This information is current as
of June 15, 2017.
Exocytosis of CTLA-4 Is Dependent on
Phospholipase D and ADP Ribosylation
Factor-1 and Stimulated during Activation of
Regulatory T Cells
Karen I. Mead, Yong Zheng, Claire N. Manzotti, Laura C. A.
Perry, Michael K. P. Liu, Fiona Burke, Dale J. Powner,
Michael J. O. Wakelam and David M. Sansom
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The Journal of Immunology is published twice each month by
The American Association of Immunologists, Inc.,
1451 Rockville Pike, Suite 650, Rockville, MD 20852
Copyright © 2005 by The American Association of
Immunologists All rights reserved.
Print ISSN: 0022-1767 Online ISSN: 1550-6606.
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J Immunol 2005; 174:4803-4811; ;
doi: 10.4049/jimmunol.174.8.4803
http://www.jimmunol.org/content/174/8/4803
The Journal of Immunology
Exocytosis of CTLA-4 Is Dependent on Phospholipase D and
ADP Ribosylation Factor-1 and Stimulated during Activation
of Regulatory T Cells1
Karen I. Mead,* Yong Zheng,* Claire N. Manzotti,* Laura C. A. Perry,* Michael K. P. Liu,*
Fiona Burke,* Dale J. Powner,† Michael J. O. Wakelam,† and David M. Sansom2*
O
ptimal 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 (5–7).
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
*Medical Research Council Centre for Immune Regulation, University of Birmingham Medical School, and †Cancer Research U.K. Institute for Cancer Studies, University of Birmingham, Birmingham, United Kingdom
Received for publication January 28, 2004. Accepted for publication February
4, 2005.
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.
[email protected]
Copyright © 2005 by The American Association of Immunologists, Inc.
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
3
Abbreviations used in this paper: Treg, regulatory T; PM, plasma membrane; ARF,
ADP ribosylation factor; PLD, phospholipase D; CHO, Chinese hamster ovary; CHOCTLA-4, CTLA-4 transfected CHO; HEK, human embryonic kidney; DAPI, 4⬘,6⬘diamidino-2-phenylindole; LAMP, lysosome-associated membrane protein.
0022-1767/05/$02.00
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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 lysosomeassociated 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. The Journal of Immunology, 2005, 174: 4803– 4811.
4804
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 ␮g/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 ⫻ 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 ⫻ 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 ␮g/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 ␮g/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 ␮g/ml in medium.
Flow cytometry
A total of 2 ⫻ 105 cells suspended in 100 ␮l of medium was treated with
butan-1-ol, butan-2-ol, or brefeldin A as required and incubated with PEconjugated 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 ⫻ 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 ⫻ 104 cells was grown on multispot slides overnight in 20-␮l
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 ␮g/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 ⫻63 water lens.
For each image, optical sections were obtained at intervals of ⬃0.5 ␮m
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
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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.
EXOCYTOSIS OF CTLA-4
The Journal of Immunology
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 antiCTLA-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.
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
201
FVKM 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 wildtype 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
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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.
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.
4805
4806
EXOCYTOSIS OF CTLA-4
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.
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.
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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.
The Journal of Immunology
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, CHOCTLA-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
4807
CTLA-4 is localized to a distinct intracellular compartment
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 ␮g/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 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
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both 4°C and 37°C. However, CTLA-4 was still expressed intracellularly, as revealed in permeabilized cells. Furthermore, Butan1-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.
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 wildtype 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.
4808
EXOCYTOSIS OF CTLA-4
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.
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
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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 DNARF-transfected cells.
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.
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
The Journal of Immunology
4809
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
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 ␮M) 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.
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.
Downloaded from http://www.jimmunol.org/ by guest on June 15, 2017
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.
4810
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.
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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
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In the case of GLUT4, this stimulation appears to be via a PI3Kdependent 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
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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,
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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
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also observed predominantly intracellular CTLA-4 (29), suggesting this is not an artifact of expression in CHO cells. Furthermore,
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it is notable that endocytosis is not actually measured. Thus, while
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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
EXOCYTOSIS OF CTLA-4
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