The study was financially supported by the National Research Foundation (NRF) and the South African Medical Research Council (SAMRC). We would like to acknowledge the National Health Laboratory Service (NHLS) for purchase of the Guava Muse Cell Analyzer. All figures in this publication were created with Biorender.com.

NOTE: This protocol describes steps in the cell preparation, cell enumeration, cell staining, and analysis of actinomycin D-treated SiHa cells using flow cytometric commercial assays measured and analyzed on a benchtop flow cytometer (Figure 2).
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/62663/62663fig2v2.jpg" /> 
Figure 2: Workflow for the detection of biochemical apoptotic biomarkers by flow cytometry. Cells are cultured and treated as described in protocol step 1.1. (1) Cells are cultured, (2) enumerated and (3) stained with fluorescent reagents, and (4) analysed by a benchtop flow cytometer. Data are further gated, and statistically analyzed. Abbreviations: 7-AAD = 7-aminoactinomycin D; PBS = phosphate-buffered saline; PE = Phycoerythrin; TMRE = tetramethylrhodamine ethyl ester. <a href="https://www.jove.com/files/ftp_upload/62663/62663fig2v2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
1. Preparation of cell culture and treatments for flow cytometry
NOTE: Ensure aseptic technique is followed when handling cell cultures.
<ol>
	<li>Grow cell cultures in a humidified CO2 environment at 37 &#176;C with 5% CO2. Ensure that cell culture is approximately 70% confluent in the parent flask before passaging cells for experiments.</li>
	<li>Remove the medium from the flask, wash cells with 1x phosphate-buffered saline (PBS) and add 500 &#181;L of trypsin to detach the cells. After the cells begin to round and detach from the flask, sharply knock the base of the flask on the bench to detach the cells. Then neutralize the trypsin by adding approximately 5 mL of fresh culture medium supplemented with 10% fetal calf serum before counting the cells.&#160;</li>
	<li>Seed cells at 15,000 cells/mL in 3 mL of cell culture medium in a 6-well cell culture plate. Incubate the culture plates overnight at 37 &#176;C with 5% CO2 to allow the attachment of the cells to the bottom of the well.</li>
	<li>Treat the experimental cultures with 100 ng/mL actinomycin D and the medium and solvent controls with fresh culture medium and dimethyl sulfoxide (DMSO), respectively, for 24 h. Collect the spent medium in a 15 mL tube and wash cells with 1x PBS. Thereafter, add the 1x PBS wash to the tube, and then add 500 &#181;L of trypsin to the well. Next, neutralize the trypsin by adding 5 mL of fresh culture medium. 
	&#8203;NOTE: (1) Prolonged trypsin incubation or incomplete neutralization may digest the cells and compromise their cell membrane, which may skew the results. In addition, it may also change cell membrane asymmetry and, thus, the accessibility to phosphatidylserine. (2) Cells that have detached and floated to the surface can be included in the subsequent analysis by centrifuging the removed medium at 300 &#215; g for 5 min and resuspending the cells in fresh culture medium.</li>
</ol>
2. Enumerating cells using the viability assay
<ol>
	<li>Pipette 450 &#181;L of propidium iodide in a microcentrifuge tube. Add 50 &#181;L of the cell suspension to the microcentrifuge tube. Incubate the tube at room temperature for 5 min. Enumerate the cells by flow cytometry (refer to section 4).</li>
	<li>Dilute all samples to a concentration of 1 &#215; 106 cells/mL with cell culture medium before proceeding to the apoptosis assays.</li>
</ol>
3. Staining cells for flow cytometry
NOTE: Sterile conditions are not required for this part of the protocol.
<ol>
	<li>Detection of phosphatidylserine externalization using Annexin V
	<ol>
		<li>Add 100 &#181;L of cell suspension into a microcentrifuge tube. Add 100 &#181;L of a 1:1 mixture of fluorescent conjugated Annexin V and 7-AAD reagent. Incubate at room temperature for 20 min, protected from light.</li>
	</ol>
	</li>
	<li>Analysis of mitochondrial membrane depolarization
	<ol>
		<li>Centrifuge 100 &#181;L of the cell suspension at 300 &#215; g for 5 min and discard the supernatant.</li>
		<li>Resuspend the cells in 1 mL of 1x PBS, add 100 &#181;L of TMRE staining solution to each sample, and mix the suspension by gentle back-pipetting.</li>
		<li>Incubate the cells for 20 min in a humidified CO2 environment at 37 &#176;C. Wrap the samples in clean aluminum foil to protect from light. 
		NOTE: Mitochondrial membrane potential is a functional marker that is sensitive to minor changes to the cell environment. Therefore, samples should be incubated and measured under identical conditions (temperature, pH, and time elapsed between the onset of incubation and fluorescent measurement) to maintain reproducibility).&#160;Also, note that inadequate protection of samples from light causes photobleaching of fluorophores resulting in falsely low emitted fluorescence.&#160;</li>
		<li>After incubation, add 5 &#956;L of the 7-AAD staining solution to each sample and mix. Incubate for 5 min at room temperature, protected from light.</li>
	</ol>
	</li>
	<li>Detection of activated terminal caspase-3 and -7 using caspase substrate DEVD 
	NOTE: The following solutions are prepared before performing this assay.
	<ol>
		<li>Dilute the DEVD-bound DNA-binding peptide in DMSO to a ratio of 1:8 with sterile 1x PBS to make the caspase-detection working solution. Store the solution on ice or at 2-8 &#176;C, protected from light. 
		NOTE: Each sample will require 5 &#181;L of this solution.</li>
		<li>Add 2 &#181;L of a 7-AAD stock solution to 148 &#181;L of 1x PBS to make the 7-AAD working solution. Store the solution on ice or at 2-8 &#176;C, protected from light. 
		NOTE: Each sample will require 150 &#181;L of this solution.</li>
		<li>Centrifuge 50 &#181;L of the cell suspension for 5 min at 300 &#215; g. Discard the supernatant. Resuspend the cells in 50 &#181;L of 1x PBS, followed by 5 &#181;L of the caspase-detection working solution. Mix thoroughly.</li>
		<li>Loosen the cap of the tubes and incubate for 30 min in a humidified CO2 environment at 37 &#176;C, protected from light. Add 150 &#181;L of the 7-AAD working solution to each sample and mix. Incubate for 5 min at room temperature, protected from light.</li>
	</ol>
	</li>
	<li>Detection of double-strand DNA breaks and total DNA damage
	<ol>
		<li>Centrifuge 50 &#181;L of the cell suspension for 5 min at 300 &#215; g. Discard the supernatant.</li>
		<li>Resuspend the cells in 500 &#181;L of 1x PBS. Add 500 &#181;L of a formaldehyde-based fixation buffer and mix. Incubate the samples on ice for 10 min.</li>
		<li>Centrifuge for 5 min at 300 &#215; g and discard the supernatant. Resuspend the cells in 90 &#181;L of 1x PBS in a microcentrifuge tube. Add 10 &#181;L of the antibody solution to the microcentrifuge tube. Incubate at room temperature for 30 min in the dark.</li>
		<li>Add 100 &#181;L of 1x PBS and centrifuge for 5 min at 300 &#215; g. Discard the supernatant. Resuspend the cells in 200 &#181;L of 1x PBS.</li>
	</ol>
	</li>
</ol>
4. Run samples on the flow cytometer.
<ol>
	<li>Verify the analytical performance of the flow cytometer by running the instrument&#39;s system check kit. Do not proceed with running samples until all checks are completed and passed.</li>
	<li>Locate the desired assay by browsing through the catalog of pre-programmed assays on the instrument and select Run Assay.
	<ol>
		<li>Mix the sample by gently back-pipetting before loading the sample onto the flow cytometer. 
		NOTE: Adequate mixing ensures cells remain in suspension and prevents low cell counts.</li>
		<li>First, load a negative control sample onto the flow cytometer, and select Run (Adjust Settings) so that the instrument begins aspirating the sample and provides a real-time preview of detected events. Refer to the Table of Materials for instrument name and details.</li>
		<li>Using the live preview, adjust the thresholds for fluorescence and cell size and draw a rectangular gate around the cell population. Drag the threshold marker to exclude cellular debris. Select Next (Set Health Profile) to proceed. 
		NOTE: It is important to know the size of the cells. Drag the sliders and observe changes in how detected events are plotted on the real-time preview, as this will inform an accurate selection of the thresholds. If cellular debris is not excluded at this stage, it cannot be removed in post-acquisition analyses.</li>
		<li>Tap and drag the quadrant markers to separate the cell populations and so that the instrument plots detected events in real-time. Use these plots to guide the end-user on the appropriate placement of the quadrant markers. Select Next (Verify Samples) to proceed so that the instrument displays a summary of the settings. After reviewing the settings, select Next (Verify Settings) to apply these settings for all samples within the experiment. 
		NOTE: The instrument provides a 2-min live preview of cells used to adjust the instrument&#39;s settings. If this time limit expires, the instrument will release the sample, and steps 4.2.2-4.2.4 will have to be repeated. Remove the sample and mix well before reloading and continuing.</li>
		<li>Gate a population of cells by drawing a region around the cell population. Adjust thresholds for fluorescence and cell size using the sliders located on the x- and y-axis of the live preview. Apply these settings for all samples within the experiment.</li>
		<li>Mix the first sample by gently back-pipetting and load the first sample onto the flow cytometer and select Next. Name the sample and select Run so that the system begins running the sample. 
		NOTE: The instrument can only run one sample at a time.</li>
		<li>Once all samples have been run, save the experiment by giving it an appropriate title. Save the settings of the current experiment for retrieval in future runs (optional).</li>
	</ol>
	</li>
</ol>
5. Post-acquisition analysis
<ol>
	<li>If required, perform fine-tuning of gates or quadrant markers post-acquisition.</li>
	<li>Locate the experiment that needs adjustment by navigating the system&#39;s file browser and open the experiment.</li>
	<li>Tap on the thumbnail preview of the plot to enlarge it. Tap on the top-left or bottom-right corners of the cell gate to adjust the dimensions of the gate. To adjust the quadrant markers, tap on the intersection of the vertical and horizontal lines to move the markers as they are. To adjust the angle of either line, tap on the line and drag the handle.</li>
	<li>Adjust the markers (as previously described in steps 4.1.3-4.1.4) as desired and apply these settings to all samples in the experiment by selecting the check-mark icon, marking all the samples, and selecting Accept.</li>
</ol>
6. Statistical analysis
<ol>
	<li>Run tests in triplicate and run an analysis of variance (ANOVA) with a post-hoc Bonferroni test to assess significant differences between the treated samples and controls. 
	NOTE: The statistical test of choice is dependent on the investigator and the variables being analyzed.</li>
</ol>

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Luminex&#174; Corporation kindly provided article processing charges.

In this study, actinomycin D-treated SiHa cells analyzed by FC revealed significant early and late biomarkers for apoptosis. Sub-optimal conditions for cell preparation, enumeration, and staining identified inaccurate results, emphasizing the need for close adherence to the manufacturer&#39;s instructions when performing FC.
This study of apoptosis by early and late biomarker identification conforms to the NCCD guidelines1 for investigating apoptosis. Actinomycin D-treated SiHa cultures demonstrated positive biomarkers for early and late apoptotic stages. The Annexin V/PI assay and the mitochondrial permeability assay showed that actinomycin D induced a PS-flip pattern and dissipation of the mitochondrial membrane transition, respectively. Once apoptotic cells reach the point of no return induced by mitochondrial perturbation, the terminal caspases are activated3,7. The activation of the terminal caspases 3 and 7 observed in this study indicates late-stage apoptosis. Furthermore, terminal caspase activation causes internucleosomal DNA cleavage and extensive DNA fragmentation, which classically has been reported as a step-ladder pattern observed by gel electrophoresis28,38.
The nuclear damage with the ATM and H2A.X FC DNA damage assay showed double-strand breaks in DNA and total DNA damage. These results confirmed classical downstream caspase-induced apoptotic nuclear damage (karyorrhexis and karyorrlysis) in experimental cultures. The use of multiple flow cytometric biomarkers thus detected the sequential multistage events in apoptosis and accurately and reproducibly identified cell populations at early and late apoptotic stages. These findings are consistent with the known pro-apoptotic features of actinomycin D in cancer treatment in humans37,39,40,41 and further support the use of actinomycin D as a positive control in FC cell culture experiments investigating apoptosis.
The gating of cell populations in this study was informed by negative medium and solvent controls, which separated apoptotic from healthy cells. Alternatively, a mixture of populations of positive and negative controls may also be used to define live and apoptotic cell populations to set cell population gates7,9. Once diseased and healthy cellular states are defined and gated, the template settings can be applied to all subsequent experimental and control cultures.
Strict adherence to the FC protocol is essential to avoid false results. During the optimization of the protocol, the following problems were observed: (1) low cell concentration, (2) high cell concentration, (3) cell clumping, (4) prolonged staining, and (5) poor sample handling. These problems can be prevented by strict adherence to optimized protocol requirements. This emphasizes the crucial nature of pre-analytical and analytical steps for FC to obtain accurate data. During cell preparation, trypsinization, pipetting, centrifugation, and dilutions must be conducted with care. Over-trypsinization and vigorous pipetting may result in chemical and mechanical shearing of cells, respectively. Prolonged and high-speed centrifugation may result in cell breakdown and high cellular debris counts. Optimal cell concentration is required to minimize the incorrect acquisition of cell events. Therefore, primary cell suspensions should be diluted to obtain optimal cell concentration.
Furthermore, while handling samples, care must be taken to prevent cell clumping and cell fragmentation and ensure that cells remain in suspension during analysis. Sample handling to prevent cell clumping allows single laminar cell flow, prevents mechanical blockage of the instrument&#39;s capillary tubing, and curbs spurious large cell size indexes. Another caveat is protecting cultures against light to avoid photo-oxidation and quenching of the fluorophores in the assays to prevent false-negative results. Care must be taken to ensure minimal light exposure at the staining step of the cells and subsequent processing steps. Moreover, prolonged immunostaining times may result in false-positive results as proteins are non-specifically stained. Therefore, adherence to manufacturer-prescribed incubation staining periods is important.
In summary, FC can accurately detect apoptosis and discriminate between early and late apoptosis biomarkers in cell culture. In addition, advances in technology have led to the manufacture of benchtop flow cytometers for non-expert scientists to study cell health and complex intracellular signaling pathways.

&#160;Apoptosis, classified as type 1 programmed cell death1, ensures an equilibrium between cell proliferation and cell death2. Apoptosis is essential during human development, post-injury, and for the prevention of diseases such as cancer3. Intrinsic and extrinsic apoptotic cell death signaling pathways4 cause sequential biochemical and morphological intracellular changes2,5,6. Morphological apoptotic features can be identified by microscopy, and the biochemical perturbation can be analyzed by biochemical assays, including flow cytometry (FC)7.
Flow cytometric analysis to identify apoptosis and understand the associated intracellular mechanisms has burgeoned over the last two decades8. FC is a scientific methodology that analyzes cells in a fluid that passes through single or multi-channel lasers (Figure 1)9,10,11. The cells in the fluid are focused into a single file by the fluidics system of the flow cytometer using hydrodynamic focusing. As cells pass through the laser, light is scattered or emitted from the cells. The scattered light can be in the forward direction (forward scatter) or toward the side (side scatter) and provides information about the cell size, and cell granularity or internal structures, respectively.
In addition, fluorescent reagents, such as fluorescent dyes or antibodies labeled with fluorophores, detect specific surface or intracellular structures or molecules. When the laser excites the fluorophores, light is emitted at a specific wavelength. Detectors&#8211;commonly photomultiplier tubes&#8211;quantify the scattered and emitted light from cell samples. The detectors produce a quantifiable current that is proportional to the light scatter and fluorescence emission. The electronic output is converted into digital signals by computing software to identify cell populations based on cell size, cell granularity, and the relative cell fluorescence of fluorophore-labeled molecules9,12,13.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/62663/62663fig1v3.jpg" /> 
Figure 1: Schematic describing the technical operation and workflow of traditional flow cytometry. Cells are stained with fluorescent reagents and probed by a laser. The fluorescence signals generated are detected and converted into an electronic output, which is further digitized and analyzed by computer software and statistical programs. Abbreviations: FSC = forward scatter; SSC = side scatter. <a href="https://www.jove.com/files/ftp_upload/62663/62663fig1v3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
FC is used in both research and health diagnostics. The two goals of FC in necrobiology are the elucidation of molecular and functional properties of cell death and the discrimination of various modes of cell death14,15,16,17,18. FC applications include cell enumeration, sorting of cell populations, immunophenotyping, biomarker detection (e.g., apoptosis biomarkers), toxicity studies, and protein engineering12. In addition, FC is commonly applied to health diagnostics to assist with the diagnosis and monitoring of patients with hematological malignancies. Advances in instrumentation, fluorophore detection, and detector systems are expanding FC applications to include imaging cytometry, mass cytometry and spectral cytometry with broader research applications12.
The flow cytometric analysis of apoptosis provides advantages over traditional techniques used in assessing cell health. FC can analyze many single cells in a heterogenous sample rapidly and reproducibly to estimate apoptosis3,5. The ability of FC to provide quantitative information on cell phenotypes on an individual cell basis, and avoid bulk analysis, offers superior sensitivity to Western blotting, enzyme-linked immunosorbent assays (ELISAs), fluorometry, and spectrophotometry techniques utilized in the analysis of apoptosis8,19. Furthermore, the relative ease of FC analysis in contrast to the cumbersome and poorly reproducible manual steps of Western blots and ELISAs is advantageous. The reproducible, accurate, and high-throughput analysis of FC therefore is beneficial in cancer research20.
FC also permits the simultaneous analysis of cell cycle parameters for healthy and abnormal apoptotic cell populations21. As apoptosis is a dynamic process, different methods can produce variable results and are dependent on the time-point at which the cells are harvested22. The simultaneous quantitative assessment of multiple parameters of cell phenotype permits the detection of minor subpopulations with high accuracy, e.g., rare cell subsets with a low frequency of 0.01% can be detected23. Multi-parametric FC analysis is especially useful as apoptotic death occurs along a spectrum of early and late biochemical changes with cells at various points along the apoptotic continuum. For example, the use of dual staining using Annexin V and propidium iodide in FC analysis of apoptotic cells enables the categorization of early apoptotic cells, late apoptotic cells, and dead cells24. The accurate detection of apoptosis at multiple stages avoids misclassification and false-negative results. Thus, multiparametric analysis by FC improves the overall specificity of detecting cell phenotypes and avoids the misclassification of minor populations. Furthermore, cell sorting by FC allows the isolation of cell populations with high purity for subsequent analysis7.
The disadvantage of FC includes the use of cells in suspension, which can be challenging in tissue analysis, as disaggregation of tissue into cells may alter cellular function19. Moreover, the lack of standardization of FC instrument setup, data analysis, and assay reports&#160;may cause variation in results19, emphasizing the need for optimally training FC operators to perform, and analyze and report&#160;data. For example, the ability of FC to discriminate true apoptotic debris from apoptotic nuclei requires i) proper acquisition settings, ii) the use of calibration beads to identify a diploid DNA peak, and iii) negative and positive cell controls that are cell-specific3. Furthermore, multiparametric analysis is limited by the number of detectors, and optimal compensation needs to be performed to avoid non-specific results and spillover of fluorescent emission when using multiple fluorescent reagents25. Advances in instrument and fluorophore technology have improved parameter detection to 30 parameters12.
The identification of apoptotic cell death is not always simple7, and sensitive and specific biomarkers should be considered. The Nomenclature Committee on Cell Death (NCCD) recommends that more than one assay be used to study and quantify the process of apoptosis26. Microscopy analysis for classical apoptotic features26 is also recommended to confirm apoptosis and avoid false-positive results7. Four cardinal biochemical features that span early and late apoptotic events are (1) loss of cell membrane asymmetry; (2) dissipation mitochondrial membrane potential (&#916;&#936;m); (3) caspase activation; and (4) DNA damage26.
During early apoptosis, phosphatidylserine is externalized to the outer cell membrane27 and can be detected by Annexin V fluorescently labeled with phycoerythrin27,28,29. Furthermore, dual staining with the fluorescent DNA-binding dye, 7-aminoactinomycin D (7-AAD), distinguishes live, late-apoptotic, and dead cells. Therefore, early-apoptotic cells stain positive for Annexin V and negative for 7-AAD, in contrast to late-apoptotic cells, that stain positive for both dyes24.
Intrinsic apoptotic signals induce dissipation of the mitochondrial membrane potential (&#916;&#936;m). The disrupted &#916;&#936;m causes the release of early pro-apoptotic proteins from the mitochondrial intermembrane space into the cytosol27,29,30. Change in &#916;&#936;m can be assessed by dual staining with the positively charged, lipophilic dye, tetramethylrhodamine ethyl ester, TMRE, and 7-AAD. The TMRE dye accumulates within the inner membrane of intact mitochondria when the membrane potential is high. Depolarized mitochondria demonstrate decreased fluorescence. Live cells with polarized mitochondria (intact mitochondrial membrane) stain positive for TMRE and negative for 7-AAD. Dead cells with depolarized mitochondria stain negative for TMRE and positive for 7-AAD31.
The caspases are a family of intracellular proteases that, when activated, signal and execute apoptosis26,27. The terminal executioner caspases (3,6,7) effect late apoptosis29,32,33. Caspase-3 and -7 activities can be measured by a fluorescently labeled substrate, which, when cleaved, binds to DNA and emits a fluorescent signal. Furthermore, any compromise to cell membrane integrity can be assessed by staining with 7-AAD. Apoptotic cells stain positive for the DNA-binding dye but negative for 7-AAD. Late-apoptotic and dead cells stain positive for both dyes34.
Late apoptosis is characterized by DNA damage27,29,35, which can be evaluated by phosphorylated ataxiatelangiectasia mutated kinase (ATM) and histone H2A.X. Double-stranded DNA breaks (DSBs) cause phosphorylation of H2A.X. Fluorescently labeled antibodies against ATM and H2A.X can determine DNA damage. Negative detection of both ATM and H2A.X indicate no DNA damage, while detection of both dyes indicates the presence of double-strand breaks in the DNA36.
Actinomycin D is a potent inducer of apoptosis and acts by binding to DNA to block transcription and translation events37. This study aimed to assess biochemical apoptosis induced by actinomycin D in the SiHa cell line by analyzing early- and late-stage biomarkers of apoptosis. Four biochemical biomarkers of apoptosis assessed the sequential steps in the apoptotic cascade that included loss of cell membrane asymmetry, change in mitochondrial membrane potential, activation of terminal caspases, and DNA damage.

<table><tbody><tr><td>6-well plates</td><td>Lasec</td><td>P1PLA044C-000006</td><td></td></tr><tr><td>Dimethyl Sulfoxide</td><td>Sigma-Aldrich</td><td>D8418</td><td></td></tr><tr><td>DMEM</td><td>ThermoFisher</td><td>41966052</td><td></td></tr><tr><td>Glutamine</td><td>Sigma-Aldrich</td><td>P10-040500</td><td></td></tr><tr><td>Guava Muse Cell Analyzer</td><td>Luminex</td><td>0500-3115</td><td></td></tr><tr><td>Microcentrifuge tubes/Eppendorf</td><td>Merck</td><td>EP0030122208-200EA</td><td></td></tr><tr><td>Muse Annexin V kit</td><td>Merck</td><td>MCH100105</td><td></td></tr><tr><td>Muse Caspase-3/7 kit</td><td>Merck</td><td>MCH100108</td><td></td></tr><tr><td>Muse Count and Viability kit</td><td>Merck</td><td>MCH600103</td><td></td></tr><tr><td>Muse DNA Damage kit</td><td>Merck</td><td>MCH200107</td><td></td></tr><tr><td>Muse MitoPotential kit</td><td>Merck</td><td>MCH100110</td><td></td></tr><tr><td>PBS Buffer</td><td>ThermoFisher</td><td>70013065</td><td></td></tr><tr><td>Pen-strep</td><td>Sigma-Aldrich</td><td>P4333</td><td></td></tr><tr><td>SiHa cells</td><td>ATCC</td><td>CRL-1550</td><td></td></tr><tr><td>T25 culture flasks</td><td>Sigma-Aldrich</td><td>C6231</td><td></td></tr><tr><td>Trypsin</td><td>Pan Biotech</td><td>P10-040500</td><td></td></tr></tbody></table>

flow cytometric analysis of apoptotic biomarkers in actinomycin d-treated siha cervical cancer cells

Apoptosis biomarkers were investigated in actinomycin D-treated SiHa cervical cancer cells using a benchtop flow cytometer. Early biomarkers (Annexin V and mitochondrial membrane potential) and late biomarkers (caspases 3 and 7, and DNA damage) of apoptosis were measured in experimental and control cultures. Cultures were incubated for 24 hours in a humidified incubator at 37 &#176;C with 5% CO2. The cells were then detached using trypsin and enumerated using a flow cytometric cell count assay. Cells were further analyzed for apoptosis using an Annexin V assay, a mitochondrial electrochemical transmembrane potential assay, a caspase 3/7 assay, and a DNA damage assay. This article provides an overview of apoptosis and traditional flow cytometry, and elaborates flow cytometric protocols for processing and analyzing SiHa cells. The results describe positive, negative, and sub-optimal experimental data. Also discussed are interpretation and caveats in performing flow cytometric analysis of apoptosis using this analytical platform. Flow cytometric analysis provides an accurate measurement of early and late biomarkers for apoptosis.

The cell count and viability (Figure 3) results showed that 95.2% of the cells in the sample were live, and 4.8% were dead. The total cell concentration in the original sample was 6.20 x 106 cells/mL.
The Annexin V and cell death assay (Figure 4) showed a significant increase (p &#60; 0.0001) in apoptotic cells in SiHa cells treated with 100 ng/mL actinomycin D in comparison to the controls. Because Annexin V staining is increased in cells during early stage apoptosis, this finding suggests that 100 ng/mL actinomycin D induced apoptosis in SiHa cells.
The mitochondrial electrochemical transmembrane potential assay (Figure 5) demonstrated a significant decrease (p &#60; 0.0001) in cell health profiles (live, depolarized and live, depolarized, and dead, dead) between actinomycin D, medium control, and solvent control. These data suggest that 100 ng/mL actinomycin D induced mitochondrial depolarization in SiHa cells.
The caspase 3/7 assay (Figure 6) demonstrated significant (p &#60; 0.0001) activation of caspases 3 and 7 in SiHa cells treated with 100 ng/mL actinomycin D compared to the controls. These finding demonstrates that 100 ng/mL actinomycin D induced apoptosis in SiHa cells.
The DNA damage assay (Figure 7) showed that 100 ng/mL actinomycin D significantly (p &#60; 0.0001) induced DNA damage markers, ATM and H2A.X, in SiHa cells. This finding suggests&#160;significant increase in DNA damage in SiHa cells treated with actinomycin D.
The results of suboptimal experiments (Figure 8) demonstrate analytical considerations across all assays. Cell concentration affects accuracy of data. In Figure 8A, the cell count is unacceptably low. The gated cell populations in all 4 quadrants of the dot plot have low signal intensity. The manufacturer optimizes&#160;the assay for 300-700 cells/&#181;L in the final sample volume.&#160;This example illustrates the importance of using the correct sample concentration prescribed by the manufacturer.
In addition, high cell concentrations also caused erroneous results (Figure 8B). Medium and experimental cultures demonstrated 99.95% and 100% live cells, respectively. The flow rate for both assays exceeded the manufacturer&#39;s optimized concentration of 100-500 cells/&#956;L and required dilution with 1x Assay Buffer to avoid inaccurate analysis.&#160;
Cell clumping should be avoided during the preparation of experimental cultures as it produces false results due to increased cell size indices, as illustrated in the Annexin V assay. Figure 8C shows a dual problem of high cell concentration that exceeds the manufacturer&#39;s instruction and cell clumping, as evidenced by cell size indices exceeding 4 in the SiHa medium control. The high cell concentration is illustrated by the sheeting of cells forming a bright red plane of cells in medium cultures, demonstrating discordantly high apoptotic cell populations.
Prolonged staining of cultures can result in non-specific binding of proteins and result in false results, as demonstrated by the Annexin V assay. Figure 8D demonstrates similar results for medium and experimental cultures due to prolonged staining.&#160;
Poor sample handling, extended trypsinization of adherent cells,&#160;vigorous pipetting during wash steps, and high-speed and prolonged centrifugation steps cause cell lysis and high amounts of cell debris. In Figure 8E, cultures analyzed by the caspase 3/7 assay demonstrate increased cell debris evidenced by a small cell size index (&#60;2.2 cell size index). Care should therefore be taken when preparing samples for data acquisition.
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/62663/62663fig03.jpg" /> 
Figure 3: Cell Count and viability assay. The population profile separates debris from live and dead cells. The two-dimensional dot plot is gated and divides live and dead cell populations. The information panel provides quantitative data on cell count, percentage of total viable cells, and the total number of viable cells in the sample. These data can be used to standardize the cell count in all samples for subsequent apoptosis analyses. <a href="https://www.jove.com/files/ftp_upload/62663/62663fig03large.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/62663/62663fig04.jpg" /> 
Figure 4: Annexin V assay. All samples show identical gating parameters, and statistical analysis of each sub-population reveals a significant increase in apoptosis in cells treated with actinomycin D. All assays were performed as three independent experiments, and each experiment was assayed in triplicate. Data are presented as mean &#177; SD, and p &#60; 0.05 was considered statistically significant. **** p &#60; 0.0001. <a href="https://www.jove.com/files/ftp_upload/62663/62663fig04large.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/62663/62663fig05.jpg" /> 
Figure 5: Mitochondrial electrochemical transmembrane potential assay. All samples show identical gating parameters, and statistical analysis of each sub-population reveals significant perturbation in mitochondrial membrane potential in cells treated with actinomycin D. All assays were performed as three independent experiments, and each experiment was assayed in triplicate. Data are presented as mean &#177; SD, and p &#60; 0.05 was considered statistically significant. **** p &#60; 0.0001. <a href="https://www.jove.com/files/ftp_upload/62663/62663fig05large.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/62663/62663fig06.jpg" /> 
Figure 6: Caspase 3/7 detection assay. All samples show identical gating parameters, and statistical analysis of each sub-population reveals a significant increase in caspase 3/7 activity in cells treated with actinomycin D. All assays were performed as three independent experiments, and each experiment was assayed in triplicate. Data are presented as mean &#177; SD, and p &#60; 0.05 was considered statistically significant. **** p &#60; 0.0001. <a href="https://www.jove.com/files/ftp_upload/62663/62663fig06large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 7" class="xfigimg" src="/files/ftp_upload/62663/62663fig07.jpg" /> 
Figure 7: DNA damage assay. All samples show identical gating parameters, and statistical analysis of each sub-population reveals a significant increase in double-strand DNA damage and total DNA damage in cells treated with actinomycin D. All assays were performed on three independent experiments, and each experiment was assayed in triplicate. Data are presented as mean &#177; SD, and p &#60; 0.05 was considered statistically significant. * p &#60; 0.05; *** p &#60; 0.001; **** p &#60; 0.0001. <a href="https://www.jove.com/files/ftp_upload/62663/62663fig07large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 8" class="xfigimg" src="/files/ftp_upload/62663/62663fig8v2.jpg" /> 
Figure 8: Sub-optimal experimental conditions yield poor results. (A) Low concentration of cells, (B) high concentration of cells, (C) cell clumping and aggregation evident by high cell size index, (D) prolonged staining of cell samples evident by increased positive staining in both samples, and (E) poor sample handling evident by increased cellular debris (cell size index &#60; 2.2). <a href="https://www.jove.com/files/ftp_upload/62663/62663fig8v2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Flow Cytometric Analysis of Apoptotic Biomarkers in Actinomycin D-Treated SiHa Cervical Cancer Cells at JoVE.com

Flow Cytometric Analysis of Apoptotic Biomarkers in Actinomycin D-Treated SiHa Cervical Cancer Cells

Apoptosis can be characterized by flow cytometric analysis of early and late apoptotic biomarkers. The cervical cancer cell line, SiHa, was analyzed for apoptosis biomarkers after treatment with Actinomycin D using a benchtop flow cytometer.

Apoptosis biomarkers were investigated in actinomycin D-treated SiHa cervical cancer cells using a benchtop flow cytometer. Early biomarkers (Annexin V ...

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Cancer Research

4.3K Views.  University of Pretoria. Flowcytometrie is een krachtige techniek die multiparametrische analyse van afzonderlijke cellen mogelijk maakt. Het kan biomarkers van vroege en late apoptose nauwkeurig en snel identificeren door miljoenen cellen te analyseren. Meerdere celmoleculen worden gekleurd en tegelijkertijd gedetecteerd.In het bijzonder maakt bench-top flowcytometrie analyse en interpretatie door niet-deskundige flowcytometristen mogelijk. Esther Zhou, een postdoctorale student uit de vitamine D-kankergroep,...

Video: Flow Cytometric Analysis of Apoptotic Biomarkers in Actinomycin D-Treated SiHa Cervical Cancer Cells

Flow Cytometric Detection of Newly-formed Breast Cancer Stem Cell-like Cells After Apoptosis Reversal

Cancer recurrence has long been studied by oncologists while the underlying mechanisms remain unclear. Recently, we and others found that a phenomenon named apoptosis reversal leads to increased tumorigenicity in various cell models under different stimuli. Previous studies have been focused on tracking this process in vitro and in vivo; however, the isolation of real reversed cells has yet to be achieved, which limits our understanding on the consequences of apoptosis reversal. Here, we take advantage of a Caspase-3/7 Green Detection dye to label cells with activated caspases after apoptotic induction. Cells with positive signals are further sorted out by fluorescence-activated cell sorting (FACS) for recovery. Morphological examination under confocal microscopy helps confirm the apoptotic status before FACS. An increase in tumorigenicity can often be attributed to the elevation in the percentage of cancer stem cell (CSC)-like cells. Also, given the heterogeneity of breast cancer, identifying the origin of these CSC-like cells would be critical to cancer treatment. Thus, we prepare breast non-stem cancer cells before triggering apoptosis, isolating caspase-activated cells and performing the apoptosis reversal procedure. Flow cytometry analysis reveals that breast CSC-like cells re-appear in the reversed group, indicating breast CSC-like cells are transited from breast non-stem cancer cells during apoptosis reversal. In summary, this protocol includes the isolation of apoptotic breast cancer cells and detection of changes in CSC percentage in reversed cells by flow cytometry.

Here, we present a protocol to isolate apoptotic breast cancer cells by fluorescence-activated cell sorting and further detect the transition of breast non-stem cancer cells to breast cancer stem cell-like cells after apoptosis reversal by flow cytometry.

Cancer recurrence has long been studied by oncologists while the underlying mechanisms remain unclear. Recently, we and others found that a phenomenon ...

Real Time Detection of In Vitro Tumor Cell Apoptosis Induced by CD8+ T Cells to Study Immune Suppressive Functions of Tumor-infiltrating Myeloid Cells

Potentiation of the tumor-killing ability of CD8+ T cells in tumors, along with their efficient tumor infiltration, is a key element of successful immunotherapies. Several studies have indicated that tumor infiltrating myeloid cells (e.g., myeloid-derived suppressor cells (MDSCs) and tumor-associated macrophages (TAMs)) suppress cytotoxicity of CD8+ T cells in the tumor microenvironment, and that targeting these regulatory myeloid cells can improve immunotherapies. Here, we present an in vitro assay system to evaluate immune suppressive effects of monocytic-MDSCs and TAMs on the tumor-killing ability of CD8+ T cells. To this end, we first cultured na&#239;ve splenic CD8+ T cells with anti-CD3/CD28 activating antibodies in the presence or absence of suppressor cells, and then co-cultured the pre-activated T cells with target cancer cells in the presence of a fluorogenic caspase-3 substrate. Fluorescence from the substrate in cancer cells was detected by real-time fluorescence microscopy as an indicator of T-cell induced tumor cell apoptosis. In this assay, we can successfully detect the increase of tumor cell apoptosis by CD8+ T cells and its suppression by pre-culture with TAMs or MDSCs. This functional assay is useful for investigating CD8+ T cell suppression mechanisms by regulatory myeloid cells and identifying druggable targets to overcome it via high throughput screening.

Real Time Detection of In Vitro Tumor Cell Apoptosis Induced by CD8+ T Cells to Study Immune Suppressive Functions of Tumor-infiltrating Myeloid Cells

We describe here a protocol to investigate cytotoxicity of pre-activated CD8+ T cells against cancer cells by detecting apoptotic cancer cells via real-time microscopy. This protocol can investigate mechanisms behind myeloid cell-induced T cell suppression and evaluate compounds aimed at replenishing T cells via blockade of immune suppressive myeloid cells.

Potentiation of the tumor-killing ability of CD8+ T cells in tumors, along with their efficient tumor infiltration, is a key element of successful ...

Assessing Cell Viability and Death in 3D Spheroid Cultures of Cancer Cells

Three-dimensional spheroids of cancer cells are important tools for both cancer drug screens and for gaining mechanistic insight into cancer cell biology. The power of this preparation lies in its ability to mimic many aspects of the in vivo conditions of tumors while being fast, cheap, and versatile enough to allow relatively high-throughput screening. The spheroid culture conditions can recapitulate the physico-chemical gradients in a tumor, including the increasing extracellular acidity, increased lactate, and decreasing glucose and oxygen availability, from the spheroid periphery to its core. Also, the mechanical properties and cell-cell interactions of in vivo tumors are in part mimicked by this model. The specific properties and consequently the optimal growth conditions, of 3D spheroids, differ widely between different types of cancer cells. Furthermore, the assessment of cell viability and death in 3D spheroids requires methods that differ in part from those employed for 2D cultures. Here we describe several protocols for preparing 3D spheroids of cancer cells, and for using such cultures to assess cell viability and death in the context of evaluating the efficacy of anticancer drugs.

Here, we present several simple methods for evaluating viability and death in 3D cancer cell spheroids, which mimic the physico-chemical gradients of in vivo tumors much better than the 2D culture. The spheroid model, therefore, allows evaluation of the cancer drug efficacy with improved translation to in vivo conditions.

Three-dimensional spheroids of cancer cells are important tools for both cancer drug screens and for gaining mechanistic insight into cancer cell biology. ...

Cell Cycle-specific Measurement of &#947;H2AX and Apoptosis After Genotoxic Stress by Flow Cytometry

The presented method or slightly modified versions have been devised to study specific treatment responses and side effects of various anti-cancer treatments as used in clinical oncology. It enables a quantitative and longitudinal analysis of the DNA damage response after genotoxic stress, as induced by radiotherapy and a multitude of anti-cancer drugs. The method covers all stages of the DNA damage response, providing endpoints for induction and repair of DNA double-strand breaks (DSBs), cell cycle arrest and cell death by apoptosis in case of repair failure. Combining these measurements provides information about cell cycle-dependent treatment effects and thus allows an in-depth study of the interplay between cellular proliferation and coping mechanisms against DNA damage. As the effect of many cancer therapeutics including chemotherapeutic agents and ionizing radiation is limited to or strongly varies according to specific cell cycle phases, correlative analyses rely on a robust and feasible method to assess the treatment effects on the DNA in a cell cycle-specific manner. This is not possible with single-endpoint assays and an important advantage of the presented method. The method is not restricted to any particular cell line and has been thoroughly tested in a multitude of tumor and normal tissue cell lines. It can be widely applied as a comprehensive genotoxicity assay in many fields of oncology besides radio-oncology, including environmental risk factor assessment, drug screening and evaluation of genetic instability in tumor cells.

The presented method combines the quantitative analysis of DNA double-strand breaks (DSBs), cell cycle distribution and apoptosis to enable cell cycle-specific evaluation of DSB induction and repair as well as the consequences of repair failure.

The presented method or slightly modified versions have been devised to study specific treatment responses and side effects of various anti-cancer ...

Far-Red Fluorescent Senescence-Associated &#946;-Galactosidase Probe for Identification and Enrichment of Senescent Tumor Cells by Flow Cytometry

Cellular senescence is a state of proliferative arrest induced by biological damage that normally accrues over years in aging cells but may also emerge rapidly in tumor cells as a response to damage induced by various cancer treatments. Tumor cell senescence is generally considered undesirable, as senescent cells become resistant to death and block tumor remission while exacerbating tumor malignancy and treatment resistance. Therefore, the identification of senescent tumor cells is of ongoing interest to the cancer research community. Various senescence assays exist, many based on the activity of the well-known senescence marker, senescence-associated beta-galactosidase (SA-&#946;-Gal). 
Typically, the SA-&#946;-Gal assay is performed using a chromogenic substrate (X-Gal) on fixed cells, with the slow and subjective enumeration of "blue" senescent cells by light microscopy. Improved assays using cell-permeant, fluorescent SA-&#946;-Gal substrates, including C12-FDG (green) and DDAO-Galactoside (DDAOG; far-red), have enabled the analysis of living cells and allowed the use of high-throughput fluorescent analysis platforms, including flow cytometers. C12-FDG is a well-documented probe for SA-&#946;-Gal, but its green fluorescent emission overlaps with intrinsic cellular autofluorescence (AF) that arises during senescence due to the accumulation of lipofuscin aggregates. By utilizing the far-red SA-&#946;-Gal probe DDAOG, green cellular autofluorescence can be used as a secondary parameter to confirm senescence, adding reliability to the assay. The remaining fluorescence channels can be used for cell viability staining or optional fluorescent immunolabeling.
Using flow cytometry, we demonstrate the use of DDAOG and lipofuscin autofluorescence as a dual-parameter assay for the identification of senescent tumor cells. Quantitation of the percentage of viable senescent cells is performed. If desired, an optional immunolabeling step may be included to evaluate cell surface antigens of interest. Identified senescent cells can also be flow cytometrically sorted and collected for downstream analysis. Collected senescent cells can be immediately lysed (e.g., for immunoassays or 'omics analysis) or further cultured.

A protocol for fluorescent, flow cytometric quantification of senescent cancer cells induced by chemotherapy drugs in cell culture or in murine tumor models is presented. Optional procedures include co-immunostaining, sample fixation to facilitate large batch or time point analysis, and the enrichment of viable senescent cells by flow cytometric sorting.

Cellular senescence is a state of proliferative arrest induced by biological damage that normally accrues over years in aging cells but may also emerge ...

Assessment of Mitochondrial Health in Cancer-Associated Fibroblasts Isolated from 3D Multicellular Lung Tumor Spheroids

Cancer-associated fibroblasts (CAFs) are among the most abundant stromal cells present in the tumor microenvironment, facilitating tumor growth and progression. Complexity within the tumor microenvironment, including tumor secretome, low-grade inflammation, hypoxia, and redox imbalance, fosters heterotypic interaction and allows the transformation of inactive resident fibroblasts to become active CAFs. CAFs are metabolically distinguished from normal fibroblasts (NFs) as they are more glycolytically active, produce higher levels of reactive oxygen species (ROS), and overexpress lactate exporter MCT-4, leading to the opening of the mitochondrial permeability transition pore (MPTP). Here a method has been described to analyze the mitochondrial health of activated CAFs isolated from the multicellular 3D tumor spheroids comprising of human lung adenocarcinoma cells (A549), human monocytes (THP-1), and human lung fibroblast cells (MRC5). Tumor spheroids were disintegrated at different time intervals and through magnetic-activated cell sorting, CAFs were isolated. The mitochondrial membrane potential of CAFs was assessed using JC-1 dye, ROS production by 2',7'-dichlorodihydrofluorescein diacetate (DCFDA) staining, and enzyme activity in the isolated CAFs. Analyzing the mitochondrial health of isolated CAFs provides a better understanding of the reverse Warburg effect and can also be applied to study the consequences of CAF mitochondrial changes, such as metabolic fluxes and the corresponding regulatory mechanisms on lung cancer heterogeneity. Thus, the present study advocates an understanding of tumor-stroma interactions on mitochondrial health. It would provide a platform to check mitochondrial-specific drug candidates for their efficacies against CAFs as potential therapeutics in the tumor microenvironment, thereby preventing CAF involvement in lung cancer progression.

Multicellular 3D tumor spheroids were prepared with lung adenocarcinoma cells, fibroblasts, and monocytes, followed by the isolation of cancer-associated fibroblasts (CAFs) from these spheroids. Isolated CAFs were compared with normal fibroblasts to assess mitochondrial health by studying the mitochondrial transmembrane potential, reactive oxygen species, and enzymatic activities.

Cancer-associated fibroblasts (CAFs) are among the most abundant stromal cells present in the tumor microenvironment, facilitating tumor growth and ...

Validation of FAM83A as a Target Gene in Cervical Cancer

Knockdown of FAM83A to Verify Its Role in Cervical Cancer Cell Growth and Cisplatin Sensitivity

The exploration of tumor target genes holds paramount importance for the prevention and treatment of cervical cancer. In this study, we outline the steps involved in the identification of a tumor target gene FAM83A in cervical cancer. First, the Cancer Genome Atlas dataset was employed to validate the expression and prognostic significance of FAM83A in women. A small interfering RNA (siRNA) was used for knockdown of the FAM83A gene in HeLa and C33a cells. Next, 5-ethynyl-2&#39;-deoxyuridine (EdU) staining was conducted to determine the effects on the proliferation capabilities of the tumor cells. Wound healing and porous membrane insert assays were performed to evaluate tumor cell migration and invasion abilities.
Western blotting was used to quantify apoptosis-related protein levels. JC-1 staining was employed to evaluate mitochondrial function alterations. Furthermore, cisplatin (diaminedichloroplatinum, DDP) intervention was used to assess the therapeutic potential of the target gene. Flow cytometry and colony formation assays were conducted to further validate the anticancer characteristics of the gene. As a result, FAM83A knockdown was shown to inhibit the proliferation, migration, and invasion of cervical cancer cells and sensitize these cells to cisplatin. These comprehensive methodologies collectively validate FAM83A as a tumor-associated target gene, holding promise as a potential therapeutic target in the prevention and treatment of cervical cancer.

Author Spotlight: Exploring the Role of FAM83A in Cervical Cancer 

Here, we show the procedures for FAM83A knockdown; the assays to detect its effects on proliferation, migration, and invasion of cervical cancer cells; and the sensitization of these cells to cisplatin. This study provides a promising target gene for cervical cancer and a reference for further drug research.

The exploration of tumor target genes holds paramount importance for the prevention and treatment of cervical cancer. In this study, we outline the steps ...

Measuring Caspase Activity Using a Fluorometric Assay or Flow Cytometry

The activation of cysteine proteases, known as caspases, remains an important process in multiple forms of cell death. Caspases are critical initiators and executioners of apoptosis, the most studied form of programmed cell death. Apoptosis occurs during developmental processes and is a necessary event in tissue homeostasis. Pyroptosis is another form of cell death that utilizes caspases and is a critical process in activating the immune system through the activation of the inflammasome, which results in the release of members of the interleukin-1 (IL-1) family. To assess caspase activity, target substrates can be assessed. However, sensitivity can be an issue when examining single cells or low-level activity. We demonstrate how a fluorogenic substrate can be used with a population-based assay or single-cell assay by flow cytometry. With proper controls, different amino acid sequences can be used to identify which caspases are active. Using these assays, the simultaneous loss of the inhibitors of apoptosis proteins upon tumor necrosis factor (TNF) stimulation has been identified, which primarily induces apoptosis in macrophages rather than other forms of cell death.

The present protocol describes two methods to measure caspase activity through a fluorogenic substrate using flow cytometry or a spectrofluorometer.

The activation of cysteine proteases, known as caspases, remains an important process in multiple forms of cell death. Caspases are critical initiators ...

Biochemistry

Modified Annexin V/Propidium Iodide Apoptosis Assay For Accurate Assessment of Cell Death

Studies of cellular apoptosis have been significantly impacted since the introduction of flow cytometry-based methods. Propidium iodide (PI) is widely used in conjunction with Annexin V to determine if cells are viable, apoptotic, or necrotic through differences in plasma membrane integrity and permeability1,2. The Annexin V/ PI protocol is a commonly used approach for studying apoptotic cells3. PI is used more often than other nuclear stains because it is economical, stable and a good indicator of cell viability, based on its capacity to exclude dye in living cells 4,5. The ability of PI to enter a cell is dependent upon the permeability of the membrane; PI does not stain live or early apoptotic cells due to the presence of an intact plasma membrane 1,2,6. In late apoptotic and necrotic cells, the integrity of the plasma and nuclear membranes decreases7,8, allowing PI to pass through the membranes, intercalate into nucleic acids, and display red fluorescence 1,2,9. Unfortunately, we find that conventional Annexin V/ PI protocols lead to a significant number of false positive events (up to 40%), which are associated with PI staining of RNA within the cytoplasmic compartment10. Primary cells and cell lines in a broad range of animal models are affected, with large cells (nuclear: cytoplasmic ratios &lt;0.5) showing the highest occurrence10. Herein, we demonstrate a modified Annexin V/ PI method that provides a significant improvement for assessment of cell death compared to conventional methods. This protocol takes advantage of changes in cellular permeability during cell fixing to promote entry of RNase A into cells following staining. Both the timing and concentration of RNase A have been optimized for removal of cytoplasmic RNA. The result is a significant improvement over conventional Annexin V/ PI protocols (&lt; 5% events with cytoplasmic PI staining).

An accurate method for the assessment of cell death is described. The protocol improves upon conventional Annexin V/ propidium iodide (PI) protocols, which display up to 40% false- positive events in cell lines and primary cells from a broad range of animal models.

Studies of cellular apoptosis have been significantly impacted since the introduction of flow cytometry-based methods. Propidium iodide (PI) is widely ...

Biology

Flow Cytometric Analysis of Biomarkers for Detecting Human Sperm Functional Defects

The conventional semen parameter analysis is widely used to assess male fertility. However, studies have found that ~15% of infertile patients show no abnormalities in conventional semen parameters. Additional technologies are needed to explain the idiopathic infertility and detect subtle sperm defects. Currently, biomarkers of sperm function, including sperm apoptosis, mitochondrial membrane potential (MMP), and DNA damage, reveal sperm physiology at the molecular level and are capable of predicting male fertility.
With flow cytometry (FCM) techniques, each of these markers can be rapidly, accurately, and precisely measured in human semen samples, but time costs substantially increase and results could be obstructed if all the biomarkers need to be tested with a single cytometer. In this protocol, after collection and immediate incubation at 37 &#176;C for liquefication, semen samples were further analyzed for sperm apoptosis using Annexin V-fluorescein isothiocyanate (FITC)/propidium iodide (PI) staining. The MMP was labeled with 5,5&#8242;,6,6&#8242;-tetrachloro-1,1&#8242;,3,3&#8242;-tetraethyl-benzimidazolylcarbocyanine iodide (JC-1) probe, and DNA damage was assessed using the sperm chromatin structure assay (SCSA) with acridine orange (AO) staining. Thus, flow cytometric analysis of sperm function markers can be a practical and reliable toolkit for the diagnosis of infertility and evaluation of sperm function at both bench and bed.

The present protocol provides a practical solution that allows for the measurement of apoptosis, mitochondrial membrane potential, and DNA damage in human sperm with a single cytometer.

The conventional semen parameter analysis is widely used to assess male fertility. However, studies have found that ~15% of infertile patients show no ...

Flow Cytometric Analysis of Apoptotic Biomarkers in Actinomycin D-Treated SiHa Cervical Cancer Cells

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