Peripheral blood lymphocytes express CCR5, a chemokine receptor for immune cell migration and calcium signaling that serves as an important coreceptor for the HIV. After in vitro stimulation, CCR5 expression is dramatically increased on mature T lymphocytes, especially on the CD45RO+ memory subset. In this study, we report that TNF-α delays the surface expression of CCR5 on PBLs after activation and diminishes CCR5 irrespective of its initial level. Functional loss of CCR5 is reflected in a decreased capability of the treated cells to migrate and signal calcium after MIP-1β stimulation. The effect is mediated via the p80 type II TNF receptor (TNFR2), which induces NF-κB among other factors, leading to an enhanced secretion of the chemokines macrophage-inflammatory protein-1α, macrophage-inflammatory protein-1β, and RANTES. Expression of these chemokines directly down-regulates CCR5. These findings reveal a new regulatory mechanism utilized by activated peripheral T cells to modulate their chemotaxis and potentially other functions mediated by CCR5, including the infection of T lymphocytes by macrophage-tropic HIV strains.

Tumor necrosis factor plays a central role in various immune and inflammatory phenomena (1, 2). The cytokine exerts its bioactivity either as a transmembrane molecule or soluble protein (3). TNF-α achieves signals ranging from cellular activation and proliferation to cytotoxicity and apoptosis. These responses may involve the differential usage of two distinct TNF receptors, p60 (TNFR1) and p80 (TNFR2) and their respective signaling cascades (4, 5, 6). A variety of cell types, both hemopoietic and stromal, have been described to produce TNF (for review, see Ref. 1). Cells of the myeloid lineage such as monocytes and macrophages are potent TNF producers. T lymphocytes also produce TNF, especially in response to recent TCR stimulation (7, 8).

TNF-α can be produced as a normal immune defense mechanism against viruses (9) and tumors (10, 11, 12). However, TNF-α is also associated with many inflammatory and autoimmune pathological processes (1). For example, the dominant role of TNF-α in the pathogenesis of rheumatoid arthritis (RA)2 and idiopathic inflammatory bowel diseases has been demonstrated by successful clinical treatment trials using anti-TNF Abs (13, 14). It was proposed that the inflammatory effect of TNF may be mediated by preferential signaling through TNFR2. TNFR2 induces a widely used gene transactivator: NF-κB, which is known for its role in the transcriptional induction of many immune mediators (for a review, see Refs. 15, 16). NF-κB has an additional important role in preventing cell death (15, 16, 17). NF-κB has been found to be a potent inducer of many chemokines (18, 19, 20).

Chemokines are proinflammatory cytokines that attract and activate specific subsets of leukocytes (21). The chemokine superfamily is divided into subgroups based on structural and genetic considerations. The CXC, CC, C, and CX3C families are characterized by the distance of the two cysteines nearest the amino terminus. The CC chemokines RANTES and macrophage-inflammatory protein (MIP-1α) promote lymphocyte activation (22, 23). These chemokines as well as MIP-1β bind to CCR5. Besides CCR5, RANTES and MIP-1α also bind to CCR1, and RANTES binds to CCR3 and CCR4. Chemokine receptors are seven-transmembrane receptors that couple to G proteins, which in turn can lead to an intracellular calcium signal and protein tyrosine kinase activation (for reviews, see Refs. 24, 25, 26).

CCR5 is an important coreceptor for HIV, and humans lacking its expression are generally resistant to HIV infection, with few exceptions (27, 28, 29, 30, 31, 32). Several reports have shown that LPS in macrophages (33) or CD40L in a macrophage/T cell system (34) can influence the HIV coreceptor expression in vitro. However, published reports give contradictory results concerning the CCR5 expression (33, 35, 36, 37). Recently, it has been suggested that the TNF-α necessary for the maturation of dendritic cells can also influence the expression of chemokine receptors on these cells and cause CCR5 down-modulation (38, 39). It has also been reported that TNF-α decreases the CCR5 expression in peripheral blood monocytes and alveolar macrophages by the production of RANTES (40). However, the effect of TNF-α on CCR5 expression by T cells is not known. This issue is particularly relevant because T cells are important producers of TNF-α and CCR5 plays a significant role in mediating T cell migration and infection by HIV. Our study addresses the interaction between TNF-α and CCR5 in activated human peripheral blood T lymphocytes.

Eagle Hanks’ amino acid (EHAA; Click’s medium) was used for PBLs (Biofluids, Rockville, MD), supplemented with 10% heat-inactivated FCS, penicillin (100 U/ml), streptomycin (100 U/ml), and l-glutamine (2 mM).

PBLs from healthy, normal individuals were obtained from buffy coats from the Department of Transfusion Medicine, Clinical Center, National Institutes of Health, collected by countercurrent elutriation from apheresed subjects from a normal donor pool or by heparin-free phlebotomy. Multiple blood samples were obtained from several CCR5 Δ32 homozygous and heterozygous individuals, who gave informed consent as part of an established clinical protocol approved by Institutional Review Board (IRB; 91-I-0140).

Goat anti-human Abs to MIP-1α, MIP-1β, RANTES, and isotype-matched control Abs were obtained from R&D Systems (Minneapolis, MN). Anti-CCR5 FITC and anti-CD45RO PE were obtained from PharMingen (San Diego, CA). Abs against TNFR2 and TNFR1 were from R&D Systems.

Blood was diluted with PBS 1/5, and Ficoll-Paque (Pharmacia, Piscataway, NJ) was carefully underlayered. After a 15-min spin (2000 rpm, room temperature), the cells at the interphase were collected, treated with ACK lysis buffer (Biofluids) and washed. Cells in complete EHAA were stimulated with 5 μg/ml Con A (Boehringer Mannheim, Indianapolis, IN) for 48–72 h. After treatment with 10 mg/ml α-methyl mannoside (Sigma, St. Louis, MO) for 30 min at 37°C, cells were washed and cultured in complete EHAA with 400 U IL-2 (Proleukin, Midwest Medical, Earth City, MO). The PBL cultures were fed twice per week until they were >2 mo old. CD4+ T cells were obtained by column separation (human T cell CD4 subset column kit; R&D Systems), following the manufacturer’s recommendations.

Stimulated PBLs were incubated with the indicated amounts of TNF-α for the indicated time periods in 6-well or 24-well plates. The supernatants were subjected to ELISA analysis for MIP-1α, MIP-1β, and RANTES (R&D Systems) using the manufacturers’ directions.

PBLs were stained with 1/50 diluted Abs in the presence or absence of TNF-α or anti-TNFR2. Stained cell populations were analyzed on a FACSCalibur with CellQuest Software (Becton Dickinson, Mountain View, CA). Live gates were determined by forward light scatter and side light scatter and verified with propidium iodide staining.

A total of 5 × 106 untreated and TNF-α-treated PBLs in 100 μl was added to the top chamber of a 5-μm pore size 24-Transwell plate (Costar, Fisher Scientific, Pittsburgh, PA) in a final volume of 1 ml in complete medium. To the bottom part edge, MIP-1β (Pepro Tech, Rocky Hill, NJ) was added at the indicated concentrations in triplicates. After 3 h, the inserts containing the cells were removed, and 200-μl aliquots of cells in the bottom chamber were subjected to flow cytometry in a constant volume for 30 s. Total cell numbers obtained were analyzed relative to the migration that had occurred without any chemokine added. Approximately 10% of the cells migrated within this time frame.

Statistical analysis was performed using Microsoft Excel (Redmond, WA).

PBLs were resuspended in HBSS with 1% FCS and 10 mM HEPES containing 10 mM CaCl2 and 10 mM MgCl2 and loaded with 2 μM Indo-1 and 150 μg/ml pluronic acid (Molecular Probes, Eugene, OR) at 30°C for 30 min with constant agitation. Cells were washed twice. The calcium flux was detected using a FACSVantage (Becton Dickinson) dual laser flow cytometer with a time zero injection module (Cytek, Fremont, CA). The data were analyzed with Flowjo Software (Tree Star, Stanford).

PBLs were stimulated in vitro with Con A, treated with TNF-α, and analyzed for CCR5 expression. We observed a decrease in CCR5 surface expression that was detectable within 24 h and almost complete by 48 h (Fig. 1,A). We found that this effect involved the CD45RO+ cells, which were the main population expressing CCR5 (Fig. 1,B) (41, 42, 43, 44). As evident in comparing Fig. 1, A and B, we found that individuals differed in the degree to which separate populations of CCR5-expressing cells were demarcated, but in all cases the CCR5 level was reduced by TNF. The decrease in CCR5 surface expression was directly correlated with the amount of TNF-α added to the culture and reached a plateau at 10 ng/ml (Fig. 1 C). A small subset of cells expressed CCR5 even at maximal doses of TNF-α. These results indicate that TNF-α is a regulator of CCR5 expression on T cells, most prominently on the CD45RO+ subset.

FIGURE 1.

TNF-α treatment leads to decreased CCR5 expression. A, Flow cytometry histograms of CCR5 expression after 24- and 48-h treatment with 30 ng/ml TNF-α compared with no treatment. The numbers indicate the percentage of CCR5-positive cells. Data from one individual with high initial CCR5 expression are depicted. B, PBLs were double stained with anti-CD45RO (PE) and anti-CCR5 (FITC). The untreated and TNF-α-treated (48 h) samples analyzed by flow cytometry are shown. An individual homozygous for CCR5 Δ32 was used as a negative control for the CCR5 gate. The numbers indicate the percentage of the total population in the quadrants. The graphs are representative of seven independent experiments that included 18 individuals. This sample depicts a different individual than shown in A. C, The decrease in the CCR5 expression is a function of TNF-α concentration. PBLs were treated with the indicated concentrations of TNF-α, and the CCR5 analysis was performed by flow cytometry. The individuals’ CCR5 expression is set to 100% without the treatment and compared with the fraction of cells that express CCR5 after TNF-α treatment for 48 h with the indicated doses. Corresponding values for results shown in A would be 75% after 24 h, and 11.75% after 48 h of TNF-α treatment.

FIGURE 1.

TNF-α treatment leads to decreased CCR5 expression. A, Flow cytometry histograms of CCR5 expression after 24- and 48-h treatment with 30 ng/ml TNF-α compared with no treatment. The numbers indicate the percentage of CCR5-positive cells. Data from one individual with high initial CCR5 expression are depicted. B, PBLs were double stained with anti-CD45RO (PE) and anti-CCR5 (FITC). The untreated and TNF-α-treated (48 h) samples analyzed by flow cytometry are shown. An individual homozygous for CCR5 Δ32 was used as a negative control for the CCR5 gate. The numbers indicate the percentage of the total population in the quadrants. The graphs are representative of seven independent experiments that included 18 individuals. This sample depicts a different individual than shown in A. C, The decrease in the CCR5 expression is a function of TNF-α concentration. PBLs were treated with the indicated concentrations of TNF-α, and the CCR5 analysis was performed by flow cytometry. The individuals’ CCR5 expression is set to 100% without the treatment and compared with the fraction of cells that express CCR5 after TNF-α treatment for 48 h with the indicated doses. Corresponding values for results shown in A would be 75% after 24 h, and 11.75% after 48 h of TNF-α treatment.

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To investigate whether the effect of TNF-α is limited to lymphocytes expressing substantial levels of CCR5, or also affects cells expressing low levels of CCR5, we analyzed individuals expressing low CCR5 due to the Δ32 genetic mutation in the CCR5 gene. The Δ32 allele is due to a gene deletion of 32 bp that produces a mutant protein that expresses a unique 32-aa sequence instead of its last two transmembrane domains and is not expressed on the cell surface (29, 30, 32). First, we determined CCR5 surface expression after the stimulation of the cells (Fig. 2,A) and compared it with the T cell activation marker CD25 (Fig. 2,B). We found that CCR5 is not significantly expressed when the cells peak in activation (between days 3 and 5; data not shown), but slowly increases as the activation marker CD25 decreases between days 10 and 20 (compare Fig. 2, A and B). In cases in which the Δ32 mutation is homozygous, the individuals lack CCR5 surface expression (Δ32/Δ32, Fig. 2,A). Heterozygotes for Δ32 had greater CCR5 expression, and wild-type individuals with two normal alleles of CCR5, although variable, generally had the greatest CCR5 expression. These differences between individuals were independent of the degree of cellular activation because all samples appeared to be comparably activated, as indicated by CD25 expression (Fig. 2,B). Table I shows the range of CCR5 expressed in repeated tests on the individuals used in this study. Following TNF-α stimulation for 48 h, we found that both wild-type individuals and Δ32 heterozygous individuals show a comparable decrease in their individual specific surface expression (Fig. 2 C). Only very little CCR5 remains on the cell surface of the heterozygous individuals after TNF-α stimulation due to their initial lower expression.

FIGURE 2.

The CCR5 surface expression of Con A-stimulated PBLs of individuals with diverse CCR5 genotypes. A, PBLs of normal, Δ32 heterozygous, and Δ32 homozygous subjects were stimulated in vitro with Con A and IL-2, as indicated in Materials and Methods. CCR5 surface expression was monitored by flow cytometry analysis on the indicated days. We chose one heterozygous individual expressing intermediate levels of CCR5 as a representative sample. B, Expression of the IL-2Rα-chain (CD25) is graphed for the corresponding samples in A. The data are representative of more than five independent experiments (at variable time points after stimulation) of a regular donor pool of 20 people. C, Average decrease in CCR5 expression after TNF-α treatment of wild-type and CCR5 Δ32 heterozygous individuals after day 20. The individuals’ CCR5 expression is set to 100% without the treatment (stippled bars), then after the 48-h TNF-α treatment (solid bars), the percentage that remains expressed is calculated. For the wild-type bars eight individuals were analyzed, and for the heterozygous bars three individuals were analyzed in two or more independent experiments. The SD of all individuals is shown.

FIGURE 2.

The CCR5 surface expression of Con A-stimulated PBLs of individuals with diverse CCR5 genotypes. A, PBLs of normal, Δ32 heterozygous, and Δ32 homozygous subjects were stimulated in vitro with Con A and IL-2, as indicated in Materials and Methods. CCR5 surface expression was monitored by flow cytometry analysis on the indicated days. We chose one heterozygous individual expressing intermediate levels of CCR5 as a representative sample. B, Expression of the IL-2Rα-chain (CD25) is graphed for the corresponding samples in A. The data are representative of more than five independent experiments (at variable time points after stimulation) of a regular donor pool of 20 people. C, Average decrease in CCR5 expression after TNF-α treatment of wild-type and CCR5 Δ32 heterozygous individuals after day 20. The individuals’ CCR5 expression is set to 100% without the treatment (stippled bars), then after the 48-h TNF-α treatment (solid bars), the percentage that remains expressed is calculated. For the wild-type bars eight individuals were analyzed, and for the heterozygous bars three individuals were analyzed in two or more independent experiments. The SD of all individuals is shown.

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Table I.

CCR5 expression in different Δ32 genotype individualsa

Genotype(+/+)(+/Δ32)(Δ32/Δ32)
% CCR5 positive cells 45.94 10.67 0.74 
Standard deviation 26.78 6.81 0.32 
No. of individuals 
Genotype(+/+)(+/Δ32)(Δ32/Δ32)
% CCR5 positive cells 45.94 10.67 0.74 
Standard deviation 26.78 6.81 0.32 
No. of individuals 
a

PBLs were stimulated in vitro with Con A followed by IL-2. After 14 days the surface expression for CCR5 was determined by flow cytometry. The average percentage of cells that showed CCR5 staining above background are indicated (Δ32 homozygous individuals or isotype Abs were used as controls with identical results). The parentheses indicate the genotype of the individuals in the group, and the results only include frequent blood donors whose CCR5 expression could be validated in more than one blood draw.

We treated PBLs early in the activation process (before CCR5 expression) with TNF-α to determine how TNF-α affects the up-regulation of CCR5. The result shown indicates that the addition of TNF-α significantly delays the expression of CCR5 on the cell surface (Fig. 3). Interestingly, the T cells eventually overcame the initial TNF-α suppression at about day 13 if no further TNF-α was added to the cell culture. Conversely, the cells again decreased CCR5 if TNF-α was administered to the cultures at day 14 (black triangle). Desensitization toward TNF-α signals is not evident at this time because the CCR5 expression is diminished to a comparable degree by TNF-α whether or not the cells had been previously treated with TNF-α (black triangle vs filled circle). Taken together, the above results suggest that TNF-α may be a physiological regulator of CCR5-mediated T cell functions.

FIGURE 3.

The presence of TNF-α delays the up-regulation of CCR5 after in vitro stimulation. Stimulated PBLs at the same density were treated with 30 ng/ml TNF-α on day 4. TNF-α was not replenished and the medium was not changed. The CCR5 surface expression was monitored flow cytometrically. The symbols indicate the percentage of cells that showed CCR5 staining above background (Δ32 homozygous control). The white square represents the untreated sample, and the black diamond the TNF-α-treated sample. The individual shown is (+/+)mb. On day 10 of the treatment (day 14 after isolation), both samples were split and TNF-α (30 ng/ml) was given to the previously untreated sample (•) and to the previously treated sample (▾). The graph is representative of studies on four individuals.

FIGURE 3.

The presence of TNF-α delays the up-regulation of CCR5 after in vitro stimulation. Stimulated PBLs at the same density were treated with 30 ng/ml TNF-α on day 4. TNF-α was not replenished and the medium was not changed. The CCR5 surface expression was monitored flow cytometrically. The symbols indicate the percentage of cells that showed CCR5 staining above background (Δ32 homozygous control). The white square represents the untreated sample, and the black diamond the TNF-α-treated sample. The individual shown is (+/+)mb. On day 10 of the treatment (day 14 after isolation), both samples were split and TNF-α (30 ng/ml) was given to the previously untreated sample (•) and to the previously treated sample (▾). The graph is representative of studies on four individuals.

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To determine whether one of the two TNF-α receptors (TNFRs) or both could mediate the signal leading to decreased CCR5 expression, we analyzed the cell surface expression of the TNFRs. We found continuously high expression of TNFR2 in all individuals tested after activation, but found no detectable TNFR1 (data not shown). Correspondingly, we found that treatment with an anti-TNFR2 agonist Ab led to a decrease in CCR5 expression, whereas TNFR1 Abs had no effect (Fig. 4,A, and data not shown). Thus, the signal leading to the decrease in CCR5 surface expression is mediated by TNFR2, which itself is not down-modulated after TNF-α treatment (data not shown). For example, we found no down-modulation of the TNFR2 during the recovery of the CCR5 expression at late times after TNF-α administration (Fig. 4 B).

FIGURE 4.

Stimulation of TNFR2 decreases CCR5 expression. A, In vitro stimulated PBLs were treated with 0.5 μg/ml anti-TNFR2 polyclonal Ab for 48 h, and the CCR5 surface expression was analyzed by flow cytometry. The individuals’ CCR5 expression is set to 100% without treatment (stippled bar), and after the anti-TNFR2 treatment for 48 h (gray bar) the percentage that remains expressed is calculated. A total of four individuals was analyzed in two independent experiments. The SD of all individuals is shown. B, The surface expression of TNFR2 on human PBLs on five individuals is graphed based on flow cytometry analysis.

FIGURE 4.

Stimulation of TNFR2 decreases CCR5 expression. A, In vitro stimulated PBLs were treated with 0.5 μg/ml anti-TNFR2 polyclonal Ab for 48 h, and the CCR5 surface expression was analyzed by flow cytometry. The individuals’ CCR5 expression is set to 100% without treatment (stippled bar), and after the anti-TNFR2 treatment for 48 h (gray bar) the percentage that remains expressed is calculated. A total of four individuals was analyzed in two independent experiments. The SD of all individuals is shown. B, The surface expression of TNFR2 on human PBLs on five individuals is graphed based on flow cytometry analysis.

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TNFR2 is well known to activate the transcription factor NF-κB (45). Furthermore, it had been reported that the chemokines RANTES, MIP-1α, and MIP-1β have NF-κB-responsive elements in their promoters (18, 46). Therefore, it seemed possible that these chemokines were induced by TNF-α stimulation. Indeed, we found up to a 10-fold increase of these chemokines released after TNF-α stimulation (Fig. 5,A). Furthermore, we analyzed whether levels of NF-κB could influence TNF-α induction of the chemokines. For this purpose, we used the NF-κB-inhibitory peptide (KBI), which strongly interferes with the translocation of the p50/p65 complex to the nucleus (44). We have shown that the KBI peptide prevents the accumulation of the nuclear NF-κB complex detectable by mobility shift electrophoresis (17). We found that KBI caused a substantial decrease in the TNF-α-stimulated chemokine secretion (Fig. 5 B). The blockade was not complete, possibly due to the limited potency of the inhibitory peptide or to alternate mechanisms of chemokine induction. Nevertheless, the results allow us to conclude that TNF-α-induced transcription factors such as NF-κB account for at least part of the increase in chemokines after TNF-α treatment, although clearly other factors also play an important role.

FIGURE 5.

KBI diminishes the TNF-α-induced expression of RANTES, MIP-1α, and MIP-1β. The result shows an ELISA assay of supernatants taken from in vitro stimulated PBLs at the same density that were untreated or treated with 30 ng/ml TNF-α for 48 h. A, The concentration of the secreted chemokines RANTES, MIP-1α, and MIP-1β is graphed without (□) and with (▪) TNF-α stimulation. The graph depicts the result of six individuals in two independent experiments, and the error bars reflect the SD of these individuals. B, □, The chemokine production of samples treated with TNF-α (30 ng/ml) and control peptide (KBC) (80 μg/ml), which is set to 100%. ▪, Samples treated with TNF-α and KBI (80 μg/ml). A total of four individuals were analyzed in two independent experiments. The SD of all individuals is shown.

FIGURE 5.

KBI diminishes the TNF-α-induced expression of RANTES, MIP-1α, and MIP-1β. The result shows an ELISA assay of supernatants taken from in vitro stimulated PBLs at the same density that were untreated or treated with 30 ng/ml TNF-α for 48 h. A, The concentration of the secreted chemokines RANTES, MIP-1α, and MIP-1β is graphed without (□) and with (▪) TNF-α stimulation. The graph depicts the result of six individuals in two independent experiments, and the error bars reflect the SD of these individuals. B, □, The chemokine production of samples treated with TNF-α (30 ng/ml) and control peptide (KBC) (80 μg/ml), which is set to 100%. ▪, Samples treated with TNF-α and KBI (80 μg/ml). A total of four individuals were analyzed in two independent experiments. The SD of all individuals is shown.

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In light of the fact that CCR5 down-regulation was greatest at about 48 h after TNF-α addition, it seemed likely that new gene synthesis could be involved. The previous result clearly indicated a role for NF-κB in the synthesis of the chemokines. Therefore, we tested the influence of KBI on CCR5 expression after TNF-α treatment and found a significantly weaker response to TNF-α in the presence of KBI compared with the control KBC peptide (Fig. 6). Thus, an NF-κB signal is important for the CCR5 down-modulation induced by TNF-α.

FIGURE 6.

The treatment with KBI significantly modifies the TNF-α-induced reduction in CCR5 expression. In vitro stimulated PBLs were treated with 30 ng/ml TNF-α for 48 h, and the CCR5 surface expression was analyzed by flow cytometry. The light bar represents the CCR5 expression of an untreated sample of each individual and is set to 100%, whereas the black bar represents the TNF-α and control peptide (KBC) (80 μg/ml)-treated samples. The striped bars indicate samples of the individuals that were treated with TNF-α and NF-κB-blocking peptide (KBI) (80 μg/ml). A total of four individuals were analyzed in three independent experiments. The SD of all individuals is shown.

FIGURE 6.

The treatment with KBI significantly modifies the TNF-α-induced reduction in CCR5 expression. In vitro stimulated PBLs were treated with 30 ng/ml TNF-α for 48 h, and the CCR5 surface expression was analyzed by flow cytometry. The light bar represents the CCR5 expression of an untreated sample of each individual and is set to 100%, whereas the black bar represents the TNF-α and control peptide (KBC) (80 μg/ml)-treated samples. The striped bars indicate samples of the individuals that were treated with TNF-α and NF-κB-blocking peptide (KBI) (80 μg/ml). A total of four individuals were analyzed in three independent experiments. The SD of all individuals is shown.

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The previous results strongly suggest a role for CCR5-binding chemokines in the CCR5 down-modulation. This hypothesis demands that a blockade of the chemokines that are induced by TNF-α should diminish the TNF-α-induced CCR5 down-regulation. To test this possibility, we incubated the TNF-α-treated samples with either control Ab (Ab ctrl) or a combination of Abs against the three CCR5-binding chemokines, RANTES, MIP-1α, and MIP-1β (Ab mix). We found that the mixture of chemokine-blocking Abs strongly prevented the down-regulation of CCR5 by TNF-α (Fig. 7). This result directly implicates chemokine production as the link between TNF-α treatment and decreased CCR5.

FIGURE 7.

The treatment with a combination of Abs against RANTES, MIP-1α, and MIP-1β prevents the TNF-α-mediated reduction of CCR5 expression. In vitro stimulated PBLs were treated with 30 ng/ml TNF-α for 48 h, and the CCR5 surface expression was analyzed by flow cytometry. The grey bar shows the CCR5 expression of the untreated samples, which is set to 100%. The black bar represents the TNF-α and 20 μg/ml control Ab. The bead-filled bar indicates samples that were treated with TNF-α and a combination of Abs against RANTES, MIP-1α, and MIP-1β (5 μg/ml each). A total of four individuals were analyzed in two independent experiments. The SD of all individuals is shown.

FIGURE 7.

The treatment with a combination of Abs against RANTES, MIP-1α, and MIP-1β prevents the TNF-α-mediated reduction of CCR5 expression. In vitro stimulated PBLs were treated with 30 ng/ml TNF-α for 48 h, and the CCR5 surface expression was analyzed by flow cytometry. The grey bar shows the CCR5 expression of the untreated samples, which is set to 100%. The black bar represents the TNF-α and 20 μg/ml control Ab. The bead-filled bar indicates samples that were treated with TNF-α and a combination of Abs against RANTES, MIP-1α, and MIP-1β (5 μg/ml each). A total of four individuals were analyzed in two independent experiments. The SD of all individuals is shown.

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We investigated whether the decrease in CCR5 induced by the TNF-α treatment has functional consequences for the CD45RO+ cells. Activated lymphocytes were treated with TNF-α for 48 h and then analyzed for their capacity to flux calcium in response to CCR stimulation. We observed a significant decrease in MIP-1β-stimulated calcium flux in cells that were pretreated with TNF-α. By contrast, the same cells gave strong calcium responses after stromal cell-derived factor (SDF) stimulation (Fig. 8,A). We also examined whether pretreatment with TNF-α affected the migration of T cells toward the MIP-1β chemokine. We found that TNF-α treatment decreased the capacity of T cells to migrate in response to MIP-1β (Fig. 8,B). A similar effect was observed when we examined purified CD4+ cells, which also expressed less CCR5 after TNF-α treatment (data not shown). Purified CD4+ cells exhibited lower mobility toward MIP-1β after TNF-α treatment (Fig. 8 C). This finding indicates that the amount of endogenous chemokines produced by CD4+ T cells themselves is sufficient to have an impact on the level of functional CCR5 and that CD8+ cells are not required for this process. Thus, TNF-α can regulate CCR5 expression in CD4+ T cells, which is a key target cell population for HIV infection.

FIGURE 8.

Calcium response and migration of PBLs due to MIP-1β are diminished after TNF-α treatment. In vitro stimulated PBLs were treated with 30 ng/ml TNF-α for 48 h. Cells were then subjected to functional analysis. A, The calcium response of the TNF-α-pretreated T cells after labeling with Indo-1 was determined. The MIP-1β and SDF-1 stimuli were always injected at 60 s after a control injection of buffer at 30 s. The experiment is representative of three experiments with four different individuals. The CCR5 surface expression was 59% without and 31% with TNF-α treatment for the individual shown. B, The migration assay was conducted in a 5 μm Transwell plate with the indicated amount of chemokine on the bottom. After 3 h, the cell numbers were analyzed and compared with background migration. A dose response for total PBLs is shown. Eighty percent of the cell population was positive for CCR5 in the untreated situation and 43% with TNF-α treatment. C, The difference in migration of CD4+ only cells is depicted. Fifty-seven percent of the cells were positive for CCR5 in this particular individual without treatment and 25% after TNF-α treatment. The experiments are representative of four experiments with three different individuals.

FIGURE 8.

Calcium response and migration of PBLs due to MIP-1β are diminished after TNF-α treatment. In vitro stimulated PBLs were treated with 30 ng/ml TNF-α for 48 h. Cells were then subjected to functional analysis. A, The calcium response of the TNF-α-pretreated T cells after labeling with Indo-1 was determined. The MIP-1β and SDF-1 stimuli were always injected at 60 s after a control injection of buffer at 30 s. The experiment is representative of three experiments with four different individuals. The CCR5 surface expression was 59% without and 31% with TNF-α treatment for the individual shown. B, The migration assay was conducted in a 5 μm Transwell plate with the indicated amount of chemokine on the bottom. After 3 h, the cell numbers were analyzed and compared with background migration. A dose response for total PBLs is shown. Eighty percent of the cell population was positive for CCR5 in the untreated situation and 43% with TNF-α treatment. C, The difference in migration of CD4+ only cells is depicted. Fifty-seven percent of the cells were positive for CCR5 in this particular individual without treatment and 25% after TNF-α treatment. The experiments are representative of four experiments with three different individuals.

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Our investigation demonstrates that TNF-α, an important inflammatory protein, potently influences the expression of CCR5, which is the receptor for the proinflammatory chemokines RANTES, MIP-1α, and MIP-1β on T cells and a critical coreceptor for HIV. CCR5 is up-regulated by Ag receptor stimulation of lymphocytes at the point when the activation marker CD25 decreases and is predominantly expressed on CD45RO+ cells. The TNF-α signal that reduces CCR5 surface expression is mainly delivered through TNFR2. TNFR2 can activate NF-κB, which participates in the production and secretion of the chemokines RANTES, MIP-1α, and MIP-1β. It is evident that the synthesis of these chemokines is necessary to achieve CCR5 down-regulation, presumably by binding to the receptor. CCR5 suppression by TNF-α treatment has clear functional effects. Fewer cells migrate toward a chemotactic signal, and treated samples were not capable of manifesting a strong calcium signal after MIP-1β stimulation. Thus, we have identified an inverse correlation between TNF-α, which can be produced by activated T cells or monocytes, and the functional level of the CCR5 chemokine receptor.

It is well established that cell surface expression of CCR5 can be modulated by internalization, which occurs as a consequence of binding of the cognate chemokines or the chemokine derivative AOP-RANTES (47, 48, 49, 50, 51, 52). The internalization mechanism is still under investigation, but appears to involve the G protein-coupled receptor kinases (GRKs) (48), of which GRK2 and GRK3 are expressed in human PBLs. The kinases associate with the receptor and cause the phosphorylation of serines 336, 337, 342, and 349, which is important for the internalization process (48, 50). Also, dynamin is associated with CCR5, causing the internalization to take place through the formation of clathrin vesicles (51). Another report shows an enhancement of tyrosine phosphorylation of CCR5 and the association of Janus kinase 1/STAT5 with CCR5 after activation by RANTES (53). Furthermore, RANTES can lead to focal adhesion kinase association with CCR5 and Zap70; Lck and Pyk2 are activated after ligand binding (53, 54, 55, 56). How these signals participate in the internalization process or other signaling events is not yet established. CCR5 down-modulation could affect signaling elicited by the chemokines or, potentially, the HIV envelope protein (55, 57), which may profoundly influence lymphocyte responses because the two crucial signal molecules Lck and Zap70 are affected. It is also important to note that under circumstances in which the TCR is strongly reengaged (for a review, see Ref. 6), TNF-α had been shown to be part of the mechanism by which these cells undergo apoptosis (8). In the system used in this study, we observed a maximum of 15% cell death with the TNF-α doses used, which is much less than the regression in CCR5 expression and does not account for the loss of CCR5+ cells. The reason for the lack in death under these circumstances is likely the fact that the sensitization or competency signal through the TCR (58, 59) was not provided, because we avoided TCR cross-linking by washing the cells after the Con A treatment.

Productive infection by HIV does not involve coreceptor internalization (47), but is strongly dependent on the level of coreceptor expression. CD4+ cells produce sufficient amounts of chemokines to achieve a coreceptor down-modulation exposed with TNF. Therefore, it is likely that macrophage-tropic HIV particles cannot efficiently enter and infect the T cells that have down-modulated CCR5 after TNF-α exposure, as was demonstrated for macrophages (33, 47, 60, 61, 62). However, TNF-α-induced NF-κB activation also leads to an induction of the HIV long terminal repeat promoter in previously infected T cells (63). Hence, a counterbalance between these effects may determine the ultimate effect of TNF-α on the progression of HIV infection.

We suggest that TNF-α not only enhances the migration of the lymphocytes toward a site of inflammation by increasing chemokine secretion, but may also influence the efficiency by which lymphocytes migrate away from a site of inflammation through the CCR5 down-modulation. In this manner, the response to a strong inflammatory stimulus is overcome and the lymphocyte may then migrate toward other signals, for example, homing signals in secondary lymphoid tissue, and away from the initial site of inflammation. The prompt exit of T cells from the site of infection may be important for a proper immune regulation because the inappropriate presence of T cells appears to be a prominent pathologic effect of autoimmune diseases. In RA, for example, the role of TNF-α in disease exacerbation has been well recognized. Only very recently a role for the chemokine receptor CCR5 in RA was confirmed, consistent with the elevated levels of RANTES found in RA patients (64, 65). The lymphocytes found in inflamed synovial joints of RA patients are CCR5+ (43, 66, 67). Studies on individuals with a variable CCR5 genotype revealed clinical amelioration in RA. In cases in which heterozygosity of the Δ32 mutation occurred, the patients lacked rheumatoid factor, and the swelling of the joints was decreased compared with wild-type individuals (68, 69). In this case, migration to the site of inflammation was reduced. In the case of the CCR5-dependent homing, the cells may persist in the inflamed location until the receptor is decreased. Therefore, the decrease in receptor may be a valuable therapeutic target if internalization could be specifically enhanced. In this case, the T cells on the site of inflammation can execute a protective function, but then do not stay long enough to do harm. One step in this direction is the study of chemokine derivatives such as AOP-RANTES, which are believed to enhance internalization and prevent receptor recycling to the surface. In vitro AOP-RANTES is able to induce CCR5 down-modulation of RA patient cells (66) and may have advantages compared with blocking the chemokines with Abs if administered locally.

Another interesting aspect we have considered is that in internalizing the receptor, the T cells actually decrease the amount of free chemokines, which could be an important termination mechanism for the inflammatory response. Such a mechanism has been suggested to be a strategy by which human CMV escapes detection by the immune system. Human CMV expresses U28, a protein homologous to chemokine receptors. Although it can induce a calcium signal (70), its main function may be the efficient internalization and disposal of secreted chemokines (71). This example of a viral pathogen that disposes of chemokines by internalization suggests that this could also be a normal homostatic regulatory mechanism. It is notable that even the low levels of CCR5 on Δ32 heterozygotes were subject to TNF-α-mediated turnover. Given that the CCR5 promoter itself has an NF-κB binding site (72), which could potentially respond to TNF-α, the T cells may have developed a negative feedback loop for limiting inflammatory responses induced by chemokines. Thus, TNF-α may play a key role in coordinating both positive and negative control of chemokine responses.

We thank Joseph Adelsberger and Ruth Swofford for excellent technical assistance; Shirley Starnes for editorial assistance; Drs. David Scott, Nancy Noben-Traut, and Lixin Zheng for reading the manuscript; and Drs. Richard Siegel and Francis Chan for critical discussions. We thank Dr. Anthony Fauci for laboratory support, and The University of Freiburg and Drs. Hans Kleinig and Edgar Wagner for continuous support. We thank Dr. Ronald Rabin for the biotinylated OKT-3 Ab and technical advice.

2

Abbreviations used in this paper: RA, rheumatoid arthritis; AOP, aminooxypentane; EHAA, Eagle Hanks’ amino acid; GRK, G protein-coupled receptor kinase; KBC, NF-κB control peptide; KBI, NF-κB-inhibitory peptide; MIP, macrophage-inflammatory protein; SDF, stromal cell-derived factor.

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