Architectural Changes in the TCR:CD3 Complex Induced by MHC:Peptide Ligation¹

T cells, at the cellular level, and the TCR, at the molecular level, play pivotal roles in the initiation and regulation of the immune response. It has been known for some time that one of the most predictable events that occurs following ligation with MHC:peptide complexes is down-modulation of surface TCR complexes (1, 2, 3). However, elucidation of the molecular events that mediate TCR down-modulation and its role in T cell signal transduction remain elusive.

Renewed interest in the potential importance of TCR down-modulation was initiated by seminal studies suggesting that a single MHC:peptide complex can serially ligate and trigger ∼200 TCR, leading to an amplified and sustained signal (3). The serial triggering model is dependent on two factors: optimal kinetics in the interaction between the TCR and MHC:peptide complexes and ligation-induced TCR down-modulation. These factors combine to ensure the rapid clearance of ligated TCR, which releases the specific MHC:peptide complexes to bind to additional TCR (3, 4). Indeed, the rapid and constitutive internalization of the TCR is likely to facilitate serial triggering (5). However, it is well established that sustained signaling is required, particularly for naive T cells, to initiate full T cell activation (6, 7, 8, 9). Thus, it is currently unclear what molecular mechanism facilitates the rapid removal of ligated TCR, which is required to drive serial triggering, while at the same time maintaining the signaling necessary to activate the T cell.

The TCR is expressed as a large multichain complex composed of at least eight polypeptides (TCRαβ, CD3εγ, εδ, and ζζ) (10, 11). The cytoplasmic tails of the CD3 molecules contain immune receptor tyrosine-based activation motifs, which are phosphorylated upon TCR ligation (12). These act as key docking sites for Src homology 2 domain-containing proteins that mediate signal transduction and T cell activation. Our previous studies have suggested that TCR down-modulation is mediated by the intracellular retention of ligated complexes rather than an increase in the internalization rate (5). However, it is not clear how the intracellular sorting machinery distinguishes between ligated and unligated complexes.

The CD3 ζ-chain has long been considered to have a unique relationship with the TCR complex and is thought to play an important role in facilitating stable TCR cell surface expression (13, 14). Studies in CD3ζ knockout mice and CD3ζ-deficient T cell variants have clearly demonstrated that the TCR is not stably expressed at the cell surface in the absence of CD3ζ (15, 16, 17, 18, 19, 20, 21). In this instance, hexameric TCRαβCD3εγεδ complexes are assembled in the endoplasmic reticulum and subsequently transported via the Golgi apparatus to the lysosomes for degradation (13). Assembly with CD3ζ is thought to prevent trafficking to the lysosomes and stabilize surface expression of the TCR by shielding a CD3γ dileucine motif, which is known to mediate targeting to lysosomes for degradation (16, 22, 23, 24, 25). Finally, CD3ζ appears to turn over independently from the rest of the TCR complex on the cell surface (26).

Collectively, these studies suggested that CD3ζ has a unique and dynamic relationship with the TCR:CD3 complex and plays a key role in regulating its assembly, transport, and cell surface expression. We hypothesized that CD3ζ may also play a central role in mediating TCR down-modulation. To examine this possibility, we explored the relationship between CD3ζ and the TCR:CD3 complex after T cell activation by MHC:peptide complexes.

All CD3ζ mutants were made by recombinant PCR, using murine CD3ζ cDNA as a template (gift from L. Samelson, National Institutes of Health, Bethesda, MD) and cloned into pCIneo (Promega, Madison, WI) (27), pCIneo-internal ribosome entry sequence (IRES)⁵-green fluorescent protein (GFP), or murine stem cell virus (MSCV)-IRES-GFP, which is an MSCV-based retroviral vector. The latter two of these vectors contain an encephalomyocarditis virus IRES and GFP. IRES allows for translation of the test protein and GFP from the same mRNA, thus facilitating verification of coexpression while avoiding attachment at the protein level. CD3ζ-FLAG was produced by attaching the FLAG peptide to the CD3ζ N terminus. The first two amino acids were duplicated to ensure correct signal sequence cleavage. Thus, the sequence of the extracellular domain was QSDYKDDDDKQSFGLLDPK (duplicated residues in italics, FLAG peptide in bold, native CD3ζ extracellular sequence underlined). CD3ζ.K9R and K9S mutants were produced using oligonucleotide primers that contained an AAA to AGA mutation (K9R) or an AAA to AGC mutation (K9S). Details of the construction strategy and oligos used will be provided on request ([email protected]).

After sequence verification, CD3ζ-wild type (CD3ζ-WT) and CD3ζ-FLAG were transfected into CD3ζ loss variants of 3A9 (3A9.ζ-7 and 3A9.ζ-4, hen egg lysozyme (HEL)_48–62-specific, H-2A^k-restricted) (21) and 2B4 (MA5.8, pigeon cytochrome c (PCC)_88–104-specific, H-2E^k-restricted) (15) by electroporation (27, 28). Transfectants were selected with G418 and in some cases cloned or bulk sorted by FACS.

CD3ζ.WT, K9R and K9S DNA constructs were cloned into MSCV.IRES.GFP. Ecotropic retrovirus was produced in 293T cells by FuGENE-mediated transfection. Briefly, 293T cells (provided by D. Baltimore; 2 × 10⁶ cells in a 10-cm tissue culture petri dish, cultured overnight) were transfected with the CD3ζ constructs (4 μg) and the helper plasmids pEQ.PAM-E (4 μg) and pVSVg (2 μg) using FuGENE transfection reagent (Roche, Basel, Switzerland) according to the manufacturer’s instructions and were incubated overnight at 37°C. Medium was changed after 24 h, and after 72 h supernatant was collected and filtered (0.45 μm pore size filter). Viral titer was determined by 3T3 transduction, and the percentage of GFP cells was determined by flow cytometry. T cell hybridomas (10⁴ in 1 ml medium plus 6 μg/ml polybrene) were transduced at a multiplicity of infection of 5:1 and were cultured at 37°C for 12 h. To overcome the low level of ecotropic receptor on the surface of hybridomas, this transduction procedure was repeated five times. Expression was assessed by flow cytometry.

Assays were performed as previously described (29). Briefly, hybridomas (5 × 10⁴/well) were stimulated with LK35.2 cells (2.5 × 10⁴/well; murine B cell lymphoma; H-2A^kd) as APC and peptides or HEL protein (Sigma-Aldrich, St. Louis, MO) at the concentration indicated. Supernatants were removed after 24 h for estimation of IL-2 secretion by culture with the IL-2-dependent T cell line, CTLL-2, as described. For some assays, IL-2 was determined using a particle-based flow cytometric assay as previously described (30).

The assay for down-modulation of TCR:CD3 was performed as described previously (5, 27). Briefly, T cell hybridomas (5 × 10⁴; 100 μl) were cultured in flat-bottom, 96-well microtiter plates with LK35.2 cells that had been prepulsed overnight with or without 10 μM peptide (HEL_48–62 for 3A9 or PCC_88–104 for 2B4). At the time points indicated, cells were transferred to V-well microtiter plates for flow cytometry, which was performed as detailed previously (27, 29). Briefly, cells were stained with anti-CD3ε-FITC or -PE (all Abs from BD PharMingen (San Diego, CA) unless stated otherwise) and anti-FLAG-biotin (cation-independent M2 mAb; Sigma-Aldrich) plus streptavidin-allophycocyanin. Cells were stained with anti-H-2A^k-PE to gate out B cells, and with propidium iodide for live/dead cell gating. The percent down-modulation of TCR:CD3 was determined from the median values using the unstimulated controls as reference.

Assays were performed as previously described with some modifications (5). Briefly, T cells (5 × 10⁶/IP) were cultured at 37°C with APC (LK35.2 ± 10 μM HEL; 5 × 10⁶/IP) for 5 or 24 h. In some experiments LK35.2 cells expressing GFP were used to facilitate enumeration and equalization of T cells (GFP⁻) between samples. In Src family protein tyrosine kinase (PTK) inhibition studies, T cells and APCs were incubated separately for 2 h at 37°C in the presence or the absence of 10 μM 4-amino-5-(4-chlorophenyl)-7-(t-butyl)pyrazolo[3,4-d]pyrimidine (PP2) (Calbiochem, San Diego, CA) before being cultured together for 24 h at 37°C. Cells were surface-biotinylated with 1 mg/ml sulfo-N-hydroxysuccinimide (NHS)-biotin (Pierce, Rockford, IL) in HBSS for 30 min on ice. Excess biotin was quenched with 20 mM l-lysine/HBSS, and the cells were washed three times. Dead cells were removed by density gradient centrifugation using Lymphocyte Separation Medium (Cappel/ICN Pharmaceuticals, Costa Mesa, CA). Cells were lysed in 1% digitonin lysis buffer (1% digitonin (Wako Biochemicals, Osaka, Japan; Calbiochem, San Diego, CA), 0.05 M Tris (pH 7.4), 0.15 M NaCl, 2 mM Pefablock (Roche Applied Science, Indianapolis, IN), 40 μg/ml aprotinin, and 20 μg/ml leupeptin) at 4°C for 30 min, and the lysate was precleared with heat-killed formalin fixed Staphylococcus aureus (Pansorbin) cells (Calbiochem) for 1 h at 4°C. The lysate was then immunoprecipitated with protein A-Sepharose (Amersham Pharmacia Biotech, Piscataway, NJ) precoated with anti-TCR-Cβ (H57-597), twice sequentially (1 h each) at 4°C, followed by protein A-Sepharose precoated with anti-CD3ζ (H146-968) for 1 h at 4°C. Sepharose beads were washed with 0.2% digitonin buffer (three times) and immunoprecipitated proteins eluted in an equal volume of 2× SDS sample buffer at 75°C for 5 min.

In some experiments proteins bound to anti-TCR-Cβ-coated protein A-Sepharose beads were eluted, and the protein complexes were disrupted by boiling for 5 min in 1% SDS. Supernatants were collected, and 900 μl of 1% Nonidet P-40 lysis buffer was added. CD3ζ-FLAG proteins were then immunoprecipitated from the lysate with anti-CD3ζ-coated protein A-Sepharose beads for 1 h at 4°C. Unbound proteins remaining in the lysate were precipitated with 12% TCA by vortexing and placing on ice for 45 min. The precipitated proteins were pelleted by centrifugation at 12,000 × g for 10 min at 4°C. The pellet was then washed with acetone (−20°C), vortexed, centrifuged again, allowed to air dry, and resuspended in 1× SDS running buffer. Samples were subsequently treated as described above.

Eluted proteins were resolved by 10% SDS-PAGE or 12% NuPage bis-Tris precast gels (Invitrogen, San Diego, CA) and transferred onto a polyvinylidene difluoride membrane. Blots were blocked with 5% milk powder in PBS-Tween 20 (1 h at room temperature), followed by streptavidin-HRP or streptavidin-alkaline phosphatase (streptavidin-AP; 60 min at room temperature). Blots were developed using ECL Plus (Amersham Pharmacia Biotech, Little Chalfont, U.K.) or Vistra ECF substrate (Amersham Pharmacia Biotech). Occasionally, membranes were stripped in 100 mM 2-ME, 2% (w/v) SDS, and 0.1 M Tris (pH 6.7) in PBS at 50°C for 30 min. Membranes were then washed in PBS/0.2% Tween 20 and incubated with a 1/4 dilution of anti-CD3ζ (H146-968) and anti-CD3ε (HMT3.1) hybridoma supernatants. Ab binding was detected using protein A/G-AP (Amersham Pharmacia Biotech). Densitometric analysis was performed with QuantityOne (Bio-Rad, Hercules, CA) or ImageQuant (Amersham Pharmacia Biotech).

Biotinylated proteins were purified from cell lysates using SoftLink Soft Release Avidin Resin (Promega). Briefly, resin was washed with 0.1 M NaPO₄ (pH 7.0) and preadsorbed in 5 mM biotin/0.1 M NaPO₄ for 20 min at 4°C with rocking. Beads were regenerated by washing three times with 1 ml of 10% acetic acid, followed by three times with 1 ml of 0.1 M NaPO₄ (pH 7.0). For the final wash, the beads were incubated for 30 min at 4°C with rocking to allow avidin to refold. Lysate containing biotinylated proteins was then added to the resin and incubated at 4°C for 30 min with rocking. The resin was washed with three times with 1 ml of 0.2% digitonin wash buffer, and biotinylated proteins were eluted with 10 mM biotin in 1% digitonin lysis buffer (three times, 500 μl).

CD3ζ loss variants of the 3A9 and 2B4 T cell hybridomas (3A9.ζ− and 2B4.ζ−) were transfected with either CD3ζ-WT or an N-terminal FLAG-tagged CD3ζ (CD3ζ-FLAG) to follow the fate of CD3ζ after TCR ligation with MHC:peptide complexes. Both CD3ζ constructs restored surface expression of the TCR:CD3 complex as well as function, as determined by IL-2 production, to levels comparable to those in the parental 3A9 and 2B4 T cell hybridomas (Fig. 1, A and B, and data not shown). Mock transfections with empty vector failed to restore TCR expression or function (data not shown). As TCR is not stably expressed on the cell surface in the absence of CD3ζ (13, 15, 16, 21), it is reasonable to assume that all surface TCR:CD3 complexes on the CD3ζ-FLAG transfectants contain FLAG-tagged CD3ζ. Indeed, both CD3ζ-WT and CD3ζ-FLAG restored TCR expression comparably (Fig. 1,A). However, minimal FLAG staining was observed, suggesting that the epitope is concealed within the complex (Fig. 1,A). As expected, both the 3A9 and 2B4 transfectants down-modulated TCR following stimulation with Ag-pulsed B cells (Fig. 1 C). Surprisingly, there was a substantial increase in FLAG staining, which rose ∼25-fold in the 2B4 transfectants and ∼400-fold in the 3A9 transfectants 10–12 h poststimulation. These data imply that after TCR ligation, the TCR:CD3 complex undergoes structural alterations, such that the extracellular portion of CD3ζ-FLAG becomes exposed for Ab binding. The high level of CD3ζ-FLAG staining compared with TCR staining at 12 h poststimulation suggests that although TCR ligation induces down-modulation of the TCR:CD3 complex, CD3ζ may remain on the cell surface. Interestingly, the increase in FLAG staining was delayed relative to maximal TCR down-modulation. Although 3A9 TCR down-modulation was maximal after 3 h, CD3ζ-FLAG increase was not evident until 5 h. The reason for this delay is unclear, but it is possible that some time is required before a sufficient quantity of free CD3ζ has accumulated to a detectable level by flow cytometry. Alternatively, CD3ζ may be down-modulated with the ligated TCR complex, separated in an intracellular compartment, and thus not be seen as free CD3ζ until it was recycled back to the cell surface. This would be synonymous with our previous suggestions regarding the transport of nonligated TCR (5). Of course, such a lag may also be a reflection of the appearance of newly synthesized CD3ζ after TCR ligation.

The relationship between CD3ζ and the TCRαβCD3εγδ complex was studied further using surface biotinylation and sequential IP. The CD3ζ-WT and CD3ζ-FLAG transfectants of the 3A9ζ-hybridoma were surface-biotinylated 5 and 24 h after incubation with B cells precultured with or without Ag. Dead cells were removed to ensure that only surface-biotinylated proteins from viable cells were included. Cell lysates were immunoprecipitated sequentially with anti-TCR-Cβ three times to remove TCR complex-associated CD3ζ, followed by anti-CD3ζ to determine the amount of free surface-biotinylated CD3ζ.

In unstimulated CD3ζ-WT transfectants, there was some labeling of TCR-associated CD3ζ; however, very little free CD3ζ was observed. In the CD3ζ-FLAG transfectants, some biotinylated free CD3ζ was seen (Fig. 2). This increased biotin label on free CD3ζ may be due to increased biotinylation efficiency, as the FLAG tag contains two additional lysine residues. The relatively small amount of free CD3ζ-FLAG is consistent with the minimal surface FLAG staining observed on resting T cells (Fig. 1,A). Interestingly, no TCR-associated CD3ζ-FLAG biotinylation was detected in unstimulated cells, suggesting that the tag is buried in the complex. Efficient biotinylation may also have been precluded if the extracellular lysine residues had formed stable interactions with adjacent residues in the complex. It is important to note that the lack of CD3ζ-FLAG biotinylation on resting cells is not due to an absence of associated protein, as the presence of CD3ζ-FLAG was confirmed by subsequent immunoblot analysis using an anti-CD3ζ Ab (data not shown and Figs. 3 and 4). After antigenic stimulation, two changes in the pattern of TCR:CD3 biotinylation were observed. First, there was an increase in the TCR-associated CD3ζ biotin signal (Fig. 2). Second, the free CD3ζ biotin signal increased significantly, which was particularly evident after 24 h. In contrast, there was little change in the level of CD3εγδ biotinylation.

Based on these observations, three key issues arose. Firstly, it was possible that the increased signal intensity at ∼16 kDa was due to recruitment of another protein(s) to the TCR:CD3 complex after TCR ligation. To assess this possibility, CD3ζ-FLAG transfectants of the 3A9.ζ-hybridoma were surface-biotinylated 24 h after incubation with B cells precultured with or without Ag. We preferentially chose to use the CD3ζ-FLAG transfectants for these experiments because it is more readily biotinylated and thus easier to detect. Cell lysates were then immunoprecipitated with anti-TCR Cβ Ab (Fig. 3,A, lanes 1 and 2). As shown in Fig. 2, increased signal intensity at ∼16 kDa was again evident after Ag stimulation without significant alteration in the biotinylation of CD3εγδ. Proteins bound to the anti-TCR Cβ Sepharose beads were eluted, and the TCR complex was disrupted by boiling in the presence of 1% SDS. CD3ζ-FLAG proteins were subsequently immunoprecipitated with anti-CD3ζ Ab (Fig. 3,A, lanes 3 and 4). The remaining proteins that were not bound by the anti-CD3ζ Ab were precipitated with TCA before electrophoresis (Fig. 3,A, lanes 5 and 6). After detection of biotinylated proteins using streptavidin-AP, the blot was stripped and reprobed with anti-CD3ζ Ab to verify that all the CD3ζ protein had been immunoprecipitated, and none remained in the unbound fraction (Fig. 3 B). It was clear from this experiment that essentially all the ∼16-kDa biotinylated protein detected after anti-TCR-Cβ IP was CD3ζ, as evidenced by its complete removal from the lysate after IP with anti-CD3ζ.

Secondly, as visualization of proteins in this system is dependent on their ability to be biotinylated, it was unclear whether the increased CD3ζ signal intensity represented an increase in the amount of CD3ζ protein or an increase in the biotinylation state of CD3ζ. Differentiation between these possibilities required selective quantitation of cell surface CD3ζ protein. Cells were surface-biotinylated and lysed as described; however, before IP with anti-TCR-Cβ, biotinylated proteins were purified from the lysate using a SoftLink monomeric avidin system. This allowed for the selective IP of cell surface proteins. The eluted biotinylated proteins were then sequentially immunoprecipitated with anti-TCR-Cβ and anti-CD3ζ as described above. Upon probing with streptavidin-AP, the characteristic pattern of increased TCR-associated and free CD3ζ signal intensity was observed after Ag stimulation (Fig. 4,A, upper panel; compare lanes 1 and 4, and lanes 3 and 6), with densitometric analyses revealing 2.4- and 8.5-fold increases in TCR-associated and free CD3ζ, respectively (Fig. 4,B, upper panels). The blots were then stripped and reprobed with a mixture of anti-CD3ζ and anti-CD3ε Abs (Fig. 4 A, lower panel). The anti-CD3ε Ab was included to enable normalization of the CD3ζ signal intensity relative to the rest of the TCR:CD3 complex. Densitometric analyses of both TCR-associated and free CD3ζ protein revealed no increase in TCR-associated CD3ζ protein. However, a 3-fold increase in free CD3ζ protein was detected after Ag stimulation. The reduced CD3γδε-biotin and CD3ε protein signal after ligation is due to TCR down-modulation. Thus, the increased CD3ζ biotin signal observed after TCR ligation corresponds to an increased biotinylation state of TCR-associated CD3ζ rather than a change in the amount of CD3ζ protein associated with the TCR complex. For free CD3ζ, the increased biotin signal appears to be the consequence of an increased amount of protein.

Thirdly, what residue(s) in CD3ζ is biotinylated? The CD3 ζ-chain has an extremely short, nine-amino acid extracellular domain (NH₂-QSFGLLDPK). The most likely candidate for biotinylation is the lysine residue at position 9 (K9). To determine whether enhanced biotinylation at K9 is responsible for the increased CD3ζ biotin signal after TCR ligation, CD3ζ constructs were generated in which the lysine residue was mutated to either arginine (K9R) or serine (K9S). 3A9 CD3ζ-hybridomas were transduced with retrovirus encoding the mutant CD3 ζ-chains. TCR expression and T cell function were comparable to those in T cells expressing wild-type CD3ζ (Fig. 5,A and data not shown). Previous studies have suggested an important role for the K9 residue in the extracellular domain of CD3ζ in signal transduction as well as in stabilizing the association of CD3ζ with the TCR:CD3 complex (31). Thus, we were surprised to find that these mutations had no effect on the function of 3A9. Although the reason for this discrepancy is unclear, it may simply represent differences in the T cell systems used. Transductants were then stimulated, surface-biotinylated, and immunoprecipitated with anti-TCR-Cβ as described. The data clearly show that mutation of K9 had no effect on CD3ζ biotinylation (Fig. 5,B). After detection of biotinylated proteins, the blot was stripped and probed with a mixture of anti-CD3ζ and anti-CD3ε Abs (Fig. 5,C). This blot clearly demonstrates that despite a weak biotin signal, CD3ζ is associated with TCR complexes in unstimulated cells. It also supports our previous finding that the increased biotin signal of TCR-associated CD3ζ is not due to an increased amount of protein. The relative increase in the CD3ζ:CD3ε biotin ratio between unstimulated and stimulated cells was essentially identical (densitometric analysis: CD3ζ-WT, 2.09-fold; CD3ζ-K9R, 2.2-fold; CD3ζ-K9S, 1.8-fold; Fig. 5 D). Thus, it appears that the increase in CD3ζ biotinylation is not mediated via the K9 residue.

Taken together, our data strongly suggest that TCR ligation by MHC:peptide causes alterations in the TCR:CD3 complex, such that the short extracellular domain of CD3ζ becomes more accessible for biotinylation. Furthermore, ligation appears to cause an increase in the amount of free CD3ζ on the cell surface. There are two possible explanations for the latter. First, de novo synthesis of CD3ζ induced by TCR ligation may escape intracellular retention and be expressed on the cell surface. Second, free CD3ζ may result from dissociation between CD3ζ and the rest of the TCR:CD3 complex.

Two members of the Src family of PTKs, p56^lck and p59^fyn, have been shown to play a critical role in T cell development and activation by mediating two of the earliest events following TCR ligation, phosphorylation of CD3 chain immune receptor tyrosine-based activation motif residues and activation of ZAP-70 (32, 33). The question arose of whether the architectural changes observed after TCR ligation were dependent on Src family PTKs. The role of these kinases in the induction of TCR:CD3 architectural changes could provide information regarding the order of events following TCR ligation. 3A9 T cell hybridomas or 3A9 CD3ζ mutant hybridomas stably transfected with FLAG-tagged CD3ζ were pretreated and stimulated in the presence or the absence of PP2, a selective inhibitor of Src family PTKs (34). Cells were surface-biotinylated, and cell lysates were immunoprecipitated twice sequentially with anti-TCR-Cβ, followed by anti-CD3ζ. Immunoprecipitated proteins were resolved by SDS-PAGE, blotted, and probed with streptavidin-AP for detection of biotinylated proteins, as described above. Parallel experiments showed that TCR down-modulation was significantly decreased, and IL-2 production was abrogated after PP2 treatment of 3A9 T cells (data not shown). After Ag stimulation, there was a substantial increase in the TCR-associated CD3ζ signal as well as an increase in free CD3ζ, as described previously (Fig. 6). Pretreatment of the cells with PP2 almost completely abrogated these changes (Fig. 6). Taken together, these data suggest that the structural changes induced in the TCR:CD3 complex after TCR ligation are highly dependent on Src family PTKs and thus presumably occur after or concurrently with kinase activation.

The expression of TCR on the surface of T cells is a dynamic process. In resting T cells, for example, a cycling reservoir of TCR is constitutively internalized and recycled back to the cell surface (5, 13, 35). T cell stimulation also results in internalization of the TCR:CD3 complex. However, this is dependent on ligation of the TCR by Ab or MHC:peptide complexes and, unlike internalization in resting cells, results in a rapid and net reduction of TCR on the cell surface (36, 37). TCR ligation-mediated down-modulation is a consequence of TCR degradation within lysosomal compartments (5, 38, 39). Some studies have proposed that TCR ligation induces the down-modulation of nonengaged receptors (40), although others argue that only specific, ligated receptors are down-modulated after ligation (4). Support for the latter theory comes from a recent study involving mathematical modeling of MHC:peptide-induced TCR down-modulation (41). In it, the researchers propose that an activated TCR remains marked for internalization after dissociating from the MHC:peptide complex. This raises an important question. How does the T cell recognize ligated TCR and subsequently direct these molecules to lysosomes for degradation? Our results suggest that MHC:peptide ligation of the TCR:CD3 complex results in exposure of the CD3ζ NH₂ terminus, which is ordinarily buried in the complex. For discussion purposes, we have generally interpreted this event as reflecting architectural changes in the TCR:CD3 complex. We would also propose, based on increased free CD3ζ levels following stimulation, that such architectural changes may ultimately lead to dissociation of the CD3ζ dimer from the TCR:CD3 complex. However, the origin of the free cell surface CD3ζ has not yet been formally demonstrated, and thus we cannot rule out the possibility that this free CD3ζ derives from de novo synthesis induced by TCR ligation. This would be surprising, because it is generally thought that the free CD3ζ seen after stimulation are products of de novo synthesis and cannot escape intracellular retention (13, 15, 16, 21). Further studies are required to resolve this issue. Regardless, we propose that the architectural changes occurring within the TCR:CD3 complex after TCR ligation may be a mechanism by which T cells recognize ligated TCR and direct them to lysosomal compartments for degradation. Recently, D’Oro and colleagues (42) proposed that the CD3 ζ-chain maintains cell surface receptor expression by sterically blocking internalization sequences in other TCR components, using chimeric CD3 ζ-chains expressing intracellular domains of varying lengths to show that the size of the intracellular tail, rather than the amino acid sequence, was all that was required to support the expression of the fully assembled receptor. We have made similar observations (A. Szymczak, H. Liu, and D. A. A. Vignali, unpublished observations). Other studies with CD16/CD3 chimeras have suggested that CD3ζ may shield the CD3γ dileucine motif (24). As this motif is known to mediate targeting to lysosomes for degradation (22, 23), structural changes in the TCR:CD3 complex, particularly those involving CD3ζ, may result in targeting of ligated TCR:CD3 complexes to lysosomes for degradation, rather than recycling back to the cell surface. It is also possible that other motifs are involved in this process, as T cells lacking the cytoplasmic tail of CD3γ and CD3δ still down-modulate TCR after Ag stimulation (43). Indeed, an endocytosis signal in CD3ε has recently been described (44). Thus, TCR down-modulation may be determined by the exposure of intracellular retention motifs that are normally concealed by CD3ζ.

Until recently, the idea that the TCR:CD3 complex undergoes conformational change after TCR ligation was speculative. However, a study by Gil and colleagues (45) demonstrated that ligation of TCR resulted in the exposure of a proline-rich sequence in CD3ε, which, in turn, leads to recruitment of the adaptor protein, Nck. Interestingly, these events occurred before and independently of tyrosine kinase activation and appeared to be involved in early signaling processes. In our study the observed architectural changes initiated by TCR ligation were heavily dependent on Src family PTK activity, suggesting that the exposure of CD3ζ and its subsequent dissociation from the remainder of the TCR:CD3 complex are relatively late events and may occur downstream of CD3ε exposure and Nck recruitment. It is therefore possible that the architectural changes observed in this study may not be directly involved in the signaling process, but, rather, are required for the downstream effects of TCR ligation, such as TCR down-modulation. Importantly, it has been shown that the expression of constitutively active p56^lck induces TCR down-modulation (39).

An interesting question remains of which residue in the short extracellular domain of CD3ζ is biotinylated upon TCR ligation? Previous studies have suggested an important role for the K9 residue in the extracellular domain of CD3ζ in signal transduction as well as in stabilizing the association of CD3ζ with the TCR:CD3 complex (31). The NHS-ester used to label proteins with biotin preferentially labels lysine residues, but can also label the protein NH₂ terminus. Although other residues, such as arginine, histidine, and cysteine, can be labeled, the frequency and efficiently of this reaction at physiological pH are very low. Thus, the K9 residue appeared to be the most likely candidate. We hypothesized that K9 may, in unligated TCR:CD3 complexes, be involved in interactions with amino acids in other TCR or CD3 chains, preventing its biotinylation. Upon TCR ligation, it is possible that this association is broken, exposing the K9 residue for biotinylation. However, our studies clearly showed that the K9R and K9S mutations had no effect on CD3ζ biotinylation. Although it remains to be determined which residue/moiety in the extracellular domain of CD3ζ is biotinylated, the most likely candidate is the NH₂ terminus, although these are usually capped in eukaryotic cells.

An interesting question raised by our data is whether there are other multichain transmembrane receptors that undergo architectural changes and/or dissociation upon ligand binding. CD3ζ is part of a family of molecules that form an integral part of many different receptors (11). For instance, CD3η (a splice variant of CD3ζ) and the FcRγ can substitute for CD3ζ in the TCR:CD3 complex (46, 47, 48, 49). In addition, CD3ζ and FcRγ form part of the FcγRIII (CD16) Ig FcR expressed on T and NK cells. FcRγ is also incorporated into several other FcRs, such as FcγRI, FcγRII, and FcεRI, on macrophages, monocytes, neutrophils, mast cells, and basophils (49). Lastly, the CD3ζ-like membrane adapter proteins, DAP12 and DAP10 (KAP10), form NK cell-activating receptors with CD94/NKG2C and NKG2D, respectively (50, 51, 52, 53). Thus, it will be interesting to determine whether the ligand-induced alteration and/or dissociation of such transmembrane adapter molecules from multichain receptor complexes are common events. Given the large number of receptors that incorporate CD3ζ-like transmembrane adapter molecules, which are represented in all leukocytes and parts of the CNS (49, 50, 51, 52, 53, 54), our demonstration of ligand-induced architectural changes in the TCR may have broad implications for our understanding of the molecular events that initiate or perpetuate signal transduction via multichain receptor complexes.

We are very grateful to Ralph Kubo for the Abs, Mark Davis for the 3A9 TCR transgenic mice, Elio Vanin and David Baltimore for the 293T cells, Emil Unanue for M12.C3.F6, and Larry Samelson for the CD3ζ cDNA. We also thank Richard Cross and Mahnaz Paktinat for assistance with the flow cytometry, Janet Gatewood for performing the particle-based flow cytometric cytokine assay, Elio Vanin for assistance and reagents for establishing retroviral transduction in our laboratory, and members of the Vignali laboratory for constructive criticisms and comments.