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The Journal of Immunology, 2004, 172: 6107-6114.
Copyright © 2004 by The American Association of Immunologists

The Size and Phenotype of Virus-Specific T Cell Populations Is Determined by Repetitive Antigenic Stimulation and Environmental Cytokines 1

Laila E. Gamadia*,{dagger}, Ester M. M. van Leeuwen*,{dagger}, Ester B. M. Remmerswaal{dagger}, Si-La Yong{dagger}, Sugianto Surachno*, Pauline M. E. Wertheim-van Dillen{ddagger}, Ineke J. M. ten Berge*,§ and René A. W. van Lier2,{dagger}

* Renal Transplant Unit, Department of Internal Medicine, {dagger} Laboratory for Experimental Immunology, {ddagger} Department of Virology, and § Division of Clinical Immunology and Rheumatology, Department of Internal Medicine, Academic Medical Centre, University of Amsterdam, Amsterdam, The Netherlands


   Abstract
Top
 Abstract
Introduction
 Materials and Methods
 Results
 Discussion
 References
 
Based on the expression of the TNFR SFP CD27, two Ag-primed CD8+ T cell subsets can be discerned in the circulation of healthy individuals: CD27+ T cells that produce a variety of cytokines but do not display immediate cytolytic activity; and cytotoxic CD27– T cells, which secrete only IFN-{gamma} and TNF-{alpha}. The mechanism that controls the generation of these different phenotypes is unknown. We show that CMV reactivation not only increases the number of virus-specific T cells but also induces their transition from a CD27+ to a CD27– phenotype. In support of a relation between pool size and phenotype in a cohort of latently infected individuals, the number of Ag-specific CD27– CD8+ T cells was found to be linearly related to the total number of CMV-specific CD8+ T cells. In vitro studies revealed that the acquisition of the CD27– phenotype on CMV-specific T cells depended on the interaction of CD27 with its cellular ligand, CD70. Expression of CD70 was proportional to the amount of antigenic stimulation and blocked by the CD4+ T cell-derived cytokine IL-21. Thus, induction of CD70, which may vary in distinct viral infections, appears to be a key factor in determining the size and phenotype of the CMV-specific T cell population in latently infected individuals.


   Introduction
Top
 Abstract
 Introduction
Materials and Methods
 Results
 Discussion
 References
 
T cell memory is established after clearance of acute infection and maintained lifelong. The generation of CD8+ memory T cells is supposedly by the survival of a number of Ag-stimulated effector T cells that are generated early in the primary adaptive immune response to the causative pathogen. What determines a primary induced CD8+ T lymphocyte to survive into a long-lived memory cell is as yet unclear (1). Primed T cells in the peripheral blood of healthy individuals show considerable phenotypic and functional heterogeneity (2). At one side of the spectrum are CD45R0+CD28+CD27+ T cells that produce a large variety of cytokines, but are unable to execute cytolysis without prior in vitro culture (3). These cells have been defined as memory-type T cells because of their ability to clonally expand in response to recall Ags in the absence of helper T cell-derived cytokines. Based on the expression of the chemokine receptor CCR7, this population may be subdivided into CCR7+ central memory T cells that have the ability to migrate to secondary lymphoid organs and CCR7– effector memory T cells that do not have this ability (4). It is unclear whether CCR7+ and CCR7– T cells are also functionally distinct (5, 6). Separate from these memory-type cells is a population of CD45RA+CCR7–CD28–CD27– T cells (further referred to as CD45RA+CD27– T cells). Based on the expression of CD95 ligand, perforin, and granzyme B and the ability to execute cytotoxicity directly ex vivo, these T cells have been designated effector-type T cells. This population, which is also CD11ahigh and CD57+, increases with age and has a restricted diversity of TCR-V{beta} genes (7, 8, 9).

T cells with specificities for different persistent viruses vary in phenotype and function. For instance, EBV-specific T cells are predominantly CD45R0+CD28+CD27+, and in asymptomatic HIV-carriers, HIV-specific T cells are mostly CD45R0+CD28–CD27+, contrasting the phenotype of CMV-reactive T cells that are predominantly CD45RA+CD27– (10). From these findings, it was postulated that viruses might induce phenotypically distinct T cells (10). However, it is clear that the selection for phenotypes is not absolute because, e.g., in healthy CMV carriers during latency, virus-specific T cells with either CD45R0+CD27+ and CD45RA+CD27– surface profiles can be found (10, 11). Analysis of virus-specific T cells during acute infections with EBV, HIV, or CMV revealed that early effector T cells are clearly distinct from the effector-type CD45RA+CD27– T cells found in latently infected persons. T cells expanding early in immune responses have CD45R0, CD28, CD27 surface expression, express the G1 phase-associated nuclear Ag Ki-67, and contain perforin and granzyme B (12, 13, 14). These observations raised questions about the generation of CD45RA+CD27– T cells and their role in protective immunity (10).

Although it was originally proposed that CD45RA+CD28–CD27– T cells had entered a state of senescence (15), telomere length analyses revealed similar replicative histories in vivo of CD45R0+CD27+ and CD45RA+CD27– CD8+ T cells (8). Moreover, it was shown that CD45RA+CD27– T cells were unable to divide when stimulatory mAbs to CD3 and CD28 are combined to activate these cells (2). Still, CMV-specific T cells with this phenotype potently expanded when stimulated by specific MHC/peptide ligands in the presence of helper T cell-derived cytokines (16, 17). These latter experiments implied that although CD45RA+CD27– T cells are fully differentiated T cells where it concerns CTL effector functions, they are not terminally differentiated (18) in the sense of having lost the ability to yield progeny.

Here, we studied patients with well-documented CMV reactivation to investigate the generation of CD27– T cells in vivo and their role in maintaining CMV latency. We show that effector-type T cells can expand in vivo when viral replication occurs. Moreover, antigenic load and helper T cell-derived cytokines are key regulators of the phenotype of virus-specific T cells during latency, likely through regulation of the CD27 ligand, CD70.


   Materials and Methods
Top
 Abstract
 Introduction
 Materials and Methods
Results
 Discussion
 References
 
Five HLA-A2-positive and 1 HLA-B7-positive CMV-seropositive renal transplant recipients of a kidney of either a CMV-seropositive donor (2) or a CMV-seronegative donor (4) were longitudinally studied. Basic immunosuppressive therapy consisted of cyclosporin A, blood trough level aimed at 150 ng/ml, mycophenolate mofetil (1000 mg twice daily), and prednisone (10 mg daily). Viral replication was monitored longitudinally by quantitative PCR from the time point of transplantation, and reactivation was determined by detectable viral loads above the cutoff point of 80 copies/ml. All subjects experienced asymptomatic CMV reactivation and no antiviral treatment was given. Heparinized peripheral blood samples were collected before transplantation and weekly during 17 wk after transplantation, whereafter samples were collected once a month. PBMC were isolated using standard density gradient centrifugation techniques and subsequently cryopreserved. In addition, CMV-specific T cells were characterized in 11 healthy individuals and 36 renal transplant recipients on basic immunosuppressive therapy either before or after reactivation as measured by PCR. All patients gave written informed consent, and the local medical ethical committee approved the study.

Peptides

The HLA-A2-binding CMV pp65-derived peptide NLVPMVATV and the HLA-B7-binding CMV pp65-derived peptide TPRVTGGGAM were purchased from the IHB-LUMC peptide synthesis library facility (Department of Immunohaematology and Blood Bank, Leids Universitair Medisch Centrum, Leiden, The Netherlands).

Generation of tetrameric complexes

Tetrameric complexes were manufactured at the tetramer facility of Sanquin, The Netherlands, and generated essentially as described by Altman et al. (19) In brief, purified HLA-A2.1 H chain or HLA-B7.2 H chain and {beta}2-microglobulin were synthesized using a prokaryotic expression system (pET; Novagen, Milwaukee, WI). The H chain was modified by deletion of the transmembrane-cytosolic tail and C-terminal addition of a sequence containing the BirA enzymatic biotinylation site. The HLA-A2.1 binding CMV pp65-derived peptide NLVPMVATV and the HLA-B7.2-derived peptide TPRVTGGGAM were used for refolding. Monomeric complexes were concentrated, biotinylated by BirA (expressed using the pET expression system, and purified using Clontech cobalt beads; Clontech Laboratories, Palo Alto, CA) in the presence of biotin (Molecular Probes, Eugene, OR), ATP (Sigma-Aldrich, St. Louis, MO), and MgC12. The biotinylated product was separated from free biotin by fast protein liquid chromatography using a Superdex 200 HR16/60 column (Amersham Pharmacia, Little Chalfont, U.K.). Streptavidin-allophycocyanin conjugate (Molecular Probes) was added in a 1:4 molar ratio, and subsequently tetramers were fast protein liquid chromatography purified using the same column.

CFSE labeling

PBMCs were pelleted and resuspended in PBS at a final concentration of 5–10 x 106 cells/ml. PBMCs were labeled with 2.5 µM CFSE (Molecular Probes) in PBS for 8 min, shaking at 37°C. Cells were washed and subsequently resuspended in IMDM supplemented with 10% human pool serum, antibiotics, and 3.57 x 10–4 % (v/v) 2-ME (Merck, West Point, PA; culture medium).

Culture and stimulation of cells

CFSE-labeled cells were cultured in culture medium for 5 days in 24-well plates at a concentration of 0.5–1 x 106 cells/ml. CMV pp65-derived peptide was added at a final concentration of 1.25 µg/ml. CMV-Ag (inactivated whole virus, 10 µl/ml; Microbix Biosystems, Toronto, Canada) was used to stimulate cells. Furthermore, for stimulation, IL-2 (50 U/ml; Biotest, Dreieich, Germany), IL-15 (3 ng/ml; R&D Systems, Abingdon, U.K.), and IL-21 (50 ng/ml; Zymogenetics, Seattle, WA), were added. For blocking experiments, we used CD70 Ab (clone 2F2; Ref.20) at concentrations of 40 µg/ml. For dose-dependent analysis of CD70 expression, 1.25 µg of peptide and 10 µl of CMV-Ag/ml were titrated. Flow cytometric analysis was performed before culture and after 5 days.

Immunofluorescent staining and flow cytometry of CMV-specific CD8+ T cells

Thawed PBMC were resuspended in RPMI containing 10% FCS and antibiotics; 200,000 PBMC were incubated with fluorescent label-conjugated mAbs (concentrations according to manufacturer’s instructions) and an appropriate concentration of tetrameric complexes. Negative controls to validate specificity of the CMV-peptide-tetrameric complexes consisted of HLA-A2.1/HLAB7.2-negative CMV-seropositive or HLA-A2.1/HLA-B7.2-positive CMV-seronegative healthy individuals and renal transplant recipients. Negative controls always showed tetramer staining of <0.01% of total lymphocytes (data not shown). For staining with the mouse anti-human CCR7 mAb, a three-step staining protocol was performed consisting of incubation with the CCR7 Ab (BD PharMingen, San Diego, CA), for 30 min, washing, incubation with biotinylated goat anti-mouse IgM (BD PharMingen) for 30 min, and incubation with 10% (v/v) normal mouse serum (Sanquin, Amsterdam, The Netherlands) followed by incubation with streptavidin-PE and directly conjugated mAbs and tetrameric complexes for 30 min. For staining with the mouse anti-human mAb CD70 (clone 2F2) a two-step staining protocol was performed consisting of incubation with the CD70 Ab for 30 min, washing, incubation with FITC-conjugated anti-mouse IgG1 Abs, washing, and incubation with 10% (v/v) normal mouse serum followed by incubation of directly conjugated mAbs. Analyses consisted of allophycocyanin-conjugated tetramers and CD8-PerCP (BD Biosciences, San Jose, CA) in combination with either CD28 (Sanquin) and CD27 (BD Biosciences), CCR7 and CD45RA, CD27 and CD45RA (BD Biosciences) and CD45RA and CD45R0 (BD Biosciences), CD38 (BD Biosciences), and CD70, all combinations in FITC and PE, and additional stainings for CD70 expression were done with CD70 in combination with CD4-PE, CD8-allophycocyanin and CD19-PerCP.

CMV-PCR

Quantitative PCR was performed in EDTA whole blood samples as described for plasma or serum (21).

Statistical analyses

Linear regression analysis using a mixed model was performed for the absolute numbers of total CMV-specific CD8+ T cells in relation with total CD27–CMV-specific CD8+ T cells, and for the percentage CMV-specific CD8+ T cells and the percentage of CD27– of CMV-specific CD8+ T cells taken as a logit. Subsequently, the regression coefficient was calculated and tested for significance. Repeated observations were analyzed by paired Student’s t test or a repeated measures ANOVA; p values < 0.05 were considered statistically significant.


   Results
Top
 Abstract
 Introduction
 Materials and Methods
 Results
Discussion
 References
 
CMV reactivation induces quantitative and qualitative changes in the CD8+ virus-specific T cell compartment

In healthy individuals and renal transplant recipients during latency, CMV-specific CD8+ T cells may have different predominant phenotypes ranging from CD45R0+CD28+CD27+ (memory-type, patient 1; Fig. 1a; t = 0) to CD45RA+CD45R0dull/–CD27– (effector-type, patient 2; Fig. 1d; t = 0) (10, 11). The fate of these distinct CMV-specific T cell populations was followed during CMV reactivation.



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  		FIGURE 1. Quantitative and qualitative changes of CMV-specific CD8+ memory cells during reactivation. a–c, Differentiation of CMV-specific CD8+ T cells in one patient with a starting population of CD45R0+CD27+CD28+ CMV-specific cells. d–f, Differentiation of CMV-specific CD8+ T cells in one patient with a starting population of CD45RA+CD27–CD28– cells. Time defined as t = 0 before reactivation, t = 1 during reactivation at peak PCR value, and t = 2 at first time point after reactivation. All plots gated on CD8+ T cells. CMV-specific CD8+ T cells as defined by specific tetramer staining plotted in black; total CD8+ T cells are plotted in gray. Percentage of CMV-specific CD8+ T cells of patients 1 and 2, respectively, at time points 0–2: Patient 1, 0.23, 0.42, and 2.55%; patient 2, 2.21, 2.83, and 2.96%. a and d: x-axis log fluorescence CD27-FITC; y-axis log fluorescence CD28-PE; percentages depicted are the percentage of CD27– cells of total tetramer+ cells. b and e: x-axis log fluorescence CD45R0-PE; y-axis log fluorescence CD45RA-FITC; c and f: x-axis log fluorescence CD38-PE; y-axis log fluorescence HLA-DR-FITC.

 
In the patients with a CD45RA–CD28+CD27+ CD8+ T cell phenotype before reactivation, a substantial number of these cells lost expression of CD28 and CD27 (Fig. 1, row a). Longitudinal analysis of CMV-specific CD8+CD27– T cells in these individuals showed that this subset increased immediately when replication of virus occurs (Fig. 2a, CD27–tetramer+ T cells in relation to viral load in one representative individual; Fig. 2b, CD27–tetramer+ T cells in all individuals; median before reactivation, 26.5%; range, 0.0 to 91.1%; median at the time point of reactivation, 45.22%) and keeps on doing so long after viral replication has ceased (median at the first time point after reactivation, 66.35%; median at the last time point measured, >6 mo after reactivation, 72.73%; p = 0.039). In the individuals with a predominant effector, CD45RA+CD28–CD27– CD8+ T cell phenotype, there was an increase in the number of CMV-specific T cells but no phenotypic change of virus-specific CD8+ T cells with regard to the expression of CD28 and CD27 (Fig. 1, row d). To analyze whether this down-regulation of CD27 was not contributable to other factors as immunosuppressive medication upon transplantation, we also analyzed EBV-specific cells in individuals who were CMV and EBV seropositive, experiencing CMV reactivation and not EBV reactivation. As is shown in Fig. 2c, CMV-specific cells increase in CD28–CD27–CD45RA+ cells, whereas EBV-specific cells, that in the individual shown were CD45R0+CD28+CD27+, do not change in surface phenotype after CMV reactivation, implying that this phenomenon is solely due to specific stimulation of CMV-specific cells.



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  		FIGURE 2. Ongoing down-modulation of CD27 on CMV-specific CD8+ T cells after reactivation. a, Longitudinal analysis of CD27 expression on CMV-specific CD8+ T cells in relation to CMV viral load in one representative patient with a CD28+CD27+CD45R0+ phenotype of CMV-specific cells before CMV reactivation. {blacktriangledown}, viral load; •, number of CD27– CMV-specific CD8+ T cells. b, Percentage of CD27– CMV-tetramer+ T cells in all patients during follow-up of reactivation. Time defined as: t = 0, before reactivation; t = 1, during reactivation at peak PCR value; t = 2, first time point after reactivation; and t = 3, >6 mo after reactivation. c, Expression of CD27, CD28, CD45RA, and CD45R0 on CMV-specific cells (left) and EBV-specific cells (right) before and 1 year after CMV reactivation in two representative patients. All plots gated on total tetramer+ T cells.

 
Irrespective of the prereactivation phenotype, the activation markers CD38 and HLA-DR were up-regulated on CMV-specific cells in all patients at the time of viral replication (Fig. 1, rows c and f), as was the expression of CD45R0, reflecting recent proliferation of these cells (Fig. 1, rows b and e). CCR7 expression, documented to be up-regulated upon activation (17, 18), did not change in any of the patients during reactivation (data not shown), although re-expression of this marker could redirect subjected cells to secondary lymphoid organs, rendering them undetectable in peripheral blood.

These data suggest that irrespective of their phenotype, CMV-specific T cells can confer protective immunity in situations of virus reactivation. Moreover, virus reactivation appears not only to increase the size of the virus-specific T cell pool but also, in individuals that start off with memory-type CMV-specific T cells, to induce a change from a predominant CD27+ to a CD27– phenotype.

The CD27– phenotype is correlated with the magnitude of the CD8+ T cell response during latency

The above findings suggested a connection between the number of virus-specific T cells in latency and their phenotype which would accord with the expansion of CMV-specific CD8+CD27– T cells in immunocompromised patients (11, 22), B cell chronic lymphocytic leukemia patients (23), and the elderly (24, 25).

Indeed, close analysis of the total percentage of CD8+ CMV-specific cells as determined by tetramer staining and the expression of CD27 by these cells shows a strong correlation between the loss of CD27 and a higher percentage of CMV-specific CD8+ T cells as determined by tetramer staining, for both renal transplant recipients and healthy control individuals. The correlation of the percentage of CMV-tetramer+CD8+ T cells and the percentage of CD27– of these cells is depicted in Fig. 3a, which shows that after the percentage of tetramer+ cells reaches ~1%, >=50% of these cells are CD27–, whereas virtually all CMV-specific cells are CD27– when a percentage of 2% or higher is reached. Also, when depicted in absolute numbers, the amount of CMV-specific cells is linearly correlated to the amount of CD27– cells (Fig. 3b).



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  		FIGURE 3. High frequencies of CMV-specific CD27– effector cells are correlated to the total amount of CMV-specific CD8+ T cells. a, Correlation of the percentage of total CMV-tetramer+ T cells and the percentage of CD27– of CMV-specific T cells. •, Values in renal transplant recipients before or after reactivation; {circ}, values in healthy control individuals; curve, functional relationship determined by the formula % CD27– = 100 * [(e**(0.226 + 0.86*ln(% tetramer))/(1 + (e**(0.226 + 0.86*ln(% tetramer))]; p = 0.005. b, Correlation of the absolute number of total CMV-tetramer+ T cells and the absolute number CD27– CMV-specific T cells. •, Values in renal transplant recipients; curve, linear relationship determined by absolute number of CMV-tetramer+ cells = 0.001 + (1.46 * absolute number of CMV-tetramer+CD27–) (p = 0.0012).

 
Generation of the CD27– phenotype of virus-specific T cells is dependent on interaction of CD27 with its cellular ligand, CD70

T cells down-modulate CD27 after interaction with its cellular ligand CD70 both in vitro and in vivo (26, 27). In this process, T cells receive costimulatory signals for expansion (26, 27) and acquisition of effector functions (28). To analyze whether virus-induced CD70 expression could account for the down-modulation of CD27 on virus-specific T cells after CMV reactivation, CD70 expression was measured in the reactivating patients. Indeed, whereas during latency most CMV tetramer-binding T cells were CD70 negative (Fig. 4, top left), CD70 transiently increased during reactivation episodes (Fig. 4. middle and right). Accordingly, viral Ag-induced CD70 expression could be induced in vitro, and the magnitude of CD70 expression showed a dose-response relationship with the virus-derived stimulus (Fig. 4, bottom).



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  		FIGURE 4. CD70 expression is induced by Ag and is dose dependent. a, CD70 is expressed on CD8+ and CMV-specific CD8+ T lymphocytes during viral replication in vivo. All plots gated on CD8+ T cells. Time defined as: t = 0, before reactivation; t = 1, during reactivation; and t = 2, after reactivation. Quadrant percentages depicted as percentage CD70+ of tetramer+ cells. b: left, Upon in vitro stimulation with CMV-Ag and CMV-peptide, CD70 expression is induced on CD8+ T cells. x-Axis log fluorescence CFSE; y-axis CD70 expression, gated on CD8+ T cells; total CD8+ T cells plotted in gray, and CMV-specific CD8+ T cells plotted in black. Right, Expression levels of CD70 on CD8+ T cells in various doses of CMV-Ag and peptide stimulation. x-Axis log fluorescence CD70; filled histogram, no stimulus; closed histogram, stimulation with 0.15625 µl/ml CMV-Ag and 0.0195 µg/ml CMV-peptide; dotted histogram, stimulation with 0.625 µl/ml CMV-Ag and 0.0781 µg/ml CMV-peptide; bold histogram, stimulation with 2.5 µl/ml CMV-Ag and 0.3125 µg/ml CMV-peptide.

 
To test whether viral Ag-induced up-regulation of CD70 dictated CD27 expression, T cells from donors with a CD45R0+CD27+ phenotype (Fig. 5a) were cultured for 5 days with specific peptide and CMV-Ag to activate helper T cells in the absence and presence of blocking CD70 mAb. In these cultures, CMV-specific T cells rapidly down-regulated CD27 (Fig. 5b). However, addition of CD70 mAb to these cultures reversed this blocking effect and yielded CD27+ CMV-specific T cells at day 5 of culture. Irrespective of CD27 down-regulation, these cells remained CD28+. The observation that CD28 is down-regulated in vivo during CMV reactivation implies that other signals induce CD28 down-regulation. Helper CD4+ T cells can directly support CD8+ T cell expansion through secretion of stimulatory cytokines and provision of membrane-bound helper factors (29). Furthermore, the homeostatic cytokine IL-15 was shown to be involved in maintenance and activation/functional differentiation of memory CD8+ T cells (30, 31, 32). To analyze a possible qualitative role of different cytokines on CD8+ T cell differentiation, the effects of cytokines, which previously were found to support proliferation of CMV-specific T cells (17), on CD70 and CD27 expression were analyzed. In conjunction with peptide, IL-2 and IL-15 induced CD70 expression on CMV-specific T cells and CD70-dependent down-modulation of CD27 (Fig. 6, a and b). Although IL-21 was reported to stimulate only CD45RA+ thymocytes (33), in our experiments, in an Ag-specific setting, CD45R0+CD8+ T cells were also responsive to IL-21. Interestingly, IL-21, a cytokine presumed to limit effector cell expansion (34), did not induce CD70 expression and supported propagation of CD27+ CMV-specific T cells without differentiation (Fig. 6c).



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  		FIGURE 5. CD70 induces differentiation of CD27+ CMV-specific CD8+ T cells. a, Expression of CD70 on total lymphocytes from the donor shown in Fig. 1a after stimulation with CMV-peptide and CMV-Ag (open histogram) and no stimulus (filled histogram). x-Axis log fluorescence CD70. b, Histogram overlay of CD27 expression on tetramer+CD8+ T cells after 5 days of stimulation with peptide and CMV-Ag with or without blocking of CD70. CD27 expression on CMV-specific CD8+ T cells without blocking of CD70 (filled histogram) and with blocking of CD70 (open histogram). c, Histogram overlay of CD28 expression on tetramer+CD8+ T cells after 5 days of stimulation with peptide and CMV-Ag with or without blocking of CD70. CD28 expression on CMV-specific CD8+ T cells without blocking of CD70 (filled histogram) and with blocking of CD70 (open histogram). One representative donor of four tested is shown.

 


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  		FIGURE 6. CD70 expression is induced by IL-2 and IL-15 but not by IL-21. Left, Expression of CD70 on total lymphocytes after stimulation with CMV-peptide and IL-2, IL-15, or IL-21, respectively (open histogram) and no stimulus (filled histogram). x-Axis log fluorescence CD70. Right, Dot plots of CD27 expression on tetramer+CD8+ T cells after 5 days of stimulation with peptide and IL-2, IL-15, or IL-21, respectively, with or without blocking of CD70. Plots gated on CD8+ T cells. x-Axis log fluorescence CMV-tetramer; y-axis log fluorescence CD27. Histogram overlays of CD27 expression on CMV-specific CD8+ T cells with different stimulations without blocking of CD70 (filled histogram) and with blocking of CD70 (open histogram). One representative donor of four tested is shown.

 

   Discussion
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
References
 
Our results show that in persistent viral infection, reactivation increases the number of memory cells and skews these cells to a CD27– phenotype. In primary infection, the amount of Ag determines the size of the murine CD8+ T cell response. Subsequently, the magnitude of this CD8+ T cell response sets the size of the Ag-specific T cell pool during the memory phase (35, 36). Recently, it was demonstrated that the contraction phase of the primary immune response is independent of the clearance of Ag (37). However, both cytokines present during initial priming and the amount of Ag present after the contraction phase are likely to have an impact on the quality and quantity of the memory cells rendered thereafter (36, 37, 38). It was proposed that different viruses elicit different human CD8+ T cell responses with respect to their surface phenotype (10). EBV and HIV predominantly elicit CD28–/+CD27+CD8+ responses. The analysis of Ag-specific memory CD8+ T cells directed against influenza virus (39, 40), which is completely cleared from the host, reveals that these cells are all CD28+CD27+ and present at low percentages compared with the frequency of CD8+ T cells directed at persistent viruses. CMV-specific responses in both healthy individuals and renal transplant recipients vary in magnitude and phenotype among individuals (10, 11, 16). The strong correlation between the percentage of virus-specific cells in the total CD8+ T cell population and the percentage of CD27– cells of the virus-specific cells indicates that a uniform mechanism takes place that drives both the expansion of virus-specific memory cells and their differentiation to cytotoxic effector-type cells.

In this study, replication of CMV, measured by PCR, could be pinpointed to a very limited period shortly after transplantation, whereas the increase in total CMV-specific cells and the down-modulation of CD27 on these cells was an ongoing process for up to >1 year after transplantation. The ongoing rise in CMV-specific CD8+ T cells therefore is possibly generated by an increase in the number of infected host cells that persistently present Ag to immune cells. The only known mechanism through which CD27 is down-regulated is after interaction with its ligand CD70. CD70 belongs to the TNF family, and its expression on B and T cells can be induced by antigenic stimulation in vitro (41) and in vivo (this study) and appears to be modulated by the cytokine milieu. We do not know at this moment where the interaction between CD27 and CD70, leading to the down-modulation of the former molecule, takes place in vivo during CMV reactivation. Moreover, although we showed that virus-specific T cells can express CD70, we also found that B cells and CD4+ T cells in patients with positive CMV PCRs express considerable amounts of CD70 (data not shown) and therefore may have the ability to trigger and modulate CD27 on CMV-reactive T cells. Our study showed that TCR triggering in combination with IL-2 and IL-15 induced CD70, whereas IL-21 did not give CD70 expression. Accordingly, IL-2 and IL-15 induced expansion of CD27– virus-specific T cells, whereas IL-21 promoted outgrowth of CD27+ cells. Although both IL-2 and IL-21 are helper cell-derived cytokines, the differentiation of CMV-specific memory CD8+ T cells to a CD27– phenotype upon activation with whole CMV-Ag in combination with peptide shows that the role of IL-21 production by CMV-specific CD4+ memory T cells, reported to have a Th1 profile (42), is limited in accordance with previous reports in which IL-21 was shown to be a Th2 cytokine (43). Whereas production of helper cell-derived cytokines as IL-21 and IL-2 is limited to the antigenic activation phase, IL-15 can be produced by a number of lymphoid and nonlymphoid cells (44). It is feasible that viruses elicit distinct helper cell cytokine profiles by differential infection of APC and by different target cell tropism, thereby also determining the number and phenotype of CD8+ T cells during the memory phase (45, 46, 47).

CD27-deficient mice have a reduced ability to form adequate numbers of Ag-specific T cells upon viral infection (48). In contrast, mice that constitutively express CD70 on B cells have a strongly enhanced capacity to form effector T cells in response to viruses, tumors, and protein Ags (Ref.49 and R. Arens, manuscript in preparation). Thus, these mouse models strongly infer that the CD27-CD70 interaction is a major determinant in setting the size of the Ag-specific pool after immunization. The way in which CD27 signaling regulates these processes is incompletely unraveled, but CD27–/– T cells show diminished survival after stimulation (48). An effect of CD27 on T cell survival would be in line with proposed functions of related TNFR family molecules such as OX40 and 4-1BB that increase expression of antiapoptotic molecules such as Bcl-2 and Bcl-xL (50, 51, 52). Indeed, recently we found strong up-regulation of Bcl-xL expression by CD27 triggering (M. van Oosterwijk, manuscript in preparation). On the basis of these observations, it can be postulated that through pathogen-induced expression of CD70 and subsequent triggering of CD27, up-regulation of Bcl-xL and increased survival of Ag-specific T cells, the contraction phase after initial expansion is altered leading to a higher setpoint in memory cell numbers. Simultaneously, through prolonged survival, T cells may receive differentiation signals for a longer period. This proposed link between survival and differentiation may explain the clear correlation we found between CMV-specific T cell numbers and their phenotype. In our in vitro studies, we did not find an effect of CD70 blockade on numbers of Ag-specific T cells, although these Abs were capable of blocking CD27 signaling (Figs. 5 and 6). The lack of effect might be explained by the fact that up-regulation of Bcl-xL may protect T cells from death by neglect, i.e., cytokine shortage, which may be essential in vivo but irrelevant in our cultures where ample cytokines are present.

Taken together, our data imply that the induction and expansion of CD8+ virus-specific T cells and their differentiation to the CD27– effector phenotype is driven by both antigenic load and helper T cell-derived or homeostatic cytokines. We submit that regulation of CD70 expression could be central in this process. Elucidating the differentials inducing and sustaining the CD27– effector phenotype could have a profound impact on immunotherapeutic strategies, especially in disease situations in which strong and persistent cytolytic responses are warranted.


   Acknowledgments
 
We thank Dr. G. J. Weverling for expert statistical analysis, Dr. D. van Baarle for expert technical assistance regarding tetramer manufacturing, and Dr. F. Miedema, Dr. E. Eldering, and Dr. R. Arens for critical reading of the manuscript.


   Footnotes
 
1 This work was supported by Dutch Kidney Foundation Grant C98-1724 (to L.E.G. and E.B.R.). Back

2 Address correspondence and reprint requests to Dr. Rene A.W. van Lier, Academic Medical Centre, University of Amsterdam, L1-152, P.O. Box 22660, 1100 DD Amsterdam, The Netherlands. E-mail address: r.vanlier{at}amc.uva.nl Back

Received for publication October 23, 2003. Accepted for publication March 15, 2004.


   References
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 References
 

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