Human T Cells - The Journal of Immunology

Cytolytic Mechanisms and Expression of Activation-Regulating Receptors on Effector-Type CD8ⴙCD45RAⴙCD27ⴚ Human T Cells Paul A. Baars,1* Laura M. Ribeiro do Couto,*1,2,3 Jeanette H. W. Leusen,† Berend Hooibrink,* Taco W. Kuijpers,* Susanne M. A. Lens,4* and Rene´ A. W. van Lier5* Circulating CD8ⴙ T cells with a CD45RAⴙCD27ⴚ phenotype resemble cytolytic effector cells because they express various cytolytic mediators and are able to execute cytotoxicity without prior stimulation in vitro. We here demonstrate that CD8ⴙCD45RAⴙCD27ⴚ T cells can use both granule exocytosis and Fas/Fas ligand pathways to induce apoptosis in target cells. The availability of these cytolytic mechanisms in circulating T cells suggests that the activity of these cells must be carefully controlled to prevent unwanted tissue damage. For this reason, we analyzed the expression of surface receptors that either enhance or inhibit T cell function. Compared with memory-type cells, effector cells were found to express normal levels of CD3⑀ and TCR␨ and relatively high levels of CD8. CTLA-4 was absent from freshly isolated effector cells, whereas a limited number of unstimulated memory cells expressed this molecule. In line with recent findings on CD8ⴙCD28ⴚ T cells, CD45RAⴙCD27ⴚ T cells were unique in the abundant expression of NK cell-inhibitory receptors, both of Ig superfamily and C-type lectin classes. Binding of NK cell-inhibitory receptors to classical and nonclassical MHC class I molecules may inhibit the activation of the cytolytic machinery induced by either Ag receptor-specific or nonspecific signals in CD8ⴙCD45RAⴙCD27ⴚ T cells. The Journal of Immunology, 2000, 165: 1910 –1917.

E

ffective immune responses against viruses depend for a large part on the induction of CTL that lyse virally infected cells. Studies initially performed in mice have shown that after infection, viral peptide-specific T cells undergo massive expansion and rapidly acquire effector function (1–3). After elimination of the pathogen, most of the CD8⫹ Ag-specific lymphocytes die as a result of apoptosis. However, a portion of specific T cells persists (1) and constitutes a pool of memory cells, which on challenge with the same or a similar pathogen is able to mount a more forceful immune response (4). Recent studies on EBV infection have shown that similar mechanisms are operational in the human system (5). In humans, cell surface marker analysis has been thoroughly used to discern functional distinct subsets of CD8⫹ T cells. Proliferative responses to viral Ags are predominantly confined to the CD45RA⫺CD45R0⫹ subset, which suggests that this population contains memory-type cells (6, 7). In line with this *Department of Immunobiology and Laboratory for Experimental and Clinical Immunology, Academic Medical Center, CLB, University of Amsterdam, Amsterdam, The Netherlands; and †Department of Cell Biology, University Medical Center Utrecht, Utrecht, The Netherlands Received for publication January 18, 2000. Accepted for publication June 6, 2000. The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked advertisement in accordance with 18 U.S.C. Section 1734 solely to indicate this fact. 1

P.A.B. and L.M.R. contributed equally to this study.

2

L.R. was supported by Fundac¸a˜o para a Cieˆncia e a Tecnologia, Portugal (Grant PRAXIS XXI/BD/9156/96). 3 Current address: Department of Molecular Immunology, National Institute of Public Health and Environmental Protection, P.O. Box 1, 3720 BA Bilthoven, The Netherlands.

assumption, cytokine secretion profile, activation requirements, and the expression of adhesion and homing receptors on CD45RA⫺CD45R0⫹ T cells resemble those of Ag-experienced T cells (8, 9). Next to the CD45RA⫺CD45R0⫹ subset, a second subpopulation with features of in vivo priming is found in human peripheral blood. This population is characterized by the absence of the costimulatory molecules CD28 (10) and CD27 (7), the presence of both CD45RA and CD57 Ags (11), and abundant expression of CD11a (12). Because CD8⫹ T cells within this subset express cytolytic mediators such as perforin, granzyme A and B, and Fas ligand (FasL) 6 mRNA and are able to execute cytotoxicity without prior in vitro stimulation, it is suggested that they represent effector cells in vivo (13). Interestingly, although these effector type cells have poor proliferative potential in vitro, it has been documented that these cells increase with age (14). The recent observation (15) that these cells lack the chemokine receptor CCR7 indicates that they do not recirculate through the secondary lymphoid organs but rather migrate to sites of inflammation. Finally, compatible with selection by specific Ag in vivo is the observation that their TCR V␤ repertoire is strongly skewed when compared with either naive or memory CD8⫹ T cells (16). Indeed, recent studies using HLA/viral-peptide tetrameric complexes have demonstrated that specific CD8⫹ T cells with CD28⫺ and CD45RA⫹CD27⫺ phenotypes can be found in EBV, hepatitis C virus, and CMV carriers (17, 18). Studies with in vitro expanded T cell clones and experiments in mutant mice have provided evidence that CTL can exert their effector functions by at least two independent pathways: granule exocytosis and the Fas/FasL pathway (19). The granule exocytosis

4 Current address: Institut Biochimi Universite´ de Lausanne, Chemin de Boveresses 155, 1066 Epalinges, Switzerland. 5

Address correspondence and reprint requests to Dr. Rene´ A. W. van Lier, Department of Immunobiology, Plesmanlaan 125, 1066 CX Amsterdam, The Netherlands. E-mail address: [email protected]

Copyright © 2000 by The American Association of Immunologists

6 Abbreviations used in this paper: FasL, Fas ligand; RT, room temperature; CMA, concanamycin A; NKRs, NK cell-inhibitory receptors.

0022-1767/00/$02.00

The Journal of Immunology pathway involves secretion of granules containing cytotoxic effector molecules onto the surface of target cells. Perforin plays a critical role in this pathway, because it can polymerize to form channel-like structures in target cell membrane, through which granzymes can enter and subsequently activate the death machinery (20). Alternatively, CTLs can use the Fas pathway to kill their target cells. This cytolytic pathway is mainly based on cell-cell interaction between Fas, expressed on the target cell, and FasL, expressed on the CTL (21). Engagement of the Fas receptor results in the aggregation of its intracellular death domains, followed by the activation of several caspases with the ultimate death of the target cell (22). We here show that both granule exocytosis and FasL pathways are readily operational in circulating effector-type CD8⫹ T cells. Furthermore, in search for potential mechanisms that could control the cytotoxic machinery of these cells in vivo, we found normal expression of CD3⑀ and CD3␨ chains and absence of CTLA-4 expression. However, in agreement with recent finding on CD28⫺ T cells (23, 24), a strong increase in the expression of various classes of killer-inhibitory receptors was observed when cells maturate into CD8⫹CD45RA⫹CD27⫺ effector T cells (25).

Materials and Methods Antibodies For the analysis of expression of cell surface and intracellular molecules, the following mouse anti-human mAbs were used: unlabeled Fas2 and CD3, CD27 FITC, and CD3 FITC and biotinylated granzyme B (GB12) (CLB, Amsterdam, The Netherlands); CD45RA PECy5 and TCR␨ FITC (Serotec Kidlington, Oxford, U.K.); CD94, NKG2a, CD158a, CD158b, and CD45RA, CTLA-4 (CD152), all PE labeled (Coulter-Immunotech, Miami, FL); CD8 peridinin chlorophyll protein, CD45R0 APC, CD16 PE, NKB1-FITC (Becton Dickinson, San Jose, CA); unlabeled FasL (Nok-2) (PharMingen, San Diego, CA); unlabeled granzyme A, perforin FITC, (Ho¨lzel Diagnostika, Cologne, Germany); and annexin V FITC (Nexins Research, Hoeven, The Netherlands). As a conjugate for unlabeled or biotinylated mAbs, goat F(ab⬘)2 anti-mouse IgG2a PE (Southern Biotechnology Associates, Birmingham, AL), goat anti-mouse FITC (CLB), and streptavidin red 670 (Life Technologies, Gaithersburg, MD) were used, respectively. Isotype-matched mAb served as controls.

Cells PBMC were isolated from buffy coats of healthy blood donors by density centrifugation with Ficoll-Isopaque (Pharmacia Biotech, Uppsala, Sweden). Subsequently, CD8⫹ T cells (⬎97% TCR␣␤⫹CD8⫹ cells as assessed by flow cytometry; data not shown) were prepared by incubating PBMC with anti-CD8 microbeads (Miltenyi Biotec, Bergisch Gladbach, Germany), followed by positive selection with the VarioMACS (Miltenyi Biotec) according to the manufacturer’s recommendations. For subset purification (7), CD8⫹ T cells were stained with CD45RA and CD27 and sorted on a FACStar (Becton Dickinson) into CD45RA⫹CD27⫹ (naive fraction), CD45RA⫹CD27⫺ (effector fraction), and CD45RA⫺CD27⫹ (memory fraction) populations (⬎95% purity as assessed by flow cytometry; data not shown). For NK isolation, PBMC were stained with CD3 and CD16 mAb and sorted on a FACStar into CD3⫺CD16⫹ cells (⬎96% purity). The Fas-sensitive subclone Ramos.FsA of the Burkitt lymphoma Ramos (26) has been previously described.

Cell culture All culture experiments were performed in IMDM (Life Technologies, Gaithersburg, MD) enriched with 10% heat-inactivated FCS (Euro Biochem, Bierges, Belgium), gentamicin (43 ␮g/ml), and 2-ME (0.0035%) (culture medium).

Confocal laser microscopy Purified CD45RA⫹CD27⫹, CD45RA⫹CD27⫺, and CD3⫺CD16⫹ populations were isolated as described above. After adhesion to poly-L-lysinecoated object glasses (30 min at room temperature (RT), cell populations were fixated in PBS containing 3% paraformaldehyde (60 min at RT). Next, cells were incubated in PBS containing 50 mM NH4Cl (5 min at RT) and subsequently permeabilized in PBS/0.1% saponin/0.5% BSA (30 min at RT). After washing, intracellular staining was performed by incubating

1911 cells with anti-human granzyme A Ab (45 min at RT). Cells were then washed three times and incubated with goat anti-mouse IgG FITC (30 min at RT). Moviol containing 2.5% diazabicyclo octane (Fluka, Buchs, Switzerland) was used to mount on the cells. For immediate analysis, we made use of a Leitz DMIRB fluorescence microscope (Leica, Voorburg, The Netherlands) interfaced with a Leica TCS4D confocal laser microscope (Leica, Heidelberg, Germany). Images were imported in Adobe Photoshop 4.0 (Adobe Systems, San Jose, CA).

Induction and measurement of granzyme release CD8⫹ subpopulations were isolated as described above. Cells were cultured in triplicate in 96-well plates (0.3 ⫻ 106 cells/well) in either the presence or the absence of 1 ng/ml PMA and 1 ␮M ionomycin (both from Sigma, St. Louis, MO). Supernatants were harvested after 1 h culture, and the levels of soluble granzyme A and B were measured by ELISA. Stimulated cells were used for intracellular evaluation of these molecules according to the protocol described below. Granzyme A and B concentrations in culture supernatants were measured by specific solid phase sandwich ELISA as previously described (27) with some modifications. The granzyme A ELISA was performed using the GA29 mAb (coating Ab; CLB) and the biotinylated GA28 mAb (detection Ab; CLB). The limits of detection of the granzyme A and B ELISA systems were 12 and 3 pg/ml, respectively.

Cytotoxicity assay To evaluate the contribution of the Fas/FasL pathway by effector CD8⫹ T cells, a redirected cytotoxic assay was performed as previously described (28) with some modifications. Briefly, to specifically block cytotoxicity mediated by perforin and granzymes, isolated CD8⫹ T cells were pretreated (or not) for 4 h with 400 nM concanamycin A (CMA) (Sigma), washed once, and cultured in the presence of CMA. Next, cells were cocultured in 24-well plates (106 cells/well) with FsA target cells (0.2 ⫻ 106 cells/well) (E:T 5:1). The redirected cytotoxic assay conditions were imitated by incubating the cells for 18 h with anti-CD3 (clone CLB-T3/4.1) to stimulate effector CD8⫹ T cells and bind to the FcR expressed on Ramos.FSA cells. To evaluate the existence of Fas/FasL-dependent cytotoxicity, blocking Abs (anti-FasL (NOK-2) or anti-Fas (Fas2)) were added to the cultures. Apoptosis of Ramos.FSA cells was evaluated using FITClabeled annexin V as described previously (26). Ramos.FSA cells were discriminated from CD8⫹ T cells by gating on forward and side scatter parameters (data not shown).

Flow cytometry Expression of cell surface molecules. Staining for NK cell-inhibitory receptors (NKRs) was performed by incubating freshly isolated PBMC with saturating amounts of directly labeled CD8, CD45RA, CD27, and an antiMHC class I killer-inhibitory receptor mAb in PBS containing 0.5% BSA (Bayer, Kankakee, IL) (30 min at 4°C). Expression of the different markers was measured on a FACScalibur (Becton Dickinson) and analyzed with the Cell Quest program (Becton Dickinson). CD8⫹ T cells were gated, and NKRs were analyzed on the different CD8 subsets. Expression of cytoplasmic molecules/epitopes. Intracellular content of FasL was measured in freshly isolated CD8⫹ T cells before and after in vitro stimulation. FasL on stimulated cells was measured in the presence of an inhibitor of protein secretion, resulting in the cytoplasmic accumulation of the synthesized FasL. After cell fixation and permeabilization, intracellular staining was performed according to a protocol originally described by Jung et al. (29) with some modifications. Briefly, isolated CD8⫹ T cells were stimulated (106 cells/ml) for 4 h with PMA (1 ng/ml) and ionomycin (1 ␮M) in the presence of the protein secretion inhibitor monensin (1 ␮M) (all from Sigma). Next, the cells were washed twice in cold PBS-0.5% BSA and stained with CD45RA and CD27 (30 min at 4°C, washed twice with cold PBS, and fixed with PBS containing 4% paraformaldehyde (5 min at 4°C). Fixation was followed by permeabilization with PBS containing 0.1% saponin (Calbiochem, La Jolla, CA) and 0.5% BSA. Nonspecific binding was blocked by incubating the cells in the same buffer supplemented with 10% human pooled serum (CLB) (20 min at 4°C). For all subsequent incubation and washing steps, PBS, 0.1% saponin, 0.5% BSA was used. Cells were then washed once and stained with 5 ␮g/ml anti-FasL (30 min at 4°C). After another washing step, cells were stained with PE-labeled goat anti-mouse IgG2a isotype-specific mAb (20 min at 4°C). Expression of the different markers was measured on a FACScalibur and analyzed with the Cell Quest program. CD8⫹ T cells were gated, and FasL was analyzed on the different CD8 subsets. Expression of CTLA-4 and TCR␨ was performed using a permeabilization and staining protocol identical with that described above.

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CYTOLYTIC PATHWAYS AND INHIBITORY RECEPTORS OF EFFECTOR CD8⫹ T CELLS

FIGURE 1. Confocal laser microscopy analysis of granzyme A expression in purified CD8⫹ subsets and NK cells. Expression is shown in the total CD8⫹ population (a) and is compared with the sorted naive CD45RA⫹CD27⫹ (b), effector CD45RA ⫹ CD27 ⫺ (c), and NK cell (CD3⫺CD16⫹) population (d).

Results Utilization of cytotoxic pathways by circulating effector cells We previously found by flow cytometry that CD8⫹CD45RA⫹CD27⫺ effector-type T cells abundantly express components of the exocytotic cytolysis pathway, i.e., granzyme A, B and perforin (Ref. 7 and data not shown). To test whether in circulating CTL these cytotoxic enzymes are contained within granules and are thereby ready for receptor-induced exocytosis, the intracellular distribution of granzyme A in CD8⫹ T cells (Fig. 1a) was studied by confocal scanning laser microscopy. In accordance with the cytotoxic ca-

pacities of the different subsets, nearly all effector CD8⫹ T cells contained granzyme A in a granular fashion (Fig. 1c), whereas in contrast, naive (CD45RA⫹CD27⫹) CD8⫹ T cells did not express granzyme A (Fig. 1b). Moreover, the expression of this molecule in CD8⫹CD45RA⫹CD27⫺ T cells parallels in both qualitative and quantitative terms the expression observed in freshly isolated NK cells (Fig. 1d). Next we investigated the capacity of CD8⫹ T cells to release granzyme A and B on stimulation with PMA and ionomycin. Considerable concentrations of both granzyme A and granzyme B could be measured in supernatants from cultures of effector CD8⫹

FIGURE 2. Release of granzyme A and B after 1 h stimulation with PMA (1 ng/ml) and ionomycin (iono) (1 ␮M). CD8⫹ total (䡺), CD45RA⫹CD27⫹ (u), CD45RA⫺CD27⫹ (1 ), CD45RA⫹CD27⫺ (f) were sorted and stimulated for 1 h. Granzymes A and B were measured in the supernatants by ELISA.

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FIGURE 3. Extracellular (top) and intracellular (bottom) staining of FasL on CD8⫹ subsets. Unstimulated cells (left) and 4-h PMA/ionomycin/ monensin-stimulated cells (right) were stained extracellularly for CD45RA and CD27. After fixation and permeabilization, cells were stained intracellularly for FasL. In the histograms, the solid line shows FasL expression, whereas the dotted line shows the isotype-matched control mAb. FasLpositive cells were colored black and plotted back into the CD45RA and CD27 dot plot, whereas FasL-negative cells were colored gray. Numbers in the dot plot give the percentage of FasL-positive within the four main subsets and correspond clockwise with the CD45RA⫹CD27⫹, CD45RA⫺ CD27⫹ , CD45RA⫺ CD27⫺ , and CD45RA⫹ CD27⫺ subset.

T cells (Fig. 2). Moderate levels of granzyme A could also be detected in supernatants of memory (CD45RA⫺CD27⫹) cells, in accordance with low levels of expression of granzyme A in this subset (data not shown). In contrast, no significant release of granzyme B was observed when naive or memory cells were stimulated in our experimental conditions. Furthermore, the presence of granzymes in the

FIGURE 4. FasL-dependent killing of CD8⫹ effector cells. Isolated CD8⫹ cells were not pretreated (A) or pretreated (B) with CMA to abolish perforin-granzyme-containing granules. A redirected killer assay was performed with the Fas-sensitive Ramos clone FsA with addition of anti-CD3. E:T ratio, 5:1. FACS staining with annexin V-FITC in combination with propidium iodide assayed the percentage of apoptotic cells. After 18 h of culture, ⬃60% of the Ramos.FSA cells specifically bound annexin V-FITC. Anti-FasL (Nok-2) and anti-Fas (Fas-2) mAbs were added to block FasL-dependent killing, whereas ␥1 and ␥2a irrelevant mAbs were added as controls. The condition with anti-CD3 alone was set to 100%, and the percentage of apoptotic cells is given as the percentage of this control.

supernatants corresponded with a decrease in the intracellular levels on stimulated cells as evaluated by flow cytometry (data not shown). These results indicate that the stored granzymes (A and B) are rapidly released from effector cells after stimulation.

FIGURE 5. Intracellular staining of TCR␨ in CD8 subsets. Purified CD8⫹ cells were stained extracellularly for CD45RA and CD27. After fixation and permeabilization, cells were stained intracellularly with TCR␨. TCR␨ brightly expressing cells were colored black and plotted back into the CD45RA and CD27 dot plot, whereas TCR␨ dull cells were colored gray. The dotted line shows the negative control. Numbers in the dot plot give the percentage of TCR␨ positive within the four main subsets and correspond clockwise with the CD45RA⫹CD27⫹, CD45RA⫺CD27⫹, CD45RA⫺CD27⫺, and CD45RA⫹CD27⫺ subset.


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Table I. CTLA-4 expression in CD8 T cell subsetsa

Stimulus

CD8 Total

“Naive” CD45RA⫹ CD27⫹

Medium CD2 CD2 ⫹ CD28

4b 24 34

1 19 10

“Memory” CD45RA⫺ CD27⫹

CD45RA⫺ CD27⫺

“Effector” CD45RA⫹ CD27⫺

11 48 50

14 42 53

2 11 32

a Purified CD8 cells were stimulated as indicated for 16 h in the presence of monensin. Extracellular staining was performed for CD45RA and CD27. After fixation with PFA, cells were intracellularly stained for CTLA-4. All isotype-matched controls were ⬍1%. One representative experiment is shown of three. b Percentage of CTLA-4⫹ cells within given subset.

FasL mRNA is readily detectable in CD45RA⫹CD27⫺ cells but hardly in one of the other circulating CD8⫹ T cell subsets (7). We evaluated the extracellular and intracellular expression of FasL protein in either unstimulated or stimulated CD8⫹ T cell subsets. Unstimulated cells did not show extracellular expression of FasL, whereas intracellular expression of FasL was abundantly found in CD45RA⫹CD27⫺ cells (Fig. 3, left). To assess the expression of FasL after stimulation, purified CD8⫹ T cells were cultured for 4 h in the presence or absence of PMA and ionomycin. Even in the presence of the metalloproteinase inhibitor KB8301 only a minimal amount of extracellular FasL could be detected (data not shown). Stimulation did, however, result in an increase in the intracellular expression of FasL in the memory compartment (Fig. 3, right). To investigate whether both granule exocytosis and the Fas/ FasL pathways contribute to the cytotoxic potential of circulating effector cells, a redirected killer assay was performed. For this, effector CD8⫹ cells were tested for their ability to induce apoptosis in a Fas-sensitive subclone of the Burkitt lymphoma cell line Ramos, i.e., Ramos.FSA (26). Cytotoxicity was initiated by adding CD3 mAb to stimulate CD8⫹ cells and bind to the FcR of the Ramos.FSA cells. Evaluation of apoptotic cell death on the target cells was performed by annexin V staining. To estimate the contribution of the exocytosis pathway to the killing by CD8⫹CD45RA⫹CD27⫺ T cells, CMA, a selective inhibitor of perforin-based killing, was used (30). Apoptosis of Ramos.FSA cells by freshly isolated effector CTLs could be blocked by ⬃55% when cells were incubated with CMA showing contribution of the granule exocytosis pathway (data not shown). The remaining cytotoxic response appeared to be largely dependent on the triggering

FIGURE 6. Extracellular staining of CD94 and CD158a in CD8⫹ subsets. Purified CD8⫹ cells were stained extracellularly for CD45RA, CD27, together with CD94 or CD158a. CD94or CD158a-positive cells were colored black and plotted back into the CD45RA and CD27 dot plot whereas negative cells were colored gray. The dotted line shows the negative control. Numbers in the dot plot give the percentage of CD94- or CD158a-positive cells within the four main subsets and correspond clockwise with the CD45RA ⫹ CD27 ⫹ , CD45RA ⫺ CD27⫹, CD45RA⫺CD27⫺, and CD45RA⫹ CD27⫺ subset.

of Fas on the Ramos.FSA cells, because addition of either Fasblocking mAb (Fas2) or FasL mAbs reduced apoptosis by 74 and 46%, respectively (Fig. 4). Because combined usage of CMA and Fas-blocking mAb reduced target cell lysis by ⬃90%, we believe that the granule exocytosis and FasL pathway account for the majority of cytolytic activity of these circulating effectors. Expression of activation-regulating molecules on effector-type T cells The abundance of cytolytic mediators in circulating effector CTLs suggests that the activity of these cells must be carefully controlled to prevent unwanted tissue damage. In a number of in vivo conditions, such as autoimmunity, tumor growth, and HIV infection (31–34), low responsiveness of T cells may be induced by downregulation of the expression of TCR␨, which is an essential signaling component in the CD3/TCR complex. To investigate whether low expression of TCR␨ would be involved in the control of the activation of cytotoxic effector cells, we analyzed the expression of TCR␨ on the different CD8⫹ T cell subsets. As shown in Fig. 5, memory and effector CTLs expressed comparable high levels of TCR␨ whereas the naive subset expressed TCR␨ less intensively. Moreover, because effector CTLs also express relatively high levels of CD3⑀ and the coreceptor CD8 (data not shown), we conclude that it is unlikely that circulating effectors are kept at a low level of activation by suboptimal signaling via TCR/CD3. CTLA-4 acts as a negative regulator of T cell activation and is vital for the control of lymphocyte homeostasis (35). To evaluate whether this molecule could be involved in the regulation of activation of circulating effector CTLs, CTLA-4 expression was

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Table II. Expression of NKRs on CD8 T cell subsets in comparison with NK cells

CD94 NKG2a CD158a CD158b NKB-1 a b

CD8 Total

CD45RA⫹ CD27⫹

CD45RA⫺ CD27⫹

CD45RA⫺ CD27⫺

CD45RA⫹ CD27⫺

NK Cell CD3⫺ CD56⫹

43 ⫾ 11a (31–59)b 6⫾3 (4–12) 11 ⫾ 14 (0.6–39) 9⫾6 (3–20) 1.9 ⫾ 0.8 (1.1–3.2)

3⫾1 (2–5) 0.6 ⫾ 0.3 (0–0.9) 0.4 ⫾ 0.4 (0.1–1.2) 1.4 ⫾ 0.8 (0.4–3) 0.3 ⫾ 0.3 (0–0.7)

26 ⫾ 14 (9–44) 7⫾2 (4–9) 0.3 ⫾ 0.2 (0.1–0.5) 2.0 ⫾ 0.8 (0.7–3) 0.1 ⫾ 0.1 (0–0.4)

64 ⫾ 24 (34–93) 12 ⫾ 11 (4–36) 3⫾6 (0.2–15) 12 ⫾ 10 (2–30) 3⫾5 (0–11)

90 ⫾ 4 (86–98) 10 ⫾ 7 (2–20) 32 ⫾ 37 (6–92) 22 ⫾ 11 (9–37) 4⫾2 (2–7)

93 ⫾ 4 (90–98) 47 ⫾ 7 (42–55) 32 ⫾ 21 (10–51) 45 ⫾ 12 (33–56) 14 ⫾ 1 (13–15)

Percentage of positive cells within given subset with SEM (n ⫽ 6). Numbers in parentheses, range.

analyzed. CD8⫹ T cells were stained for CD45RA, CD27, and CTLA-4 before and after stimulation with CD2 and CD28 mAbs. On unstimulated cells, this T cell regulator was almost absent in effector CTLs whereas ⬃10% of memory CTLs expressed this molecule (Table I). On activation with anti-CD2 mAbs, both effector CTLs and memory-type cells had comparable but moderate expression, i.e., 32 and 50% positive cells, respectively (Table I). The absence on freshly isolated cells and the low expression on in vitro activated T cells render it unlikely that CTLA-4 acts as a major negative regulator of effector CTL activation. MHC class I inhibitory receptors have been found on CD28⫺ T cells, and studies in which T cell clones were activated by (super) Ags have shown that inhibitory receptors can modulate effector CTL function (36). We analyzed the expression of NKRs during proposed stages of CD8⫹ T cell differentiation (25). Naive (CD45RA⫹CD27⫹) CD8⫹ T cells virtually lacked any of the NKRs analyzed. In contrast, circulating effector (CD45RA⫹CD27⫺) CD8⫹ T cells showed a strong expression of all NKRs analyzed which was with respect to CD94 and CD158a,b comparable with the expression levels on NK cells (Fig. 6, Table II). Interestingly, within the CD45RA⫺ subset, CD27⫺ cells had a relatively high NKR expression. The present data corroborate that the differentiation of human CD8⫹ T cells toward effector CTL is accompanied by an increase in the expression of NKRs.

Discussion Class I-restricted effector CTL specifically recognize cells that express peptides derived from intracellular replicating pathogens and may subsequently induce death of these cells. The data presented here show that CTL that have developed in vivo and that can be identified by the CD8⫹CD45RA⫹CD27⫺ surface phenotype have two separate death-inducing mechanisms at their disposal, the granule exocytosis and the FasL pathway. The coexpression of these systems suggests that, dependent on properties of the infected cell, distinct elimination strategies can be used by these effectors in vivo. In addition, IFN-␥ and TNF-␣ that are highly produced by these cells (7) likely aid in coping adequately with infections, e.g., by limiting virus replication (37). Although circulating effector T cells are well equipped with these cell death-inducing pathways, they do not, without stimulation, release granzymes or perforin, nor do they express FasL on the plasma membrane. Short term stimulation in vitro, however, induces a very rapid release of the constituents of the exocytosis pathway in culture supernatants. On the other hand, although CD8⫹CD45RA⫹CD27⫺ T cells contain FasL mRNA (7) and express limited but discernible amounts of FasL intracellularly, we were unable, even in the presence of specific proteinase inhibitors,

to reliably demonstrate FasL on the surface of effector cells. However, the contribution of FasL in the induction of apoptosis in a Fas-sensitive Burkitt lymphoma line demonstrates that, in accordance with previous findings in mice (38, 39), in vivo matured human CTLs can make use of the FasL effector pathway The strong expression of these cytolytic molecules and the functional availability of these compounds upon activation suggest that the activity of these circulating effectors must be carefully controlled to prevent unwanted damage to healthy cells, e.g., after the recognition of cross-reactive peptides. NKRs, which have been orginally described on NK cells, can after binding to MHC class I molecules transduce inhibitory signals for cellular cytotoxicity (40). Concordant with previous findings on CD28⫺ T cells, we found that CD8⫹CD45RA⫹CD27⫺ T cells express a variety of NKR, both of the C-type lectin and Ig superfamily classes (23, 24). In healthy individuals the majority of CD8⫹CD27⫺ T cells express CD45RA but lack CD28 showing that under physiological conditions CD27⫺ and CD28⫺ subsets represent largely overlapping populations of CD8⫹ T cells (7). However, when patients with acute viral infections are being analyzed a very considerable portion of CD45RA⫺CD28⫺ T cells does express CD27 (25, 47). We have postulated that by combined analysis of CD27, CD28, and CD45RA expression, distinct stages of differentiated CD8 T cells can be identified (25). We here show that the differentiation toward CD8⫹CD45RA⫹CD27⫺ “effector-type” T cell is not only accompanied by an enhancement of intracellular cytolytic mediators but also coincides with an strong increase in the expression of various NKRs. A complicating finding in appreciating the functional consequences of the expression of these molecules is that the mAb that are being used to detect NKR of the Ig family will also bind to splice variants of these molecules that can act as activating receptors, i.e., killer cell activating receptor (41). Differing with Mingari et al. (23), we find highest expression of these regulating receptors on CD45RA⫹CD27⫺ and therefore CD28⫺ T cells (7) and not on CD45RA⫺CD27⫺ T cells. Although it has recently been reported that IL15 can induce CD94/NKG-2A expression on mitogenically activated T cells (42), it is less clear which specific signals do induce the expression of Ig superfamily type NKRs on T cells. T cell cloning experiments have indicated that the expression of NKRs is a stable phenotypic trait. In some donors, very high percentages of both C-type lectin and Ig superfamily NKRs were found specifically in the CD27⫺ T cell fraction. This finding could suggest that concomitant with their differentiation toward differentiated effector cells, CD8⫹CD45RA⫹CD27 receive specific signals that induce up-regulation of these cell activation-regulating receptors.

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Multiple studies have documented that CD8⫹CD28⫺, CD8⫹CD45RA⫹CD27⫺, or NKR⫹ subsets have a restricted and selected usage of the TCR repertoire suggestive of selection by Ag (10, 16). Indeed, more recently, it was shown that virus-specific and tumor Ag-specific CD8⫹CD45RA⫹CD27⫺ T cells can be found in hepatitis C virus- and CMV-infected persons and melanoma patients, respectively (18, 43). Several reports have documented that NKRs can inhibit both superantigen- and Ag-induced activation of T cells (36, 44, 45). Interestingly, Lee et al. (43) reported that despite the presence of Ag-specific CD8⫹CD45RA⫹CD27⫺ in the circulation of a melanoma patient, no lysis of melanoma cell line in vitro was observed. Because our data suggest that these cells will also express high levels of NKRs, inhibition of TCR-induced killing can be envisaged. For this reason, it is important to more precisely define the signals that induce expression of NKRs on effector T cells, because these receptors will seriously interfere with vaccination strategies for, e.g., cancer. The question remains on the physiological function of circulating effector-type cells. The functional relevance of these cells appears to be supported by the fact that circulating effector-type cells increase with age (46). Although the presence of differentiated and functionally competent CTL could provide a first line of defense against invading pathogens to which a protective response has already been mounted once, the expression of NKR will preclude rapid activation by these cells by any dose of peptide. Yet, either with decreasing amounts of MHC molecules (which may limit the inhibitory signal) or with increasing specific peptide concentration (which will augment the stimulatory signal), these cells may become activated. In fact, the integration between these positive and negative signals allows the effector CTL to detect the specific loading of MHC molecules with foreign peptides irrespective of the number of MHC molecules expressed on a target cell.

Acknowledgments We thank Jo¨rg Hamann, Do¨rte Hamann, and Claire Boog for critical reading of and comments on the manuscript. We also thank Angela Wolbink and Erik Hack for performing granzyme A and B ELISAs.

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