structure and enzymatic properties are adapted for the particular function they execute in cells. Myosin 5a processively walks on F-actin to transport melanosomes and vesicles in cells. Conversely, nonmuscle myosin 2b operates as a bipolar filament containing approximately 30 molecules. It moves F-actin of opposite polarity toward the center of the filament, where the myosin molecules work asynchronously to bind actin, impart a power stroke, and dissociate before repeating the cycle. Nonmuscle myosin 2b, along with its other nonmuscle myosin 2 isoforms, has roles that include&#160;cell adhesion, cytokinesis, and tension maintenance. The mechanochemistry of myosins can be studied by performing in vitro motility assays using purified proteins. In the gliding actin filament assay, the myosins are bound to a microscope coverslip surface and translocate fluorescently labeled F-actin, which can be tracked. In the single molecule/ensemble motility assay, however, F-actin is bound to a coverslip and the movement of fluorescently labeled myosin molecules on the F-actin is observed. In this report, the purification of recombinant myosin 5a from Sf9 cells using affinity chromatography is outlined. Following this, we outline two fluorescence microscopy-based assays: the gliding actin filament assay and the inverted motility assay. From these assays, parameters such as actin translocation velocities and single molecule run lengths and velocities can be extracted using the image analysis software. These techniques can also be applied to study the movement of single filaments of the nonmuscle myosin 2 isoforms, discussed herein in the context of nonmuscle myosin 2b. This workflow represents a protocol and a set of quantitative tools that can be used to study the single molecule and ensemble dynamics of nonmuscle myosins.

Presented here is a procedure to express and purify myosin 5a followed by a discussion of its characterization, using both ensemble and single molecule in vitro fluorescence microscopy-based assays, and how these methods can be modified for the characterization of nonmuscle myosin 2b.

Myosin proteins bind and interact with filamentous actin (F-actin) and are found in organisms across the phylogenetic tree. Their structure and enzymatic ...

Inner Mitochondrial Membrane Sensitivity to Na+ Reveals Partially Segmented Functional CoQ Pools

Ubiquinone (CoQ) pools in the inner mitochondrial membrane (IMM) are partially segmented to either complex I or FAD-dependent enzymes. Such subdivision can be easily assessed by a comparative assay using NADH or succinate as electron donors in frozen-thawed mitochondria, in which cytochrome c (cyt c) reduction is measured. The assay relies on the effect of Na+ on the IMM, decreasing its fluidity. Here, we present a protocol to measure NADH-cyt c oxidoreductase activity and succinate-cyt c oxidoreductase activities in the presence of NaCl or KCl. The reactions, which rely on the mixture of reagents in a cuvette in a stepwise manner, are measured spectrophotometrically during 4 min in the presence of Na+ or K+. The same mixture is performed in parallel in the presence of the specific enzyme inhibitors in order to subtract the unspecific change in absorbance. NADH-cyt c oxidoreductase activity does not decrease in the presence of any of these cations. However, succinate-cyt c oxidoreductase activity decreases in the presence of NaCl. This simple experiment highlights: 1) the effect of Na+ in decreasing IMM fluidity and CoQ transfer; 2) that supercomplex I+III2 protects ubiquinone (CoQ) transfer from being affected by lowering IMM fluidity; 3) that CoQ transfer between CI and CIII is functionally different from CoQ transfer between CII and CIII. These facts support the existence of functionally differentiated CoQ pools in the IMM and show that they can be regulated by the changing Na+ environment of mitochondria.

Inner Mitochondrial Membrane Sensitivity to Na+ Reveals Partially Segmented Functional CoQ Pools

This protocol describes a comparative assay, using mitochondrial complex activities CI+CIII and CII+CIII in the presence or absence of Na+, to study the existence of partially segmented functional CoQ pools.

Ubiquinone (CoQ) pools in the inner mitochondrial membrane (IMM) are partially segmented to either complex I or FAD-dependent enzymes. Such subdivision ...

A Semi-Automated and Reproducible Biological-Based Method to Quantify Calcium Deposition In Vitro

Vascular calcification involves a series of degenerative pathologies, including inflammation, changes to cellular phenotype, cell death, and the absence of calcification inhibitors, that concomitantly lead to a loss of vessel elasticity and function. Vascular calcification is an important contributor to morbidity and mortality in many pathologies, including chronic kidney disease, diabetes mellitus, and atherosclerosis. Current research models to study vascular calcification are limited and are only viable at the late stages of calcification development in vivo. In vitro tools for studying vascular calcification use end-point measurements, increasing the demands on biological material and risking the introduction of variability to research studies. We demonstrate the application of a novel fluorescently labeled probe that binds to in vitro calcification development on human vascular smooth muscle cells and determines the real-time development of in vitro calcification. In this protocol, we describe the application of our newly developed calcification assay, a novel tool in disease modeling that has potential translational applications. We envisage this assay to be relevant in a broader spectrum of mineral deposition research, including applications in bone, cartilage, or dental research.

A Semi-Automated and Reproducible Biological-Based Method to Quantify Calcium Deposition In Vitro

Cardiovascular disease is the leading cause of death worldwide. Vascular calcification contributes substantially to the burden of cardiovascular morbidity and mortality. This protocol describes a simple method to quantify vascular smooth muscle cell-mediated calcium precipitation in vitro by fluorescent imaging.

Vascular calcification involves a series of degenerative pathologies, including inflammation, changes to cellular phenotype, cell death, and the absence ...