MONITORING IMMUNE RESPONSES IN CANCER PATIENTS RECEIVING TUMOR VACCINES

Transcription

1 Intern. Rev. Immunol., 22: , 2003 Copyright # 2003 Taylor & Francis /03 $ DOI: / MONITORING IMMUNE RESPONSES IN CANCER PATIENTS RECEIVING TUMOR VACCINES EDWIN B. WALKER Providence Portland Medical Center, Earle A. Chiles Research Institute, Robert W. Franz Cancer Research Center, Portland, Oregon, USA MARY L. (NORA) DISIS University of Washington, Department of Medicine=Oncology, Seattle, Washington, USA Clinical evaluation of therapeutic tumor vaccines has resulted in examination and comparison of the types of immune function assays required to monitor tumor antigen stimulated T cell effector function in immunized patients. Three of the most commonly used assays include ELISPOT, tetramer assay, and cytokine flow cytometry (CFC). Discussed are the method and principles for each assay and an assessment of important methodological, reagent, and data acquisition issues that are relevant for the accurate and effective use of the assays. The sensitivity and utility of the assays and present arguments advocating their integrated use in future immunomonitoring studies are also discussed. Keywords: immunomonitoring assays, cancer vaccines The century-long effort to understand the mechanisms of tumoractivated host immune response has resulted in the current database of sequence and molecular structural information for human tumorspecific antigens (TSA) and tumor-associated antigen (TAA) systems [1]. This information is one of the cornerstones of the extensive recent effort to develop effective strategies of tumor vaccine immunotherapy. The present effort to test the clinical efficacy of therapeutic cancer vaccines is also bolstered by the convergence of data from several other venues of basic immunological research. Chief among these are the emerging details of the regulatory role (both tolerogenic and Address correspondence to Edwin B. Walker, 4805 NE Glisan, Providence Portland Medical Center, Earle A. Chiles Research Institute, Robert W. Franz Cancer Research Center, Portland, OR 97213, USA. edwalker@providence.org 283

2 284 E. B. Walker and M. L. (Nora) Disis stimulatory) played by professional antigen-presenting cells such as dendritic cells (DC), and the correlated functional and phenotype heterogeneity of DC subpopulations [2 4]; a more detailed understanding of the way potent T cell and DC-derived cytokine immunomodulators such as IL-12, IL-15, IL-2, GM-CSF, and IFN-g control the antigen-specific response of both CD4 and CD8 positive memory=effector T cells [5,6]; mechanistic details of MHC I- and MHC II-regulated T cell receptor (TCR) engagement, and the role that costimulatory molecules such as CD40, CD86, CD80, CD28, OX-40, and CTLA-4 play in T cell activation or tolerance [7 10]; the details of the critical role of the innate immune system in regulating the antigen-driven adaptive immune response through the interface of the dendritic cell [11 13]; and a more comprehensive understanding of the many mechanisms of tumor-mediated and T cell mediated inhibition of antitumor response, including the newly understood relevance of the inhibitory effects of Natural Killer Cells (NKT) cells and CD4 þ CD25 þ regulatory T cells [14,15]. Also, the recent successful development of immunomonitoring assay systems with markedly increased levels of specificity and sensitivity, capable of detecting low to moderate levels of T cell activation in Peripheral Blood Mononuclear Cells (PBMC) or in lymphocytes harvested from tumors or lymph nodes [16,17], will play a critical role in all future efforts in cancer vaccine development. There is now a rapidly accelerating clinical effort to test the effects of many different approaches to the development and delivery of therapeutic cancer vaccines. These diverse vaccine modalities include whole tumor or tumor lysate antigens admixed with a range of different adjuvants; tumor cells genetically modified to express immunomodulating cytokines such as granulocyte=macrophage-colony stimulating factor (GM-CSF) or costimulatory molecules; protein or peptide vaccines admixed with adjuvants and=or combined with concomitant administration of cytokines such as IL-2 or GM-CSF; dendritic cell vaccines comprised of DC expanded in ex vivo culture systems and pulsed with tumor lysate, TAA peptides, or whole proteins; DC fused with tumor cells or transduced to express tumor antigen systems, and the use of naked recombinant plasmid DNA or recombinant viruses to deliver TAA [1]. Many of these model tumor vaccine systems have recently been tested in clinical trials [1,18 20]. Effective assays to measure the host immune response(s) to these tumor vaccine interventions are becoming increasingly important tools to evaluate the efficacy of a given vaccine strategy and to promote a candidate vaccine into phase II or phase III clinical trials.

3 Immunomonitoring of Patients Receiving Cancer Vaccines 285 The ultimate goal of immunomonitoring assays designed to quantitate tumor-antigen specific effector T cell function is that of defining surrogate immune function parameters which correlate with changes in disease progression and are predictive of clinical outcome. However, this objective remains an elusive goal given the fact that, with few exceptions, there has been poor correlation between measured amplification of tumor-antigen stimulated T cell responses and clinical regression of disease [21 23]. There may be many explanations for the lack of correlation between a given tumor-antigen driven immune response and clinical outcome, including the most obvious one that the immunomonitoring assay employed measures an irrelevant T cell response that is not an effective mechanism for tumor destruction. Additional explanations could involve the inability of antigen-specific effector T cells to traffic to the tumor site or to survive in the often highly immunosuppressive microenvironment of the tumor [24]. This central paradox of tumor vaccine therapy does not, however, negate the fundamental importance of refining and optimizing immunomonitoring assays as tools to measure the potency of clinical vaccine candidates to stimulate tumor-associated antigen-specific T cell responses. A given vaccine candidate or vaccine strategy may not prove clinically effective in its initial application, but in the absence of any data characterizing its ability to stimulate antigen-specific immune function, there is no criteria basis for continued refinement and further testing of a potentially valuable cancer vaccine. Within the last two years there have been several reviews of major issues pertaining to the development and use of various in vitro assay systems to monitor immune function of cancer immunotherapy patients [16,17,24 26]. These reviews have focused primarily on giving a brief description of the various assay procedures, their strengths and weaknesses, a brief overview of how they have been used in clinical trials, and their relative sensitivity and practical utility. In this review we will recapitulate a discussion of the operational details of selected assay systems for the purpose of continuity. Thus, we will describe the operational principles and method for three effector T cell assays: the ELISPOT assay, MHC I restricted, peptidespecific tetramer staining and flow cytometry analysis, and the cytokine flow cytometry (CFC) assay. We have omitted any discussion of the delayed type hypersensitivity (DTH) test, the family of limiting dilution assays (LDA), and the measurement of T cell cytokines by real time quantitative polymerase chain reaction (PCR) analysis (qrt-pcr). As discussed in one recent review [17], in some vaccine settings it is questionable whether or not positive DTH responses are truly antigen specific, and overall the assay is at best semi-quantita-

4 286 E. B. Walker and M. L. (Nora) Disis tive. The qrt-pcr assay is a relatively new approach to immunomonitoring in cancer vaccine patients, and while it may be very valuable in measuring T cell responses in certain types of hypocellular samples, such as fine-needle aspirates, more extensive studies are required to determine its relative sensitivity compared to other techniques. qrt-pcr also yields very little useful information regarding the number or phenotype of cells producing antigen-stimulated cytokine mrna; it is not a particularly data-rich assay system [17,27]. LDA system involve the serial dilution of T cells in a large number of wells followed by in vitro stimulation and the subsequent measurement of T cell proliferation, cytolytic function, or cytokine synthesis. Poisson statistical analysis is used to determine the number of wells in the dilution series with one or less antigen-specific cells at the start of the stimulation and to determine the overall precursor frequency in the test sample. This assay, while quantitative, is extremely cumbersome, labor intensive, and fairly operator dependent [17]; there are also questions concerning its sensitivity compared to both the ELI- SPOT and the CFC assays [28,29]. While we will concentrate on the quantitative methods employed in the ELISPOT, CFC, and tetramer assays, we will also review in some detail key reagent and data analysis questions not generally covered in previous reviews, as well as important, relatively unresolved methodological issues common to the successful performance of all three assay formats. These methodological issues include a discussion of the methods used to determine the relative sensitivity of the assays, the problems associated with the use of frozen cells in the assay procedures, and the utility of combining the application of different assay systems in the integrated analysis of patient responses. We will additionally address the issue of assay validation and some of the unique challenges involved in validating patient memory=effector T cell immune response assay systems. ELISPOT ASSAY Operational Method The ELISPOT assay originally developed for quantitation of monoclonal antibody secreting cells [30] was later adapted for the enumeration of antigen-specific cytokine-secreting T cells [31,32]. The ELISPOT assay consists of six sequential steps: (1) coating of a flatbottom 96-well nitrocellulose-bottomed microtiter plate with a monoclonal antibody directed against the cytokine of interest; (2) the plate is then blocked to prevent nonspecific adherence of secreted proteins; (3) the test population of cytokine-secreting T cells is added to the plate

5 Immunomonitoring of Patients Receiving Cancer Vaccines 287 (usually PBMC or separated CD4 þ or CD8 þ T cells) and incubated with stimulator cells and=or an antigen source for variable time periods (usually h); (4) cells are lysed and washed off the plate; (5) an enzyme-linked, cytokine-specific antibody is added to the plate, incubated, and washed off; and (6) final resolution of the foci of antibodycytokine complexes by the addition of an enzyme-substrate reagent, which generates a colored spot at each point on the filter where a single cell or clusters of cells have secreted cytokine. These spots are then counted most reliably by some form of automated, computer-interfaced image analysis system. Data is usually calculated by ratioing the number of spots per well divided by the total number of cells plated in the well. Additional methodological adaptations that have been used to augment weak responses include the addition of costimulatory antibodies such as CD28=CD49d or cytokines such as IL-2 [33]. Methodological and Reagent Issues Most reported studies have used the ELISPOT assay to analyze MHC I restricted CD8 þ T cell responses to peptide antigen systems, although CD4 T cell responses against intact protein antigens have also been reported [34]. When whole protein antigen stimulation is used, effector cells such as PBMC are frequently preincubated for varying lengths of time with the protein antigen prior to plating the cells for the final assay procedure. In vitro stimulated cells (IVS) can be analyzed directly or after reconstitution with autologous antigenpresenting cells (APC) or a transporter associated with antigen processing (TAP)-deficient T2 cell line [35]. However, IVS effector cells frequently give high background cytokine expression if they are used after a very recent restimulation cycle. As with many biological assays, the serum source is often critical since many serum types or lots can stimulate high background effects usually a screened lot of human AB serum (heat inactivated) is preferable. The selection of the anticytokine antibody pair i.e., the capture or plate-coating antibody and the enzyme-linked resolution antibody is also critical to optimizing the assay. Antibody pairs should be tested carefully to identify a set that will give optimal antigen-specific binding. In order to ensure reproducible performance, a positive stimulation control such as the use of SEB of reagents and assay manipulation is frequently employed. Freezing of PBMC has been reported to have no effect on the quantitation of IFN-g secretion by effector T cells [134]. However, often reports of the use of cryopreserved PBMCs involve stimulating the cells with potent mitogens like PHA or PMA þ ionomyacin [33], or powerful infectious disease antigens such as varicella-zoster virus [37]

6 288 E. B. Walker and M. L. (Nora) Disis or influenza virus peptides [36]. It is still unclear if the T cell responses to stimulation by relatively weak TAA antigens such as gp100, tyrosinase, and MART-1 are similarly unaffected by cryopreservation. Currently, most laboratory environments actively engaged in immunomonitoring studies of cancer immunotherapy patients primarily analyze the functional and phenotype characteristics of archived collections of cryopreserved PBMC harvested from whole blood, leukapheresis collections, or lymphocytes recovered from lymph nodes or tumor samples [134]. The predominant use of cryopreserved cell samples results from the considerable logistical and methodological constraints, which frequently make the use of freshly collected cells impractical in most clinical research settings. However, it is generally understood that the use of cryopreserved cells in in vitro effector function assays, such as the ELISPOT or the CFC assay, frequently results in overall lower functional responses; this may especially be true when measuring T cell responses to intact complex protein recall antigens [38]. In recent years more emphasis has been placed on trying to understand the cellular biochemical and physiological events, which may be as important as the physical-chemical or osmotic forces that contribute to the attrition of cryopreserved cells. Thus, there is now a growing consensus that cellular hypothermia and cryopreservation of many different cell types results in increased levels of delayed-onset cell death due to apoptosis [39 45]. The freeze-thaw cycle appears to introduce sublethal stress or danger signals including oxidative stress, adenylate depletion, and nitric oxide upregulation [42], which can result in activation of proteolytic cleavage of apoptosis-related proteins such as caspase-3 and caspase-8 [41] and thus result in apoptosis and delayed onset cell death within h depending on the cell type. In recent years there have been many successful efforts to reduce apoptotic-dependent loss of viability and function of frozen cells using a variety of new cryopreservation procedures. These have included the use of caspase inhibitors [45], the combined use of glucose-supplemented freezing medium and programmable controlledrate freezers [40,43], and the use of dextran-based freezing medium such as Hypothermosol [42]. The continued exploration and development of more optimal freezing protocols will be key to the effective use of cryopreserved cells in immunomonitoring studies. A second potentially valuable approach to the preservation of viability and function of cryopreserved cells involves the possible addition of various cytokines with antiapoptotic potential to resting cultures of freshly thawed cells for short periods of time (24 48 h) prior to their use in immunomonitoring procedures. Two cytokine families, the type

7 Immunomonitoring of Patients Receiving Cancer Vaccines 289 I interferons (IFN-a and IFN-b) [46,47], and the IL-2 receptor family of cytokines including IL-2, IL-4, IL-7, and IL-15 [48 52] have been shown to prevent apoptosis in antigen-activated T cells; and, in the case of IL-2 and IL-15, to promote proliferative expansion of memory T cells in vivo even in the absence of antigen stimulation. IL-4, IL-7, and IL-15 have also been shown to prevent apoptosis of antigen-naïve, resting CD4 þ T cells [53], and in mature memory=effector CD4 þ cells IL-15 promotes stability and proliferative quiescence in the absence of TCR engagement [48]. IL-15 primarily sustains the low steady-state proliferation of CD8 þ memory T cells in vivo, and, in the absence of specific recall antigen, does not promote CD8 þ functional activation [6,47,54]. IL-15 is produced by dendritic cells after CD40 ligand stimulation [55] and may also prevent apoptosis of dendritic cells [56]. Thus, a second rational experimental approach to preventing apoptosis of memory T cell populations and dendritic cells in frozen PBMC or leukapheresis preparations might involve the relatively short-term (overnight) resting of freshly thawed cells in in vitro culture systems containing type I interferons or cytokines such as IL-7 and IL-15. Clinical Application and Correlation The ELISPOT assay has been used extensively to measure the frequencies of antiviral effector T cells there have been fewer studies using the assay to analyze T cell responses to tumor antigens [57 60]. Notably, there have been limited examples of correlation between clinical outcome and T cell responses measured by ELISPOT. In one study of patients immunized with a polyvalent melanoma vaccine, it was found that all ten patients responding to therapy also demonstrated measurable T cell responses to HLA class I binding peptides from MAGE 3 and=or from MART-1. Conversely, no T cell response to these antigens as measured by ELISPOT was found in the clinically nonresponding patients [61]. In a recent study by Banchereau and colleagues [62], melanoma patients vaccinated with HLA-A2 melanoma peptide-loaded DC demonstrated an overall positive correlation between clinical outcome and effector T cell response as measured by ELISPOT analysis. Subsets of patients vaccinated with a CEA antigen system showed correlation between increasing CEA-specific T cells and stable disease [63]. In another clinical study in which melanoma patients were vaccinated with tyrosinase peptides and GM-CSF, increased levels of peptide-specific T cells were found in four out of fifteen patients who also showed measurable clinical response. These responses included one patient with a mixed response, a patient with stable disease, and two patients with prolonged freedom from disease

8 290 E. B. Walker and M. L. (Nora) Disis recurrence [59]. In a Phase I trial reported by Marshall and coworkers [64], patients were immunized with a CEA antigen construct, and ELISPOT analysis was performed to determine if changes in CEAspecific T cell responses were prognostic for survival. The statistical analysis demonstrated that both higher post-treatment CEA-specific T cell responses as well as higher post-pretreatment CEA-specific T cell ratios correlated with increased survival time after accounting for disease status. Enk and coworkers demonstrated that MAGE-specific T cell numbers detected by ELISPOT correlated with improved survival in melanoma patients who were resected and had followup IFN-a therapy [65]. Sensitivity Comparison to Other Assays The limit of detection sensitivity of the ELISPOT assay has been shown to be at least 1 cell in 100,000. This result was reported in a study by Asai and colleagues in which a gp100 peptide-specific HLA- A2 þ CTL line was titered at decreasing ratios into culture wells containing normal HLA-A2 þ PBMC (10 5 per well) and cognate gp100 peptide antigen [33]. This type of limit of detection titration analysis using a very potent IFN-g producing cell line titered against a very quiescent background environment allowed for the detection of one IFN-g positive cell in 100,000 plated cells. Arguably, this type of limit of detection analysis of assay sensitivity may be somewhat artificial since more frequently the antigen-unstimulated background levels of reactive lymphocytes from patients with active or even subclinical infectious disease, or in situations where lymphocytes (PBMC) from vaccinated cancer patients are analyzed after in vitro cytokine-mediated IVS expansion, are relatively high. Thus, antigen-unstimulated background levels that reflect the more frequently encountered real world use of the ELISPOT assay are often substantially higher than one in 100,000 cells more commonly those background values can be as high as 0.01% to 0.02% of the plated negative control wells thus lowering the practical limits of detection for this assay. Additional confusion concerning the determination of ELISPOT assay sensitivity results from the fact that positive events ( spots ) are counted as a percentage of total plated cells. If the plated cells are intact PBMC then the total T cell number is a much smaller component of the total cell sample; therefore, positive events represent a much higher percent of the T cell subpopulation. By contrast, the CFC assay usually enumerates cytokine-positive cells as a percent of total gated CD8 þ or CD4 þ T cells. Thus, accurate comparative studies of ELISPOT and

9 Immunomonitoring of Patients Receiving Cancer Vaccines 291 CFC assay sensitivity must use techniques to normalize positive event counts in each assay to a common cellular standard. To date there have been few studies directly comparing ELISPOT results to other immunomonitoring assays, such as tetramer staining procedures or cytokine flow cytometry analysis (CFC). In one study where ELISPOT and MHC-specific peptide tetramer staining were compared for the detection of EBV antigen-specific CTLs, values obtained by tetramer analysis were approximately four times greater than those measured by ELISPOT analysis [29]. Similarly, other recent comparative studies have demonstrated higher tetramer detection values than those enumerated by ELISPOT analysis for the quantitation of influenza-specific and Melan-A specific effector T cells [66], and for the detection of tyrosinase-specific T cell clones [67]. In another very recent report, the frequency of influenza-reactive CD8 þ T cells in normal subjects was assessed by ELISPOT and by CFC analysis. The frequency of influenza-peptide responsive effector T cells as measured by each assay showed high interassay reproducibility and a close correlation between both assay systems [68]. Conversely, Kuzushima and colleagues, using both PBMC from EBV-positive subjects and established EBV-specific CTL lines, demonstrated that the CFC assay could detect fourfold higher EBV-specific T cell responses than those measured by ELISPOT analysis [28]. The ELISPOT assay has been widely used recently with good effect. With proper attention to screening and optimization of reagent stocks, the use of computer-interfaced image analysis systems to eliminate operator variability, the proper validation of the assay procedure, and the consistent use of internal system suitability positive controls, the assay procedure appears to be sensitive and accurate. Perhaps the major deficit of the assay is the fact that it is not a very data-rich system compared to the multiple parameter flow cytometry based environments of the tetramer and CFC assays. There are, however, new technological developments that have recently been described that will allow for simultaneous detection of two cytokines [69] and the estimation of the strength of cytokine expression from a single cell by measuring the size and density of an individual spot [70]. The ELI- SPOT assay can be semiautomated, and, on a practical level, can be performed fairly quickly in the laboratory. If further comparative studies demonstrate more conclusively its sensitivity is equal to or greater than the CFC assay, the ELISPOT assay could prove valuable as an initial screening system for high sample throughput analysis prior to the more detailed types of integrated quantitative and qualitative (phenotype) analysis that are afforded by the CFC and=or tetramer assay environments.

10 292 E. B. Walker and M. L. (Nora) Disis CYTOKINE FLOW CYTOMETRY (CFC) ASSAY Operational Method Cytokine flow cytometry (CFC) analysis is based on the principle that activated cells such as antigen-stimulated CD4 þ and CD8 þ T cells can be detected by directly measuring pooled, internal cytoplasmic levels of functionally important cytokines such as IFN-g and IL-2 using cytokine-specific, fluorochrome-conjugated monoclonal antibodies and correlated multiparameter phenotype analysis by flow cytometry. Thus, the functional phenotype of an antigen-triggered cell as measured by the synthesis of a specific cytokine can be correlated with different lineage and lineage subset cell surface phenotypes determined through the simultaneous use of at least three other cell surface antibodies and standard flow cytometry techniques. Cell populations of interest such as whole blood, Ficoll-separated PBMC, or cell suspensions harvested from lymph nodes are stimulated in vitro for relatively short periods of time (1 2 h) with recall antigens prior to the addition of drugs such as monesin or brefeldin A (BFA), which inhibit the subsequent secretion of synthesized cytokines [71,72]. Cells are incubated with brefeldin A for an additional 4 5 h and subsequently collected, washed, and fixed in paraformaldehyde or similar fixatives. Permeabilization of cell membranes is achieved using mild nonionic detergents to allow internalization of cytokine-specific monoclonal antibodies that have been selected and formulated for their binding specificity to formalin fixed forms of various functionally important cytokines. Subsequently, cells are washed and stained with relevant lineage-specific monoclonal antibodies, which delineate T cells and T cell subsets (i.e., CD3, CD8, and CD4) and with antibodies specific for activation markers such as CD69. Finally, cells are fixed in paraformaldehyde at 4 C prior to analysis. The CFC assay has been used recently to measure memory CD4 þ T cell responses to infectious disease recall antigens, demonstrating predominantly memory responses to protein antigens [72,73] and some CD8 memory response at higher concentrations of protein antigens [74]. By comparison, cognate recall peptides or peptide mixtures have been demonstrated to be very effective at stimulating memory CD8 T cell responses [75 79]. Methodological and Reagent Issues A critical methodological feature of the CFC assay system is the requirement that all antibodies selected for intracellular cytokine staining must have high binding affinity and specificity for cytokinespecific epitopes that are formed subsequent to the particular fixation

11 Immunomonitoring of Patients Receiving Cancer Vaccines 293 and permeabilization conditions used in the assay procedure [134]. Several different commercial sources of prepackaged fixation and permeabilization buffer kits are available, as well as anticytokine and anticell, surface antigen monoclonal antibodies optimized and selected for use with these vendor-specific fixation and permeabilization reagents. To date there has been no careful comparison of different sources of vendor-specific reagents (i.e., fixation=permeabilization buffers plus associated antibody sets) to each other, and there is no published data systematically comparing the different commercial reagent systems. However, in the absence of this type of empirical comparative analysis it is generally advisable to pair selected anticytokine antibodies from a given vendor to the fixation= permeabilization buffers from the same vendor. Often small alterations in the fixation and permeabilization steps, such as changes in buffer concentration or time course of the processing steps, can also alter the optimal titer and performances of the anticytokine and cell surface specific antibodies used. More commonly, antibodies to different cell surface lineage-specific, lineage subset, or activation antigens from several different vendors can be used on cells previously processed with a particular selected set of fixation and permeabilization buffers. Where this proves not to be the case, it is usually necessary to stain the cells with all the surface phenotype antibodies prior to the fixation and permeabilization step [134]. The CFC assay can be performed on whole blood samples as well as Ficoll-separated PBMC and cell suspensions collected from lymph nodes. Whole blood collection should be done in sodium heparin since other anticoagulants can chelate calcium and significantly lower the functional cytokine responses of stimulated lymphocytes. Although whole blood can be stored prior to use (room temperature), storage times beyond 8 h severely reduce antigen-presenting cell (APC) function, and this can significantly compromise CFC responses when using whole protein antigens. The effective use of frozen PBMC in the CFC assay, as with the ELISPOT assay, remains problematic for some types of samples and antigens. Freezing PBMC lowers the T cell response to protein antigens, presumably due to attenuation of APC numbers and=or function [38]. MHC-matched, selected peptides or peptide cocktails do not require processing before binding to MHC molecules, and thus can still be successfully used to trigger both CD4 þ and CD8 þ responses from frozen archived cells [38]. The stimulation culture system itself can be adapted to provide more optimal costimulation of antigen-specific T cell responses by the addition of costimulatory antibodies such as CD28 and CD49d [80]. The antigen stimulated culture system can also be extended up to

12 294 E. B. Walker and M. L. (Nora) Disis h with the use of lower brefeldin A concentrations (5 mg=ml), which are maintained throughout the last h of the culture period. This longer (overnight) culture period is often logistically more practical when patient samples are received late in the day, and the results are equivalent to those observed using the shorter induction cycle. In some laboratories, computer-regulated thermocycling devices or water baths have recently been employed to maintain the antigen plus brefeldin A induction cycle for 4 h at 37 C, followed by the overnight maintenance of the sample at 4 C. This automated approach allows for effective harvest and processing of the samples the following day [81]. As with any biological assay, both negative and positive system suitability controls are normally used. For the CFC assay the positive control system used to trigger T cell responses serves the dual purpose of demonstrating the immune competence of the donor cells and is used to help set gain and voltage settings on the instrument to define the two-color fluorescence regions in which antigen-stimulated cells should be found in the final acquisition and analysis histograms. The selection of the best positive control system is still an unresolved subject of much debate, but various T cell mitogens or superantigens such as SEB [71] are favored approaches to this problem. The CFC assay was demonstrated to be very reproducible in a recent report. In a CMV-specific CFC assay system the overall coefficient of variation was within 5% for intraassay variability and within 20% for interassay variability [81]. Importantly, where intersite variability has been observed it could be attributed to operatordependent differential gating during analysis of the flow cytometry data [134]. Two data analysis procedures are important in maintaining reproducibility of data acquisition and analysis using the CFC assay. First, it is important to note that both CD4 þ and CD8 þ T cells will downmodulate CD4 and CD8 cell surface molecules subsequent to recent antigen stimulation [134]. Frequently CD4 dim or CD8 dim lymphocytes are minor populations of the total CD4 þ and CD8 þ subsets, but they are often highly enriched for activated effector T cells, which are positive for cytokine expression. Exclusion of those dim CD4- and CD8- positive T cells can significantly lower the overall percentage of cytokine-positive T cells enumerated [38,82]. A second procedure, which significantly improves the robust character of the CFC assay for whole blood analysis, is the use of pan-platelet (CD62p) and panmonocyte (CD33) monoclonal antibodies in any multiparameter (multifluorescence) acquisition and analysis. Background cytokine staining is often increased due to activation of platelets and monocytes. Using an exclusion or dump channel in the acquisition

13 Immunomonitoring of Patients Receiving Cancer Vaccines 295 and=or analysis of CFC data to eliminate CD62p- and CD33- positive cells simplifies and makes the quantitation of cytokine-positive cells more uniform [81]. Clinical Application and Correlation Until recently the CFC assay has been utilized primarily to analyze human T lymphocyte responses to different infectious disease agents such as HIV [83 86], CMV [72,73,81,87,88], EBV [28,89], and several others, including hepatitis C virus [90], mycobacterium [91], and herpes virus [92]. Examples of clinical correlation with CFC data have been described in this now extensive literature. Thus, Pitcher and coworkers [83] demonstrated that HIV patients could be stratified into populations that maintained a detectable CD4 response to HIV antigens by CFC analysis and those that did not. In this study, all HIVpositive nonprogressors were in the test population with measurable anti-hiv CD4 þ T cell responses, while about half of the patients with progressive disease were not responsive to HIV antigen stimulation as measured by CFC. In another series of human studies, HIV-infected patients with cytomegalovirus (CMV)-associated end organ disease showed a correlated loss of CD4 þ T cell IFN-g responses to CMV antigen stimulation [88,93,94]. The CFC assay has also been used to monitor immune response in cancer vaccine settings. Several recent reports have demonstrated that CFC assay procedures can be effectively employed to detect MART-1 and tyrosinase stimulated CD8 þ T cell responses in melanoma patients [95], MUC-1 mucin antigen-triggered CD4 þ T cell responses in patients with various solid tumors [96], and CD4 þ T cell cytokine responses in multiple myeloma patients vaccinated with an immunoglobulin idiotype immunogen [97]. There have been few examples of the correlation of antigen-specific CFC responses and clinical outcome in cancer immunotherapy patients. Patients with stage IV malignant melanoma, vaccinated with an SRL 172 vaccine, were monitored by CFC analysis for antigen-specific lymphocyte activation [98]. In this study the expression of intracellular IL-2 production was associated with improved survival. Reinartz and colleagues analyzed antigen-directed intracellular cytokine synthesis at different time points during immunization of ovarian cancer patients with the anti-idiotype vaccine ACA125. Late in the vaccine course they observed a Th2 pattern of cytokine production that correlated with anti-anti-idiotype antibody production and prolonged survival [99]. The increased use of the CFC assay to monitor immune function of cancer patients in immunotherapy clinical trials will provide new

14 296 E. B. Walker and M. L. (Nora) Disis opportunities to analyze the statistical correlation between clinical outcome and CFC characterization of effector T cell responses in the near future. Sensitivity Comparison to Other Assays To date there have been few well controlled studies comparing CFC versus ELISPOT analysis using the same test population(s) of cells. As mentioned previously, where this has been reported the results suggest that the two assay procedures produce either equivalent results [68] or else demonstrate that the CFC assay is generally more sensitive, in the sense of counting more cytokine-positive cells above the background levels measured in appropriate negative controls [28,100]. Thus, while the background levels of cell-associated cytokine synthesis may minimally be in the % (percentange of gated CD8 þ or CD4 þ T cells) range for the fluorescence-based CFC assay, it also appears the assay may be more sensitive than the ELISPOT assay by virtue of its higher efficiency of detection and counting of cytokinepositive events including weakly positive events. The limit of detection studies used to establish background (noise) levels of cytokine detection of less than 0.01% for the ELISPOT assay may prove to be artificial by virtue of their design since they measured cytokine secretion by potent cytokine-producing cell lines titered at increasingly lower concentrations into quiescent syngeneic PBMC. This experimental approach may artificially provide for an optimally potent test population of effector T cells titered against an optimally quiescent background population of unresponsive PBMCs. As previously indicated, it is also questionable whether detected background or noise levels of cytokine expression by antigen-unstimulated negative controls in the ELISPOT assay are normally below a range of % of T cells in assay procedures using more commonly employed test populations of cells, such as PBMC from vaccinated donors or combined cultures of IVS T cells and antigen-pulsed, autologous dendritic cells. Additional well-controlled comparative studies need to be carried out to determine if the relative detection sensitivity of the two assays truly differs. Ultimately the question of comparative assay sensitivity may prove to be somewhat unimportant since it has recently been estimated that high frequencies of circulating tumor antigen specific memory T cells (1% of CD8 þ T cells) may be required for protective antitumor immunity [17,26,27]. Both the CFC and ELISPOT assays are fully capable of detecting antigen-specific T cell responses above 0.05% of circulating CD8 þ T lymphocytes. Presently the limited amount of data comparing CFC and ELISPOT sensitivity

15 Immunomonitoring of Patients Receiving Cancer Vaccines 297 in monitoring T cell responses of cancer immunotherapy patients suggests this will be an active area of interest in the near future. Both assays can now be somewhat automated, the time course and logistics required to perform them are very manageable, and both assays appear reproducible when proper controls and validation procedures are used. However, only the CFC assay provides a data acquisition environment, which allows for the simultaneous, detailed, multiparameter phenotype analysis of cytokine-positive effector T cells. TETRAMER ASSAY Operational Method The fluorescence-based tetramer binding assay is mechanistically possible because T lymphocytes recognize processed antigenic peptides in the context of syngeneic MHC class I (CD8 þ T cells), or MHC class II (CD4 þ T cells) molecules. Thus, in the case of CD8 þ T cells, the TCR recognizes an MHC class I complex composed of an allele-specific HLA heavy chain, a specific nine-amino-acid peptide, and a b2 microglobulin light chain. Early developmental studies were based on the concept that if such HLA allele-restricted, antigen-specific molecular complexes could be synthesized and conjugated with appropriate fluorochromes they could, in principle, be used to selectively bind to MHC-restricted, peptide-antigen specific effector T cells. The first step in designing such a high avidity soluble MHC molecular complex was achieved by Garboczi and coworkers [101], who were successful in folding recombinant MHC heavy and light (b2 microglobulin) chains into a TCR-binding complex with noncovalently bound 8 10 amino acid peptide antigens. The binding of such peptide sequences is allele dependent, and thus a peptide sequence that binds optimally with a particular HLA allele (HLA-A2) may not bind effectively to a disparate HLA allele (HLA-B35). Tetramer reagent development was further advanced when it was demonstrated that the binding affinity of many such monomeric MHC complex arrays for their putative cognate TCR molecular structures was not strong enough to allow detection by conventional assays. Work by Altman and coworkers [102] modified the recombinant heavy chain to include an enzymatic biotinylation site. With the addition of strepavidin, high avidity tetrameric molecular complexes (tetravalent) were formed. The conjugation of fluorochromes like phycoerytherin (PE) to this quaternary TCRbinding complex resulted in a powerful new approach to the quantitative assessment of antigen-specific T lymphocytes using flow cytometry analysis. Until recently most tetramer flow cytometry

16 298 E. B. Walker and M. L. (Nora) Disis analysis has been performed using MHC class I peptide tetramers for CD8 þ T lymphocyte staining, but recent reports also describe quantitation of antigen-specific CD4 þ T cell responses using MHC class II peptide tetramer molecular constructs [ ]. The use of fluorochrome-conjugated tetramer reagents for flow cytometry analysis is an especially powerful immunophenotyping technique in the context of three- and four-color flow cytometry, where CD8 þ cells can be interrogated for the simultaneous expression of other memory, trafficking, or activation cell surface markers. MHC class I restricted, tetramer-based flow cytometry analysis typically involves staining a minimum of 10 6 PBMC per test condition with primarily PE-conjugated tetramers using an optimal predetermined concentration of reagent for 30 to 60 min at room temperature. Cells are washed thoroughly and subsequently stained with fluorochrome-conjugated anti-cd8 monoclonal antibody, and (in the case of four-color analysis) two other cell surface, epitope-specific monoclonal antibodies of interest. Frequently, a cocktail of lineagespecific antibodies, such as CD14=CD19=CD56 (all conjugated to a single common fluorochrome), will be added to allow for exclusion or negative selection of unwanted cells, which may nonspecifically bind low levels of the tetramer reagent stain. Lymphocytes are subsequently gated based on standard forward angle and 90 light scatter criteria to eliminate aggregates and apoptotic=necrotic cells; and non- CD8 þ T cells are excluded based on staining by the lineage-specific cocktail stain. Propidium iodide (PI) may also be added to more definitively exclude dead cells if the sample is to be analyzed without fixation. Commonly, tetramer-stained, multiparameter cell samples can be fixed in paraformaldehyde and analyzed up to 48 h later. PEconjugated, tetramer-specific staining is typically very bright, and truly positive cells are generally well clustered with up to 2 3 logs separation from the negative control regions of the usual two parameter histogram (CD8 versus tetramer staining) used for final data analysis. Methodological and Reagent Issues It is important to periodically confirm the specificity and level of staining of putative MHC class I peptide-specific tetramer constructs using known antigen-specific T cell clones, since in many cases degradation of noncovalent tetramer complexes can occur within a few weeks [134]. In addition to the intrinsic instability of many MHC class I peptide combinations, there are several other key technical issues that can affect tetramer staining. Staining temperature significantly

17 Immunomonitoring of Patients Receiving Cancer Vaccines 299 influences the level of tetramer staining [ ]. Tetramer staining of T cells at 4 C results in low avidity, cross-reactive binding interactions, which reduces the overall level of tetramer-specific fluorescence [134]. Conversely, tetramer staining at 23 C (room temperature) or 37 C favors higher avidity tetramer-tcr binding interactions. Since 37 C staining also promotes tetramer-tcr complex internalization, optimal staining effects are most often observed at room temperature for min. A very critical variable that is frequently overlooked or ignored is the fact that counterstaining with a particular anti-cd8 or anti-cd3 or -CD4 monoclonal antibody, or changes in the concentration of a given anti-cd8 antibody, can significantly alter the level of tetramer binding in a multiple parameter (multiple antibody) staining procedure [109, 110, 134]. Depending on the CD8 epitope recognized by a given anti-cd8 monoclonal reagent and the antibody concentration, the correlated tetramer staining intensity can be inhibited, unaffected, or even increased [109, 110, 134]. Tetramer reagents should be titered to determine optimal staining concentrations. Using predetermined optimal tetramer concentrations, it can be demonstrated by subsequent titration analysis of the CD8 monoclonal antibody employed that the quantitation of tetramer-positive events measured by flow cytometry analysis can vary over as much as a two- to fourfold range, depending on the concentration and epitope specificity of a given counter-staining anti-cd8 monoclonal antibody. Presently there are very limited commercial sources of MHC class I peptide tetramers these reagents are either custom made using HLA allele and peptide specificities of interest to the laboratory purchasing the product, or a limited number of more commonly used HLA-A2 restricted peptide combinations are sold off the shelf. As a result, this reagent platform is very expensive and limited in its range of application. Even with the use of commercially available tetramers produced under fairly stringent conditions of specificity and titration analysis, there can be significant levels of nonspecific background staining. As mentioned, it is often critical to use an exclusion or dump channel in flow cytometry analysis to gate out cells such as monocytes, which are frequently very sticky for tetramer stains and produce high background levels of nonspecific staining. This is particularly true of analysis performed on freshly collected intact PBMC and thawed PBMC. The use of frozen samples greatly facilitates analysis of multiple samples from a clinical trial since cells can be stored and batched for analysis, thus lowering the effects of inter-assay variability. Several studies demonstrate that tetramer analysis is very effective and reproducible using cryopreserved cells; but the use of frozen cells must be carefully controlled to prevent overcounting of

18 300 E. B. Walker and M. L. (Nora) Disis positive cells due to higher overall levels of nonspecific tetramer binding. Clinical Application and Correlation Fluorescent tetramer complexes were initially used primarily to measure antiviral T cell responses. AIDS patients were monitored using tetramers specific for HIV gag, pol, and env epitopes the resulting studies demonstrated a strong inverse relationship between HIV-specific CTLs and viral load [111] and showed the need for sustained viral antigen expression to maintain long-term circulating CTL numbers [112,113]. Hepatitis C antiviral CD8 þ T cell responses have been followed with HCV peptide-specific tetramers both in the peripheral blood and liver [114], and in HTVLI patients tetramer analysis detected expansion of a CTL subset specific for an HLA-A2 restricted peptide epitope of the HTLVI tax protein [115]. Tetramer studies in EBV [116] and influenza [117] demonstrate high levels of virus-specific CTL expansion in acute infections and long-term persistence of lower virus-specific CTL frequencies in healthy donors. Romero and coworkers reported the first study of the ex vivo quantitation of tumor antigen specific T cells in cancer patients using fluorescenated tetramer reagents [118]. In this report, HLA- A2 restricted tetramers containing either the tyrosinase peptide or the Melan-A peptide analogue A27L were used in combination with anti-cd8 and anti-cd3 mabs to detect tumor antigen specific T cells present in metastatic lymph nodes. In this study no tyrosinase-reactive CD8 þ T lymphocytes were detected; however, high levels of Melan-A responsive CD8 þ T cells were found in nine lymph nodes resected from six HLA-A2 positive melanoma patients. There were no detectable levels of tetramer-positive CD8 þ T cells in normal or micrometastatic lymph nodes. This result clearly suggested the ability of the assay to detect the accumulation of tumor-antigenreactive CD8 þ T lymphocytes in metastatic foci within the lymph node. Interestingly, CD8 þ T cells detected using HLA-A2 binding Melan-A=MART-1 peptide-specific tetramers have been found in both melanoma patients and normal controls [83]. Notably, the Melan- A specific T cells in normal donors displayed a so-called naïve cell surface phenotype (CD45RA hi =CD45RO lo ), while Melan-A=MART-1 tetramer-positive CD8 þ T lymphocytes from melanoma patients were of the memory phenotype, CD45RA lo =CD45RO hi [83]. Very recently there have been a series of published reports clearly demonstrating that HLA-restricted, peptide-specific tetramers can be effectively used to detect upregulation of antigen-specific T cell responses in

19 Immunomonitoring of Patients Receiving Cancer Vaccines 301 vaccinated cancer patients [108, ]. However, although tetramer analysis can detect postvaccine therapy increases in circulating antigen-specific T cells, these measured increases are seldom associated with clinical outcome [78,121], including any real correlation with tumor regression [23,78,119]. In a recent clinical study where more detailed phenotyping of tetramer-positive CD8 þ T cells was carried out in unimmunized melanoma patients, both CD45RA hi =CD45RO lo functionally anergic T cells and CD45RO hi =CD45RA lo memory=effector T cells were detected by tetramer staining in the same patient. In the same report, other unimmunized cancer patients exhibited either CD45RA hi =CD45RO lo, or CD45RO hi =CD45RA lo CD8 þ T lymphocytes that were also positive for MART-1 or tyrosinase tetramer staining [95]. In related studies, unimmunized melanoma patients had either predominantly CD45RA lo, Melan-A=MART-1 tetramer-specific CD8 þ T cells, or they exhibited both CD45RA lo and CD45RA hi Melan-A=MART-1 tetramerspecific CD8 þ T cells prior to immunization [124,125]. After Melan- A=MART-1 immunization the majority of Melan-A tetramer-positive T cells expressed the CD45RA lo memory phenotype in immunized patients [124,125]. Sensitivity Comparison to Other Assays Various groups have published experimental results showing the lower effective limit for the detection of tetramer staining of CD8 þ T cells in PBMC to be 0.01% [106,114,115,126,127], 0.02% [117], and 0.04% [66], respectively. These values are comparable to the lower limit of detection reported for the CFC assay. They are due in part to the highly specific binding characteristics of the tetramer reagents, and the fact that the effective use of flow cytometry dump channels (to exclude nonspecific binding to PBMC subsets such as monocytes) significantly reduces nonspecific background staining. The characteristic bright fluorescence of the PE-conjugated tetramer reagents usually gives a one to two log separation between positive and negative events measured by flow cytometry, and this increases sensitivity (as with the CFC assay) by increasing the efficiency of counting of positively stained cells especially cells that are weakly positive. Evaluating the ultimate importance of tetramer staining as an immunomonitoring procedure to characterize anti-taa immune response in cancer vaccine patients is at present a complex issue. This is due to the observations made in a series of reports in which tetramer quantitation of antigen-specific T cells was compared to LDA and ELISPOT analysis [17,29,117,128], and to CFC enumeration of T