Exocytosis of CTLA-4 Is Dependent on Phospholipase D and ADP Ribosylation Factor-1 and Stimulated during Activation of Regulatory T Cells
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免疫学杂志 2005年第8期
Abstract
CTLA-4 is an essential protein in the regulation of T cell responses that interacts with two ligands found on the surface of APCs (CD80 and CD86). CTLA-4 is itself poorly expressed on the T cell surface and is predominantly localized to intracellular compartments. We have studied the mechanisms involved in the delivery of CTLA-4 to the cell surface using a model Chinese hamster ovary cell system and compared this with activated and regulatory human T cells. We have shown that expression of CTLA-4 at the plasma membrane (PM) is controlled by exocytosis of CTLA-4-containing vesicles and followed by rapid endocytosis. Using selective inhibitors and dominant negative mutants, we have shown that exocytosis of CTLA-4 is dependent on the activity of the GTPase ADP ribosylation factor-1 and on phospholipase D activity. CTLA-4 was identified in a perinuclear compartment overlapping with the cis-Golgi marker GM-130 but did not colocalize strongly with lysosomal markers such as CD63 and lysosome-associated membrane protein. In regulatory T cells, activation of phospholipase D was sufficient to trigger release of CTLA-4 to the PM but did not inhibit endocytosis. Taken together, these data suggest that CTLA-4 may be stored in a specialized compartment in regulatory T cells that can be triggered rapidly for deployment to the PM in a phospholipase D- and ADP ribosylation factor-1-dependent manner.
Introduction
Optimal activation of T cells requires the coordinated engagement of multiple receptors expressed on both T cells and APCs. Of these, binding of the TCR to specific peptide-MHC complexes and costimulation of the T cell through the CD28 receptor are important activating events. CD28 is stimulated by two ligands on the APCs CD80 and CD86, which provide signals that enhance T cell proliferation, cytokine production, and survival (1, 2). However, this system is complicated by the fact that a related receptor, CTLA-4, also interacts with CD80 and CD86 but has inhibitory effects on T cell function and cell cycle progression (3, 4).
The role of CTLA-4 in regulating T cell activation is evident from CTLA-4-deficient mice. These mice exhibit severe lymphoproliferative disease and die 3–4 wk after birth, indicating that CTLA-4 is critical for maintaining tolerance to self-tissues (5, 6, 7). There are several possibilities for the mechanism of CTLA-4 function. These include direct inactivating signals to T cells, competition with CD28 for binding to ligands on APCs, enhancement of suppression by specialized regulatory T (Treg)3 cells, and stimulation of tryptophan catabolism in APCs (1, 2). Recently, genetic studies have suggested that soluble CTLA-4 or even forms of CTLA-4, which cannot bind ligands, may be involved in disease susceptibility (8, 9). Thus, as yet, no single model of CTLA-4 function has emerged, and it is possible that more than one mechanism exists.
One major difference between CD28 and CTLA-4 lies in their expression patterns in T cells. CD28 is expressed constitutively on the surface of both resting and activated T cells. In contrast, CTLA-4 expression is undetectable in resting T cells. In activated T cells, despite equivalent mRNA levels to CD28, surface expression of CTLA-4 is much lower. This is because the majority of CTLA-4 molecules appears to be intracellular as a result of rapid endocytosis from the cell surface (10, 11). Internalization is thought to be controlled by a tyrosine-based motif, YxxM, in the cytoplasmic domain of CTLA-4 that interacts with the clathrin adaptor complex AP-2 (12, 13). Furthermore, it has been suggested that phosphorylation of tyrosine (Y)201 in this motif by Src kinases disrupts interaction with AP-2, resulting in stabilization of CTLA-4 at the cell surface (12). Interestingly, CD28 also contains a YxxM motif in its cytoplasmic tail yet is not endocytosed, suggesting that regulation of CTLA-4 expression is likely to be complex. Indeed, the observation that the entire cytoplasmic domain of CTLA-4 is absolutely conserved across a number species suggests that much of the sequence may be critical for its correct localization, membrane expression, and function. Interestingly, a specialized subset of CD4+ T cells (Treg) retain a constitutive intracellular pool of CTLA-4, suggesting that there may be a storage compartment from which CTLA-4 can undergo regulated exocytosis, possibly in a similar manner to proteins such as GLUT4 (14).
Despite numerous studies on CTLA-4-AP-2 interaction, little is known regarding the intracellular trafficking pathways used by CTLA-4 to gain access to the plasma membrane (PM). Typically, such transport of molecules through the cell is controlled by recruitment of proteins into coated vesicles that are transported from the trans-Golgi network toward the PM (15). Clathrin adaptor complexes, such as AP-1 and AP-3, are thought to be required for these vesicles and target them to the PM or to lysosomes, respectively (16, 17). Recently, the AP-3 adaptor complex has been shown to be required for the microtubule-mediated movement of lytic granules in CTLs (18). Consistent with its location in intracellular vesicles, CTLA-4 is also thought to interact with AP-1 via its cytoplasmic YVKM motif (19). As with CD28 (20), CTLA-4 has also been reported to interact with PI3K via this motif; however, the significance of this interaction is unclear at present (21).
The budding of clathrin-coated vesicles depends on a number of additional components, including the small ADP ribosylation factor (ARF) GTPases and the enzyme phospholipase D (PLD). ARF-1 is the most studied member of the ARF family and has been found to be required for assembly of coated vesicles at a variety of transport steps (22, 23). PLD is widely distributed and catalyzes the conversion of the membrane phospholipid, phosphatidylcholine, into phosphatidic acid and choline. This leads to high levels of phosphatidic acid within local membranes, resulting in increased binding of coat proteins and subsequent vesicle formation. Accordingly, PLD had been implicated at several stages of vesicle trafficking (24, 25).
Given the critical functions of CTLA-4 within the immune system and its unusual pattern of cellular expression, we have sought to characterize the controls that regulate its PM expression. We have established a model system of CTLA-4 trafficking in Chinese hamster ovary (CHO) cells and compared this with normal human T cells. Using this model, we have examined the requirement for PLD-1, PLD-2, and ARF-1 in exocytosis of CTLA-4 using selective inhibitors and dominant negative mutants. Our data demonstrate that PLD-1, PLD-2, and ARF-1 are required for transport of CTLA-4 proteins from the Golgi apparatus to the PM and that inhibition of these pathways prevents cell surface expression of CTLA-4. Furthermore, using stimuli that result in T cell activation, we observed a substantial increase in exocytosis of CTLA-4 that was prevented by inhibitors of both ARF and PLD. Despite this increase in traffic to the PM, only small increases of steady-state levels of surface CTLA-4 were observed, indicating that increased exocytic traffic was also accompanied by continued endocytosis. Our data identify several key points in trafficking pathways that regulate surface expression of CTLA-4 and indicate that regulated exocytosis is a major mechanism for controlling cell surface expression.
Materials and Methods
CTLA-4-transfected cells
CTLA-4-transfected CHO (CHO-CTLA-4) cells were generated by electroporation using the full-length and mutated forms of human CTLA-4 cDNA cloned into a CMV expression vector pCDNA-3. Cells were grown in DMEM as described previously (26). Cells expressing the plasmid were selected using G418 (500 μg/ml) treatment and sorted by FACS. Cultures were maintained at 37°C in a humid atmosphere containing 5% CO2 and were trypsinized and passaged every other day at 75% confluence.
Stable cultures of CHO-CTLA-4 cells were retransfected with wild-type or dominant negative GFP-tagged PLD and ARF plasmids (27). These cells (2 x 106) were transfected transiently by electroporation using an Amaxa nucleofector device, according to the manufacturer’s instructions, for CHO cell transfections and analyzed at 24 h.
Where human embryonic kidney (HEK) cells (HEK-293) were used for colocalization studies, the cells were cultured and transfected as for CHO cells with the exception that the Amaxa conditions for HEK cells were followed.
T cells
Buffy coats from blood donated by healthy donors were obtained from the National Blood Service. PBMCs were isolated using Ficoll density gradient centrifugation at 800 x g for 30 min. The buoyant layer was removed and washed twice in RPMI 1640 medium (supplemented with 10% FCS, 100 U/ml penicillin, and 100 μg/ml streptomycin). CD4+ T cells were purified by negative selection using magnetic beads, and CD4+CD25+ regulatory cells were purified subsequently by positive selection using CD25-immunomagnetic beads, according to the manufacturer’s instructions (Miltenyi Biotec).
Inhibitors
Brefeldin A was used at a concentration of 1 μg/ml. Cells were incubated at 37°C in a humid atmosphere containing 5% CO2 for 3 h before Ab staining. Butan-1-ol and butan-2-ol were used at a concentration of 1.5% in transfected CHO cells and 1.0% in T cell blasts. Cells were incubated for 1 h before Ab staining. Cells treated with cycloheximide were incubated for the times shown with 10 μg/ml in medium.
Flow cytometry
A total of 2 x 105 cells suspended in 100 μl of medium was treated with butan-1-ol, butan-2-ol, or brefeldin A as required and incubated with PE-conjugated anti-CTLA-4 (BN13; BD Pharmingen). Surface expression was detected by incubating cells at 4°C for 1 h, and recycled CTLA-4 was detected by incubation at 37°C. Total CTLA-4 expression was determined by fixing cells in 2% paraformaldehyde for 5 min at 4°C. Cells were washed once in PBS followed by anti-CTLA-4 Ab in PBS containing 0.1% saponin. Cells were washed in PBS/saponin and analyzed. Control samples were stained using the same protocol using a PE-labeled isotype control Ab. Where indicated, surface bound Abs were removed by acid washing for 2 min in PBS adjusted to pH 2.0. Percentage internalization was calculated by mean fluorescence intensity acid wash/mean fluorescence intensity PBS wash x 100. Analysis was conducted using a FACScan flow cytometer (BD Biosciences), and data for 10,000 cells were collected.
Confocal microscopy
A total of 5 x 104 cells was grown on multispot slides overnight in 20-μl drops. Cells were fixed either in ice-cold methanol for 5 min or in 2% formaldehyde for 5 min, followed by 0.1% saponin. Cells were incubated with 10% FCS in PBS before Ab addition to block nonspecific binding. Anti-GM130 was obtained from BD Biosciences and used at 1:100. Mouse anti-human CD63 and rabbit anti-human lysosome-associated membrane protein (LAMP) were generous gifts from Dr. G. Griffiths (University of Oxford, Oxford, U.K.). Mouse anti-human CD71 (transferrin receptor) was obtained from D. Hardie (University of Birmingham, Birmingham, U.K.). Anti-EEA-1 was a gift from B. Reaves (University of Bath, Bath, U.K.). Primary Abs were visualized using anti-mouse or anti-rabbit Abs conjugated to Alexa Fluor 488 or 594 (Molecular Probes). Anti-CTLA-4 Ab (11G1; a gift from Dr. J. Allison, University of California, Berkeley, CA) was directly conjugated to Alexa Fluor 594 (Molecular Probes) in our laboratory and used at 1/100 dilution. Staining for CTLA-4 was conducted at 4°C, 37°C, or after fixing and permeabilization as indicated. 4',6'-Diamidino-2-phenylindole (DAPI) was added to fixed cells at a concentration of 10 μg/ml for 1 min at room temperature to stain nuclei. Cells were washed twice and mounted onto slides. Fluorescence was examined using a Zeiss Axiovert 100M confocal microscope and a Zeiss LSM 510 scan module. All images were obtained with a C-Apochromat x63 water lens. For each image, optical sections were obtained at intervals of 0.5 μm through the cell, and the images shown are representative of sections through the centre of the cell.
Results
CTLA-4 trafficking is conserved in CHO cells and activated T cells
To facilitate the study of CTLA-4 intracellular trafficking and surface expression, we used transfected CHO cells constitutively expressing CTLA-4 as a model system. The expression pattern of CTLA-4 was compared with activated peripheral blood T cells using three different staining protocols. Staining at 4°C measured CTLA-4 expressed at the cell surface only. In contrast, staining at 37°C labeled all molecules arriving at the cell surface during the incubation period but did not distinguish whether they remained at the cell surface or were subsequently endocytosed. Therefore, labeling at 37°C gives an accurate indication of the total amount of CTLA-4 that reaches the PM over a given period of time, although this will include some newly synthesized protein. Finally, the total pool of CTLA-4 protein was measured in cells that were fixed and permeabilized before Ab staining. As shown in Fig. 1, this analysis revealed strong similarities between CTLA-4 expressed in CHO cells and in activated T cells. In both cases, much lower levels of cell surface CTLA-4 expression were observed in cells stained at 4°C when compared with cells stained at 37°C (Fig. 1a). Furthermore, cells that were fixed and permeabilized revealed similar levels of expression to cells stained at 37°C. Taken together, these data suggested that CTLA-4 was continually transported to the cell surface in both CHO and activated T cells but that steady-state levels of surface CTLA-4 were low due to continual endocytosis.
FIGURE 1. The pattern of CTLA-4 expression is similar in T cell blasts and CHO-CTLA-4 cells. a, CHO-CTLA-4 cells and T cells (unstimulated and stimulated with PMA and ionomycin) were stained for cell surface CTLA-4 (4°C staining), endocytosed CTLA-4 (37°C staining), or total CTLA-4 (F + P) using anti-CTLA-4-PE. A total of 5000 cells was analyzed by flow cytometry. b, FACS analysis of CHO-CTLA-4 cells in the presence of cycloheximide (CHX). Transfectants were stained 4°C, 37°C, and in fixed cells (total) after CHX treatment for the times shown. c, Confocal analysis of CTLA-4 in T cells and transfectants determined at 37°C and after fixing and permeabilizing (F + P) using anti-CTLA-4 conjugated to Alexa 594.
To ensure that the increased levels obtained at 37°C were not simply a reflection of newly synthesized CTLA-4, these experiments were repeated using transfectants in the presence of cycloheximide to inhibit new protein synthesis. This data (Fig. 1b) revealed that although the total level of CTLA-4 expression diminished over time with cycloheximide, the difference between staining at 4°C and 37°C remained, clearly demonstrating that new protein synthesis does not account for the difference between 4°C and 37°C, which supports the conclusion that intracellular CTLA-4 is continually transported to the PM at 37°C.
Finally, we analyzed both T cells and CHO cells by confocal microscopy (Fig. 1c). Staining at 37°C revealed that in both CHO and T cells CTLA-4 expression was located predominantly in intracellular vesicles; again, consistent with the concept that CTLA-4 was being exported to the PM and then rapidly endocytosed, contributing to the low steady-state level seen at the cell surface. Likewise, in fixed and permeabilized cells, a perinuclear compartment was observed containing high levels of CTLA-4 molecules representing either newly synthesized CTLA-4 or protein that had been endocytosed and targeted back to this compartment. Taken together, these observations suggested that CHO cells and activated T cells had very similar patterns of CTLA-4 expression.
Tyrosine 201 is not essential for CTLA-4 endocytosis
Endocytosis of CTLA-4 has been suggested to require a cytoplasmic 201YVKM motif for interaction with the clathrin adaptor AP-2. Therefore, we investigated the requirement for this motif in regulating CTLA-4 expression. Although mutation 201YVKM to 201FVKM appears to result in higher steady-state levels of CTLA-4 at the cell surface compared with wild type, analysis by confocal microscopy still revealed the presence of endocytosed CTLA-4 in intracellular vesicles. This appeared similar to wild-type CTLA-4 but was in clear contrast to a chimeric construct expressing CTLA-4 with a CD28 cytoplasmic domain, which was stably expressed at the cell surface (Fig. 2a). This suggested that endocytosis was not abolished in the 201FVKM mutant. Furthermore, the intracellular vesicles observed were not an artifact of nonspecific endocytosis or macropinocytosis of the CTLA-4 Ab because this did not occur with a hybrid molecule containing CTLA-extracellular and transmembrane domains but with a CD28-cytoplasmic domain. We also determined the kinetics of endocytosis by flow cytometry using an acid wash to remove surface CTLA-4 staining (Fig. 2b). This revealed that although endocytosis of CTLA-4 was slower (presumably leading to higher levels of surface CTLA-4 at steady state), by 20 min all of the surface-labeled CTLA-4 201FVKM was internalized. Thus, our data revealed that this motif is not essential for endocytosis of CTLA-4 in a cellular context. This information combined with the finding that activated human T cells only express weak cell surface CTLA-4 despite substantial CTLA-4 traffic to the PM at 37°C (see Fig. 1a) strongly suggests that during T cell activation changes in CTLA-4 expression might be regulated through exocytosis and not only by inhibiting endocytosis.
FIGURE 2. Mutation of tyrosine 201 does not prevent clathrin-mediated endocytosis. a, Expression patterns of wild-type (WT) and Y201F and CTLA-4-CD28 molecules in CHO cells were determined by confocal microscopy in fixed cells. Cells were stained using anti-CTLA-4-Alexa 594. b, Rates of CTLA-4 endocytosis were determined for WT and Y201F CTLA-4 and for CD28 in CHO cells. Cells were labeled at 4°C with anti-CTLA-4-PE, washed, and raised to 37°C for the time shown. Nonendocytosed Ab was removed by acid washing, and the levels of internalized (acid protected) CTLA-4 were measured by flow cytometry. c, The effect of sucrose was compared on internalization of anti-CTLA-4-PE after 30 min. Cells were labeled and washed as above either in the presence or absence of 0.4 M sucrose. Cells were washed in acid buffer and analyzed by FACS.
Because endocytosis of CTLA-4 has been reported previously to occur via an AP-2-mediated, clathrin-dependent pathway (12), we also treated CHO-CTLA-4 with hypertonic sucrose, which is known to inhibit the formation of functional clathrin-coated vesicles (28). As shown in Fig. 2c, cells treated in this way were unable to endocytose surface CTLA-4 molecules, resulting in accumulation of CTLA-4 at the PM. In contrast, in the absence of sucrose, both wild-type and CTLA-4 201FVKM transfectants internalized CTLA-4 but not the chimeric CTLA-4-CD28 molecule. Overall, these data demonstrated that CHO-CTLA-4 provided a model of CTLA-4 trafficking that required information resident in the CTLA-4-cytoplasmic domain and proceeded via clathrin-mediated endocytosis. Nonetheless, the YVKM-AP-2-binding motif was not essential to this process.
Inhibition of PLD-1 and ARF-1 blocks CTLA-4 exocytosis
We next initiated studies on the role of exocytosis in control of CTLA-4 expression in CHO-CTLA-4 cells. Initially, CHO-CTLA-4 cells were treated with inhibitors of PLD signaling (butan-1-ol) to determine the involvement of this pathway in trafficking of CTLA-4 containing vesicles (Fig. 3a). Strikingly, this treatment resulted in a very substantial reduction of staining at both 4°C and 37°C. However, CTLA-4 was still expressed intracellularly, as revealed in permeabilized cells. Furthermore, Butan-1-ol had no significant effect on the expression of CD28, which is not thought to interact with either AP-1 or AP-2. In control experiments, butan-2-ol, which does not inhibit PLD signaling, had no effect on CTLA-4 expression. A similar picture was observed using brefeldin A, which inhibits ARF proteins (Fig. 3b). Taken together, these results demonstrated that disruption of either ARF or PLD function was sufficient to prevent trafficking of CTLA-4 to the cell surface but did not affect CD28 expression.
FIGURE 3. PLD and ARF proteins are required for transport of CTLA-4 to the PM. a, CTLA-4- or CD28-transfected CHO cells were treated with 1.5% butan-1-ol, butan-2-ol, or 1 μg/ml brefeldin A (BFA) in DMEM for 3 h at 37°C. Cells were then stained for CTLA-4 or CD28 expression for 1 h at either 4°C, 37°C, or for total CTLA-4 expression after fixing and permeabilization (F + P). Each histogram shows the isotype control staining (shaded histogram) or specific Ab with (bold line) or without (thin line) butanol. b, The effect of BFA on CTLA-4 and CD28 expression was compared at 37°C. In all panels, 10,000 events were collected and analyzed by flow cytometry.
To confirm that the results obtained from inhibitors were not due to nonspecific or toxic effects and to investigate which specific ARF and PLD proteins were involved, a genetic approach was also adopted. CTLA-4-CHO cells were transfected with plasmids expressing either GFP-tagged, wild-type or dominant negative PLD-1 and ARF-1 proteins and analyzed by flow cytometry (Fig. 4a). This showed that dominant negative mutants of both PLD-1 and ARF-1 inhibited expression of cell surface CTLA-4 proteins detected at 4°C or at 37°C. In contrast, although GFP fluorescence was reduced by the permeabilization protocol, the total levels of CTLA-4 expression were unaffected by dominant negative ARF-1 or PLD-1. These effects were most obvious at 37°C where wild-type PLD and ARF did not inhibit CTLA-4 staining (cells are seen in Fig. 4a, upper right quadrant), whereas with the dominant negative GFP, proteins prevented CTLA-4 staining (cells are seen in Fig. 4a, lower right quadrant). Data for dominant negative PLD-2 was indistinguishable from PLD-1 (data not shown). Thus, the expression of both dominant negative PLD-1, PLD-2, or ARF-1 proteins prevented CTLA-4 from reaching the PM. When using dominant negative constructs, we did observe a slight drop in CTLA-4 staining cells in apparently untransfected cells. However, these transfections were also associated with slightly lower overall levels of CTLA-4 staining, suggesting this may be a result of decreased staining in these cells. However, it is possible that transfections with the dominant negative plasmids had a slight nonspecific inhibitory effect. Nonetheless, the data in the GFP-positive populations were unequivocal, and we concluded that ARF-1 and PLD-1 were required for effective transport of CTLA-4 to the cell surface. Consistent with these observations, confocal microscopy on cells transfected with dominant negative and wild-type ARF-1 revealed a lack of staining of CTLA-4 at both 4°C and 37°C with dominant negative ARF-1 (Fig. 4b). However, once again, total staining was unaffected. Taken together, these data indicate that CTLA-4 transport to the PM is dependent on both ARF and PLD pathways.
FIGURE 4. Surface expression of CTLA-4 is blocked by dominant negative (DN) PLD-1 and ARF-1. a, Stable lines of CHO-CTLA-4 cells were transfected transiently with GFP-tagged wild-type (WT) or DN PLD-1 and ARF-1. Expression of GFP proteins was detected along the x-axis and compared with CTLA-4 expression (BN13-PE) on the y-axis using flow cytometry. CTLA-4 staining was determined at 4°C and 37°C and in permeabilized cells to determine effects on CTLA-4 expression. b, Confocal analysis of cells transfected with DN-ARF-1 as in a, with the exception that CTLA-4 was detected using Alexa 594-conjugated Ab, and nuclei were counterstained using DAPI (white). Arrowheads highlight the lack of CTLA-4 staining in DN-ARF-transfected cells.
CTLA-4 is localized to a distinct intracellular compartment
To characterize the intracellular location of CTLA-4, we performed colocalization experiments. Initially, these were performed in CHO cells; however, the lack of suitable colocalization reagents for these cells prompted us to transfect CTLA-4 into HEK-293. This allowed us to study the localization of several endogenous proteins, including CD63 and transferrin receptor, both of which are known to be internalized by clathrin-mediated endocytosis. Importantly, the staining pattern of CTLA-4 in both CHO and HEK systems was indistinguishable by confocal microscopy and retained the same characteristics at 37°C and 4°C by FACS. In fixed HEK cells, CTLA-4 colocalized significantly in a perinuclear location with GM130, which represented Golgi staining. However, despite reports that CTLA-4 may be stored in secretory lysosomes (29), the amount of CTLA-4 that colocalized with the lysosomal markers LAMP-2 and CD63 was very limited (Fig. 5). We also performed staining for CD71 and for the early endosome marker EEA-1. However, once again, while there was some colocalization with these compartments, in particular with EEA-1, this was clearly incomplete. In control experiments, CD63 and LAMP demonstrated complete colocalization. These results indicated that CTLA-4 is found in a both a perinuclear compartment (that most likely contains both pre- and postendocytic vesicles), as well as distinct cytoplasmic vesicles; some of which are EEA-1 positive. Surprisingly, CTLA-4-containing vesicles did not appear to be strongly CD63 or CD71 associated, despite the fact that both proteins contain an AP-2-sorting motif, indicating that CTLA-4 has a distinct trafficking itinerary.
FIGURE 5. Colocalization of CTLA-4. CTLA-4-transfected HEK-293 cells were fixed and permeabilized and stained with anti-CTLA-4 Alexa 594 and analyzed by confocal microscopy. Cells were costained (green) for LAMP-1, GM130, EEA-1, or CD71 as shown. Control colocalization for CD71 and CD63 with LAMP is shown. Sections shown are representative staining of an optical section through the middle of the cell. Nuclei were counterstained with DAPI (blue). Colocalization is represented as yellow.
Up-regulation of CTLA-4 on T cells results from increased exocytosis and is dependent on ARF and PLD activity
Because data in CHO cells indicated a critical role for ARF-1 and PLD in the efficient export of CTLA-4 to the PM, we examined this process in T cells. Purified human T cells were stimulated using PMA and ionomycin for 4 h to induce CTLA-4 expression and the effect of butanol treatment studied as before (Fig. 6a). As observed with CHO cells. treatment with butan-1-ol abolished PM traffic with only minor effects on the total level of CTLA-4. Similar effects were also observed with brefeldin A (data not shown), indicating that data derived from the CHO-CTLA-4 was consistent with human T cells.
FIGURE 6. PLD and ARF are required for stimulated exocytosis of CTLA-4 in CD4+ and CD4+CD25+ human T cells. a, Purified resting CD4+ T cells were stimulated for 4 h with PMA (5 ng/ml) and ionomycin (500 μM) to induce CTLA-4 expression in the presence (bold line) or absence (thin line) of butanol. CTLA-4 expression was detected at 4°C and 37°C and in permeabilized cells by flow cytometry using anti-CTLA-4-PE. b, Treg cells were stimulated for 2 h with PMA (5 ng/ml) and stained at 37°C (dashed line) in the presence of butan-1-ol (bold line) and butan-2-ol (thin line). Control Treg cells stained for anti-CTLA-4 but not stimulated are shown (shaded histogram). Brefeldin A (BFA) treatment is shown as a bold line.
We also studied this process using CD4+CD25+ Treg cells, which constitutively express intracellular CTLA-4. Because PMA is a well-established activator of PLD, we predicted that PMA alone should increase CTLA-4 membrane traffic in the absence of full T cell activation. This experiment (Fig. 6b) revealed that unstimulated Treg did not constitutively recycle CTLA-4 to the PM as detected by labeling at 37°C. However, stimulation with PMA resulted in a rapid and marked up-regulation of CTLA-4 within 2 h that was abolished by inhibiting PLD or ARF proteins. These data directly demonstrate that up-regulation of CTLA-4 in human T cells is driven by regulated exocytosis in a PLD-dependent manner. Furthermore, these data indicate that in Treg CTLA-4 is stored in a compartment that is sensitive to such regulated exocytosis.
Discussion
CTLA-4 is an essential protein for immune regulation, the absence of which leads to fatal autoimmune tissue destruction. A major feature of this protein is its unusual pattern of intracellular expression and its highly conserved cytoplasmic domain, which is as yet without a clearly established function. To better understand how CTLA-4 expression is controlled, we developed a model of CTLA-4 trafficking in CHO cells that is amenable to confocal analysis and genetic manipulation. We have validated this model against human T cells and found no obvious differences in the patterns of CTLA-4 expression, with the exception that trafficking of CTLA-4 to the PM was constitutive in CHO cells and whereas it was stimulation dependent in T cells. However, in both cases, our data establish that trafficking is dependent on ARF-1 and PLD activity.
The data presented suggest a critical role for PLD and ARF-1 in movement of CTLA-4-containing vesicles from a perinuclear region to the PM. This compartment was observed during labeling at both 37°C and in fixed cells, suggesting it contains both postendocytic vesicles as well as newly budding vesicles. Interestingly, clathrin-coated vesicles that use the adapter protein AP-1 are generally associated with the trans-Golgi network, and consistent with this finding, yeast two-hybrid studies have shown that CTLA-4 can indeed interact with AP-1 via its YVKM motif (19). In support of this, we observed colocalization with GM-130, which is a marker of the cis-Golgi.
Several studies have suggested that CTLA-4 is located in lysosomes or secretory granules and may be translocated via secretory lysosomes (11, 29, 30). In our model, we only observed limited colocalization with lysosomal markers LAMP-1 and CD63, suggesting that the majority of CTLA-4 is not within lysosomes. Furthermore, despite it being relatively clear that in CD8+ T cells and other specialized secretory cells—and secretory lysosomes represent a significant mechanism of exocytosis (31)—it is not clear whether this is a major mechanism for CTLA-4 in CD4+ cells and, in particular, Treg cells. In our hands, the levels of surface CTLA-4 seen following ionomycin (a stimulus for lysosome secretion) are substantially less than that seen with PMA, which may stimulate generalized vesicle traffic via PLD. We believe this is more consistent with a nonlysosomal store as the major source of membrane-translocated CTLA-4. Furthermore, given that lysosomes are clearly capable of degrading CTLA-4 rapidly (32), this seems unlikely to be the location of long-term CTLA-4 storage for cells such as Treg. Thus, although CTLA-4 can clearly be detected in lysosomes, additional studies are needed to clarify the role of this compartment in stimulated exocytosis.
The pattern of CTLA-4 trafficking observed in our studies appears similar to that of the glucose transporter GLUT4 (14). GLUT4 is a recycling receptor that interacts with both AP-1 and AP-2 and recycles to a trans-Golgi network 38-negative compartment (33). Furthermore, PLD activity has also been implicated in translocation of GLUT4 to the PM (34). Perhaps most interestingly, both CTLA-4 and GLUT4 appear to use a storage compartment from which exocytosis is achieved rapidly upon stimulation. In the case of GLUT4, this stimulation appears to be via a PI3K-dependent mechanism. Although the pathway responsible for stimulating for CTLA-4 exocytosis in T cells remains to be elucidated, our data show that activation of PLD using phorbol ester is sufficient for translocation. Interestingly, the major known physiological ligands that drive T cell activation (TCR and CD28) are known to activate PI3K (20, 35) and PLD (36), suggesting this mechanism may be applicable during normal T cell stimulation. However, although there is a report that wortmannin can up-regulate CTLA-4 expression (37), we have observed no consistent effects of PI3K in our studies, which leaves the role of PI3K in CTLA-4 trafficking needing additional investigation.
The data presented here suggest that despite delivery to the cell surface CTLA-4 is continually endocytosed. This seems to conflict somewhat with previous suggestions that expression of CTLA-4 at the cell surface is stabilized by phosphorylation of its cytoplasmic domain, thereby disrupting AP-2-mediated endocytosis. However, direct data measuring endocytosis of CTLA-4 are somewhat limited. Shiratori et al. (12) showed clearly that CTLA-4 interacts via its cytoplasmic domain with AP-2 using the YVKM motif. However, data directly showing that activation of T cells caused significant phosphorylation of CTLA-4, resulting in stable cell surface CTLA-4, are missing. Indeed, increased CTLA-4 at the cell surface is generally only seen using pervanadate as a phosphatase inhibitor and not under normal conditions of T cell activation (12, 38). Interestingly, our own experiments with pervanadate in T cells (K. Mead, unpublished observations) reveal that CTLA-4 expression is markedly more enhanced at 37°C compared with 4°C, which raises the possibility that this increase is due to enhanced delivery rather than decreased endocytosis. Therefore, it may be significant that pervanadate can act by stimulating PLD activation and thereby possibly enhance exocytosis (39). In other studies, kinases such as lck and Fyn have been found to significantly phosphorylate CTLA-4 (40), yet without the use of pervanadate, this does not affect levels of CTLA-4 at the cell surface, suggesting that simple phosphorylation of CTLA-4 does not block endocytosis (41).
Thus, one possibility consistent with our experience and data is that CTLA-4 endocytosis continues even under normal conditions of T cell activation and that mutation of Y201 does not inhibit endocytosis. Interestingly, others have made similar mutants and also observed predominantly intracellular CTLA-4 (29), suggesting this is not an artifact of expression in CHO cells. Furthermore, while it is clear that CTLA-4 may be phosphorylated on tyrosine 201, even in response to TCR stimulation (42), there is no direct evidence that this prevents internalization. Although increased surface staining is attributed to disrupted endocytosis in many studies, it is notable that endocytosis is not actually measured. Thus, while our results contrast with the current perception that disruption of endocytosis results in increased surface expression of CTLA-4, we believe it is more likely that regulated exocytosis of CTLA-4 is in fact the critical regulatory step.
Although we have yet to define how PLD is involved in CTLA-4 translocation, the inhibition by both the catalytically inactive form of PLD and butan-1-ol implicates the generation of phosphatidic acid in the regulatory process. We have previously observed in RBL-2H3 cells that trafficking of secretory lysosomes is not prevented by inhibition of PLD activity but that fusion of the vesicles with the PM was ablated (23). Thus, one possible role for the generated phosphatidic acid may be in controlling the fusion of the vesicles with the PM.
At the present time, the mechanism of CTLA-4 inhibitory action is not understood, and several modes of action are possible. Given the exceptional degree of conservation of the CTLA-4-cytoplasmic domain, the importance of regulated trafficking of CTLA-4 cannot be underestimated. CTLA-4 is thought to promote T cell anergy (unresponsiveness) (43). It is interesting to note that differential expression studies of anergic T cells have identified ARF-6, as well as the exchange factor GRP-1, as differentially expressed under anergic conditions (44, 45). Furthermore, anergy induction has recently been associated with up-regulation of proteins such as Grail, which has strikingly similar expression patterns to CTLA-4 (46). Taken together, this may well suggest that regulation of vesicle trafficking may be a target in T cell anergy. The present studies now provide the basis for more detailed analysis of the control of CTLA-4 trafficking.
Disclosures
The authors have no financial conflict of interest.
Footnotes
The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked advertisement in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
1 This work was supported by the Arthritis Research Campaign (ARC) (to D.M.S., C.N.M. and M.K.P.L.), the Biotechnology and Biological Sciences Research Council (to Y.Z.), and the Wellcome Trust (to M.J.O.W. and D.J.P.). D.M.S. is an ARC Senior Research Fellow. K.I.M. is a Medical Research Council PhD student.
2 Address correspondence and reprint requests to Dr. David M. Sansom, Medical Research Council Centre for Immune Regulation, University of Birmingham Medical School, Vincent Drive, Birmingham B15 2 TT, U.K. E-mail address: d.m.sansom{at}bham.ac.uk
3 Abbreviations used in this paper: Treg, regulatory T; PM, plasma membrane; ARF, ADP ribosylation factor; PLD, phospholipase D; CHO, Chinese hamster ovary; CHO-CTLA-4, CTLA-4 transfected CHO; HEK, human embryonic kidney; DAPI, 4',6'-diamidino-2-phenylindole; LAMP, lysosome-associated membrane protein.
Received for publication January 28, 2004. Accepted for publication February 4, 2005.
References
Sansom, D. M., C. N. Manzotti, Y. Zheng. 2003. What’s the difference between CD80 and CD86?. Trends Immunol. 24:313.
Sansom, D. M.. 2000. CD28, CTLA-4 and their ligands: who does what and to whom?. Immunology 101:169.
Walunas, T. L., C. Y. Bakker, J. A. Bluestone. 1996. CTLA-4 ligation blocks CD28-dependent T cell activation. J. Exp. Med. 183:2541.
Krummel, M. F., J. P. Allison. 1995. CD28 and CTLA-4 have opposing effects on the response of T cells to stimulation. J. Exp. Med. 182:459
Waterhouse, P., J. M. Penninger, E. Timms, A. Wakeham, A. Shahinian, K. P. Lee, C. B. Thompson, H. Griesser, T. W. Mak. 1995. Lymphoproliferative disorders with early lethality in mice deficient in CTLA-4. Science 270:985
Tivol, E. A., F. Borriello, A. N. Schweitzer, W. P. Lynch, J. A. Bluestone, A. H. Sharpe. 1995. Loss of CTLA-4 leads to massive lymphoproliferation and fatal multiorgan tissue destruction, revealing a critical negative regulatory role of CTLA-4. Immunity 3:541.
Chambers, C. A., T. J. Sullivan, J. P. Allison. 1997. Lymphoproliferation in CTLA-4-deficient mice is mediated by costimulation-dependent activation of CD4+ cells. Immunity 7:885.
Ueda, H., J. M. Howson, L. Esposito, J. Heward, H. Snook, G. Chamberlain, D. B. Rainbow, K. M. Hunter, A. N. Smith, G. Di Genova, et al 2003. Association of the T-cell regulatory gene CTLA4 with susceptibility to autoimmune disease. Nature 423:506.
Vijayakrishnan, L., J. M. Slavik, Z. Illes, R. J. Greenwald, D. Rainbow, B. Greve, L. B. Peterson, D. A. Hafler, G. J. Freeman, A. H. Sharpe, et al 2004. An autoimmune disease-associated CTLA-4 splice variant lacking the B7 binding domain signals negatively in T cells. Immunity 20:563.
Alegre, M.-L., P. J. Noel, B. J. Eisfelder, E. Chuang, M. R. Clark, S. L. Reiner, C. B. Thompson. 1996. Regulation of surface and intracellular expression of CTLA-4 on mouse T cells. J. Immunol. 157:4762.
Linsley, P. S., J. Bradshaw, J. Greene, R. Peach, K. L. Bennett, R. S. Mittler. 1996. Intracellular trafficking of CTLA-4 and focal localisation towards sites of TCR engagement. Immunity 4:535.
Shiratori, T., S. Miyatake, H. Ohno, C. Nakaseko, K. Isono, J. S. Bonifacino, T. Saito. 1997. Tyrosine phosphorylation controls internalization of CTLA-4 by regulating its interaction with clathrin-associated adaptor complex AP-2. Immunity 6:583.
Zhang, Y., J. P. Allison. 1997. Interaction of CTLA-4 with AP-50, a clathrin-coated pit adaptor protein. Proc. Natl. Acad. Sci. USA 94:9273.
Bryant, N. J., R. Govers, D. E. James. 2002. Regulated transport of the glucose transporter GLUT4. Nat. Rev. Mol. Cell Biol. 3:267.
Beraud-Dufour, S., W. Balch. 2002. A journey through the exocytic pathway. J. Cell Sci. 115:1779
Dell’Angelica, E. C.. 2001. Clathrin-binding proteins: got a motif: join the network!. Trends Cell Biol. 11:315.
Dell‘Angelica, E. C., V. Shotelersuk, R. C. Aguilar, W. A. Gahl, J. S. Bonifacino. 1999. Altered trafficking of lysosomal proteins in Hermansky-Pudlak syndrome due to mutations in the 3A subunit of the AP-3 adaptor. Mol. Cell 3:11.
Clark, R. H., J. C. Stinchcombe, A. Day, E. Blott, S. Booth, G. Bossi, T. Hamblin, E. G. Davies, G. M. Griffiths. 2003. Adaptor protein 3-dependent microtubule-mediated movement of lytic granules to the immunological synapse. Nat. Immunol. 4:1111.
Schneider, H., M. Martin, F. A. Agarraberes, L. Yin, I. Rapoport, T. Kirchhausen, C. E. Rudd. 1999. Cytolytic T lymphocyte-associated antigen-4 and the TCR /CD3 complex, but not CD28, interact with clathrin adaptor complexes AP-1 and AP-2. J. Immunol. 163:1868.
Ward, S., J. Westwick, N. Hall, D. Sansom. 1993. CD28 ligation elevates PtdIns(3,4)P2 and PtdIns(3,4,5)P3 in T cells. Eur. J. Immunol. 23:2572.
Schneider, H., V. S. Prasad, S. E. Shoelson, C. E. Rudd. 1995. CTLA-4 binding to lipid linkase phosphatidyl-3-kinase in T cells. J. Exp. Med. 181:351
Colombo, M. I., J. Inglese, C. D’Souza-Schorey, W. Beron, P. D. Stahl. 1995. Heterotrimeric G proteins interact with the small GTPase ARF: possibilities for the regulation of vesicular traffic. J. Biol. Chem. 270:24564.
Dascher, C., W. E. Balch. 1994. Dominant inhibitory mutants of ARF1 block endoplasmic reticulum to Golgi transport and trigger disassembly of the Golgi apparatus. J. Biol. Chem. 269:1437.
Brown, F. D., N. Thompson, K. M. Saqib, J. M. Clark, D. Powner, N. T. Thompson, R. Solari, M. J. Wakelam. 1998. Phospholipase D1 localises to secretory granules and lysosomes and is plasma-membrane translocated on cellular stimulation. Curr. Biol. 8:835.
Freyberg, Z., A. Siddhanta, D. Shields. 2003. Slip, sliding away: phospholipase D and the Golgi apparatus. Trends Cell Biol. 13:540.
Boshell, M., J. McLeod, L. Walker, N. Hall, Y. Patel, D. Sansom. 1996. Effect of antigen presentation on superantigen induced apoptosis mediated by Fas/Fas ligand interactions in human T cells. Immunology 87:586.
Tzachanis, D., L. J. Appleman, A. A. Van Puijenbroek, A. Berezovskaya, L. M. Nadler, V. A. Boussiotis. 2003. Differential localization and function of ADP-ribosylation factor-6 in anergic human T cells: a potential marker for their identification. J. Immunol. 171:1691
Korthauer, U., W. Nagel, E. M. Davis, M. M. Le Beau, R. S. Menon, E. O. Mitchell, C. A. Kozak, W. Kolanus, J. A. Bluestone. 2000. Anergic T lymphocytes selectively express an integrin regulatory protein of the cytohesin family. J. Immunol. 164:308.
Anandasabapathy, N., G. S. Ford, D. Bloom, C. Holness, V. Paragas, C. Seroogy, H. Skrenta, M. Hollenhorst, C. G. Fathman, L. Soares. 2003. GRAIL: an E3 ubiquitin ligase that inhibits cytokine gene transcription is expressed in anergic CD4+ T cells. Immunity 18:535.(Karen I. Mead, Yong Zheng)
CTLA-4 is an essential protein in the regulation of T cell responses that interacts with two ligands found on the surface of APCs (CD80 and CD86). CTLA-4 is itself poorly expressed on the T cell surface and is predominantly localized to intracellular compartments. We have studied the mechanisms involved in the delivery of CTLA-4 to the cell surface using a model Chinese hamster ovary cell system and compared this with activated and regulatory human T cells. We have shown that expression of CTLA-4 at the plasma membrane (PM) is controlled by exocytosis of CTLA-4-containing vesicles and followed by rapid endocytosis. Using selective inhibitors and dominant negative mutants, we have shown that exocytosis of CTLA-4 is dependent on the activity of the GTPase ADP ribosylation factor-1 and on phospholipase D activity. CTLA-4 was identified in a perinuclear compartment overlapping with the cis-Golgi marker GM-130 but did not colocalize strongly with lysosomal markers such as CD63 and lysosome-associated membrane protein. In regulatory T cells, activation of phospholipase D was sufficient to trigger release of CTLA-4 to the PM but did not inhibit endocytosis. Taken together, these data suggest that CTLA-4 may be stored in a specialized compartment in regulatory T cells that can be triggered rapidly for deployment to the PM in a phospholipase D- and ADP ribosylation factor-1-dependent manner.
Introduction
Optimal activation of T cells requires the coordinated engagement of multiple receptors expressed on both T cells and APCs. Of these, binding of the TCR to specific peptide-MHC complexes and costimulation of the T cell through the CD28 receptor are important activating events. CD28 is stimulated by two ligands on the APCs CD80 and CD86, which provide signals that enhance T cell proliferation, cytokine production, and survival (1, 2). However, this system is complicated by the fact that a related receptor, CTLA-4, also interacts with CD80 and CD86 but has inhibitory effects on T cell function and cell cycle progression (3, 4).
The role of CTLA-4 in regulating T cell activation is evident from CTLA-4-deficient mice. These mice exhibit severe lymphoproliferative disease and die 3–4 wk after birth, indicating that CTLA-4 is critical for maintaining tolerance to self-tissues (5, 6, 7). There are several possibilities for the mechanism of CTLA-4 function. These include direct inactivating signals to T cells, competition with CD28 for binding to ligands on APCs, enhancement of suppression by specialized regulatory T (Treg)3 cells, and stimulation of tryptophan catabolism in APCs (1, 2). Recently, genetic studies have suggested that soluble CTLA-4 or even forms of CTLA-4, which cannot bind ligands, may be involved in disease susceptibility (8, 9). Thus, as yet, no single model of CTLA-4 function has emerged, and it is possible that more than one mechanism exists.
One major difference between CD28 and CTLA-4 lies in their expression patterns in T cells. CD28 is expressed constitutively on the surface of both resting and activated T cells. In contrast, CTLA-4 expression is undetectable in resting T cells. In activated T cells, despite equivalent mRNA levels to CD28, surface expression of CTLA-4 is much lower. This is because the majority of CTLA-4 molecules appears to be intracellular as a result of rapid endocytosis from the cell surface (10, 11). Internalization is thought to be controlled by a tyrosine-based motif, YxxM, in the cytoplasmic domain of CTLA-4 that interacts with the clathrin adaptor complex AP-2 (12, 13). Furthermore, it has been suggested that phosphorylation of tyrosine (Y)201 in this motif by Src kinases disrupts interaction with AP-2, resulting in stabilization of CTLA-4 at the cell surface (12). Interestingly, CD28 also contains a YxxM motif in its cytoplasmic tail yet is not endocytosed, suggesting that regulation of CTLA-4 expression is likely to be complex. Indeed, the observation that the entire cytoplasmic domain of CTLA-4 is absolutely conserved across a number species suggests that much of the sequence may be critical for its correct localization, membrane expression, and function. Interestingly, a specialized subset of CD4+ T cells (Treg) retain a constitutive intracellular pool of CTLA-4, suggesting that there may be a storage compartment from which CTLA-4 can undergo regulated exocytosis, possibly in a similar manner to proteins such as GLUT4 (14).
Despite numerous studies on CTLA-4-AP-2 interaction, little is known regarding the intracellular trafficking pathways used by CTLA-4 to gain access to the plasma membrane (PM). Typically, such transport of molecules through the cell is controlled by recruitment of proteins into coated vesicles that are transported from the trans-Golgi network toward the PM (15). Clathrin adaptor complexes, such as AP-1 and AP-3, are thought to be required for these vesicles and target them to the PM or to lysosomes, respectively (16, 17). Recently, the AP-3 adaptor complex has been shown to be required for the microtubule-mediated movement of lytic granules in CTLs (18). Consistent with its location in intracellular vesicles, CTLA-4 is also thought to interact with AP-1 via its cytoplasmic YVKM motif (19). As with CD28 (20), CTLA-4 has also been reported to interact with PI3K via this motif; however, the significance of this interaction is unclear at present (21).
The budding of clathrin-coated vesicles depends on a number of additional components, including the small ADP ribosylation factor (ARF) GTPases and the enzyme phospholipase D (PLD). ARF-1 is the most studied member of the ARF family and has been found to be required for assembly of coated vesicles at a variety of transport steps (22, 23). PLD is widely distributed and catalyzes the conversion of the membrane phospholipid, phosphatidylcholine, into phosphatidic acid and choline. This leads to high levels of phosphatidic acid within local membranes, resulting in increased binding of coat proteins and subsequent vesicle formation. Accordingly, PLD had been implicated at several stages of vesicle trafficking (24, 25).
Given the critical functions of CTLA-4 within the immune system and its unusual pattern of cellular expression, we have sought to characterize the controls that regulate its PM expression. We have established a model system of CTLA-4 trafficking in Chinese hamster ovary (CHO) cells and compared this with normal human T cells. Using this model, we have examined the requirement for PLD-1, PLD-2, and ARF-1 in exocytosis of CTLA-4 using selective inhibitors and dominant negative mutants. Our data demonstrate that PLD-1, PLD-2, and ARF-1 are required for transport of CTLA-4 proteins from the Golgi apparatus to the PM and that inhibition of these pathways prevents cell surface expression of CTLA-4. Furthermore, using stimuli that result in T cell activation, we observed a substantial increase in exocytosis of CTLA-4 that was prevented by inhibitors of both ARF and PLD. Despite this increase in traffic to the PM, only small increases of steady-state levels of surface CTLA-4 were observed, indicating that increased exocytic traffic was also accompanied by continued endocytosis. Our data identify several key points in trafficking pathways that regulate surface expression of CTLA-4 and indicate that regulated exocytosis is a major mechanism for controlling cell surface expression.
Materials and Methods
CTLA-4-transfected cells
CTLA-4-transfected CHO (CHO-CTLA-4) cells were generated by electroporation using the full-length and mutated forms of human CTLA-4 cDNA cloned into a CMV expression vector pCDNA-3. Cells were grown in DMEM as described previously (26). Cells expressing the plasmid were selected using G418 (500 μg/ml) treatment and sorted by FACS. Cultures were maintained at 37°C in a humid atmosphere containing 5% CO2 and were trypsinized and passaged every other day at 75% confluence.
Stable cultures of CHO-CTLA-4 cells were retransfected with wild-type or dominant negative GFP-tagged PLD and ARF plasmids (27). These cells (2 x 106) were transfected transiently by electroporation using an Amaxa nucleofector device, according to the manufacturer’s instructions, for CHO cell transfections and analyzed at 24 h.
Where human embryonic kidney (HEK) cells (HEK-293) were used for colocalization studies, the cells were cultured and transfected as for CHO cells with the exception that the Amaxa conditions for HEK cells were followed.
T cells
Buffy coats from blood donated by healthy donors were obtained from the National Blood Service. PBMCs were isolated using Ficoll density gradient centrifugation at 800 x g for 30 min. The buoyant layer was removed and washed twice in RPMI 1640 medium (supplemented with 10% FCS, 100 U/ml penicillin, and 100 μg/ml streptomycin). CD4+ T cells were purified by negative selection using magnetic beads, and CD4+CD25+ regulatory cells were purified subsequently by positive selection using CD25-immunomagnetic beads, according to the manufacturer’s instructions (Miltenyi Biotec).
Inhibitors
Brefeldin A was used at a concentration of 1 μg/ml. Cells were incubated at 37°C in a humid atmosphere containing 5% CO2 for 3 h before Ab staining. Butan-1-ol and butan-2-ol were used at a concentration of 1.5% in transfected CHO cells and 1.0% in T cell blasts. Cells were incubated for 1 h before Ab staining. Cells treated with cycloheximide were incubated for the times shown with 10 μg/ml in medium.
Flow cytometry
A total of 2 x 105 cells suspended in 100 μl of medium was treated with butan-1-ol, butan-2-ol, or brefeldin A as required and incubated with PE-conjugated anti-CTLA-4 (BN13; BD Pharmingen). Surface expression was detected by incubating cells at 4°C for 1 h, and recycled CTLA-4 was detected by incubation at 37°C. Total CTLA-4 expression was determined by fixing cells in 2% paraformaldehyde for 5 min at 4°C. Cells were washed once in PBS followed by anti-CTLA-4 Ab in PBS containing 0.1% saponin. Cells were washed in PBS/saponin and analyzed. Control samples were stained using the same protocol using a PE-labeled isotype control Ab. Where indicated, surface bound Abs were removed by acid washing for 2 min in PBS adjusted to pH 2.0. Percentage internalization was calculated by mean fluorescence intensity acid wash/mean fluorescence intensity PBS wash x 100. Analysis was conducted using a FACScan flow cytometer (BD Biosciences), and data for 10,000 cells were collected.
Confocal microscopy
A total of 5 x 104 cells was grown on multispot slides overnight in 20-μl drops. Cells were fixed either in ice-cold methanol for 5 min or in 2% formaldehyde for 5 min, followed by 0.1% saponin. Cells were incubated with 10% FCS in PBS before Ab addition to block nonspecific binding. Anti-GM130 was obtained from BD Biosciences and used at 1:100. Mouse anti-human CD63 and rabbit anti-human lysosome-associated membrane protein (LAMP) were generous gifts from Dr. G. Griffiths (University of Oxford, Oxford, U.K.). Mouse anti-human CD71 (transferrin receptor) was obtained from D. Hardie (University of Birmingham, Birmingham, U.K.). Anti-EEA-1 was a gift from B. Reaves (University of Bath, Bath, U.K.). Primary Abs were visualized using anti-mouse or anti-rabbit Abs conjugated to Alexa Fluor 488 or 594 (Molecular Probes). Anti-CTLA-4 Ab (11G1; a gift from Dr. J. Allison, University of California, Berkeley, CA) was directly conjugated to Alexa Fluor 594 (Molecular Probes) in our laboratory and used at 1/100 dilution. Staining for CTLA-4 was conducted at 4°C, 37°C, or after fixing and permeabilization as indicated. 4',6'-Diamidino-2-phenylindole (DAPI) was added to fixed cells at a concentration of 10 μg/ml for 1 min at room temperature to stain nuclei. Cells were washed twice and mounted onto slides. Fluorescence was examined using a Zeiss Axiovert 100M confocal microscope and a Zeiss LSM 510 scan module. All images were obtained with a C-Apochromat x63 water lens. For each image, optical sections were obtained at intervals of 0.5 μm through the cell, and the images shown are representative of sections through the centre of the cell.
Results
CTLA-4 trafficking is conserved in CHO cells and activated T cells
To facilitate the study of CTLA-4 intracellular trafficking and surface expression, we used transfected CHO cells constitutively expressing CTLA-4 as a model system. The expression pattern of CTLA-4 was compared with activated peripheral blood T cells using three different staining protocols. Staining at 4°C measured CTLA-4 expressed at the cell surface only. In contrast, staining at 37°C labeled all molecules arriving at the cell surface during the incubation period but did not distinguish whether they remained at the cell surface or were subsequently endocytosed. Therefore, labeling at 37°C gives an accurate indication of the total amount of CTLA-4 that reaches the PM over a given period of time, although this will include some newly synthesized protein. Finally, the total pool of CTLA-4 protein was measured in cells that were fixed and permeabilized before Ab staining. As shown in Fig. 1, this analysis revealed strong similarities between CTLA-4 expressed in CHO cells and in activated T cells. In both cases, much lower levels of cell surface CTLA-4 expression were observed in cells stained at 4°C when compared with cells stained at 37°C (Fig. 1a). Furthermore, cells that were fixed and permeabilized revealed similar levels of expression to cells stained at 37°C. Taken together, these data suggested that CTLA-4 was continually transported to the cell surface in both CHO and activated T cells but that steady-state levels of surface CTLA-4 were low due to continual endocytosis.
FIGURE 1. The pattern of CTLA-4 expression is similar in T cell blasts and CHO-CTLA-4 cells. a, CHO-CTLA-4 cells and T cells (unstimulated and stimulated with PMA and ionomycin) were stained for cell surface CTLA-4 (4°C staining), endocytosed CTLA-4 (37°C staining), or total CTLA-4 (F + P) using anti-CTLA-4-PE. A total of 5000 cells was analyzed by flow cytometry. b, FACS analysis of CHO-CTLA-4 cells in the presence of cycloheximide (CHX). Transfectants were stained 4°C, 37°C, and in fixed cells (total) after CHX treatment for the times shown. c, Confocal analysis of CTLA-4 in T cells and transfectants determined at 37°C and after fixing and permeabilizing (F + P) using anti-CTLA-4 conjugated to Alexa 594.
To ensure that the increased levels obtained at 37°C were not simply a reflection of newly synthesized CTLA-4, these experiments were repeated using transfectants in the presence of cycloheximide to inhibit new protein synthesis. This data (Fig. 1b) revealed that although the total level of CTLA-4 expression diminished over time with cycloheximide, the difference between staining at 4°C and 37°C remained, clearly demonstrating that new protein synthesis does not account for the difference between 4°C and 37°C, which supports the conclusion that intracellular CTLA-4 is continually transported to the PM at 37°C.
Finally, we analyzed both T cells and CHO cells by confocal microscopy (Fig. 1c). Staining at 37°C revealed that in both CHO and T cells CTLA-4 expression was located predominantly in intracellular vesicles; again, consistent with the concept that CTLA-4 was being exported to the PM and then rapidly endocytosed, contributing to the low steady-state level seen at the cell surface. Likewise, in fixed and permeabilized cells, a perinuclear compartment was observed containing high levels of CTLA-4 molecules representing either newly synthesized CTLA-4 or protein that had been endocytosed and targeted back to this compartment. Taken together, these observations suggested that CHO cells and activated T cells had very similar patterns of CTLA-4 expression.
Tyrosine 201 is not essential for CTLA-4 endocytosis
Endocytosis of CTLA-4 has been suggested to require a cytoplasmic 201YVKM motif for interaction with the clathrin adaptor AP-2. Therefore, we investigated the requirement for this motif in regulating CTLA-4 expression. Although mutation 201YVKM to 201FVKM appears to result in higher steady-state levels of CTLA-4 at the cell surface compared with wild type, analysis by confocal microscopy still revealed the presence of endocytosed CTLA-4 in intracellular vesicles. This appeared similar to wild-type CTLA-4 but was in clear contrast to a chimeric construct expressing CTLA-4 with a CD28 cytoplasmic domain, which was stably expressed at the cell surface (Fig. 2a). This suggested that endocytosis was not abolished in the 201FVKM mutant. Furthermore, the intracellular vesicles observed were not an artifact of nonspecific endocytosis or macropinocytosis of the CTLA-4 Ab because this did not occur with a hybrid molecule containing CTLA-extracellular and transmembrane domains but with a CD28-cytoplasmic domain. We also determined the kinetics of endocytosis by flow cytometry using an acid wash to remove surface CTLA-4 staining (Fig. 2b). This revealed that although endocytosis of CTLA-4 was slower (presumably leading to higher levels of surface CTLA-4 at steady state), by 20 min all of the surface-labeled CTLA-4 201FVKM was internalized. Thus, our data revealed that this motif is not essential for endocytosis of CTLA-4 in a cellular context. This information combined with the finding that activated human T cells only express weak cell surface CTLA-4 despite substantial CTLA-4 traffic to the PM at 37°C (see Fig. 1a) strongly suggests that during T cell activation changes in CTLA-4 expression might be regulated through exocytosis and not only by inhibiting endocytosis.
FIGURE 2. Mutation of tyrosine 201 does not prevent clathrin-mediated endocytosis. a, Expression patterns of wild-type (WT) and Y201F and CTLA-4-CD28 molecules in CHO cells were determined by confocal microscopy in fixed cells. Cells were stained using anti-CTLA-4-Alexa 594. b, Rates of CTLA-4 endocytosis were determined for WT and Y201F CTLA-4 and for CD28 in CHO cells. Cells were labeled at 4°C with anti-CTLA-4-PE, washed, and raised to 37°C for the time shown. Nonendocytosed Ab was removed by acid washing, and the levels of internalized (acid protected) CTLA-4 were measured by flow cytometry. c, The effect of sucrose was compared on internalization of anti-CTLA-4-PE after 30 min. Cells were labeled and washed as above either in the presence or absence of 0.4 M sucrose. Cells were washed in acid buffer and analyzed by FACS.
Because endocytosis of CTLA-4 has been reported previously to occur via an AP-2-mediated, clathrin-dependent pathway (12), we also treated CHO-CTLA-4 with hypertonic sucrose, which is known to inhibit the formation of functional clathrin-coated vesicles (28). As shown in Fig. 2c, cells treated in this way were unable to endocytose surface CTLA-4 molecules, resulting in accumulation of CTLA-4 at the PM. In contrast, in the absence of sucrose, both wild-type and CTLA-4 201FVKM transfectants internalized CTLA-4 but not the chimeric CTLA-4-CD28 molecule. Overall, these data demonstrated that CHO-CTLA-4 provided a model of CTLA-4 trafficking that required information resident in the CTLA-4-cytoplasmic domain and proceeded via clathrin-mediated endocytosis. Nonetheless, the YVKM-AP-2-binding motif was not essential to this process.
Inhibition of PLD-1 and ARF-1 blocks CTLA-4 exocytosis
We next initiated studies on the role of exocytosis in control of CTLA-4 expression in CHO-CTLA-4 cells. Initially, CHO-CTLA-4 cells were treated with inhibitors of PLD signaling (butan-1-ol) to determine the involvement of this pathway in trafficking of CTLA-4 containing vesicles (Fig. 3a). Strikingly, this treatment resulted in a very substantial reduction of staining at both 4°C and 37°C. However, CTLA-4 was still expressed intracellularly, as revealed in permeabilized cells. Furthermore, Butan-1-ol had no significant effect on the expression of CD28, which is not thought to interact with either AP-1 or AP-2. In control experiments, butan-2-ol, which does not inhibit PLD signaling, had no effect on CTLA-4 expression. A similar picture was observed using brefeldin A, which inhibits ARF proteins (Fig. 3b). Taken together, these results demonstrated that disruption of either ARF or PLD function was sufficient to prevent trafficking of CTLA-4 to the cell surface but did not affect CD28 expression.
FIGURE 3. PLD and ARF proteins are required for transport of CTLA-4 to the PM. a, CTLA-4- or CD28-transfected CHO cells were treated with 1.5% butan-1-ol, butan-2-ol, or 1 μg/ml brefeldin A (BFA) in DMEM for 3 h at 37°C. Cells were then stained for CTLA-4 or CD28 expression for 1 h at either 4°C, 37°C, or for total CTLA-4 expression after fixing and permeabilization (F + P). Each histogram shows the isotype control staining (shaded histogram) or specific Ab with (bold line) or without (thin line) butanol. b, The effect of BFA on CTLA-4 and CD28 expression was compared at 37°C. In all panels, 10,000 events were collected and analyzed by flow cytometry.
To confirm that the results obtained from inhibitors were not due to nonspecific or toxic effects and to investigate which specific ARF and PLD proteins were involved, a genetic approach was also adopted. CTLA-4-CHO cells were transfected with plasmids expressing either GFP-tagged, wild-type or dominant negative PLD-1 and ARF-1 proteins and analyzed by flow cytometry (Fig. 4a). This showed that dominant negative mutants of both PLD-1 and ARF-1 inhibited expression of cell surface CTLA-4 proteins detected at 4°C or at 37°C. In contrast, although GFP fluorescence was reduced by the permeabilization protocol, the total levels of CTLA-4 expression were unaffected by dominant negative ARF-1 or PLD-1. These effects were most obvious at 37°C where wild-type PLD and ARF did not inhibit CTLA-4 staining (cells are seen in Fig. 4a, upper right quadrant), whereas with the dominant negative GFP, proteins prevented CTLA-4 staining (cells are seen in Fig. 4a, lower right quadrant). Data for dominant negative PLD-2 was indistinguishable from PLD-1 (data not shown). Thus, the expression of both dominant negative PLD-1, PLD-2, or ARF-1 proteins prevented CTLA-4 from reaching the PM. When using dominant negative constructs, we did observe a slight drop in CTLA-4 staining cells in apparently untransfected cells. However, these transfections were also associated with slightly lower overall levels of CTLA-4 staining, suggesting this may be a result of decreased staining in these cells. However, it is possible that transfections with the dominant negative plasmids had a slight nonspecific inhibitory effect. Nonetheless, the data in the GFP-positive populations were unequivocal, and we concluded that ARF-1 and PLD-1 were required for effective transport of CTLA-4 to the cell surface. Consistent with these observations, confocal microscopy on cells transfected with dominant negative and wild-type ARF-1 revealed a lack of staining of CTLA-4 at both 4°C and 37°C with dominant negative ARF-1 (Fig. 4b). However, once again, total staining was unaffected. Taken together, these data indicate that CTLA-4 transport to the PM is dependent on both ARF and PLD pathways.
FIGURE 4. Surface expression of CTLA-4 is blocked by dominant negative (DN) PLD-1 and ARF-1. a, Stable lines of CHO-CTLA-4 cells were transfected transiently with GFP-tagged wild-type (WT) or DN PLD-1 and ARF-1. Expression of GFP proteins was detected along the x-axis and compared with CTLA-4 expression (BN13-PE) on the y-axis using flow cytometry. CTLA-4 staining was determined at 4°C and 37°C and in permeabilized cells to determine effects on CTLA-4 expression. b, Confocal analysis of cells transfected with DN-ARF-1 as in a, with the exception that CTLA-4 was detected using Alexa 594-conjugated Ab, and nuclei were counterstained using DAPI (white). Arrowheads highlight the lack of CTLA-4 staining in DN-ARF-transfected cells.
CTLA-4 is localized to a distinct intracellular compartment
To characterize the intracellular location of CTLA-4, we performed colocalization experiments. Initially, these were performed in CHO cells; however, the lack of suitable colocalization reagents for these cells prompted us to transfect CTLA-4 into HEK-293. This allowed us to study the localization of several endogenous proteins, including CD63 and transferrin receptor, both of which are known to be internalized by clathrin-mediated endocytosis. Importantly, the staining pattern of CTLA-4 in both CHO and HEK systems was indistinguishable by confocal microscopy and retained the same characteristics at 37°C and 4°C by FACS. In fixed HEK cells, CTLA-4 colocalized significantly in a perinuclear location with GM130, which represented Golgi staining. However, despite reports that CTLA-4 may be stored in secretory lysosomes (29), the amount of CTLA-4 that colocalized with the lysosomal markers LAMP-2 and CD63 was very limited (Fig. 5). We also performed staining for CD71 and for the early endosome marker EEA-1. However, once again, while there was some colocalization with these compartments, in particular with EEA-1, this was clearly incomplete. In control experiments, CD63 and LAMP demonstrated complete colocalization. These results indicated that CTLA-4 is found in a both a perinuclear compartment (that most likely contains both pre- and postendocytic vesicles), as well as distinct cytoplasmic vesicles; some of which are EEA-1 positive. Surprisingly, CTLA-4-containing vesicles did not appear to be strongly CD63 or CD71 associated, despite the fact that both proteins contain an AP-2-sorting motif, indicating that CTLA-4 has a distinct trafficking itinerary.
FIGURE 5. Colocalization of CTLA-4. CTLA-4-transfected HEK-293 cells were fixed and permeabilized and stained with anti-CTLA-4 Alexa 594 and analyzed by confocal microscopy. Cells were costained (green) for LAMP-1, GM130, EEA-1, or CD71 as shown. Control colocalization for CD71 and CD63 with LAMP is shown. Sections shown are representative staining of an optical section through the middle of the cell. Nuclei were counterstained with DAPI (blue). Colocalization is represented as yellow.
Up-regulation of CTLA-4 on T cells results from increased exocytosis and is dependent on ARF and PLD activity
Because data in CHO cells indicated a critical role for ARF-1 and PLD in the efficient export of CTLA-4 to the PM, we examined this process in T cells. Purified human T cells were stimulated using PMA and ionomycin for 4 h to induce CTLA-4 expression and the effect of butanol treatment studied as before (Fig. 6a). As observed with CHO cells. treatment with butan-1-ol abolished PM traffic with only minor effects on the total level of CTLA-4. Similar effects were also observed with brefeldin A (data not shown), indicating that data derived from the CHO-CTLA-4 was consistent with human T cells.
FIGURE 6. PLD and ARF are required for stimulated exocytosis of CTLA-4 in CD4+ and CD4+CD25+ human T cells. a, Purified resting CD4+ T cells were stimulated for 4 h with PMA (5 ng/ml) and ionomycin (500 μM) to induce CTLA-4 expression in the presence (bold line) or absence (thin line) of butanol. CTLA-4 expression was detected at 4°C and 37°C and in permeabilized cells by flow cytometry using anti-CTLA-4-PE. b, Treg cells were stimulated for 2 h with PMA (5 ng/ml) and stained at 37°C (dashed line) in the presence of butan-1-ol (bold line) and butan-2-ol (thin line). Control Treg cells stained for anti-CTLA-4 but not stimulated are shown (shaded histogram). Brefeldin A (BFA) treatment is shown as a bold line.
We also studied this process using CD4+CD25+ Treg cells, which constitutively express intracellular CTLA-4. Because PMA is a well-established activator of PLD, we predicted that PMA alone should increase CTLA-4 membrane traffic in the absence of full T cell activation. This experiment (Fig. 6b) revealed that unstimulated Treg did not constitutively recycle CTLA-4 to the PM as detected by labeling at 37°C. However, stimulation with PMA resulted in a rapid and marked up-regulation of CTLA-4 within 2 h that was abolished by inhibiting PLD or ARF proteins. These data directly demonstrate that up-regulation of CTLA-4 in human T cells is driven by regulated exocytosis in a PLD-dependent manner. Furthermore, these data indicate that in Treg CTLA-4 is stored in a compartment that is sensitive to such regulated exocytosis.
Discussion
CTLA-4 is an essential protein for immune regulation, the absence of which leads to fatal autoimmune tissue destruction. A major feature of this protein is its unusual pattern of intracellular expression and its highly conserved cytoplasmic domain, which is as yet without a clearly established function. To better understand how CTLA-4 expression is controlled, we developed a model of CTLA-4 trafficking in CHO cells that is amenable to confocal analysis and genetic manipulation. We have validated this model against human T cells and found no obvious differences in the patterns of CTLA-4 expression, with the exception that trafficking of CTLA-4 to the PM was constitutive in CHO cells and whereas it was stimulation dependent in T cells. However, in both cases, our data establish that trafficking is dependent on ARF-1 and PLD activity.
The data presented suggest a critical role for PLD and ARF-1 in movement of CTLA-4-containing vesicles from a perinuclear region to the PM. This compartment was observed during labeling at both 37°C and in fixed cells, suggesting it contains both postendocytic vesicles as well as newly budding vesicles. Interestingly, clathrin-coated vesicles that use the adapter protein AP-1 are generally associated with the trans-Golgi network, and consistent with this finding, yeast two-hybrid studies have shown that CTLA-4 can indeed interact with AP-1 via its YVKM motif (19). In support of this, we observed colocalization with GM-130, which is a marker of the cis-Golgi.
Several studies have suggested that CTLA-4 is located in lysosomes or secretory granules and may be translocated via secretory lysosomes (11, 29, 30). In our model, we only observed limited colocalization with lysosomal markers LAMP-1 and CD63, suggesting that the majority of CTLA-4 is not within lysosomes. Furthermore, despite it being relatively clear that in CD8+ T cells and other specialized secretory cells—and secretory lysosomes represent a significant mechanism of exocytosis (31)—it is not clear whether this is a major mechanism for CTLA-4 in CD4+ cells and, in particular, Treg cells. In our hands, the levels of surface CTLA-4 seen following ionomycin (a stimulus for lysosome secretion) are substantially less than that seen with PMA, which may stimulate generalized vesicle traffic via PLD. We believe this is more consistent with a nonlysosomal store as the major source of membrane-translocated CTLA-4. Furthermore, given that lysosomes are clearly capable of degrading CTLA-4 rapidly (32), this seems unlikely to be the location of long-term CTLA-4 storage for cells such as Treg. Thus, although CTLA-4 can clearly be detected in lysosomes, additional studies are needed to clarify the role of this compartment in stimulated exocytosis.
The pattern of CTLA-4 trafficking observed in our studies appears similar to that of the glucose transporter GLUT4 (14). GLUT4 is a recycling receptor that interacts with both AP-1 and AP-2 and recycles to a trans-Golgi network 38-negative compartment (33). Furthermore, PLD activity has also been implicated in translocation of GLUT4 to the PM (34). Perhaps most interestingly, both CTLA-4 and GLUT4 appear to use a storage compartment from which exocytosis is achieved rapidly upon stimulation. In the case of GLUT4, this stimulation appears to be via a PI3K-dependent mechanism. Although the pathway responsible for stimulating for CTLA-4 exocytosis in T cells remains to be elucidated, our data show that activation of PLD using phorbol ester is sufficient for translocation. Interestingly, the major known physiological ligands that drive T cell activation (TCR and CD28) are known to activate PI3K (20, 35) and PLD (36), suggesting this mechanism may be applicable during normal T cell stimulation. However, although there is a report that wortmannin can up-regulate CTLA-4 expression (37), we have observed no consistent effects of PI3K in our studies, which leaves the role of PI3K in CTLA-4 trafficking needing additional investigation.
The data presented here suggest that despite delivery to the cell surface CTLA-4 is continually endocytosed. This seems to conflict somewhat with previous suggestions that expression of CTLA-4 at the cell surface is stabilized by phosphorylation of its cytoplasmic domain, thereby disrupting AP-2-mediated endocytosis. However, direct data measuring endocytosis of CTLA-4 are somewhat limited. Shiratori et al. (12) showed clearly that CTLA-4 interacts via its cytoplasmic domain with AP-2 using the YVKM motif. However, data directly showing that activation of T cells caused significant phosphorylation of CTLA-4, resulting in stable cell surface CTLA-4, are missing. Indeed, increased CTLA-4 at the cell surface is generally only seen using pervanadate as a phosphatase inhibitor and not under normal conditions of T cell activation (12, 38). Interestingly, our own experiments with pervanadate in T cells (K. Mead, unpublished observations) reveal that CTLA-4 expression is markedly more enhanced at 37°C compared with 4°C, which raises the possibility that this increase is due to enhanced delivery rather than decreased endocytosis. Therefore, it may be significant that pervanadate can act by stimulating PLD activation and thereby possibly enhance exocytosis (39). In other studies, kinases such as lck and Fyn have been found to significantly phosphorylate CTLA-4 (40), yet without the use of pervanadate, this does not affect levels of CTLA-4 at the cell surface, suggesting that simple phosphorylation of CTLA-4 does not block endocytosis (41).
Thus, one possibility consistent with our experience and data is that CTLA-4 endocytosis continues even under normal conditions of T cell activation and that mutation of Y201 does not inhibit endocytosis. Interestingly, others have made similar mutants and also observed predominantly intracellular CTLA-4 (29), suggesting this is not an artifact of expression in CHO cells. Furthermore, while it is clear that CTLA-4 may be phosphorylated on tyrosine 201, even in response to TCR stimulation (42), there is no direct evidence that this prevents internalization. Although increased surface staining is attributed to disrupted endocytosis in many studies, it is notable that endocytosis is not actually measured. Thus, while our results contrast with the current perception that disruption of endocytosis results in increased surface expression of CTLA-4, we believe it is more likely that regulated exocytosis of CTLA-4 is in fact the critical regulatory step.
Although we have yet to define how PLD is involved in CTLA-4 translocation, the inhibition by both the catalytically inactive form of PLD and butan-1-ol implicates the generation of phosphatidic acid in the regulatory process. We have previously observed in RBL-2H3 cells that trafficking of secretory lysosomes is not prevented by inhibition of PLD activity but that fusion of the vesicles with the PM was ablated (23). Thus, one possible role for the generated phosphatidic acid may be in controlling the fusion of the vesicles with the PM.
At the present time, the mechanism of CTLA-4 inhibitory action is not understood, and several modes of action are possible. Given the exceptional degree of conservation of the CTLA-4-cytoplasmic domain, the importance of regulated trafficking of CTLA-4 cannot be underestimated. CTLA-4 is thought to promote T cell anergy (unresponsiveness) (43). It is interesting to note that differential expression studies of anergic T cells have identified ARF-6, as well as the exchange factor GRP-1, as differentially expressed under anergic conditions (44, 45). Furthermore, anergy induction has recently been associated with up-regulation of proteins such as Grail, which has strikingly similar expression patterns to CTLA-4 (46). Taken together, this may well suggest that regulation of vesicle trafficking may be a target in T cell anergy. The present studies now provide the basis for more detailed analysis of the control of CTLA-4 trafficking.
Disclosures
The authors have no financial conflict of interest.
Footnotes
The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked advertisement in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
1 This work was supported by the Arthritis Research Campaign (ARC) (to D.M.S., C.N.M. and M.K.P.L.), the Biotechnology and Biological Sciences Research Council (to Y.Z.), and the Wellcome Trust (to M.J.O.W. and D.J.P.). D.M.S. is an ARC Senior Research Fellow. K.I.M. is a Medical Research Council PhD student.
2 Address correspondence and reprint requests to Dr. David M. Sansom, Medical Research Council Centre for Immune Regulation, University of Birmingham Medical School, Vincent Drive, Birmingham B15 2 TT, U.K. E-mail address: d.m.sansom{at}bham.ac.uk
3 Abbreviations used in this paper: Treg, regulatory T; PM, plasma membrane; ARF, ADP ribosylation factor; PLD, phospholipase D; CHO, Chinese hamster ovary; CHO-CTLA-4, CTLA-4 transfected CHO; HEK, human embryonic kidney; DAPI, 4',6'-diamidino-2-phenylindole; LAMP, lysosome-associated membrane protein.
Received for publication January 28, 2004. Accepted for publication February 4, 2005.
References
Sansom, D. M., C. N. Manzotti, Y. Zheng. 2003. What’s the difference between CD80 and CD86?. Trends Immunol. 24:313.
Sansom, D. M.. 2000. CD28, CTLA-4 and their ligands: who does what and to whom?. Immunology 101:169.
Walunas, T. L., C. Y. Bakker, J. A. Bluestone. 1996. CTLA-4 ligation blocks CD28-dependent T cell activation. J. Exp. Med. 183:2541.
Krummel, M. F., J. P. Allison. 1995. CD28 and CTLA-4 have opposing effects on the response of T cells to stimulation. J. Exp. Med. 182:459
Waterhouse, P., J. M. Penninger, E. Timms, A. Wakeham, A. Shahinian, K. P. Lee, C. B. Thompson, H. Griesser, T. W. Mak. 1995. Lymphoproliferative disorders with early lethality in mice deficient in CTLA-4. Science 270:985
Tivol, E. A., F. Borriello, A. N. Schweitzer, W. P. Lynch, J. A. Bluestone, A. H. Sharpe. 1995. Loss of CTLA-4 leads to massive lymphoproliferation and fatal multiorgan tissue destruction, revealing a critical negative regulatory role of CTLA-4. Immunity 3:541.
Chambers, C. A., T. J. Sullivan, J. P. Allison. 1997. Lymphoproliferation in CTLA-4-deficient mice is mediated by costimulation-dependent activation of CD4+ cells. Immunity 7:885.
Ueda, H., J. M. Howson, L. Esposito, J. Heward, H. Snook, G. Chamberlain, D. B. Rainbow, K. M. Hunter, A. N. Smith, G. Di Genova, et al 2003. Association of the T-cell regulatory gene CTLA4 with susceptibility to autoimmune disease. Nature 423:506.
Vijayakrishnan, L., J. M. Slavik, Z. Illes, R. J. Greenwald, D. Rainbow, B. Greve, L. B. Peterson, D. A. Hafler, G. J. Freeman, A. H. Sharpe, et al 2004. An autoimmune disease-associated CTLA-4 splice variant lacking the B7 binding domain signals negatively in T cells. Immunity 20:563.
Alegre, M.-L., P. J. Noel, B. J. Eisfelder, E. Chuang, M. R. Clark, S. L. Reiner, C. B. Thompson. 1996. Regulation of surface and intracellular expression of CTLA-4 on mouse T cells. J. Immunol. 157:4762.
Linsley, P. S., J. Bradshaw, J. Greene, R. Peach, K. L. Bennett, R. S. Mittler. 1996. Intracellular trafficking of CTLA-4 and focal localisation towards sites of TCR engagement. Immunity 4:535.
Shiratori, T., S. Miyatake, H. Ohno, C. Nakaseko, K. Isono, J. S. Bonifacino, T. Saito. 1997. Tyrosine phosphorylation controls internalization of CTLA-4 by regulating its interaction with clathrin-associated adaptor complex AP-2. Immunity 6:583.
Zhang, Y., J. P. Allison. 1997. Interaction of CTLA-4 with AP-50, a clathrin-coated pit adaptor protein. Proc. Natl. Acad. Sci. USA 94:9273.
Bryant, N. J., R. Govers, D. E. James. 2002. Regulated transport of the glucose transporter GLUT4. Nat. Rev. Mol. Cell Biol. 3:267.
Beraud-Dufour, S., W. Balch. 2002. A journey through the exocytic pathway. J. Cell Sci. 115:1779
Dell’Angelica, E. C.. 2001. Clathrin-binding proteins: got a motif: join the network!. Trends Cell Biol. 11:315.
Dell‘Angelica, E. C., V. Shotelersuk, R. C. Aguilar, W. A. Gahl, J. S. Bonifacino. 1999. Altered trafficking of lysosomal proteins in Hermansky-Pudlak syndrome due to mutations in the 3A subunit of the AP-3 adaptor. Mol. Cell 3:11.
Clark, R. H., J. C. Stinchcombe, A. Day, E. Blott, S. Booth, G. Bossi, T. Hamblin, E. G. Davies, G. M. Griffiths. 2003. Adaptor protein 3-dependent microtubule-mediated movement of lytic granules to the immunological synapse. Nat. Immunol. 4:1111.
Schneider, H., M. Martin, F. A. Agarraberes, L. Yin, I. Rapoport, T. Kirchhausen, C. E. Rudd. 1999. Cytolytic T lymphocyte-associated antigen-4 and the TCR /CD3 complex, but not CD28, interact with clathrin adaptor complexes AP-1 and AP-2. J. Immunol. 163:1868.
Ward, S., J. Westwick, N. Hall, D. Sansom. 1993. CD28 ligation elevates PtdIns(3,4)P2 and PtdIns(3,4,5)P3 in T cells. Eur. J. Immunol. 23:2572.
Schneider, H., V. S. Prasad, S. E. Shoelson, C. E. Rudd. 1995. CTLA-4 binding to lipid linkase phosphatidyl-3-kinase in T cells. J. Exp. Med. 181:351
Colombo, M. I., J. Inglese, C. D’Souza-Schorey, W. Beron, P. D. Stahl. 1995. Heterotrimeric G proteins interact with the small GTPase ARF: possibilities for the regulation of vesicular traffic. J. Biol. Chem. 270:24564.
Dascher, C., W. E. Balch. 1994. Dominant inhibitory mutants of ARF1 block endoplasmic reticulum to Golgi transport and trigger disassembly of the Golgi apparatus. J. Biol. Chem. 269:1437.
Brown, F. D., N. Thompson, K. M. Saqib, J. M. Clark, D. Powner, N. T. Thompson, R. Solari, M. J. Wakelam. 1998. Phospholipase D1 localises to secretory granules and lysosomes and is plasma-membrane translocated on cellular stimulation. Curr. Biol. 8:835.
Freyberg, Z., A. Siddhanta, D. Shields. 2003. Slip, sliding away: phospholipase D and the Golgi apparatus. Trends Cell Biol. 13:540.
Boshell, M., J. McLeod, L. Walker, N. Hall, Y. Patel, D. Sansom. 1996. Effect of antigen presentation on superantigen induced apoptosis mediated by Fas/Fas ligand interactions in human T cells. Immunology 87:586.
Tzachanis, D., L. J. Appleman, A. A. Van Puijenbroek, A. Berezovskaya, L. M. Nadler, V. A. Boussiotis. 2003. Differential localization and function of ADP-ribosylation factor-6 in anergic human T cells: a potential marker for their identification. J. Immunol. 171:1691
Korthauer, U., W. Nagel, E. M. Davis, M. M. Le Beau, R. S. Menon, E. O. Mitchell, C. A. Kozak, W. Kolanus, J. A. Bluestone. 2000. Anergic T lymphocytes selectively express an integrin regulatory protein of the cytohesin family. J. Immunol. 164:308.
Anandasabapathy, N., G. S. Ford, D. Bloom, C. Holness, V. Paragas, C. Seroogy, H. Skrenta, M. Hollenhorst, C. G. Fathman, L. Soares. 2003. GRAIL: an E3 ubiquitin ligase that inhibits cytokine gene transcription is expressed in anergic CD4+ T cells. Immunity 18:535.(Karen I. Mead, Yong Zheng)