Flow Cytometry

Flow cytometry, a laser-based technology enabling single-cell analysis, was employed to investigate apoptosis and oxidative stress in response to soluble factors in vitro. In flow cytometry, the characteristics of each cell in suspension are investigated as they flow through a hydrodynamically-focused stream of liquid (sheath fluid); each particle or cell (“event”) is interrogated by a single-wavelength light beam. Detectors surrounding the laser beam measure a number of characteristics of each event at the point where the liquid stream and light beam meet. Forward light scatter (FSC), a measure of light refraction directly proportional to cell size, is measured by a detector in line with the light beam. Side light scatter (SSC), a measure of cell granularity, is measured by a number of detectors perpendicular to the light beam (as intracellular components can disperse light in all directions). Thus, FSC and SSC measurements help to identify cell populations based on size and internal complexity.

Figure 2.11. The flow cell of a flow cytometer. The cell suspension, which may be fluorescently labelled, enters the flow cell and is hydrodynamically focused in a stream of sheath fluid. Cells pass

Sample

(stained cells in suspension) Sheath fluid

Nozzle

Laser light source

Hydrodynamic focusing (cells pass through in single file)

Fluorescence emitted from stained cells detected

Forward and side scattered light from all cells detected

In addition, cells can be specifically interrogated for the presence of fluorophores when fluorescent dyes (e.g. intercalating dyes for nucleic acids, fluorescence-linked antibodies) are employed in sample preparation. These fluorophores can be excited by the laser as the event passes through the beam, and fluorescent detectors measure scattered emissions at a longer wavelength. Optimised laser parameters (e.g. voltage and gain), specific FSC/SSC settings, and a strategic gating technique help to develop a biochemical profile of the cell suspension, as the signals are amplified and visualised digitally on the FACSAria™ flow cytometer. The flow cell of a flow cytometer is depicted in Figure 2.11.

2.2.7.1 Apoptosis Assay

The Dead Cell Apoptosis Kit with Annexin V and PI (Thermo Fisher) was employed as in previous publications (Rochfort et al., 2014) for apoptosis and viability measurements in HAECs and HASMCs following exposure to soluble factors. This kit not only distinguishes between live and dead cells, but measures the degree of apoptosis at the point of interrogation.

This kit employs two fluorescent labels; first, PI, a red fluorescent dye that intercalates with available nucleic acid is used to distinguish between live and dead cells. Using the same logic as for cell counting (Section 2.2.1.5), PI cannot penetrate undamaged cell membranes and therefore is a suitable measure of the non-viable cell population. The second fluorescent dye, green Alexa Fluor®-488, is conjugated to recombinant Annexin V anticoagulant. In healthy live cells, a protein called phosphatidylserine (PS) is expressed on the inner surface cell membrane, while in apoptotic cells, PS is translocated to the extracellular surface. Annexin V phospholipid-binding protein has a high affinity for PS, and thus fluorescence-conjugated Annexin V acts as a suitable identifier of apoptotic cells. Using this dye combination in flow cytometry, viable non-apoptotic cells can be identified as having negligible fluorescence, viable apoptotic cells will have green fluorescence, and non-viable post-apoptotic cells have both red and green fluorescence (Figure 2.12). These distinct cell populations can be visualised using the 488 nm laser on the FACSAria™, exciting both PI and Alexa Fluor®-488 dyes which have differing emission spectra.

For the apoptosis assay, cells were first exposed to experimental treatments as required; two untreated negative controls (stained, unstained) and one apoptotic positive control (20%

DMSO, 30 minutes) were also included. Cells were trypsinised and washed in PBS while the Annexin V binding buffer (BB) (1/5 dilution of stock in dH2O) and 100 µg/mL PI (1/20 dilution

100 µL BB. Alexa Fluor®-488 Annexin V (5 µL) and PI (1 µL) were added to the suspension and incubated for 15 minutes at room temperature in the dark, with the exception of one untreated control. BB (400 µL) was then added and the suspension transferred to sterile Falcon® tubes. Tubes were kept on ice in the dark until ready to read.

The negative unstained control was first processed to determine the optimal FSC/SSC settings for each cell type. The positive control was run to ensure all three cell populations (healthy, apoptotic and non-viable) were clearly within range. All samples were read on the FITC (~530 nm) and PE-Texas Red (~610 nm) detection channels to 10,000 events. As PI and Alexa Fluor®-488 have relatively close emission maxima, a compensation protocol was employed to remove bleed-through fluorescence between channels. To do this, single-stained cell suspensions for both PI and Alexa Fluor®-488 were analysed alongside a double-stained sample. The auto-compensation settings on the Becton-Dickinson FACSDIVA™ software automatically calculates and eliminates the percentage of overlapping fluorescence, improving result specificity. Quadrant and gating strategies (e.g. doublet discrimination) were employed as required for data analysis via FACSDIVA™ and freely available Cyflogic software.

Unstained population

Double-stained population

Single-stained population

Figure 2.12. Example output of the apoptosis assay. (A) unstained live cells. (B) Alexa Fluor®-488 stained apoptotic cells. (C) Alexa Fluor®-488 and PI stained post-apoptotic non-viable cells. Image adapted from www.thermofisher.com.

2.2.7.2 Oxidative Stress

As for IF microscopy (Section 2.2.6.2), ROS levels were investigated using DHE staining by flow cytometry. Unlike qualitative microscopic analyses, ROS levels can be quantified using this technique by measuring the relative intensity of ethidium fluorescence, after DHE is oxidised, in control versus treated samples. In this case, HAECs were again exposed to 3 µM DHE prior to the completion of the incubation period (an unstained control was also included).

HAECs were then trypsinised and centrifuged to gather the cell pellet, washed in FACS buffer, re-centrifuged and re-suspended in 500 µL FACS buffer. After transferring to Falcon® tubes, samples were excited using the 488 nm laser and emission spectra read on the PE-Texas Red channel for 10,000 events. No compensation settings were required for DHE single stain analysis. Gating strategies were employed as required and data analysed via FACSDIVA™

and Cyflogic.