The TCR-Binding Region of the HLA Class I α2 Domain Signals Rapid Fas-Independent Cell Death: A Direct Pathway for T Cell-Mediated Killing of Target Cells?

Rolf D. Pettersen, Gustav Gaudernack, Mette Kløvstad Olafsen, Sverre O. Lie and Kjetil Hestdal
J Immunol May 1, 1998, 160 (9) 4343-4352;

Abstract

TCR binding to an MHC class I/peptide complex is a central event in CTL-mediated elimination of target cells. In this study, we demonstrate that specific activation of the TCR-binding region of the HLA-A2 class I α2 domain induces apoptotic cell death. mAbs to this region rapidly induced apoptosis of HLA-A2-expressing Jurkat E11 cells, as determined by morphologic changes, phosphatidylserine exposure on the cell surface, and propidium iodide uptake. In contrast, apoptosis was not induced following culture with mAbs directed to other regions of the class I molecule. Death signaling by class I molecules is apparently dependent on coreceptor activation, as apoptosis is also signaled by HLA-A2 molecules, where the intracytoplasmic residues were deleted. HLA class I α2-mediated cell death appeared to proceed independent of the Fas pathway. Compared with apoptotic signaling by Fas ligation, HLA class I α2-mediated responses displayed a faster time course and could be observed within 30 min. Furthermore, class I α2-induced cell death did not involve observable DNA fragmentation. The apoptotic response was not affected significantly by peptide inhibitors of IL-1β converting enzyme (ICE)-like proteases and CPP32. Taken together, activation of the TCR-binding domain of the class I α2 helix may result in apoptotic signaling apparently dependent on a novel death pathway. Thus, target HLA class I molecules may directly signal apoptotic cell death following proper ligation by the TCR.

Two major functions have been assigned to MHC class I molecules in directing the host immunity to transformed and virally infected cells. Expressed on the cell surface, class I molecules present a wide variety of peptide Ags for CTL surveillance and activation (1, 2, 3). MHC class I molecules also control NK cell activity, as these cells express specialized inhibitory receptors that recognize different class I structures. Thus, abnormal cells deficient in class I expression are lysed by NK cells (4, 5).

Recognition of a target cell induces a complex array of responses in the CTL (6, 7). TCR binding to peptide ligand and allele-specific α1 and α2 residues of the class I molecule initiates signaling through the TCR/CD3 complex. These signals partly activate CD8 coreceptor binding to the α3 domain of the same MHC class I molecule, thereby increasing TCR-ligand interactions (8, 9, 10, 11, 12). Furthermore, other auxiliary molecules also seem to coregulate CTL adhesion to target cells (13, 14).

Following proper activation of the CTL, the effector or cytolytic phase is activated to eliminate target cells (15). In this context, perforin- and Fas-mediated pathways have been recognized as the two major complementary cytotoxic pathways used by CTLs (16, 17). The Ca2+-dependent perforin pathway has been implicated in virtually all forms of cell-mediated cytotoxicity (15, 16, 17, 18, 19). Most effector CTL have perforin and granzymes stored in cytoplasmic granules that are released upon TCR signaling (6, 20). Perforin forms channels in the target cell, and it is believed that transmitted perforin and granzymes (mainly granzyme B) in concert induce apoptotic cell death (21, 22). NK cells seem to mainly, but not entirely, rely on the perforin-mediated pathway to eliminate class I-deficient target cells (18, 19, 23). In addition, highly potent peritoneal exudate lymphocytes seem to exert their cytotoxic function independent of the perforin pathway, as they contain no lytic granules (24).

Transmembrane signaling by Fas molecules is the other recognized major pathway in CTL-mediated induction of apoptotic target cell death (16, 17). Activation of CTL rapidly induces CD95 ligand expression, and thus enables these cells to induce cell death in Fas-expressing and sensitive target cells (25). In contrast to the perforin pathway, Fas-mediated apoptosis is Ca2+ independent (26). Furthermore, in some situations CTL may also use TNF-α to kill target cells (27). However, this pathway is slower than Fas- and perforin-dependent pathways (15).

Recent studies demonstrate that HLA class I molecules may have other biologic functions in addition to Ag presentation, as class I molecules may transduce regulatory signals. Thus, activation of class I molecules may apparently result in cell activation (28, 29, 30), growth inhibition (31, 32, 33, 34, 35, 36, 37), and cell aggregation (37, 38, 39). Furthermore, engagement of class I α3 residues may cosignal or directly induce apoptosis (40, 41). We have reported recently that the TCR-accessible region of the HLA class I α2 domain may have a unique function in class I-mediated signaling (37). It is believed that the perforin/granzyme and Fas pathways account for all acute cytolytic activity of CTL, and that there is no distinction between autologous and allogeneic CD8+CTL-mediated cytotoxicity (16, 17). There is presently no direct evidence for alternative pathways of CTL-mediated target cell elimination. However, studies with perforin-less mice show that these are still able to reject allogeneic CD95− tumor cells injected i.p. as efficiently as perforin normal mice (15). Furthermore, murine CD8+ T cells apparently clear rotavirus infection independent of perforin and Fas (42). Thus, these and other observations may suggest additional cytotoxic pathway(s).

We asked whether direct activation of epitopes encompassing TCR contact residues on HLA molecules could represent an alternative death pathway. In this study, we demonstrate that HLA class I molecules can signal Fas-independent cell death following specific engagement of the TCR-binding region of the class I α2 domain.

Materials and Methods

mAbs and chemicals

Hybridomas MA2.1 (anti-A2, B17, IgG1) (43), PA2.1 (anti-A2, w69, IgG1) (44), BB7.2 (anti-A2, w69, IgG2b) (45), W6/32 (anti-monomorphic HLA, IgG2a) (46) and OKT3(anti-CD3, IgG2a) were obtained from American Type Culture Collection (Rockville, MD). The CR11-351 hybridoma (anti-A2, 28, IgG1) (47) was generously provided by Dr. Carlo Russo (Cornell University, New York, NY). The supertypic anti-HLA-A mAb RG1 (IgG1) has been described previously (48). Anti-CD95 (phycoerythrin-conjugated DX2, IgG1) was obtained from PharMingen (San Diego, CA). Apoptosis-inducing anti-CD95 (CH11, IgM) and apoptosis-inhibitory anti-CD95 (ZB4, IgG1) were from Immunotech (Marseseille Cedex, France). Murine IgG1 (MOPC-21) control was purchased from Sigma Chemical (St. Louis, MO), and murine IgM control (TEPC 183) was obtained from BiosPacific (Emeryville, CA). IgG1-phycoerythrin control was purchased from Becton Dickinson (San Jose, CA). Human rFas-Fc chimera was obtained from R&D Systems (Abingdon, U.K.). IL-1β-converting enzyme (ICE)3 inhibitor II Ac-Tyr-Val-Ala-Asp-chloromethyl ketone (Ac-YVAD-CMK) was from Calbiochem (La Jolla, CA). ICE inhibitor Z-Val-Ala-DL-Asp-fluoromethylketone (Z-VAD-FMK) and CPP32 inhibitor Ac-Asp-Glu-Val-Asp-aldehyde (Ac-DEVD-CHO) were purchased from BACHEM Feinchemikalien (Bubendorf, Switzerland). Streptavidin-FITC was from Dako (Glostrup, Denmark).

Cell culture

Hybridomas were maintained in DMEM supplemented with 17% FCS (Life Technologies, Gaithersburg, MD), 100 U/ml penicillin, and 100 μg/ml streptomycin. Human cells were cultured in RPMI 1640 supplemented with 10% FCS, 1.5 mM glutamine, 100 U/ml penicillin, and 100 μg/ml streptomycin. Jurkat cells transfected with either native (Jurkat E11) or truncated HLA-A2 (Jurkat E13) genes or native (Jurkat E10) or truncated (Jurkat E12) HLA-B27 genes (49) were kindly provided by Dr. Peter E. Lipsky (The University of Texas Southwestern Medical Center at Dallas, TX). Jurkat cells were subcultured routinely every 2 to 3 days to maintain cell densities between 1 × 105 and 1 × 106 cells/ml. Experiments were performed on 5 × 105 cells/ml, unless otherwise indicated.

Human PBMC were isolated by Lymphoprep (Nycomed Pharma, Oslo, Norway) centrifugation. All cultures were grown at 37°C in a humidified 5% CO2 atmosphere.

Flow cytometry

Ag expression was determined with flow cytometry, as described (37). Cells were finally washed twice in PBS and fixed with 1% paraformaldehyde in PBS. Samples were analyzed using a FACScan (Becton Dickinson), and data were collected on 10,000 cells.

Flow-cytometric determination of apoptosis and cell death

Apoptosis was determined by monitoring changes in cell size and granularity by flow cytometry and assessment of phosphatidylserine exposure by Annexin V-FITC binding using the ApoAlert Annexin V apoptosis kit (Clontech Laboratories, Palo Alto, CA). DNA fragmentation was determined with the TUNEL assay kit from Boehringer Mannheim (Mannheim, Germany), according to the manufacturer’s instructions. Cell membrane permeability was assessed by determining uptake of the DNA-binding fluorescent dye propidium iodide (PI; 2.5 μg/ml) after incubation for 10 min.

Results

Activation of TCR-binding HLA class I α2 epitopes signals apoptosis

CTL recognition of target cells involves TCR interactions with peptides and specific HLA class I residues (2). X-ray crystallographic data have recently identified amino acid residues within the class I α2 domain that make contact with the TCR (50). These residues may be responsible for TCR/MHC class I cross-talk and conveying signals to the target cell. Activation of class I α2 residues that interact with the TCR may initiate target cell signaling, as evidenced by cell aggregation of most normal and leukemic cells and growth inhibition of mitogen-stimulated T cells following mAb stimulation of this class I region (37). In this study, we addressed the involvement of the TCR-binding region of the HLA class I α2 domain in signaling cell death. HLA-A2-expressing Jurkat E11 cells (49) were used as a model system, as Jurkat cells can be induced to undergo apoptosis by different pathways (51, 52, 53, 54) and rapidly respond with cell signaling (i.e., cell aggregation) following engagement of residues within the TCR contact region of HLA-A2 molecules (data not shown).

CTL targets are believed to die from programmed cell death or apoptosis (20). This is an active process with characteristic morphologic changes, including cell shrinkage and nuclear condensation (55). Thus, to examine whether apoptosis could be signaled by activation of the class I α2 domain, we first examined the impact of anti-HLA class I on cell size and granularity using flow cytometry. Jurkat E11 cells were cultured with mAbs to positions within (RG1 and CR11-351) (37, 48, 56, 57) the TCR-accessible α2 domain of HLA-A2 molecules (2, 50) or other distant class I epitopes (MA2.1, PA2.1, BB7.2, and W6/32) (57). These studies clearly demonstrated that both RG1 (epitope involves residues 144 and 151) (37, 48) and CR11-351 (epitope involves residues 142, 145, and 149) (56, 57) induced a major impact on cell morphology, as evidenced by a shift in cell size and granularity (Fig. 1⇓). Within 3 h, up to 50% of the cells could be recognized as a second population of smaller cells compared with control cultures (Fig. 1⇓A). In contrast, no significant influence on cell size or granularity was observed with the anti-HLA class I mAbs MA2.1 (involving position 62) (57, 58), PA2.1, BB7.2 (both specific for positions 107 and 161) (57, 59, 60, 61), W6/32 (recognizing a conformational determinant involving conserved residues on the α1 and α2 domains) (62), or isotype control mAb MOPC-21 after 2, 3, or 5 h of cell culture (Fig. 1⇓ and data not shown). The effect of CR11-351 was readily observable within 60 min (Fig. 1⇓B). For both RG1 and CR11-351, the effect was stronger after prolonged culture, reaching a maximum after 5 to 6 h (Fig. 1⇓B and data not shown). Thus, engagement of TCR-accessible epitopes on the class I α2 domain rapidly leads to cell shrinkage and changes in granularity, as observed with cells undergoing apoptosis.

FIGURE 1.
FIGURE 1.

Stimulation of the TCR-binding region of the HLA class I α2 domain induces morphologic cell changes. Jurkat E11 cells were cultured with the indicated anti-HLA class I mAbs or isotype control mAb MOPC-21 (1 μg/ml) for different time periods and examined for changes in cell size and granularity on the basis of changes in the light-scattering properties of the cells. A, Light-scattering properties of the cells after 3 h. The numbers indicate the percentage of cells in the region R1 defining viable cells. Ten thousand cells were analyzed under each condition. B, Kinetics of morphologic cell changes expressed by the number of cells in R1. Each data point was derived from an analysis of 10,000 cells. Data in A and B are from two different experiments.

During the early stages of apoptosis, phosphatidylserine translocates from the interior to the exterior part of the plasma membrane and becomes exposed at the cell surface to enable recognition by macrophages (51, 63). Annexin V binds with high affinity to phosphatidylserine and can thus be used to identify cells in all stages of programmed cell death (51). PI only stains cells with a disrupted cell membrane and can be used to identify late apoptotic and dead cells. To assess the impact of HLA class I signaling on phosphatidylserine exposure and cell viability, Jurkat E11 cells were cultured with anti-HLA class I mAbs. Annexin V-FITC and PI staining were performed to distinguish between early/intermediate apoptotic and late apoptotic/dead cells. This allowed us to follow the dynamics of the process. Assessment of Annexin V-FITC binding and PI uptake was performed by two-color flow cytometry. These experiments demonstrated that both RG1 and CR11–351 rapidly induced phosphatidylserine exposure on Jurkat E11 cells, as shown in Figure 2⇓. Furthermore, determination of PI uptake disclosed an increase in both the number of dead (Annexin V-positive/PI-positive) and apoptotic (Annexin V-positive/PI-negative) cells, as compared with control cultures (Fig. 2⇓), indicating a rapid progress through the apoptotic stage. In contrast, no phosphatidylserine exposure, nor changes in cell distribution or PI uptake were observed with the anti-HLA class I mAbs PA2.1, MA2.1, BB7.2 or W6/32 after 2, 3, of 5 h of cell culture (Fig. 2⇓ and data not shown). In addition, in experiments using the HLA-A2-negative Jurkat E10 cell line as control, no RG1 or CR11-351 effects were observed (Fig. 2⇓). Thus, class I molecules may apparently transduce apoptotic signals following specific engagement of the TCR-accessible α2 region of HLA class I molecules.

FIGURE 2.
FIGURE 2.

Phosphatidylserine externalization and PI uptake are induced by mAbs RG1 and CR11-351 on HLA-A2-expressing Jurkat cells. Jurkat cells expressing HLA-A2 or HLA-B27 were cultured with anti-HLA class I mAbs or control mAb MOPC-21 (1 μg/ml), as indicated, and examined for Annexin V-FITC binding and PI uptake by flow cytometry. Ten thousand cells were analyzed in each case. The numbers at the top indicate the percentage of cells in the lower right (LR) and upper right (UR) regions, respectively.

HLA-A2 molecules signal apoptosis independent of the cytoplasmic domain

The role of the cytoplasmic domain of class I molecules in signaling apoptotic responses has not been determined. To examine a possible involvement of this domain in signaling programmed cell death, RG1 and CR11-351 responses were assessed using Jurkat E13 cells expressing truncated HLA-A2 (A21→312) (49) molecules. These experiments showed that Jurkat E13 cells rapidly responded to RG1 and CR11-351 incubation with both phosphatidylserine exposure and a shift in the cell population with reduction of cell size (Fig. 3⇓ and data not shown). However, no responses were observed in control cultures with MOPC-21 or PA2.1, nor with the HLA-A2-negative Jurkat cell line E12 (Fig. 3⇓). Thus, mAbs RG1 and CR11-351 induce apoptosis by activating an HLA class I signaling pathway that seems to be independent of the cytoplasmic region of class I molecules.

FIGURE 3.
FIGURE 3.

Truncated HLA-A2 molecules signal apoptosis. Jurkat transfectants expressing HLA-A2 or HLA-B27 molecules where the intracytoplasmic residues were deleted, were cultured with mAb RG1, CR11-351, PA2.1 or the control mAb MOPC-21 (1 μg/ml) for 3 h, as indicated. Annexin V-FITC binding and PI uptake were examined by flow cytometry. Ten thousand cells were analyzed in each condition. The numbers at the top indicate the percentage of cells in the lower right (LR) and upper right (UR) regions, respectively.

Class I α2-induced apoptosis is independent of CD95 signaling

CD95 and perforin are the two recognized major pathways for CTL-induced target cell destruction (16, 17). To assess whether CD95 was involved in HLA class I-mediated apoptosis, we first examined the expression of CD95 on Jurkat E11 cells using flow cytometry. These studies revealed that Jurkat E11 constitutively expressed CD95 (data not shown). Furthermore, to address a possible apoptotic response due to CD95-CD95L interactions, the impact of RG1 and CR11-351 was examined on Jurkat E11 cells preincubated with control mAb MOPC-21, neutralizing anti-Fas mAb ZB4, or human rFas-Fc chimera. Control cultures with CH11 (anti-Fas mAb-inducing apoptosis) were included. These experiments clearly demonstrated that whereas ZB4 and human rFas-Fc chimera blocked CH11 responses, they had no effect on the ability of RG1 or CR11-351 to induce apoptosis (Fig. 4⇓). As target cells are believed to die rapidly following recognition by CTLs, we also compared the kinetics of CR11-351- and CH11-induced cell death to evaluate the relative importance of the two pathways. These assessments demonstrated that CR11-351 induced the most rapid and pronounced induction of phosphatidylserine exposure and cell death (Fig. 5⇓). Thus, after 1 h, the levels of Annexin V binding and cell death were similar to control cultures with CH11 compared with strong induction of Annexin V binding and cell death with CR11-351. Interestingly, a relative constant proportion of cells (23–27%) was observed in the early/intermediate (transit) apoptotic stage during the 2- to 4-h period. In contrast, a steady accumulation of dead cells (27–41%) was observed in the same period. During the same time interval, an apoptotic picture emerged with Fas activation. In this study, cells accumulated in the early/intermediate (transit) apoptotic stage with no significant increase in the number of dead cells. The data in Figure 5⇓ indicate that the class I pathway may be a major pathway in CTL-mediated target cell killing.

FIGURE 4.
FIGURE 4.

The HLA class I α2 domain signals apoptosis independent of CD95-CD95L interactions. Jurkat E11 cells were preincubated for 30 min with 10 μg/ml control mAb MOPC-21, neutralizing anti-Fas mAb ZB4, or human rFas-Fc chimera, and further cultured for 4 h with 1 μg/ml RG1, CR11-351 or CH11, as indicated. Apoptosis was assessed by examining Annexin V-FITC binding with flow cytometry. Ten thousand cells were analyzed in each case. Percentage of Annexin V-positive cells is indicated in parentheses. Results are representative of two separate experiments.

FIGURE 5.
FIGURE 5.

Kinetics of phosphatidylserine exposure and cell death after HLA class I α2 and CD95 activation. Jurkat E11 cells were cultured with CR11-351, anti-Fas mAb CH11, or isotype control mAbs MOPC-21 or TEPC-183 (1 μg/ml), as indicated, and examined for Annexin V-FITC binding and PI uptake by flow cytometry. Ten thousand cells were analyzed at each time point and under each condition. A, Assessments of Annexin V-FITC binding vs PI uptake from one representative experiment. The numbers at the top indicate the percentage of cells in the lower right (LR) and upper right (UR) regions, respectively. B, Assessments of percentage of Annexin V binding and PI-positive cells at each time point. Results represent the mean of three different experiments. SD ranges for assessments of PI uptake were 1.9 to 5.7 with CR11-351, and 1.6 to 3.0 with CH11. SD ranges for determinations of Annexin V-FITC binding were 1.8 to 7.7 with CR11-351, and 2.4 to 10.1 with CH11.

Programmed cell death may proceed with or without DNA cleavage into oligonucleosomes. While Fas-induced apoptosis of Jurkat cells typically results in DNA fragmentation (51), CD45 and CD99 ligation results in a different kind of programmed cell death that does not involve DNA fragmentation into oligonucleosomes (52, 53). Compared with observations with CH11, the effects of RG1 and CR11-351 on induction of phosphatidylserine exposure and changes in cell size and granularity were very rapid and evident already within the first hour (Figs. 1⇑, 2⇑, and 5⇑, and data not shown). To further compare Fas and HLA class I-induced apoptosis of Jurkat E11 cells, the cells were stimulated with CH11, RG1, CR11-351, or isotype control mAbs and analyzed for DNA nicks with the TUNEL assay. Due to a more rapid effect of RG1 and CR11-351 compared with CH11, cultures with anti-class I mAbs were analyzed after 1, 2, 3, and 4 h, while the effect of CH11 was determined after 3 and 4 h. These studies showed that whereas CH11 clearly induced DNA fragmentation, only a minor increase in fluorescence staining was observed with the anti-class I mAbs RG1 and CR11-351 (Fig. 6⇓ and data not shown). Taken together, these data suggest that activation of the TCR-binding α2 region of HLA class I molecules rapidly signals apoptosis independent of the CD95 pathway. This class I pathway may contribute significantly to target cell killing by CTL.

FIGURE 6.
FIGURE 6.

mAb CR11-351 induced apoptosis proceeds without observable DNA fragmentation. Jurkat E11 cells were cultured with CR11-351, anti-Fas mAb CH11, or isotype control mAbs MOPC-21 or TEPC-183 (1 μg/ml), as indicated, for 4 h. DNA fragmentation was examined with the TUNEL assay. Incorporated biotin-16-dUTP was determined using streptavidin-FITC and flow cytometry.

Role of ICE and CPP32

ICE and CPP32 are proteases that are rapidly activated in cells induced by different stimuli to undergo apoptosis. Following Fas ligation, ICE-like and CPP32-like proteases are sequentially activated (64). Fas-mediated signaling in Jurkat cells and cytoplasts, as measured by phosphatidylserine exposure, is inhibited by peptide inhibitors of ICE activity (65). To determine the role of ICE and CPP32 activity in signaling by the class I α2 domain, we assessed the impact of specific ICE and CPP32 peptide inhibitors on RG1- and CR11-351-induced responses. Control cultures with CH11 were also included. Jurkat E11 cells were preincubated for 2 h with the ICE inhibitors Ac-Tyr-Val-Ala-Asp-chloromethyl ketone or Z-Val-Ala-DL-Asp-fluoromethylketone or the CPP32 inhibitor Ac-Asp-Glu-Val-Asp-aldehyde before addition of RG1, CR11-351, CH11, or isotype control mAb, and further cultured for 4 h. Apoptosis was determined by assessment of Annexin V-FITC binding with flow cytometry. These experiments showed that whereas peptide inhibitors of ICE and CPP32 profoundly inhibited Fas-mediated apoptosis, the apoptotic responses induced by RG1 and CR11-351 were not influenced significantly by blocking ICE or CPP32 activity (Table I⇓). Thus, HLA class α2 mediated death signaling proceeds apparently independent of ICE and CPP32 involvement.

View this table:
Table I.

Effects of ICE and CPP32 peptide inhibitors on RG1 and CR11-351 induced Annexin V bindinga

RG1 and CR11-351 induce cell death in mitogen-stimulated, but not resting normal T cells

The experiments reported above have all been performed on transformed cells. To establish a model system for normal cells, we investigated the impact of HLA signaling on resting and activated normal T cells. We have demonstrated previously that mAbs RG1 and CR11-351 inhibit the proliferation of mitogen-stimulated lymphocytes, indicating some form of negative signaling (37). To determine whether negative signaling by the TCR-binding region of the class I α2 domain required cell proliferation, the impact of RG1 and CR11-351 on unstimulated and mAb OKT3-stimulated PBMC was examined. Cell size and granularity (forward scatter vs side scatter) were examined with flow cytometry, and regions representing living (R1) and dead or dying lymphocytes (R2) were defined based on Annexin V-FITC binding and PI uptake (data not shown). The effects of RG1 and CR11-351 were assessed based on the increase in the number of lymphocytes in R2 compared with cultures with control mAb MOPC-21.

In three different experiments, no significant increase in cell numbers was observed in region R2 when resting PBMC were incubated with RG1 or CR11-351 for 24 h (Fig. 7⇓ and data not shown). In contrast, with cells prestimulated for 18 h with OKT3 and then further activated for 6 h with RG1 or CR11-351, we observed an increasing number of dead and dying T lymphocytes compared with similar cultures with isotype control mAb (Fig. 7⇓ and data not shown). On the average, we found an increase in R2 of 43.5% (range, 37.3–51.2%) with RG1, 46.5% (range, 44.8–49.2%) with CR11-351, and 10.7% (range, 7.1–15.4%) with PA2.1 relative to MOPC-21. Taken together, the cell’s state of activation appears to determine sensitivity or resistance to death signaling by the TCR-binding region of the class I α2 domain.

FIGURE 7.
FIGURE 7.

mAb CR11-351 induces cell death of mitogen-stimulated, but not resting T cells. PBMC (1 × 106 cells/ml) were incubated with CR11-351 or isotype control mAb MOPC-21 (2 μg/ml), as indicated, and examined for the impact on cell viability. Regions R1 and R2 represent living and apoptotic/dead cells, respectively. Under each condition, 10,000 cells were collected, and the number of cells in region R2 was determined. The numbers above each plot represent the number of cells in region R2 and the percentage of cells in this region compared with control culture with MOPC-21. A, Resting PBMC were incubated with mAbs for 24 h and examined for the number of cells in R2. B, PBMC stimulated with the mAb OKT3 (1 μg/ml) for 18 h were finally cultured for an additional 6 h with CR11-351 or MOPC-21 and then analyzed.

Discussion

We have found that activation of specific TCR-binding residues on the HLA class I α2 domain can rapidly induce morphologic changes, cell surface phosphatidylserine exposure, and cell death independent of the Fas signaling pathway. These data support a unique functional role of this class I region, and furthermore, suggest that TCRs interacting with MHC class I/peptide ligands may directly impose negative signaling in the target cell. This may represent a novel pathway to target cell killing by CTL.

CTL activation and effector functions are subjected to a complex array of control mechanisms and signaling events that only partly have been determined. In the initial phase, CTL contact with the APC is determined by both TCR affinity for peptide and the level of presented foreign peptide/MHC complexes (6, 7, 66). Polymorphic residues on the α1 and α2 domains in adjacent positions to the peptide-binding groove will also contribute to determine the overall binding capacity of the TCR (2).

Sufficient TCR-MHC/peptide interaction signals activation and adhesion of CD8 to a conserved region on the α3 domain of class I molecules. Thus, CD8 contributes to increase the overall TCR avidity to MHC/peptide and also acts as a cosignaling receptor (6, 7, 8, 9, 10, 11, 12). Other costimulatory and adhesion molecules are also required for CTL activation and cytolytic activity (6, 7, 13, 14). In this context, we have shown recently that engagement of TCR-accessible epitopes on the class I α2 domain induces profound cell aggregation responses independent of LFA-1/ICAM interactions (37). Unidentified adhesion molecules may therefore further participate in coregulation of CTL responses.

We have focused on the role of HLA class I molecules in delivering death signals to a potential target cell. In this context, the signaling capacity of the TCR-binding region of the class I α2 domain has been further assessed. MHC class I molecules have been implicated both in positive and negative signaling (28, 29, 30, 31, 32, 33, 34, 35, 36, 37, 40, 41). Sambahra and Miller (40) showed that the α3 domain cosignals induction of apoptosis upon TCR stimulation of immature T cells. Moreover, class I α3 domain activation alone may also be sufficient to induce apoptotic cell death (41).

The mAbs RG1 and CR11-351 bind epitopes in the class I α2 region encompassing amino acids 149–151 (37, 48, 56, 57). These residues have recently been demonstrated to be directly involved in TCR contact with HLA-A2 using x-ray crystallography (50). We have shown previously that this region of class I molecules may be unique in cell signaling (37). Data presented in this work show that activation of HLA-A2 molecules with mAbs RG1 and CR11-351 directly signals rapid morphologic changes, phosphatidylserine exposure, and cell death in Jurkat cells expressing HLA-A2 molecules. In contrast, none of these effects was observed with mAbs against distant class I α1 or α2 epitopes. Thus, these results and previous observations (35, 37) clearly demonstrate a unique ability of the TCR-accessible region of the class I α2 domain to transduce regulatory signals. Importantly, this strongly suggests that the TCR can impose regulatory signals and induce cell death upon binding the target MHC class I/peptide complex.

Previously we observed that all normal PBMC and most, but not all, leukemic hemopoietic cell lines expressing HLA-A2 specifically responded to RG1 and CR11-351 with profound cell aggregation. Furthermore, mitogen-stimulated T cells responded significantly to these mAbs with growth inhibition. In contrast, the majority of tested leukemic cell lines were unaffected (37 and data not shown). Thus, activation of the TCR-binding class I α2 region gave different responses in normal and leukemic cells. The Jurkat E11 T cell line expressing HLA-A2 molecules was found to respond very rapidly to RG1 and CR11-351 activation with cell aggregation and morphologic changes, thus representing a model system for class I α2 signaling.

Assessments with RG1 and CR11-351 showed a rapid and profound induction of phosphatidylserine exposure in HLA-A2-transfected Jurkat cells, as determined by Annexin V-FITC binding. These studies were in agreement with the observed changes in cell size and granularity. Thus, activation of the TCR-binding region of the HLA class I α2 domain imposed responses in the Jurkat transfectants characteristic of apoptotic cell death.

The role of the cytoplasmic class I region in signaling apoptosis has not been determined. We have shown previously that RG1 and CR11-351 induce cell aggregation in Jurkat E13 transfectants expressing HLA-A2 where the cytoplasmic residues are deleted (37). Similarly, we now describe that RG1 and CR11-351 induce rapid morphologic changes and phosphatidylserine exposure in Jurkat E13 cells. Thus, class I α2-mediated cell aggregation responses and apoptotic responses are apparently signaled independent of the cytoplasmic class I domain. Importantly, these data strongly indicate that class I α2-mediated signaling requires engagement of a membrane-bound coreceptor.

Fas activation is implicated in controlling the level of activated T cells and as a major pathway in mediating CD8+CTL target cell lysis. We showed that Jurkat E11 cells constitutively expressed CD95, and it was therefore of interest to determine whether the CD95 pathway was involved in class I signaling and also to compare the effects of the anti-Fas mAb CH11 with RG1 and CR11-351. Blocking of possible CD95-CD95L interactions with neutralizing anti-Fas mAb or human rFas-Fc chimera did not influence the ability of RG1 or CR11-351 to induce apoptosis. Furthermore, we found that class I activation more rapidly induced changes in cell morphology and phosphatidylserine exposure compared with CD95 activation. We also assessed the mAb’s potential to induce DNA fragmentation in Jurkat E11 with the TUNEL assay. Interestingly, whereas CH11 induced a significant response within 4 h, only a minor shift in staining intensity was observed following RG1 and CR11-351 stimulation. Thus, a direct or indirect activation of the CD95 pathway appeared not to be involved in HLA class I α2-mediated signaling of cell death. Furthermore, in contrast to Fas- and perforin-mediated pathways, MHC class I α2-mediated signaling of cell death appears to proceed independent of DNA fragmentation into oligonucleosomes. Triggering of programmed cell death without observable DNA fragmentation into oligonucleosomes has also been observed following activation of other cell surface Ags (52, 53). Ligation of CD45 on T and B lymphocytes rapidly induces this kind of apoptosis (52), and it would therefore be of great interest to determine whether CD45 is activated in HLA class I α2-induced apoptosis.

The basis for the fundamental differences in apoptotic responses is not known. The Fas-mediated pathway involves ICE and CPP32 activation, as demonstrated by assessments with inhibitory peptides to these enzymes (64, 65). In contrast, we found no requirements for these proteases in class I α2-mediated signaling of apoptosis. Thus, CD95 and HLA class I α2-mediated death signaling clearly depend on different signaling pathways.

It has been demonstrated that susceptibility to CTL-induced apoptosis is a function of the proliferative status of the target (67). The cell cycle status may also be important in relation to responses to HLA class I α2 activation. Whereas anti-class I α2 activation rapidly induced apoptosis in more than 60% of Jurkat E11 cells, a resistant subpopulation was always observed. Interestingly, with unstimulated PBMC we observed no mAb RG1 or CR11-351 impact on cell morphology or viability. However, the mAbs clearly induced cell aggregation (data not shown). In contrast, cells preactivated with OKT3 were clearly sensitive to RG1 and CR11-351. Thus, normal nonproliferating PBMC may signal cell aggregation, but apparently not cell death upon activation of the TCR-accessible region of the class I α2 domain. This may favor a model in which the TCR participate directly on two levels: 1) to activate new adhesion molecules and 2) to deliver a lethal hit to transformed (i.e., proliferating) cells. Thus, resting cells with latent virus infections may potentially escape this type of death response.

Recent studies have implicated a functional role of the class I α3 domain in signaling apoptosis (40, 41). In the case of Jurkat cells, these were shown insensitive to apoptotic responses induced by class I α3 activation alone (41), indicating distinct roles for the α2 and α3 domains in signaling. It is therefore possible that class I α2 and α3 activation by the respective ligands TCR and CD8 may coregulate apoptotic responses in a target cell.

In conclusion, our data suggest that peptide-directed TCR interaction with target cell MHC class I molecules may directly induce phosphatidylserine exposure for target clearance by phagocytes, and furthermore, effect cell disintegration by programmed cell death. Thus, MHC class I-mediated signaling responses may in some target cells directly determine sensitivity or resistance to CD8+CTL-mediated lysis.

Our results suggest a model in which the physical binding of CTL TCR with target peptide/MHC class I complex per se represents the first line of cytotoxic defense against virus-infected or transformed cells. The success of this strategy will depend on the ability of the target cell to respond with sufficient activation of the HLA class I adhesion and death-related signaling pathways.

Acknowledgments

We thank Dr. Peter E. Lipsky for providing the Jurkat transfectants and Dr. Carlo Russo for providing the CR11-351 hybridoma.

Footnotes

  • ↵1 This study was supported financially by the Norwegian Cancer Society.

  • ↵2 Address correspondence and reprint requests to Dr. Rolf D. Pettersen, Department of Pediatric Research, Rikshospitalet, The National Hospital, N-0027 Oslo, Norway.

  • 3 Abbreviations used in this paper: ICE, IL-1β-converting enzyme; CD95L, CD95 ligand; PI, propidium iodide; TUNEL, terminal deoxynucleotidyl transferase-mediated dUTP-biotin nick end labeling.

  • Received August 18, 1997.
  • Accepted January 7, 1998.

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The Journal of Immunology
Vol. 160, Issue 9
1 May 1998
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