IFN-α-2b, known as potent immune modulator, can either inhibit or enhance immune cell activity within the tightly regulated microenvironment of inflammation, depending upon the concentration of the cytokine and the activation stage of the cell. Chemokine receptors, which not only mediate chemotaxis of immune cells to the site of inflammation but also affect cellular activation by transferring corresponding signals, represent yet another level of immune regulation. Here we demonstrate that IFN-α increases the expression of CCR1 and CCR3 in primary mononuclear phagocytes, as well as in the monocytoid cell line U937. Enhanced receptor mRNA expression correlated with functional readouts such as increased intracellular calcium mobilization and cell migration in response to ligands. Expression of CCR2b, CCR4, CCR5, and CXCR4 was unchanged or decreased after IFN-α treatment. These observations indicate a differentially regulated cellular signaling relationship of IFN-α pathways and chemokine receptor expression. We also provide evidence that, under these conditions, IFN-α treatment increased the expression of CD95 (Fas, Apo1), resulting in enhanced susceptibility to apoptosis. Taken together, these data add important information for the rational application of IFN-α (2b) in immune and cancer therapies.

Interferon-α-2b, a member of the cytokine family that was first defined by its ability to establish an antiviral state in cells, was also shown to modulate a variety of physiologic processes that appear to be divergent. Whether a cell becomes more activated and will be induced to proliferate and differentiate or whether antiproliferative, anergic, and apoptotic processes are mediated instead critically depends upon the preactivation stage of a cell (e.g., IFN receptor balance, costimulatory signals) and the concentration of IFN-α within a microenvironment (1). Convergently, IFN-α has been shown to have proinflammatory effects, but has also been used to inhibit chronic inflammations (2). Whereas in vitro systems can facilitate dissection of the different cellular pathways involved, these results might be difficult to interpret with regard to physiologic relevance.

Although often quite complex, clinical data, for which IFN-α-2b was successfully used as a treatment modality in cancer, chronic inflammation, and viral infection, provide a most interesting source of information. Opinions vary markedly about the underlying mechanism that leads to the commonly observed IFN-α-mediated clinical effects. Experimental data provide evidence for the support of several hypotheses. Whether the antitumor effects of IFN-α relate to induced cell differentiation into a stage accessible to immune surveillance or to the control of cell cycling and therefore cell growth inhibition remains unresolved. However, there is also experimental evidence that IFN-α enhances the activation of a cell so as to affect apoptosis as mediated through the CD95 (Fas, Apo1) signaling pathway (3, 4).

It has been shown in some experimental systems that IFN-α can induce the differentiation of cells and enhance the expression of integrin (β1) and integrin receptors (5), thereby facilitating cell adhesion and affecting cell trafficking. Chemokines and chemokine receptors (CRs)3 are important mediators of signals that regulate leukocyte trafficking as well (6, 7). The level of CR expression on a cell affects responses to its specific ligands, because resting immune cells with a low density of appropriate receptors may not be competent to migrate and adhere even in the presence of high concentrations of chemokines. We were interested to find out whether IFN-α-2b is involved in the process of modulating CR expression and signaling by a cell. Because the modulation of APCs by IFN-α results in a complex alteration of their Ag-presenting functions (8, 9, 10), we specifically investigated human monocyte-derived cell subsets as well as the monocytoid cell line U937 to assess the expression and function of CRs subsequent to IFN-α-2b priming. In addition, we investigated whether changes in CD95 apoptosis receptor expression and changes in susceptibility to apoptosis as induced through CD95 are linked to these events.

Peripheral blood samples were obtained from healthy volunteers according to the protocols approved by the Institutional Review Board. PBMCs were isolated by automated Ficoll/Hypaque density-gradient centrifugation using a CS-3000 Plus Blood Cell Separator (Fenwal Division, Baxter Healthcare, Deerfield, IL). Monocytes were isolated from PBMCs by counterflow-centrifugal elutriation using a Beckman JE-5.0 rotor and a type A chamber (Beckman Instruments, Palo Alto, CA). The purity of the separated monocyte fraction was ∼98% as determined by cytomorphology in Pappenheim stain and ∼96% as determined by the expression of CD14 Ag (LeuM3; Becton Dickinson, Mountain View, CA) and measured by flow cytometry. U937 cells were obtained from the American Type Culture Collection (Manassas, VA). Cells were cultured in complete medium (RPMI 1640, containing 10% heat-inactivated FCS, low endotoxin grade; all Life Technologies, Gaithersburg, MD) at a concentration of 0.8 × 106 cells/ml. IFN-α-2b (Schering, Kenilworth, NJ) was added to the cultures in varying concentrations so as to assess dose-dependent effects. A concentration of 300 IU/ml was determined to be sufficient for mediating effects on CR expression without toxic side effects and was therefore used in most of the experiments. Aliquots of differently treated cells were collected for mRNA analysis, calcium flux assays, cell surface staining, and FACS analysis or migration assays. In apoptosis experiments, the anti-CD95 mAb CH11 (Kamyia, San Francisco, CA), an Ab that mimics the Fas ligand by triggering the CD95 receptor, or mAb ZB4, a CD95 blocking Ab (AMAC, Westbrook, ME), were used at concentrations ranging from 0.01 μg/ml to 0.5 μg/ml. Cell viability was measured using a standard trypan blue (Life Technologies) exclusion test. In some experiments, an automated colorimetric assay based on tetrazoliumbromide (MTT) reduction was used to determine cell number and cell metabolic rates. Levels of the chemokine RANTES were determined in culture supernatants derived from monocyte-derived mononucleated phagocytes (MDMs) or U937 cells using an ELISA system (R&D Systems, Minneapolis, MN) according to the manufacturer’s recommendations. Samples were stored at <80°C until they were assayed.

A method that has been described previously was applied (11). Briefly, aliquots of 106 cells were incubated for 20 min at 37°C with Fluo-3, reconstituted with Pluronic F-127 (Molecular Probes, Eugene, OR) according to the manufacturer’s directions. After incubation, the samples were washed once with RPMI 1640 (without phenol red, without sodium carbonate, containing 25 mM HEPES) and finally resuspended in 1 ml of the same medium. Data were acquired with a FACScalibur (Becton Dickinson), with excitation at 488 nm. Cells were gated by forward and side scatter properties. Calcium mobilization was determined by a two-parameter density plot, measuring linear emission at 530 nm in the FL-1 window for the gated cell population over time. After 20 s of acquisition, the appropriate chemokine (25 ng/ml of RANTES, macrophage inflammatory protein-1α (MIP-1α), MIP-1β, or stromal cell-derived factor-1α (SDF-1α); all from R&D Systems) or 100 ng/ml of the ionophore ionomycin (Sigma, St. Louis, MO) were added.

Cell migration assays were performed as described previously (12). Briefly, 1.5 × 105 U937 cells in 150 μl of RPMI 1640 medium containing 0.25% human serum albumin were transmigrated through 5-μm pore size bare filter transwell inserts (Costar, Cambridge, MA) for 12 h. Migrated cells were counted by FACS analysis scatter-gating on lymphocytes. Chemotaxis was conducted in the presence of optimized ligand concentrations (RANTES, 100 ng/ml; MIP-1α, 100 ng/ml; eotaxin, 200 ng/ml; MIP-1β, 250 ng/ml; SDF-1α, 250 ng/ml; all R&D Systems). Three independent experiments were performed in duplicate cultures; mean values (± SD) were determined as expressed as a percentage as compared with untreated controls.

Semiquantitative mRNA assays were performed using end-point dilution and direct comparison of partner samples as is recommended under circumstances in which no internal control/competitor is available. Aliquots of differently treated cells were collected at days 2 and 5. Total cellular RNA was isolated by TRIzol LS reagent (Life Technologies) according to the manufacturer’s protocol. Samples of RNA were treated by DNase I (amplification grade, Life Technologies). Synthesis of first-strand cDNA was performed in 20 μl of reaction mix (50 mM Tris-HCl (pH 8.3), 75 mM KCl, 3 mM MgCl, 10 mM DTT, 500 μM each of dNTPs, and 2.5 μM of random hexamer primer, Perkin-Elmer Applied Biosystems, Branchburg, NJ), 1.5 μg of RNA, and 200 U of SuperScript II RT (Life Technologies) for 1 h at 42°C, followed by heating for 5 min at 99°C. Amplification was performed with 2 μl of RT mixture in a total volume of 20 μl, using 2 μM dNTPs, 8 pmol of primers, and 2.5 U of AmpliTaq AGold polymerase (Perkin-Elmer Applied Biosystems) for 35 cycles (94°C for 45 s, 60°C for 1.30 min, 72°C for 2 min) after an initial 9-min denaturation at 94°C. The resulting PCR products were separated on 2% SeaKem GTG agarose gel (FMC BioProducts, Rockland, ME). As a control for DNA contamination, equal amounts of RNA extraction products were used for each sample assessed, and PCR amplification was performed without the addition of RT to the first-strand synthesis. The conditions for the semiquantitative mRNA assay were specifically defined so as to exclude the possibility of amplifying contaminating genomic DNA. The sets of primers used have been described previously (13).

For a qualitative and quantitative determination of cell surface molecules, flow cytometry was conducted by incubating cells at 4°C in PBS containing 1% BSA and 5 mM EDTA for 30 min with Abs at concentrations recommended by the manufacturer after cells had been washed and incubated with 10% normal goat serum in PBS to block unoccupied sites when indicated. Abs to CXCR4 (mouse IgG2a, clone 12G5, obtained from the AIDS Research and Reference Reagents Program, Rockville, MD) were developed with anti-IgG2a (goat-anti-mouse, FITC-labeled, human-adsorbed; Caltag Laboratories, Burlingame, CA). The mouse IgG2a isotype control Abs were obtained from Coulter (Hialeah, FL). PE-conjugated mAbs to CD4 and CD14, as well as appropriate isotype control Abs, were obtained from Becton Dickinson. The FITC-conjugated anti-CD95 mAb UB2 and appropriate isotype control Abs were obtained from PharMingen (San Diego, CA). The receptor density for MIP-1α was determined by flow cytometry using the Fluorokine staining kit (R&D systems) according to the manufacturer’s recommendations.

DNA analysis of cell cycle status was performed using flow cytometry. After culture, cells were harvested, washed with PBS containing 2% FCS, fixed with ice-cold 70% ethanol, rewashed, and resuspended with 10 μg/ml propidium iodide solution (Boehringer Mannheim, Indianapolis, IN) containing 10 μg/ml bovine RNase (Boehringer Mannheim). After 30 min, DNA content was measured by flow cytometry. Collected data were analyzed using M-Cycle software, version 2.5 (Phoenix Flow Systems, Phoenix, AR). This method allows determination of cell cycle status and quantification of apoptotic cells by measurement of the corresponding peak. A TUNEL assay (Apotag; Oncor, Gaithersburg, MD) also was used to quantitate by flow cytometry the number of cells undergoing apoptosis. In some experiments, apoptosis was also evaluated by using the Cell Death Detection ELISAplus (Boehringer Mannheim), which quantitates cytoplasmic histone-associated DNA fragments.

We first tested the hypothesis that IFN-α-2b influences the expression of RANTES receptors (CCR1, CCR3, CCR4, and CCR5) on primary MDMs. Elutriated monocytes were primed in the presence of IFN-α-2b for 6 days as described in Materials and Methods. Because CRs are 7-transmembrane, G-coupled proteins that mediate intracellular calcium mobilization, the adherent cell subpopulations (MDMs) were analyzed for their ability to respond to the ligand RANTES, using an intracellular calcium flux assay. Subsequent to IFN stimulation, an increase in the level of signal (Fig. 1, upper right panel) was observed in MDMs when compared with untreated controls (Fig. 1, upper left panel). The calcium mobilization response to ionomycin remained unchanged (Fig. 1, compare lower left panel with lower right panel), indicating a specific regulation of RANTES receptors by the treatment. Simultaneously, we observed IFN-α-induced changes in cell morphology as determined by size and granularity using flow cytometry (FACS) (Fig. 2, compare A and B). A distinct subpopulation of larger cells with a comparatively high granularity (Fig. 2 B, defined as R1) was detected only in IFN-α-treated monocyte cultures.

FIGURE 1.

IFN-α-2b treatment of elutriated monocytes increases transient calcium flux in response to RANTES. Monocytes (untreated, left panel) were compared with IFN-α-treated monocytes (right panel) derived from the same donor in their response to RANTES (upper panel) and ionomycin (IONO, lower panel) as assessed by flow cytometry. Characteristic differences in chemokine response were detected after 3 days of treatment, and the illustrated pattern of the response is representative of cells derived from four different individuals.

FIGURE 1.

IFN-α-2b treatment of elutriated monocytes increases transient calcium flux in response to RANTES. Monocytes (untreated, left panel) were compared with IFN-α-treated monocytes (right panel) derived from the same donor in their response to RANTES (upper panel) and ionomycin (IONO, lower panel) as assessed by flow cytometry. Characteristic differences in chemokine response were detected after 3 days of treatment, and the illustrated pattern of the response is representative of cells derived from four different individuals.

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FIGURE 2.

IFN-α treatment of monocytes results in a morphologically distinct subpopulation of cells that is highly responsive to RANTES. Monocytes were treated with 300 IU/ml of IFN-α for 5 days and subsequently analyzed by flow cytometry as described in Materials and Methods. Untreated (A) vs treated (B) cells were compared by scattergram, and the R1 region was analyzed for transient calcium flux in response to RANTES. One representative experiment of four is illustrated.

FIGURE 2.

IFN-α treatment of monocytes results in a morphologically distinct subpopulation of cells that is highly responsive to RANTES. Monocytes were treated with 300 IU/ml of IFN-α for 5 days and subsequently analyzed by flow cytometry as described in Materials and Methods. Untreated (A) vs treated (B) cells were compared by scattergram, and the R1 region was analyzed for transient calcium flux in response to RANTES. One representative experiment of four is illustrated.

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These data provide evidence that IFN-α, produced as a primary response to inflammation/infection, modifies CR-mediated signals in MDMs, a subset of monocyte-derived APCs. To extend these observations, we used an in vitro model in which the majority of the conditions are standardized and the data can be reproduced, without the variability associated with primary cells. Thus, we analyzed the monocytoid (CD14+, CD4+, HLA-DR+) cell line U937 to determine whether the observed changes in transient calcium flux responses to RANTES were comparable with those observed in primary cells. Only a small proportion (∼5%) of unprimed U937 cells responded minimally to RANTES and MIP-1α (ligands to CCR1, CCR3, CCR4, and CCR5), whereas no response was observed with MIP-1β (ligand to CCR5). After IFN-α priming, an increase in transient intracellular calcium flux response was observed to both RANTES (Fig. 3,A) and MIP-1α, but not to MIP-1β (data not shown). IFN-α treatment caused >30% of the cells to respond to RANTES, as determined by transient calcium flux. This effect was dose-dependent (Fig. 3, compare A, B, and C in the upper panel) and was reduced to baseline levels by neutralizing (polyclonal) anti-IFN-α Abs (data not shown). Untreated and IFN-treated cells responded in a similar manner to the Ca2+ ionophore ionomycin (Fig. 3, lower panel, compare A, B, and C). This confirmed the receptor/ligand-specific modulation of calcium mobilization by IFN-α.

FIGURE 3.

Increased transient calcium flux in response to RANTES as primed by IFN-α is dose-dependent. Response to RANTES (upper panel), as determined by flow cytometry in U937 cells treated with 30 IU/ml (3 ng/ml, B) and 300 IU/ml (30 ng/ml, C) of IFN-α-2b for 3 days, was compared with untreated controls (A). The lower panel illustrates that calcium flux responses to ionomycin (IONO) are not affected by IFN-α treatment (left to right: untreated, 30 IU/ml, and 300 IU/ml of IFN-α-2b). Data are representative of three independent experiments.

FIGURE 3.

Increased transient calcium flux in response to RANTES as primed by IFN-α is dose-dependent. Response to RANTES (upper panel), as determined by flow cytometry in U937 cells treated with 30 IU/ml (3 ng/ml, B) and 300 IU/ml (30 ng/ml, C) of IFN-α-2b for 3 days, was compared with untreated controls (A). The lower panel illustrates that calcium flux responses to ionomycin (IONO) are not affected by IFN-α treatment (left to right: untreated, 30 IU/ml, and 300 IU/ml of IFN-α-2b). Data are representative of three independent experiments.

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To further define the specificity of IFN-α-induced changes in CR expression and to possibly distinguish responses to different ligands, cell migration assays were used to determine whether treatment with IFN-α enhances chemotaxis in response to the chemokines RANTES, MIP-1α, eotaxin, MIP-1β, and SDF-1α. In this assay, the ability of U937 cells to migrate in culture through a membrane in response to different stimuli is determined by enumerating the cells in both sides of the culture chamber after 12 h. IFN-α-2b pretreatment (300 IU/ml, for 4 days) induced an increase in cell migration to ∼300% in response to RANTES (100 ng/ml), to ∼200% in response to MIP-1α (100 ng/ml), and to ∼150% in response to eotaxin (200 ng/ml) when compared with untreated controls that were exposed to the same chemokine. No significant changes in cell migration in response to MIP-1β (250 ng/ml) or to SDF-1α (250 ng/ml) were observed (Fig. 4).

FIGURE 4.

IFN-α treatment selectively modulates chemotaxis of APC subsets. Chemotaxis was analyzed using transwell migration of U937 cells in response to RANTES (ligand to CCR1, CCR3, and CCR5), MIP-1α (CCR1, CCR5), eotaxin (CCR3), MIP-1β (CCR5), and SDF-1α (CXCR4), as described in Materials and Methods. Changes in migration are illustrated as the percent increase over the control cultures that were not treated with IFN-α but were exposed to the corresponding chemokine. Results of four independent experiments were combined; mean values ± SEM are depicted.

FIGURE 4.

IFN-α treatment selectively modulates chemotaxis of APC subsets. Chemotaxis was analyzed using transwell migration of U937 cells in response to RANTES (ligand to CCR1, CCR3, and CCR5), MIP-1α (CCR1, CCR5), eotaxin (CCR3), MIP-1β (CCR5), and SDF-1α (CXCR4), as described in Materials and Methods. Changes in migration are illustrated as the percent increase over the control cultures that were not treated with IFN-α but were exposed to the corresponding chemokine. Results of four independent experiments were combined; mean values ± SEM are depicted.

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To determine whether increased intracellular calcium mobilization and enhanced migration ability correlate with CR expression, mRNA levels were assessed in IFN-treated monocytes. Application of a standardized RT-PCR method revealed differential modulation of CR mRNA expression by IFN-α-2b. The cytokine induced a marked increase in CCR1 and CCR3 expression as determined at 2 days after the initiation of treatment. The level of CXCR4 and CCR5 expression remained unchanged (Fig. 5,A). Similar results were obtained in experiments using U937 cells. The expression of CCR1 and CCR3 was increased, whereas the expression of CCR5 and CXCR4 remained unchanged under IFN-α-2b treatment (Fig. 5 B). No specific messages for CCR2b and CCR4 were detectable by means of RT-PCR amplification, independent from treatments with IFN-α (data not shown).

FIGURE 5.

IFN-α treatment increases the levels of CCR1 and CCR3 mRNA, as determined by standardized RT-PCR. RNA was purified from the differently treated samples (containing the same amount of cells) after 2 days and was assessed as described in Materials and Methods. The influence of possible genomic DNA contamination was excluded by control amplifications of all samples in the absence of retro transcription steps. Amplification of the 18S-ribosomal RNA band was used to ensure comparability of samples. Results from primary MDMs (A) and U937 cells (B) are shown. Depicted data are representative of four independent experiments in which cells from three different donors were analyzed.

FIGURE 5.

IFN-α treatment increases the levels of CCR1 and CCR3 mRNA, as determined by standardized RT-PCR. RNA was purified from the differently treated samples (containing the same amount of cells) after 2 days and was assessed as described in Materials and Methods. The influence of possible genomic DNA contamination was excluded by control amplifications of all samples in the absence of retro transcription steps. Amplification of the 18S-ribosomal RNA band was used to ensure comparability of samples. Results from primary MDMs (A) and U937 cells (B) are shown. Depicted data are representative of four independent experiments in which cells from three different donors were analyzed.

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To evaluate whether increased Ca2+ flux, mRNA expression, and chemotaxis correlated with the surface expression of CRs, we stained IFN-treated monocytes with fluorescein-labeled MIP-1α (specific ligand to CCR1 and CCR5) and compared these samples with untreated cells by flow cytometry. A marked increase in fluorescence intensity was observed in cell populations that had been treated with IFN-α-2b (Fig. 6, left panel), indicating a higher cell surface density of CCR1 and/or CCR5. Unchanged surface staining with MIP-1β (ligand to CCR5) indicated that the difference in the labeling with MIP-1α is due to an increase in CCR1 (data not shown). To assess the effects of IFN-α treatment on the surface receptor density of CXCR4, a receptor-specific Ab stain was used. FACS analyses revealed a slight decrease in CXCR4 expression in IFN-α treated cells (Fig. 6, right panel).

FIGURE 6.

Flow cytometry assessment of CR expression on the surface of monocyte-derived cells. Expression of CCR1 was increased after 4 days of treatment with IFN-α-2b, as determined by labeling with MIP-1a (left panel). No significant differences between IFN-treated and control cells were observed in CXCR4 surface expression (right panel). Four independent experiments were performed with similar results.

FIGURE 6.

Flow cytometry assessment of CR expression on the surface of monocyte-derived cells. Expression of CCR1 was increased after 4 days of treatment with IFN-α-2b, as determined by labeling with MIP-1a (left panel). No significant differences between IFN-treated and control cells were observed in CXCR4 surface expression (right panel). Four independent experiments were performed with similar results.

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Because we observed relative decreases in cell density that evolved in cultures of U937 cells under IFN-α-2b treatment (when compared with untreated cultures) that were not able to be explained by toxicity, the cell numbers (using trypan blue exclusion) as well as cell metabolic rates (using the MTT assay (14)) were determined. After 5 days, cultures that were treated with 300 IU/ml IFN-α contained 29 ± 4% fewer cells than the controls. However, when the metabolic rates as determined by MTT assays were compared, only a 17 + 6% reduction was observed in IFN-α-treated cultures (data not shown). To determine whether the lower cell number in primed U937 cultures was due to apoptosis-related cell death, we applied a TUNEL assay and found no significant difference in the percentage of apoptotic cells in the two differently treated groups under the described culture conditions (p > 0.5, determined in two independent experiments). Application of the Cell Death Detection ELISAplus (Boehringer Mannheim) to cell culture supernatants and to cell lysates confirmed that lower cell density in IFN-α-treated U937 cell cultures was not caused by induction of cell death.

The cell viability data and the metabolic assays indicated that IFN-α-2b decreases cell proliferation rates. Therefore, we performed cell cycle analysis using flow cytometry. Propidium iodide staining of the cells allowed assessment of DNA content during the cell cycle, distinguishing G0/G1-phase, S-phase, and G2/M-phase peaks. Patterns of DNA content of cells (identified as viable by gating) were compared between treatment groups, and the relative amount of cells within the different peaks was calculated. IFN-α treatment of U937 cells for 3 days (300 IU/ml IFN-α) resulted in a 13 ± 3% increase of cells that were at G0/G1 phase (data not shown).

We were also interested in evaluating whether expression of the activation marker CD95, a receptor known to mediate signals that can lead to apoptosis, was modulated by IFN-α-2b in our system, especially because different researchers reported distinct outcomes in various experimental systems (3, 4). Flow cytometry analysis revealed a slight increase in CD95 expression on U937 cells that were treated with IFN-α (300 IU/ml, for 4 days; Fig. 7,A). In comparison, however, this was a far less intense increase in CD95 receptor density than that induced by IFN-γ (30 ng/ml, 3 days of treatment) (Fig. 7,B). When both IFN-α and IFN-γ were combined at similar concentrations, a further increase in CD95 expression was noted (Fig. 7 C). Changes in CD95 expression as induced by IFN-α-2b or IFN-γ were dose-dependent and were not detectable by means of FACS analysis when IFN-α was used as single treatment at concentrations of <10 IU/ml (1 ng/ml for IFN-γ, data not shown).

FIGURE 7.

Increase of CD95 expression on U937 cells as mediated by IFN-α and IFN-γ. Changes in cell surface expression of CD95 were detected by flow cytometry at 3 days after treatment with IFN-α-2b (300 IU/ml) (A), IFN-γ (30 ng/ml) (B), and the combination of both IFN-α and IFN-γ (C). Experiments were repeated three times with similar results.

FIGURE 7.

Increase of CD95 expression on U937 cells as mediated by IFN-α and IFN-γ. Changes in cell surface expression of CD95 were detected by flow cytometry at 3 days after treatment with IFN-α-2b (300 IU/ml) (A), IFN-γ (30 ng/ml) (B), and the combination of both IFN-α and IFN-γ (C). Experiments were repeated three times with similar results.

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An increase in CD95 expression per se does not indicate whether a cell becomes more susceptible to signals mediated by the receptor. Consequently, a CD95 agonistic mAb (CH11, 0.25 μg/ml) was used to mimic CD95 ligand binding and signaling. Exposure to CH11 for 3 days increased the rate of apoptosis as detected by MTT assays from 28 ± 7% in unprimed cell cultures to 63 ± 12% in IFN-α-2b-pretreated U937 cells (500 IU/ml, for 4 days) when compared with untreated controls. When cells were primed with both IFN-α and IFN-γ (50 ng/ml) under these culture conditions, only 8 ± 2% of cells remained viable. Titration experiments in which IFN-α, IFN-γ, and CH11 were evaluated separately and in combination revealed that IFN-α and IFN-γ have at least additive effects on CD95-mediated apoptosis induction in U937 cells (data not shown).

Because inflammatory responses are regulated by “pro and contra” signals and concepts of autocrine loops as well as negative feed backs have been described for a variety of cellular receptors upon interaction with their ligands, we investigated the effect of IFN-α-2b on chemokine production. Theoretically, the RANTES receptor CCR1 and CCR3 expression and signaling (that we found to be increased) could be modulated by RANTES production as induced by IFN-α. We measured RANTES in cell culture supernatants after 6 days of IFN-α treatment and found levels that were reduced to 74 ± 15% of levels in untreated controls (37 ± 8 ng RANTES/106cells).

The differential regulation of CR expression and related signaling represents one of the means of modulating leukocyte trafficking and inflammation. Known immune-modulatory functions of IFN-α include the enhancement of cellular adhesion molecule expression (15). It was not known, however, that the modulation of cell adhesion by IFN-α as observed in monocytoid cells involves the differential regulation of CR expression. We have now demonstrated that IFN-α-2b can enhance CCR1 and CCR3 expression on APC subsets. These events appear to be specifically related to the pathways involved in IFN-α signal transduction. Alternatively, CR modulation could be the result of a feedback regulation to the IFN-α-induced reduction of chemokine production. Although our experimental setting did not allow for the exclusion of one or the other possibility, our results illustrate that CR expression and signaling can be differentially regulated by IFN-α, because receptors such as CCR2b, CCR4, CCR5, or CXCR4 were not affected or found to be decreased.

Our study supports a paradigm in which locally produced cytokines can regulate the quantity and the quality of leukocytes trafficking to the site of action as mediated by both the chemokines and their receptors (16). Moreover, increased systemic levels of cytokines such as IFN-α may change the overall activation of leukocytes by priming more cells into a stage of enhanced susceptibility to chemokines that are produced at defined anatomic sites. Our data provide evidence that the recruitment and maturation of APC precursors is regulated in a similar manner, in that increased levels of IFN-α act by differentially increasing CR subset expression. This priming effect, in turn, may also contribute to some differentiation steps of monocytes in the development toward mature APCs. In support of our hypothesis, involvement of IFN-γ in the maturation of monocytes that includes modulation of CCR5 has recently been demonstrated (13, 17). Thus, morphological changes in primary monocyte-derived cells treated with IFN-α, which we detected by flow cytometry, may indicate comparable processes. A better functional and phenotypic characterization of these cell subsets remains an issue of future investigations.

A number of cytokines and growth factors (including IL-2, IL-4, IL-10, IL-12, and IL-13, as well as GM-CSF and IFN-γ (13, 18, 19, 20, 21, 22, 23, 24)) were found to have CR-modulating properties. The experimental systems that were used to detect changes in CR expression and signaling, however, vary. Effects on the regulation of gene transcription, the CR mRNA stability, or the cell surface receptor expression and the rate of internalization were reported to take place within minutes but can require up to several days. In our experimental system, the maximal effect of IFN-α on enhanced CCR1 and CCR3 expression was detected between days 2 and 5 posttreatment. This time interval necessary for response may indicate an indirect modulation, possibly as a feedback mechanism, especially because levels of newly produced RANTES, a ligand for CCR1, CCR3, and CCR5 were found to be slightly decreased compared with cultures that were not treated with IFN-α-2b. Consequently, further studies are required to define the direct connection between intracellular pathways of IFN-α-induced signals and CR expression.

We also provide evidence that IFN-α may be involved in the process of cell survival-regulation by mediating susceptibility to apoptosis in activated immune cells of monocytoid origin. This was demonstrated by the enhanced CD95 expression and increased susceptibility of U937 cells to CD95-triggered apoptosis. A further increase of CD95 expression and response to related signaling was observed by a combination of IFN-α-2b and IFN-γ treatment, perhaps an indication of partially overlapping signal transduction for both type I and type II IFN (25, 26). A relationship of IFN-γ-induced signals and CD95 expression is well established (27, 28). We have now demonstrated such an interaction for IFN-α-2b and the CD95 system in monocytoid cells in vitro, supporting the rational of using the augmenting effects of both type I and type II IFN for developing new combination treatment approaches to cancer.

It has been proposed that IFN-α-induced apoptosis may occur in cells during all cell cycle stages; therefore, apoptosis and cell cycle arrest would represent independent responses to IFN-α (29, 30). The high degree of cell death that was inducible through CD95 receptor triggering in our experimental system supports the previously observed independence of some apoptosis pathways from cell cycle stages. In addition to confirming that IFN-α affects cell cycling, our results also indicate that these events are related to the APC differentiation and maturation process by involving changes in CR expression and function. Although further studies are necessary to elucidate the mechanisms that control the differential regulation of CR expression in greater detail, the newly identified immunoregulatory effect of IFN-α-2b on APCs as related to CR expression and function may help in understanding the complex pathophysiologic role of this cytokine and may contribute to the development of alternative immune-modulating therapeutic strategies in malignancies.

1

This work was supported in part by National Institutes of Health Grant R03-AI43204.

3

Abbreviations used in this paper: CR, chemokine receptor; MDM, monocyte-derived mononucleated phagocyte; MIP, macrophage inflammatory protein; SDF, stromal cell-derived factor.

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