Experimental protocol to study cell viability and apoptosis

Experimental protocol to study cell viability and apoptosis

Materials and equipment:

• Cell dissociation agent (e.g., trypsin) – for adherent cell lines only

• Cell culture medium containing FBS or trypsin inhibitors

• PBS or HBSS supplemented with calcium chloride

• Benchtop centrifuge

• Fluorescent annexin V probe

• Viability dye (e.g., propidium iodide)

• Flow cytometer

Experimental procedure

Cell viability experiments using flow cytometry are performed quite commonly. They are used to study the effect of different agents, e.g., cytotoxic drugs, on the cell viability of various cell lines. They can be used to establish the minimum required concentration of antibiotics causing cytotoxic effects, which is useful for stable cell generation using antibiotic markers. They are often used when studying apoptosis and other forms of cell death. Viability dyes are recommended in flow cytometry experiments performed on unfixed cells in order to exclude dead cells from the analysis.

Live cells and early apoptotic cells are impermeable to viability dyes because they have an integral cell membrane and therefore are negative (Figure 6). Annexin V binds to phosphatidylserine that is found on the outer layer of early apoptotic cells. Annexin V also binds to dead cells, where it binds to a pool of phosphatidylserine present on the outer and inner layers because of their compromised cell membrane integrity. Viability dyes are able to penetrate dead cells and bind to their DNA.

Figure 6. Analysis of cell viability and apoptosis by flow cytometry.

Experimental protocol to study cell viability and apoptosis

1. Adherent cells:

   1. Remove medium and wash cell monolayer with PBS

   2. Add dissociating agent (e.g., trypsin) and incubate at 37°C for 5 min or until cells detach from the cell culture dish.

   3. Add serum-containing medium or trypsin inhibitors to inactivate dissociating agent.

   4. Transfer cells into a microcentrifuge tube.

Suspension cells: Transfer cells into a microcentrifuge tube.

2. Pellet cells by centrifugation (300 G’s, 5 min at room temperature). Remove medium and resuspend in PBS or HBSS containing calcium ions.

Note: Annexin V requires calcium for interaction with phospholipids. Supplement buffers with calcium salts and avoid chelating agents such as EDTA or EGTA.

3. Count cells and take 1 million (10⁶) cells per condition:

   1. Unstained sample – for establishing autofluorescence levels

   2. Sample stained with a viability dye

   3. Sample stained with an annexin V probe

   4. Sample stained with a viability dye and an annexin V probe

Note 1: Annexin V and a viability dye have to have different excitation spectra in order to distinguish their staining. Please refer to section 4 for tips on multicolor design experiments.

Note 2: Consider including a positive control sample, where cells are treated with an agent inducing cell death to validate the performance of used probes.

4. Add fluorescently labeled annexin V and incubate for 15 min at room temperature.

Annexin V preferentially binds to phosphatidylserine, which is found on the inner layer of the plasma membrane in living cells. In early apoptotic cells, phosphatidylserine is translocated to the outer layer, making it accessible for annexin V binding.

5. Pellet cells by centrifugation (300 G’s, 5 min at room temperature). Remove medium and resuspend in PBS or HBSS.

6. Add a viability dye (e.g., propidium iodide) and incubate for 5-20 min at room temperature.

Propidium iodide (PI) is a DNA intercalating agent – it is used as a fluorescent dye that binds to DNA. Dead cells lose cell membrane integrity, which allows the dye to reach the nucleus and bind to nucleic acids.

7. Analyze cells on a flow cytometer.

Note: Do not wash cells prior to analysis to avoid washing out the viability dye accumulated in dead cells.