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The Journal of Immunology, 2008, 180, 6713 -6724
Copyright © 2008 by The American Association of Immunologists, Inc.
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The Chemokine Receptor CXCR3 Is Degraded following Internalization and Is Replenished at the Cell Surface by De Novo Synthesis of Receptor1

Andrea Meiser2, Anja Mueller2,3, Emma L. Wise, Ellen M. McDonagh, Sarah J. Petit, Namita Saran, Peter C. Clark, Timothy J. Williams and James E. Pease4

Leukocyte Biology Section, National Heart and Lung Institute, Faculty of Medicine, Imperial College London, South Kensington Campus, London SW7 2AZ, United Kingdom


   Abstract
Top
 Abstract
Introduction
 Materials and Methods
 Results
 Discussion
 Disclosures
 References
 
The chemokine receptor CXCR3 is expressed on the surface of both resting and activated T lymphocytes. We describe in this study the endocytosis of CXCR3 using T lymphocytes and CXCR3 transfectants. Chemokine-induced CXCR3 down-regulation occurred in a rapid, dose-dependent manner, with CXCL11 the most potent and efficacious ligand. Endocytosis was mediated in part by arrestins, but appeared to occur independently of clathrin and caveolae. In contrast to other chemokine receptors, which are largely recycled to the cell surface within an hour, cell surface replenishment of CXCR3 occurred over several hours and was dependent upon mRNA transcription, de novo protein synthesis, and transport through the endoplasmic reticulum and Golgi. Confocal microscopy and Western blotting confirmed the fate of endocytosed CXCR3 to be degradation, mediated in part by lysosomes and proteosomes. Site-directed mutagenesis of the CXCR3 C terminus revealed that internalization and degradation were independent of phosphorylation, ubiquitination, or a conserved LL motif. CXCR3 was found to be efficiently internalized in the absence of ligand, a process involving a YXXL motif at the extreme of the C terminus. Although freshly isolated T lymphocytes expressed moderate cell surface levels of CXCR3, they were only responsive to CXCL11 with CXCL9 and CXCL10 only having significant activity on activated T lymphocytes. Thus, the activities of CXCR3 are tightly controlled following mRNA translation. Because CXCR3+ cells are themselves a source of IFN-{gamma}, which potently induces the expression of CXCR3 ligands, such tight regulation of CXCR3 may serve as a control to avoid the unnecessary amplification of activated T lymphocyte recruitment.


   Introduction
Top
 Abstract
 Introduction
Materials and Methods
 Results
 Discussion
 Disclosures
 References
 
The chemokine receptor CXCR3 is expressed on a wide variety of cells including activated T lymphocytes, NK cells, malignant B lymphocytes, endothelial cells, and thymocytes (1, 2, 3, 4, 5, 6). Three major CXCR3 ligands, CXCL9, CXCL10, and CXCL11, have been identified, all of which are induced by IFN-{gamma} and are therefore thought to promote Th1 immune responses (7, 8, 9). Recent studies have shown that the CXCR3 ligands exhibit unique temporal and spatial expression patterns, suggesting that they have nonredundant functions in vivo. Moreover, the CXCR3 ligands share low sequence homology (around 40% amino acid identity) and exhibit differences in their potencies and efficacies at CXCR3 with CXCL11 being the dominant ligand in several assays (8, 10). CXCR3 and its ligands have been implicated as playing an important role in the induction and perpetuation of several human inflammatory disorders including atherosclerosis (11), autoimmune diseases (12), transplant rejection (13, 14), and viral infections (15). Consequently, the mechanisms underlying the regulation of CXCR3 expression at the cell surface are of considerable interest.

The number of receptors on a cell surface results from a balance between the rate of internalization and the rate of replacement (recycling and synthesis of nascent receptor). Following ligand binding, there are two major routes whereby G protein-coupled receptors (GPCRs),5 as typified by chemokine receptors, are internalized into cells. The first and most well-defined route involves the binding of arrestin to the phosphorylated receptor, which in turn initiates the internalization process by binding to clathrin. The receptor-arrestin complex is then sequestered in clathrin-coated pits. This pathway is often considered a default system for degradation and recycling of receptors (16, 17). The second pathway involves invaginations of the cell membrane known as caveolae and functions independently of clathrin-coated pits (18). Although the rate of internalization of a receptor is an important factor in determining its level at the cell surface, the rate of recycling and the rate of synthesis of new receptors are also important. Until recently, the mechanisms of the recycling process were poorly understood, and internalized receptors were thought to have several potential fates. The concept of two different classes of receptor (as distinguished by their recycling) has been introduced recently, in which class A receptors traffic to recycling endosomes and are rapidly returned to the cell surface (16). In contrast, class B receptors are dephosphorylated in endosomes followed by slow recycling back to the plasma membrane. Sequentially, the receptors pass through late endosomes and the Golgi and finally are transported back to the cell surface. Another potential fate is that of degradation, which may be perceived to down-regulate receptor expression. To date, protein synthesis has not been shown to play a role in GPCR replenishment (19, 20, 21).

In this study we show that CXCR3 is internalized both constitutively and following incubation with CXCL11, resulting in degradation of the receptor. We also show that in the absence of detectable recycling, cell surface replenishment of CXCR3 is dependent upon de novo protein synthesis.


   Materials and Methods
Top
 Abstract
 Introduction
 Materials and Methods
Results
 Discussion
 Disclosures
 References
 
Materials

All chemicals unless otherwise stated were purchased from Sigma-Aldrich. Chemokines were purchased from PeproTech. Filipin, sucrose, nystatin, monensin, and nocodazole were purchased from Sigma-Aldrich or Calbiochem. Brefeldin A, actinomycin D, and bafilomycin A1 were obtained from Tocris. Cycloheximide was from ICN Biomedicals. The mouse anti-human CXCR3 mAb (clone 49801.111) and the mouse isotype-matched control IgG1 (MOPC 21 clone) were obtained from Sigma-Aldrich. The anti-HA.11 Ab was from Covance, and the anti-{alpha}-tubulin Ab was from Abcam. Secondary Abs were obtained from DakoCytomation. Plasmids encoding dominant negative mutants of β-arrestin 1 and β-arrestin 2 were gifts of Dr. M. Caron (Duke University Medical Center, Durham, NC). Plasmids encoding the fusion proteins GFP-DIII and GFP-DIII{Delta}2 were gift of Dr. A. Benmarah (Institute Cochin, Paris, France).

Cell culture and transient transfection

The murine pre-B cell line L1.2 was maintained as previously described in RPMI 1640 supplemented medium (22). L1.2 cells stably transfected with pCDNA3 containing the CXCR3A cDNA hemagglutinin (HA)-tagged at the N terminus (10) were cultured in the same medium with the addition of 1 mg/ml geneticin (G418) to maintain selection. Mutant CXCR3 constructs were generated by site-directed mutagenesis using the QuikChange mutagenesis kit (Stratagene) with the pCDNA3 HA-CXCR3A plasmid as template. Transient transfection of L1.2 cells with plasmids was conducted by electroporation as previously described (23). Cells were cultured for 24 h in medium supplemented with 10 mM sodium butyrate before use to enhance cell surface receptor expression. Mouse embryonic fibroblasts (MEFs) derived from both wild-type (WT) mice and mice deficient in β-arrestins 1 and 2 were a gift of Dr. R. Lefkowitz (Duke University Medical Center, Durham NC) and were maintained as previously described (24). Transfection of MEFs was by electroporation as previously described (25). Cells were cultured overnight in medium supplemented with 10 mM sodium butyrate before use to enhance cell surface receptor expression.

For the generation of activated T lymphocytes, PBMC were isolated from blood sampled from healthy donors, according to the Royal Brompton Hospital Ethics Committee approved protocol as previously described (26). Lymphocytes were separated from monocytes by allowing the latter to adhere to a tissue culture flask for 2 h at 37°C and were activated by culture in the presence of 100 IU/ml IL-2 and 2 mg/ml Con A for 7–10 days. Purified T lymphocytes were isolated from whole blood using the Rosette-Sep Human T cell Enrichment Cocktail kit (StemCell Technologies), which typically gave a population >95% pure. Nucleofection of purified T lymphocytes with plasmids encoding GFP-DIII and GFP-DIII{Delta}2 was achieved by using an Amaxa nucleofector, according to the manufacturer’s instructions, using program U-014, which typically gave around 60% cell viability as deduced by staining with the dye ToPro3 (Invitrogen).

Internalization assay and flow cytometry analysis

Internalization assays were essentially conducted as previously described by Sauty et al. (27). Activated T lymphocytes or L1.2-CXCR3 cells were incubated with serum-free medium for 1 h at 37°C and then resuspended in medium without serum at 5 x 106 cells/ml. Cells were then incubated with chemokines (50 nM) for various times at 37°C, and washed in ice-cold PBS containing 1% FCS and 1% NaN3 before flow cytometry analysis. Cell surface-expressed CXCR3 was detected using an anti-CXCR3 Ab and FITC conjugated anti-mouse IgG. Samples were quantified on a FACSCalibur, and data processed with CellQuest software (version 3.1; BD Biosciences) with dead cells excluded from analysis. The relative CXCR3 surface expression was calculated as a percentage using the following: 100 x (mean channel of fluorescence (stimulated) – mean channel of fluorescence (negative control))/(mean channel of fluorescence (medium) – mean channel of fluorescence (negative control)). Pilot experiments staining CXCR3 transfectants on ice in either the presence or absence of 50 nM CXCL11 confirmed that binding of ligand by CXCR3 did not significantly reduce detection by the primary Ab (data not shown). Where inhibitors were used, cells were incubated for 1 h at 37°C with filipin (5 µg/ml), nystatin (50 µg/ml), monensin (50 µM), sucrose (0.4 M), or cycloheximide (10 µg/ml) before assays of receptor down-regulation were performed.

Recovery of cell surface receptor levels

Receptor down-regulation was initiated as described. After 30 min incubation with chemokines, the cells were washed three times in medium without FCS and resuspended in medium without FCS and incubated at 37°C. To remove CXCL11 from endogenous glycosaminoglycans, activated T lymphocytes were washed once in prewarmed 0.5 M NaCl/RPMI 1640 as previously described (27) then twice in RPMI 1640 and incubated at 37°C. Samples were taken at different time points and cells were washed in PBS buffer containing 1% FCS and 1% NaN3. Cells were stained with Abs as described. Where inhibitors were used, brefeldin A (5 µM), actinomycin D (5 µM), bafilomycin A1 (100 nM), or cycloheximide (10 µg/ml) were added to the cells during the recovery phase, following induction of CXCR3 down-regulation by ligand.

Confocal analysis

The H9 human T cell lymphoma line was washed in RPMI 1640 and resuspended at a concentration of 5 x 106 cells/ml in serum-free RPMI 1640 and were incubated at 37°C with 50 nM CXCL11. Samples were removed either before or at the indicated times following the addition of CXCL11. Internalization buffer (1% FCS, 1% NaN3 in PBS) was added to the samples removed, and tubes were incubated on ice until all time points were collected. Each time point was divided into either isotype control or Ab-staining tubes. Cells were washed twice in cold PBS before fixation in 4% paraformaldehyde for 20 min on ice. Cells were then washed in PBS and permeabilized in 0.5% saponin buffer containing anti-human lysosome-associated membrane protein (LAMP)-1 Ab (1/20 dilution; BD Pharmingen) or an equal concentration of mouse IgG1 isotype control. After incubation with the primary Ab, cells were washed in saponin buffer before incubation with goat anti-mouse IgG Alexa Fluor 568 (1/100 dilution in 0.5% saponin buffer; Invitrogen). Cells were washed and then preblocked with mouse IgG before being incubated with mouse anti-human CXCR3 FITC (1/10 dilution in 0.5% saponin; R&D Systems) or isotype control. Cells were then resuspended in 4% paraformaldehyde and were spun onto poly-L-lysine-coated glass coverslips in 24-well tissue culture plates at 1200 rpm for 5 min. The supernatant was removed and the coverslips washed twice in PBS and once in deionized water before being removed from the wells, allowed to air dry, and mounted onto slides in VectorShield Hardset fluorescence mounting medium (Vector Laboratories). Analysis was conducted by confocal microscopy using a Leica TCS NT confocal microscope with a x40 oil objective. Image analysis was conducted using Leica LCS Lite software version 2.61 and the images manipulated for presentation using Adobe Photoshop version 6.0.

SDS-PAGE and Western blot analysis

L1.2 transfectants expressing WT CXCR3, and CXCR3-AAA, CXCR3-K324R, CXCR3-{Delta}4, and CXCR3-{Delta}34 constructs were washed in RPMI 1640 and resuspended at 5 x 106 cells/ml in serum-free RPMI 1640 containing 10 µg/ml cycloheximide. Where indicated, 40 µM MG132 or 200 µM chloroquine was also added. The cells were preincubated for 30 min at 37°C before the addition of 50 nM CXCL11. Samples were taken at the indicated time points, the cells washed in ice-cold PBS and resuspended in lysis buffer containing 1% N-dodecyl-β D-maltoside, 10% glycerol, 1/1000 protease inhibitor cocktail in PBS (28). Equal quantities of cell lysates were separated on 4–12% SDS-PAGE gels and were electrophoretically transferred onto a nitrocellulose membrane, which was subsequently blocked with 5% milk in 0.01 M PBS with 0.05% Tween 20. The blots were independently probed with either anti-HA (1/1000 dilution; Covance) or anti-{alpha}-tubulin (1/10,000 dilution; Abcam) as a loading control. Following washing and probing with a secondary HRP-conjugated polyclonal goat anti-mouse Ig (1/1000 dilution), blots were developed by ECL (GE Healthcare).

Chemotaxis

Chemotaxis assays using either CXCR3 transfectants or purified T lymphocytes were performed essentially as previously described (10, 29) using ChemoTX plates with a 5-µm pore size, purchased from NeuroProbe. For T lymphocyte migration, enumeration was conducted using a hemocytometer and cell migration to buffer alone was subtracted from the resulting data, with individual results expressed as a percentage of the total cells applied to the filter. For L1.2 transfectant chemotaxis, the same apparatus was used, although at the end of the assay, cells were transferred from the lower chamber to a white 96-well microtiter plate using a funnel plate (NeuroProbe), and cells were detected with CellTiter Glo (Promega). Luminescence was measured using a TopCount microplate scintillation and luminescence counter (PerkinElmer). Data described are expressed as a chemotactic index, relative to migration observed to medium alone.

Ligand binding assays

125I-CXCL11 and 125I-CXCL10 were purchased from PerkinElmer Life Sciences. Ligand binding was performed as previously described using centrifugation through oil to separate bound chemokine from free chemokine (26). Data are presented following the subtraction of nonspecific binding, taken as the counts obtained when the labeled chemokine was displaced by a 1000-fold excess of homologous cold chemokine.

Data analysis

Data were analyzed using Prism 4.0 (GraphPad Software) by ANOVA with Bonferroni’s Multiple comparisons test.


   Results
Top
 Abstract
 Introduction
 Materials and Methods
 Results
Discussion
 Disclosures
 References
 
Down-regulation of cell surface CXCR3 following incubation with ligand

We initially used activated PBMCs to investigate the process of CXCR3 down-regulation following incubation of the cells with the three natural ligands described to date for CXCR3, namely CXCL9/Mig, CXCL10/IP-10, and CXCL11/I-TAC. Activated PBMCs, cultured for 7–10 days with Con A and IL-2, readily expressed CXCR3 on their cell surface as detected by flow cytometry using a specific mAb. Incubation of PBMCs with all three CXCR3 ligands induced a dose-dependent loss of CXCR3 from the cell surface (Fig. 1A) as deduced by staining with the same CXCR3-specific mAb.


Figure 1
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  		FIGURE 1. Internalization of CXCR3 by PBMCs and L1.2 transfectants. A and B, Dose-dependent nature and the respective kinetics of ligand-induced CXCR3 internalization in PBMCs as determined by flow cytometry using a specific anti-CXCR3 mAb. The ligands CXCL11 ({blacksquare}), CXCL10 (•) and CXCL9 ({blacktriangleup}) are shown. C and D, The levels of CXCR3 internalization in PBMCs and L1.2 transfectants, respectively, following incubation with 50 nM CXCL11 after pretreatment in the presence or absence of 0.4 M sucrose, 50 µM monensin, 5 µg/ml filipin, and 50 µg/ml nystatin. Cell surface CXCR3 levels were measured as described and the untreated control ({square}) is shown. ***, p < 0.001 compared with CXCL11 treatment alone. E, The effect of filipin pretreatment on the specific binding of 125I-CXCL11 to L1.2 CXCR3 transfectants. Data represent the mean ± SEM of at least three different experiments.

 
Notably, CXCL11 was the most efficacious ligand, with a 50 nM concentration of CXCL11 reducing cell surface staining to <20% of their starting levels. We subsequently examined the kinetics of this response, using 50 nM concentrations of each ligand (Fig. 1B). Loss of cell surface CXCR3 occurred rapidly, with optimal down-regulation observed by 30 min in agreement with a previous study (27). Similar data was also obtained using a previously described L1.2 cell line stably expressing CXCR3 (data not shown) with CXCL11 again the most efficacious of the ligands, although the maximum level of receptor down-regulation observed was reduced to ~50% of starting levels as we have previously observed with L1.2 cells expressing the related receptor CCR3 (30). This reduction likely reflects less efficacious coupling of the human receptor to murine intracellular machinery in the transfectants. In all subsequent experiments, we therefore incubated cells with 50 nM CXCL11 to achieve optimal CXCR3 down-regulation.

Pathways of CXCR3 internalization

Two major pathways are known by which chemokine receptors are internalized; either via clathrin-coated vesicles following the clathrin-mediated endocytic pathway or via caveolae. Hypertonic sucrose medium has been shown to block the assembly of clathrin-coated pits (31), whereas internalization via caveolae can be inhibited with filipin or nystatin (32). Monensin is an inhibitor of vesicle acidification, a process essential for the sorting events occurring during endocytosis of GPCRs such as the β2-adrenergic receptor (33). We assessed the activity of these inhibitors on CXCR3 down-regulation in PBMCs and L1.2 CXCR3 transfectants (Fig. 1, C and D). Neither filipin nor nystatin had any inhibitory effect on CXCR3 down-regulation in either cell type, suggesting that caveolae are not involved in the endocytosis of CXCR3. Although sucrose had little effect on ligand-induced CXCR3 down-regulation in PBMCs (Fig. 1C), it was observed to significantly reduce the levels of CXCR3 internalization in L1.2 CXCR3 cells following treatment with CXCL11 (Fig. 1D). In L1.2 transfectants, monensin treatment significantly reduced CXCL11-induced internalization of the receptor, suggesting that vesicular acidification is necessary for the sorting events occurring following CXCR3 endocytosis. In PBMCs, monensin had a modest inhibitory effect on CXCR3 endocytosis that did not reach statistical significance. Because it has been previously demonstrated that cholesterol and lipid rafts are required for the maintenance of chemokine receptor conformation (34, 35), we also sought to examine the effects of filipin and nystatin on ligand binding. Although nystatin treatment altered the density of CXCR3 transfectants making them unable be centrifuged through oil in our ligand binding assay (data not shown) treatment of cells with filipin was observed to have little effect on CXCL11 binding (Fig. 1E).

CXCR3 can be internalized independently of clathrin and β-arrestin 1 and β-arrestin 2

Because sucrose and monensin were without effect on CXCR3 internalization in PBMCs we sought to confirm our findings by using an alternative strategy to inhibit clathrin. T lymphocytes were purified from blood and underwent nucleofection either in buffer alone (mock nucleofection) or in buffer containing plasmid encoding a GFP-tagged construct, DIII (DIII transfection). This construct inhibits clathrin-coated pit assembly and therefore clathrin-dependent internalization (36, 37). Twenty-four hours after nucleofection, cells were harvested and incubated at 37°C either in the presence or absence of CXCL11 before staining for CXCR3 expression. A significant percentage of mock-nucleofected T lymphocytes were shown to express CXCR3 (Fig. 2A, top left quadrant), which was seen to be reduced following CXCR3 treatment (Fig. 2C, top left quadrant). Nucleofection of T lymphocytes led to the identification of two populations of CXCR3-positive cells, a major population not expressing the DIII-GFP fusion protein (Fig. 2B, top left quadrant) and a minor population expressing the DIII-GFP fusion protein (Fig. 2B, top right quadrant). CXCL11 treatment was seen to significantly reduce the number of cells within both populations (Fig. 2D, top right and left quadrants). A similar lack of effect upon CXCL11-induced CXCR3 endocytosis was also seen following nucleofection of T lymphocytes with the control protein GFP-DIII{Delta}2, which corresponds to the GFP-DIII construct lacking all AP-2 binding sites (data not shown). Collectively this suggests that ligand-driven endocytosis of CXCR3 in T lymphocytes occurs independently of clathrin.


Figure 2
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  		FIGURE 2. Neither clathrin nor arrestins are required for the CXCL11-induced internalization of CXCR3. A–D, CXCL11-induced internalization of CXCR3 in purified T lymphocytes as deduced by flow cytometry following nucleofection in the presence or absence (mock) of a plasmid containing a dominant negative GFP-tagged DIII construct that inhibits clathrin-coated pit assembly. Nucleofection was conducted 24 h before the induction of CXCR3 internalization with 50 nM CXCL11. The percentage of cells is shown in each quadrant and the experiment shown is representative of three separate experiments. E, The relative internalization of CXCR3 in WT MEF cells ({square}) and β-arrestin 2- or β-arrestin 3-deficient MEF cells ({blacksquare}), transiently transfected with CXCR3, following incubation with 50 nM CXCL11 or buffer alone. Data represent the mean ± SEM of four different experiments. F, The relative internalization of CXCR3 in response to 50 nM CXCL11 or buffer alone in a stable L1.2 transfectant cell line following transfection 24 h previously with either plasmid pcDNA3 (mock) or pcDNA3 containing the V53D and V54D dominant negative mutants of β-arrestin 1 and β-arrestin 2. Data represent the mean ± SEM of three different experiments **, p < 0.01 and ***, p < 0.001 compared with buffer-treated controls.

 
The clathrin-dependent pathway for endocytosis of GPCRs typically involves the binding of arrestins to the intracellular face of the phosphorylated receptor. To examine whether CXCR3 internalization is dependent upon arrestins, we transiently transfected MEFs obtained from both WT and β-arrestin 1- and β-arrestin 2-deficient mice. Internalization was then induced by incubation with CXCL11 and CXCR3 down-regulation assessed as before by flow cytometry. CXCR3 down-regulation in WT MEFs was similar to that seen in L1.2 transfectants, with CXCL11 reducing cell surface levels to around 50% of their starting levels (Fig. 2E). In MEFs from β-arrestin 1- and β-arrestin 2-deficient mice, CXCR3 down-regulation in response to ligand was observed, but at a reduced level, with only a 20% reduction of CXCR3 cell surface levels in response to CXCL11, suggestive of an incomplete requirement for β arrestin in the down-modulation process. Similarly, transfection of L1.2 CXCR3 transfectants with either empty plasmid or plasmids encoding the V53D and V54D dominant form of β-arrestin 1 and β-arrestin 2 were without effect upon CXCR3 down-regulation induce by CXCL11 (Fig. 2F). Collectively, the data suggest the existence of a β-arrestin-independent pathway for the endocytosis of CXCR3.

Inhibition of CXCR3 cell surface replenishment

After the induction of down-regulation by ligand, the recovery of cell surface CXCR3 levels was relatively slow in both PBMCs and L1.2 transfectants (Fig. 3, A and B), with only ~70–80% recovery of the original CXCR3 cell surface levels observed a full 3 h after incubation with CXCL11. This was in contrast to another Th1-expressed chemokine receptor CXCR6, which showed 100% recovery of cell surface levels within 1 h of ligand-induced down-regulation (Fig. 3C) and is typical of receptor recycling to the cell surface as described for other chemokine receptors (19, 38, 39). The slow recovery of cell surface CXCR3 levels suggested to us that upon ligand-induced internalization, CXCR3 is either slowly recycled, as is the case for class B GPCRs such as the vasopressin type 2 receptor (40) or alternatively, is degraded. In the case of degradation, cell surface replenishment would therefore require de novo synthesis of receptor.


Figure 3
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  		FIGURE 3. CXCR3 cell surface replenishment is dependent upon de novo protein synthesis. Activated PBMCs (A), L1.2 CXCR3 transfectants (B), and L1.2 CXCR6 transfectants (C) were incubated for 1 h in serum-free medium in the presence ({blacksquare}) or absence ({circ}) of 10 µg/ml cycloheximide. Following incubation with CXCL11 (PBMCs and CXCR3 transfectants) or CXCL16 (CXCR6 transfectants), cells were further incubated with cycloheximide and samples taken at the indicated time points for the determination of cell surface expression of CXCR3 by flow cytometry. Data represent the mean ± SEM of at least three independent experiments. **, p < 0.01 and ***, p < 0.001 compared with cycloheximide-treated cells.

 
To examine this latter hypothesis, we preincubated the cells for 1 h with cycloheximide, induced CXCR3 internalization with CXCL11, and let the cells recover in the presence of cycloheximide. Treatment with cycloheximide ablated the recovery of CXCR3 in both PBMCs and L1.2 transfectants, whereas recovery of cell surface CXCR6 levels to 80% of the starting levels was observed at the 2-h point. This latter value was approximately half of the level of staining seen at the same time point with CXCR6 transfectants that had not been treated with cycloheximide, suggesting that both recycling and de novo synthesis cooperate in maintaining CXCR6 cell surface levels in the transfectant system used (Fig. 3C).

Thus, in contrast to CXCR6 and other chemokine receptors described in the literature (19, 20, 41), cell surface replenishment of CXCR3 is dependent upon de novo protein synthesis. If this postulate is true, then CXCR3 cell surface replenishment should also be dependent upon mRNA transcription and efficient transport through the endoplasmic reticulum (ER) and Golgi. We therefore preincubated PBMCs or L1.2 transfectants for 1 h in the presence or absence of actinomycin D (an inhibitor of transcription), or brefeldin A and bafilomycin A1, which have been shown to inhibit function of the ER and Golgi apparatus, respectively, and therefore inhibit transport of receptors through these compartments (41, 42). Internalization was induced with 50 nM CXCL11 and the expression of CXCR3 was monitored at 3 h post-internalization. In PBMCs, cell surface replenishment of CXCR3 was also significantly inhibited, although not reduced to basal levels (Fig. 4A). In L1.2 transfectants, cell surface CXCR3 levels remained at baseline following incubation with CXCL11 in the presence of actinomycin D, brefeldin A, or bafilomycin A1 (Fig. 4B). Collectively, the data suggest that the observed recovery of CXCR3 at the cell surface is dependent upon newly synthesized receptor trafficking through functional Golgi apparatus in the cell, in contrast to chemokine receptors such as CCR4, CCR5, and CXCR6, which appear to be replenished by a recycling mechanism (19, 20).


Figure 4
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  		FIGURE 4. Cell surface expression of CXCR3 is dependant upon functional ER and Golgi. Activated PBMCs (A) and L1.2 CXCR3 cells (B) were incubated with 50 nM CXCL11 for 30 min, washed three times with serum-free medium, and then incubated in serum-free medium for up to 3 h in the presence or absence of actinomycin D (5 µM), brefeldin A (5 µM), and bafilomycin A1 (100 nM). Cell surface CXCR3 was assessed by flow cytometry. The extent of CXCR3 internalization following the initial 30 min incubation with CXCL11 ({square}) and CXCR3 cell surface level recovery ({blacksquare}) of an untreated control after 3 h are presented. Data represent the mean ± SEM of at least three independent experiments. ***, p < 0.001 compared with untreated control.

 
The role of the CXCR3 C terminus in internalization

Because the C terminus of several GPCRs has been implicated in the internalization process, we sought to examine the role of this motif in the internalization of CXCR3. Site-directed mutagenesis of the CXCR3 cDNA was performed to generate four mutant constructs (Fig. 5A). The first of these mutated a triple LLL motif to AAA, thereby losing two potential LL motifs previously reported to be involved in CXCR2 internalization (43). The second mutation targeted the sole intracellular lysine residue, K324. Ubiquitination of internalized GPCRs has been shown to target them for degradation, a process whereby the 74 aa ubiquitin is covalently attached to intracellular lysine residues. The two remaining mutations introduced premature stop codons within the cDNA, truncating the receptor by either 4 aa ({Delta}4 construct) or 34 aa ({Delta}34 construct). These mutations removed a YXXL motif at the extreme C terminus and the entire repertoire of C-terminal serine and threonine residues, respectively. The latter construct allowed us to examine the requirement for phosphorylation of CXCR3 in the internalization process. All four mutants were transiently expressed in L1.2 cells, and cell surface expression was monitored by flow cytometry. All four mutants trafficked to the cell membrane, although the {Delta}34 mutant was expressed at levels significantly below those of WT CXCR3 (Fig. 5B). Conversely, the {Delta}4 mutant was consistently expressed at greater levels than found in WT CXCR3, although this did not reach significance. All four constructs were able to mediate chemotaxis of cells in response to CXCL11, with the typical bell-shaped responses optima around the 3 nM concentration (Fig. 5C). Likewise, internalization of CXCR3 in response to CXCL11 was unimpaired by mutation, with the 50 and 100 nM concentrations of ligand inducing significant internalization compared with untreated cells (Fig. 5D). Because the {Delta}4 construct appeared to be expressed at higher levels than WT CXCR3 (Fig. 5B), we postulated that CXCR3 might be internalized constitutively, i.e., in the absence of ligand, and the loss of the four most C-terminal residues might inhibit this process. We subsequently examined the expression of both WT CXCR3 and the {Delta}4 construct over a 6-h period, following pretreatment with cycloheximide to inhibit de novo synthesis. WT CXCR3 was seen to be quite rapidly lost from the cell surface in the absence of ligand, with approximately half of the original cell surface levels of CXCR3 observed after 4 h of incubation. In comparison, internalization of the {Delta}4 construct was less efficacious, with the remaining cell surface levels of mutant receptor at the 6-h time point significantly greater than receptor levels of WT CXCR3.


Figure 5
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  		FIGURE 5. The effects of C-terminal mutation upon CXCR3 function. A, The amino acid identity of the C-terminal CXCR3 mutants analyzed. B, The relative expression levels of the constructs following the transient transfection of L1.2 cells, compared with cell surface staining observed with the WT CXCR3 construct. *, p < 0.001. C, The relative chemotactic responses to CXCL11 of the same CXCR3 mutants. D, depicts the internalization of the CXCR3 mutants in response to increasing concentrations of CXCL11. Comparisons were made with untreated transfectants in each case.*, p < 0.001 and ***, p < 0.01 compared with controls. E, Constitutive internalization in the absence of ligand of both WT CXCR3 and the {Delta}4 construct over a 6-h period following treatment with cycloheximide to inhibit de novo synthesis of receptor. All data represent the mean ± SEM of at least three experiments.

 
Endocytosed CXCR3 passes through late endosomes or lysosomes before its degradation

We subsequently turned our attention to the fate of CXCR3 following its internalization, using confocal microscopy to examine intracellular localization of the receptor in permeablized T lymphocytes. A predominantly granular intracellular staining pattern for CXCR3 was evident in untreated cells (Fig. 6A), identical with that previously described by Gasser and colleagues (44). Likewise, a similar pattern was seen for staining with the late endosome marker LAMP-1 (Fig. 6B) with little colocalization of signals seen (Fig. 6C). Treatment with CXCL11 for 15 min resulted in clustering of LAMP-1+ vesicles with apparent colocalization of CXCR3 with LAMP-1 in some but not all cells (Fig. 6, E and F). This result may reflect either a rapid loss of CXCR3 immunoreactivity following trafficking to lysosomes or the fact that this pathway is not the sole route of CXCR3 degradation. Little, if any, colocalization of CXCR3 with LAMP-1 staining was observed in cells 60 min following treatment with CXCL11, suggesting that degradation of CXCR3 may be complete by this point (Fig. 6, G–I).


Figure 6
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  		FIGURE 6. Internalized CXCR3 traffics to late endosomes. T lymphocytes were processed for confocal microscopy either in the absence of treatment (A–C) or following incubation with CXCL11 for 15 min (D–F) or 60 min (G–I). Following processing, T lymphocytes were stained with Abs directed against CXCR3 (green) and LAMP-1 (red) before analysis by confocal microscopy. A, D, and G show the CXCR3 signal. B, E, and H show the LAMP-1 signal. C, F, and I show the two signals overlaid. Scale bar represents 50 µm. F, Colocalization of CXCR3 and LAMP-1 signals in cells is highlighted (arrowheads). Data shown are representative of at least three independent experiments.

 
To further examine CXCR3 degradation, we used Western blotting methodologies. HA-tagged CXCR3 was expressed transiently in L1.2 cells and following pretreatment with cycloheximide to inhibit de novo synthesis, the cells were incubated for varying time periods in the presence or absence of CXCL11. Cell lysates were then examined by SDS-PAGE, followed by Western blotting. As can be seen in Fig. 7A, CXCR3 appears as a band of ~50 kDa. Following 3 h of incubation at 37°C, either in buffer alone or supplemented with CXCL11, the band representing CXCR3 was seen to reduce considerably in intensity, suggestive of a degradative fate. Additional pretreatment of CXCR3 transfectants with either the proteosome inhibitor MG132 or the lysosomal inhibitor chloroquine, before treatment with CXCL11, was observed to inhibit the degradative process (Fig. 7B). We subsequently examined the panel of four C-terminal CXCR3 mutants to examine the effects of mutation upon degradation. Compared with untreated cells, obvious degradation of each construct was observed, suggesting that none of the C-terminal motifs we examined are critical for targeting CXCR3 for degradation (Fig. 7C). Thus it appears that CXCR3 is readily degraded in the presence or absence of ligand, by pathways involving both the proteosomes and lysosomes, and that ubiquitination of CXCR3 is not a fundamental part of this process.


Figure 7
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  		FIGURE 7. CXCR3 is degraded following constitutive or CXCL11-induced internalization. L1.2 cells transiently transfected with plasmids encoding CXCR3 were preincubated for 30 min at 37°C with 10 µg/ml cycloheximide and an aliquot was reserved (0 h time point). Incubation of the remaining cells was then allowed to proceed in the presence or absence of 50 nM CXCL11 for 3 h. Cell lysates were generated and analyzed by Western blotting using anti-HA mAb (top). Blots were subsequently stripped and reprobed with an anti-{alpha}-tubulin (aT) Ab as a loading control (bottom). A, Both constitutive (untreated) and CXCL11-induced degradation of WT CXCR3 over the 3-h time period as deduced by a loss of immunoreactivity. B, The effects of preincubation of either 40 µM MG132 or 200 µM chloroquine on CXCL11-induced degradation. C, The CXCL11-induced degradation of WT CXCR3 and C-terminal CXCR3 mutants. Data shown are from one experiment representative of three different experiments.

 
Previous reports have detailed the strict control of CXCR3 mRNA expression in freshly isolated PBMCs. Although a significant proportion of freshly isolated PBMCs express CXCR3 at the cell surface, mRNA transcripts remain undetectable and the cells are unresponsive to CXCL9 and CXCL10 in assays of chemotaxis and calcium flux (1, 2). This unresponsive phenotype is reversed following culture of PBMCs for several days in a medium containing IL-2 and a mitogen such as PHA, and correlates with mRNA induction and increased levels of CXCR3 at the cell surface. We revisited these data using freshly isolated T lymphocytes used either immediately after isolation or following culture for 10 days in medium supplemented with IL-2 and Con A. As previously described for PBMCs, freshly isolated T lymphocytes expressed modest levels of cell surface CXCR3, which were significantly up-regulated following 10 days of culture (Fig. 8A). Modest numbers of freshly isolated cells were seen to migrate in response to increasing concentrations of CXCL9 and CXCL10, responses which were significantly enhanced following 10 days of culture (Fig. 8, B and C). In contrast, chemotactic responses of both freshly isolated and cultured T lymphocytes to CXCL11 were robust, notably at the optimal concentration of 10 nM (Fig. 8D). The greater efficacy of CXCL11 on freshly isolated cells was also evident when internalization assays were performed, with CXCL11 but neither CXCL9 nor CXCL10 inducing significant internalization of CXCR3 (Fig. 8E). Radioligand binding assays competing 125I-CXCL10 and 125I-CXCL11 with homologous unlabelled ligand (Fig. 8F) suggested a greater number of binding sites for CXCL11 on freshly isolated T lymphocytes than were evident for CXCL10, despite both ligands reporting to have similar nanomolar affinities at CXCR3 (8, 10, 45). Culture for 10 days in the presence of IL-2 and Con A resulted in a trend for an increase in the number of binding sites for both ligands, although this increase was not statistically significant. Thus, despite freshly isolated cells expressing CXCR3 at the cell surface, the responses to CXCR3 ligands appear to be muted, with only CXCL11 inducing significant biological function.


Figure 8
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  		FIGURE 8. CXCR3 function in T lymphocytes is strictly controlled. A–F, A series of experiments to compare CXCR3 expression and function in freshly isolated T lymphocytes (day 0) and T lymphocytes cultured for 10 days in medium supplemented with IL-2 and Con A (day 10). A, The relative expression levels of CXCR3 on T lymphocytes as deduced by flow cytometry using an anti-CXCR3 Ab. **, p < 0.01. B–D, The relative chemotactic responses of T lymphocytes to increasing concentrations of CXCL9, CXCL10, and CXCL11. E, The relative induction of CXCR3 internalization on freshly isolated T lymphocytes as deduced by flow cytometry, following incubation with 50 nM of CXCL9, CXCL10, or CXCL11. *, p < 0.05 compared with controls. F, The relative levels of specific binding of 0.1 nM 125I-CXCL10 and 125I-CXCL11 by T lymphocytes. All data represent the mean ± SEM of at least three experiments.

 

   Discussion
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
Disclosures
 References
 
Although there is growing information regarding the mechanisms of GPCR internalization and recycling, no data concerning the fate of internalized CXCR3 have been published to date. Once internalized, a GPCR can experience one of two fates, namely dissociation of ligand and recycling of functional receptor back to the plasma membrane or degradation. These fates are not mutually exclusive, as CXCR4 has been shown to undergo both processes following engagement with ligand (46, 47). In this study, we provide several lines of evidence to suggest that the fate of endocytosed CXCR3 is predominantly one of degradation, with de novo synthesis of CXCR3 required for the recovery of CXCR3 cell surface levels. Indeed, little if any CXCR3 was seen to reappear at the cell surface when transcription, translation, or the trafficking of nascent proteins through the golgi-ER was perturbed with appropriate pharmacological inhibitors.

Although the related receptor CXCR4 has been shown to undergo lysosomal degradation in a ubiquitin-dependent manner (47, 48), both the lysosome and the proteosome appear to facilitate the degradation of CXCR3, as deduced by sensitivity to both MG132, a 26 S proteasome inhibitor (49), and chloroquine, an inhibitor of intralysosomal catabolism (50). Taken in the context of our confocal microscopy data, we suggest that internalized CXCR3 traffics to late endosomes or lysosomes that may communicate in part with the proteosome for CXCR3 degradation. Cooperation between both the lysosomal and proteosomal pathways has been described for the degradation of other receptors including the growth hormone receptor (51) and the IL-2R/IL-2R ligand complex (52). In the case of the growth hormone receptor, endocytosis occurs in the absence of receptor ubiquitination but still requires intact proteasomal activity, suggesting that an adaptor protein targets the receptor to the proteosome (53). In the case of CXCR3, ubiquitination of the receptor appears not to be required for either internalization or degradation, as mutation of the sole intracellular lysine residue had little effect upon either process. This may suggest the existence of an additional motif within the CXCR3 intracellular regions that targets it for degradation by this route. Alternatively, CXCR3 may be envisaged to interact with an adapter protein, which itself undergoes ubiquitination, targeting both proteins to the proteosome. Such a process has been described for the lectin Siglec-7, which is targeted to the proteosome as a complex with SOCS-3 (54).

Two main routes have been described for the internalization of GPCRs following their activation. The best-characterized pathway uses clathrin-coated pits. In this pathway the phosphorylated receptor is bound by arrestins and located to clathrin-coated pits, where the complex is internalized in vesicles. These vesicles are subsequently released from the plasma membrane by dynamin and transported to endosomes, where dephosphorylation of the receptor occurs and resensitized receptor is recycled to the plasma membrane (55). The clathrin-mediated pathway has been demonstrated for the internalization of other chemokine receptors of the CXC class, notably CXCR1 (56), CXCR2 (57), and CXCR4 (38). A second pathway of internalization depends on caveolae (58), cholesterol rich, highly organized membrane structures that have been shown to be involved in the internalization of other GPCRs including the chemokine receptors CCR4 and CCR5 (19, 20). Although caveolae have been described in macrophages (59), there is still some debate as to whether lymphocytes contain caveolae (60, 61). Evidence for the use of either pathway of receptor internalization is often provided through the overexpression of dominant negative constructs (e.g., arrestin, dynamin, and clathrin mutants) or the use of pharmacological inhibitors to invoke or preclude the use of a particular pathway (62). In both human PBMCs and an established transfectant system expressing the human ortholog of CXCR3 (10), cell surface levels of CXCR3 were rapidly reduced in a concentration- and time-dependant manner following exposure to ligand. In CXCR3 transfectants, use of inhibitors suggested that the pathway mediating ligand-induced endocytosis did not appear to involve caveolae but involved clathrin. In contrast, treatment of activated PBMCs with hypertonic sucrose did not inhibit the internalization of CXCR3, and the use of an inhibitor of clathrin-coated pit assembly had no effect upon the down-regulation of CXCR3 in purified T lymphocytes. Likewise, we found no absolute requirement for arrestin in the internalization process. In MEFs from mice deficient in β-arrestins 1 and 2 (24), internalization of CXCR3 was significantly reduced, but not completely abolished, whereas the use of dominant negative arrestin mutants was without effect upon CXCR3 internalization in our transfectant system. Collectively, our results suggest an alternative pathway for the endocytosis of CXCR3, one that is independent of clathrin or arrestin or a combination. This finding is in agreement with a previous study in which CXCL11-induced internalization of CXCR3 in a transfectant system was found to occur in a dynamin and β-arrestin 2-independent manner (28).

The cellular motifs controlling ligand-driven internalization and targeting it for subsequent degradation remain elusive. Removal of potential phosphorylation sites in the C terminus by truncation had no effect on CXCL11-induced internalization, as previously described for CXCR3 transfectants in both HEK-293 and 300-19 cell lines (28, 63). Likewise, mutation of the LLL motif was without effect on CXCL11-induced internalization again in agreement with a study using 300-19 transfectants (28) but in disagreement with a study using HEK-293 transfectants in which some inhibition of CXCL11-induced internalization was observed (63). This likely reflects differences in the intracellular machinery to which CXCR3 is coupled in either cell system.

Of interest was the finding that CXCR3 is constitutively degraded in the absence of ligand, a robust process that was mediated to a significant extent by a canonical YXX{varphi} motif at the extreme of the C terminus. Because such motifs have been implicated in the sorting of transmembrane proteins to endosomes and lysosomes (64), we hypothesize that the YSGL motif interacts with currently unknown intracellular proteins and controls the constitutive internalization of CXCR3. Supportive of our hypothesis, a distal YKKL motif within the C terminus of the GPCR PAR1 directs constitutive receptor internalization that is clathrin- and dynamin-dependent but independent of arrestins (65, 66).

Cell surface levels of CXCR3 are tightly regulated by both constitutive and ligand-driven degradation and the replenishment of cell surface CXCR3 appears not to be dependent upon recycling as has been shown for other chemokine receptors, but upon de novo synthesis of CXCR3 protein and its subsequent transportation through the Golgi apparatus. To our knowledge, this is the first example of a GPCR in which protein synthesis is essential for the replenishment of the receptor on the cell surface following stimulation with ligand. This strict control is in addition to other mechanisms of posttranslational regulation of CXCR3 function. Although expressing significant amounts of CXCR3 on the cell surface, freshly isolated T lymphocytes were poorly responsive to the CXCR3 ligands CXCL9 and CXCL10 as previously described (2), with the exception of the ligand CXCL11. This phenotype was corrected upon activation of the T lymphocytes by prolonged incubation with IL-2 and a mitogen such as Con A, a process that corresponds with CXCR3 mRNA induction (2), increased cell surface expression of the receptor, and the acquisition of robust functional responses to all three ligands. CXCL10 and CXCL11 have previously been described as allotopic ligands of CXCR3, with activated T cells expressing a significant population of CXCR3 molecules that can bind 125I-CXCL11 but not 125I-CXCL10 (45). The binding of CXCL10 is thought to be controlled at the level of G protein coupling because treatment of cell membranes with GTP{gamma}S (guanosine 5'-O-(3-thiotriphosphate) or pertussis toxin resulted in a total loss of CXCL10 binding. In contrast, CXCL11 can bind to both coupled and uncoupled CXCR3 (45). Supportive of this idea, both resting and activated T lymphocytes were observed to bind significantly more 125I-CXCL11 than125I-CXCL10. Recently published data from studies using mice deficient in the G protein {alpha} subunits G{alpha}i2 and G{alpha}i3 found that although T lymphocytes from mice lacking G{alpha}i2 subunits exhibited no chemotaxis to CXCR3 ligands, T lymphocytes from mice lacking G{alpha}i3 displayed significant increases in both migration and GTP{gamma}S binding and migration as compared with WT T lymphocytes (67). This suggests that in mice, G{alpha}i2 subunits are crucial for CXCR3 signaling, and that G{alpha}i3 subunits can act as intracellular inhibitors of CXCR3 function, thereby modulating CXCR3 responsiveness. Examining our findings in the light of these data, we can hypothesize that up-regulation of CXCR3 itself does not necessarily result in responsiveness to CXCL10 and that CXCR3 function in the human is likely be modulated at the intracellular level by interaction with G proteins.

CXCR3 has previously been reported to be expressed in an intracellular compartment within T lymphocytes, which can rapidly be mobilized to the cell surface by treatment with arachidonic acid (44). This rapid, transient mobilization of receptor has been postulated to enable the cells to respond timely to changes in the microenvironment in vivo. Such a capacity for increased cell surface expression is likely to be counterbalanced by the degradative fate of CXCR3 we describe in this study. It is noteworthy that the CXCR3 ligands are all readily induced by IFN-{gamma} (7, 8, 9) and that the Th1-polarized lymphocytes specifically attracted by these chemokines are themselves a source of IFN-{gamma} (68). It can be postulated that such fine tuning of CXCR3 activity by degradation of internalized receptor serves to avoid the unnecessary amplification of T lymphocyte recruitment in vivo, which would have undesirable consequences for the host. Generation of an artificial CXCR3 ligand that promotes the cellular degradation of CXCR3 in the absence of intracellular signaling may represent an alternative strategy for the therapeutic modulation of CXCR3 with potential benefit in a wide variety of disease processes.


   Acknowledgments
 
We are grateful to Dr. Robert Lefkowitz for providing the WT and β-arrestin-deficient MEF, to Dr. Marc Caron for the provision of plasmids encoding arrestin mutants, and to Dr. Alexandre Benmerah for providing plasmids encoding the DIII and DIII{Delta}2 constructs. We thank Professor Mark Marsh, University College London, and Dr. Richard Colvin, Massachusetts General Hospital, for helpful discussions.


   Disclosures
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 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 Disclosures
References
 
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 Grants PG/2000055 and FS/05/021 from the British Heart Foundation, Grant 174240 from the Arthritis Research Campaign, and Project Grant 076036/Z/04/Z from the Wellcome Trust. Back

2 A. Meiser and A. Mueller contributed equally to the study. Back

3 Current address: School of Chemical Sciences and Pharmacy, University of East Anglia, Norwich NR4 7TJ, U.K. Back

4 Address correspondence and reprint requests to Dr. James E. Pease, Leukocyte Biology Section, Faculty of Medicine, National Heart and Lung Institute, Sir Alexander Fleming Building, Imperial College London, South Kensington Campus, London SW7 2AZ, U.K. E-mail address: j.pease{at}imperial.ac.uk Back

5 Abbreviations used in this paper: GPCR, G protein-coupled receptor; HA, hemagglutinin; LAMP, lysosome-associated membrane protein; MEF, mouse embryonic fibroblast; ER, endoplasmic reticulum; WT, wild type. Back

Received for publication March 14, 2008. Accepted for publication March 14, 2008.


   References
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 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 Disclosures
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