The assessment of antigen-specific CD8+ T cells through the combination of MHC class I tetramer and intracellular staining

Journal of Immunological Methods 268 (2002) 9 – 19 www.elsevier.com/locate/jim

Review

The assessment of antigen-specific CD8+ T cells through the combination of MHC class I tetramer and intracellular staining Victor Appay *, Sarah L. Rowland-Jones MRC Human Immunology Unit, Weatherall Institute of Molecular Medicine, John Radcliffe Hospital, Headington, Oxford OX3 9DS, UK Received 14 October 2001; accepted 14 January 2002

Abstract Peptide-bound histocompatibility leukocyte antigen (HLA) class I tetramers enable a precise identification of antigenspecific CD8+ T cells using flow cytometry. The combination of this technology with intracellular staining techniques opens up significantly better ways of studying these cells than previously possible, allowing immunologists to look at their life cycle (activation and proliferation), manner of death (aging and apoptosis) and effector function (cytotoxic potential and cytokine production). In this review, we hope to provide an overview of these possibilities, as well as making specific suggestions about the use of intracellular staining techniques in the study of antigen-specific T cells. Understanding how antigen-specific cells develop and function in different circumstances and pathologies will be the key to unravelling the secrets of our cellular immune system. D 2002 Elsevier Science B.V. All rights reserved. Keywords: Tetramers; CD8+ T cells; Intracellular; Function

1. Introduction The introduction of peptide– major histocompatibility complex (MHC) class I tetrameric complex technology initiated a profound revolution in the field of cellular immunology (Altman et al., 1996). In 1996, after more than 20 years of abortive efforts to measure

Abbreviations: BrdU, bromodeoxyuridine; CFSE, carboxyfluorescein diacetate succinimidyl ester; FITC, fluorescein; FISH, fluorescence in situ hybridization; HIV, human immunodeficiency virus; HLA, histocompatibility leukocyte antigen; GMP, granule membrane protein; ICS, intracellular cytokine staining; MHC, major histocompatibility complex; PMA, phorbol myristate acetate. * Corresponding author. Tel.: +44-1865-222-312; fax: +441865-222-502. E-mail address: [email protected] (V. Appay).

the CD8+ T cell response with precision, immunologists at last had at their disposal a tool which enabled a direct visualization of antigen-specific CD8+ T cells by flow cytometry. CD8+ T cells are one of the major components of cell-mediated immunity and play a key role in the elimination of virus-infected, tumour and allograft cells. The analysis of tetramer-positive CD8+ T-cells has provided a method that reliably quantitates the number of specific CD8+ T cells present in peripheral blood and secondary lymphoid organs (Ogg et al., 1998; Romero et al., 1998). The benefit of such a technology is not only limited to the precise quantification and cloning of antigen-specific T cells. The next step in the use of tetramers is the detailed analysis of the characteristics of the antigen-specific T-cell population. Through the reliable identification of antigen-specific CD8+ T

0022-1759/02/$ - see front matter D 2002 Elsevier Science B.V. All rights reserved. PII: S 0 0 2 2 - 1 7 5 9 ( 0 2 ) 0 0 1 9 5 - 3

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cells, tetramers offer the possibility of obtaining additional information about their phenotype through costaining for cell surface markers, including markers of activation (Callan et al., 1998; Murali-Krishna et al., 1998), T-cell receptor Vh usage (Wilson et al., 1998), and other differentiation and homing markers (e.g. CD family and chemokine receptors) (Faint et al., 2001). However, a cell is not characterized so much by what lies on its surface, but by what resides inside, where most of its biology takes place and which is likely to reflect its functional phenotype. The study of the functional state of T cells is crucial to understand T-cell responses against pathogens or self-antigens. As flow cytometry technology has advanced to include such developments as multiparameter analysis and techniques to look at intracellular features (Koester and Bolton, 2000), it is now possible to assess practically all aspects of the life of the cell. The combination of intracellular labeling techniques with the use of tetramers enables us to characterize in detail the function of antigen-specific CD8+ T cells. This review provides instances where these techniques were or could be successfully used in human or murine studies, with the aim of providing an overall picture of the possibilities that exist using these technologies to assess the features of antigenspecific CD8+ T cells.

2. Studying the features of antigen-specific CD8+ T cells 2.1. Proliferation Proliferation is the principal consequence of cellular activation; in vivo, infection by pathogens leads to a marked expansion of T cells. Assessing the proliferation of antigen-specific CD8+ T cells is essential to understand the dynamics of the T-cell response, and to examine questions of the relationship between bystander and specific T-cell activation. For many years, the incorporation of 3H-thymidine during DNA replication has been the gold standard to monitor proliferation of T cells. However, this technique is limited to the measurement of bulk cell division, a limitation overcome by flow cytometry, which enables analysis at a single-cell level. Recently, new flow cytometry-based methods have been successfully

developed to look at cellular proliferation. Similar to the incorporation of 3H-thymidine, proliferating lymphocytes can directly incorporate bromodeoxyuridine (BrdU), a thymidine analogue, which can be detected by flow cytometry using anti-BrdU antibodies following cell permeabilization (Gratzner, 1982; Tough et al., 1996). An important recent technical advance has been the possibility of monitoring cell division, for up to eight discrete cycles, by following the serial halving of the vital dye carboxyfluorescein diacetate succinimidyl ester (CFSE) (Lyons, 2000). CFSE can permeate membranes and lymphocytes can therefore be loaded into by simple incubation; moreover, normal cell function is maintained throughout the assay. Following diffusion into the intracellular environment, the molecule loses its permeant ability and becomes highly fluorescent. CFSE is then partitioned with remarkable fidelity between daughter cells: proliferation results in progressive dilution and ultimately extinction of the dye, which can be observed by flow cytometry (excited at 488 nm) with detection of green light-FL1. A major advantage of these two methods is that cell proliferation can be followed in vivo in mouse models: either by feeding mice with BrdU throughout the period of antigen challenge, or by transferring CFSE-loaded cells into mice. Using multiparameter FACS analysis, these two methods (BrdU and CFSE) can easily be combined with tetramer staining. The combination of tetramer staining and BrdU/CFSE labeling has recently been used in studies of the dynamics of virus-specific T-cell responses and of the establishment of memory cells (Doherty and Christensen, 2000; Sakai et al., 2001; Kaech and Ahmed, 2001). However, as these two methodologies both require the initial loading of cells with a label, they are limited to in vitro experiments in human studies. Other techniques have been developed to enable the assessment of T-cell proliferation ex vivo. They employ the measurement of intracellular markers that correlate with proliferation, such as DNA content (e.g. labeled with propidium iodine, which, when excited at 488 nm, emits red fluorescence that can be recorded on FL2), and proteins expressed differentially in cycling and noncycling cells [e.g. Ki67, proliferating cell nuclear antigen (PCNA), phosphorylated histone H3, cyclins] (Darzynkiewicz et al., 2001). The nuclear antigen Ki67 is one of the most commonly used

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markers to assess cell proliferation. Although its function remains unknown, its expression has been associated with cell cycling, increasing rapidly during the S and G2 phases, reaching a maximum during the M phase, so that Ki67 positive cells are frequently defined as proliferating cells (Gerdes et al., 1984). Following permeabilization, tetramer-positive T-lymphocytes can easily be stained with anti-Ki67 antibodies, thereby providing a measurement of the proliferation of antigen-specific CD8+ T cells, for example, during acute viral infection or viral reactivation (Oxenius et al., 2001; Champagne et al., 2001). 2.2. Apoptosis The fate of T cells is dictated by a complex combination of factors (such as the levels of activation, co-stimulation, stress and other environmental conditions), but in most cases, will ultimately end in apoptosis (programmed cell death) of the cells. For instance, the majority of antigen-specific T cells will undergo apoptosis following primary infection (Ahmed and Gray, 1996). CD8+ T cells can also apoptose following lysis of their targets (Xu et al., 2001). Understanding the balance between survival and apoptosis of antigenspecific CD8+ T cells is a central issue in immunology. In mammals, apoptosis is conducted through two main pathways. The first involves the engagement of death receptors such as Fas (also referred as CD95) through interactions with their ligands (e.g. Fas ligand) and has been referred to as ‘‘death by design’’ (Strasser et al., 2000). The second pathway, referred to as ‘‘death by neglect,’’ is governed by the Bcl-2 family, which includes both proapoptotic (e.g. Bax, Bik) and antiapoptotic (e.g. Bcl-2, Bcl-xL) members, promoting or preventing death signals from diverse cytotoxic stimuli (such as cytokine deprivation, DNA or mitochondrial damage) (Adams and Cory, 1998; Strasser et al., 2000). These two pathways lead to the activation of a range of caspases, which in turn triggers apoptosis. A number of methods exist for the detailed study of the induction of these pathways. A common way to look at apoptosis involves staining with annexin V. Early during apoptosis, intracellular phosphatidylserine becomes exposed at the cell surface and can be bound with high affinity by fluorochrome-conjugated annexin V (van Engeland et al., 1998). Staining with annexin V enables the iden-

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tification of apoptotic cells and can easily be combined with tetramer staining for ex vivo analysis (Tan et al., 1999; Oxenius et al., 2001). Additional methods have been developed to study the apoptotic pathways in more detail, and to gain information regarding the susceptibility of cells to apoptosis. Not only is it possible to measure the expression of cell surface death receptors such as CD95 ex vivo, but various intracellular mediators of the apoptotic pathway can also be studied. These include members of the Bcl-2 family, which can be stained using monoclonal antibodies following permeabilization of the cells. Recently, the antiapoptotic factor Bcl-2 was shown to be up-regulated in resting memory cells, suggesting that memory cells have a better capacity to survive (Grayson et al., 2000). Other possible options to assess apoptosis and survival reside in the staining of intracellular activated caspases, using anti-caspase antibodies (Campos et al., 1999) or fluorochromeconjugated caspase ligands or inhibitors (Bedner et al., 2000), and of apoptosis-associated proteins, such as the mitochondrial membrane protein APO2.7 (Koester et al., 1997). Although these latter methods may have particular relevance for our understanding of T-cell responses, they have not yet been described in combination with tetramer staining. 2.3. Activation and aging The activation status of antigen-specific CD8+ T cells ex vivo and in vitro can easily be monitored by staining for cell surface activation markers [e.g. CD69, CD38, Human leukocyte antigen (HLA) class II molecules] (Callan et al., 1998; Appay et al., 2000). Cellular activation results from ligand-receptor mediated signal transduction events, which affect the nuclear transcriptional machinery. The eventual consequences of activation can be as diverse as proliferation or apoptosis of the cells. Flow cytometry-based methods have been described which look at the signalling mechanisms and pathways (such as intracellular protein phosphorylation). For instance, following permeabilization, it is possible to stain lymphocytes with anti-phosphotyrosine antibodies (Muller et al., 1998) or antibodies specific for phosphorylated forms of signalling proteins, such as STAT (Fleisher et al., 1999). Although currently feasible, these methods should be used with caution in combi-

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nation with tetramer staining, as the use of tetramer staining rapidly triggers various signalling pathways (reviewed elsewhere in this issue), so could lead to erroneous conclusions. In certain conditions, chronic activation can drive the aging of lymphocytes, which may eventually reach a state of senescence (Globerson and Effros, 2000). Although there is little data available in this area, the aging of lymphocytes involved in antigenspecific CD8+ T-cell responses will be important to study. One possible way to address the aging of cells resides in the measure of telomere length (Effros and Pawelec, 1997). Cellular expansion, involving extensive numbers of divisions, results in shortening of hexameric DNA sequences found at the end of chromosomes referred to as telomeres. With each division, 50– 100 base pairs of telomeric DNA are lost due to the inability of DNA polymerase to replicate fully the extreme ends of chromosomes. Telomeres are responsible for maintaining chromosomal stability and integrity, and their shortening may eventually lead to cell cycle arrest and replicative senescence (Blackburn, 1991). The measurement of telomere length by flow cytometry is now possible by means of a recently developed technique, named flow-FISH (fluorescence in situ hybridization by flow cytometry) (Rufer et al., 1998, 1999). This technique involves the permeabilization of lymphocytes, followed by hybridization with fluorescein (FITC)-labeled telomere-specific probes, prior to FACS analysis. The problem in combining this method with tetramer staining resides in the need to heat the samples to 80 jC for 10 min during the hybridization, which presents difficulties regarding the stability of fluorochromes (other than FITC), and therefore reduces the potential for doing multicolor analysis. However, a recent study has described the measurement of telomere length associated with tetramer staining, by selecting a heat stable fluorochrome (Cy5) to label the tetramers (Plunkett et al., 2001). The increase numbers of fluorochromes and fluorochrome combinations available is likely to render possible the development of multicolor flow-FISH in the near future. 2.4. Effector function A major issue in the study of antigen-specific CD8+ T cells is the assessment of the effector char-

acteristics of these cells. Antigen-specific CD8+ T cells can mobilize two main effector mechanisms: cytolysis of infected or malignant cells and production of cytokines, chemokines and microbicidal molecules. The exploration of their capacity to execute these functions is particularly important in pathologies in which failure of immune surveillance is postulated to be significant (e.g. human immunodeficiency virus [HIV] infection and melanoma). Antigen-specific CD8+ T cells mediate cytotoxicity through two main pathways: the perforin-dependent cytotoxic pathway and the Fas-dependent pathway (Trapani et al., 1999). Perforin is a 70-kDa protein contained in lytic granules, which forms pores in the target cell membranes, enabling the entry of granzymes, which activate an apoptotic cascade resulting in cell death (Liu et al., 1995; Smyth and Trapani, 1995). Both perforin and granzymes are found preformed inside the cells and can easily be stained ex vivo by flow cytometry following permeabilization, in combination with tetramer staining (Appay et al., 2000; Chen et al., 2001; Gamadia et al., 2001). The presence of these cytotoxic proteins provides an indication of the cytotoxic potential of CD8+ T cells. It is also possible to monitor the presence of lytic granules using monoclonal antibodies specific for TIA-1 (also called GMP-17), a protein found in the granule membrane (Medley et al., 1996). CD8+ T cells can also mediate cytotoxicity through the Fas/FasL pathway: FasL (Fas ligand) present on the CD8+ T-cell surface binds to target cell surface death receptor Fas (or CD95), which induces apoptosis of the target cell. FasL has also been shown to be located in lytic granules (Bossi and Griffiths, 1999). Although fluorochrome-conjugated anti-FasL antibodies are available, the ex vivo measure of FasL by flow cytometry remains difficult, likely to be due to a lack of sensitivity of the method or low levels of expression in the cells. CD8+ T cells produce also various soluble factors, including cytokines and chemokines, which play an active role in the immune response. For instance, IFNg plays an important role in the induction of cellular antiviral proteins (Guidotti and Chisari, 1996) and through its ability to activate macrophages (Dayton et al., 1985); IL-2 is a strong inducer of T-cell proliferation; and TNF-a is able to inhibit viral gene expression and replication (Guidotti and Chisari, 1996). The CC-chemokines RANTES, MIP-1a and MIP-1h

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coordinate chemoattraction of lymphocytes and macrophages to the infection site, and have direct inhibitory effects against HIV infection (Cocchi et al., 1995). CD8+ T cells do not contain preformed cytokines, therefore, direct ex vivo intracellular staining is not possible. However, CD8+ T cells are able to produce cytokines rapidly upon stimulation (Slifka et al., 1999), and over the past few years reliable methods have been developed to measure intracellular cytokines by flow cytometry (Pala et al., 2000). These methods require stimulation of the lymphocytes and their incubation for a few hours in the presence of a protein transport inhibitor (such as Brefeldin A and Monensin), so that the cytokines produced are retained inside the cells and can therefore be stained using anti-cytokine antibodies after permeabilization. Intracellular cytokine staining (ICS) for IFN-g, TNFa, IL-2 and MIP-1h has been successfully combined with tetramer staining (Appay et al., 2000; GeaBanacloche et al., 2000; Kostense et al., 2001; Sandberg et al., 2001). The production by tetramer-positive cells of other cytokines (e.g. IL-4, IL-10 or IL-13) has not yet been described, probably because of the difficulty of detecting these cytokines by intracellular staining, as well as the low frequency of CD8+ T cells producing them. Although this technique does not permit ex vivo staining, it is currently the only way to measure cytokine production in tetramer-positive cells. The novel FlowMetrix system (Vignali, 2000) does not allow cytokine-producing populations to be distinguished, while the detection of secreted cytokines by flow cytometry does not appear to be particularly reliable (Asemissen et al., 2001).

3. Experimental recommendations 3.1. General approach The general protocol used in our laboratory is presented in Appendix A. Intracellular staining requires fixation of the cells (to maintain structural integrity) before permeabilization (to allow entry of antibodies). The levels of tetramer staining that can be achieved on fixed cells are significantly reduced and so this step should be completed before fixation and permeabilization of the samples. The choice of an intracellular antigen to study should not necessarily be

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limited by the availability of monoclonal antibodies already tested for flow cytometry. The extended selection of antibodies specific for intracellular antigens used in immunoblotting or immunochemistry opens up abundant possibilities for flow cytometry; once these antibodies are optimized. This wide variety of potential intracellular antigens for study also implies that a diversity of intracellular locations can be assessed, and therefore, differences in permeabilization efficiency according to the fixation/permeabilization agent should be considered. This point has been discussed recently by Koester and Bolton (2000). Incomplete permeabilization can lead to the obtaining of false negatives and incorrect results, so the study of a completely new cellular antigen should be approached with caution. In our experience to date (measuring intracellular IFNg, TNFa, MIP-1h, IL-2, perforin, granzymes A and B, GMP-17, Ki-67, Bcl-2 and CD3~ in tetramer-positive cells), the FACSk permeabilizing solution from Becton Dickinson has generally led to satisfactory results. The determination of correct specific fluorescence staining limits (negative – positive, low – high) is an important point in intracellular staining, especially considering the observation of small populations such as tetramer-positive cells. This can be facilitated by the comparison of positive and negative controls, and the addition of isotype controls in the assays. However, the use of isotype controls is not always of great value (Baumgarth and Roederer, 2000) and certain stainings can present particular difficulties in interpretation (e.g. a range of levels of expression, small shifts between positive and negative expression). In these less obvious situations, the evaluation of antigen expression in a population can be facilitated by dissecting this population into several subsets by means of other markers, as exemplified in Fig. 1. Consistent associations between the expression of intracellular markers and particular cell subsets can often be found, which can then validate the intracellular staining limits. 3.2. Use of whole blood The use of whole blood for ex vivo staining is a definite advantage, for both clarity of staining and consequently the interpretation of results, and should be preferred whenever possible. In addition to requir-

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Fig. 1. Example of the dissection of the lymphocyte population into several subsets in order to determine the correct specific fluorescence staining limits. Leukocytes were stained for the cell surface markers CD8, CD28 and CD45RO, and for intracellular perforin. The partition of the lymphocytes in distinct subsets using several phenotypic markers enables a clear resolution of three levels of perforin expression: negative, low and high.

ing only small amounts of sample, cells directly stained from whole blood are untouched (so remain unaffected by the Ficoll cell separation), which leads to reduced backgrounds and yields a cleaner separation of the stained populations for cell surface and tetramer stainings. This is a particular advantage for intracellular staining (Fig. 2). Intracellular staining of cryopreserved PBMC may present an additional disadvantage: using a similar protocol to that for whole blood, permeabilization of the cells may be incomplete, which may lead to erroneous observations, as illustrated in Fig. 2. The staining of intracellular granzyme A in whole blood shows that granzyme A is expressed in most CD8+ T cells with the exception of naı¨ve cells (CD27 positive high), and that all the gated tetramer-positive cells express granzyme A. However, the staining of the same population from cryopreserved PBMC results in totally different observations, with a different distribution of granzyme A in the CD8+ T-cell subsets and a majority of the gated tetramer-positive cells expressing no granzyme A. Permeabilization of the cells results in a slight but significant reduction of the cell size (observed on FSC-height). On the cryopreserved PBMC, by gating on the smaller (low FSC) and therefore permeabilized lymphocytes, a similar pattern of granzyme A expression as in whole blood can be obtained, so that all the tetramer-positive cells express granzyme A. We have observed similar permeabilization problems with sev-

eral intracellular markers (including IFNg, perforin, granzyme A, Bcl-2 and CD3~). This situation is not absolute, and consistent staining can often be obtained from cryopreserved PBMC (substantial vortexing of the cells during permeabilization can help). Nevertheless, it is an important point to consider and to check carefully whenever intracellular staining is carried out, in order to avoid any misjudgment and misinterpretation of intracellular FACS staining. 3.3. Tetramers and T-cell stimulation The general protocol used in our laboratory is presented in Appendix B. Staining for cytokines such as IFNg or IL-2 requires stimulation of the cells to induce cytokine production, which presents particular difficulties. For instance, the in vivo relevance of an in vitro stimulation-based assay can be debated. The common ICS protocol uses phorbol myristate acetate (PMA) and ionomycin (Pala et al., 2000), which induce an efficient but rather artificial stimulation. Apart from the use of mitogens, antigen-specific CD8+ T cells can also be stimulated by means of specific peptides, which represent a more physiological way of stimulation. However, cytokine production is activation level dependent, and can therefore differ as a function of the concentration of peptide used, which appears maximal at 10 AM (Kostense et al., 2001; Appay et al., 2000). Also, the higher the

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Fig. 2. Example of intracellular granzyme A staining in tetramer-positive cells from whole blood or cryopreserved PBMC. Four color stainings (granzyme A, tetramer, CD8 and CD27) were carried out in a single donor directly from whole blood or on cryopreserved PBMC. The use of whole blood provides significantly clearer and more reliable stainings.

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concentration used, the more likely activation-induced cell death (AICD) is to result (Kostense et al., 2001). It remains therefore open to debate which peptide concentration is the most appropriate and how results with different peptide concentrations should be interpreted. Cellular activation to trigger cytokine production results also in the down-regulation of the T-cell receptor (Valitutti et al., 1996), which can make it difficult to perform tetramer staining and to identify the relevant antigen-specific populations. It was shown that staining with tetramers prior to activation can help significantly in preserving a reasonable level of tetramer staining, even following the incubation time for cytokine production (Appay et al., 2000); it is possible that tetramers are internalized during this period, but can still be detected in permeabilized cells (Whelan et al., 1999). This method has been used in a number of recent studies (Appay et al., 2000; Callan et al., 2000; Oxenius et al., 2001; Hislop et al., 2001). Although the binding of tetramers by itself can induce partial stimulation of the cells, this does not seem to have any major consequences for the readout of cytokine production (Appay et al., 2000). Nevertheless, tetramer staining for the no-stimulation control should be done at the end of the cytokine production incubation time to avoid any background increase resulting per se from the tetramer staining. A limitation of the ICS technique is the need for high cell numbers in order to carry out the assay. Considering the generally small size of the tetramerpositive cell populations, and the treatment undergone by the cells (i.e. activation, permeabilization), 1  105 to 5  106 PBMC per staining, with at least 0.5% of tetramer-positive CD8+ T cells, are generally needed to start with, in order to ensure a satisfactory result. Whole blood can also be used for ICS, which produces notably clearer stainings and enables the detection of low frequencies of cytokine-producing cells (Suni et al., 1998). However, in preliminary assays, we were not entirely satisfied with the combination of tetramer staining and ICS in whole blood, as, following the cytokine production incubation time, tetramer staining did not seem as reliable as with freshly purified PBMC. An optimized protocol for tetramer-ICS in whole blood would definitely represent a significant advance.

4. Concluding remarks The access to an increasing number of intracellular markers and the rapid developments in flow cytometry has opened seemingly endless possibilities for study of the characteristics and functions of antigen-specific CD8+ T cells in different pathologies. In the near future, we are likely to witness the arrival of even more elaborate applications that combine tetramer technology with other flow cytometry-based techniques, such as the direct measure of cytotoxicity [e.g. in vivo killing assay (Nelson et al., 2000), FATAL assay (Sheehy et al., 2001)] or the measure of calcium mobilization (Greimers et al., 1996), in order to provide further insights in the mechanisms regulating T-cell responses. Moreover, the introduction of new technologies such as multicolor (more than four) flow cytometry (Baumgarth and Roederer, 2000), laser scanning cytometry (Kamentsky, 2001) and HLA class II tetramers (McMichael and Kelleher, 1999) (see the present issue) is very promising. The development and combination of these technologies has already offered a better understanding of basic immunology and is likely to change wellestablished concepts in the future.

Appendix A. Intracellular staining in whole blood or PBMC (tested for intracellular perforin, granzymes A and B, GMP-17, Ki-67, Bcl-2 and CD3~) 

 

 

Add relevant titrated tetramers to 150 Al of heparinised whole blood (or 2  105 to 5  105 PBMC) and incubate for 15 min at 37 jC. Add conjugated monoclonal antibodies specific for extracellular markers and incubate for 15 min. For whole blood, add 3 ml of FACSk lysing solution (Becton Dickinson, San Jose, CA) (diluted 1:10 ratio in water), vortex and leave for 15 min at room temperature in the dark. Centrifuge (5 min at 1500 rpm) and discard supernatant. Wash  1 with 3 ml of PBE (PBS; 0.5% BSA; 0.5 mM EDTA). Add 300 Al FACSk permeabilizing solution (Becton Dickinson) (diluted 1:10 ratio in water), vortex and leave for 15 min at room temperature in the dark.

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Add 3 ml of PBE, spin and discard supernatant. Add conjugated monoclonal antibodies specific for intracellular markers and incubate in the dark for 15 min.  Wash  1 with 3 ml of PBE, discard supernatant.  Resuspend in 200 Al of FACS fix (PBS with 2% formaldehyde).  Keep at 4 jC until analysis. 

Appendix B. ICS in PBMC (tested for intracellular IFNg, TNFa, MIP-1h, IL2) 



    



  

Add relevant titrated tetramers to 1  105 –5  106 PBMCs (for peptide stimulated cells), incubate for 15 min at 37 jC (leave nonstimulated controls untreated). Add specific peptides (10 AM final concentration) and anti-CD28 antibodies (1 Ag/ml) (optional) and incubate at 37 jC for 1 h (final volume 200 – 500 Al). (Use SEB as positive control at 2 Ag/ml.) Add Brefeldin A (10 Ag/ml final concentration). Incubate for 5 h. Add relevant tetramers to nonstimulated cells and incubate 15 min at 37 jC. Wash all tubes with PBE (PBS; 0.5% BSA; 0.5 mM EDTA), spin, and discard supernatant. Add 300 Al of FACSk permeabilizing solution (Becton Dickinson) (diluted 1:10 ratio in water) to the sample, vortex and incubate in the dark at room temperature for 15 min. Prepare antibody mix and incubate with titrated antibodies for 15 min at room temperature in the dark. Wash  1 with 3 ml of PBE, discard supernatant. Resuspend in 200 Al of FACS fix (PBS with 2% formaldehyde). Keep at 4 jC until analysis.

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