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Cytokine flow cytometry: a multiparametric approach for assessing cellular immune responses to viral antigens
Clinical Immunology 110 (2004) 222 – 231 www.elsevier.com/locate/yclim
Cytokine flow cytometry: a multiparametric approach for assessing cellular immune responses to viral antigens Vernon C. Maino * and Holden T. Maecker BD Biosciences, San Jose, CA 95131, USA Received 7 November 2003; accepted with revision 10 November 2003
Abstract Considerable attention has been focused on CD8 and CD4 T cell responses as a major element of the cellular immune response to viral infections including human immunodeficiency virus (HIV) and hepatitis C virus (HCV). However, increasing evidence based on the recent introduction of more quantitative assays for measuring antigen-specific T cells has suggested that the role of these cells in the development of a protective immune response to a particular viral pathogen may be determined by a complex interplay of multiple virologic and cellular factors. Thus, measurements of only the frequencies of the T cell subsets participating in the response to viral pathogens may be an incomplete reflection of efficacy. In this review, we suggest that some measurable factors may influence the role of T cell immunity in conferring protection, including functional avidity, epitope breadth and specificity, proliferative capacity, cytokine repertoire, degree of anergy, and differentiation phenotype, as well as magnitude, of viral-specific CD4 and CD8 T cells. We suggest that automated cytokine flow cytometry (CFC) is an efficient approach to the measurement of the complex interplay of multiple immune parameters involved in immune protection. These ideas are discussed in the context of new developments in sample preparation and analysis automation. D 2004 Published by Elsevier Inc. Keywords: CFC; Viral pathogens; Immune response
Introduction The generation and maintenance of immunity to most viral antigens involve a complex balance of both humoral and cellular immunity. It is likely that successful control of viral infection requires the integrated and effective action of multiple components of innate and adaptive immunity. Until recently, the potency of the immune response to viral antigens was largely based on the strength of antibody responses as measured by radioimmunoassay (RIA) or enzyme-linked immunosorbent assay (ELISA) [1– 4]. More recently, it has become clear that both CD4 and CD8 T cell responses to infectious pathogen or vaccine may also play Abbreviations: CFC, cytokine flow cytometry; CMV, cytomegalovirus; CTL, cytotoxic T lymphocyte; ELISA, enzyme-linked immunosorbent assay; ELISPOT, enzyme-linked immunospot assay; HCV, hepatitis C virus; HIV, human immunodeficiency virus; MHC, major histocompatibility complex; RIA, radioimmunoassay. * Corresponding author. BD Biosciences, 2350 Qume Drive, San Jose, CA 95131. Fax: +1-408-954-2156. E-mail address: [email protected] (V.C. Maino). 1521-6616/$ - see front matter D 2004 Published by Elsevier Inc. doi:10.1016/j.clim.2003.11.018
significant roles in the protection from progressive viral infection and disease [5– 9]. Thus, quantitation of these T cell responses has become a focus of increasing interest in the quest to understand protective immunity in many infectious diseases. In recent years, labor-intensive, nonquantitative assays measuring bulk culture T cell proliferation and cytotoxicity have given way to short-term highly quantitative single-cell assays measuring cytokine secretion or major histocompatibility complex (MHC) oligomer binding. As a result, one can now measure populations of antigen-specific T cells of particular phenotypes and/or functions at very precise frequencies [10 – 13]. Although this has provided more quantitative (and qualitative) information about T cell responses to viral infection, it is not yet known what combination of phenotypes and functions, and what frequencies of antigen-specific T cells, provide protection from viral infections such as human immunodeficiency virus (HIV) [6,14 – 16]. In this review, we will examine the critical features of cytokine flow cytometry compared with other single-cell
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immune function techniques and we will describe several new developments in the flow cytometric analysis of cell surface and intracellular markers of antigen-responsive cells that can provide the capability to rapidly examine multiple markers of cellular immunity from small clinical samples. This capability has the potential to accelerate the discovery of new combinations of cellular and soluble markers that might constitute protective cellular immunity to viral antigens such as HIV and hepatitis C virus (HCV).
Methodological considerations Cytokine flow cytometry (CFC) combines the advantages of multiparameter analysis by flow cytometry with the functional readout of cytokine production as a unique method to characterize antigen-specific T cell responses. Like enzyme-linked immunospot (ELISPOT) assays, CFC can measure the response to an entire protein, viral lysate, or peptide. In contrast to analysis of antigen-specific T cells with MHC-peptide oligomers, which require known peptide sequences matched to specific class I or class II MHC, T cell cytokine responses to intact proteins or peptide mixtures can be determined with cells of unknown MHC origin. In addition to cytokines, other intracellular markers can be assessed including chemokines, perforin, and granzymes [17 – 20]. Furthermore, both CD4 and CD8 T cell subset responses can be simultaneously detected and distinguished within the same sample when peptide pools are used for stimulation [21] (Fig. 1). In general, peptides containing 15 amino acid residues each, and overlapping by 11 amino acid residues, elicit equivalent CD4 responses to the intact
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proteins from which the peptide sequences were derived [21]. The basic CFC methodology involves short-term in vitro activation (frequently 6 h) with protein or peptide antigens or mitogens, in the presence of a secretion inhibitor such as brefeldin A to enhance the accumulation of cytokines or other intracellular markers. For peptides or mitogens, which do not require processing by host antigenpresenting cells, the secretion inhibitor can be present during the entire incubation period with no reduction in response [21,22]. For protein antigens, an initial period (1 –2 h) of incubation is performed with antigen alone to allow for optimal antigen processing and presentation, which could otherwise be compromised by the presence of brefeldin A or monensin [13,22,23]. The entire incubation period is deliberately short to maintain cell viability and to accurately capture the frequency of responsive cells in whole blood or PBMC samples. Longer incubation times may compromise the quantitative nature of the assay because results would be altered by unpredictable proliferative and/or apoptotic responses which begin to be significant after 12 or more hours of stimulation [13,24]. For multiparametric staining of cell-surface and intracellular determinants, a surface-staining step is often added before cell fixation. However, some antibodies to cellsurface determinants retain avidity to epitopes subjected to fixation and permeabilization conditions. In these systems, all of the staining can be performed after fixation and permeabilization. Both whole blood and PBMC sample preparations may be used to assess frequencies of antigen-specific T cell cytokine responses to whole protein antigens, viral lysates,
Fig. 1. Quantitation of CD4 and CD8 T cell responses from the same activated sample. Because peptide mixes of 15 amino acid residues can stimulate both CD4 and CD8 T cells, gating on the population of interest (left panel) can reveal both responses from the same activated blood or PBMC sample. Responses to an HIV p55 peptide mix and a CMV pp65 peptide mix are shown for an HIV+ blood sample (right panels).
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Fig. 2. IFNg CFC responses measured in percentage of CD3+ CD4+ or CD3+ CD4 responding T cells from whole blood versus PBMC washed into medium (RPMI + 10% FCS).
or peptides. Whole blood activation obviates the need for cell processing before stimulation and yields results equivalent to those obtained with PBMC preparations [23,25] (Fig. 2). Fractionated PBMC may offer advantages when utilizing cryopreserved specimens, however, it is important to validate freezing and thawing procedures when performing functional assays of this type to insure sensitivity and reproducibility.
How does CFC compare with alternative measures of T cell immunity? Proliferation Traditional measurements of CD4 and CD8 T cell responses to antigen have relied on long-term assays which require days to weeks of tissue culture and only provide semiquantitative information that is dependent on population survival behavior in tissue culture. The lymphoproliferative assay (LPA), which largely measures the in vitro proliferative capacity of CD4 T cells, became the early standard measure of antigen-specific memory CD4 T cell response and is still widely used today [26 –29]. Typically, PBMC are placed in culture with recombinant protein antigens for 5 – 7 days and then pulsed with 3H-thymidine to detect proliferation. In reality, this is an integrative assay, as it is dependent upon the following parameters: (1) antigen processing and presentation; (2) the ability of T cells to proliferate in culture; and (3) the initial frequency of antigen-specific T cells [28,30]. Of course, there is no way to separate these three parameters, or even to know what type(s) of cells are proliferating.
Although in some examples the frequency of CD4+ T cells expressing cytokine in response to antigen exhibits some correlation with stimulation indices obtained using 3 H-thymidine-based proliferation assays [31 –33], this relationship may vary depending upon in vitro culture conditions which may influence the relative rates of proliferation and apoptosis. More recently, single-cell flow cytometric methods been employed to measure proliferative T cell responses to various stimuli including antigen [34 –36] These procedures include intracellular markers of cell division including Ki67 [35,37 – 39] in vitro or in vivo incorporation of BrdU, a thymidine analog, into DNA, which can be detected by antibody [33] and vital membrane dyes, for example, CFSE [40], which can be employed to determine the number of division cycles. Although in general, singlecell analyses of cellular proliferation demonstrate higher correlations with short-term single-cell assays of T cell activation, the requirement to subject T cells to longer term tissue culture (>24 h) may still compromise accurate measurements of ex vivo frequencies of proliferating antigen-specific cells. CTL assays Just as proliferation has been employed as a measure of CD4 T cell function, the cytotoxic T lymphocyte (CTL) assay has been a traditional measurement of the CD8 T cell response. The methods used to assess CTL function typically rely on long-term culture of PBMC with antigen followed by incubation of activated T cells with 51chromium-labeled MHC-matched target cells expressing the presented peptide or peptides of interest. Activity is typically based on the ratio of effector to target cells which promotes an arbitrary percentage of cell lysis (e.g. 50%) relative to a positive control for 51chromium release. The procedure is time consuming, uses large amounts of biological sample, and because it measures population responses, is poorly quantitative. A potentially more valuable assessment of the CD8 T cell response would be to measure the frequency of CTL precursor frequencies, that is, antigen-specific T cells capable of killing target cells expressing cognate antigen. In this regard, several markers have been associated with cytotoxic activity including granzymes [41 –43], perforin [43 – 46], interferon-g [36,44], and TNF-a [47,48]. In a recent study, Ghanekar et al. [44] utilized CFC to correlate the frequency of cytomegalovirus (CMV) peptide-induced CD8+ T cells expressing INFg and/or perforin with CTL activity as measured by traditional 51Cr-release assays. Strong positive correlation was observed between specific lysis of peptide-pulsed targets in a 51Cr-release assay and frequencies of peptide-activated CD8+ T cells expressing IFN-g at 6 h (r2 = 0.72) or 7 days (r2 = 0.91, P < 0.005) (Fig. 3). Enumeration of responding cells expressing perforin, another marker associated with CTL, did not improve this correlation.
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MHC oligomers The binding of MHC class-I-restricted tetramer complexes to cognate epitope-specific T cells has been suggested as an alternative method to identify CTL precursor frequencies [11]. In comparative studies, the frequencies of cytokine-positive cells obtained by CFC have been similar to those obtained by MHC tetramer analysis (Refs. [49,51] and our unpublished results). Fig. 4 shows the results of dual flow cytometric analysis of CD8+ T cells recognizing a specific HLA-A2 matched peptide derived from the pp65 CMV antigen and cytokine-positive cells following stimulation with the same cognate pp65 peptide. The correlated expression of IFN-g and the binding of MHC-matched tetramer on peptide-activated CD8+ T cells demonstrate that these two methods typically detect overlapping populations of peptide-specific CD8+ T cell populations. It should be noted, however, that the use of MHC-peptide oligomers is limited to systems with defined peptide epitopes and MHC restriction—something not common in human clinical trial settings. ELISPOT
Fig. 3. Correlation of IFNg production as measured by CFC assay with cytotoxicity as measured by 51Cr-release assay in 12 HLA-A2+ CMV seropositive donors. (A) Six-hour CFC assay on day 0 (immediately ex vivo) is correlated with 51Cr release assay after 7 days restimulation in vitro. (B) Six-hour CFC assay after 7 days restimulation in vitro is even more highly correlated with 51Cr release assay done on the same day.
An alternate widely used assay for the determination of frequencies of antigen-specific T cell responses is the ELISPOT, or enzyme-linked immunospot assay, which measures cytokine secretion by PBMC in filter-bottom wells using immunochemical detection [52 – 55]. While this procedure allows for single-cell quantitation and a functional readout (cytokine production), it has the disadvantage of being a single-parameter (or at most dual-parameter) analysis, without the ability to easily and definitively discriminate subset phenotypes such as CD4+ and CD8+ T cells.
Fig. 4. Relationship of tetramer staining and IFNg production. (A) Example of a combined analysis of HLA-A2-CMV peptide tetramer staining and IFNg production after stimulation with the same CMV peptide. The percentage of cells producing IFNg is almost equal to the initial tetramer-positive population. Note the down-modulation of tetramer staining upon activation with the relevant peptide. (B) Correlation of separate tetramer and IFNg CFC assays done on eight HLA-A2+ CMV seropositive donors. The two assays are highly correlated, with a slope close to 1.
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Fig. 5. Correlation of ELISPOT and CFC assays using CMV lysate or CMV pp65 peptide mix stimulation in healthy CMV seropositive donors. Results are expressed as number of positive cells per 105 PBMC for both assays to allow quantitative comparison. Note from the X- and Y-axis scales that the CFC assay detects on average three to four times as many positive cells as the ELISPOT assay.
The advantage of this system, however, is that multiple samples can be rapidly screened for positive cytokine responses. A widely used application for this technique, for example, is T cell epitope mapping [56 – 58]. Although CFC and ELISPOT both measure single-cell cytokine responses to stimuli, they differ in various technical aspects. The instrument of detection for CFC is flow cytometry, whereas the readout for ELISPOT is microscopy or image analysis. In addition, these methods also differ in other respects including incubation times, number of parameters analyzed, and sample compatibility [59 –61]. In general, the T cell frequencies observed with either method correlate fairly well. Fig. 5 compares CFC and ELISPOT measurements of CMV-specific T cell responses from identical CMV seropositive samples. In this example, frequencies for both determinations were calculated as a percentage of total PBMC to better assess the relative sensitivities of each procedure. Both procedures were able to detect lowfrequency and high-frequency responses to CMV antigens, although the slope of the correlative relationship in this example suggests that CFC may detect higher numbers (2 to 5) of antigen-specific CD4 and CD8 T cell responses to CMV. Thus, while the limit of detection is generally lower with ELISPOT, in practice, flow cytometry may enable the detection of greater numbers of cells expressing cytokine [52,62].
Limitations of CFC Although CFC compares well with other quantitative and qualitative measures of T cell immunity in terms of sensitivity, breadth of response, and capability to define multiple phenotypes, there remain some limitations. The major limitations of CFC include: (1) the complexity of the assay is still relatively high because it involves live cell stimulation, cell fixation, permeabilization, and staining; (2) throughput (i.e. number of samples processed per unit time) of CFC assays is limited by the capability of the flow
cytometer; and (3) the sensitivity of the assay is limited by the upper limit of spontaneous cytokine-producing cells, which average around 0.05 – 0.1% for IFN-g production in CMV+ donors [63]. The first and second limitations can be addressed by use of standardized reagents and increased use of automation to be discussed below. The third limitation is only problematic if we believe that immunologically protective responses will fall within this range (0.05 – 0.1% of peripheral blood CD4 and CD8 T cells, respectively). In fact, the natural T cell immune response to many peptides derived from viruses like CMV is at least one log higher than this [63], and protective responses elicited by vaccines may need to generate similar frequencies [64]. Functional T cell assays are subject to higher variation than assays that employ phenotypic staining by FACS due to assay complexity and the biological variability associated with in vitro cell activation. Nevertheless, recent evaluations have reported an overall coefficient of variation in a CMVspecific CFC assay to be within 25% in a multisite study [59].
Clinical relevance of CFC assays Using techniques like CFC, it is clear that chronic viral infections including HIV, HCV, and CMV induce readily detectable CD4 and CD8 T cell responses, at least during acute infection [16,65 –67]. However, it is not clear what type of cellular response differentiates protective and nonprotective responses in these infections. The ability to quantitate frequencies of functional antigen-specific T cells has enabled investigators to assess the relationship between the strength of CD4+ and CD8+ T cell responses and immune protection in many disease models. Thus, the response to infection of LCMV in mice is associated with strong CD4 T cell cytokine responses, which are correlated with long-term memory [68]. In a murine model of Leishmania, protective vaccination is associated with the development of IFN-g-producing CD8 T cells, as measured by
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CFC [69,70]. In a primate model of AIDS, control of viral rebound after structured therapy interruption is strongly correlated with antiviral CD8 T cell cytokine responses [68]. Such responses are also correlated with protection from mucosal challenge after vaccination [71]. In humans, susceptibility to CMV-associated end organ disease in HIVinfected individuals was shown to correlate with the loss of CD4+ T cell IFN-g responses to CMV antigens [72 –74]. While such studies demonstrate correlation of CFC responses with disease outcome, there have been no proven thresholds established for protective immunity as measured by a functional assay in any disease system. Most likely, the magnitude of CD4 and CD8 responses alone will not be as predictive of protection from disease as a combination of magnitude and other factors such as epitope breadth and affinity, phenotype, and function of antigen-specific cells [61]. However, the measurement of a CFC response is likely to be a necessary, if not sufficient, component of protective immunity for many infectious diseases.
Rationale for enhanced throughput of flow-based antigen-specific assays Although flow cytometric assays provide a powerful approach to define surrogate markers of clinical responses in drug and vaccine studies, the throughput limitations described above have limited the utility of this platform in large clinical studies. These concerns and the perceived need to provide enhanced capability to assess biological complexity with the introduction of polychromatic flow cytometry (>4 fluorescent parameters) have stimulated interest in the development of hardware and software modifications for enhanced throughput performance for flowbased assays including CFC. The development of platforms that allow the simultaneous analysis of multiple immune response parameters from small amounts of biological sample should provide a much broader and more sophisticated evaluation of host responses to disease or vaccine challenge. In this context four- to eight-color multiparametric flow cytometric assays can assess multiple functional attributes of complex cellular phenotypes. These assays can be performed in 96-well plates using only 100 Al of either whole blood or PBMC cell preparations. With less than 10 ml of whole blood, an entire 96-well plate can be formatted to run 96 complex functional cellular assays with the potential to evaluate antigen dose, cytokine repertoire, antigen-specific T cell subset responses, and antigen-presenting cell function. A multiparametric examination of phenotypic and functional profiles of antigen-specific cells will most likely be necessary to generate enough data to determine which parameters best predict protective immunity to viral infection. Along with phenotypic and functional profiles, responses to specific epitopes may relate to protection, as may the overall breadth of epitopic responses. All of these
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factors can be compared, particularly with flow cytometrybased assays, but they will require highly standardized and automated procedures to be applied in large clinical trial settings.
Automation—a key capability in the development of standardized cellular assays Many processes contribute to standardization and automation of flow-based cellular assays. These processes include sample preparation, cell activation, cell surface and intracellular immunofluorescent staining, sample loading, and data analysis. Recent developments have enabled all of the procedures involved in cell activation and cell staining to be performed in 96-well plates [75]. Fig. 6 demonstrates comparable responses of whole blood and PBMC activated and stained in tubes versus 96-well plates. This advance greatly simplifies the challenge to integrate the flow-based assays with standard 96-well plate based automation platforms. The sample-handling component of a fully automated system can be addressed with various robotic solutions. Fig. 7 illustrates one example of an integrated sample processing robot which is capable of performing activation, staining, and washing of 96-well plate-based cellular assays including four-color antigen-specific cytokine flow cytometry. In this example, a robotic arm moves plates among the activation station, the fluid handler, an indexed plate centrifuge, and the flow cytometer. Additional stations can be added as desired including bar code readers, plate hotels, and incubators. The ability to process large numbers of multiplexed samples greatly expands the amount of data that must be analyzed. Manual four-color data analysis performed by individual operators, even experienced individuals, is both time-consuming and is a potential source of error and variation in results. To address this issue, automated gating algorithms have been developed which can perform hierarchical gating and statistical ‘‘best fit’’ analysis for a variety of experimental data. These algorithms are capable of quantitating rare event populations, as for example is required for the determination of frequencies of antigenspecific T cells obtained using CFC or MHC-oligomer analysis. In addition, these algorithms can work with annotated experimental protocols to provide easy access to the data and compatibility with complex database archiving. Use of automated analysis protocols integrated into current flow cytometric analysis software programs can enhance the sample processing throughput by up to 100fold and dramatically reduce hands-on technical time depending upon the number of samples analyzed. In addition, automated analysis applications introduce much needed objectivity for the analysis of flow cytometric data. Correlations of manual and automated analysis of CD4+ and CD8+ T cell responses from multiple subjects to CMV
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Fig. 6. Equivalence of tube- and plate-based CFC assays. PBMC (left panels) or whole blood samples (right panels) were activated and processed in tubes (top panels) or plates of the indicated type (bottom panels). Equivalent results for both unstimulated and pp65 peptide mix-stimulated samples are seen. A different donor is shown in the PBMC versus whole blood experiments.
peptide and staphylococcus enterotoxin B (SEB) antigens have been performed [75]. These comparisons demonstrate a very high correlation (r2 > 0.95; P < 0.0001) between frequencies of response determined by standard manual methods of analysis performed by an expert and automated
gating algorithms. Combined with front-end robotic sample handling solutions, automated gating and analysis of multiparametric flow cytometry data provide a powerful platform for high throughput analysis of complex biological processes involved in cellular immunity. Another signifi-
Fig. 7. An automated system for activation, processing, and acquisition of plate-based CFC samples. The system is linked by a central robotic arm and software which communicates with the various components as labeled.
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cant benefit of automated methodologies for flow cytometry, especially in the context of evaluation of clinical trials, is the opportunity to establish standardized quantitative assays for measuring cellular immunity in multiple patient samples.
The future: an expanded role for multiparameter flow cytometry in monitoring immune responses in clinical samples Although it is generally agreed that T cells play a central role in the generation of protective immune responses for many bacterial and viral infections, it has been difficult in humans to develop prognostic cellular markers of immune protection to chronic viruses like HIV and HCV in vaccinated or infected individuals. It is likely that multiple markers of cellular immunity will finally constitute the best predictor of immune protection to chronic viral infection in human subjects. The ability to measure multiple parameters of cellular and humoral immunity made possible with new developments in flow cytometry hardware and software will greatly extend the throughput and utility of this platform in the analysis of complex processes of cellular immunity. Automation will also allow flow cytometric evaluations to be standardized across clinical trial sites and importantly will enable sample tracking and error-free data annotation. To optimize the value of the impressive amounts of data flow cytometry can generate, it will also be important to marry data processing capability to the automated data analysis algorithms. Using meta-file analysis, data can be mined from multiple database sources to attempt to correlate any combination of parameters (which can include the dimensions of both subset frequencies and antigen density) with other experimental or clinical parameters, for example, clinical end points of immune protection. Armed with automated data handling capability, highly annotated flexible databases, and the more recently introduced flow cytometers with higher analysis rates and capability to analyze eight or more parameters, the clinical investigator can look forward to vastly more comprehensive evaluations of cellular and humoral immunity from standard clinical samples. One might consider, for example, examining CD4+ and CD8+ T cell responses to antigen in the context of antigen dose (functional avidity), peptide pools (epitope specificities), cytokine and chemokine expression, and identification of proliferative subsets. It is a reasonable hypothesis that some combination of these parameters will correlate with protective immunity to a given pathogen, such as HIV. It is also possible that different immune parameters will apply to different pathogens, and these parameters will require large clinical trials to validate them for a given disease. If one considers each analysis (from 100 to 200 Al of whole blood) as capable of evaluating 6 –10 separate parameters, it is possible to assess of 300 –500
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Cytokine flow cytometry: a multiparametric approach for assessing cellular immune responses to viral antigens Vernon C. Maino * and Holden T. Maecker BD Biosciences, San Jose, CA 95131, USA Received 7 November 2003; accepted with revision 10 November 2003
Abstract Considerable attention has been focused on CD8 and CD4 T cell responses as a major element of the cellular immune response to viral infections including human immunodeficiency virus (HIV) and hepatitis C virus (HCV). However, increasing evidence based on the recent introduction of more quantitative assays for measuring antigen-specific T cells has suggested that the role of these cells in the development of a protective immune response to a particular viral pathogen may be determined by a complex interplay of multiple virologic and cellular factors. Thus, measurements of only the frequencies of the T cell subsets participating in the response to viral pathogens may be an incomplete reflection of efficacy. In this review, we suggest that some measurable factors may influence the role of T cell immunity in conferring protection, including functional avidity, epitope breadth and specificity, proliferative capacity, cytokine repertoire, degree of anergy, and differentiation phenotype, as well as magnitude, of viral-specific CD4 and CD8 T cells. We suggest that automated cytokine flow cytometry (CFC) is an efficient approach to the measurement of the complex interplay of multiple immune parameters involved in immune protection. These ideas are discussed in the context of new developments in sample preparation and analysis automation. D 2004 Published by Elsevier Inc. Keywords: CFC; Viral pathogens; Immune response
Introduction The generation and maintenance of immunity to most viral antigens involve a complex balance of both humoral and cellular immunity. It is likely that successful control of viral infection requires the integrated and effective action of multiple components of innate and adaptive immunity. Until recently, the potency of the immune response to viral antigens was largely based on the strength of antibody responses as measured by radioimmunoassay (RIA) or enzyme-linked immunosorbent assay (ELISA) [1– 4]. More recently, it has become clear that both CD4 and CD8 T cell responses to infectious pathogen or vaccine may also play Abbreviations: CFC, cytokine flow cytometry; CMV, cytomegalovirus; CTL, cytotoxic T lymphocyte; ELISA, enzyme-linked immunosorbent assay; ELISPOT, enzyme-linked immunospot assay; HCV, hepatitis C virus; HIV, human immunodeficiency virus; MHC, major histocompatibility complex; RIA, radioimmunoassay. * Corresponding author. BD Biosciences, 2350 Qume Drive, San Jose, CA 95131. Fax: +1-408-954-2156. E-mail address: [email protected] (V.C. Maino). 1521-6616/$ - see front matter D 2004 Published by Elsevier Inc. doi:10.1016/j.clim.2003.11.018
significant roles in the protection from progressive viral infection and disease [5– 9]. Thus, quantitation of these T cell responses has become a focus of increasing interest in the quest to understand protective immunity in many infectious diseases. In recent years, labor-intensive, nonquantitative assays measuring bulk culture T cell proliferation and cytotoxicity have given way to short-term highly quantitative single-cell assays measuring cytokine secretion or major histocompatibility complex (MHC) oligomer binding. As a result, one can now measure populations of antigen-specific T cells of particular phenotypes and/or functions at very precise frequencies [10 – 13]. Although this has provided more quantitative (and qualitative) information about T cell responses to viral infection, it is not yet known what combination of phenotypes and functions, and what frequencies of antigen-specific T cells, provide protection from viral infections such as human immunodeficiency virus (HIV) [6,14 – 16]. In this review, we will examine the critical features of cytokine flow cytometry compared with other single-cell
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immune function techniques and we will describe several new developments in the flow cytometric analysis of cell surface and intracellular markers of antigen-responsive cells that can provide the capability to rapidly examine multiple markers of cellular immunity from small clinical samples. This capability has the potential to accelerate the discovery of new combinations of cellular and soluble markers that might constitute protective cellular immunity to viral antigens such as HIV and hepatitis C virus (HCV).
Methodological considerations Cytokine flow cytometry (CFC) combines the advantages of multiparameter analysis by flow cytometry with the functional readout of cytokine production as a unique method to characterize antigen-specific T cell responses. Like enzyme-linked immunospot (ELISPOT) assays, CFC can measure the response to an entire protein, viral lysate, or peptide. In contrast to analysis of antigen-specific T cells with MHC-peptide oligomers, which require known peptide sequences matched to specific class I or class II MHC, T cell cytokine responses to intact proteins or peptide mixtures can be determined with cells of unknown MHC origin. In addition to cytokines, other intracellular markers can be assessed including chemokines, perforin, and granzymes [17 – 20]. Furthermore, both CD4 and CD8 T cell subset responses can be simultaneously detected and distinguished within the same sample when peptide pools are used for stimulation [21] (Fig. 1). In general, peptides containing 15 amino acid residues each, and overlapping by 11 amino acid residues, elicit equivalent CD4 responses to the intact
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proteins from which the peptide sequences were derived [21]. The basic CFC methodology involves short-term in vitro activation (frequently 6 h) with protein or peptide antigens or mitogens, in the presence of a secretion inhibitor such as brefeldin A to enhance the accumulation of cytokines or other intracellular markers. For peptides or mitogens, which do not require processing by host antigenpresenting cells, the secretion inhibitor can be present during the entire incubation period with no reduction in response [21,22]. For protein antigens, an initial period (1 –2 h) of incubation is performed with antigen alone to allow for optimal antigen processing and presentation, which could otherwise be compromised by the presence of brefeldin A or monensin [13,22,23]. The entire incubation period is deliberately short to maintain cell viability and to accurately capture the frequency of responsive cells in whole blood or PBMC samples. Longer incubation times may compromise the quantitative nature of the assay because results would be altered by unpredictable proliferative and/or apoptotic responses which begin to be significant after 12 or more hours of stimulation [13,24]. For multiparametric staining of cell-surface and intracellular determinants, a surface-staining step is often added before cell fixation. However, some antibodies to cellsurface determinants retain avidity to epitopes subjected to fixation and permeabilization conditions. In these systems, all of the staining can be performed after fixation and permeabilization. Both whole blood and PBMC sample preparations may be used to assess frequencies of antigen-specific T cell cytokine responses to whole protein antigens, viral lysates,
Fig. 1. Quantitation of CD4 and CD8 T cell responses from the same activated sample. Because peptide mixes of 15 amino acid residues can stimulate both CD4 and CD8 T cells, gating on the population of interest (left panel) can reveal both responses from the same activated blood or PBMC sample. Responses to an HIV p55 peptide mix and a CMV pp65 peptide mix are shown for an HIV+ blood sample (right panels).
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Fig. 2. IFNg CFC responses measured in percentage of CD3+ CD4+ or CD3+ CD4 responding T cells from whole blood versus PBMC washed into medium (RPMI + 10% FCS).
or peptides. Whole blood activation obviates the need for cell processing before stimulation and yields results equivalent to those obtained with PBMC preparations [23,25] (Fig. 2). Fractionated PBMC may offer advantages when utilizing cryopreserved specimens, however, it is important to validate freezing and thawing procedures when performing functional assays of this type to insure sensitivity and reproducibility.
How does CFC compare with alternative measures of T cell immunity? Proliferation Traditional measurements of CD4 and CD8 T cell responses to antigen have relied on long-term assays which require days to weeks of tissue culture and only provide semiquantitative information that is dependent on population survival behavior in tissue culture. The lymphoproliferative assay (LPA), which largely measures the in vitro proliferative capacity of CD4 T cells, became the early standard measure of antigen-specific memory CD4 T cell response and is still widely used today [26 –29]. Typically, PBMC are placed in culture with recombinant protein antigens for 5 – 7 days and then pulsed with 3H-thymidine to detect proliferation. In reality, this is an integrative assay, as it is dependent upon the following parameters: (1) antigen processing and presentation; (2) the ability of T cells to proliferate in culture; and (3) the initial frequency of antigen-specific T cells [28,30]. Of course, there is no way to separate these three parameters, or even to know what type(s) of cells are proliferating.
Although in some examples the frequency of CD4+ T cells expressing cytokine in response to antigen exhibits some correlation with stimulation indices obtained using 3 H-thymidine-based proliferation assays [31 –33], this relationship may vary depending upon in vitro culture conditions which may influence the relative rates of proliferation and apoptosis. More recently, single-cell flow cytometric methods been employed to measure proliferative T cell responses to various stimuli including antigen [34 –36] These procedures include intracellular markers of cell division including Ki67 [35,37 – 39] in vitro or in vivo incorporation of BrdU, a thymidine analog, into DNA, which can be detected by antibody [33] and vital membrane dyes, for example, CFSE [40], which can be employed to determine the number of division cycles. Although in general, singlecell analyses of cellular proliferation demonstrate higher correlations with short-term single-cell assays of T cell activation, the requirement to subject T cells to longer term tissue culture (>24 h) may still compromise accurate measurements of ex vivo frequencies of proliferating antigen-specific cells. CTL assays Just as proliferation has been employed as a measure of CD4 T cell function, the cytotoxic T lymphocyte (CTL) assay has been a traditional measurement of the CD8 T cell response. The methods used to assess CTL function typically rely on long-term culture of PBMC with antigen followed by incubation of activated T cells with 51chromium-labeled MHC-matched target cells expressing the presented peptide or peptides of interest. Activity is typically based on the ratio of effector to target cells which promotes an arbitrary percentage of cell lysis (e.g. 50%) relative to a positive control for 51chromium release. The procedure is time consuming, uses large amounts of biological sample, and because it measures population responses, is poorly quantitative. A potentially more valuable assessment of the CD8 T cell response would be to measure the frequency of CTL precursor frequencies, that is, antigen-specific T cells capable of killing target cells expressing cognate antigen. In this regard, several markers have been associated with cytotoxic activity including granzymes [41 –43], perforin [43 – 46], interferon-g [36,44], and TNF-a [47,48]. In a recent study, Ghanekar et al. [44] utilized CFC to correlate the frequency of cytomegalovirus (CMV) peptide-induced CD8+ T cells expressing INFg and/or perforin with CTL activity as measured by traditional 51Cr-release assays. Strong positive correlation was observed between specific lysis of peptide-pulsed targets in a 51Cr-release assay and frequencies of peptide-activated CD8+ T cells expressing IFN-g at 6 h (r2 = 0.72) or 7 days (r2 = 0.91, P < 0.005) (Fig. 3). Enumeration of responding cells expressing perforin, another marker associated with CTL, did not improve this correlation.
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MHC oligomers The binding of MHC class-I-restricted tetramer complexes to cognate epitope-specific T cells has been suggested as an alternative method to identify CTL precursor frequencies [11]. In comparative studies, the frequencies of cytokine-positive cells obtained by CFC have been similar to those obtained by MHC tetramer analysis (Refs. [49,51] and our unpublished results). Fig. 4 shows the results of dual flow cytometric analysis of CD8+ T cells recognizing a specific HLA-A2 matched peptide derived from the pp65 CMV antigen and cytokine-positive cells following stimulation with the same cognate pp65 peptide. The correlated expression of IFN-g and the binding of MHC-matched tetramer on peptide-activated CD8+ T cells demonstrate that these two methods typically detect overlapping populations of peptide-specific CD8+ T cell populations. It should be noted, however, that the use of MHC-peptide oligomers is limited to systems with defined peptide epitopes and MHC restriction—something not common in human clinical trial settings. ELISPOT
Fig. 3. Correlation of IFNg production as measured by CFC assay with cytotoxicity as measured by 51Cr-release assay in 12 HLA-A2+ CMV seropositive donors. (A) Six-hour CFC assay on day 0 (immediately ex vivo) is correlated with 51Cr release assay after 7 days restimulation in vitro. (B) Six-hour CFC assay after 7 days restimulation in vitro is even more highly correlated with 51Cr release assay done on the same day.
An alternate widely used assay for the determination of frequencies of antigen-specific T cell responses is the ELISPOT, or enzyme-linked immunospot assay, which measures cytokine secretion by PBMC in filter-bottom wells using immunochemical detection [52 – 55]. While this procedure allows for single-cell quantitation and a functional readout (cytokine production), it has the disadvantage of being a single-parameter (or at most dual-parameter) analysis, without the ability to easily and definitively discriminate subset phenotypes such as CD4+ and CD8+ T cells.
Fig. 4. Relationship of tetramer staining and IFNg production. (A) Example of a combined analysis of HLA-A2-CMV peptide tetramer staining and IFNg production after stimulation with the same CMV peptide. The percentage of cells producing IFNg is almost equal to the initial tetramer-positive population. Note the down-modulation of tetramer staining upon activation with the relevant peptide. (B) Correlation of separate tetramer and IFNg CFC assays done on eight HLA-A2+ CMV seropositive donors. The two assays are highly correlated, with a slope close to 1.
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Fig. 5. Correlation of ELISPOT and CFC assays using CMV lysate or CMV pp65 peptide mix stimulation in healthy CMV seropositive donors. Results are expressed as number of positive cells per 105 PBMC for both assays to allow quantitative comparison. Note from the X- and Y-axis scales that the CFC assay detects on average three to four times as many positive cells as the ELISPOT assay.
The advantage of this system, however, is that multiple samples can be rapidly screened for positive cytokine responses. A widely used application for this technique, for example, is T cell epitope mapping [56 – 58]. Although CFC and ELISPOT both measure single-cell cytokine responses to stimuli, they differ in various technical aspects. The instrument of detection for CFC is flow cytometry, whereas the readout for ELISPOT is microscopy or image analysis. In addition, these methods also differ in other respects including incubation times, number of parameters analyzed, and sample compatibility [59 –61]. In general, the T cell frequencies observed with either method correlate fairly well. Fig. 5 compares CFC and ELISPOT measurements of CMV-specific T cell responses from identical CMV seropositive samples. In this example, frequencies for both determinations were calculated as a percentage of total PBMC to better assess the relative sensitivities of each procedure. Both procedures were able to detect lowfrequency and high-frequency responses to CMV antigens, although the slope of the correlative relationship in this example suggests that CFC may detect higher numbers (2 to 5) of antigen-specific CD4 and CD8 T cell responses to CMV. Thus, while the limit of detection is generally lower with ELISPOT, in practice, flow cytometry may enable the detection of greater numbers of cells expressing cytokine [52,62].
Limitations of CFC Although CFC compares well with other quantitative and qualitative measures of T cell immunity in terms of sensitivity, breadth of response, and capability to define multiple phenotypes, there remain some limitations. The major limitations of CFC include: (1) the complexity of the assay is still relatively high because it involves live cell stimulation, cell fixation, permeabilization, and staining; (2) throughput (i.e. number of samples processed per unit time) of CFC assays is limited by the capability of the flow
cytometer; and (3) the sensitivity of the assay is limited by the upper limit of spontaneous cytokine-producing cells, which average around 0.05 – 0.1% for IFN-g production in CMV+ donors [63]. The first and second limitations can be addressed by use of standardized reagents and increased use of automation to be discussed below. The third limitation is only problematic if we believe that immunologically protective responses will fall within this range (0.05 – 0.1% of peripheral blood CD4 and CD8 T cells, respectively). In fact, the natural T cell immune response to many peptides derived from viruses like CMV is at least one log higher than this [63], and protective responses elicited by vaccines may need to generate similar frequencies [64]. Functional T cell assays are subject to higher variation than assays that employ phenotypic staining by FACS due to assay complexity and the biological variability associated with in vitro cell activation. Nevertheless, recent evaluations have reported an overall coefficient of variation in a CMVspecific CFC assay to be within 25% in a multisite study [59].
Clinical relevance of CFC assays Using techniques like CFC, it is clear that chronic viral infections including HIV, HCV, and CMV induce readily detectable CD4 and CD8 T cell responses, at least during acute infection [16,65 –67]. However, it is not clear what type of cellular response differentiates protective and nonprotective responses in these infections. The ability to quantitate frequencies of functional antigen-specific T cells has enabled investigators to assess the relationship between the strength of CD4+ and CD8+ T cell responses and immune protection in many disease models. Thus, the response to infection of LCMV in mice is associated with strong CD4 T cell cytokine responses, which are correlated with long-term memory [68]. In a murine model of Leishmania, protective vaccination is associated with the development of IFN-g-producing CD8 T cells, as measured by
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CFC [69,70]. In a primate model of AIDS, control of viral rebound after structured therapy interruption is strongly correlated with antiviral CD8 T cell cytokine responses [68]. Such responses are also correlated with protection from mucosal challenge after vaccination [71]. In humans, susceptibility to CMV-associated end organ disease in HIVinfected individuals was shown to correlate with the loss of CD4+ T cell IFN-g responses to CMV antigens [72 –74]. While such studies demonstrate correlation of CFC responses with disease outcome, there have been no proven thresholds established for protective immunity as measured by a functional assay in any disease system. Most likely, the magnitude of CD4 and CD8 responses alone will not be as predictive of protection from disease as a combination of magnitude and other factors such as epitope breadth and affinity, phenotype, and function of antigen-specific cells [61]. However, the measurement of a CFC response is likely to be a necessary, if not sufficient, component of protective immunity for many infectious diseases.
Rationale for enhanced throughput of flow-based antigen-specific assays Although flow cytometric assays provide a powerful approach to define surrogate markers of clinical responses in drug and vaccine studies, the throughput limitations described above have limited the utility of this platform in large clinical studies. These concerns and the perceived need to provide enhanced capability to assess biological complexity with the introduction of polychromatic flow cytometry (>4 fluorescent parameters) have stimulated interest in the development of hardware and software modifications for enhanced throughput performance for flowbased assays including CFC. The development of platforms that allow the simultaneous analysis of multiple immune response parameters from small amounts of biological sample should provide a much broader and more sophisticated evaluation of host responses to disease or vaccine challenge. In this context four- to eight-color multiparametric flow cytometric assays can assess multiple functional attributes of complex cellular phenotypes. These assays can be performed in 96-well plates using only 100 Al of either whole blood or PBMC cell preparations. With less than 10 ml of whole blood, an entire 96-well plate can be formatted to run 96 complex functional cellular assays with the potential to evaluate antigen dose, cytokine repertoire, antigen-specific T cell subset responses, and antigen-presenting cell function. A multiparametric examination of phenotypic and functional profiles of antigen-specific cells will most likely be necessary to generate enough data to determine which parameters best predict protective immunity to viral infection. Along with phenotypic and functional profiles, responses to specific epitopes may relate to protection, as may the overall breadth of epitopic responses. All of these
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factors can be compared, particularly with flow cytometrybased assays, but they will require highly standardized and automated procedures to be applied in large clinical trial settings.
Automation—a key capability in the development of standardized cellular assays Many processes contribute to standardization and automation of flow-based cellular assays. These processes include sample preparation, cell activation, cell surface and intracellular immunofluorescent staining, sample loading, and data analysis. Recent developments have enabled all of the procedures involved in cell activation and cell staining to be performed in 96-well plates [75]. Fig. 6 demonstrates comparable responses of whole blood and PBMC activated and stained in tubes versus 96-well plates. This advance greatly simplifies the challenge to integrate the flow-based assays with standard 96-well plate based automation platforms. The sample-handling component of a fully automated system can be addressed with various robotic solutions. Fig. 7 illustrates one example of an integrated sample processing robot which is capable of performing activation, staining, and washing of 96-well plate-based cellular assays including four-color antigen-specific cytokine flow cytometry. In this example, a robotic arm moves plates among the activation station, the fluid handler, an indexed plate centrifuge, and the flow cytometer. Additional stations can be added as desired including bar code readers, plate hotels, and incubators. The ability to process large numbers of multiplexed samples greatly expands the amount of data that must be analyzed. Manual four-color data analysis performed by individual operators, even experienced individuals, is both time-consuming and is a potential source of error and variation in results. To address this issue, automated gating algorithms have been developed which can perform hierarchical gating and statistical ‘‘best fit’’ analysis for a variety of experimental data. These algorithms are capable of quantitating rare event populations, as for example is required for the determination of frequencies of antigenspecific T cells obtained using CFC or MHC-oligomer analysis. In addition, these algorithms can work with annotated experimental protocols to provide easy access to the data and compatibility with complex database archiving. Use of automated analysis protocols integrated into current flow cytometric analysis software programs can enhance the sample processing throughput by up to 100fold and dramatically reduce hands-on technical time depending upon the number of samples analyzed. In addition, automated analysis applications introduce much needed objectivity for the analysis of flow cytometric data. Correlations of manual and automated analysis of CD4+ and CD8+ T cell responses from multiple subjects to CMV
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Fig. 6. Equivalence of tube- and plate-based CFC assays. PBMC (left panels) or whole blood samples (right panels) were activated and processed in tubes (top panels) or plates of the indicated type (bottom panels). Equivalent results for both unstimulated and pp65 peptide mix-stimulated samples are seen. A different donor is shown in the PBMC versus whole blood experiments.
peptide and staphylococcus enterotoxin B (SEB) antigens have been performed [75]. These comparisons demonstrate a very high correlation (r2 > 0.95; P < 0.0001) between frequencies of response determined by standard manual methods of analysis performed by an expert and automated
gating algorithms. Combined with front-end robotic sample handling solutions, automated gating and analysis of multiparametric flow cytometry data provide a powerful platform for high throughput analysis of complex biological processes involved in cellular immunity. Another signifi-
Fig. 7. An automated system for activation, processing, and acquisition of plate-based CFC samples. The system is linked by a central robotic arm and software which communicates with the various components as labeled.
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cant benefit of automated methodologies for flow cytometry, especially in the context of evaluation of clinical trials, is the opportunity to establish standardized quantitative assays for measuring cellular immunity in multiple patient samples.
The future: an expanded role for multiparameter flow cytometry in monitoring immune responses in clinical samples Although it is generally agreed that T cells play a central role in the generation of protective immune responses for many bacterial and viral infections, it has been difficult in humans to develop prognostic cellular markers of immune protection to chronic viruses like HIV and HCV in vaccinated or infected individuals. It is likely that multiple markers of cellular immunity will finally constitute the best predictor of immune protection to chronic viral infection in human subjects. The ability to measure multiple parameters of cellular and humoral immunity made possible with new developments in flow cytometry hardware and software will greatly extend the throughput and utility of this platform in the analysis of complex processes of cellular immunity. Automation will also allow flow cytometric evaluations to be standardized across clinical trial sites and importantly will enable sample tracking and error-free data annotation. To optimize the value of the impressive amounts of data flow cytometry can generate, it will also be important to marry data processing capability to the automated data analysis algorithms. Using meta-file analysis, data can be mined from multiple database sources to attempt to correlate any combination of parameters (which can include the dimensions of both subset frequencies and antigen density) with other experimental or clinical parameters, for example, clinical end points of immune protection. Armed with automated data handling capability, highly annotated flexible databases, and the more recently introduced flow cytometers with higher analysis rates and capability to analyze eight or more parameters, the clinical investigator can look forward to vastly more comprehensive evaluations of cellular and humoral immunity from standard clinical samples. One might consider, for example, examining CD4+ and CD8+ T cell responses to antigen in the context of antigen dose (functional avidity), peptide pools (epitope specificities), cytokine and chemokine expression, and identification of proliferative subsets. It is a reasonable hypothesis that some combination of these parameters will correlate with protective immunity to a given pathogen, such as HIV. It is also possible that different immune parameters will apply to different pathogens, and these parameters will require large clinical trials to validate them for a given disease. If one considers each analysis (from 100 to 200 Al of whole blood) as capable of evaluating 6 –10 separate parameters, it is possible to assess of 300 –500
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