Flow cytometric analysis of cultured T-cell viability via Annexin V and propidium iodide staining

Introduction
Cell viability is a critical factor during any experiment in flow cytometry. The proportions of dead or dying cells can vary greatly due to growth state of in vitro culture, time spent ex vivo, exposure to reagents and protocol procedures in addition to myriad other factors. Not only can dying cells lack the functional responses that may be being assessed in the experiment, but they also generally have increased non-specific binding due to sugar and nucleic acid based ‘stickiness’ which may lead to false positives. In addition, necrotic cells can also induce cell activation in samples and may lead to skewed results. Fortunately, dead or dying cell proportions can be analyzed using viability stains to assess impact on results, or to gate them out and analyze only live cells. Herein we demonstrate the use of an annexin V and propidium iodide assay to distinguish live from necrotic and apoptotic Jurkat T cells.

Materials and Methods

Lymphocyte staining with annexin V and propidium iodide.
Jurkat T lymphocytes were cultured and processed following the MMI 590 Lab 2 protocol to be run on a BD FACS Canto II flow cytometer. Briefly, cultured cells were spun down at 200 x g for 5 minutes and resuspended in fresh complete RPMI media. 1x10⁶ cells were seeded into FACS tubes with varying concentrations of ethanol and incubated at 37°C, 5% CO₂ for 40 minutes. One 0% ethanol tube was held on ice for 2 hours prior to incubation. Cells were then washed twice and resuspended in 1x annexin V binding buffer. Annexin V-FITC and propidium iodide were then added to each tube and incubated for 20 minutes in the dark at room temperature. Extra tubes incubated with 25% ethanol were stained with Annexin V or propidium iodide only for compensation controls. Fluorescence signals were compensated then experimental samples were analyzed on a BD FACS Canto II, gating out debris and doublets using the gating strategy described in MMI 490 Lab 1. Extra tubes incubated with 25% ethanol were stained with Annexin V or propidium iodide only for compensation controls.

Results and Discussion

Annexin V and propidium iodide can be used to distinguish specific states of cell death.
Cell viability is important to any experiment, as it can greatly impact results. Fortunately, this can easily analyzed using flow cytometry with an Annexin V- propidium iodide stain. Phosphatidylerine (PS) is a phospholipid anchored into the cytoplasmic side of the plasma membrane of healthy cells. When apoptosis is triggered, flippases translocate PS to the outer membrane surface where it acts as an apoptotic signal for macrophages to clear the cell. Annexin V binds PS with anti-body like affinity and is used in this experiment to distinguish cells undergoing apoptosis by high FITC and low propidium iodide (PI) staining (Figure 1, Q4). Propidium iodide is non-membrane permeable and will stain any cells with compromised membrane integrity, namely necrotic cells (Figure 1, Q1). Because Annexin V is a large protein, it generally cannot pass to the cytoplasm without a large amount of membrane damage. Thus double positive (Figure 1, Q2) cells are either late apoptotic and have extracellular facing PS with membrane degradation to allow PI staining, or they are late necrotic and have so much membrane damage that Annexin V can enter the cell to bind cytoplasmic facing PS. Herein, this assay is able to discern viable cells (Figure 1, Q3) from apoptotic, necrotic, and late necrotic/apoptotic populations. These two latter populations could be distinguished by use of a TUNEL kit to look for apoptosis triggered DNA fragmentation. The downside of Annexin-PI staining being that annexin requires a specific calcium concentration to bind PS, and may not be compatible with buffers needed for other fluorescent stains. This could be remedied by the use of an array of commercially available live/dead stains.

Figure 1. Annexin V and propidium iodide stains can distinguish live and dead cell populations. FACS plot of Jurkat T-cells incubated in 5% ethanol at 37°C in 5% CO2­­ ­for 40 minutes then stained with Annexin V-FITC and propidium iodide. Samples were acquired on a BD FACS Canto II. Percent cell population indicated in the corner of each gate.

Increasing concentrations of ethanol induces different states of cell death in cultured cells.
Experimental protocols can have varying effects on live cells, and it is important to know whether or not your cells are still viable. If a large proportion of cells are non- viable, it may be a clue that a protocol may have to be further optimized for those specific cells, or that a different assay should be used all together. In this experiment, we exposed Jurkat cells to increasing concentrations of ethanol to interesting effect. 5% ethanol exposure for 40 minutes showed little effect, with comparable cell viability to 0% controls (Figure 2). Incubation in 25% ethanol shifted almost the entire the population (93.2%) to double Annexin-PI positive, indicating a late necrotic/ apoptotic state. Interestingly 50% ethanol shifted most of this double positive population to PI positive or necrotic cells making up 22.8% and 74.4% of the cell population respectively. Perhaps, some property of 25% but not 50% ethanol allows for PS to reach the outer leaflet of the membrane, or destabilizes the membrane such that Annexin can enter. This experiment highlights the use of viability stains to assess the effects of protocols on cells, as 25% and 50% ethanol samples are useless for assaying live cells. In addition, although 5% ethanol protocol induced some cell death, viability stains allow the capacity to gate on only viable cells to garnish the most robust data possible.

Figure 2. Viability of Jurkat T-cells incubated with varying concentrations of ethanol. Cultured Jurkat T-cells were seeded into complete RPMI containing varying concentrations of ethanol prior to incubation at 37°C in 5% CO2­­ ­for 40 minutes. Ice shocked cells were held on ice for 2h before incubation with 0% ethanol. Cells were then washed and stained with Annexin V and propidium iodide prior to analysis on a BD FACS Canto II (n=1). Colors correspond populations scene in figure 1.

Light scatter properties of cultured murine macrophage and human T cells

Light scatter properties of cultured murine macrophage and human T cells.

Introduction
Flow cytometry has been a cornerstone in biological advancement, and in immunology in particular over the last 30 years. This technique achieves high levels of statistical robustness via high throughput analysis of individual cells using lasers, light scatter and fluorescent markers. Light scatter is a crucial component of flow cytometry that uses forward scatter to discern relative size and side scatter to measure relative internal complexity of individual cells. These properties can be used to gate out doublets, clumps and debris, identify distinct cell populations, or to look at any relative morphological changes within a population such as activation. Here in we will analyse the light scatter properties of Jurkat T cells and RAW macrophage cells using flow cytometry to understand how we can employ side and forward scatter to produce the best possible data in our future experiments.

Materials and Methods

Forward and side scatter analysis of in vitro cultured cell lines
RAW murine macrophage and Jurkat T lymphocytes were processed following the MMI 590 Lab 1 protocol to be run on a BD FACS Canto II flow cytometer. Briefly, cultured cells were spun down and resuspended in FACS Buffer. Cells were counted and due to low cell concentrations, 1.6x10⁵ RAW cells and 5.7x10⁵ Jurkat cells (in 1mL) were seeded into their respective FACS tube. The combined tube held  7.9x10⁴ and 2.9x10⁵. Combined results used in the data portion were obtained from Group 2 due to long sample acquisition times from low cell concentrations. FACS Canto II lasers were set at 330mV and 450mV for FSC and SSC respectively.

Results and Discussion

Light scatter can be employed to remove debris and doubletsCellular light scatter properties are an important tool when setting up any good flow cytometry experiment, as it can help to remove events that may skew results and thus the conclusions garnished from the experiment. The gating strategy used in this experiment (Figure 1a) displays this importance with 11.8% of total events (black dots) gated out in the P1 gate which is mostly debris of low size (forward scatter). The population remaining contains both the RAW and Jurkat cell populations within them, but there are still likely clumps and doublets that may skew data, especially in experiments looking for co-expression or other double-positive events since two single positive cells may appear as a double positive event if not gated out properly. To achieve this, cells with a low ratio of forward scatter area vs height are gated out (P2, red dots). When two cells are counted as a single event, their area to height ratio will be roughly twice that of a single cell event pulling the respective dot out of the bottom of the P2 gate and disregarding it from the dataset. In the mixed population, this removes an additional 12.4% of total events captured for a total of 24.2% of events removed with 52.8% and 20.7% of events removed for RAW and Jurkat only samples respectively (Figure 1b). Removing these events makes the dataset and thus and conclusions made more robust.

Figure 1. Light scatter profiles of RAW and Jurkat cell lines. a) Gating strategies on a mixed RAW and Jurkat cell population included excluding low side scatter and forward scatter events (P1 red and green). High FSC area to FSC height ratio events (red) were then excluded from this population to yield a population for final analysis (green). b) Forward and side scatter plots for post-gated RAW macrophage, Jurkat T- lymphocytes and a mixed population containing both cultures. Proportion of total captured events remaining for analysis is indicated at the top right of each plot.

Light scatter can be used to distinguish distinct cell populations
Light scatter can also be used to distinguish cell populations that are morphologically different by relative size measured by forward scatter, or relative internal complexity measured by side scatter.
We saw that the RAW macrophage sat, on average, at a lower forward scatter than the Jurkat cells indicating that they are on average smaller, although the Jurkat only population had proportion of cells that sat at the smaller range of RAW cells (figure 1b). These could be apoptotic cells and could possibly be gated out by a viability assay. Unfortunately, the use of some kind of fluorescent marker would be needed to separate these two cell populations in flow analysis because of their overlapping forward and side scatter profiles.