Analysis of cell cycle and apoptosis

Protocol for the detection of apoptotic cells during cell cycle analysis.

This method is described in the publication by Riccardi and Nicoletti, ‘Analysis of apoptosis by propidium iodide staining and flow cytometry’ (Nature Protocols, 2006).

Principle of the method: apoptotic cells are characterized, amongst other things, by features such as fragmented DNA and, consequently, a reduction in nuclear DNA content. The fluorochrome propidium iodide (PI) binds (almost!) stoichiometrically to nucleic acids, so that the PI fluorescence is proportional to the cellular DNA (and RNA, if not removed) content.

The hypotonic, surfactant-containing Nicoletti buffer (containing sodium citrate and the detergent Triton X-100) causes lysis of the cell membrane, but not of the nuclear membrane. Due to the hypotonic shock, most of the RNA is removed, thereby circumventing the problem of RNA staining. PI staining ‘according to Nicoletti’ thus enables rapid and precise analysis of cellular DNA content as well as the identification of hypodiploid cell nuclei by flow cytometry.

When measuring PI fluorescence in terminally differentiated cells (such as human primary neutrophils or differentiated neutrophil-like HL-60 cells) by flow cytometry, two peaks appear in the histogram:

•   hypodiploid DNA peak (a broad, weaker-fluorescing ‘sub-G1 peak’, characteristic of apoptotic cells)

•   diploid DNA peak (narrower, more intensely fluorescent G1 peak, characteristic of intact cells).

When measuring PI fluorescence in proliferating cells (such as immortalized human HL-60 myeloblasts/promyelocytes from acute myeloid leukaemia) by flow cytometry, the histogram shows not only the hypodiploid DNA ‘sub-G1 peak’ – characteristic of apoptotic cells – but also G1, S and G2 peaks, characteristic of dividing cells in the cell cycle.

Here is the step-by-step protocol for Nicoletti staining (direct DNA staining with PI in a hypotonic solution, "quick method”):

• Induce apoptosis with the desired substances for a specific duration 

• Once the incubation period has elapsed, transfer approximately 5-10 × 10⁵ cells from the wells into the FACS tubes. 

Non-adherent cells (such as neutrophils or myeloid cell line HL-60) do not need to be detached from the surface. By gently pipetting up and down, the cells in the wells can be brought into suspension and transferred to the FACS tubes.

Adherent cells (such as keratinocyte cell line HaCaT) must be gently detached from the surface, for example by incubation with mild trypsin (0.05%) containing EDTA. Before adding trypsin, collect the old medium from the wells (can contain apoptotic cells!), wash the cells briefly with PBS and incubate them with trypsin at 37 °C for as short a time as possible (approx. 5 min). Stop the reaction with cell culture medium or PBS containing FCS, and transfer the cell suspension (together with the old medium that has been collected from the wells) into the FACS tubes.

• Centrifuge the cell suspension in the FACS tubes for 5 minutes at 390x g at room temperature (18-23 °C), then decant the supernatant

• Gently resuspend the pellets in 300 µl Nicoletti buffer (containing 0.1% sodium citrate, 0.1% Triton X-100 and 50 µg/ml propidium iodide), then vortex

• For optimal staining, incubate the cells for at least 1 hour (maximum 4 hours) at 4 °C in the dark

• Then carry out FACS analysis immediately, measuring at least 10.000 cells per sample.

An example of flow cytometric measurement and the gating strategy can be found in Fig. 1e–f of the publication “Differentiation of HL-60 cells into primed neutrophils for the evaluation of antiapoptotic effects of poorly soluble nanoparticles”, 2005 PLoS ONE.