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Cytometry A. Author manuscript; available in PMC Nov 17, 2009.
Published in final edited form as:
PMCID: PMC2778017
Measuring Antigen-Specific Immune Responses: 2008 Update
Jan W. Gratama,1* Florian Kern,2 Fabrizio Manca,3 and Mario Roederer4
1Erasmus MC, Rotterdam, The Netherlands
2Charité, Universitätsmedizin, Berlin, Germany
3G. Gaslini Institute, Genoa, Italy
4Vaccine Research Center, Bethesda, Maryland
* Correspondence to: Jan W. Gratama, Department of Medical Oncology, Erasmus MC—Daniel den Hoed Cancer Center, P.O. Box 5201, 3008 AE ROTTERDAM, The Netherlands. j.w.gratama/at/
“Measuring antigen-specific immune responses” (MASIR) is the theme of a series of small conferences (approximately 200 participants) dedicated to a broad range of topics tied together by the common need for quantifying and characterizing antigen-specific lymphocyte responses. The overall goal of the MASIR conferences is to define the antigen-specific assays that can provide clinical correlates of vaccine efficacy, disease morbidity, or treatment efficacy. Previous MASIR conferences were held in 2005 in Courmayeur (Italy) and in 2006 in Santorini (Greece); reviews of the state-of-the-art of the field as presented at these conferences were published (1,2). This issue of Cytometry (Part A) presents 16 papers selected from the third MASIR conference held in early 2008 in La Plagne (France). These papers deal with basic methods (intracellular staining, MHC multimers), “live cell” assays, as well as applications and scientific outcomes (specifically, epitope mapping, and T-cell subsets).
Intracellular cytokine staining (ICS), also known as “cytokine flow cytometry,” combined with antigen-specific T-cell stimulation is a widely applied technique that lends itself well to the multiparameter analysis of complex cellular samples; Nomura et al. addressed the use of polychromatic flow-cytometry (i.e. using more than five fluorescence colors) in a multicenter setting (3). In situations where multiple cytokines and surface markers need to be studied in parallel—the so-called “immune function signatures” in vaccination studies—ICS is superior to the ELISPOT technology (which can, however, be performed at lower costs and is amenable to high throughput screening). In the multicenter studies reviewed by Nomura, standardized reagents were used, such as cryopreserved cells, antibodies, beads, and peptides. As would be expected in this situation, the main sources of variation between sites were differences in instrument set-up and gating.
Class I MHC tetramers entered the cytometry arena in 1996, whereas class II MHC multimers did so in 2002. Brooimans et al. (4) describe the standard assay approach for PE-conjugated tetramers carrying CMV epitopes from pp50 and pp65. Under routine conditions, it is possible to detect CMV-specific CD8+ T cells at frequencies as low as 1 cell/μl. Chattopadhyay et al. (5) focused on situations in which low-frequency CD8+ T cells bind to cognate antigen with low avidity. Here, discrimination between signal and noise can be improved by the use of a “dump channel” (i.e., the use of specific stains to identify confounding events to be excluded from analysis), or the use of the same MHC multimer reagents labeled with different fluorochromes. Signal amplification can be achieved by pretreatment of the cells with the protein kinase inhibitor, dasatinib, or the use of class I MHC multimer reagents that bind CD8 with enhanced affinity.
Similar requirements, that is, to reduce nonspecific staining in the setting of rare event detection, apply to class II MHC tetramers as outlined by Cecconi et al. (6). Typically, antigen-specific CD4+ T cells occur in peripheral blood in frequencies much less than 0.2%. The staining efficiency of class II MHC tetramers is strongly correlated to the TCR affinity for its cognate peptide-class II MHC complex. This situation is in particular important for the ability of class II MHC tetramers to identify CD4+ T cells specific for autoantigens and cancer, which are mostly characterized by the expression of low-affinity TCR. Of note, CD4+ T-cell activation, rather than the overall quantitative display of TCR molecules on the cell membrane, determines the accessibility of the TCR to class II MHC tetramers. This accessibility further relies on active cell metabolism and membrane trafficking, as indicated by the requirement for a temperature of 37°C to achieve optimal staining results.
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 51Cr-release assay.
The breadth of an antigen-specific T- or B-cell response to a pathogen or vaccine can be measured by mapping T- or B-cell epitopes. Significant efforts have been made to improve epitope mapping in basic research and clinical settings. The Immune Epitope Database and Analysis Resource (IEDB) catalogues antibody and T-cell epitopes derived from infectious agents, allergens, and autoantigens in a diverse range of hosts. Vita et al. described how this complex set of data is curated to allow continued growth and adaptation (12). This project has been charged to a team of doctoral level curators that provides quality control and insight under the support of a “curation manual,” the latter being a reference in living document format designed to assist in accurate and consistent data curation.
Mapping a complete set of epitopes from a vaccine or an entire pathogen requires a complex method for screening hundreds to thousands of peptides with a limited amount of donor sample. Commonly, mapping is performed using a functional readout (typically ELISPOT) following stimulation with overlapping peptides (or pools of such peptides) that span a given protein and presumably encompass all possible T-cell epitopes within that protein. This approach is rapid, powerful, efficient, and unrestricted by HLA type; however, the process, termed “deconvolution,” requires a large amount of sample to perform. Precopio et al. (13) present an optimized deconvolution process to identify peptide epitopes in a robust fashion, using a minimum amount of sample material.
In search of T-cell epitopes for the onconeural protein HuD which is associated with paraneoplastic neurological syndrome, De Graaf et al. (14) used the same principle only to find reproducible false-positive CD8+ T-cell responses in three of 127 individuals, which all shared HLA-A*2402 and HLA-B*1801. The same three 15-mers yielded the CD8+ T-cell response in the three individuals. This highly unusual result could not be reproduced when using new batches of peptide with a higher level of purity. Mass spectroscopical analysis of the 15-mers showed the presence of a cytomegalovirus (CMV)-encoded peptide as a contaminant. The three responding individuals were all CMV-seropositive and positive for HLA-B*1801. These findings illustrate that synthetic protein-spanning peptide pools must be handled with great care both by the manufacturer and the end-user to avoid contamination with other peptides that may be present in small amounts on reusable equipment.
Moody and Haynes (15) addressed epitope-specific B-lymphocyte responses, as studied testing secreted antibodies and via the B-cell surface receptor (BCR). A crucial point for the study of secreted antibodies is that all antibody isotypes have at least two antigen recognition sites per molecule, on the basis of which antibodies have the ability to form immune complexes. ELISPOT assays permit the quantitation of antigen-specific B cells and form a bridge between studies of secreted antibodies and individual B cells. For BCR-based studies of B cells, techniques fall into three broad categories: (i) haptens on carriers; (ii) labeled proteins or whole organisms; and (iii) epitopes presented by a display system. In all cases, a detection reagent is used to label or capture the B cell by using the cell surface-bound immune complex.
Willemsen et al. (16) made use of the selection power of phage display technology allowing the screening of tens of billions of individual clones by high-throughput selection for antigen receptors with binding capacity for the peptide-MHC complex. This strategy was used to select antibody-type receptors for a panel of class I and II-restricted tumor-specific epitopes to be used for immuno-gene therapy of cancer.
Five speakers [Appay, Van Lier, Sallusto, and Roederer (17), as well as Fuhrmann (18)] addressed the complicated and confusing issue of dividing antigen dependent T-cell development into compartments characterized by certain T-cell phenotypes and functions. Our knowledge of T cells has increased tremendously with the ever-increasing numbers of cell surface and intracellular markers along with the progress of flow cytometry. Nevertheless, recent studies have generated more questions than answers. There is consensus that T cells are heterogenous, but there is no harmonization of nomenclature and the use of markers to define T-cell subsets may further contribute to confusion. Although there may be as many T-cell subsets as there are marker combinations, the CD4+ and CD8+ T cells are considered as the main subsets, on which CD45RA and CD45R0 are involved in activation, CD27 and CD28 in costimulation and PD-1 in regulation. There are similarities between CD4+ and CD8+ T cells in phenotype, functional attributes, telomere shortening, and gene expression patterns, but CD4+ T-cell differentiation is viewed as much more complex than that of CD8+ T cells with the existence of TH1, TH2, TH17, and regulatory T cells within the CD4+ subset. Also, different viral pathogens are associated with the predominance of distinct T-cell response profiles, such as for HCV, EBV, HIV, and CMV based on the coexpression of CD27, CD28, and CCR7. Distinct T-cell profiles may reflect the differential requirements for cellular immune responses to control these pathogens.
T cells also exhibit complex functional responses; the combinatorial expression of different T-cell functions is referred to as the “quality” of the T-cell response (19). Importantly, good T-cell quality is associated with clinical relevant parameters such as delay of HIV progression or excellent control of latent infection (CMV), in contrast to measurements of the magnitude or phenotype of the T-cell response, which do not have clinical correlates. T-cell characteristics may change upon activation and inflammation, which leads to alterations in the T-cell environment and hence on their behavior in terms of homing and migration. A number of inconclusive points still remain, such as knowledge of the pathway of T-cell differentiation (is it linear, branched, reversible, or a combination?). Although murine models have allowed the performance of longitudinal studies, such studies are scarce in humans but are important to get a more comprehensive insight into T-cell immunity. As pointed out by Kern, changes in the phenotype of antigen specific T-cells following persistent or repeated activation after infection for example might have potential for new diagnostic tests (18,20).
Overall, the last 10 years have seen an explosion in the field of antigen-specific immune response monitoring. As summarized in this issue of Cytometry and at the MASIR conferences, these technologies have provided new insights into the basic biology of the immune system and are beginning to provide useful clinical information. These papers highlight the areas that are being actively explored.
1. Kern F, Li Pira G, Gratama JW, Manca F, Roederer M. Measuring Ag-specific immune responses: understanding immunopathogenesis and improving diagnostics in infectious disease, autoimmunity and cancer. Trends Immunol. 2005;26:477–484. [PubMed]
2. Li Pira G, Kern F, Gratama JW, Roederer M, Manca F. Measurement of antigen specific immune responses: 2006 update. Cytometry Part B. 2007;72B:77–85. [PubMed]
3. Nomura L, Maino VC, Maecker HT. Standardization and optimization of multipara-meter intracellular cytokine staining. Cytometry Part A. 2008;73A(this issue):984–991. [PubMed]
4. Brooimans RA, Boyce CS, Popma J, Broyles DA, Gratama JW, Southwick PC, Keeney M. Analytical performance of a standardized single-platform MHC tetramer assay for the indentification and enumeration of CMV-specific CD8+ T lymphocytes. Cytometry Part A. 2008;73A(this issue):992–1000. [PubMed]
5. Chattopadhyay PK, Melenhorst JJ, Ladell K, Gostick E, Scheinberg P, Barrett JA, Wooldridge L, Roederer M, Sewell AK, Price DA. Techniques to improve the direct ex vivo detection of low frequency antigen-specific CD8+ T cells with peptide-major histocompatibility complex class I tetramers. Cytometry Part A. 2008;73A(this issue):1001–1009. [PMC free article] [PubMed]
6. Cecconi V, Moro M, Del Mare S, Dellabona P, Casorati G. Use of MHC Class II tetramers to investigate CD4+ T cell responses: problems and solutions. Cytometry Part A. 2008;73A(this issue):1010–1018. [PubMed]
7. Wallace PK, Tario JD, Jr, Fisher JL, Wallace SS, Ernstoff MS, Muirhead KA. Tracking antigen-driven responses by flow cytometry: Monitoring proliferation by dye dilution. Cytometry Part A. 2008;73A(this issue):1019–1034. [PubMed]
8. Meier S, Stark R, Frentsch M, Thiel A. The influence of different stimulation conditions on the assessment of antigen-induced CD154 expression on CD4+ T cells. Cytometry Part A. 2008;73A(this issue):1035–1042. [PubMed]
9. Wölfl M, Kuball J, Eyrich M, Schlegel PG, Greenberg PD. Full repertoire of CD8+ T cells without the need to know epitope specificities using CD137. Cytometry Part A. 2008;73A(this issue):1043–1049. [PMC free article] [PubMed]
10. Hersperger AR, Makedonas G, Betts MR. Flow cytometric detection of perforin upregulation in human CD8 T cells. Cytometry Part A. 2008;73A(this issue):1050–1057. [PubMed]
11. van Baalen CA, Gruters RA, Berkhoff EGM, Osterhaus ADME, Rimmelzwaan GF. Cytometry Part A. 2008;73A(this issue):1058–1065. [PubMed]
12. Vita R, Peters B, Sette A. The curation guidelines of the immune epitope database and analysis resource. Cytometry Part A. 2008;73A(this issue):1066–1070. [PMC free article] [PubMed]
13. Precopio ML, Butterfield TR, Casazza JP, Little SJ, Richman DD, Koup RA, Roederer M. Optimizing peptide matrices for indentifying T-cell antigens. Cytometry Part A. 2008;73A(this issue):1071–1078. [PMC free article] [PubMed]
14. de Graaf MT, de Beukelaar JW, Burgers PC, Luider TM, Kraan J, Sillevis Smitt PA, Gratama JW. Contamination of synthetic HuD protein spanning peptide pools with a CMV-encoded peptide. Cytometry Part A. 2008;73A(this issue):1079–1085. [PubMed]
15. Moody MA, Haynes BF. Antigen-specific B cell detection reagents: Use and quality control. Cytometry Part A. 2008;73A(this issue):1086–1092. [PMC free article] [PubMed]
16. Willemsen R, Chames P, Schooten E, Gratama JW, Debets R. Selection of human antibody fragments directed against tumor T-cell epitopes for adoptive T-cell therapy. Cytometry Part A. 2008;73A(this issue):1093–1099. [PubMed]
17. Appay V, van Lier RAW, Sallusto F, Roederer M. Phenotype and function of human T lymphocyte subsets: consensus and issues. Cytometry Part A. 2008;73A(this issue):975–983. [PubMed]
18. Fuhrmann S, Streitz M, Kern F. How flow cytometry is changing the study of TB immunology and clinical diagnosis. Cytometry Part A. 2008;73A(this issue):1100–1106. [PubMed]
19. Seder RA, Darrah PA, Roederer M. T-cell quality in memory and protection: implications for vaccine design. Nat Rev Immunol. 2008;8:247–258. [PubMed]
20. Streitz M, Tesfa L, Yildirim V, Yahyazadeh A, Ulrichs T, Lenkei R, Quassem A, Liebetrau G, Nomura L, Maecker H, Volk HD, Kern F. Loss of receptor on tuberculin-reactive T-cells marks active pulmonary tuberculosis. PLoS ONE. 2007:e735, 1–10. [PMC free article] [PubMed]