Platelet Factor 4 Differentially Modulates CD4CD25 (Regulatory) versus CD4CD25– (Nonregulatory) T Cells1
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免疫学杂志 2005年第5期
Abstract
Active suppression mediated by CD4CD25 T regulatory (Tr) cells plays an important role in the down-regulation of T cell responses to both foreign and self-Ags. Platelet factor 4 (PF4), a platelet-derived CXC chemokine, has been shown to strongly inhibit T cell proliferation as well as IFN- and IL-2 release by isolated T cells. In this report we show that human PF4 stimulates proliferation of the naturally anergic human CD4CD25 Tr cells while inhibiting proliferation of CD4CD25– T cells. In coculture experiments we found that CD4CD25 Tr cells exposed to PF4 lose the ability to inhibit the proliferative response of CD4CD25– T cells. Our findings suggest that human PF4, by inducing Tr cell proliferation while impairing Tr cell function, may play a previously unrecognized role in the regulation of human immune responses. Because platelets are the sole source of PF4 in the circulation, these findings may be relevant to the pathogenesis of certain immune-mediated disorders associated with platelet activation, such as heparin-induced thrombocytopenia and autoimmune thrombocytopenic purpura.
Introduction
Autoreactive T and B cells can be detected in healthy individuals, but are normally kept in check by regulatory mechanisms. Among those is an active suppression of naive T cells by T regulatory (Tr)4 cells (1, 2, 3). Several types of Tr cells exist, including CD4 T cells that express the IL-2R -chain (CD25) constitutively, do not secrete IL-10, and suppress immune responses via direct cell-to-cell interactions; type 1 T regulatory (Tr1) cells, which function via secretion of IL-10; and TGF-1-producing Th3 cells (4, 5, 6). CD4CD25 T regulatory cells represent 5–10% of the endogenous CD4 T cell subset (7). Like their mouse counterparts, human CD4CD25 Tr cells are anergic when stimulated in vitro with anti-CD3 mAbs (8) and are able to suppress CD4 and CD8 T cell responses in vitro and in vivo upon TCR ligation (9).
Tr cells may be required for maintenance of homeostasis in the immune system (10). In addition, Tr cells can prevent pathology in autoimmune diseases (1), intestinal inflammatory diseases (11), and allograft rejection (12).
Human PF4, a heparin-binding protein contained in platelet -granules, is secreted upon activation of platelets (13). PF4 belongs to the CXC family of chemokines, but does not share certain proinflammatory properties of other CXC family members because it is missing a critical N-terminal Glu-Leu-Arg sequence, the ELR motif, that precedes the first cysteine residue (14). Several reports have identified PF4 as an inhibitor of hemopoietic progenitor and endothelial cell proliferation and angiogenesis (14, 15). Furthermore, PF4 has been shown to strongly inhibit T cell proliferation as well as IFN- and IL-2 release by isolated T cells (16).
In this report we confirm that human PF4 is a potent inhibitor of CD4CD25– T cell proliferation and show for the first time that this chemokine acts quite differently on CD4CD25 Tr cells, in that it induces proliferation and subsequent loss of regulatory function in this T cell subset. We suggest that additional studies of the latter activity may provide an explanation for the high incidence of Abs specific for PF4:heparin complexes in patients treated with unfractionated heparin and insights into a previously unexpected role for PF4 in regulation of human immune responses.
Materials and Methods
Chemicals and reagents
These were obtained from the following sources: heparin derived from porcine intestinal mucosa (10,000 IU/ml; sp. act., 176 IU/mg) and protamine sulfate (10 mg/ml) from Wyeth/Lederle/ESI; [methyl-3H]thymidine (37 MBq/ml) from PerkinElmer; BSA, phytohemagglutinin (PHA), and propidium iodide from Sigma-Aldrich; recombinant human IL-2 and IL-8 from R&D Systems; mouse mAb anti-CD3 (OKT3) from Ortho Biotech; mAb anti-CD28 (clone CD 28.2) from BD Pharmingen; and rabbit anti-mouse IgG from Jackson ImmunoResearch Laboratories. All other chemicals were reagent grade.
Isolation of T cell populations
Human peripheral blood, anticoagulated with citrate, was obtained from six healthy donors and three patients experiencing heparin-induced thrombocytopenia (HIT). Results obtained using cells derived from normal donors or HIT patients were fully comparable. The human studies were approved by The Blood Center of SE Wisconsin and University at Buffalo institutional review board. PBMC were prepared by centrifugation over Ficoll-Hypaque gradient (Amersham Biosciences). All T cell subpopulations were obtained by positive and/or negative selection of PBMCs on magnetic beads (Miltenyi Biotec) according to the manufacturer’s instructions and previously described methods (17) (Fig. 2A) or FACS sorting on a FACStar (BD Biosciences; Fig. 2B). Briefly, CD3 T cells were purified by positive selection with CD3 microbeads (20 μl/107 cells). CD4 T cells were isolated by positive selection with the CD4 MultiSort kit (20 μl/107 cells). Eluted/released CD4 T cells were washed once in cold PBS containing 0.5% BSA and 2 mM EDTA and were separated with CD25 microbeads (10 μl/107 cells) into CD4CD25– and CD4CD25 T cell fractions. Alternatively, CD4 T cells isolated by positive selection with the CD4 MultiSort kit were labeled with PE-conjugated anti-CD4 and FITC-conjugated anti-CD25 and sorted on a FACStar to obtain a subset of CD4CD25 Tr cells expressing CD25 at high levels (CD4CD25high) (7, 18). All purified T cell subpopulations were used immediately after isolation.
FIGURE 2. Isolation and purity of human CD4CD25–, CD4CD25, and CD4CD25high T cells. Total CD4 T cells were freshly isolated from HIT patients or normal donors’ PBMCs (positive selection) and separated into CD25 (positive selection) and CD25– (negative selection) fractions by magnetic beads (A) or by anti-CD4 magnetic bead enrichment followed by FACS sorting to recover CD4 expressing high levels of CD25 (CD4CD25high; B). The purity of the T cell subpopulations was determined by flow cytometry (A: CD4, >97%; CD25–, >95%; CD25, >94%; B: CD4, >95%; CD25high, >98%). Results are representative of nine independent experiments for A and three for B.
Culture conditions and proliferation/suppression assays
Isolated T cells, in triplicate wells (1 x 105/well, in a 200-μl final volume), were cultured in RPMI 1640 medium supplemented with 100 U/ml penicillin/streptomycin, 2 mM L-glutamine, 5 mM HEPES (Mediatech), and 10% heat-inactivated human AB serum (Cambrex; complete RPMI 1640) in 96-well, U-bottom plates (Corning Life Sciences). Purified T cells were incubated in the absence or the presence of various concentrations of PF4, heparin, PF4:heparin complexes, protamine, and IL-8. Cells were stimulated with PHA (2.5 μg/ml) or with immobilized (4°C, overnight) mAb anti-CD3 (OKT3) at 0.5 μg/ml, either alone or in combination with IL-2 (100 U/ml), soluble (nonimmobilized) anti-CD28 mAb (1 μg/ml), soluble anti-CD28 mAb (1 μg/ml) and IL-2 (100 U/ml), or soluble anti-CD28 mAb (1 μg/ml) together with secondary rabbit anti-mouse IgG (10 μg/ml) as a cross-linker. Human PF4, either isolated from human platelets or expressed as a recombinant protein, was purified in our laboratory as previously described (19, 20). In preliminary experiments we determined that recombinant human PF4 is indistinguishable from human PF4 isolated from platelets with respect to its effects on stimulated T cell subpopulations. All subsequent experiments were performed using human recombinant PF4 at a concentration of 5 μg/ml, unless otherwise indicated. PF4 preparations were shown to be free of endotoxin using a Limulus amebocyte lysate gel clot assay (Cambrex).
Cell proliferation/DNA synthesis was assessed by the level of TdR incorporation. After 72 h of culture, [3H]TdR (1 μCi/well) was added for an additional 16 h. The medium was then discarded, and the cells were harvested using a FilterMate 196 harvester (Packard). Radioactivity was measured using a Matrix 9600 (Packard) gas scintillation beta counter. Assays were performed in triplicate. To analyze the effect of PF4 on the suppressive activity of CD4CD25 Tr cells or CD4CD25– T cells, a constant number of CD4CD25– T cells (5 x 104) was cocultured with different ratios of CD4CD25 Tr cells (from 1/1, e.g., 5 x 104 cells, to 1/32, e.g., 1.5 x 103 cells) as indicated in the figures. Cocultures were stimulated with allogeneic APCs (PBMCs that had been depleted of CD3 T cells by negative selection on CD3 microbeads; Miltenyi Biotec) and exposed to 3000 rad of gamma irradiation. Alternatively, immobilized anti-CD3, in the absence or the presence of PF4, was used as the stimulus.
Flow cytometric analysis
T cells (1 x 105) in 0.02 M PBS, pH 7.4, containing 0.1% sodium azide and 1% BSA were stained for 30 min at 4°C in a 50-μl final volume with optimal dilution of FITC-conjugated anti-CD3 (UCHT1) or anti-CD4 (RPA-T4), PE-conjugated anti-CD25 (M-A251), and mouse subclass-specific isotype controls (BD Pharmingen). Cells were then washed and acquired by flow cytometry (FACScan, FACSCalibur, and CellQuest software (BD Biosciences)). Data were analyzed with WinList software (Verity Software House).
CFSE labeling and flow cytometric analysis
CFSE (Molecular Probes) labeling and analysis were performed according to the manufacturer’s recommendations and previously described methods (21, 22). Briefly, freshly purified CD4CD25 and CD4CD25– T cells (1 x 107/ml) were labeled with 5 μM CFSE for 15 min at 37°C in PBS containing 0.1% BSA. Cells were quenched with ice-cold PBS containing 10% autologous plasma for 5 min, then washed extensively three times. CFSE-labeled T cells (5 x 104) were cultured in 96-well, U-bottom plates coated with mAb anti-CD3 (OKT3; 0.5 μg/ml) in the absence or the presence of 5 μg/ml PF4. In coculture experiments, CFSE-labeled CD4CD25– T cells (5 x 104) were mixed with the same number (1:1 ratio) of unlabeled CD4CD25 (and vice versa). Only viable cells, as determined by propidium iodide staining exclusion, were analyzed. Briefly, after 96-h culture, cells were harvested and washed three times in ice-cold PBS containing 1% BSA. After the last washing step, cells were resuspended in 50 μl of PBS containing 1% BSA and 10 μg of propidium iodide, and acquired by flow cytometry. Data were analyzed with WinList software (Verity Software House). Division was characterized by sequential halving of CFSE fluorescence, generating equally spaced peaks on a logarithmic scale. Six individual peaks and corresponding regions were identified (0–5, delimited by dashed lines, as shown in Fig. 5A). The proportion of cells responding to the different stimuli was calculated according to published methods (21, 22). Briefly, the percentage of events (corresponding to cells) in a given cycle (n) is divided by 2 raised to the power n to calculate the percentage of original, undivided cells from which they arose (precursor number). The sums of these give the total precursor cells for each culture (Table II). As shown in Fig. 5A, the numbers appearing above each peak (0–5) are indicated with the undivided T cells (0) residing in the rightmost peak, and T cells that have divided five times (5) residing in the leftmost peak, ending at the first minor tick mark between the 101–102 scale. Therefore, the sum of cohorts from divisions 1–5 represents the number of precursor T cells that have been proliferating within the 96-h culture. The leftmost peak represents unlabeled CD4CD25 (Fig. 5, C and D) or CD4CD25– T cells (Fig. 5, G and H).
FIGURE 5. PF4 induces the proliferation of CD4CD25 Tr cells and impairs their ability to suppress the response of CD4CD25– T cells to anti-CD3. Freshly isolated CD4CD25– and CD4CD25 T cells were labeled with CFSE and stimulated with immobilized mAb anti-CD3. CFSE-labeled CD4CD25– T cells (5 x 104) were cultured in the absence (A) or the presence of PF4 (B). CFSE-labeled CD4CD25– T cells (5 x 104) mixed with the same number (1:1 ratio) of unlabeled CD4CD25 Tr cells were cultured in the absence (C) or the presence (D) of PF4. CFSE-labeled CD4CD25 T cells (5 x 104) were cultured in the absence (E) or the presence of PF4 (F). CFSE-labeled CD4CD25 Tr cells (5 x 104) mixed with the same number (1:1 ratio) of unlabeled CD4CD25– T cells were cultured in the absence (G) or the presence (H) of PF4. After 96-h culture, cells were harvested and analyzed by flow cytometry. Only viable cells, as determined by propidium iodide staining exclusion, are shown. Dashed lines (shown only in A) indicate the boundaries of each division cycle. The numbers appearing above each peak (0–5) denote each division population, with the undivided T cells (0) residing in the rightmost peak, and the T cells that have divided five times (5) residing in the leftmost peak ending at the first minor tick mark between the 101–102 scale. The leftmost peak represents unlabeled CD4CD25 (C and D) and CD4CD25– T cells (G and H). Results are representative of four independent experiments.
Table II. Responder frequency of CD4CD25– and CD4CD25 T cells stimulated with anti-CD3 mAb, in the absence or the presence of PF4a
Cytokine measurement
T cells were stimulated with immobilized anti-CD3 (OKT3; 0.5 μg/ml) and soluble anti-CD28 (1 μg/ml) mAbs in complete RPMI 1640. Supernatants from triplicate culture wells were collected after 72 h and frozen at –80°C until use. Cytokine concentrations in supernatants were assessed using a commercially available ELISA kit for human TGF-1 (R&D Systems) and a cytometric bead array (Th1/Th2 Cytokine CBA 1; BD Pharmingen) for IL-2, IL-4, IL-5, IL-10, IFN-, and TNF-, following the manufacturer’s protocols. The limits of sensitivity were as follows: 3 pg/ml for IL-2, IL-4, IL-5, IL-10, and TNF-; and 7 pg/ml for IFN- and TGF-1.
Statistical analysis
All analyses for statistically significant differences were performed with Student’s paired t test with a two-tailed distribution. A value of p < 0.05 was considered significant. All cultures were performed in triplicate, and error bars represent 1 SD. Reported results were obtained from multiple experiments with similar results. Only a representative experiment is reported for each figure.
Results
PF4 inhibits proliferation of purified CD3 T cells
CD3 T cells were isolated from buffy coats of HIT patients or normal donors using magnetic bead separation. The resulting purity of CD3 T cells was >95% as determined by flow cytometry (data not shown). The freshly purified CD3 T cells were stimulated with immobilized anti-CD3 mAb (0.5 μg/ml) in the absence or the presence of increasing concentrations of PF4, PF4:heparin complexes, heparin, or protamine. Consistent with a previous report (16), PF4 strongly inhibited T cell proliferation in a dose-dependent manner (Fig. 1). Lesser, but statistically significant, inhibition was also seen with PF4:heparin complexes. Conversely, we found that heparin alone and protamine, a positively charged heparin-binding protein used as a control for any putative interaction mediated only by positive charges, were without effect. In three independent experiments, an average of 82.4 ± 4.1% (p 0.0006) inhibition was observed in the presence of 5 μg/ml PF4, 4.8 ± 6.5% (p 0.64) in the presence of 5 μg/ml protamine, 3.9 ± 3.0% (p 0.40) in the presence of 5 U/ml heparin, and 53.4 ± 2.4% (p 0.002) in the presence of 5 μg/ml PF4:0.25 U/ml heparin complexes. These findings indicate that PF4 and PF4:heparin complexes inhibit the proliferation of human T cells in response to TCR cross-linking. The absence of APCs, such as B cells and monocytes, in these cultures shows that APCs are not required for this effect.
FIGURE 1. PF4 inhibits the proliferation of purified CD3 T cells stimulated by anti-CD3 mAb. Total CD3 T cells were freshly isolated from HIT patients or normal donors’ PBMCs (purity, >95%). CD3 T cells (1 x 105 cells/well) were stimulated with immobilized anti-CD3 mAb (0.5 μg/ml) in the absence or the presence of the indicated concentrations of PF4, PF4:heparin complexes, heparin, or protamine. After 72-h culture, [3H]thymidine was added for an additional 16 h. Individual data points represent the average of triplicate determinations ± 1 SD. *, p < 0.05 vs medium (by t test). Results are representative of three independent experiments.
PF4 inhibits proliferation of CD4CD25– T cells, but stimulates proliferation of CD4CD25 Tr cells
It has been convincingly demonstrated that CD4CD25 Tr cells are present in the peripheral blood of humans and exert inhibitory regulatory function on naive T cells in a cell-cell contact-dependent manner (8, 17). To investigate whether PF4 acts on a specific T cell subset, we isolated CD4CD25– nonregulatory T cells and CD4CD25 regulatory T cells from HIT patients and normal healthy donors using a two-step magnetic separation. Purity >94% was achieved for both T cell subpopulations, as determined by flow cytometry (Fig. 2). Freshly isolated CD4CD25– T cells and CD4CD25 Tr cells were cultured and activated with different stimuli in the absence or the presence of 5 μg/ml PF4. As shown in Fig. 3A, PF4 strongly inhibited the proliferation of CD4CD25– T cells in response to anti-CD3 mAb alone or combined with anti-CD28 mAb, but was without effect on the response to anti-CD3/anti-CD28 cross-linked (with rabbit anti-mouse IgG), anti-CD3 plus IL-2, and anti-CD3/CD28 plus IL-2. In five independent experiments (three with T cells from normal donors and two from HIT patients), in the presence of PF4 an average of 14.5 ± 2.6% (p 0.02) inhibition was observed when these T cells were activated by PHA, 52.3 ± 5.1% (p 0.0008) when activated by anti-CD3 alone (anti-CD3), and 39.7 ± 1.5% (p 0.002) when activated by anti-CD3 together with anti-CD28 (CD3/28). Addition of anti-CD3, anti-CD28, and rabbit anti-mouse IgG as a cross-linker (CD3/28/CL), or anti-CD3 together with IL-2 (CD32), resulted in a slight, but not statistically significant, inhibition mediated by PF4 (12.9 ± 5.2% (p 0.16) and 11.5 ± 3.9% (p 0.11), respectively). When this T cell subpopulation was activated by anti-CD3 together with anti-CD28 and IL-2 (CD3/282), a slight, but not statistically significant, increase in the proliferation in the presence of PF4 was observed (11.7 ± 4.6%; p 0.20). These data indicate that PF4 has inhibitory properties on CD4CD25– T cell proliferation, as observed for unfractionated T cells (Fig. 1). Moreover, IL-2 and cross-linker interfere with PF4 inhibitory effects on T cell proliferation.
FIGURE 3. PF4 inhibits the proliferation of CD4CD25– T cells, but stimulates the proliferation of CD4CD25 Tr cells. CD4CD25– (A) and CD4CD25 (B) T cell populations, in the absence (–) or the presence () of PF4 (5 μg/ml), were tested (1 x 105 cells/well) for their ability to proliferate in response to different stimuli, including PHA (2.5 μg/ml), immobilized mAb anti-CD3 at 0.5 μg/ml, either alone (CD3) or in combination with soluble anti-CD28 mAb at 1 μg/ml (CD3/28), soluble anti-CD28 mAb at 1 μg/ml together with secondary rabbit anti-mouse IgG (10 μg/ml) used as a cross-linker (CD3/28/CL), IL-2 at 100 U/ml (CD32), IL-2 at 100 U/ml, and soluble anti-CD28 mAb at 1 μg/ml (CD3/282). CD4CD25– (C) or CD4CD25high (D) T cell populations were stimulated with immobilized mAb anti-CD3 at 0.5 μg/ml either alone (CD3) or in combination with soluble anti-CD28 mAb at 1 μg/ml (CD3/28) in the absence (medium only (M)) or the presence of PF4 (5 g/ml), PF4:heparin complexes (PF4:H; 5 μg/ml:0.25 U/ml) or IL-8 (5 μg/ml). After 72-h culture, [3H]thymidine was added for an additional 16 h. Individual data points represent the average of triplicate determinations ± 1 SD. *, p < 0.05 vs medium (by t test). Results are representative of five independent experiments for A and B, and three for C and D.
CD4CD25 Tr cells do not ordinarily respond to PHA or anti-CD3 mAb (8, 23). As shown in Fig. 3B, in contrast to what was observed for CD4CD25– T cells, PF4 (5 μg/ml) caused CD4CD25 Tr cells to become responsive to anti-CD3 and promoted their increased response to the other agents used for stimulation, except for PHA.
In five independent experiments (three using T cells isolated from normal donors and two using cells from HIT patients), in the presence of PF4 an average of 86.8 ± 13.1% (p 0.009) up-regulation (increased proliferation vs medium) was observed when these T cells were activated by anti-CD3 alone (CD3; 47.2 ± 7.0%; p 0.002), when stimulated by anti-CD3 together with anti-CD28 (CD3/28; 24.7 ± 6.6%; p 0.03), with the addition of anti-CD3 together with anti-CD28 and rabbit anti-mouse IgG as a cross-linker (CD3/28/CL; 42.7 ± 1.9%; p 0.003) when anti-CD3 and IL-2 (CD32) were combined together, and 30.2 ± 3.1% (p 0.01) when IL-2 was added to anti-CD3 and anti-CD28 (CD3/282). These data provide the first evidence that PF4 differentially modulates human T cell subsets.
It has been reported that CD4CD25 Tr cells can be distinguished on the basis of the CD25 levels of expression (high and low) being the CD25high subset a homogeneous population consisting of Tr cells (7, 18). We therefore repeated the experiment shown above with the CD4CD25high Tr cell subset obtained by FACS sorting. The antithetic effects of PF4 on CD4CD25– T cells and CD4CD25high Tr cells in response to anti-CD3 and anti-CD3/28 mAbs were comparable to those obtained with CD4CD25 Tr cells isolated by a two-step magnetic separation and appear to be specific because these effects were not mimicked by equivalent quantities of IL-8, a CXC chemokine that is significantly homologous with PF4 in amino acid composition (Fig. 3, C and D). In three independent experiments (two using T cells isolated from normal donors and one from HIT patients) confirming our observations, PF4 strongly inhibited proliferation of CD4CD25– T cells (Fig. 3C) induced by anti-CD3 (48.4 ± 6.7%; p 0.01) or induced by anti-CD3 together with anti-CD28 (36.5 ± 6.5%; p 0.02), whereas heparin partially neutralized the inhibitory effects of PF4 (41.1 ± 6.2% (p 0.04) for -CD3; 25.6 ± 6.2% (p 0.04) for -CD3/28). Similarly to what is shown in Fig. 3B, PF4 strongly up-regulated the proliferation of CD4CD25high Tr cells (Fig. 3D) induced by anti-CD3 (81.5 ± 15.7%; p 0.007) or induced by anti-CD3 together with anti-CD28 (48.3 ± 8.0%; p 0.01), whereas in the presence of heparin the up-regulatory effects of PF4 were reduced, but were still significant (50.7 ± 16.1% (p 0.05) for -CD3; 31.7 ± 6.3% (p 0.04) for -CD3/28). IL-8 failed to affect the proliferation of either T cell population.
PF4 inhibits cytokine release of CD4CD25– T cells, but up-regulates cytokine release of CD4CD25 Tr cells
Cytokine profiles of CD4CD25– T cells and CD4CD25 Tr cells stimulated with immobilized anti-CD3 and soluble anti-CD28 mAbs are summarized in Table I (numbers represent the average values (picograms per milliliter) of pooled data from three independent experiments; SDs were <20%). Mirroring the proliferation results (Fig. 3, A and B), PF4 strongly inhibited the release of IL-2, IL-4, IL-5, IL-10, IFN-, and TNF- by CD4CD25– T cells without affecting TGF-1 production. In contrast, PF4 significantly increased the release of IL-2, IFN-, and TNF-, but not IL-10 or TGF-1, by CD4CD25 Tr cells. Taken together, these findings demonstrate a previously unrecognized role for PF4 in modulating the response of human T cell subpopulations to TCR cross-linking.
Table I. Cytokine production (picograms per milliliter) by CD4CD25– and CD4CD25 T cells in the absence or the presence of PF4a
PF4 induces proliferation, rather than suppression, of cocultures of CD4CD25– and CD4CD25 T cells
It is known that CD4CD25 Tr cells suppress the proliferation and cytokine production of CD4CD25– T cells (8, 17). To further characterize the suppressive effect of PF4 on mixed T cell populations, we studied the effect of PF4 on the response to allogenic APCs (Fig. 4A; representative of four independent experiments) or immobilized anti-CD3 (Fig. 4B; representative of three independent experiments) of T cell mixtures containing known ratios of CD4CD25 and CD4CD25– T cells. As expected, in the absence of PF4, CD4CD25 Tr cells, at a 1:1 ratio, suppressed the proliferative response of CD4CD25– T cells to alloantigens (40.2 ± 6.3%; p 0.028; Fig. 4A) and to anti-CD3 (52.9 ± 3.5%; p 0.029; Fig. 4B).
FIGURE 4. CD4CD25 regulatory T cells suppress the proliferation of CD4CD25– T cells in a dose-dependent manner. In the presence of PF4, the proliferation of mixed CD4CD25– and CD4CD25 cell culture increases. Freshly purified CD4CD25 Tr cells, at the indicated ratio, were added to autologous CD4CD25– T cells (5 x 104) stimulated with allogeneic APCs (A; 1 x 105) or immobilized mAb anti-CD3 (B; 0.5 μg/ml) in the absence or the presence of PF4 (5 μg/ml). After 72-h culture, [3H]thymidine was added for an additional 16 h. Individual data points represent the average of triplicate determinations ± 1 SD. *, p < 0.05 (by t test). Results are representative of four independent experiments for A and three for B.
Unexpectedly, however, PF4 accentuated the proliferative response of a 1:1 coculture of CD4CD25 and CD4CD25– T cells to alloantigens (74.9 ± 6.3%; p 0.004) and to anti-CD3 (34.3 ± 5.4%; p 0.006). Because in a coculture system, TdR incorporation cannot discriminate between CD4CD25– and CD4CD25 T cell proliferation, the increased proliferation observed in the presence of PF4 can be ascribed to either an increased response of CD4CD25 Tr cells to PF4 (as our previous findings could suggest) or a loss of the suppressive activity of CD4 CD25 Tr cells on CD4CD25– T cells, which, in turn, proliferate.
Suppression of the proliferative response of CD4CD25– T cells to anti-CD3, mediated by CD4CD25 Tr cells, is impaired in the presence of PF4
To dissect which subset of T cells proliferated in cocultures of CD4CD25 and CD4CD25– T cells exposed to PF4, we repeated the study shown in Fig. 4B using CFSE labeling to track the proliferation of the two T cell subpopulations. In these studies, CFSE-labeled CD4CD25– (5 x 104) T cells were cultured alone (Fig. 5, A and B) or with an equal number of unlabeled CD4CD25 Tr cells (Fig. 5, C and D). In the converse experiment, CFSE-labeled CD4CD25 (5 x 104) Tr cells were cultured alone (Fig. 5, E and F) or with an equal number of unlabeled CD4CD25– T cells (Fig. 5, G and H). Each of the cell mixtures was stimulated with immobilized anti-CD3 mAb in the absence or the presence of PF4. After a 96-h culture, cells were harvested and analyzed by flow cytometry. Allogeneic APCs alone were not used in this assay, because the levels of autofluorescence they generated and the relatively weak stimulus provided greatly limited the number of cell cycles that could be resolved by flow cytometry (data not shown).
In four independent experiments, as expected, CD4CD25– T cells proliferated in response to anti-CD3 (Fig. 5A and Table II), and this response was inhibited by PF4 (Fig. 5B and Table II). Unexpectedly, however, in the presence of unlabeled CD4CD25 Tr cells, the proliferation of CD4CD25– T cells was moderately enhanced by PF4, rather than suppressed (Fig. 5D and Table II). CD4CD25 Tr cells were unresponsive to anti-CD3 (Fig. 5E and Table II); however, they proliferated in response to this stimulus when PF4 was present (Fig. 5F and Table II). This response was unaffected by the presence of unlabeled CD4CD25– T cells (Fig. 5H and Table II). Together, these data show that PF4 acts on CD4CD25 Tr cells to stimulate their proliferation (Fig. 5H) and that this response, paradoxically, impairs the ability CD4CD25 Tr cells to inhibit the proliferation of CD4CD25– T cells (Fig. 5D) in response to anti-CD3.
Discussion
CD4CD25 Tr cells are essential for the maintenance of self-tolerance in mouse models (1, 11, 12). A large body of evidence suggests that human CD4CD25 T cells share many of the characteristics of murine CD4CD25 Tr cells (18). In our experimental conditions, PF4 stimulates the proliferation of CD4CD25 regulatory T cells (Fig. 3, B and D). Moreover, in the presence of PF4, CD4CD25 Tr cells lose their potent suppressive capacity on conventional CD4CD25– T cells in response to alloantigen or TCR engagement (anti-CD3; Figs. 4 and 5). Therefore, in this study we report that human PF4 reverts the anergic Tr cell phenotype and impairs their suppressive activity, suggesting a previously unrecognized role of PF4 in the regulation of human immune responses.
Although it is known that CD4CD25 Tr cells exert their suppressive effect by inhibiting IL-2 production and promoting cell cycle arrest (7, 24), the specific mechanism governing the suppressive activity of CD4CD25 Tr cells is still undefined. Signaling through a few receptors appears to be critical for the regulatory activity of CD4CD25 Tr cells. A glucocorticoid-induced TNF receptor (TNFRSF18) is highly expressed on mouse CD4CD25 Tr cells (25). In a mouse model, glucocorticoid-induced TNF receptor stimulation abrogated CD4CD25 Tr cell-mediated suppression in vitro and in vivo (25, 26). 4-1BB, another TNF receptor superfamily member, has been shown to modulate the suppressor function of activated, but not resting, CD4CD25 Tr (27). Ligation of CD28 or blockade of CTLA-4 expressed on CD4CD25 Tr cells can also abrogate suppression (28, 29).
How PF4 modulates CD25CD4 Tr cell-mediated suppression remains to be determined. As a possible mechanism of action, direct signaling of PF4 via CXCR3B, a recently identified spliced variant of CXCR3 to which PF4 binds with high affinity (30), or another yet to be identified receptor may interfere with the capacity of CD25CD4 Tr cells to deliver to other T cells a negative signal for activation and proliferation. Alternatively, PF4 could physically interfere (in an antagonistic or synergistic manner) with the interaction of costimulatory molecules on T cells and APCs by competing for glycosaminoglycan binding. In our experiments, heparin partially affected the proliferative response induced by PF4 on the two T cell subsets. It is known that PF4 has a high affinity for cell surface heparan sulfate proteoglycans, on which it recognizes a specific structural motif (31). Although proteoglycans generally serve as coreceptors (32), an intrinsic signaling capacity has been reported for syndecan-4 expressed on lymphocytes and monocytes (33). Therefore, direct signaling through proteoglycans cannot be ruled out for PF4. It should be considered that PF4 could also affect T cell function indirectly by induction of immunosuppressive cytokines. In line with previous observations (17), we found that isolated CD4CD25 T cells produced TGF-1, but not IL-10. The levels of both TGF-1 and IL-10 were unchanged in the presence of PF4 (Table I), suggesting that neither IL-10 nor TGF-1 mediates PF4 effects on Tr cells. Future investigation into how PF4 signaling is initiated is necessary to fully characterize the mechanism of its effects.
In our experimental conditions, the effects of PF4 were seen using concentrations of PF4 >1 μg/ml (Fig. 1). Subsequent experiments on regulatory and nonregulatory CD4 T cells were performed using PF4 at a concentration of 5 μg/ml. This is a high concentration, considering that in physiological conditions only minute quantities (14 ng/ml) of PF4 are found in human plasma (34). However, activated platelets, from 1 ml of whole blood, rapidly release PF4 levels in the range of 10 μg/ml (35). Moreover, significant amounts of PF4 are normally associated with endothelial cell proteoglycans (36). Our results are in agreement with those of Fleischer et al. (16), who reported half-maximal inhibition of T cell proliferation at 0.7 μM PF4 (e.g., 5 μg/ml). The finding by Taub et al. (37) that PF4 lacks any capacity to modulate purified human T cells does not appear to contradict our present observations, because the authors used PF4 at physiological concentrations (<1 μg/ml). Interestingly, PF4-mediated modulation of T cell proliferation occurs at the same concentration range (2–10 μg/ml) as that required for inhibition of endothelial cells (38) and activation of human neutrophils (39) and monocytes (40) by this chemokine. Such plasma concentrations of PF4 could occur during heparin therapy, because heparin releases PF4 from endothelial cell glycosaminoglycans and directly from circulating platelets (41). Moreover, platelet activation that occurs during and after specific invasive procedures, such as cardiopulmonary bypass (conventionally associated with heparin administration) (42), also leads to increased PF4 release into the circulation. Therefore, a transient impairment of Tr suppressor activity might be postulated in otherwise healthy individuals experiencing acute and severe platelet destruction, hence possibly explaining why different patient groups exposed to heparin become sensitized at such different rates (43, 44).
Of interest, the immune response in HIT is peculiar in that Ag formation results from noncovalent interaction of a specific protein (PF4) with a specific drug (heparin) (20). In normal subjects, peptides derived from PF4 are undoubtedly presented regularly to T cells without triggering an immune response, either because T cells reactive with such peptides have been deleted in the thymus or because they have been tolerized by exposure to PF4 peptides in the context of class II MHC in the absence of costimulatory molecules. We propose that in HIT a transient impairment of the suppressive capacity of CD4CD25 Tr cells primes naive T cells, triggering this unusual immune response. Therefore, we hypothesize that in HIT, PF4 is not only the target for the Ab (when complexed with heparin), but is also a modulator of T cell activation.
Our observation that PF4 modulates, in an opposite manner, regulatory vs nonregulatory T cells, thus affecting their function, provides new insights into a previously unexpected role for this chemokine in regulation of the human immune response. Therefore, our findings could be relevant to immune complex-mediated inflammatory and autoimmune disorders associated with platelet activation and subsequent release of PF4, among those HIT, systemic lupus erythematosus, and autoimmune thrombocytopenic purpura, in which the trigger for the primary immune response is poorly understood.
Disclosures
The authors have no financial conflict of interest.
Acknowledgments
We are indebted to Dr. Mortimer Poncz (University of Pennsylvania) for the generous gift of the cDNA clone encoding human PF4, and to Dr. Carolyn Keever-Taylor (Medical College of Wisconsin) for kindly providing the OKT3 mAb.
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 Grant HL64704 from the National Heart, Lung, and Blood Institute.
2 Current address: University of Ulsan College of Medicine, Department of Laboratory Medicine, Ulsan University Hospital, 290-3 Jeonha-dong, Dong-gu, Ulsan 682-714, South Korea.
3 Address correspondence and reprint requests to Dr. Gian Paolo Visentin, Department of Pediatrics, University at Buffalo-State University of New York, Biomedical Research Building, Room 422, Buffalo, NY 14214. E-mail address: visentin{at}buffalo.edu
4 Abbreviations used in this paper: Tr, T regulatory; HIT, heparin-induced thrombocytopenia; PF4, platelet factor 4; PHA, phytohemagglutinin.
Received for publication August 26, 2004. Accepted for publication December 13, 2004.
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Active suppression mediated by CD4CD25 T regulatory (Tr) cells plays an important role in the down-regulation of T cell responses to both foreign and self-Ags. Platelet factor 4 (PF4), a platelet-derived CXC chemokine, has been shown to strongly inhibit T cell proliferation as well as IFN- and IL-2 release by isolated T cells. In this report we show that human PF4 stimulates proliferation of the naturally anergic human CD4CD25 Tr cells while inhibiting proliferation of CD4CD25– T cells. In coculture experiments we found that CD4CD25 Tr cells exposed to PF4 lose the ability to inhibit the proliferative response of CD4CD25– T cells. Our findings suggest that human PF4, by inducing Tr cell proliferation while impairing Tr cell function, may play a previously unrecognized role in the regulation of human immune responses. Because platelets are the sole source of PF4 in the circulation, these findings may be relevant to the pathogenesis of certain immune-mediated disorders associated with platelet activation, such as heparin-induced thrombocytopenia and autoimmune thrombocytopenic purpura.
Introduction
Autoreactive T and B cells can be detected in healthy individuals, but are normally kept in check by regulatory mechanisms. Among those is an active suppression of naive T cells by T regulatory (Tr)4 cells (1, 2, 3). Several types of Tr cells exist, including CD4 T cells that express the IL-2R -chain (CD25) constitutively, do not secrete IL-10, and suppress immune responses via direct cell-to-cell interactions; type 1 T regulatory (Tr1) cells, which function via secretion of IL-10; and TGF-1-producing Th3 cells (4, 5, 6). CD4CD25 T regulatory cells represent 5–10% of the endogenous CD4 T cell subset (7). Like their mouse counterparts, human CD4CD25 Tr cells are anergic when stimulated in vitro with anti-CD3 mAbs (8) and are able to suppress CD4 and CD8 T cell responses in vitro and in vivo upon TCR ligation (9).
Tr cells may be required for maintenance of homeostasis in the immune system (10). In addition, Tr cells can prevent pathology in autoimmune diseases (1), intestinal inflammatory diseases (11), and allograft rejection (12).
Human PF4, a heparin-binding protein contained in platelet -granules, is secreted upon activation of platelets (13). PF4 belongs to the CXC family of chemokines, but does not share certain proinflammatory properties of other CXC family members because it is missing a critical N-terminal Glu-Leu-Arg sequence, the ELR motif, that precedes the first cysteine residue (14). Several reports have identified PF4 as an inhibitor of hemopoietic progenitor and endothelial cell proliferation and angiogenesis (14, 15). Furthermore, PF4 has been shown to strongly inhibit T cell proliferation as well as IFN- and IL-2 release by isolated T cells (16).
In this report we confirm that human PF4 is a potent inhibitor of CD4CD25– T cell proliferation and show for the first time that this chemokine acts quite differently on CD4CD25 Tr cells, in that it induces proliferation and subsequent loss of regulatory function in this T cell subset. We suggest that additional studies of the latter activity may provide an explanation for the high incidence of Abs specific for PF4:heparin complexes in patients treated with unfractionated heparin and insights into a previously unexpected role for PF4 in regulation of human immune responses.
Materials and Methods
Chemicals and reagents
These were obtained from the following sources: heparin derived from porcine intestinal mucosa (10,000 IU/ml; sp. act., 176 IU/mg) and protamine sulfate (10 mg/ml) from Wyeth/Lederle/ESI; [methyl-3H]thymidine (37 MBq/ml) from PerkinElmer; BSA, phytohemagglutinin (PHA), and propidium iodide from Sigma-Aldrich; recombinant human IL-2 and IL-8 from R&D Systems; mouse mAb anti-CD3 (OKT3) from Ortho Biotech; mAb anti-CD28 (clone CD 28.2) from BD Pharmingen; and rabbit anti-mouse IgG from Jackson ImmunoResearch Laboratories. All other chemicals were reagent grade.
Isolation of T cell populations
Human peripheral blood, anticoagulated with citrate, was obtained from six healthy donors and three patients experiencing heparin-induced thrombocytopenia (HIT). Results obtained using cells derived from normal donors or HIT patients were fully comparable. The human studies were approved by The Blood Center of SE Wisconsin and University at Buffalo institutional review board. PBMC were prepared by centrifugation over Ficoll-Hypaque gradient (Amersham Biosciences). All T cell subpopulations were obtained by positive and/or negative selection of PBMCs on magnetic beads (Miltenyi Biotec) according to the manufacturer’s instructions and previously described methods (17) (Fig. 2A) or FACS sorting on a FACStar (BD Biosciences; Fig. 2B). Briefly, CD3 T cells were purified by positive selection with CD3 microbeads (20 μl/107 cells). CD4 T cells were isolated by positive selection with the CD4 MultiSort kit (20 μl/107 cells). Eluted/released CD4 T cells were washed once in cold PBS containing 0.5% BSA and 2 mM EDTA and were separated with CD25 microbeads (10 μl/107 cells) into CD4CD25– and CD4CD25 T cell fractions. Alternatively, CD4 T cells isolated by positive selection with the CD4 MultiSort kit were labeled with PE-conjugated anti-CD4 and FITC-conjugated anti-CD25 and sorted on a FACStar to obtain a subset of CD4CD25 Tr cells expressing CD25 at high levels (CD4CD25high) (7, 18). All purified T cell subpopulations were used immediately after isolation.
FIGURE 2. Isolation and purity of human CD4CD25–, CD4CD25, and CD4CD25high T cells. Total CD4 T cells were freshly isolated from HIT patients or normal donors’ PBMCs (positive selection) and separated into CD25 (positive selection) and CD25– (negative selection) fractions by magnetic beads (A) or by anti-CD4 magnetic bead enrichment followed by FACS sorting to recover CD4 expressing high levels of CD25 (CD4CD25high; B). The purity of the T cell subpopulations was determined by flow cytometry (A: CD4, >97%; CD25–, >95%; CD25, >94%; B: CD4, >95%; CD25high, >98%). Results are representative of nine independent experiments for A and three for B.
Culture conditions and proliferation/suppression assays
Isolated T cells, in triplicate wells (1 x 105/well, in a 200-μl final volume), were cultured in RPMI 1640 medium supplemented with 100 U/ml penicillin/streptomycin, 2 mM L-glutamine, 5 mM HEPES (Mediatech), and 10% heat-inactivated human AB serum (Cambrex; complete RPMI 1640) in 96-well, U-bottom plates (Corning Life Sciences). Purified T cells were incubated in the absence or the presence of various concentrations of PF4, heparin, PF4:heparin complexes, protamine, and IL-8. Cells were stimulated with PHA (2.5 μg/ml) or with immobilized (4°C, overnight) mAb anti-CD3 (OKT3) at 0.5 μg/ml, either alone or in combination with IL-2 (100 U/ml), soluble (nonimmobilized) anti-CD28 mAb (1 μg/ml), soluble anti-CD28 mAb (1 μg/ml) and IL-2 (100 U/ml), or soluble anti-CD28 mAb (1 μg/ml) together with secondary rabbit anti-mouse IgG (10 μg/ml) as a cross-linker. Human PF4, either isolated from human platelets or expressed as a recombinant protein, was purified in our laboratory as previously described (19, 20). In preliminary experiments we determined that recombinant human PF4 is indistinguishable from human PF4 isolated from platelets with respect to its effects on stimulated T cell subpopulations. All subsequent experiments were performed using human recombinant PF4 at a concentration of 5 μg/ml, unless otherwise indicated. PF4 preparations were shown to be free of endotoxin using a Limulus amebocyte lysate gel clot assay (Cambrex).
Cell proliferation/DNA synthesis was assessed by the level of TdR incorporation. After 72 h of culture, [3H]TdR (1 μCi/well) was added for an additional 16 h. The medium was then discarded, and the cells were harvested using a FilterMate 196 harvester (Packard). Radioactivity was measured using a Matrix 9600 (Packard) gas scintillation beta counter. Assays were performed in triplicate. To analyze the effect of PF4 on the suppressive activity of CD4CD25 Tr cells or CD4CD25– T cells, a constant number of CD4CD25– T cells (5 x 104) was cocultured with different ratios of CD4CD25 Tr cells (from 1/1, e.g., 5 x 104 cells, to 1/32, e.g., 1.5 x 103 cells) as indicated in the figures. Cocultures were stimulated with allogeneic APCs (PBMCs that had been depleted of CD3 T cells by negative selection on CD3 microbeads; Miltenyi Biotec) and exposed to 3000 rad of gamma irradiation. Alternatively, immobilized anti-CD3, in the absence or the presence of PF4, was used as the stimulus.
Flow cytometric analysis
T cells (1 x 105) in 0.02 M PBS, pH 7.4, containing 0.1% sodium azide and 1% BSA were stained for 30 min at 4°C in a 50-μl final volume with optimal dilution of FITC-conjugated anti-CD3 (UCHT1) or anti-CD4 (RPA-T4), PE-conjugated anti-CD25 (M-A251), and mouse subclass-specific isotype controls (BD Pharmingen). Cells were then washed and acquired by flow cytometry (FACScan, FACSCalibur, and CellQuest software (BD Biosciences)). Data were analyzed with WinList software (Verity Software House).
CFSE labeling and flow cytometric analysis
CFSE (Molecular Probes) labeling and analysis were performed according to the manufacturer’s recommendations and previously described methods (21, 22). Briefly, freshly purified CD4CD25 and CD4CD25– T cells (1 x 107/ml) were labeled with 5 μM CFSE for 15 min at 37°C in PBS containing 0.1% BSA. Cells were quenched with ice-cold PBS containing 10% autologous plasma for 5 min, then washed extensively three times. CFSE-labeled T cells (5 x 104) were cultured in 96-well, U-bottom plates coated with mAb anti-CD3 (OKT3; 0.5 μg/ml) in the absence or the presence of 5 μg/ml PF4. In coculture experiments, CFSE-labeled CD4CD25– T cells (5 x 104) were mixed with the same number (1:1 ratio) of unlabeled CD4CD25 (and vice versa). Only viable cells, as determined by propidium iodide staining exclusion, were analyzed. Briefly, after 96-h culture, cells were harvested and washed three times in ice-cold PBS containing 1% BSA. After the last washing step, cells were resuspended in 50 μl of PBS containing 1% BSA and 10 μg of propidium iodide, and acquired by flow cytometry. Data were analyzed with WinList software (Verity Software House). Division was characterized by sequential halving of CFSE fluorescence, generating equally spaced peaks on a logarithmic scale. Six individual peaks and corresponding regions were identified (0–5, delimited by dashed lines, as shown in Fig. 5A). The proportion of cells responding to the different stimuli was calculated according to published methods (21, 22). Briefly, the percentage of events (corresponding to cells) in a given cycle (n) is divided by 2 raised to the power n to calculate the percentage of original, undivided cells from which they arose (precursor number). The sums of these give the total precursor cells for each culture (Table II). As shown in Fig. 5A, the numbers appearing above each peak (0–5) are indicated with the undivided T cells (0) residing in the rightmost peak, and T cells that have divided five times (5) residing in the leftmost peak, ending at the first minor tick mark between the 101–102 scale. Therefore, the sum of cohorts from divisions 1–5 represents the number of precursor T cells that have been proliferating within the 96-h culture. The leftmost peak represents unlabeled CD4CD25 (Fig. 5, C and D) or CD4CD25– T cells (Fig. 5, G and H).
FIGURE 5. PF4 induces the proliferation of CD4CD25 Tr cells and impairs their ability to suppress the response of CD4CD25– T cells to anti-CD3. Freshly isolated CD4CD25– and CD4CD25 T cells were labeled with CFSE and stimulated with immobilized mAb anti-CD3. CFSE-labeled CD4CD25– T cells (5 x 104) were cultured in the absence (A) or the presence of PF4 (B). CFSE-labeled CD4CD25– T cells (5 x 104) mixed with the same number (1:1 ratio) of unlabeled CD4CD25 Tr cells were cultured in the absence (C) or the presence (D) of PF4. CFSE-labeled CD4CD25 T cells (5 x 104) were cultured in the absence (E) or the presence of PF4 (F). CFSE-labeled CD4CD25 Tr cells (5 x 104) mixed with the same number (1:1 ratio) of unlabeled CD4CD25– T cells were cultured in the absence (G) or the presence (H) of PF4. After 96-h culture, cells were harvested and analyzed by flow cytometry. Only viable cells, as determined by propidium iodide staining exclusion, are shown. Dashed lines (shown only in A) indicate the boundaries of each division cycle. The numbers appearing above each peak (0–5) denote each division population, with the undivided T cells (0) residing in the rightmost peak, and the T cells that have divided five times (5) residing in the leftmost peak ending at the first minor tick mark between the 101–102 scale. The leftmost peak represents unlabeled CD4CD25 (C and D) and CD4CD25– T cells (G and H). Results are representative of four independent experiments.
Table II. Responder frequency of CD4CD25– and CD4CD25 T cells stimulated with anti-CD3 mAb, in the absence or the presence of PF4a
Cytokine measurement
T cells were stimulated with immobilized anti-CD3 (OKT3; 0.5 μg/ml) and soluble anti-CD28 (1 μg/ml) mAbs in complete RPMI 1640. Supernatants from triplicate culture wells were collected after 72 h and frozen at –80°C until use. Cytokine concentrations in supernatants were assessed using a commercially available ELISA kit for human TGF-1 (R&D Systems) and a cytometric bead array (Th1/Th2 Cytokine CBA 1; BD Pharmingen) for IL-2, IL-4, IL-5, IL-10, IFN-, and TNF-, following the manufacturer’s protocols. The limits of sensitivity were as follows: 3 pg/ml for IL-2, IL-4, IL-5, IL-10, and TNF-; and 7 pg/ml for IFN- and TGF-1.
Statistical analysis
All analyses for statistically significant differences were performed with Student’s paired t test with a two-tailed distribution. A value of p < 0.05 was considered significant. All cultures were performed in triplicate, and error bars represent 1 SD. Reported results were obtained from multiple experiments with similar results. Only a representative experiment is reported for each figure.
Results
PF4 inhibits proliferation of purified CD3 T cells
CD3 T cells were isolated from buffy coats of HIT patients or normal donors using magnetic bead separation. The resulting purity of CD3 T cells was >95% as determined by flow cytometry (data not shown). The freshly purified CD3 T cells were stimulated with immobilized anti-CD3 mAb (0.5 μg/ml) in the absence or the presence of increasing concentrations of PF4, PF4:heparin complexes, heparin, or protamine. Consistent with a previous report (16), PF4 strongly inhibited T cell proliferation in a dose-dependent manner (Fig. 1). Lesser, but statistically significant, inhibition was also seen with PF4:heparin complexes. Conversely, we found that heparin alone and protamine, a positively charged heparin-binding protein used as a control for any putative interaction mediated only by positive charges, were without effect. In three independent experiments, an average of 82.4 ± 4.1% (p 0.0006) inhibition was observed in the presence of 5 μg/ml PF4, 4.8 ± 6.5% (p 0.64) in the presence of 5 μg/ml protamine, 3.9 ± 3.0% (p 0.40) in the presence of 5 U/ml heparin, and 53.4 ± 2.4% (p 0.002) in the presence of 5 μg/ml PF4:0.25 U/ml heparin complexes. These findings indicate that PF4 and PF4:heparin complexes inhibit the proliferation of human T cells in response to TCR cross-linking. The absence of APCs, such as B cells and monocytes, in these cultures shows that APCs are not required for this effect.
FIGURE 1. PF4 inhibits the proliferation of purified CD3 T cells stimulated by anti-CD3 mAb. Total CD3 T cells were freshly isolated from HIT patients or normal donors’ PBMCs (purity, >95%). CD3 T cells (1 x 105 cells/well) were stimulated with immobilized anti-CD3 mAb (0.5 μg/ml) in the absence or the presence of the indicated concentrations of PF4, PF4:heparin complexes, heparin, or protamine. After 72-h culture, [3H]thymidine was added for an additional 16 h. Individual data points represent the average of triplicate determinations ± 1 SD. *, p < 0.05 vs medium (by t test). Results are representative of three independent experiments.
PF4 inhibits proliferation of CD4CD25– T cells, but stimulates proliferation of CD4CD25 Tr cells
It has been convincingly demonstrated that CD4CD25 Tr cells are present in the peripheral blood of humans and exert inhibitory regulatory function on naive T cells in a cell-cell contact-dependent manner (8, 17). To investigate whether PF4 acts on a specific T cell subset, we isolated CD4CD25– nonregulatory T cells and CD4CD25 regulatory T cells from HIT patients and normal healthy donors using a two-step magnetic separation. Purity >94% was achieved for both T cell subpopulations, as determined by flow cytometry (Fig. 2). Freshly isolated CD4CD25– T cells and CD4CD25 Tr cells were cultured and activated with different stimuli in the absence or the presence of 5 μg/ml PF4. As shown in Fig. 3A, PF4 strongly inhibited the proliferation of CD4CD25– T cells in response to anti-CD3 mAb alone or combined with anti-CD28 mAb, but was without effect on the response to anti-CD3/anti-CD28 cross-linked (with rabbit anti-mouse IgG), anti-CD3 plus IL-2, and anti-CD3/CD28 plus IL-2. In five independent experiments (three with T cells from normal donors and two from HIT patients), in the presence of PF4 an average of 14.5 ± 2.6% (p 0.02) inhibition was observed when these T cells were activated by PHA, 52.3 ± 5.1% (p 0.0008) when activated by anti-CD3 alone (anti-CD3), and 39.7 ± 1.5% (p 0.002) when activated by anti-CD3 together with anti-CD28 (CD3/28). Addition of anti-CD3, anti-CD28, and rabbit anti-mouse IgG as a cross-linker (CD3/28/CL), or anti-CD3 together with IL-2 (CD32), resulted in a slight, but not statistically significant, inhibition mediated by PF4 (12.9 ± 5.2% (p 0.16) and 11.5 ± 3.9% (p 0.11), respectively). When this T cell subpopulation was activated by anti-CD3 together with anti-CD28 and IL-2 (CD3/282), a slight, but not statistically significant, increase in the proliferation in the presence of PF4 was observed (11.7 ± 4.6%; p 0.20). These data indicate that PF4 has inhibitory properties on CD4CD25– T cell proliferation, as observed for unfractionated T cells (Fig. 1). Moreover, IL-2 and cross-linker interfere with PF4 inhibitory effects on T cell proliferation.
FIGURE 3. PF4 inhibits the proliferation of CD4CD25– T cells, but stimulates the proliferation of CD4CD25 Tr cells. CD4CD25– (A) and CD4CD25 (B) T cell populations, in the absence (–) or the presence () of PF4 (5 μg/ml), were tested (1 x 105 cells/well) for their ability to proliferate in response to different stimuli, including PHA (2.5 μg/ml), immobilized mAb anti-CD3 at 0.5 μg/ml, either alone (CD3) or in combination with soluble anti-CD28 mAb at 1 μg/ml (CD3/28), soluble anti-CD28 mAb at 1 μg/ml together with secondary rabbit anti-mouse IgG (10 μg/ml) used as a cross-linker (CD3/28/CL), IL-2 at 100 U/ml (CD32), IL-2 at 100 U/ml, and soluble anti-CD28 mAb at 1 μg/ml (CD3/282). CD4CD25– (C) or CD4CD25high (D) T cell populations were stimulated with immobilized mAb anti-CD3 at 0.5 μg/ml either alone (CD3) or in combination with soluble anti-CD28 mAb at 1 μg/ml (CD3/28) in the absence (medium only (M)) or the presence of PF4 (5 g/ml), PF4:heparin complexes (PF4:H; 5 μg/ml:0.25 U/ml) or IL-8 (5 μg/ml). After 72-h culture, [3H]thymidine was added for an additional 16 h. Individual data points represent the average of triplicate determinations ± 1 SD. *, p < 0.05 vs medium (by t test). Results are representative of five independent experiments for A and B, and three for C and D.
CD4CD25 Tr cells do not ordinarily respond to PHA or anti-CD3 mAb (8, 23). As shown in Fig. 3B, in contrast to what was observed for CD4CD25– T cells, PF4 (5 μg/ml) caused CD4CD25 Tr cells to become responsive to anti-CD3 and promoted their increased response to the other agents used for stimulation, except for PHA.
In five independent experiments (three using T cells isolated from normal donors and two using cells from HIT patients), in the presence of PF4 an average of 86.8 ± 13.1% (p 0.009) up-regulation (increased proliferation vs medium) was observed when these T cells were activated by anti-CD3 alone (CD3; 47.2 ± 7.0%; p 0.002), when stimulated by anti-CD3 together with anti-CD28 (CD3/28; 24.7 ± 6.6%; p 0.03), with the addition of anti-CD3 together with anti-CD28 and rabbit anti-mouse IgG as a cross-linker (CD3/28/CL; 42.7 ± 1.9%; p 0.003) when anti-CD3 and IL-2 (CD32) were combined together, and 30.2 ± 3.1% (p 0.01) when IL-2 was added to anti-CD3 and anti-CD28 (CD3/282). These data provide the first evidence that PF4 differentially modulates human T cell subsets.
It has been reported that CD4CD25 Tr cells can be distinguished on the basis of the CD25 levels of expression (high and low) being the CD25high subset a homogeneous population consisting of Tr cells (7, 18). We therefore repeated the experiment shown above with the CD4CD25high Tr cell subset obtained by FACS sorting. The antithetic effects of PF4 on CD4CD25– T cells and CD4CD25high Tr cells in response to anti-CD3 and anti-CD3/28 mAbs were comparable to those obtained with CD4CD25 Tr cells isolated by a two-step magnetic separation and appear to be specific because these effects were not mimicked by equivalent quantities of IL-8, a CXC chemokine that is significantly homologous with PF4 in amino acid composition (Fig. 3, C and D). In three independent experiments (two using T cells isolated from normal donors and one from HIT patients) confirming our observations, PF4 strongly inhibited proliferation of CD4CD25– T cells (Fig. 3C) induced by anti-CD3 (48.4 ± 6.7%; p 0.01) or induced by anti-CD3 together with anti-CD28 (36.5 ± 6.5%; p 0.02), whereas heparin partially neutralized the inhibitory effects of PF4 (41.1 ± 6.2% (p 0.04) for -CD3; 25.6 ± 6.2% (p 0.04) for -CD3/28). Similarly to what is shown in Fig. 3B, PF4 strongly up-regulated the proliferation of CD4CD25high Tr cells (Fig. 3D) induced by anti-CD3 (81.5 ± 15.7%; p 0.007) or induced by anti-CD3 together with anti-CD28 (48.3 ± 8.0%; p 0.01), whereas in the presence of heparin the up-regulatory effects of PF4 were reduced, but were still significant (50.7 ± 16.1% (p 0.05) for -CD3; 31.7 ± 6.3% (p 0.04) for -CD3/28). IL-8 failed to affect the proliferation of either T cell population.
PF4 inhibits cytokine release of CD4CD25– T cells, but up-regulates cytokine release of CD4CD25 Tr cells
Cytokine profiles of CD4CD25– T cells and CD4CD25 Tr cells stimulated with immobilized anti-CD3 and soluble anti-CD28 mAbs are summarized in Table I (numbers represent the average values (picograms per milliliter) of pooled data from three independent experiments; SDs were <20%). Mirroring the proliferation results (Fig. 3, A and B), PF4 strongly inhibited the release of IL-2, IL-4, IL-5, IL-10, IFN-, and TNF- by CD4CD25– T cells without affecting TGF-1 production. In contrast, PF4 significantly increased the release of IL-2, IFN-, and TNF-, but not IL-10 or TGF-1, by CD4CD25 Tr cells. Taken together, these findings demonstrate a previously unrecognized role for PF4 in modulating the response of human T cell subpopulations to TCR cross-linking.
Table I. Cytokine production (picograms per milliliter) by CD4CD25– and CD4CD25 T cells in the absence or the presence of PF4a
PF4 induces proliferation, rather than suppression, of cocultures of CD4CD25– and CD4CD25 T cells
It is known that CD4CD25 Tr cells suppress the proliferation and cytokine production of CD4CD25– T cells (8, 17). To further characterize the suppressive effect of PF4 on mixed T cell populations, we studied the effect of PF4 on the response to allogenic APCs (Fig. 4A; representative of four independent experiments) or immobilized anti-CD3 (Fig. 4B; representative of three independent experiments) of T cell mixtures containing known ratios of CD4CD25 and CD4CD25– T cells. As expected, in the absence of PF4, CD4CD25 Tr cells, at a 1:1 ratio, suppressed the proliferative response of CD4CD25– T cells to alloantigens (40.2 ± 6.3%; p 0.028; Fig. 4A) and to anti-CD3 (52.9 ± 3.5%; p 0.029; Fig. 4B).
FIGURE 4. CD4CD25 regulatory T cells suppress the proliferation of CD4CD25– T cells in a dose-dependent manner. In the presence of PF4, the proliferation of mixed CD4CD25– and CD4CD25 cell culture increases. Freshly purified CD4CD25 Tr cells, at the indicated ratio, were added to autologous CD4CD25– T cells (5 x 104) stimulated with allogeneic APCs (A; 1 x 105) or immobilized mAb anti-CD3 (B; 0.5 μg/ml) in the absence or the presence of PF4 (5 μg/ml). After 72-h culture, [3H]thymidine was added for an additional 16 h. Individual data points represent the average of triplicate determinations ± 1 SD. *, p < 0.05 (by t test). Results are representative of four independent experiments for A and three for B.
Unexpectedly, however, PF4 accentuated the proliferative response of a 1:1 coculture of CD4CD25 and CD4CD25– T cells to alloantigens (74.9 ± 6.3%; p 0.004) and to anti-CD3 (34.3 ± 5.4%; p 0.006). Because in a coculture system, TdR incorporation cannot discriminate between CD4CD25– and CD4CD25 T cell proliferation, the increased proliferation observed in the presence of PF4 can be ascribed to either an increased response of CD4CD25 Tr cells to PF4 (as our previous findings could suggest) or a loss of the suppressive activity of CD4 CD25 Tr cells on CD4CD25– T cells, which, in turn, proliferate.
Suppression of the proliferative response of CD4CD25– T cells to anti-CD3, mediated by CD4CD25 Tr cells, is impaired in the presence of PF4
To dissect which subset of T cells proliferated in cocultures of CD4CD25 and CD4CD25– T cells exposed to PF4, we repeated the study shown in Fig. 4B using CFSE labeling to track the proliferation of the two T cell subpopulations. In these studies, CFSE-labeled CD4CD25– (5 x 104) T cells were cultured alone (Fig. 5, A and B) or with an equal number of unlabeled CD4CD25 Tr cells (Fig. 5, C and D). In the converse experiment, CFSE-labeled CD4CD25 (5 x 104) Tr cells were cultured alone (Fig. 5, E and F) or with an equal number of unlabeled CD4CD25– T cells (Fig. 5, G and H). Each of the cell mixtures was stimulated with immobilized anti-CD3 mAb in the absence or the presence of PF4. After a 96-h culture, cells were harvested and analyzed by flow cytometry. Allogeneic APCs alone were not used in this assay, because the levels of autofluorescence they generated and the relatively weak stimulus provided greatly limited the number of cell cycles that could be resolved by flow cytometry (data not shown).
In four independent experiments, as expected, CD4CD25– T cells proliferated in response to anti-CD3 (Fig. 5A and Table II), and this response was inhibited by PF4 (Fig. 5B and Table II). Unexpectedly, however, in the presence of unlabeled CD4CD25 Tr cells, the proliferation of CD4CD25– T cells was moderately enhanced by PF4, rather than suppressed (Fig. 5D and Table II). CD4CD25 Tr cells were unresponsive to anti-CD3 (Fig. 5E and Table II); however, they proliferated in response to this stimulus when PF4 was present (Fig. 5F and Table II). This response was unaffected by the presence of unlabeled CD4CD25– T cells (Fig. 5H and Table II). Together, these data show that PF4 acts on CD4CD25 Tr cells to stimulate their proliferation (Fig. 5H) and that this response, paradoxically, impairs the ability CD4CD25 Tr cells to inhibit the proliferation of CD4CD25– T cells (Fig. 5D) in response to anti-CD3.
Discussion
CD4CD25 Tr cells are essential for the maintenance of self-tolerance in mouse models (1, 11, 12). A large body of evidence suggests that human CD4CD25 T cells share many of the characteristics of murine CD4CD25 Tr cells (18). In our experimental conditions, PF4 stimulates the proliferation of CD4CD25 regulatory T cells (Fig. 3, B and D). Moreover, in the presence of PF4, CD4CD25 Tr cells lose their potent suppressive capacity on conventional CD4CD25– T cells in response to alloantigen or TCR engagement (anti-CD3; Figs. 4 and 5). Therefore, in this study we report that human PF4 reverts the anergic Tr cell phenotype and impairs their suppressive activity, suggesting a previously unrecognized role of PF4 in the regulation of human immune responses.
Although it is known that CD4CD25 Tr cells exert their suppressive effect by inhibiting IL-2 production and promoting cell cycle arrest (7, 24), the specific mechanism governing the suppressive activity of CD4CD25 Tr cells is still undefined. Signaling through a few receptors appears to be critical for the regulatory activity of CD4CD25 Tr cells. A glucocorticoid-induced TNF receptor (TNFRSF18) is highly expressed on mouse CD4CD25 Tr cells (25). In a mouse model, glucocorticoid-induced TNF receptor stimulation abrogated CD4CD25 Tr cell-mediated suppression in vitro and in vivo (25, 26). 4-1BB, another TNF receptor superfamily member, has been shown to modulate the suppressor function of activated, but not resting, CD4CD25 Tr (27). Ligation of CD28 or blockade of CTLA-4 expressed on CD4CD25 Tr cells can also abrogate suppression (28, 29).
How PF4 modulates CD25CD4 Tr cell-mediated suppression remains to be determined. As a possible mechanism of action, direct signaling of PF4 via CXCR3B, a recently identified spliced variant of CXCR3 to which PF4 binds with high affinity (30), or another yet to be identified receptor may interfere with the capacity of CD25CD4 Tr cells to deliver to other T cells a negative signal for activation and proliferation. Alternatively, PF4 could physically interfere (in an antagonistic or synergistic manner) with the interaction of costimulatory molecules on T cells and APCs by competing for glycosaminoglycan binding. In our experiments, heparin partially affected the proliferative response induced by PF4 on the two T cell subsets. It is known that PF4 has a high affinity for cell surface heparan sulfate proteoglycans, on which it recognizes a specific structural motif (31). Although proteoglycans generally serve as coreceptors (32), an intrinsic signaling capacity has been reported for syndecan-4 expressed on lymphocytes and monocytes (33). Therefore, direct signaling through proteoglycans cannot be ruled out for PF4. It should be considered that PF4 could also affect T cell function indirectly by induction of immunosuppressive cytokines. In line with previous observations (17), we found that isolated CD4CD25 T cells produced TGF-1, but not IL-10. The levels of both TGF-1 and IL-10 were unchanged in the presence of PF4 (Table I), suggesting that neither IL-10 nor TGF-1 mediates PF4 effects on Tr cells. Future investigation into how PF4 signaling is initiated is necessary to fully characterize the mechanism of its effects.
In our experimental conditions, the effects of PF4 were seen using concentrations of PF4 >1 μg/ml (Fig. 1). Subsequent experiments on regulatory and nonregulatory CD4 T cells were performed using PF4 at a concentration of 5 μg/ml. This is a high concentration, considering that in physiological conditions only minute quantities (14 ng/ml) of PF4 are found in human plasma (34). However, activated platelets, from 1 ml of whole blood, rapidly release PF4 levels in the range of 10 μg/ml (35). Moreover, significant amounts of PF4 are normally associated with endothelial cell proteoglycans (36). Our results are in agreement with those of Fleischer et al. (16), who reported half-maximal inhibition of T cell proliferation at 0.7 μM PF4 (e.g., 5 μg/ml). The finding by Taub et al. (37) that PF4 lacks any capacity to modulate purified human T cells does not appear to contradict our present observations, because the authors used PF4 at physiological concentrations (<1 μg/ml). Interestingly, PF4-mediated modulation of T cell proliferation occurs at the same concentration range (2–10 μg/ml) as that required for inhibition of endothelial cells (38) and activation of human neutrophils (39) and monocytes (40) by this chemokine. Such plasma concentrations of PF4 could occur during heparin therapy, because heparin releases PF4 from endothelial cell glycosaminoglycans and directly from circulating platelets (41). Moreover, platelet activation that occurs during and after specific invasive procedures, such as cardiopulmonary bypass (conventionally associated with heparin administration) (42), also leads to increased PF4 release into the circulation. Therefore, a transient impairment of Tr suppressor activity might be postulated in otherwise healthy individuals experiencing acute and severe platelet destruction, hence possibly explaining why different patient groups exposed to heparin become sensitized at such different rates (43, 44).
Of interest, the immune response in HIT is peculiar in that Ag formation results from noncovalent interaction of a specific protein (PF4) with a specific drug (heparin) (20). In normal subjects, peptides derived from PF4 are undoubtedly presented regularly to T cells without triggering an immune response, either because T cells reactive with such peptides have been deleted in the thymus or because they have been tolerized by exposure to PF4 peptides in the context of class II MHC in the absence of costimulatory molecules. We propose that in HIT a transient impairment of the suppressive capacity of CD4CD25 Tr cells primes naive T cells, triggering this unusual immune response. Therefore, we hypothesize that in HIT, PF4 is not only the target for the Ab (when complexed with heparin), but is also a modulator of T cell activation.
Our observation that PF4 modulates, in an opposite manner, regulatory vs nonregulatory T cells, thus affecting their function, provides new insights into a previously unexpected role for this chemokine in regulation of the human immune response. Therefore, our findings could be relevant to immune complex-mediated inflammatory and autoimmune disorders associated with platelet activation and subsequent release of PF4, among those HIT, systemic lupus erythematosus, and autoimmune thrombocytopenic purpura, in which the trigger for the primary immune response is poorly understood.
Disclosures
The authors have no financial conflict of interest.
Acknowledgments
We are indebted to Dr. Mortimer Poncz (University of Pennsylvania) for the generous gift of the cDNA clone encoding human PF4, and to Dr. Carolyn Keever-Taylor (Medical College of Wisconsin) for kindly providing the OKT3 mAb.
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 Grant HL64704 from the National Heart, Lung, and Blood Institute.
2 Current address: University of Ulsan College of Medicine, Department of Laboratory Medicine, Ulsan University Hospital, 290-3 Jeonha-dong, Dong-gu, Ulsan 682-714, South Korea.
3 Address correspondence and reprint requests to Dr. Gian Paolo Visentin, Department of Pediatrics, University at Buffalo-State University of New York, Biomedical Research Building, Room 422, Buffalo, NY 14214. E-mail address: visentin{at}buffalo.edu
4 Abbreviations used in this paper: Tr, T regulatory; HIT, heparin-induced thrombocytopenia; PF4, platelet factor 4; PHA, phytohemagglutinin.
Received for publication August 26, 2004. Accepted for publication December 13, 2004.
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