Human Macrophages Constitute Targets for Immunotoxic Inorganic Arsenic

Anthony Lemarie, Claudie Morzadec, Emilie Bourdonnay, Olivier Fardel and Laurent Vernhet
J Immunol September 1, 2006, 177 (5) 3019-3027; DOI: https://doi.org/10.4049/jimmunol.177.5.3019

Abstract

Chronic exposure to inorganic arsenic, a widely distributed environmental contaminant, can lead to toxic effects, including immunosuppression. Owing to the established roles of human macrophages in immune defense, we determined, in the present study, whether inorganic arsenic can affect these major immune cells. Our results demonstrate that noncytotoxic concentrations of arsenic trioxide (As2O3), an inorganic trivalent form, markedly impair differentiated features of human blood monocyte-derived macrophages. First, treatment of macrophages with 1 μM As2O3 induced a rapid cell rounding and a subsequent loss of adhesion. These morphologic alterations were associated with a marked reorganization of actin cytoskeleton, which includes retraction of peripheral actin extensions and formation of a cortical actin ring. In addition, As2O3 reduced expression of various macrophagic surface markers, enhanced that of the monocytic marker CD14, and altered both endocytosis and phagocytosis; unexpectedly, exposure of macrophages to the metalloid also strongly potentiated expression of TNFα and IL-8 induced by LPS. Finally, like monocytes, As2O3-treated macrophages can be differentiated into dendritic-like cells. Impairment of macrophage function by As2O3 mainly resulted from activation of a RhoA/Rho-associated kinase pathway; indeed, pretreatment of macrophages with the Rho-associated kinase inhibitor Y-27632 prevented metalloid effects on cytoskeleton and phagocytosis. Moreover, As2O3 was found to increase level of the active GTP-bound form of RhoA and that of phosphorylated-Moesin, a major cytoskeleton adaptor protein involved in RhoA regulation. Taken together, our results demonstrated that human macrophages constitute sensitive targets of inorganic arsenic, which may contribute to immunotoxicity of this environmental contaminant.

Inorganic arsenic (iAs)3 is a potent environmental toxic to which millions of people are exposed over the world, mainly through contaminated drinking water (1). Epidemiological studies have demonstrated that long-term exposure to iAs can favor development of cardiovascular diseases such as atherosclerosis (2); it can also induce cancer of skin, bladder, lung, and, possibly, of other internal organs (3, 4). iAs particularly increases incidence of nonmelanoma skin cancers, a known complication of organ transplantation along with the use of immunosuppression (5, 6). Interestingly, it was recently reported that iAs can exert immunosuppressive effects that may contribute to its general toxicity. Indeed, this metalloid was recently demonstrated to affect normal function of lymphocytes in chronically exposed humans (7). In these individuals, increased urine arsenic levels were associated with reduced CD4 lymphocyte population, proliferative response to PHA, and IL-2 secretion. Moreover, low micromolar concentrations of arsenite, an inorganic trivalent arsenical compound, delays in vitro proliferation of activated human T lymphocytes by reducing production and secretion of IL-2 (8). Alterations of lymphocyte functions have also been observed in animal models. Lymphocytes, from BALB/c mice exposed to 50 mg/L arsenic in the drinking water for 4 wk, exhibited reduced proliferative responses following T cell mitogen stimulation (9); in addition, treatment of B6C3F1 mice with gallium arsenide (200 mg/kg) decreased lymphocyte populations (10).

Besides lymphocytes, monocytes/macrophages may also constitute major targets of iAs. Indeed, we and others have recently demonstrated that low micromolar concentrations of iAs inhibit in vitro macrophagic differentiation of human blood-derived monocytes (11, 12). Particularly, we showed that arsenic trioxide (As2O3), an iAs trivalent form, triggers apoptosis of human monocytes during their differentiation by down-regulating NF-κB-dependent survival pathways (11). In addition, in vivo experiments have reported that iAs can alter macrophage functions. Thus, treatment of Swiss albino mice with arsenite (0.5 mg/kg) for 15 days significantly increased bacterial load in blood and delayed bacterial clearance by spleen (13); these effects were associated with marked reduction of adhesion property, chemotactic migration, and phagocytic activity of splenic macrophages. Mice exposure to gallium arsenide affected proteolytic cathepsin activities and Ag processing of splenic macrophages (14). Together, these results suggest that, in addition to lymphocytes, human macrophages may constitute by themselves sensitive targets of iAs. The present study was designed to investigate this hypothesis.

In this work, we demonstrate that low concentrations of As2O3 (0.25–1 μM), in the range of arsenic blood levels measured in chronically exposed humans (10–60 μg/L) (15, 16), alter differentiation features of human primary macrophages such as morphology, surface marker expression, or phagocytic activity. These effects were initiated by a rapid reorganization of F-actin cytoskeleton, resulting from a sustained and persistent activation of a RhoA-Rho-associated kinase (ROCK) pathway. Our results highlight human macrophages as marked sensitive targets of iAs, which may contribute to the immunosuppressive properties of this environmental contaminant.

Materials and Methods

Chemical reagents and Abs

As2O3, FITC-conjugated dextran (molecular mass, 40,000 kDa), FITC-phalloidin, and LPS were purchased from Sigma-Aldrich. The WST-1 tetrazolium salt 4-[3–4(iodophenyl)-2-(4-nitrosophenyl)-2H-5-tetrazole]-1,3-benzene disulfonate was purchased from Roche Diagnostics. Y-27632 was from Calbiochem (VWR). GM-CSF (specific activity, 1.2 × 108 UI/mg) was obtained from Shering Plough, whereas IL-4 (specific activity, 2 × 108 UI/ml) was from Promocell. Rabbit polyclonal Abs against phospho- and total Ezrin/Radixin/Moesin (ERM) proteins were purchased from Cell Signaling Technology (Ozyme). Mouse mAb raised against Rho guanine dissociation inhibitors (RhoGDI) was from Santa Cruz Biotechnology (Tebu-bio). FITC-conjugated mAbs against CD1a, CD11c, CD14, CD29, and CD71, and PE-conjugated mAb against CD11b were purchased from Immunotech. Fluorescent latex microspheres were provided by Polysciences.

Cell cultures

PBMC were first isolated from bloody buffy coats of healthy donors through Ficoll gradient centrifugation. Human monocytes were then prepared by a 2-h adhesion step, which routinely obtained >90% of adherent CD14-positive cells as assessed by immunostaining. These monocytic cells were next cultured for 6 days in RPMI 1640 medium supplemented with 10% FCS, 2 mM l-glutamine, 20 UI/ml penicillin, and 20 μg/ml streptomycin, in the presence of 800 UI/ml GM-CSF to get macrophages as previously reported (17, 18, 19). To obtain dendritic cells, monocytic cells were differentiated in the presence of 800 UI/ml GM-CSF and 500 UI/ml IL-4, as previously reported (20). Once differentiated, macrophages were cultured in GM-CSF-free RPMI 1640 medium in the absence or presence of As2O3 for indicated time intervals. In some experiments, macrophages were first pretreated with 20 μM Y-27632 for 2 h, and then exposed to As2O3. Y-27632 was dissolved in distilled water. For cell death assays and flow cytometric studies, both floating and adherent macrophages were analyzed. Adherent macrophages were collected after 15 min of incubation at 37°C in PBS supplemented with 100 μM EDTA.

Cellular adhesion assay

Adherence of macrophages was analyzed using the WST-1 assay in 96-well microplates, as previously described (18). The yellow formazan product formed by viable adherent cells was quantified by detection of its absorbance at 450 nm using a Titertek Multiskan spectrophotometer (Flow Laboratories).

Determination of apoptosis and necrosis

To analyze alterations of the plasma membrane structure linked to apoptosis, exposition of phosphatidylserine to the extracellular environment was studied. We determined binding of annexin V (A5), a calcium-dependent protein with high affinity for phosphatidylserine, using A5 conjugated to the fluorescent label Alexa 568 (Roche Diagnostics). Simultaneously, necrotic cells, which have lost their plasma membrane integrity, were detected with the green fluorescent DNA dye sytox green (SG) (Interchim). After treatment, cells were collected, washed, and incubated with dyes as previously described (21). Apoptotic (A5+SG−) and necrotic (A5−SG+ and A5+SG+) cells were quantified using a fluorescence Olympus BX60 microscope, in comparison with living cells. At least 200 cells were counted for each cell suspension.

Flow cytometric immunolabeling assays

After treatments, macrophages were collected and centrifuged. Then, phenotypic analysis of cells was performed using flow cytometric direct immunofluorescence assays (20); cells were first incubated for 30 min in PBS with 5% human AB serum at 4°C to avoid nonspecific mAb binding, and then incubated with appropriate FITC- or PE-conjugated mAbs for 20 min at 4°C. Fluorescence related to immunolabeling was measured using a FACSCalibur flow cytometer (BD Biosciences).

Endocytosis assay

Cells were incubated with 1 mg/ml FITC-dextran for 30 min at 37°C. Cellular uptake of FITC-dextran was then monitored by flow cytometry at 525 nm. A negative control was performed in parallel by incubating cells with FITC-dextran at 4°C instead of 37°C. Uptake of FITC-dextran was expressed as Δ mean fluorescence intensity (MFI), i.e., MFI (uptake at 37°C) – MFI (uptake at 4°C).

Phagocytosis of latex microspheres and FITC-labeled Escherichia coli

Cells were incubated with 15 μl of fluorescent latex microspheres for 30 min at 37°C. Cellular phagocytosis of latex beads was then monitored by flow cytometry at 525 nm. A negative control was performed in parallel by incubating cells with latex beads at 4°C instead of 37°C.

For studying uptake of E. coli bacteria by macrophages, a Phagotest (BD Biosciences) was used according to the instructions of manufacturer. Briefly, 20 μl of precooled FITC-labeled E. coli bacteria were added to a 100-μl suspension of macrophages. The resulting cell suspension was next incubated for 3 h at 37°C or at 4°C (controls). Cells were then washed, and 100 μl of quenching solution was added. After washings, cells were analyzed on FACSCalibur. Phagocytosis of latex microspheres or FITC-labeled E. coli was expressed as Δ MFI, i.e., MFI (uptake at 37°C) – MFI (uptake at 4°C).

Total RNA isolation and reverse transcription-real-time quantitative PCR (RT-qPCR) assay

Total RNAs were extracted from macrophages using the TRIzol method (Invitrogen Life Technologies) and then subjected to RT-qPCR analyses as described previously (22). RT-qPCR assays were performed using the fluorescent dye SYBR Green methodology and an ABI Prism 7000 detector (Applied Biosystem). Specific primers were as follows: TNFα, forward, AACCTCCTCTCTGCCATCAA; TNFα, reverse, ATGTTCGTCCTCCTCACAGG; IL-8, forward, AAGAAACCACCGGAAGGAAC; IL-8, reverse, AAATTTGGGGTGGAAAGGTT. The specificity of each gene amplification was checked up at the end of qPCR through analysis of dissociation curves of the PCR products. Amplification curves were read with ABI Prism 7000 SDS software using the comparative cycle threshold method. Relative quantification of the steady-state target mRNA levels was calculated after normalization of the total amount of cDNA to an 18S RNA endogenous reference.

Determination of cytokine levels

Levels of TNFα and IL-8 in the supernatants of macrophage cultures were quantified using Duoset ELISA development system kits obtained from R&D Systems. Briefly, 96-well plates, initially coated with 1 μg/ml mouse anti-human TNFα or IL-8 Ab overnight at room temperature, were incubated for 2 h with macrophage culture supernatants or recombinant human TNFα or IL-8 standard. After washing, plates were processed according to the manufacturer’s instructions.

Analysis of F-actin expression

Monocytes were first differentiated with GM-CSF into macrophages on glass coverslips for 6 days, and then treated with As2O3. After washing, cells were fixed on coverslips with 3% paraformaldehyde in PBS for 30 min at 4°C and washed three times with PBS. Fixed cells were subsequently incubated with a blocking and permeabilizing solution (PBS, 2 mg/ml BSA, 0.2 mg/ml saponin) for 1 h at room temperature. Cells were then incubated with FITC-phalloidin (1.5 μM), to detect F-actin filaments, in blocking solution for 2 h at room temperature and washed in blocking solution. Thereafter, cells were costained with blocking solution containing 1 μg/ml 4′,6′-diamidino-2-phenylindole, a fluorescent dye specific for DNA, during 15 min. After washings, coverslips were mounted with PBS-glycerol-Dabco. Fluorescent-labeled cells were captured with a DMRXA2 Leica microscope and a COOLSNAP HQ CCD camera, using Metavue software (Molecular Devices).

RhoA-GTP pull-down assay

RhoA-GTP levels were measured using the RhoA activation assay kit from Cytoskeleton (Tebu-bio). Briefly, cells were rapidly lysed at 4°C and incubated with Rhotekin-RBD affinity beads to specifically pull-down RhoA-GTP. After washing, RhoA levels were quantified by running bead/protein complexes in Laemmli buffer containing 0.1 M DTT and probing with the mouse monoclonal anti-RhoA Ab as recommended by manufacturers.

Western blot immunoassays

After treatment, macrophages were harvested and lysed on ice with lysis buffer (60 mM β-glycerophosphate, 15 mM paranitrophenylphosphate, 25 mM MOPS (pH 7.2), 15 mM EGTA, MgCl2, 1 mM NaF, 1 mM phenylphosphate, 2 mM DTT, 1 mM sodium orthovanadate, 1% protease inhibitor mixture (Roche)). Then, lysates were sonicated on ice. Protein concentration was quantified using the Bradford’s method (23). A total of 50 μg of each sample was heated for 5 min at 100°C, and then analyzed by 12% SDS-PAGE, and electroblotted overnight onto nitrocellulose membranes (Bio-Rad). After blocking, membranes were hybridized with primary Abs overnight at 4°C and incubated with appropriate HRP-conjugated secondary Ab (DakoCytomation). Immunolabeled proteins were visualized by chemiluminescence.

Statistical analysis

The results are presented as means ± SEM. Significant differences were evaluated with the Student’s t test. Criterion of significance of the difference between means was p < 0.05.

Results

As2O3 alters morphology, adhesion, and actin organization of human macrophages

Blood monocyte-derived macrophages are tightly adherent cells displaying a “fried-egg”-like morphology, i.e., large spread cells with distinct nuclei (17) (Fig. 1⇓A); these macrophages survived in GM-CSF-free culture medium for several days without alteration of their morphology. In contrast, a 6-day treatment of cultures with 1 μM As2O3 induced a time-dependent rounding of the macrophage (Fig. 1⇓A); rounded and contracted morphology was first observed in some macrophages after 8 h; the amount of rounded macrophages was nearly maximal and stable after 72 h. Morphological effects of arsenic were dose-dependent and detectable from 0.25 μM (Fig. 1⇓B). After a 6-day treatment with As2O3, macrophage rounding was associated with a loss of adhesion as assessed by the WST-1 assay; indeed, only 63.2 ± 6.2 and 31.3 ± 4.5% of macrophages, comparatively to untreated cells, were still adherent after treatment with 0.5 and 1 μM As2O3, respectively. Adhesion and spreading of macrophages are mainly related to reorganization of the actin cytoskeleton. In our cellular model, untreated macrophages displayed extended F-actins resembling filopodia, in their periphery, as previously reported (24). Interestingly, As2O3 induced a time-dependent reorganization of F-actin cytoskeleton in macrophages treated for 72 h (Fig. 1⇓C). Peripheral extensions of F-actin began to retract after 8 h and formed a cortical actin ring after 24 h. As2O3-induced reorganization of F-actin was then stable for at least 72 h.

FIGURE 1.
FIGURE 1.

As2O3 alters morphology and F-actin organization of human primary macrophages. Macrophages were cultured at day 0 (d0) in the absence or presence of 1 μM As2O3 for the indicated time intervals (A and C) or for 6 days (d6) at the indicated concentrations (B). A and B, Phase-contrast microscopy of macrophages from d0 to d6 (magnification, ×40). C, After fixation and permeabilization, immunostaining of cells was performed with FITC-phalloidin to specifically detect F-actin. Cells were costained with 4′,6′-diamidino-2-phenylindole to detect nuclei and subsequently viewed by fluorescence microscopy (magnification, ×400). Scale bar, 10 μm. Data are representative of four independent experiments.

As2O3-induced morphological alterations do not result from cell death

Contribution of direct cytotoxicity to morphological effects of As2O3 was then evaluated using the fluorescent A5 and SG probes, which detected apoptosis and necrosis, respectively. As shown in Fig. 2⇓A, >85% of macrophages were viable after 6, 12, and 18 days of cultures in the absence or presence of 1 μM As2O3. In contrast, higher concentrations of metalloid (2–4 μM) decreased cell viability and specifically increased the percentage of apoptotic macrophages (A5+SG−) after a 6-day treatment (Fig. 2⇓B).

FIGURE 2.
FIGURE 2.

As2O3-induced morphological alterations are not associated with cell death. Human macrophages were cultured in the absence or presence of As2O3 for the indicated time intervals (A) and concentrations (B). Then, cells were costained with annexin V-Alexa 568 (A5) and SG to detect apoptotic (A5+SG−) and necrotic cells (A5−SG+ and A5+SG+), respectively, and viewed by fluorescence microscopy. In A, only viable cells (A5−SG−) are represented in the graph. Values are means ± SEM of three (A) and eight (B) independent experiments.

As2O3 alters phenotype of human macrophages

Besides morphology and adhesion, we next determined whether As2O3 could affect other specific features of differentiated macrophages. We first analyzed expression of some typical surface markers. Table I⇓ shows that treatment of macrophages for 6 days with 1 μM As2O3, markedly reduced expression of CD11b and CD29, two integrins involved in macrophage adhesion and spreading (25). As2O3 also decreased expression of CD71, a well-known macrophagic differentiation marker (18). In contrast, it markedly increased that of CD14, a typical marker of human blood monocytes, down-regulated by GM-CSF during macrophagic differentiation (26). However, a concentration of 1 μM As2O3 failed to modify CD11c levels, which indicates that metalloid effects were not due to a general alteration of surface markers. We then evaluated effects of As2O3 on endocytosis and phagocytosis, two major functions of macrophages. Exposure of macrophages to 1 μM As2O3 for 6 days markedly reduced endocytosis of FITC-dextran (Fig. 3⇓A) and phagocytosis of fluorescent microspheres (Fig. 3⇓B), when compared with untreated cells. Effect of metalloid on macrophagic functions was also evaluated using Phagotest, a true functional assay investigating phagocytosis of bacteria. As expected, uptakes of FITC-labeled E. coli by macrophages incubated at 4°C were negligible (MFI = 2). Fig. 3⇓C demonstrates that 1 μM As2O3 significantly decreased the uptake of FITC-labeled E. coli after a 4- or 6-day treatment. We next determined whether metalloid could alter the ability of macrophages to secrete inflammatory cytokines in response to LPS, which constitutes another major function of APCs. LPS markedly enhanced secretion of both TNFα and IL-8 after 8 and 24 h (Fig. 4⇓); concomitantly, treatment with LPS for 8 h strongly induced TNFα and IL-8 mRNA levels, as assessed using RT-qPCR assays (Table II⇓). A treatment of 6 days with 1 μM As2O3 had no obvious effect on cytokine secretion, whereas it marginally increased TNFα mRNA levels of untreated macrophages. By contrast, it strongly potentiated secretion and mRNA level of TNFα and IL-8 induced by LPS (Fig. 4⇓, Table II⇓).

FIGURE 3.
FIGURE 3.

As2O3 decreases endocytosis and phagocytosis by macrophages. Differentiated macrophages were cultured in the absence or presence of 1 μM As2O3 for 4 (d4) (C) or 6 days (d6) (A–C). Macrophages were incubated with FITC-dextran (A), fluorescent latex microbeads (B), or FITC-labeled E. coli (C) at 4°C (negative control) or 37°C to measure endocytosis (A) and phagocytosis (B and C). Cellular uptakes of FITC-dextran, microbeads, or fluorescent E. coli were then determined by flow cytometry. Uptakes, expressed as ΔMFI (ΔMFI = MFI37°C – MFI4°C), are means ± SEM of, at least, three independent experiments. ∗, p < 0.05 compared with untreated macrophages.

FIGURE 4.
FIGURE 4.

As2O3 enhances TNFα and IL-8 secretion induced by LPS. Human macrophages were cultured in the absence (control) or presence of 1 μM As2O3 for 6 days. Cells were then untreated or exposed to 200 ng/ml LPS for 8 or 24 h in the absence or presence of As2O3. Secretion of cytokine in supernatants were then determined by ELISA. ∗, p < 0.05.

View this table:
Table I.

Phenotypic analysis of macrophages exposed to 1μM As2O3a

View this table:
Table II.

As2O3 potentiates TNFα and IL-8 mRNA levels induced by LPSa

As2O3 effects toward human macrophages are reversible

To determine whether effects of metalloid were reversible, we first treated macrophages with 1 μM As2O3 for 6 days; cells were then washed and cultured with GM-CSF in arsenic-free medium for another 6 days. Fig. 5⇓A shows that withdrawal of iAs, associated with addition of GM-CSF, induced readhesion and spreading of As2O3-treated macrophages. Moreover, reversion of cell rounding was also associated with changes in surface marker expression (Fig. 5⇓, B–E). Indeed, after withdrawal of iAs and addition of GM-CSF, expressions of CD11b, CD29, and CD71, decreased in As2O3-treated macrophages, returned to levels observed in untreated macrophages; in addition, CD14 levels, up-regulated in As2O3-treated cells, were concomitantly reduced.

FIGURE 5.
FIGURE 5.

As2O3-induced alterations of macrophages are reversible. Human macrophages were cultured in the absence (control) or presence of 1 μM As2O3 for 6 days (d0 to d6). Then, macrophages were washed, centrifuged, and reincubated in fresh medium for another 6 days (d6 to d12) in the absence or presence of GM-CSF. In A, phase-contrast microscopy was performed at d12. Experiments were repeated five times, with similar results. B–E, Cells were stained with FITC- or PE-conjugated mAbs directed against the surface markers CD14 (B), CD71 (C), CD11b (D), and CD29 (E), and analyzed by flow cytometry. Results are expressed as means ± SEM of, at least, three independent experiments. ∗, p < 0.05.

As2O3-treated macrophages could be differentiated into dendritic-like cells

Monocytes are not only able to differentiate into macrophages but also into immature dendritic cells, in response to appropriate stimuli such as treatment with GM-CSF and IL-4 (20). Because the above results suggest that As2O3 “de-differentiated” macrophages into monocytic cells, we next analyzed whether As2O3-treated macrophages could be differentiated into dendritic cells as monocytic cells. Fig. 6⇓A shows that macrophages previously exposed to As2O3 for 6 days and then cultured with GM-CSF and IL-4 in iAs-free medium for an additional 6 days, expressed high levels of the typical dendritic marker CD1a. This CD1a expression was similar to that found in blood monocyte-derived dendritic cells (Fig. 6⇓A); in contrast, treatment of control macrophages with GM-CSF and IL-4 did not significantly increased CD1a expression, indicating that fully differentiated macrophages failed to yield dendritic-like cells. In addition, like blood monocyte-derived dendritic cells, and in contrast to untreated macrophages, macrophages first exposed to As2O3 and next cultured with GM-CSF and IL-4 in the absence of iAs were nonadherent (Fig. 6⇓B).

FIGURE 6.
FIGURE 6.

As2O3-treated cells can be differentiated into dendritic-like cells. Human blood monocytes and macrophages previously untreated (control d6) or exposed to 1 μM As2O3 for 6 days (d6) were cultured in the absence or presence of GM-CSF plus IL-4, in As2O3-free RPMI 1640 medium for 6 days (d6 to d12). A, Cells were then stained with FITC-conjugated mAbs directed against the dendritic cell marker CD1a and analyzed by flow cytometry. Results are expressed as means ± SEM of three independent experiments. B, Phase-contrast microscopy of cell cultures (magnification, ×40). Experiments were repeated three times, with similar results.

Inhibition of ROCK prevents As2O3-induced morphological alterations

The Rho-associated kinase ROCK is a major target of the small GTP-binding protein RhoA and a key signaling molecule involved in cytoskeleton regulation (27). Pretreatment of cultures with Y-27632, a specific inhibitor of ROCK (28), prevented both F-actin reorganization and cellular rounding in macrophages treated with As2O3 for 24 h (data not shown) and 72 h (Fig. 7⇓A). In addition, Fig. 7⇓B indicates that the ROCK inhibitor significantly prevented, although partially, alteration of phagocytosis due to As2O3 treatment. We next evaluated the effect of As2O3 on levels of the active GTP-bound form of RhoA and found that 1 μM As2O3 strongly increased its level after a treatment of 8 h (Fig. 8⇓A). RhoA-GTP levels remained increased for at least 72 h. We finally analyzed expression of RhoGDI and ERM proteins, which are involved in the regulation of RhoA activity (29, 30). Fig. 8⇓B shows that As2O3 did not alter protein levels of RhoGDI; in contrast, it increased levels of phospho-Moesin without affecting those of total ERM proteins (Fig. 8⇓C). In fact, ezrin and radixin are only slightly expressed in macrophages and their phosphorylated forms were barely detected, if at all, by Western blotting. Effects of metalloid on phospho-Moesin levels were rapid and detectable as soon as 4 h.

FIGURE 7.
FIGURE 7.

The ROCK inhibitor Y-27632 prevents As2O3 effects on macrophage phenotype. A, Human macrophages were pretreated or not for 2 h with a selective inhibitor of ROCK, Y-27632 (20 μM), and then cultured in the absence or presence of 1 μM As2O3 for 72 h. F-actin immunolocalization by fluorescence microscopy (magnification, ×400; scale bar, 10 μm) and phase-contrast microscopy were then achieved as previously described. B, Macrophages were incubated with FITC-labeled E. coli at 4°C (negative control) or 37°C to measure phagocytosis. Cellular uptakes of fluorescent E. coli were then determined by flow cytometry. Uptakes, expressed as ΔMFI (ΔMFI = MFI37°C – MFI4°C), are means ± SEM of four independent experiments. ∗, p < 0.05 compared with As2O3-treated macrophages.

FIGURE 8.
FIGURE 8.

As2O3 increases RhoA-GTP and phospho-Moesin levels in macrophages. Macrophages were cultured in the absence or presence of 1 μM As2O3 for the indicated time intervals. A, The GTP-binding fraction of RhoA was pulled down as described in Materials and Methods. The bead/protein complexes and whole-cell lysates were then immunoblotted to detect RhoA levels. Western blot analysis of Rho-GDI (B) and phospho-ERM protein (C) expressions were performed with whole-cell lysates. Equal gel loading and transfer efficiency were checked by protein hybridization with anti-RhoA and anti-ERM Ab, respectively. Experiments in A–C were repeated, at least three times, with similar results.

Discussion

The present results demonstrated that As2O3 markedly alters differentiation features of human macrophages, likely through a mechanism involving a RhoA-ROCK pathway. Such effects result in the acquisition by As2O3-treated macrophages of phenotypic properties usually exhibited by monocytic cells.

First, our results show that treatment of macrophages with 0.5–1 μM As2O3, two noncytotoxic concentrations, induced rapid cellular rounding and thereafter loss of adhesion. Cell rounding was tightly associated with a time-dependent reorganization of F-actin in macrophages. In fact, As2O3 induced retraction of peripheral F-actin extensions and stable formation of a cortical actin ring; interestingly, organization of F-actin cytoskeleton in arsenic-treated macrophages was very similar to that observed in freshly adherent monocytes (31). In addition, As2O3 decreased expression of the CD11b and CD29 integrins, which play a major role in adhesion, spreading, and differentiation of monocytes (32). As2O3 also decreased expression of the CD71 macrophagic marker, increased that of the CD14 monocyte marker, and markedly reduced both endocytosis and phagocytosis. Finally, like blood monocytes, As2O3-treated macrophages could be differentiated into dendritic-like cells by GM-CSF and IL-4. Our study therefore clearly demonstrated that metalloid effects on morphology and surface marker expression were reversible, thus confirming that they did not reflect a general unspecific toxicity toward macrophages.

Macrophages are known to possess functional plasticity, which allows them to adapt to changing microenvironment, notably after cytokine or LPS treatment (33). The present results suggest that human macrophages can also “de-differentiate” into monocyte-like cells when exposed to an appropriate chemical compound. This study and other recent reports highlight a marked influence of iAs on cell differentiation. Besides macrophages, iAs partially de-differentiates murine adipocytes (34); arsenite inhibits differentiation of preadipocytes and human epidermal cells likely by maintaining proliferative cell capacity during differentiation program (35, 36). We have also recently demonstrated that As2O3 totally blocks differentiation of human blood monocytes into macrophages through down-regulation of survival pathways (11). On the other hand, As2O3 was shown to promote differentiation of breast cancer cells (37) and that of acute promyelocytic leukemia cells (38). Together, these results demonstrate that iAs exerts complex effects on differentiation according to the cell type.

Different arguments support the idea that As2O3 impaired human macrophage function, at least in part, through activation of a RhoA/ROCK signaling pathway. First, our results demonstrate that the ROCK inhibitor Y-27632 prevented F-actin reorganization, cell rounding and phagocytosis inhibition in As2O3-exposed macrophages. In addition, we report that As2O3 activates the small G protein RhoA, a potent ROCK inducer. Indeed, As2O3 induces a rapid and sustained increase of RhoA-GTP levels which preceded its effects on morphology and cytoskeleton. Involvement of RhoA/ROCK in As2O3 toxicity is further strengthened by the fact that this pathway is known 1) to induce cell contraction through reorganization of actin cytoskeleton, notably in macrophages (24) and 2) to constitute a negative regulator of monocyte spreading (39). In addition, the recent observation that RhoA-activated ROCK negatively regulated phagocytosis of apoptotic cells (40) argues for a role of ROCK in As2O3-induced down-regulation of phagocytic activity. RhoA is regulated by several proteins and particularly by RhoGDI proteins, which maintain RhoA in its inactive GDP-bound form (30). DNA microarrays have revealed that arsenite can down-regulate mRNA levels of RhoGDI proteins (41, 42). However, our results show that As2O3 had no effect on RhoGDI protein levels in macrophages; in contrast, it increased those of phospho-Moesin, a member of the ERM protein family. Phosphorylation of ERM proteins favors their activation by stabilizing the active open conformation (29). Once phosphorylated, these proteins can promote dissociation of RhoGDI from RhoA and subsequent activation of RhoA (29, 30). The fact that phosphorylation of Moesin seems to precede increase of RhoA-GTP levels suggests that this ERM protein was involved in RhoA activation in As2O3-treated macrophages. Besides its effects on RhoA, iAs was also recently demonstrated to regulate actin organization through activation of the small G protein Cdc42 in murine vascular cells (43). These results thus indicate that such small G proteins constitute new molecular targets of iAs which merit further exploration.

Together with our previous study (11), the present results suggest that low micromolar concentrations (0.25–1 μM) of As2O3, which are relevant of arsenic blood levels measured in chronically exposed humans (15), can target the monocyte/macrophage system in humans. Indeed, our data likely indicate that iAs can reduce the pool of human functional macrophages by 1) inhibiting macrophagic differentiation of blood monocytes (11) and 2) impairing features of differentiated monocyte-derived macrophages, notably endocytosis and phagocytosis; at higher concentrations (4 μM), As2O3 also induces apoptosis of differentiated macrophages. Relevance of these mechanisms is strengthened by in vivo studies reporting a decrease of phagocytic activity and antibacterial response in mice treated with arsenite (13). However, toxicity of iAs toward macrophages appears to be complex, because As2O3 strongly increases the macrophage response to LPS. In fact, although it had no significant effect on untreated macrophages, As2O3 appears to “prime” macrophages for LPS-induced secretion of TNFα and IL-8, two major proinflammatory cytokines. The superinduction of TNFα and IL-8 mRNA levels in macrophages coexposed to LPS and iAs suggests that iAs can modulate LPS intracellular signals. Such effects may contribute to the development of iAs-related inflammation observed in chronically exposed humans (15).

Interestingly, iAs, used under its trioxide form, is an effective antileukemic drug and its concentrations in blood patients are close to those acting in vitro on macrophages (44). This indicates that macrophages, in addition to malignant cells, may also constitute targets in As2O3-treated patients, and therefore supports the idea that a careful analysis of the monocyte/macrophage compartment may be of interest in such subjects.

In conclusion, our results demonstrate that noncytotoxic concentrations of As2O3 down-regulate differentiation features of human primary macrophages likely through activation of a RhoA/ROCK pathway. Moreover, they show that human macrophages constitute by themselves sensitive targets of iAs, which may contribute to immunotoxicity of this major environmental contaminant.

Acknowledgments

We are grateful to Dr. C. Leberre (Etablissement Français du Sang, Rennes, France) for providing us with blood buffy coats. Thanks to the Institut Fédératif de Recherche 140 microscopy platform in Rennes.

Disclosures

The authors have no financial conflict of interest.

Footnotes

  • The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked advertisement in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

  • ↵1 This work was supported by grants from Association pour la Recherche sur le Cancer and Ligue Contre le Cancer (Comité d’Ille-et-Vilaine). A.L. and E.B. are recipients of a fellowship from Ligue Nationale Contre le Cancer and Ligue Contre le Cancer (Comité d’Ille-et-Vilaine), respectively.

  • ↵2 Address correspondence and reprint requests to Dr. Laurent Vernhet, Institut National de la Santé et de la Recherche Médicale, Unité 620, Faculté des Sciences Pharmaceutiques et Biologiques, Institut Fédératif de Recherche 140, Université de Rennes-1, 2 avenue du Professeur Léon Bernard, 35043 Rennes, France. E-mail address: Laurent.Vernhet{at}rennes.inserm.fr

  • ↵3 Abbreviations used in this paper: iAs, inorganic arsenic; ERM, Ezrin/Radixin/Moesin; RhoGDI, Rho guanine dissociation inhibitor; A5, annexin V; SG, sytox green; MFI, mean fluorescence intensity; RT-qPCR, reverse transcription-real-time quantitative PCR; ROCK, Rho-associated kinase.

  • Received November 29, 2005.
  • Accepted June 8, 2006.

References

  1. ↵
    Rossman, T. G.. 2003. Mechanism of arsenic carcinogenesis: an integrated approach. Mutat. Res. 533: 37-65.
    OpenUrlCrossRefPubMed
  2. ↵
    Navas-Acien, A., A. R. Sharrett, E. K. Silbergeld, B. S. Schwartz, K. E. Nachman, T. A. Burke, E. Guallar. 2005. Arsenic exposure and cardiovascular disease: a systematic review of the epidemiologic evidence. Am. J. Epidemiol. 162: 1037-1049.
    OpenUrlAbstract/FREE Full Text
  3. ↵
    Brown, K. G., G. L. Ross. 2002. Arsenic, drinking water, and health: a position paper of the American Council on Science and Health. Regul. Toxicol. Pharmacol. 36: 162-174.
    OpenUrlCrossRefPubMed
  4. ↵
    Abernathy, C. O., D. J. Thomas, R. L. Calderon. 2003. Health effects and risk assessment of arsenic. J. Nutr. 133: 1536S-1538S.
    OpenUrlAbstract/FREE Full Text
  5. ↵
    Abel, E. A.. 1989. Cutaneous manifestations of immunosuppression in organ transplant recipients. J. Am. Acad. Dermatol. 21: 167-179.
    OpenUrlCrossRefPubMed
  6. ↵
    Boukamp, P.. 2005. Non-melanoma skin cancer: what drives tumor development and progression?. Carcinogenesis 26: 1657-1667.
    OpenUrlAbstract/FREE Full Text
  7. ↵
    Soto-Pena, G. A., A. L. Luna, L. Acosta-Saavedra, P. Conde, L. Lopez-Carrillo, M. E. Cebrian, M. Bastida, E. S. Calderon-Aranda, L. Vega. 2006. Assessment of lymphocyte subpopulations and cytokine secretion in children exposed to arsenic. FASEB J. 20: 779-781.
    OpenUrlAbstract/FREE Full Text
  8. ↵
    Galicia, G., R. Leyva, E. P. Tenorio, P. Ostrosky-Wegman, R. Saavedra. 2003. Sodium arsenite retards proliferation of PHA-activated T cells by delaying the production and secretion of IL-2. Int. Immunopharmacol. 3: 671-682.
    OpenUrlCrossRefPubMed
  9. ↵
    Patterson, R., L. Vega, K. Trouba, C. Bortner, D. Germolec. 2004. Arsenic-induced alterations in the contact hypersensitivity response in BALB/c mice. Toxicol. Appl. Pharmacol. 198: 434-443.
    OpenUrlCrossRefPubMed
  10. ↵
    Sikorski, E. E., L. A. Burns, M. L. Stern, M. I. Luster, A. E. Munson. 1991. Splenic cell targets in gallium arsenide-induced suppression of the primary antibody response. Toxicol. Appl. Pharmacol. 110: 129-142.
    OpenUrlCrossRefPubMed
  11. ↵
    Lemarie, A., C. Morzadec, D. Merino, O. Micheau, O. Fardel, L. Vernhet. 2006. Arsenic trioxide induces apoptosis of human monocytes during macrophagic differentiation through nuclear factor-κB-related survival pathway down-regulation. J. Pharmacol. Exp. Ther. 316: 304-314.
    OpenUrlAbstract/FREE Full Text
  12. ↵
    Sakurai, T., T. Ohta, K. Fujiwara. 2005. Inorganic arsenite alters macrophage generation from human peripheral blood monocytes. Toxicol. Appl. Pharmacol. 203: 145-153.
    OpenUrlCrossRefPubMed
  13. ↵
    Bishayi, B., M. Sengupta. 2003. Intracellular survival of Staphylococcus aureus due to alteration of cellular activity in arsenic and lead intoxicated mature Swiss albino mice. Toxicology 184: 31-39.
    OpenUrlCrossRefPubMed
  14. ↵
    Lewis, T. A., C. B. Hartmann, K. L. McCoy. 1998. Gallium arsenide modulates proteolytic cathepsin activities and antigen processing by macrophages. J. Immunol. 161: 2151-2157.
    OpenUrlAbstract/FREE Full Text
  15. ↵
    Wu, M. M., H. Y. Chiou, I. C. Ho, C. J. Chen, T. C. Lee. 2003. Gene expression of inflammatory molecules in circulating lymphocytes from arsenic-exposed human subjects. Environ. Health Perspect. 111: 1429-1438.
    OpenUrlCrossRefPubMed
  16. ↵
    Pi, J., Y. Kumagai, G. Sun, H. Yamauchi, T. Yoshida, H. Iso, A. Endo, L. Yu, K. Yuki, T. Miyauchi, N. Shimojo. 2000. Decreased serum concentrations of nitric oxide metabolites among Chinese in an endemic area of chronic arsenic poisoning in inner Mongolia. Free Radic. Biol. Med. 28: 1137-1142.
    OpenUrlCrossRefPubMed
  17. ↵
    Young, D. A., L. D. Lowe, S. C. Clark. 1990. Comparison of the effects of IL-3, granulocyte-macrophage colony-stimulating factor, and macrophage colony-stimulating factor in supporting monocyte differentiation in culture: analysis of macrophage antibody-dependent cellular cytotoxicity. J. Immunol. 145: 607-615.
    OpenUrlAbstract
  18. ↵
    van Grevenynghe, J., S. Rion, E. Le Ferrec, M. Le Vee, L. Amiot, R. Fauchet, O. Fardel. 2003. Polycyclic aromatic hydrocarbons inhibit differentiation of human monocytes into macrophages. J. Immunol. 170: 2374-2381.
    OpenUrlAbstract/FREE Full Text
  19. ↵
    Lecureur, V., E. L. Ferrec, M. N’Diaye, M. L. Vee, C. Gardyn, D. Gilot, O. Fardel. 2005. ERK-dependent induction of TNFα expression by the environmental contaminant benzo(a)pyrene in primary human macrophages. FEBS Lett. 579: 1904-1910.
    OpenUrlCrossRefPubMed
  20. ↵
    Laupeze, B., L. Amiot, L. Sparfel, E. Le Ferrec, R. Fauchet, O. Fardel. 2002. Polycyclic aromatic hydrocarbons affect functional differentiation and maturation of human monocyte-derived dendritic cells. J. Immunol. 168: 2652-2658.
    OpenUrlAbstract/FREE Full Text
  21. ↵
    Lemarie, A., D. Lagadic-Gossmann, C. Morzadec, N. Allain, O. Fardel, L. Vernhet. 2004. Cadmium induces caspase-independent apoptosis in liver Hep3B cells: role for calcium in signaling oxidative stress-related impairment of mitochondria and relocation of endonuclease G and apoptosis-inducing factor. Free Radic. Biol. Med. 36: 1517-1531.
    OpenUrlCrossRefPubMed
  22. ↵
    Le Vee, M., E. Jigorel, D. Glaise, P. Gripon, C. Guguen-Guillouzo, O. Fardel. 2006. Functional expression of sinusoidal and canalicular hepatic drug transporters in the differentiated human hepatoma HepaRG cell line. Eur. J. Pharm. Sci. 28: 109-117.
    OpenUrlCrossRefPubMed
  23. ↵
    Bradford, M. M.. 1976. A rapid and sensitive method for the quantitation of microgram quantities of protein utilizing the principle of protein-dye binding. Anal. Biochem. 72: 248-254.
    OpenUrlCrossRefPubMed
  24. ↵
    Allen, W. E., G. E. Jones, J. W. Pollard, A. J. Ridley. 1997. Rho, Rac and Cdc42 regulate actin organization and cell adhesion in macrophages. J. Cell Sci. 110: 707-720.
    OpenUrlAbstract/FREE Full Text
  25. ↵
    Berton, G., C. A. Lowell. 1999. Integrin signalling in neutrophils and macrophages. Cell. Signal. 11: 621-635.
    OpenUrlCrossRefPubMed
  26. ↵
    Kruger, M., J. G. Van de Winkel, T. P. De Wit, L. Coorevits, J. L. Ceuppens. 1996. Granulocyte-macrophage colony-stimulating factor down-regulates CD14 expression on monocytes. Immunology 89: 89-95.
    OpenUrlCrossRefPubMed
  27. ↵
    Riento, K., A. J. Ridley. 2003. Rocks: multifunctional kinases in cell behaviour. Nat. Rev. Mol. Cell Biol. 4: 446-456.
    OpenUrlCrossRefPubMed
  28. ↵
    Ishizaki, T., M. Uehata, I. Tamechika, J. Keel, K. Nonomura, M. Maekawa, S. Narumiya. 2000. Pharmacological properties of Y-27632, a specific inhibitor of Rho-associated kinases. Mol. Pharmacol. 57: 976-983.
    OpenUrlAbstract/FREE Full Text
  29. ↵
    Ivetic, A., A. J. Ridley. 2004. Ezrin/radixin/moesin proteins and Rho GTPase signalling in leucocytes. Immunology 112: 165-176.
    OpenUrlCrossRefPubMed
  30. ↵
    Olofsson, B.. 1999. Rho guanine dissociation inhibitors: pivotal molecules in cellular signalling. Cell. Signal. 11: 545-554.
    OpenUrlCrossRefPubMed
  31. ↵
    Williams, L. M., A. J. Ridley. 2000. Lipopolysaccharide induces actin reorganization and tyrosine phosphorylation of Pyk2 and paxillin in monocytes and macrophages. J. Immunol. 164: 2028-2036.
    OpenUrlAbstract/FREE Full Text
  32. ↵
    Shi, C., X. Zhang, Z. Chen, K. Sulaiman, M. W. Feinberg, C. M. Ballantyne, M. K. Jain, D. E. Simon. 2004. Integrin engagment regulates monocyte differentiation through the forkhead transcription factor Foxp1. J. Clin. Invest. 114: 408-418.
    OpenUrlCrossRefPubMed
  33. ↵
    Stout, R. D., J. Suttles. 2004. Functional plasticity of macrophages: reversible adaptation to changing microenvironments. J. Leukocyte Biol. 76: 509-513.
    OpenUrlAbstract/FREE Full Text
  34. ↵
    Wauson, E. M., A. S. Langan, R. L. Vorce. 2002. Sodium arsenite inhibits and reverses expression of adipogenic and fat cell-specific genes during in vitro adipogenesis. Toxicol. Sci. 65: 211-219.
    OpenUrlAbstract/FREE Full Text
  35. ↵
    Trouba, K. J., E. M. Wauson, R. L. Vorce. 2000. Sodium arsenite inhibits terminal differentiation of murine C3H 10T1/2 preadipocytes. Toxicol. Appl. Pharmacol. 168: 25-35.
    OpenUrlCrossRefPubMed
  36. ↵
    Patterson, T. J., T. V. Reznikova, M. A. Phillips, R. H. Rice. 2005. Arsenite maintains germinative state in cultured human epidermal cells. Toxicol. Appl. Pharmacol. 207: 69-77.
    OpenUrlCrossRefPubMed
  37. ↵
    Baj, G., A. Arnulfo, S. Deaglio, R. Mallone, A. Vigone, M. G. De Cesaris, N. Surico, F. Malavasi, E. Ferrero. 2002. Arsenic trioxide and breast cancer: analysis of the apoptotic, differentiative and immunomodulatory effects. Breast Cancer Res. Treat. 73: 61-73.
    OpenUrlCrossRefPubMed
  38. ↵
    Miller, W. H., Jr, H. M. Schipper, J. S. Lee, J. Singer, S. Waxman. 2002. Mechanisms of action of arsenic trioxide. Cancer Res. 62: 3893-3903.
    OpenUrlAbstract/FREE Full Text
  39. ↵
    Aepfelbacher, M., M. Essler, E. Huber, A. Czech, P. C. Weber. 1996. Rho is a negative regulator of human monocyte spreading. J. Immunol. 157: 5070-5075.
    OpenUrlAbstract
  40. ↵
    Tosello-Trampont, A. C., K. Nakada-Tsukui, K. S. Ravichandran. 2003. Engulfment of apoptotic cells is negatively regulated by Rho-mediated signaling. J. Biol. Chem. 278: 49911-49919.
    OpenUrlAbstract/FREE Full Text
  41. ↵
    Andrew, A. S., A. J. Warren, A. Barchowsky, K. A. Temple, L. Klei, N. V. Soucy, K. A. O’Hara, J. W. Hamilton. 2003. Genomic and proteomic profiling of responses to toxic metals in human lung cells. Environ. Health Perspect. 111: 825-835.
    OpenUrlPubMed
  42. ↵
    Rea, M. A., J. P. Gregg, Q. Qin, M. A. Phillips, R. H. Rice. 2003. Global alteration of gene expression in human keratinocytes by inorganic arsenic. Carcinogenesis 24: 747-756.
    OpenUrlAbstract/FREE Full Text
  43. ↵
    Qian, Y., K. J. Liu, Y. Chen, D. C. Flynn, V. Castranova, X. Shi. 2005. Cdc42 regulates arsenic-induced NADPH oxidase activation and cell migration through actin filament reorganization. J. Biol. Chem. 280: 3875-3884.
    OpenUrlAbstract/FREE Full Text
  44. ↵
    Shen, Z. X., G. Q. Chen, J. H. Ni, X. S. Li, S. M. Xiong, Q. Y. Qiu, J. Zhu, W. Tang, G. L. Sun, K. Q. Yang, et al 1997. Use of arsenic trioxide (As2O3) in the treatment of acute promyelocytic leukemia (APL). II. Clinical efficacy and pharmacokinetics in relapsed patients. Blood 89: 3354-3360.
    OpenUrlAbstract/FREE Full Text
Back to top

In this issue

Print
Download PDF
Article Alerts
Email Article
Citation Tools

Jump to section