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. 2006 Dec 1;119(4):551–561. doi: 10.1111/j.1365-2567.2006.02470.x

The strength of the chemotactic response to a CCR5 binding chemokine is determined by the level of cell surface CCR5 density

Caroline Desmetz 1,2, Yea-Lih Lin 1, Clément Mettling 1, Pierre Portalès 2, Herisoa Rabesandratana 3, Jacques Clot 2, Pierre Corbeau 1,2
PMCID: PMC2265826  PMID: 17177831

Abstract

We have shown that the intensity of expression of the C-C chemokine receptor CCR5 at the single CD4+ cell level strongly determines the efficiency of its function as a coreceptor for human immunodeficiency virus type 1. By analogy, we examined if the number of CCR5 molecules at the cell surface might determine its chemotactic response to CCR5 ligands. To test this hypothesis, we measured by flow cytometry the migration of primary human T cells towards the CCR5-binding chemokine CCL5 in vitro. First, we observed a dose-dependent blockage of this migration exerted by an anti-CCR5 monoclonal antibody. Second, we sorted peripheral blood mononuclear cells into five subpopulations expressing various cell surface CCR5 densities, and observed a correlation between the intensity of migration towards CCL5 and the level of CCR5 expression on these subpopulations. Third, we transduced CCR5+ peripheral blood mononuclear cells with the CCR5 gene, and observed that the CCR5 over-expression induced an over-migration towards CCL5. Finally, we observed in healthy donors a correlation between the chemotactic response of peripheral blood CD8+ T cell to CCL5 and their level of surface CCR5 expression. T-cell surface CCR5 density, which is constant over time for a given individual, but varies drastically among individuals, might therefore be an important personal determinant of T-cell migration in many biological situations where CCR5-binding chemokines play a role, such as graft rejection, T helper 1-mediated auto-immune diseases, and infectious diseases involving CCR5. Moreover, our data highlight the therapeutic potential of CCR5 antagonists in these situations.

Keywords: T lymphocytes, AIDS, chemokines, chemotaxis, CCR5

Introduction

CCR5 (C-C chemokine receptor 5) is a chemokine receptor able to bind C-C chemokines, including macrophage inflammatory protein (MIP)-1α (CCL3), MIP-1β (CCL4), and RANTES (regulated on activation, normal, T-cell expressed, and secreted; CCL5).1 Chemokines are grouped into subfamilies based on the position of conserved cysteine residues in the N-terminal part of the protein and exert their action through seven transmembrane G-protein coupled receptors. They regulate leucocyte trafficking to normal and inflamed tissues2. In several inflammatory conditions, CCR5-expressing immune cells are recruited from the blood across the vascular endothelium. Consequently, in many pathological situations, such as rheumatoid arthritis (RA), multiple sclerosis, graft rejection, CCR5 expressing cells, mainly effector T cells, are found in great numbers into the inflammation sites, where CCR5 binding chemokines are produced.3–9 They participate in the inflammatory reaction, thus maintaining and potentially worsening the situation. Besides CCL5, the CCR5 ligand the most abundant in the plasma in physiological conditions, which is produced by CD8+ T cells, natural killer cells, epithelial cells, fibroblast and platelets, is a particular feature of inflammation. Increased CCL5 expression has been associated with a wide range of inflammatory disorders and pathologies, including RA, allogeneic transplant rejection, atherosclerosis, atopic dermatitis, inflammatory airway disorders (such as asthma), some neurological disorders (such as Alzheimer's disease) and malignancies. It is thought to act by promoting leucocyte infiltration to sites of inflammation.10

In addition to its chemotactic function, CCR5, as well as CXCR4, also serves as a coreceptor for the so-called R5 human immunodeficiency virus type 1 (HIV-1) strains.11 We have previously shown that the number of CCR5 molecules at the surface of a target cell drastically determines its infectibility by R5 HIV-1 strains.12 Thus, a sevenfold difference in cell surface CCR5 density results in a 60-fold difference in virus production after a single HIV life cycle. Moreover, our previous work in HIV patients showed that CD4+ T-cell surface CCR5 density is correlated with viral load13 and disease progression.14

Interestingly, we have shown that CD4+ T-cell surface CCR5 density is constant over time for a given individual, but varies from 4000 to over 24 000 molecules per cell among individuals.13 CCR5-Δ32 mutation is a 32 bp deletion in the coding region of the CCR5 gene resulting in the production of a truncated CCR5 molecule that is not expressed at the cell surface.15,16 CCR5-Δ32 homozygotes do not express the receptor at the cell surface, while heterozygous individuals for CCR5-Δ32 express intermediate levels of CCR5 on their T-cell surface.15,16 In RA patients, the frequency of the CCR5-Δ32 allele is reduced.17–19 Moreover, heterozygous individuals develop less aggressive disease than homozygous wild type CCR5 gene carriers.19–21 In patients with Sjögren's syndrome, the frequency of Δ32/CCR5 genotype is significantly decreased and suggests that carrier status for the CCR5-Δ32 allele may contribute to protection from the development of this disease.22

We hypothesized that, if the level of expression of the CCR5 receptor at the surface of T cells determines their migratory capacity in response to CCR5 ligands, there might be a polymorphism in the capacity of individuals to respond to C-C chemokines, and thereby in the occurrence and/or the course of the diseases where these chemokines are involved. Therefore, we wondered whether the functional efficiency of CCR5 function as a chemokine receptor might be influenced by its cell surface density, and tested this hypothesis on human peripheral blood T cells.

Materials and methods

Cell culture and reagents

Peripheral blood mononuclear cells (PBMC) from healthy donors were isolated by Ficoll-Paque density centrifugation and cultured at 2 × 106 cells/ml in RPMI-1640 medium supplemented with 2 mm glutamax-1, 10% fetal calf serum (FCS), 100 U/ml penicillin and 100 µg/ml streptomycin (Gibco, Grand Island, NY). Recombinant human CCL5/RANTES was purchased from R & D Systems (Minneapolis, MN). Purified anti-human CCR5 antibody, clone 2D7, was purchased from PharMingen (Becton Dickinson, San Jose, CA). Anti-CCR5 monoclonal antibody (mAb) PA12 was purchased from Progenics Pharmaceuticals, Inc (Tarrytown, NY, USA). Fluorescein isothiocyanate (FITC)-conjugated anti-CD4 mAb and anti HLA-DR (anti-CD4 FITC and anti-HLA-DR FITC), phycoerythrin (PE)-conjugated anti-CD8 and anti-CD3 mAb (anti-CD8 PE and anti-CD3 PE), phycoerythrin cyanin 5-conjugated anti-CD69 (anti-CD69 PC5), and their respective isotype controls were purchased from Beckman-Coulter (Roissy, France). FITC-conjugated goat anti-mouse (GAM) immunoglobulin probe (H + l) (GAM FITC) was purchased from Jackson (West Grove, PA). Fluorescent beads for the cytometry quantification were purchased from Beckman-Coulter.

Chemotaxis assay

The chemotaxis of peripheral blood lymphocytes (PBL) in response to recombinant human CCL5 was measured across 3 µm pore-size cell culture inserts incorporating polyethylene terephthalate membranes (Falcon, Becton Dickinson) in 24-well companion plates (Falcon cell culture insert system). Five hundred µl of culture medium (RPMI-1640 as described above), supplemented or not with recombinant human CCL5 were placed in the lower chamber of the plate. Peripheral blood from healthy volunteers was collected into an ethylenediamine tetra-acetic acid (EDTA) tube, diluted 1 : 5 in culture medium and placed into culture inserts (total volume of 100 µl per insert). The plates were then incubated for 4 hr at 37° (5% CO2). Then, the cells that had crossed the membrane were collected from the lower chamber, pelleted down, resuspended in 150 µl of culture medium, and incubated for 20 min at 20° with 10 µl of anti-CD4 FITC mAb and 10 µl of anti-CD8 PE mAb. After washing in phosphate-buffered saline (PBS), red blood cells were lysed, and the remaining cells were fixed (fluorescence-activated cell sorting (FACS) lysing solution, Becton Dickinson). After an additional washing, cells were resuspended in 250 µl of PBS and 30 µl of fluorescent beads were added. Flow cytometric analysis was performed on a FACScalibur flow cytometer (Becton Dickinson). Lymphocytes were gated on the basis of forward- and side-scatter, and the percentage of CD4+ and CD8+ T cells was determined according to thresholds for positive cell surface staining set at 1% using negative isotype-matched controls. Numeration of the absolute number of T cells was normalized after the number of fluorescent beads counted by the cytometer. To quantify the specific migration of T cells towards CCL5, the number of T cells that migrated in absence of the chemokine was subtracted from the number of T cells that migrated in presence of the chemokine.

Inhibition of migration with the anti-CCR5 2D7

Cells were preincubated at room temperature with the anti-CCR5 mAb 2D7 at indicated concentrations or with an immunoglobulin G2a (IgG2a) isotype control antibody (Beckman-Coulter) for 1 hr in RPMI-1640 supplemented with 10% FCS and were then subjected to a chemotaxis assay as described above.

Cell sorting

Sixty million PBMC separated from whole blood of normal volunteers by density centrifugation were incubated 1 hr at 4° under rotation with 3 µg of the anti-CCR5 mAb PA12 in a final volume of 3 ml of PBS supplemented with 0·1% bovine serum albumin (PBS−0·1% BSA). After washing, cells were incubated for 1 hr at 4° under rotation with 300 × 106 magnetic beads coated with an anti-mouse immunoglobulin by a DNA linker (CELLection Pan Mouse IgG, Dynal, Compiègne, France) in 7·5 ml of PBS−0·1% BSA. The tube was then placed in a magnetic device (Dynal MPC, Dynal) and cells not bound to the beads were collected. The tube was then removed from the magnet and 1·5 ml of RPMI-1640 supplemented with 1% FCS was added to the cells bound to the beads. The two last steps were repeated twice, free cells being collected each time. Finally, cells remained bound to the beads after these washings were released with DNAse according to the manufacturer's instructions. CCR5 expression at the surface of the various cell subpopulations thus sorted was determined by flow cytometry after labelling with the PA12 mAb and a FITC goat anti-mouse probe (GAM-FITC). We verified that the anti-mouse immunoglobulin coated on the beads and released after DNAse treatment did not interfere with the PA12 antibody during the labelling with GAM-FITC and thus did not modify the determination of CCR5 expression (data not shown).

Cell transduction

To produce CCR5- and LacZ-harbouring HIV vectors, VCCR5EGFP12 and pHRCMVlacZ23 plasmids were cotransfected with the HIV packaging plasmid p8.2 and the plasmid pMD.G which encodes the vesicular stomatitis virus glycoprotein envelope in 293T cells, as described.23 PBMC from healthy donors were isolated as described above, and cultured in RPMI-1640 medium supplemented with glutamax-1, 10% FCS, antibiotics, 3 µg/ml of phytohaemagglutinin P (Difco Laboratories, Becton Dickinson), and 100 U/ml interleukin-2 (Boehringer-Mannheim, Mannheim, Germany) for 3 days at 37° and 5% CO2. One million cells were then washed once and resuspended in 300 µl of culture medium in a 24-well plate. Lentiviral vectors were then added to the cell cultures at a multiplicity of infection of 30, along with 8 µg/ml of polybrene (Sigma, St Louis, MO). Cell cultures were then centrifuged at 300 g for 90 min at 30°, incubated for 18 hr at 37°, and washed twice in PBS.

Flow cytometry

PBMC transduced with the CCR5 or the LacZ gene were directly labelled with an anti-CD69 PC5 mAb, and an anti-HLA-DR FITC mAb. CCR5 density at the surface of T cells was quantitated as previously described.13 Blood was collected in EDTA tubes and processed immediately. Cells were directly labelled with an anti-CD4 PE mAb or a an anti-CD8 PE mAb, and indirectly labelled with the anti-CCR5 mAb (2D7) and the GAM-FITC antibody. After gating on CD4+ or on CD8+ cells, the intensity of CCR5 expression on CCR5 expressing cells was analysed by converting FITC fluorescence into the mean number of cell surface-bound mAb molecules per cell, using populations of standard microbeads precoated with different well-defined quantities of mAb (QIFIKIT; Dako, Glostrup, Denmark) and concurrently labelled with the same GAM-FITC probe.

Results

Dose-dependent inhibition of T-cell migration towards CCL5 by an anti-CCR5 mAb

In order to study the migration of human primary T cell towards CCL5 in accordance with their cell surface CCR5 density, we designed a chemotactic assay performed with whole blood. In this assay, blood was loaded into the upper part of a transwell chamber, which lower part contained human recombinant CCL5. We chose CCL5 because it is the most efficient of the major CCR5-binding chemokine, and the most abundant in plasma. Cells were then harvested from the lower chamber, labelled with anti-CD4 and anti-CD8 fluorescent mAb, and analysed by flow cytometry after red blood cell lysis. In this assay, the optimal CCL5 concentration was 100 ng/ml (Fig. 1a). The optimal duration of migration at this concentration was 4 hr (Fig. 1b), the difference between the migrations in presence and in absence of CCL5 being lower after 8 hr because of an increase in the non specific migration at that time. These parameters were chosen for the rest of this study. Pertussis toxin, an inhibitor of the Gαi signalling pathway which is involved in CCR5-dependent chemotaxis, inhibited 70% of the migration (data not shown). Using this assay, we measured the inhibitory effect on CCL5-induced chemotaxis of increasing concentrations of a mAb specific for the chemokine binding site on CCR5. For this purpose, peripheral blood cells were incubated with 0·001–10 µg/ml of the anti-CCR5 mAb 2D7, and submitted to the chemotaxis assay. We observed a sigmoid curve of inhibition of T-cell, CD4+ T-cell and CD8+ T-cell chemotaxis consecutive to anti-CCR5 mAb exposition (Fig. 2a, b, c). In contrast, no inhibition was observed when the anti-CCR5 antibody was replaced with an isotype control antibody.

Figure 1.

Figure 1

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(a) Chemokine dose-dependent migration of peripheral blood T cells towards CCL5. Whole blood diluted 1 : 5 in RPMI medium (100 µl per well) was subjected to a chemotaxis assay toward increasing doses of CCL5. Values are the mean ± SD (n = 3). (b) Time-dependent migration of peripheral blood CD4+ T cells (+), CD8+ T cells (○), and T cells (•) towards CCL5. In the two experiments, the cells that had migrated into the lower chamber after incubation at 37° were collected and labelled with an anti-CD4, CD8 or an isotype control, and counted with a flow cytometer. Results are expressed as the mean number of migrated cells from triplicate wells. The specific migration was quantified as described in material and methods. One representative experiment out of three performed is shown.

Figure 2.

Figure 2

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Effect of increasing concentrations of an anti-CCR5 mAb on T-cell migration towards CCL5. Peripheral blood cells were incubated with various concentrations of the anti-CCR5 mAb 2D7 and subjected to a chemotaxis assay towards CCL5. (a) The intensity of CCL5-induced migration of T cells; (b) CD4+ T cells; and (c) CD8+ T cells was measured by flow cytometry after cell labelling with anti-CD4 and anti-CD8 fluorescent mAb.

T-cell migration intensity towards CCL5 correlates with cell surface CCR5 density

In order to test directly the influence of the level of CCR5 density on the chemokine receptor function, we compared the chemotactic responses to CCL5 of subsets of mononuclear cells expressing various cell surface CCR5 densities. PBMC were incubated with a mouse anti-CCR5 mAb (PA12), then, with magnetic beads coated with antibodies specific for mouse immunoglobulins in order to sort them with a magnetic device. We were able to collect five subpopulations expressing increasing cell surface CCR5 densities after consecutive washing of the cells and final release of the cells tightly bound to the beads. The cytometry profile of these populations is represented in Fig. 3. Afterwards, the migration capacity of these subpopulations towards CCL5 was analysed in the chemotactic assay as described above. We observed a clear correlation between the cell surface CCR5 density of each cell subpopulation and its chemotactic response to the CCR5-binding chemokine (Fig. 4).

Figure 3.

Figure 3

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CCR5 expression on mononuclear cell subpopulations sorted in accordance with their surface CCR5 densities. PBMC were incubated with an anti-CCR5 mAb and thereafter with anti-immunoglobulin-coated magnetic beads. Five cell subpopulations released from the beads after sequential washings were labelled with a fluorescent anti-immunoglobulin probe and analysed by flow cytometry (b, c, d, e, f). As a negative control, non-sorted mononuclear cells were incubated with isotype control IgG2a and the probe (a).

Figure 4.

Figure 4

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Migration towards CCL5 of T-cell subpopulations sorted in accordance with their surface CCR5 densities. Mononuclear cells sorted as described in Fig. 3 were analysed in a chemotaxis assay as described in Fig. 1. Migration of T cells (a), CD4+ T cells (b) and CD8+ T cells (c), is represented for the five different mononuclear cell subpopulations.

Cell surface CCR5 density determines T cell migration capacity towards CCL5

As T-cell activation may induce CCR5 over-expression24–26 it could be argued that the T-cell subpopulations expressing the highest cell surface CCR5 densities are the most activated, and therefore respond the most to CCL5 stimulation. Therefore, we decided to transduce CCR5 positive mononuclear cells with CCR5 and to study the effect of the CCR5 over-expression thus induced at the surface of cells naturally expressing CCR5 on CCL5-induced cell migration. As CCR5– cells may differ from CCR5+ cells by various parameters in addition to CCR5 expression, we chose to work with CCR5-expressing cells and not with non-sorted mononuclear cells.

For this purpose, we sorted PBMC with magnetic beads and collected CCR5 positive cells. We then transduced the cells with an HIV-1 vector harbouring the CCR5 gene. As a negative control, we transduced these CCR5 positive mononuclear cells with the LacZ gene. Figure 5 shows the CCR5 over-expression we obtained in CCR5-transduced CCR5 positive mononuclear cells, as compared with CCR5 density on LacZ-transduced CCR5 positive mononuclear cells. Fifty-one percent of the cells were transduced, and we observed a threefold increase in cell surface CCR5 density between the two populations. Moreover, we verified that CCR5 transduction did not induce a state of higher cell activation than LacZ transduction. For this purpose, we labelled LacZ- and CCR5-transduced peripheral blood mononuclear cells with antibodies specific for the activation markers HLA-DR, and CD69. As shown in Fig. 6, the transfer of the CCR5 gene did not induce higher cell activation than the transfer of the LacZ gene. Next, we compared the migration capacity towards CCL5 of these LacZ-transduced and CCR5-transduced CCR5 positive mononuclear cells in vitro. CCR5 over-expression resulted in an increase in T cells, CD4+ T cells and CD8+ T cells response to the chemokine (Fig. 7a). In a second experiment, we observed a 1·5-fold increase in cell surface CCR5 density between LacZ-transduced and CCR5-transduced (data not shown) and the difference in migration was lower than in the first experiment as shown in Fig. 7(b). To further ascertain that a difference in cell activation is not the cause of the difference in cell migration we observed between LacZ- and CCR5-transduced cells, we focused on the migration of non-activated T cells, that do not express HLA-DR. Figure. 7(c, d) shows that non-activated CCR5-transduced cells actually respond better to CCL5 than non-activated LacZ-transduced cells. Of note, CD8+ T-cell migration was more intense than CD4+ T-cell migration among LacZ-transduced cells. This result is consistent with those reported in Figs 2 and 4, where CD8+ T cells always migrated more than CD4+ T cells. This might be because, as shown in Fig. 8, in peripheral blood, cell surface CCR5 density on CD8+ T cells is higher than on CD4+ T cells (arithmetic means of 15 595 [95% CI 14 057, 17 134] and 7125 [95% CI 6552, 7697] CCR5 molecules per cell, respectively, P < 0·0001). Likewise, the same difference in migration capacity was observed in CCR5-transduced cells between CD8+ T cells and CD4+ T cells (Fig. 7). As the transduction efficiency is similar in CD8+ T cells and in CD4+ T cells (data not shown), this might also be the consequence of a difference in CCR5 density between CD8+ and CD4+CCR5-transduced T cells.

Figure 5.

Figure 5

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CCR5 over-expression induced by transduction of PBMC with the CCR5 gene. CCR5+ mononuclear cells, sorted with an anti-CCR5 mAb and anti-immunoglobulin-coated magnetic beads, were transduced using HIV vectors harbouring either the LacZ gene used as a negative control (a, b) or the CCR5 gene (c, d). Thereafter, cells were labelled either with isotype control IgG2a (a, c) or an anti-CCR5 mAb (b, d) and a fluorescent anti-immunoglobulin probe, and analysed by flow cytometry.

Figure 6.

Figure 6

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Cell activation upon transduction with the CCR5 or the LacZ gene. PBMC were transduced with either the CCR5 or the LacZ gene, labelled 72 hr later with anti-HLA-DR (a) or anti-CD69 (b) monoclonal antibodies, and analyzed by flow cytometry.

Figure 7.

Figure 7

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T-cell surface CCR5 over-expression results in an increase in cell response to CCL5. Migration of CCR5- (closed bar) or LacZ- (open bar) transduced CCR5+ mononuclear cells towards CCL5 was analysed in a chemotaxis assay as described in Fig. 1. Two representative experiments (a, b) are shown, each made on PBMC isolated from a different healthy volunteer. In a third experiment, the migration of either HLA-DR+ or HLA-DR– cells was measured (c, d) In (b), (c) and (d), values are the mean ± SD (n = 3).

Figure 8.

Figure 8

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Cell surface CCR5 density on CD4+ and CD8+ peripheral blood T cells from healthy donors.

The chemotactic response of peripheral blood T cells to CCL5 correlates with their cell-surface CCR5 density

In order to ascertain the physiological relevance of our in vitro findings, we compared peripheral blood CD8+ T-cell migration capacity towards CCL5 with cell surface CCR5 expression levels. For this purpose, we determined peripheral blood CD8+ T cell surface CCR5 densities from 22 healthy donors by quantitative flow cytometry. Concurrently, we measured the chemotaxis capacity of these cells in presence of CCL5. As shown in Fig. 9, we observed a correlation between these two parameters (r = 0·605, P = 0·002). In contrast, there was no correlation between CD8+ T-cell surface CCR5 densities and the intensity of spontaneous cell migration in absence of CCL5 (data not shown).

Figure 9.

Figure 9

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Peripheral blood CD8+ T-cell migration capacity towards CCL5 is linked to cell surface CCR5 density. Chemotaxis is expressed as the ratio between the number of cells migrating in presence of CCL5 and the number of cells migrating in absence of CCL5.

Discussion

In the present study, we show that T- cell surface CCR5 density determines the efficiency of CCR5 as a chemokine receptor. First, we show a dose-dependent inhibition of CCR5-mediated chemotaxis by a mAb specific for the chemokine binding site on CCR5. Second, we observed a strong correlation between T-cell surface CCR5 density and migration intensity toward CCL5. Finally, the increase in cell surface CCR5 density specifically induced by gene transfer which resulted in an overmigration towards CCL5 is the definitive proof that our hypothesis was correct.

Our results are consistent with previous reports of a correlation between the cell surface level of expression of a chemokine receptor and the efficiency of this receptor in mediating chemotaxis, as for instance between CCR2 cell-surface expression and chemotactic responsiveness to the monocyte chemotactic protein.27 Yet, the fact that it is the level of cell-surface expression of a chemokine receptor that determines its efficiency had never been proven so far. It would be interesting to see whether our observation could be extended to other receptors and whether it might correspond to a general biological rule. For instance, it has been shown that the number of T-cell receptors at the surface of a T cell determines the intensity of phosphorylation of the CD3ζ chain, and of cell activation consecutive to antigen stimulation.28

This phenomenon could have many clinical implications. We have previously shown that, although CD4+ T-cell surface CCR5 density may vary from 4000 to 24 000 CCR5 molecules per cell among individuals, it is constant over time for a given individual.13 Because CCR5 efficiency as a chemokine receptor depends on the level of its cell-surface expression, there is an interindividual variability in T-cell response to CCR5-binding chemokines (Fig. 9). It means that there should be a polymorphism in the intensity of the chemotactic response of individuals in the many circumstances where these cytokines are involved. A striking situation is represented by T helper 1 (Th1)-mediated autoimmune diseases. In RA for instance, Th1 cells, interleukin-2- and interferon-γ-producing CD4+ T cells that express CCR5, migrate towards inflamed joints where synovial cells produce large amounts of CCL5, and are thought to amplify the inflammatory process.29 As the intensity of this Th1 cell recruitment is determined by Th1 cell-surface CCR5 density, individuals expressing high CCR5 densities should be more at risk of developing more severe RA than individuals expressing low CCR5 densities. Interestingly, individuals heterozygous for the 32-bp deletion in the CCR5 gene, who express low CCR5 densities, develop less aggressive RA than individuals with wild type CCR5 gene.17,18,20,21,30 We are currently testing the hypothesis that T-cell surface CCR5 density might determine the chemotactic response of T cells to the CCR5-binding chemokines produced by activated synovial cells from patients with RA. Likewise, individual level of cell-surface CCR5 expression might also influence other situations in which CCR5 is involved, such as graft rejection, infection or cancer. Moreover, we have recently shown that cell-surface CCR5 density determines the intensity of the signals triggered by CCL5.31 Thus, not only cells with high surface CCR5 densities will be recruited more efficiently than cells with low surface CCR5 densities, but once attracted they will be more activated. At the end, the level of cell-surface CCR5 expression should have a strong impact on the participation of CCR5-expressing cells to the inflammation.

Now the factors that determine the level of cell surface CCR5 expression in individuals remain to be unveiled. Some of these factors are genetic. Thus, the above-mentioned 32-bp deletion in the CCR5 gene is responsible for a low cell-surface CCR5 density in heterozygous subjects. Yet, these subjects represent only 16% of the population among Caucasians, and even less among non-Caucasians. A few mutations in the CCR5 promoter have also been linked to low CCR5 expressions.32 An interesting hypothesis is that, as the chemokines that bind to CCR5 induce its internalization, the constitutive level of production of these chemokines could modulate the ratio between the number of CCR5 receptors present on the cells and the number of CCR5 receptors present in the cells. We are currently testing this possibility. Another consequence of our findings is that the reduction of T-cell surface CCR5 density should result in a reduction of T-cell response to CCR5-binding chemokines. Thus, compounds able to decrease CCR5 mRNA expression, such as antioxidants33 or rapamycin,34 could have a therapeutic interest in pathological conditions where the down-modulation of CCR5-mediated chemotaxis is desirable. More directly, CCR5 antagonists, which effect on HIV infection is being successfully tested, should also have this therapeutic potential, even at non-saturating doses, by reducing the functional T-cell surface CCR5 density. Recently, Vierboom and colleagues have shown that a CCR5 antagonist treatment, SCH-X, inhibits the development of collagen-induced arthritis in Rhesus monkeys, a model for RA.35 The treatment suppressed joint inflammation, reduced joint destruction and erosion in four out of five animals. Moreover, CCR5 is a very interesting therapeutic target, as CCR5 knockout mice are healthy, as well as humans homozygous for CCR5-Δ32. This lack of apparent adverse consequences is probably because CCR5 is part of a highly redundant chemokine network which shares many overlapping functions.36 However, our results suggest that the reduction of the functional CCR5 cell-surface density secondary to the administration of CCR5 antagonists should impair the attraction exerted by CCR5-binding chemokines that could result in an immune deficiency in situations where CCR5 plays a positive role, as during infections with Listeria monocytogenes,37 Toxoplasma gondii38 or influenza A virus.39

In conclusion, the cell-surface CCR5 density is an important factor which determines the migration intensity of T lymphocytes toward CCL5.

Acknowledgments

The study was supported by the Association pour la Recherche sur la Polyarthrite. We are grateful to Progenics Pharmaceuticals, Inc. for the gift of mAb PA12, and we thank Jérome Estaquier for help with chemotaxis assay.

Abbreviations

CCR5

C-C chemokine receptor 5

FCS

fetal calf serum

FITC

fluorescein isothyocyanate

GAM

goat anti-mouse

HIV-1

human immunodeficiency virus type-1 Th1, T lymphocyte helper 1

RA

rheumatoid arthritis

PBMC

peripheral blood mononuclear cells

PBL

peripheral blood lymphocytes

PC5

phycoerythrin cyanin 5

PE

phycoerythrin

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