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Flow cytometric counting of apoptotic adhering cells

Flow cytometric counting of apoptotic adhering cells



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I have been trying to measure rates of apoptosis in epithelial cells by marking apoptotic cells with annexin V conjugated with phycoerythrin (PE) and sorting using flow cytometry. Unfortunately, the cells are adherent, and have been cultured in a collagen coated plate. Thus, to suspend them for flow cytometry, I have been using a trypsin treatment. Unfortunately, my results have been quite poor. I believe that there are several problems. First, the trypsin treatment is likely to induce the disruption of integral membrane proteins, potentially causing the externalization of phosphatidylserine (PS) (to which the annexin binds) on the outer plasma membrane of cells. Since the cells were incubated with the annexin V following suspension with trypsin, this could have led to a substantial increase in apparent rates of apoptosis, and masked real differences between rates of apoptosis in WT and KO cells. Furthermore, I was informed that epithelial cells, in order to maintain their structural integrity, have a relatively large ratio of cytoplasm to nucleus, and a prominent cytoskeleton resulting in increased background noise (greenish in color). In order to solve these problems I was considering several alternatives, such as changing the fluorescent marker that annexin V is conjugated with to reduce autofluorescent interference, using a different marker of apoptosis (in the place of annexin V), or, if I were to continue using annexin V, I think that incubating with annexin V followed by washing off of excess annexin, followed only then by trypsin treatment could potentially solve one of my various problems (or then again, it might not). Please help me find a suitable protocol for flow cytometric analysis of apoptotic markers in adherent epithelial cells. Additionally, if you know of any studies which utilize such methods, please direct me to them. Thanks in advance.


Here some resources,

  1. Cell harvesting effects on Annexin V staining.
  2. Protocol for apoptotic analysis using Annexin V.
  3. Quite a long discussion about this issue.
  4. Another protocol.

Cheers,

Pedro


How To Use Flow Cytometry To Measure Apoptosis, Necrosis, and Autophagy

Murder is a common theme in the mystery suspense genre. The detectives who solve these murders use a combination of observation and deduction to identify the guilty party. This metaphor suits measuring cell death.

In biology, there are four major pathways for cell death.

The study of the different ways cells die has become known as Cell Necrobiology, as coined by Darzynkiewicz and coworkers in their 1997 review article.

Flow cytometry is ideally suited as a tool to study Cell Necrobiology and, with its plethora of reagents, it is even possible to follow the different steps in these processes.

The four major ways a cell can die are:

    — programmed cell death — traumatic cell death — ordered degradation of cellular components (autophagocytosis) — ordered necrosis

Cell death is so important, that it has been the center of several Nobel prizes including one awarded in 2002 to Sydney Brenner, Robert Horvitz, and John Sulston who discovered the genes involved in apoptosis, and again in 2016 when Yoshinori Ohsumi was recognized for his work on the mechanisms of autophagy.

Of these mechanisms, apoptosis is probably the most readily studied using flow cytometry.

There are many assays that can be performed to measure apoptosis in cells. These can be grouped by the state of the cell as it dies.


Cell Biology

In cell biology research, controlling cell counting procedures at the earliest stage encourages reproducible results. Using an automated cell counter is recommended to ensure equal numbers of cells are used in downstream analysis, and to determine cell concentration in an accurate and precise manner.

Solutions

In cell biology research, whether you need cell count and viability, or to perform advanced fluorescence-based cell analysis with high reproducibility, we have a solution to suit your needs. Our instruments are automated fluorescence image cytometers designed for ease of use, precision and accuracy.

Obtaining high-quality research depends on your data’s reproducibility and consistency. NucleoCounter ® instruments perform a wide range of functions, from precise cell count and viability determinations to advanced cell sample analysis, with minimal user interference.

Plug-and-play assays include five different apoptosis assays, a cell cycle analysis and a GFP-transfection assay. In addition, a platform for user-definable assays is also available. Our instruments are designed for dedicated cell biology research and development laboratories.

Unbiased Cell Count & Viability Determinations

Manual cell counting using a hemocytometer and trypan blue exclusion method is time-consuming and depends heavily on the user’s perception and pipetting skills.

To determine cell count and viability using the NucleoCounter ® NC-202™ and NC-3000™ instruments, first load the cell suspension into the Via2-Cassette™ or Via1-Cassette™, respectively. Cells are automatically stained by the two fluorophores in the cassette, acridine orange and DAPI, staining total cells and dead cells, respectively. The volume of the counting chamber is pre-calibrated, ensuring high-precision and reproducibility.

High-speed Cell Viability & Counts

The multi-chamber NC-Slide A8™ enables high-speed viability and cell count determinations of insect and mammalian cells, measuring up to eight samples in less than three minutes. Mix the cell suspension with acridine orange and DAPI to stain all cells and the dead cells, respectively. Then, load the A8-slide™ chambers with the sample and insert the slide into the NucleoCounter ® NC-3000™.

We offer optimized counting protocols for aggregated cells, cells growing on microcarriers and in spheroids. Read more about cell count and viability using the NC-Slide A8™ for mammalian cells.

Superior Data Visuals & Analysis

The NucleoView™ software is the user interface for the NucleoCounter ® NC-200™ and NC-3000™. It handles data acquisition and presentation, along with image analysis. NucleoView™ includes a variety of data handling features that are perfectly suited to regulated environments.

The software enables you to visually inspect the fluorescence image and gives you the opportunity to verify the counting. You can select specific event populations in the scatter plots and examine and determine the validity of their inclusion or exclusion from the final counting results. Gating can be adjusted and you can save this adapted protocol for future measurements.

Counting Aggregated Cells

The NucleoCounter ® NC-202™ (and NC-200™) includes a wide range of specialized assays making it the perfect cell counter. Sphere cultures or highly aggregated cell types pose great challenges in cell counting, but with our specialized assays, these instruments can count even the most aggregated cell culture samples.

Standardized Apoptosis Assays

For advanced research into cell death, it is essential to study the mechanisms of apoptosis. Using the NucleoCounter ® NC-3000™, a series of five plug-and-play assays including Annexin V, mitochondrial potential with JC-1, caspase signaling, DNA fragmentation and the unique one-minute vitality assay, enabling you to fully investigate cell death mechanisms.

APOPTOSIS ASSAY PHYSIOLOGICAL CHANGES DETECTED STAGE
Mitochondrial potential Assay (JC-1) Collapse of the mitochondrial membrane potential Early
Annexin V Assay Collapse of plasma membrane lipid asymmetry Early-mid
Caspase Assay Caspase activation signals downstream apoptotic events Early-mid
Vitality (VB48™) Assay Decrease in cellular levels of reduced thiols e.g. GSH Late
DNA Fragmentation Assay Break-down and fragmentation of DNA Late

Covering early to late stages of apoptosis, these assays help you to carry out in-depth studies of apoptosis progression in mammalian cells. You can perform advanced data analysis effortlessly with NucleoView™. The platform presents links between images, histograms and scatter plots, giving you access to detailed and precise data analysis that identifies percentages of apoptotic, necrotic and living cells.

Monitoring Viability & GFP Transfection Efficiency

When transfecting cells with genes of interest, one way to determine the transfection efficiency is to co-express a fluorescent protein, such as green fluorescent protein (GFP) under the same promoter and determine the amount of GFP.

The NucleoCounter ® NC-3000™ offers a fast and easy to use assay to test GFP transfection efficiency and expression levels. By staining cells with Hoechst 33342 and propidium iodide (PI), you can define the total cell population and the dead cell population together with the population expressing GFP.

Cell staining in NucleoView™ software using GFP transfection efficiency assay with the NucleoCounter ® NC-3000™

A – Cells are located using Hoechst 33342 (blue) and the percentage of GFP expressing cells (green) can easily be determined. Non-viable cells are stained with propidium iodide (PI red).
B – All cells stained with Hoechst 33342 (blue) are identified.
C – GFP expressing cells are identified in green and non-viable cells in red by PI staining.

Easy coupling between the image obtained and the scatter plots or histograms allow you to determine the precise gate settings used to investigate the staining pattern for specific cell populations. The NucleoCounter ® NC-3000™ offers an all-in-one platform for evaluating GFP expression efficiency.

Non-viable cells stained with PI can be easily located with the image overlay function in the NucleoView™ software (A). GFP-transfected cells can be immediately identified (B).

Fast Cell Cycle Analysis

Investigating the impact of a treatment on cell division is one of the most powerful tools within cell biology. The NucleoCounter ® NC-3000™ provides fast and easy cell cycle analysis in under five minutes.

After adding a lysis buffer, all cell nuclei are stained, and the sample is measured using the NC-3000™. A cell cycle profile displays in the accompanying Plot Manager in the NucleoView™ software. Events in the sub-G1-phase, G0/G1-phase, S-phase and G2/M-phase are identified.

With the FlexiCyte™ software package, the NucleoCounter ® NC-3000™ can even be used to study cell proliferation with BrdU and EdU incorporation, detected with fluorescently labeled antibodies allowing for advanced studies of cell proliferation.

Illustration: Two-step cell cycle assay of untreated and camptothecin-treated (CPT) Jurkat cells. The histograms display intensity of the DNA-stain DAPI and can be used to define cell cycle events in the sub-G1-phase, G0/G1-phase, S-phase and G2/M-phase. After CPT treatment, the cell cycle is arrested in the G2/M-phase.

Advanced Cell Analysis

The FlexiCyte™ module available for the NucleoCounter ® NC-3000™ enables users to perform detailed advanced cell analysis of their own choice in mammalian and insect cells. The combination of LEDs, from UV to far red, together with a carefully chosen set of emission filters, allows you to detect a broad range of fluorescent antibodies and proteins.

The Protocol Adaptation Wizard feature guides the user through a selection of optimal settings. After image acquisition, in the Plot Manager, cell data is presented beside the fluorescent image as either scatter plots, histograms, or both. By linking images with plots, you are armed with everything you need to perform detailed data analyses.

The FlexiCyte™ software package enables detailed biomarker analysis of a broad range of fluorescent antibodies and proteins.


Induction Phase of Apoptosis

While it is not possible to measure intrinsically mediated apoptosis induction by flow cytometry, measuring extrinsically mediated apoptosis induction is relatively straightforward as it involves the analysis of the so-called death receptors (DR) CD95, CD261 (DR4), CD262 (DR5), CD120a, and CD120b, which can be detected using antibodies. Our Apoptosis Overview page goes into more detail about the two main apoptosis pathways.

Understanding the balance between death receptor expression patterns and anti-apoptotic mechanisms, signaling pathways and upregulation of these receptors is important in the fields of cancer, inflammation, tissue healing, transplantation and cell death. Flow cytometry is ideal to determine expression levels of these receptors in cell populations from various samples. Receptor levels e.g. CD95/FAS (Figure 1) can be determined in various cell types simply by FSC (forward) and SSC (side scatter)profiles where lymphocytes, monocytes and granulocytes can be identified. Alternatively specific lineage markers can be used like CD3 for T cells, CD19 for B cells and CD14 for monocytes in an immunophenotyping panel. This enables you to obtain information on the levels of induction of apoptosis in specific cells.


Table 1 below lists Bio-Rad&rsquos death receptor antibodies validated in flow cytometry. Note that care should be taken when immunophenotyping apoptotic cells. Dead cells can bind antibodies non-specifically so a viability dye is a must to avoid false positives. Staining protocols may also need to be optimized. Long incubation times may alter the apoptosis levels and intracellular staining requires fixation and permeabilization of cells.

Table 1. Flow cytometry validated antibodies against receptors of the extrinsic apoptosis pathway.

Receptor Ligand

Positive Control

Further Information

Human peripheral lymphocytes

Soluble version of CD95 (sCD95) acts as a decoy receptor inhibiting apoptosis

Human peripheral granulocytes

Cross-reacts with mouse CD266

Abbreviations: TRAIL, TNF-related apoptosis-inducing ligand TNF-R, tumor necrosis factor receptor TNFRSF12A, tumor necrosis factor receptor superfamily 12A TWEAK, TNF-like weak inducer of apoptosis.


Discussion

In the present study we have shown that a rather small fraction of peripheral blood CD34 + cells enter into apoptosis during the process of DMSO addition, freezing and thawing. On the other hand, the fraction of apoptosis and necrosis was high in the total cell population (Figure 1 and Table 2).

To our knowledge this is the first time apoptosis and necrosis have been measured in human cryopreserved peripheral blood CD34 + cells, using the well-documented annexin V flow cytometric method. Anthony et al 18 have stained fresh autologous PBPC concentrates with annexin V (without the double staining with nonvital dye), and report that 87.6% (range 30–96.6) of the CD34 + cells were annexin V-negative, which is almost identical to our figure of 81% (range 49–97). However, the figures are not strictly comparable, since the cells in our study were investigated after DMSO addition and freeze/thaw. De Boer et al 5 did flow cytometric measurements of frozen/thawed MACS-isolated CD34 + cells and showed that both loss of L-selectin expression and emergence of apoptosis occurred after freeze–thawing. Recently, Schuurhuis et al, 7 have extended this work by measuring early apoptosis in both purified and nonpurified CD34 + cells from cryopreserved leukapheresis samples. Using the early apoptotic marker Syto R 16 in combination with actinomycin D, they report that in unpurified frozen/thawed CD34 + PBPC from 15 patients, as many as 37% of the CD34 + cells were early apoptotic and 29% were necrotic. Their figure of apoptotic CD34 + cells is clearly higher than the figure in our investigation. Their measurements of apoptosis and necrosis in the CD34 + cells are more in accordance with our measurements of apoptosis and necrosis in the total cell population. According to Schuurhuis et al, Syto R 16 detects a somewhat higher fraction of early apoptotic cells than annexin V. However, we do not believe this can be the only reason for the large discrepancy of measurements. The addition of DMSO, freeze/thaw process, dilution and handling of the sample post thaw may influence the post-thaw viability. More flow cytometric studies of post-freeze/thaw PBPC apoptosis are needed to clarify the difference between the two studies.

The exposure of PS on the outer cell membrane acts as a recognition signal for macrophages to phagocytose the apoptotic cells. Thus, the annexin V-positive CD34 cells will probably be eliminated from the hematopoietic system and thus have no effect on the long-term engraftment following myeloablative chemotherapy. Quantitative measurements of annexin V-negative CD34 cells prior to reinfusion of PBPC may be important in the clinical setting to identify patients at risk of graft failure. This may be especially important for precursor cell concentrates with a low CD34 + cell count or for concentrates that have been manipulated by purging/positive selection. In the present study, none of the 11 patients were reinfused with PBPC concentrates containing less than 1.7 × 10 6 annexin V negative CD34 + cells/kg, and all the patients had a normal hematological recovery.

As shown in Figure 2, the present study indicates that one can get a rough estimate of the combined fraction of apoptotic + necrotic CD34 + cells, just by looking at the scatter characteristics. This is in accordance with other studies. 5,12 The observation that the apoptotic cells had a somewhat higher actinomycin D uptake than the viable cells and lower actinomycin D uptake than the necrotic cells, is also in accordance with other flow cytometric investigations. 16,20

We were surprised to find that as many as 31% (mean) of the total cell population in the PBPC concentrate (mainly mature blood cells), was in early apoptosis after cryopreservation. The late apoptotic/necrotic total cell fraction in our study was 33%, which is not too far from the findings of others. 21 Possible unwanted clinical implications of reinfusing a concentrate with 49–81% apoptotic and necrotic cells are unknown. The common clinical side-effects of having a thawed cryopreserved PBPC concentrate reinfused are flushing, nausea, dyspnea, post-infusion rigors and hypotension. These symptoms are usually attributed to DMSO-induced histamine release. However, a small clinical study has shown that these symptoms were also present in patients who had DMSO removed from the PBPC product before reinfusion, suggesting a possible harmful effect of reinfusing poorly cryopreserved non-viable cells. 22 On the other hand, one could speculate whether contaminating tumor cells in the PBPC concentrate may also become apoptotic and necrotic after the freeze/thaw process, suggesting a possible advantageous purging effect of the freezing procedure.

In conclusion, this study confirms the relative robustness of human CD34 + cells during the freeze/thaw procedures which are carried out in daily clinical practice. The flow cytometric three-color measurements of peripheral blood precursor cell concentrates stained with anti-CD34, annexin V and actinomycin D is a simple, fast and reproducible method. This investigation might be useful to assess the potential of progenitor cells for in vivo hematopoietic engraftment and further ex vivo manipulations, such as precursor cell expansion and gene therapy.


Discussion

It is becoming increasingly clear that the formation of ApoBDs during apoptosis is a highly regulated process 4,13,15 . However, the distribution of intracellular contents into ApoBDs is not well defined, possibly due to the lack of methodologies that could rapidly and accurately study this process. The data presented here demonstrate the ability to monitor and quantify the distribution of intracellular contents, including DNA, RNA and mitochondria, into ApoBDs efficiently by flow cytometry. It should be noted that other intracellular contents (e.g. other organelles or specific molecules) in ApoBDs could also be monitored using a similar approach if suitable staining or tracking methods are available. Likewise, similar approaches could be used for both adherent and non-adherent cells, as well as primary cells. Furthermore, utilizing the described flow cytometry-based approach, this work also establishes the concept that ApoBDs are not just one homogeneous identity, whereby subsets of ApoBDs can be defined based on their intracellular contents. This is particularly relevant for downstream functions of ApoBDs as different subsets of ApoBDs may exhibit different functions. For example, not all ApoBDs could mediate the transfer of autoantigens, cytokines or microRNAs to recipient cells. Furthermore, whether phagocytes could recognize and clear different subsets of ApoBDs differently remains to be determined.

As described in our recent studies 4,15,16 , apoptotic cell disassembly is regulated by three distinct morphological steps, namely membrane blebbing (Step 1), apoptotic protrusion formation (Step 2) and fragmentation to generate ApoBDs (Step 3). Importantly, apoptotic cells can utilize different mechanisms to generate ApoBDs depending on the activity of certain molecular factors 13,15,17 . In particular, manipulating the activity of key regulators of cell disassembly, such as ROCK1 and PANX1, can determine whether apoptotic cells will generate ApoBDs via apoptopodia (separate membrane blebs to form ApoBDs) or beaded apoptopodia (fragmentation of the apoptotic membrane protrusion to form ApoBDs) 13 . Using the flow cytometry method described in this study, we have provided evidence to support the idea that the content in ApoBDs could be influenced by the mechanism of their formation. For example, ApoBDs generated via beaded apoptopodia are less likely to contain a substantial amount of DNA compared with ApoBDs generated via apoptopodia. Thus, the distribution of intracellular contents into ApoBDs is not simply a stochastic process and could be regulated by the mechanism of apoptotic cell disassembly. Since different cell types can utilize different mechanisms to undergo cell disassembly during apoptosis 4,13,14,15,16 , ApoBDs generated by different cell types are likely to have different effects on physiological processes as well as the progression of certain diseases. It is interesting to note that previous studies have also proposed the packaging of intracellular contents such as DNA and RNA into ApoBDs is regulated, and DNA and RNA are segregated into different ApoBDs during apoptosis 22 . However, the data presented in this study suggest that although the distribution of intracellular contents into ApoBDs is a regulated process, DNA and RNA are not partitioned into separate ApoBDs. The discrepancy between previous work 22 and our current study can be attributed to how the ApoBDs were handled prior to analysis (e.g. fixed versus not fixed, cytocentrifuged versus not cytocentrifuged), and how ApoBDs were analysed (e.g. microscopy analysis based on ‘body-like structures’ versus flow cytometry analysis based on A5 staining as well as relative size and complexity/granularity).

Until now, the identification of ApoBDs in cell culture samples has largely been based on characteristics like phosphatidylserine exposure, as well as particle size and granularity 19 . In order to further study and characterize the function of ApoBDs in various disease settings, it is necessary to be able to accurately identify these vesicles. Here, we demonstrate that in addition to intracellular contents, ApoBDs can also be identified and subclassed based on the cell of origin from which they arise. Through analysis of surface marker expression on HUVEC, THP-1 monocytes and Jurkat T cells, it was observed that the expression of cell type specific markers (e.g. CD146, CD45 and CD3) are reduced on apoptotic cells and ApoBDs compared to viable cells. Although why these cell type specific markers are reduced on the surface of apoptotic cells is unclear, the expression of these markers was sufficient to identify different subsets of ApoBDs in mixed culture based on their cell origin. Notably, it is possible to identify the origin of ApoBDs from body fluids by flow cytometry if appropriate surface markers and gating strategies are established.

Collectively, the flow cytometry-based approaches described in this study could be used to monitor the formation of different subsets of ApoBDs, and better understand their function under physiological and pathological conditions.


MitoStatus TMRE

Panel 1. Flow cytometric analysis of TMRE staining in Jurkat Cells. Jurkat cells were treated with 0.025% DMSO (solid line histograms), 5 μM camptothecin (Left Plot, dashed line histogram) for 4 hr or 50 μM FCCP (20 min, Right Plot, dashed line histogram), and then stained with 100 nM BD Pharmingen™ MitoStatus TMRE (15 min, Cat. No. 564696).

Panel 2. Two-color flow cytometric analysis of TMRE staining in Jurkat Cells. Jurkat cells were treated with 0.025% DMSO (Left Plot) or 5 μM camptothecin (Right Plot) for 4 hr, then stained with 100 nM MitoStatus TMRE (15 min). Cells were resuspended in Annexin V Binding Buffer (Cat. No. 556454) and stained with APC Annexin V (15 min, Cat. No. 556419). Co-staining shows two main populations: healthy cells that are MitoStatus TMRE-positive and Annexin V-negative, and apoptotic or dead cells that are MitoStatus TMRE-negative and APC Annexin V-positive. Compared to the DMSO-vehicle treated control, camptothecin treatment increased the MitoStatus TMRE-negative and APC Annexin V-positive population, indicating that more cells are undergoing apoptosis. Loss of mitochondrial membrane potential is an early apoptotic marker so a small population of cells in transition is MitoStatus TMRE-negative and APC Annexin V-negative. Flow cytometric analysis was performed using a BD LSRFortessa™ Cell Analyzer System.

Panel 3. Immunofluorescent imaging of TMRE in HeLa Cells. HeLa cells were treated with 0.02% DMSO (Left Image) or 1 uM staurosporine (Right Image) for 3 hr. Cells were stained (30 min) with 200 nM MitoStatus TMRE and 5 ug/mL Hoechst. Staining media was removed and replaced with DPBS. Compared to the vehicle-treated control, staurosporine-treated cells show a decrease in mitochondrial staining with TMRE, as well as pyknotic nuclei characteristic of apoptotic cells. Cells were imaged on a BD Pathway™ 435 Cell Analyzer and merged using BD Attovision™ software.

MitoStatus TMRE has been tested on mouse (data not shown).

Panel 1. Flow cytometric analysis of TMRE staining in Jurkat Cells. Jurkat cells were treated with 0.025% DMSO (solid line histograms), 5 μM camptothecin (Left Plot, dashed line histogram) for 4 hr or 50 μM FCCP (20 min, Right Plot, dashed line histogram), and then stained with 100 nM BD Pharmingen™ MitoStatus TMRE (15 min, Cat. No. 564696).

Panel 2. Two-color flow cytometric analysis of TMRE staining in Jurkat Cells. Jurkat cells were treated with 0.025% DMSO (Left Plot) or 5 μM camptothecin (Right Plot) for 4 hr, then stained with 100 nM MitoStatus TMRE (15 min). Cells were resuspended in Annexin V Binding Buffer (Cat. No. 556454) and stained with APC Annexin V (15 min, Cat. No. 556419). Co-staining shows two main populations: healthy cells that are MitoStatus TMRE-positive and Annexin V-negative, and apoptotic or dead cells that are MitoStatus TMRE-negative and APC Annexin V-positive. Compared to the DMSO-vehicle treated control, camptothecin treatment increased the MitoStatus TMRE-negative and APC Annexin V-positive population, indicating that more cells are undergoing apoptosis. Loss of mitochondrial membrane potential is an early apoptotic marker so a small population of cells in transition is MitoStatus TMRE-negative and APC Annexin V-negative. Flow cytometric analysis was performed using a BD LSRFortessa™ Cell Analyzer System.

Panel 3. Immunofluorescent imaging of TMRE in HeLa Cells. HeLa cells were treated with 0.02% DMSO (Left Image) or 1 uM staurosporine (Right Image) for 3 hr. Cells were stained (30 min) with 200 nM MitoStatus TMRE and 5 ug/mL Hoechst. Staining media was removed and replaced with DPBS. Compared to the vehicle-treated control, staurosporine-treated cells show a decrease in mitochondrial staining with TMRE, as well as pyknotic nuclei characteristic of apoptotic cells. Cells were imaged on a BD Pathway™ 435 Cell Analyzer and merged using BD Attovision™ software.

MitoStatus TMRE has been tested on mouse (data not shown).

Panel 1. Flow cytometric analysis of TMRE staining in Jurkat Cells. Jurkat cells were treated with 0.025% DMSO (solid line histograms), 5 μM camptothecin (Left Plot, dashed line histogram) for 4 hr or 50 μM FCCP (20 min, Right Plot, dashed line histogram), and then stained with 100 nM BD Pharmingen™ MitoStatus TMRE (15 min, Cat. No. 564696).

Panel 2. Two-color flow cytometric analysis of TMRE staining in Jurkat Cells. Jurkat cells were treated with 0.025% DMSO (Left Plot) or 5 μM camptothecin (Right Plot) for 4 hr, then stained with 100 nM MitoStatus TMRE (15 min). Cells were resuspended in Annexin V Binding Buffer (Cat. No. 556454) and stained with APC Annexin V (15 min, Cat. No. 556419). Co-staining shows two main populations: healthy cells that are MitoStatus TMRE-positive and Annexin V-negative, and apoptotic or dead cells that are MitoStatus TMRE-negative and APC Annexin V-positive. Compared to the DMSO-vehicle treated control, camptothecin treatment increased the MitoStatus TMRE-negative and APC Annexin V-positive population, indicating that more cells are undergoing apoptosis. Loss of mitochondrial membrane potential is an early apoptotic marker so a small population of cells in transition is MitoStatus TMRE-negative and APC Annexin V-negative. Flow cytometric analysis was performed using a BD LSRFortessa™ Cell Analyzer System.

Panel 3. Immunofluorescent imaging of TMRE in HeLa Cells. HeLa cells were treated with 0.02% DMSO (Left Image) or 1 uM staurosporine (Right Image) for 3 hr. Cells were stained (30 min) with 200 nM MitoStatus TMRE and 5 ug/mL Hoechst. Staining media was removed and replaced with DPBS. Compared to the vehicle-treated control, staurosporine-treated cells show a decrease in mitochondrial staining with TMRE, as well as pyknotic nuclei characteristic of apoptotic cells. Cells were imaged on a BD Pathway™ 435 Cell Analyzer and merged using BD Attovision™ software.

MitoStatus TMRE has been tested on mouse (data not shown).

BD Pharmingen™ MitoStatus TMRE

Regulatory Status Legend

Any use of products other than the permitted use without the express written authorization of Becton, Dickinson and Company is strictly prohibited.

Recommended Assay Procedures

Bring BD Pharmingen™ MitoStatus TMRE dye powder and fresh cell culture-grade Dimethyl Sulfoxide (DMSO eg, Sigma D2650) to room temperature. Reconstitute TMRE dye powder in DMSO at a stock concentration of 0.2-1 mM. For example, 1 mg of BD Pharmingen™ MitoStatus TMRE can be dissolved in 1.94 mL of DMSO to yield a 1 mM stock solution (MW = 514.96 g/mol).

Upon arrival, store the dye powder desiccated and protected from light at ≤ -20°C until use. The dye powder is stable for at least 12 months if stored as indicated. After reconstitution with DMSO, store the stock solution at ≤ -20°C in small aliquots. The stock solution is stable for at least 6 months if stored as indicated.

Flow Cytometry Requirements

Before staining with this reagent, please confirm that your flow cytometer is capable of exciting the fluorochrome and discriminating the resulting fluorescence. Flow Cytometers (eg, BD FACSCanto™ II, BD LSRFortessa™, BD™ LSR II, or BD Accuri™ C6) equipped with a blue (eg, 488 nm) or yellow-green (eg, 561 nm) laser can be used. TMRE dye fluorescence can be detected with filters commonly used for phycoerythrin (PE) (eg, 575/26 or 582/15 nm).

Fluorescence compensation is best achieved using the cell samples of interest. When designing multicolor fluorescent staining panels, please be aware of high fluorescence spillover into the following fluorochromes' detectors: BD Horizon™ PE-CF594, BV605, BV650, or PerCP-Cy™5.5. We recommend titrating the dye and using the lowest concentration that provides adequate resolution of polarized and depolarized cell populations to reduce fluorescence spillover.

BD Pharmingen ™ MitoStatus TMRE labeling of suspended cells for flow cytometric analysis

1. Count cells to determine cell density. Adjust cell density to 1 × 10^6 cells/mL or less in fresh, pre-warmed cell culture media.

2. Stain cells in fresh, pre-warmed cell culture media or desired stain buffer with 20-200 nM BD Pharmingen ™ MitoStatus TMRE in a polypropylene container by adding the stock solution directly to the cells at the desired concentration.

a. Note: BD Pharmingen™ MitoStatus TMRE may also be added directly to the culture instead of staining in fresh media. Staining may also be performed in pre-warmed BD Pharmingen™ Stain Buffer (FBS) (Cat. No. 554656) or pre-warmed PBS. PBS may provide increased resolution for some cell types, but can also result in increased background staining.

b. Note: TMRE is known to stick to polystyrene. Samples should be stained in polypropylene containers.

c. Note: To aid in flow cytometric gating, we recommend using control cells treated with vehicle alone and/or cells treated with a mitochondrial uncoupler, such as FCCP [Carbonyl cyanide 4-(trifluoromethoxy)phenylhydrazone, eg, Sigma Cat. No. C2920].

3. Incubate samples for 15-30 minutes at 37°C protected from light.

4. Wash cells twice with BD Pharmingen™ Stain Buffer (FBS).

5. Decant the supernatant and gently mix to disrupt the cell pellet.

6. Resuspend the cells in Stain Buffer (FBS).

7. Analyze cells by flow cytometry. Alternatively, counterstain cells with compatible fluorescent dyes or antibodies as desired and then analyze.

BD Pharmingen™ MitoStatus TMRE labeling of adherent cells for flow cytometric analysis

1. Adherent cells should be stained in situ at ≤ 70% confluence. Stain cells in fresh, pre-warmed cell culture media or desired stain buffer with 20-200 nM BD Pharmingen ™ MitoStatus TMRE by adding the stock solution directly to the cells at the desired concentration.

a. Note: BD Pharmingen™ MitoStatus TMRE may also be added directly to the culture instead of staining in fresh media. Staining may also be performed in pre-warmed BD Pharmingen™ Stain Buffer (FBS) or pre-warmed Dulbecco's PBS (DPBS). DPBS may provide increased resolution for some cell types, but can also result in increased background staining.

b. Note: To aid in flow cytometric gating, we recommend using control cells treated with vehicle alone and/or cells treated with a mitochondrial uncoupler, such as FCCP..

2. Incubate samples for 15-30 minutes at 37°C protected from light.

3. Wash cells twice with BD Pharmingen™ Stain Buffer (FBS).

4. Remove cells from the growth medium. We recommend using BD™ Accutase™ Cell Detachment Solution (Cat. No. 561527).

5. Wash cells twice with BD Pharmingen™ Stain Buffer (FBS) or the equivalent.

6. Decant the supernatant and gently mix to disrupt the cell pellet.

7. Resuspend the cells in Stain Buffer (FBS).

8. Analyze cells by flow cytometry. Alternatively, counterstain cells with compatible fluorescent dyes or antibodies as desired and then analyze.

1. This dye is not compatible with cellular fixation.

2. Each user should determine the optimal concentrations of reagents, cells, and conditions for the assay of interest. We recommend titrating the reagent in early experiments to obtain optimal results.

3. Cells may be stained in bulk prior to staining with fluorescent antibodies.

BD Pharmingen™ MitoStatus TMRE labeling of cells for fluorescent imaging

1. Stain cells in fresh, pre-warmed media with 20-200 nM BD Pharmingen™ MitoStatus TMRE.

2. Incubate samples for 15-30 minutes at 37°C protected from light.

3. Remove the staining solution and replace with pre-warmed DPBS.

4. Analyze cells by fluorescence imaging. Alternatively, counterstain cells with compatible fluorescent dyes or antibodies as desired and then analyze.


Measuring Apoptosis

One of the most common features of apoptosis that can be measured by flow cytometry is externalization of phosphatidylserine (PS), a phospholipid found in the inner membrane of healthy cells. Annexin V binds to phosphatidyl serine and thus annexin V labeled with fluorophores allow apoptosis to be assessed, usually in combination with a viability dye such as propidium iodide (PI) to distinguish apoptotic from necrotic cells. Healthy cells are negative for both markers, apoptotic cells are positive for annexin V and necrotic cells are positive for both markers. When Jurkat T cells are treated with staurosporine they undergo apoptosis, followed by necrosis. The time course in Figure 33 below shows the increase in annexin V staining after 1 hour as they apoptose, followed by a subsequent increase in necrotic cells by 6 hrs.

Fig. 33. Annexin V staining to measure apoptosis. Jurkat cells were treated with staurosporine at 1 &muM for 0 hr, 1 hr and 6 hr to induce apoptosis. The cells were then stained with annexin V FITC (ANNEX300F) and ReadiDrop&trade propidium iodide (1351101). Apoptotic cells positive for annexin V can be seen in the bottom right quadrant and dead cells positive for both annexin and PI in the top right quadrant. Healthy cells are negative for both stains.

Because phosphatidylserine externalization is a dynamic, reversible process until a cell is committed to apoptosis after mitochondrial outer membrane permeabilization (MOMP), annexin V conjugates are unable to distinguish early apoptotis from late apoptosis. Polarity-sensitive indicator of viability and apoptosis (pSIVA) probes are biosensors that reversibly bind to PS and thus turn on and off as PS flips from the outer membrane to the inner membrane. This allows easy comparison of differences in apoptosis rates in response to different experimental treatments in real time.


Why You Need To Remove Dead Cells

Dead cells should be removed from all flow cytometry experiments that aim to evaluate live cell lineage and functionality.

Below are two simple examples of why you need to remove non-viable cells prior to implementing your flow cytometry gating strategies.

The two plots display mesenchymal stromal cells (MSC) produced in a Good Manufacturing Practices lab. The lab in question is very good at producing these cells from bone marrow and routinely generates 100% CD45-negative cells after their analysis.

In the panel on the left, the sample is stained with a dead cell marker only. Here, you can easily see what appears to be CD45 contamination of their MSC product in the dead cell (dead cell marker-positive) fraction. This would NOT be possible without the dead cell marker. Also, as discussed above, the left panel reveals the higher levels of autofluorescence in the dead cells. Finally, by adding anti-CD45 antibody to the sample (right panel)you see both the autofluorescent population, as well as a separate CD45-positive population that’s taking up the antibody.

Fortunately for scientists and flow cytometrists like you, there are multiple ways to label and identify dead cells so they can be removed from your flow cytometry analysis and cell sorting experiments.


DISCUSSION

Although the molecular mechanisms involved in phagocytosis of apoptotic cells are becoming better understood, little is known about the recognition and uptake of necrotically dying cells. Here, we used differentially labeled macrophages and target cells to study the uptake of apoptotic and necrotic cells in a quantitative flow cytometry phagocytosis assay. The L929 fibrosarcoma cellular system provided us with well-characterized models of apoptosis and necrosis mediated by a death domain receptor (Denecker et al., 2001b), distinct from cell death caused by physical damage such as rupture of the cell membrane by douncing, freeze thawing, or exposure to high temperature (Sauter et al., 2000 Cocco and Ucker, 2001 Fadok et al., 2001a).

Previous studies showed that PS is a prerequisite for the uptake of apoptotic cells by macrophages. PS exposed on the apoptotic cell membrane is recognized directly by the macrophage PS receptor (Fadok et al., 2000) or indirectly via binding of PS to a soluble intermediate (milk fat globule-epidermal growth factor-factor 8), which is secreted by macrophages, followed by recognition of the complex by the αvβ3 integrin on the phagocyte's surface (Hanayama et al., 2002). Our results demonstrate that both apoptotic and necrotic target cells are recognized and phagocytosed by macrophages, whereas viable control cells are not. Uptake of apoptotic and necrotic cells occurred irrespective of the stimulus inducing cell death. Phagocytosis of necrotic cells is a feature common to different macrophage cell lines and also occurs in vivo, as demonstrated in Figure 8. Apoptotic cells are taken up very efficiently as soon as PS exposure becomes evident, at a time when their membrane is still intact. Uptake of necrotic cells takes place when cells loose the integrity of the membrane and become PS positive. Moreover, recombinant annexin V can inhibit phagocytosis of both types of dying cells, suggesting that externalization of PS is a general signal indicating to phagocytes that a dying cell should be cleared, irrespective of the way the cell has died. Indeed, PS exposure is an old “eat me signal” also used for the recognition and uptake of aged erythrocytes (Schlegel and Williamson, 2001). Although the recognition of dying L929sA cells seems to be dependent on PS for both apoptotic and necrotic cells, the efficiency of uptake is higher for the former. Moreover, when given the choice, macrophages prefer to clear early apoptotic cells, as demonstrated by the competition experiment (Figure 6), even though these target cells present a lower mean annexin V staining level. The lower efficiency of uptake of necrotic cells might be due to physical constraints. It is possible that the PS exposed on necrotic and late apoptotic cells is less accessible to the macrophages. Moreover, although apoptotic cells fragment into many small-contained particles that are easy to recognize and engulf, necrotic cells swell and remain as large single entities for a long time. Similar observations were made in Caenorhabditis elegans, where necrotic corpses linger for much longer than apoptotic ones, probably due to their larger volume, although uptake of both types of dying cells requires the same engulfment genes (Chung et al., 2000). Indeed, electron microscopy analysis presented here suggests that the mechanism used by macrophages to engulf necrotic cells is morphologically different from that used for apoptotic cells. Macrophages that engulf necrotic cells protrude into the swollen ghost-like structures of the dying cells, grasping only small volumes of the cellular debris, whereas apoptotic cells are readily engulfed as contained distinct apoptotic bodies (Krysko et al., 2003).

Phagocytosis of apoptotic cells does not induce inflammation and is often referred to as a silent event (Fadok et al., 1998 Cocco and Ucker, 2001 Fadok et al., 2001a). A possible consequence of the greater difficulty apparently encountered by macrophages in clearing necrotic cells is spillage of the cellular contents of the dying cells, leading to more histotoxicity and development of proinflammatory (Haslett, 1992 Haslett et al., 1994 Wiegand et al., 2001 Medan et al., 2002) or autoimmune responses (Rovere et al., 2000 Chan et al., 2001 Magnus et al., 2001). Indeed, apoptotic neutrophils and PS-exposing membranes were reported to elicit an antiinflammatory effect, whereas incubation of macrophages with physically lysed neutrophils, but not with lysed lymphocytes, significantly stimulated the production of macrophage-inflammatory protein 2, IL-8, TNF-á, and IL-10 (Fadok et al., 2001a). However, the latter effect was attributed to the cleavage of PS-receptor on the surface of the macrophages by elastase, a protease released upon the lysis of neutrophils. The cleavage of PS-receptor probably also impairs the phagocytic capacity of the macrophages and may cause them some stress (Vandivier et al., 2002). Our results show that phagocytosis of apoptotic or necrotic cells by Mf4/4 macrophages does not induce the expression of IFNβ, TGFβ, TNF, or IL-6, neither at the mRNA nor at the protein level. On the other hand, exposure of the same macrophages to LPS strongly increased the expression of TNF and IL-6 and moderately increased the levels of IFNβ mRNA. Moreover, phagocytosis of apoptotic or necrotic L929 cells did not affect this LPS-induced response. Several reports are in agreement with these findings. Cocco and Ucker (2001) observed that heat-killed necrotic and late apoptotic thymoma and T-cell hybridoma target cells did not induce J774A.1 and RAW 264.7 macrophages to secrete TNF or IL-6. Furthermore, no signs of inflammation accompanied the necrotic interdigital cell death occurring in Apaf-1–deficient mice and further development proceeded normally (Chautan et al., 1999). Although inflammation often accompanies necrosis, the above-mentioned observations and our results indicate that this is not due to the induction of expression of proinflammatory cytokines in the macrophages clearing the dying cells. Therefore, it is conceivable that the release of cytokines or other factors from the necrotic cells themselves may be crucial for an inflammatory response. Additionally, our results demonstrate that the clearance of primary and probably also secondary necrotic cells is clearly less efficient and more difficult and time consuming than that of apoptotic cells. This process may cause the macrophages to remain at the same site longer, thus heightening the inflammatory state. These results suggest that prevention of necrosis and secondary necrosis, and promotion of apoptosis may allow a more rapid and efficient clearance of the dying cells and decrease the damage to the surrounding tissue both in injury and in antitumor cancer treatment.


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