Dye dilution-based proliferation
assays [Wallace et al. (7
)] have proven extremely useful because of their ability to correlate a critical functional outcome (i.e., expansion of antigen-responsive cells) with earlier events. An important feature of the dye dilution technique is that division-dependent changes in the expression of cell surface markers, intracellular proteins, antigen binding, or other properties of interest can be readily quantified by flow cytometry. Among these dyes, the so-called general protein labels are reactive compounds that form random covalent bonds with amino groups on cellular proteins. Their advantages include rapid and simple use, bright fluorescence signals, and often the ability to clearly visualize distinct generations of daughter cells. Limitations arising from dye dilution can be proliferation-dependent or -independent.
The so-called general membrane labels are lipophilic compounds that integrate stably into the plasma cell membrane without binding covalently. They are simple to use and—unlike with general protein labels—no waiting period is required for the staining intensity to stabilize; also, bright staining can be achieved without altering protein (and cell) function. On the downside, dye labeling assays typically take longer to complete than the measurement of population frequencies based on early activation events because cells must be cultured sufficiently long for proliferation to occur. When using cryopreserved samples, acceptable results are obtained only when good recovery (>70%) and high viability are obtained; the discrimination of apoptotic and necrotic cells is essential. Data may be reported in descriptive ways, semiquantitatively (stimulation index, mitotic index) or quantitatively (proliferation index, precursor frequency, time course analysis).
Meier et al. reported on the use of the activation marker, CD40L (CD154) upregulation, for the identification of antigen specific CD4 T cells, for example, against the hexon protein of human adenovirus (8
). As expected, only very low proportions of reactive (i.e., CD4+,CD154+) T cells (typically <0.2%) were found. Nevertheless, the test was robust, sensitive, and had low intra-assay variability. Moreover, since CD40L up-regulation because of bystander activation is very low, this approach was more specific than the use of other activation markers such as CD25, CD134, CD69, CD137, or HLA-DR. Combination with intracellular cytokine staining (IL-2 and IFNγ) was possible.
Three presentations focused on the activation of CD8+ T cells. Selection of CD137+ cells after a short TCR-mediated antigen stimulation results in enrichment of the specific T cells regardless of individual functional characteristics, making it possible to study the complete reactive T-cell repertoire [Wölfl et al. (9
)]. By virtue of this characteristic, CD137 analysis provides a more comprehensive detection method of activated CD8+ T cells than analysis for the production of a single cytokine. The low-frequency CD8+, CD137+ T cells can be selected either by flow-sorting or by the use of paramagnetic beads. CD137 permits better discrimination of positive (activated) cells than CD69 due to a higher and more robust expression level, and is more specific than CD25, which is often already detectable at an intermediate expression level on cultured T cells without recent stimulation through the TCR.
The main killing pathway of CD8+ T and NK cells makes use of preformed granules containing lytic proteins—mainly perforins and granzymes—that are secreted by exocytosis following target recognition. The secondary pathway involves the engagement of receptors on target cells that induce classical caspase-dependent apoptosis, such as those specific for Fas ligand. Perforin is the key player in the arsenal that CD8+ T cells use to kill target cells. Historically, measurement of perforin as marker for the antiviral capacity of CTL has largely been limited to staining for baseline levels in unstimulated cells using a mAb recognizing the granule-associated conformation of perforin (clone δG9). Recently, the B-D48 clone, raised against recombinant perforin, was discovered that detected perforin in multiple forms. Using a combination of these 2 antibodies, Hersperger et al. (10
) observed that CD8+ T cells can rapidly up-regulate perforin production following target cell recognition without the need for cellular proliferation. This up-regulation can be detected by flow cytometry, preferably in combination with another functional parameter to accurately assess de novo perforin synthesis.
The fluorescent antigen-transfected target cell (FATT)-CTL assay [Van Baalen et al. (11
)] combines nonradioactive quantification of target cell killing with an efficient and versatile plasmid transfection technology to achieve transient antigen expression in target cells. Plasmid vectors encoding antigen-green fluorescent protein (GFP) fusion proteins are used directly to electroporate target cells. Hence, no viruses, recombinant viral vectors, or radioactive isotopes are needed to generate target cells that present naturally processed epitopes. The elimination of antigen-GFP expressing cells is then enumerated flow cytometrically. This cytotoxicity test can be directly combined with effector cell markers, including degranulation or production of perforin, granzymes, or cytokines. One advantage specific for the FATT-CTL assay is the direct coupling between expression of antigens and fluorescent markers. This characteristic increases the dynamic range of the assay and the sensitivity due to the possibility to focus on target-cell killing among the cells that express the antigen. The FATT-CTL assay uses target cells presenting naturally processed epitopes. It measures, therefore, in a single assay, the successful completion of all the events required for antigen-specific cell-mediated cytotoxicity and is a convenient and versatile alternative to the classic 51