Functional Analysis of B and T Lymphocyte Attenuator Engagement on CD4⁺ and CD8⁺ T Cells¹

T cell specificity results from TCR-mediated recognition of specific Ag in conjunction with MHC on APCs (1). However, the functional outcome of this initial TCR engagement is modulated by secondary signals, which can have costimulatory or coinhibitory functions. The group of coinhibitory molecules includes CTLA-4 and programmed cell death-1 interacting with CD80 and CD86 or PD-L1 and PD-L2, respectively (reviewed in Ref.2).

We recently cloned a gene from differentiating thymocytes that encodes a member of the Ig superfamily of cell surface proteins (3). This protein, independently isolated from Th1 cells, has been named B and T lymphocyte attenuator (BTLA)³ (4). BTLA is most highly expressed by B cells, followed by T cells and APC (3), although its expression on macrophages has recently been reported to differ between mouse strains (5). The cytoplasmic tail of BTLA contains ITIM motifs, suggesting an inhibitory function for the protein. Indeed, lymphocytes from BTLA-deficient mice were found to be hyper-responsive to anti-CD3-mediated proliferation (3, 4). In contrast to CTLA-4 and programmed cell death-1, which are induced during T cell activation, BTLA is constitutively expressed on naive T cells and is further up-regulated upon T cell activation (3). Although BTLA was thought originally to be an additional member of the B7 family-recognizing T cell surface proteins, it has recently been demonstrated that herpes virus entry mediator (HVEM), a known ligand for the TNF family member LIGHT (derived from homologous to lymphotoxin, inducible expression), is the ligand for BTLA (6, 7). Like BTLA, HVEM is expressed by T and B cells although the hierarchy of expression on naive cells is reversed from that of BTLA (i.e., T>B) (8). A soluble form of HVEM has been shown to inhibit anti-CD3-induced proliferation of CD4⁺ T cells, and this effect could be reversed by anti-BTLA Ab (7). In addition, T cells from HVEM-deficient mice are hyperproliferative in response to Con A (9). Together, these data are consistent with a coinhibitory signal mediated by a BTLA-HVEM interaction.

We previously generated a panel of BTLA-specific mAb that preferentially react with the BTLA.2 allelic form of the protein (3). In this study we further characterize the functional properties of one of these Abs, PK18. This Ab has agonistic properties that allow analysis of the effects of BTLA activation and proliferation of naive CD4⁺ and CD8⁺ T cells.

C57BL/6J (B6), BALB.K, BTLA-deficient (BTLA^−/−), and AND TCR-transgenic (Tg) mice on an H-2^b background (10) were used in these studies. All animals were bred at The Scripps Research Institute and maintained under specific pathogen-free conditions. Experiments were conducted in accordance with National Institutes of Health Guidelines for the Care and Use of Animals and with an approved animal protocol from The Scripps Research Institute animal care and use committee.

BTLA-specific rat mAb PK3 (IgMκ), PK18 (IgG1κ), and mouse mAb PJ196 (IgG1κ) were used in these studies. The production of mAb was described previously (3). The following additional Abs were used (all from eBioscience, unless stated otherwise): FITC- or APC-conjugated anti-CD8 (clone 53-6.7), PE-conjugated anti-CD3 (clone 145-2C11), PE- or allophycocyanin-conjugated anti-CD4 (clone RM4-5), biotinylated anti-CD69 (clone 1.2F3), and allophycocyanin-conjugated anti-CD25 (clone PC61.5). Secondary staining reagents included PE-conjugated anti-rat IgM (clone G5-238; BD Pharmingen) and PE-labeled streptavidin (Biomeda). Cell surface staining was performed as described previously (3). In some instances cells were preincubated with anti-CD16/CD32 to block FcRs. For the study of T cell apoptosis, the annexin V-FITC apoptosis detection kit was used according to manufacturer’s instructions (BD Pharmingen). Briefly, T cells were washed with ice-cold PBS (Invitrogen Life Technologies), resuspended in annexin V-binding buffer (10 mM HEPES/NaOH (pH 7.4), 140 mM NaCl, and 2.5 mM CaCl₂), stained with annexin V-FITC and -propidium iodide, and assayed within 1 h by flow cytometry. As a positive control, T cell apoptosis was induced by dexamethasone (100 nM; Sigma-Aldrich). At least 20,000 viable cells were live-gated on a FACSCalibur or digital LSRII using CellQuest software (BD Biosciences) and were analyzed using FlowJo software (TriStar).

All culture experiments were performed in Click’s medium (Irvine) containing 10% heat-inactivated FCS (Invitrogen Life Technologies), penicillin/streptomycin (Invitrogen Life Technologies), 200 mM glutamine, and 50 μM 2-ME (Sigma-Aldrich). In some cultures anti-CD28 (clone 37.51; eBioscience) was used at a concentration of 10 μg/ml, or recombinant murine IL-2 (eBioscience) was added at various concentrations. Cells were cultured under standard conditions at 37°C in 5% CO₂. T cells from spleens or lymph nodes were enriched by negative selection using anti-B220 (clone RA3-6B2; eBioscience), anti-A^b class II MHC (clone Y3P) Abs, and anti-rat IgG magnetic beads (Qiagen). For Ag-specific proliferation assays, AND TCR-Tg mice were used as an enriched source of naive CD4⁺ T cells. For CD4⁺ T cell enrichments anti-CD8 (clone 53-6.7; eBioscience), Ab was added to the mixture. The purity of the resultant cell populations was >95%. In some experiments, naive CD4⁺CD25⁻ T cells were purified by sorting on a FACSDiva or FACSAria (BD Biosciences) to a purity of >99%.

Total T cells were washed with PBS, resuspended at a density of 1 × 10⁷ cells/ml in PBS containing 2 μM CFSE (Molecular Probes), and incubated for 5 min at room temperature. Labeling was terminated by addition of FCS to a final concentration of 10%, and cells were washed once in culture medium. The frequency of CD4⁺ or CD8⁺ T cell precursors that were activated to divide was calculated as the number of CFSE⁺ cells in each division/2ⁿ, where n is the number of divisions, divided by the input number of cells.

A total of 1 × 10⁶ T cells/ml were cultured in 100 μl or 1 ml in 96- or 24-well, flat-bottom plates, respectively, coated with 1:1 ratios of anti-CD3 (clone 145-2C11; eBioscience) in combination with rat IgG (Jackson ImmunoResearch Laboratories), PK18, or PJ196 at the indicated total Ig concentration. In some experiments, T cells were stimulated with a combination of 50 ng/ml PMA and 500 ng/ml ionomycin (Sigma-Aldrich). For analysis of Ag-specific T cell proliferation, CD4⁺ T cells were cultured at 5 × 10⁵ cells/ml with 2 × 10⁶ cells/ml irradiated (3300 rad) BALB.K splenocytes in the presence of 100 nM pigeon cytochrome c peptide. For proliferation studies, T cells were cultured in 96-well plates for a total of 72 h and pulsed with 1 μCi/well [³H]thymidine (PerkinElmer) for the last 16 h. Cells were harvested onto glass-fiber filters (Brandel), and [³H]thymidine incorporation was determined. To analyze T cell activation markers, cultures were harvested at various times and were analyzed by flow cytometry. In other experiments, CFSE-labeled total T cells were stimulated for 72 h and then analyzed.

Culture supernatants were assayed for mouse IL-2 using a standard sandwich ELISA (eBioscience). Maxisorb plates (Nunc) were coated with anti-IL-2 mAb (clone JES6-1A12), and detection was performed using the corresponding biotinylated mAb. The plates were developed using streptavidin-HRP and tetramethxylbenzidine as substrate. Plates were read on a plate reader at 450 nm, and data were analyzed using Softmax software (reader and software from Molecular Devices). Samples were performed in duplicate. The assay was standardized with recombinant murine IL-2, and detection ranged from 1 to 1000 pg/ml.

Jurkat T cells were transfected with a construct encoding murine BTLA-yellow fluorescent protein (YFP) fusion protein (base vector pEYFP-N1; BD Clontech) by electroporation and were stained and analyzed by flow cytometry on day 2. PK18 does not stain untransfected Jurkat T cells. Site-specific mutations were made using the GeneTailor Site-Directed Mutagenesis System (Invitrogen Life Technologies) according to the manufacturer’s instructions. All mutant constructs were sequence verified (TSRI Center for Nucleic Acids Research).

We previously generated BTLA-specific mAb by immunizing rats or mice with a recombinant soluble form of the protein (3). All Abs generated reacted preferentially with BTLA expressed on cells derived from B6, but not BALB/c, mice, the results of allelic variation in the BTLA external domain (3). We also demonstrated that coimmobilized rat anti-BTLA mAb PK18 is a potent inhibitor of anti-CD3 Ab-induced T cell proliferation (3) (Fig. 1,A). Soluble anti-BTLA Ab PK18 also significantly inhibited proliferation of CD4⁺ T cells at limiting concentrations of plate bound anti-CD3 Ab (Fig. 1,B). As a control, PK18 had no effect on the proliferation of CD4⁺ T cells derived from BTLA-deficient mice (Fig. 1,B). The PK18-mediated inhibition of T cell proliferation is not due to altered kinetics of the response (Fig. 1,C). Seven additional mAb (one rat- and six mouse-derived) had no effect on (or, in some experiments, enhanced) T cell proliferation (for example, Ab PJ196 in Fig. 1 A). The inability of other anti-BTLA Abs to inhibit T cell proliferation is not due to poor binding. In fact, PJ196 stains cells brightly even at extremely low concentrations of Ab, whereas PK18 is, somewhat surprisingly, a relatively poor staining reagent (data not shown). This suggested that PK18 and other Abs, such as PJ196, might differ in the respective epitopes on BTLA recognized.

We took advantage of the ability of PK18 and the other mAb to distinguish BTLA.2 (B6-derived) and BTLA.1 (BALB/c-derived) to map residues in the external domain of BTLA that are important for binding. BTLA.2 and BTLA.1 differ in 11 aa in the external domain of the cell surface protein (positions 41, 45, 47, 52, 55, 63, 74, 85, 91, 102, and 143, with the start methionine designated position 1). Murphy et al. (5) reported that residue 74 was an asparagine in both B6- and BALB/c-derived BTLA, whereas other strains had a histidine at this position. Our BALB/c-derived clone also had a histidine at this position, but whether this reflects differences in substrains is not known. A construct encoding a fusion of BTLA.2 and YFP was subjected to site-directed mutagenesis to allow the expression of mutants of BTLA.2-YFP that had individual residues changed to those in BTLA.1 (with the exception of codons encoding residues 45 and 47, which were mutagenized together). Individual BTLA-YFP constructs were transfected into Jurkat cells, the cells from each transfection were aliquoted for staining with various BTLA mAb, and the cells were subjected to two-color FACS analysis.

The cell surface expression of BTLA-YFP was quite variable between individual transfected cells despite similar expression of YFP (data not shown). Whether this reflects tight regulation of cell surface transport of the BTLA protein or a property of this particular fusion protein is not known. Nevertheless, cells were gated for a narrow range of expression of YFP, and then anti-BTLA mAb staining was assessed. All 10 mutant proteins were expressed on the cell surface (i.e., at least one anti-BTLA Ab could bind at near wild-type (wt) levels), but only three had significantly reduced PK18 staining compared with cells expressing wt BTLA-YFP (Fig. 1,D and data not shown). Mutation of residue 41 from BTLA.2 to BTLA.1 (P41E) completely abolished binding of PK18, whereas C85W and T45N/T47K mutations had lesser effects on Ab binding. Binding of PJ196 to mutant 45/47 was also reduced, as was binding to mutant 41, although to a somewhat lesser extent. A more modest reduction in binding of PJ196 to the 85 mutant was also observed. Staining of Ab PK3 to all three mutants is also shown for comparison, with little reduction in staining to mutants 41 and 45/47, and modest reduction in staining to mutant 85 (Fig. 1 D). Together, the data indicate that PK18 and PJ196 have overlapping, but distinct, epitopes that probably account for their distinct biological activities.

To better understand the functional consequences of BTLA engagement on CD4⁺ and CD8⁺ T cells, we took advantage of the agonistic ability of the PK18 mAb. T cells were labeled with CFSE and stimulated for 3 days with anti-CD3 in combination with PK18 or control rat IgG. The CFSE profile of total T cells revealed an almost complete block of T cell proliferation, consistent with results obtained by thymidine uptake (Fig. 2,A). However, when CD4⁺ and CD8⁺ T cells were individually analyzed in these assays, there was a near complete block of CD4⁺ T cell proliferation, whereas a significant proportion of CD8⁺ T cells had divided (Fig. 2,A). From these data we also calculated the precursor frequency of cells that were activated to divide in these cultures (Fig. 2 A). Only 2.2% of CD4⁺ precursor T cells underwent proliferation when activated by anti-CD3 in the presence of PK18, compared with 33.3% in control cultures. Moreover, the few CD4⁺ cells that did proliferate underwent a single division after anti-CD3 and PK18 stimulation, compared with up to three divisions in control cultures. In contrast, 57.9 and 28.5% of CD8⁺ precursors proliferated in the presence of anti-CD3 and rat IgG or PK18, respectively, and some cells underwent three or four divisions even in the presence of PK18.

Both CD4⁺ and CD8⁺ T cells express BTLA, although CD4⁺ T cells have modestly higher cell surface expression (3). In addition, TCR-mediated activation of T cells up-regulates BTLA (3). Thus, the more profound inhibition of proliferation of CD4⁺ compared with CD8⁺ T cells could result from differences in expression. To address this, we stained T cells activated by anti-CD3 in the presence or the absence of anti-CD28-mediated costimulation for expression of BTLA (Fig. 2 B). Up-regulation of BTLA was detected as early as day 1 after activation of CD4⁺ T cells and reached maximal levels by day 3. Costimulation had little effect early, but overall produced an increase in the number of responding cells. A similar pattern was observed for CD8⁺ T cells, although up-regulation lagged behind that in CD4⁺ T cells, and the level of expression was ∼2-fold lower. Thus, differential effects of PK18 on the proliferation of CD4⁺ or CD8⁺ T cells could, at least in part, result from differential amounts of cell surface protein and differing kinetics of up-regulation of BTLA surface expression. However, the fact that anti-BTLA can inhibit cytokine production before BTLA up-regulation (see below) suggests that this may not be the only factor involved in the differential effects of BTLA engagement on CD4⁺ and CD8⁺ T cells.

It was possible that the failure of CD4⁺ T cells to proliferate in the presence of PK18 was due to the induction of cell death. However, no significant loss of cell number was observed in PK18-treated cultures (data not shown). In addition, no increase in apoptotic cells as assessed by propidium iodide/annexin V staining was observed after activation of CD4⁺ T cells for 6 h with anti-CD3 in the presence of PK18 compared with that in cells activated in control cultures (Fig. 3 A). Analysis 16 h after stimulation yielded similar results (data not shown).

We have previously reported that BTLA is expressed by CD4⁺CD25⁺ T cells, a subset that includes regulatory cells (T_regs) (reviewed in Ref.11). Thus, we also considered the possibility that PK18 was activating T_regs, which then acted to suppress the proliferation of CD4⁺ T cells. To test this, we purified CD4⁺CD25⁻ T cells and tested the effect of BTLA engagement on anti-CD3-mediated proliferation as before. As observed for unseparated CD4⁺ T cells, proliferation of CD4⁺CD25⁻ T cells was inhibited by PK18 (Fig. 3 B). Thus, the ability of anti-BTLA to inhibit CD4⁺ T cell proliferation is a direct effect on the CD4⁺ T cell itself.

Induction of proliferation is a relatively late event after T cell activation. To determine whether PK18 blocked all signals mediated by TCR engagement, we asked whether PK18 prevented the induction of other T cell activation markers. Interestingly, the induction of CD69 expression on both CD4⁺ and CD8⁺ T cells was not inhibited by PK18 (Fig. 4). In contrast, BTLA engagement led to inhibition of subsequent IL-2R (CD25) expression on both CD4⁺ and CD8⁺ T cells (Fig. 4). However, the decrease in CD25 expression was more pronounced on CD4⁺ T cells, consistent with the greater sensitivity of this population to PK18-mediated inhibition of proliferation. The increased frequency of CD25⁻ T cells, particularly pronounced among CD8⁺ T cells, was due to reduced proliferation in the presence of PK18 and, thus, increased frequency of unactivated cells in these cultures. Because IL-2 is necessary for T cell proliferation in vitro as well as for maintenance of IL-2R expression (12), these results suggested that the primary defect in anti-BTLA engagement could be cytokine production. Indeed, anti-CD3- and PK18-stimulated T cells showed a 80-fold reduction in IL-2 secretion compared with anti-CD3- and rat IgG-stimulated T cells (Fig. 5 A).

To determine whether the decrease in IL-2 production was the sole cause of the inhibition of T cell proliferation in these cultures, we performed add-back experiments with exogenous IL-2 (Fig. 5,B). The addition of 80 pg/ml recombinant murine IL-2, comparable to that produced by anti-CD3-activated T cells themselves in these cultures (Fig. 5,A), was not sufficient to rescue the proliferation of T cells in the presence of PK18 (Fig. 5,B), nor did this concentration of IL-2 restore CD25 to wt levels (Fig. 5,C). Thus, the failure to produce IL-2 is not the sole cause of T cell inhibition mediated by PK18 under these conditions. Nevertheless, high concentrations (800 pg/ml) of IL-2 are sufficient to restore nearly 70% of the T cell proliferative response achieved in control cultures (Fig. 5,B). Despite this, high concentrations of IL-2 were not sufficient to return the expression of CD25 to wt levels on CD4⁺ or CD8⁺ T cells activated in the presence of PK18, although, again, the effects of PK18 were more pronounced on the CD4⁺ T cell population (Fig. 5 C).

CD28 costimulation is a strong enhancer of IL-2 production in T cells (13) and thus might be expected to overcome PK18 inhibition. At low concentrations of anti-CD3, PK18 was still inhibitory, even in the presence of CD28 (Fig. 6,A). However, at higher concentrations of anti-CD3, addition of CD28 costimulation restored T cell proliferation to near normal levels even in the presence of PK18 (Fig. 6,A). Similar results were found when tracking individual divisions of CFSE-labeled CD4⁺ and CD8⁺ T cells stimulated with saturating amounts of anti-CD3 (Fig. 6,B). Under these conditions, CD28 costimulation was also able to restore CD25 expression on CD4⁺ and CD8⁺ T cells activated in the presence of PK18 (Fig. 6,C). As expected, the addition of anti-CD28 increased IL-2 production in these cultures ∼50-fold (Figs. 5,A and 6,D). Thus, the ability of anti-CD28 to overcome PK18 inhibition mirrors that observed after adding high concentrations of exogenous IL-2 (Fig. 5,C). However, even in the presence of costimulation and a high concentration of anti-CD3, the presence of PK18 caused a nearly 13-fold reduction in IL-2 secretion (Fig. 6 D). Thus, CD28-mediated costimulation does not ablate the inhibitory signal initiated by BTLA engagement.

Given that soluble PK18 could inhibit the proliferation of CD4⁺ T cells at limiting concentrations of plate-bound anti-CD3 in the presence of costimulation, we also tested the effect of PK18 on Ag-stimulated CD4⁺ T cells. AND TCR-Tg CD4⁺ T cells were stimulated in the presence of irradiated BALB.K splenocytes as a source of APCs. We have previously shown that BTLA is expressed on T cells and APC, and that PK18 binds preferentially to B6-derived splenocytes, but not BALB.K splenocytes (3). Thus, in these experiments PK18 would target the T cells and not the APC. Addition of PK18 to these cultures significantly inhibited T cell proliferation in response to peptide Ag (Fig. 6 E).

Our results demonstrated that BTLA engagement at the time of initial TCR activation was a potent inhibitor of aspects of T cell activation. However, the fact that CD69 induction was not affected by PK18 raised the possibility that the inhibitory effect of BTLA engagement might be a relatively late event. Thus, we isolated naive CD4⁺ T cells from B6 mice, stimulated them overnight with plate-bound anti-CD3, and then transferred the cells to fresh plates coated with additional anti-CD3 in combination with PK18 or rat IgG. Surprisingly, PK18 was able to inhibit CD4⁺ T cell proliferation even after an initial overnight activation with anti-CD3, similar to the inhibition observed for cells exposed to PK18 for the entire culture period (Fig. 7).

We have generated a panel of mAb specific for BTLA, a recently discovered member of the Ig superfamily of cell surface proteins (4). Only one of these Abs, PK18, was found to act as an agonist for this coinhibitory molecule, thereby inhibiting facets of T cell activation.

We were able to take advantage of the fact that PK18 preferentially binds the allele expressed in B6 mice, but not that in BALB mice (3), to perform initial epitope-mapping studies. Binding of PK18 to BTLA.2, but not BTLA.1, is critically dependent on residue 41, because the P41E substitution completely abolished Ab binding. Other Abs, for example, PK3, bound to the position 41 mutant at wt levels, indicating that the inability of PK18 to bind the mutant was not caused by a lack of surface expression. In addition, the binding of PK3 to mutant 41 indicates that other epitopes on the protein were not grossly distorted. The replacement of the proline at this position with a glutamic acid is likely to alter the local structure of the protein, but might also prevent Ab binding due to the addition of the negative charge.

Of the nine additional BTLA.2 mutants (encompassing 10 aa changes), only two, the substitution at position 85 and the double substitution at positions 45 and 47, had any effect on binding of PK18. Another allele-specific mAb, PJ196, which lacks inhibitory activity, was also affected to varying degrees by substitutions at these same positions. Substitution at 85 had a small effect on binding of PJ196, whereas mutants 41 and 45/47 had significantly greater effects on binding. Overall, however, the rank order of binding of PK18 and PJ196 to these mutants differed (for PK18 mutants, 41<45/47 and 85; for PJ196, 45/47<41≪85), consistent with overlapping, but distinct, epitopes recognized by the two Abs. Given that PJ196 is a better binder to cells than PK18 (data not shown), it seems likely that the dramatic difference in biological activity between these two mAb is due to the relatively subtle differences in the epitopes bound.

These data should be helpful in producing and screening for additional Abs that have agonist function. In an Ig fold, residues 41, 45, and 47 would be predicted to be on a solvent-exposed face of the protein (data not shown). Whether this region of BTLA is also involved in binding to HVEM, and whether there could be allelic differences in that binding, remain to be determined. Interestingly, two BTLA.2-specific Abs generated by Murphy et al. (5) appear to recognize epitopes distinct from that of PK18 or PJ196, although substitution at position 85 (our numbering) does eliminate binding of one of their Abs.

When assessed by uptake of thymidine, PK18 was found to be a potent inhibitor of T cell proliferation. However, single-cell analysis of cell division by CFSE staining revealed that CD4⁺ T cells were more susceptible to the inhibitory effects of PK18 than were CD8⁺ T cells. Similarly, PK18 also inhibited CD25 up-regulation on both CD4⁺ and CD8⁺ T cells, but the inhibition was again more pronounced on CD4⁺ T cells. Resting CD4⁺ T cells express modestly higher levels of BTLA than do CD8⁺ T cells (3) (Fig. 2 B). Moreover, as shown in this study, BTLA is up-regulated with faster kinetics on activated CD4⁺ compared with CD8⁺ T cells. Thus, we cannot rule out that differences in expression may be one factor in the increased susceptibility of CD4⁺ T cells to PK18-mediated inhibition. Alternatively, CD8⁺ T cells may be intrinsically less sensitive to the inhibitory effects of PK18, especially during their initial phase of proliferation, which may be IL-2 independent (14). We have seen similar results using a soluble form of HVEM (data not shown); thus, it is unlikely that these results are peculiar to the PK18 mAb.

Soluble PK18 also inhibited the proliferation of CD4⁺ T cells at limiting concentrations of plate-bound anti-CD3 (Fig. 1 B). This was somewhat surprising, because it was previously reported that BTLA phosphorylation, and thus presumably signaling, requires co-cross-linking with the TCR (4). In these latter experiments, T cell hybridomas overexpressing a Myc-tagged version of BTLA lacking the external domain were treated with anti-Myc and anti-CD3 Abs and cross-linked with a cross-reactive secondary Ab. Whether the lack of the BTLA external domain or the use of hybridoma cells affected the requirement for co-cross-linking in this system is not known. A soluble recombinant form of the BTLA receptor HVEM, however, was also reported to inhibit T cell proliferation induced by limiting concentrations of plate-bound anti-CD3 Ab, similar to the activity of PK18 (15, 16). Thus, PK18 appears to be a good mimic of the BTLA ligand.

We have found that BTLA engagement by PK18 is a potent inhibitor of IL-2 secretion. However, the addition of exogenous IL-2 at a concentration that is normally produced in these cultures was not able to rescue proliferation or CD25 expression on T cells activated in the presence of PK18. Thus, BTLA engagement has additional IL-2-independent inhibitory effects on T cell activation. In contrast to the effects on cytokine secretion, PK18 had no effect on the induction of CD69, indicating that BTLA engagement does not prevent initial activation of the T cell, but modifies the consequence of such activation. That BTLA might function as a response modifier is also supported by kinetic studies of susceptibility to a PK18-mediated inhibitory signal (see below). Although IL-2 is a potent mediator of initial T cell proliferation in vitro, IL-2 may play a role in vivo in subsequent phases of T cell clonal expansion and as a development of effector function in addition to its effects on T_regs (reviewed in Ref.17).

CD28-mediated costimulation lowers the threshold of T cell activation (18), potently enhances IL-2 production (13, 19) and IL-2R expression (20), and promotes progression through the cell cycle (21). At limiting concentrations of anti-CD3, PK18 was inhibitory even in the presence of anti-CD28. CD28 costimulation was able to counteract the inhibitory effects of PK18 on cell division in the presence of high concentrations of anti-CD3. Even under these conditions, however, IL-2 production was significantly inhibited by PK18. These findings are consistent with the ability of APC that coexpress HVEM and CD80 to inhibit the proliferation of TCR-Tg CD4⁺ T cells in response to low, but not high, concentrations of antigenic peptide (6). Similarly, high concentrations of PK18 can inhibit Ag-specific T cell responses (Fig. 6 D). This inhibitory effect was due to binding of PK18 to the T cell, because we used APC that would not be targeted by PK18. Thus, BTLA engagement under the limiting conditions of costimulation or TCR activation that may occur in vivo may deliver an inhibitory signal. In the presence of anti-CD3, -CD28, and -PK18, the remaining IL-2 produced was probably sufficient to induce T cell proliferation to near control levels, because similar concentrations of exogenously added IL-2 rescued the proliferation of T cells activated in the presence of PK18. It is also possible that the ability of CD28 costimulation to regulate progression through the cell cycle in an IL-2-independent manner (22) plays a role in counteracting the inhibitory effects of BTLA engagement on cell division.

BTLA up-regulation occurs over several days after T cell activation. CD28-mediated costimulation increased the frequency of cells that up-regulated BTLA, but did not alter these kinetics. Because the production of cytokine is a relatively early event, the ability of PK18 to modify this response suggests that BTLA engagement on naive T cells before up-regulation of the cell surface protein can still deliver an inhibitory signal. Nevertheless, up-regulation of BTLA in vivo may be an important factor in making T cells more susceptible to BTLA-mediated inhibition. Interestingly, the BTLA ligand HVEM, also expressed by naive T cells, is down-regulated with similar kinetics as those of BTLA up-regulation (9) (data not shown). This might suggest that a BTLA-HVEM interaction is unlikely to autonomously regulate T cell function.

Because PK18 failed to block CD69 up-regulation, we asked whether BTLA engagement on preactivated CD4⁺ T cells would still be inhibitory. Surprisingly, cell proliferation was inhibited when CD4⁺ T cells were exposed to PK18 as late as 16 h after initial culture with anti-CD3. Intravital imaging has recently shown that during the first hours of initial CD4⁺ T cell activation, serial short-lived interactions with APC bearing cognate peptide are sufficient to up-regulate CD69 (23). Subsequent phases associated with more stable T cell-APC contacts appear to be required for induction of cytokine and subsequent cell division. The fact that inhibitory signals mediated by BTLA can be temporally separated from initial TCR activation raises the possibility that BTLA engagement during this secondary phase of T cell-APC interaction could modify the outcome of the T cell response.

We thank Parinaz Aliahmad for critically reading this manuscript, and Alan Saluk, Cheryl Kim, and Kurt Van Gust for excellent technical assistance with cell sorting.

The authors have no financial conflict of interest.