Fluorescence activated cell sorting (FACS) and TUNEL-assay

Flow cytometry or fluorescence-activated-cell-sorting (FACS) provides a highly sensitive method to analyse multiple characteristics of single cells. The information obtained is both qualitative and quantitative. Applications of FACS analyses and cell sorting in biochemical and biomedical fields are numerous. Conjugation of fluorescent dyes to ligands and to mono- or polyclonal antibodies has facilitated the study of density and distribution of cell-surface and cytoplasmic determinants as well as receptors, but also the identification of certain sub-populations of cells. Moreover, a wide range of fluorescent probes, such as propidium iodide, phycoerythrin or flourescein, is available for directly measuring cellular parameters, such as nucleic acid content, pH, intracellular calcium flux, enzyme activity or membrane potential (Carter and Ormerod, 2003⁷).

7 N.P. Carter and M. G. Ormerod, “Introduction to the principles of flow cytometry”, in Flow Cytometry, chap. 1, ed. M. G. Ormerod (Oxford, UK, 2003), pp. 1-22.

Flow cytometry measures optical and fluorescence features on single cell level. Physical properties are measured in two different modi, firstly by the forward angle light scatter, with the help of which cell sizes can be determined, secondly by the right-angle scatter, through which granularity and internal complexity of a cell are displayed.

Cells labelled in any of the ways described above, are passed by a light source, where the bound fluorescent molecules are excited to a higher energy state. The fluorochromes, once they return to their resting state, emit light energy at higher wavelengths. The fact that numerous fluorochromes show similar excitation wavelengths but different emission wavelengths, allows several cell properties to be measured at the same time.

During the actual screening process inside a flow cytometer, cells in suspension are drawn into a stream which is caused by a surrounding sheath of isotonic fluid that generates a laminar flow. The flow guarantees that only single cells pass through the interrogation point, where via a beam of monochromatic light, usually generated by a laser, the cells are intersected. Excited fluorochromes then emit light into all directions, which is collected by optics. These direct the light to a series of filters and dichroic mirrors, which isolate particular wavelength bands. All light signals are then detected by a system of photomultiplier tubes and digitised for computer analysis (Brown and Wittwer, 2000). Results are generally presented in histograms and two-dimensional dot-plots.

Chromatin condensation and cleavage of apoptotic linker DNA between nucleosome core particles are biochemical hallmarks of apoptosis. Especially the latter is counted for one of the later processes in apoptosis, which is caused by the activation of DNA endonucleases.

These cleave the higher order chromatin structure first into medium size fragments of ~300 kb and later on into small pieces of 50 bp in length. A general, very efficient method to detect DNA fragmentation is the TUNEL-assay (terminal deoxynucleotidyltransferase dUTP nick end labelling), a reaction that is catalysed by exogenous TdT.

For the analysis of H2AX serine 139 phosphorylation in HL-60 cells after the onset of apoptosis with relation to DNA double strand breaks and fragmentation, 1×10⁶ cells were treated with topotecan^® and incubated for 8h. Untreated cells served as controls. After incubation, the cells were washed twice with 3 ml sterile PBS and centrifuged for 5 min at 500×g in a Heraeus centrifuge (4°C). The supernatant was carefully discarded and the cell pellet was suspended in the remaining fluid. The cell suspension was injected with a syringe (gauge 0.4 mm) into a FACS tube containing 3 ml of 100% ice-cold ethanol and stored overnight at -20°C. Next, the tubes were centrifuged at 500×g for 5 min. The ethanol was quickly discarded and the cells were washed twice in PBS/0.5% BSA wash buffer and centrifuged once again at 500×g for 5 min at 4°C. Subsequently, the cells were incubated with 100 µl of the primary antibody anti-H2AX-phosphate S139 (2µg/ml) overnight with gentle agitation at 10 min intervals. Control cells were incubated in parallel in PBS/0.5% BSA wash

buffer. After that all samples were washed twice with PBS/0.5% BSA wash buffer and centrifuged at 500×g for 5 min (4°C). The supernatant was quickly removed and the cells were incubated with 100 µl of the secondary antibody AlexaFluor488 anti-mouse antibody (1:1,000) for 1h at 37°C. In a last step the cells were washed and centrifuged twice again as described and finally suspended in 500 µl sterile PBS.

For the TUNEL-assay 1×10⁶ cells were harvested and washed as described. Afterwards the samples were stained according to the protocol of the APO-DIRECT^™ KIT from BD Bioscience. The following table shows the contents of the master mix for a single sample.

staining solution 1 assay [µl]

reaction buffer 10.00

TdT enzyme 0.75

FITC-dUTP 8.00

distilled H2O (HPLC grade) 32.25

total volume 51.00

A volume of 51 µl was added to the samples that were to be analysed via TUNEL assay. The cells were incubated for 1h at 37°C. In a following step, the suspension was centrifuged at 500×g for 5 min (4°C) and the supernatant was carefully removed. The cells were washed twice with 1 ml rinse buffer provided from the KIT, centrifuged and finally suspended in 500 µl PBS. Cells just treated with rinse buffer served as control cells. All samples were analysed in the flow cytometer. The following table gives a list of all samples prepared:

FACS samples A) control cells

B) cells incubated with H2AX-phosphate S139 antibody and AlexaFluor555 labelled secondary antibody

C) cells stained with a FITC-labelled anti-BrdU monoclonal antibody (TUNEL-assay) and H2AX-phosphate S139 antibody with an AlexaFluor555 labelled secondary antibody D) cells stained with a FITC-labelled anti-BrdU monoclonal antibody (TUNEL-assay)

The results were displayed in histograms and two-dimensional dot-plots.

3 Results

The aim of this project was in the first place, to analyse the molecular basis and effects of the apoptosis-related chromatin condensation and fragmentation. To further elucidate this matter, putative apoptosis-related changes in core histone modifications, as well as histone modifications in fragmented and non-fragmented regions of the chromatin should be analysed. Moreover, the release of histones from fragmented chromatin was also investigated.

3.1 Monitoring the ongoing process of apoptosis: apoptosis markers for HL-60 cells