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Introduction

Due to a wide range of industrial applications, cadmium (Cd) is substantially distributed in various environments (1). Absorption of Cd can be achieved through numerous routes, including ingestion, smoking and inhalation of polluted air, and Cd may accumulate in the liver, kidney, lung and heart for 20–30 years (2). Cd exposure is associated with various diseases, such as renal failure, anemia, itai-itai disease and cancer (3–5), and has therefore long been considered a threat to human health (6,7). Cd elicits toxicity through various pathways, including oxidative stress and initiation of apoptosis (8,9). Numerous studies have documented the cytotoxicity of Cd under various settings. However, there has thus far been no research that has evaluated the biological effects of Cd at non-toxic concentrations, such as chronic low-dose environmental exposure, which is likely coupled to cellular dysfunction and even morbidity. As a divalent metal ion, cellular intrusion of Cd has been suspected to result in systemic disorder of iron homeostasis; however, the mechanisms underlying Cd-induced effects on iron metabolism remain unclear.

Iron is a necessary metal for all cells, and a fine-tuned regulatory system has evolved to maintain an elaborate balance for systemic iron homeostasis (10). The hepcidin-ferroportin axis has a central role in regulating iron flow (11). Hepcidin binds and induces degradation of its receptor ferroportin, the iron exporter, which leads to intracellular iron retention (12). Ferroportin is the only known iron exporter in mammals, which is mainly expressed in enterocytes and macrophages, and its concentration in splenic macrophages has been shown to determine the iron levels in serum and other organs (11,12). Ferroportin concentration is largely regulated at the post-transcriptional level, through the iron responsive element and iron regulatory protein (IRE-IRP) regulatory system (13–15); however, it may also be regulated at the transcriptional level (16,17). Ferroportin mutations or dysfunction results in iron metabolism disorders, known as ferroportin diseases, which are often associated with iron overload (18,19). Type B ferroportin disease is characterized by increased concentrations of ferroportin, due to failure to respond to hepcidin, which causes continuous iron export from the cells into the plasma; this is associated with hyperferritinemia and excess iron in hepatocytes (20). The aim of the present study was to elucidate the regulation of ferroportin in macrophages by Cd at non-toxic concentrations.

Materials and methods

Cell culture

The THP-1 human macrophage, J774A.1 mouse macrophage and HEK293T human embryonic kidney cell lines were purchased from the Shanghai Cell Bank of Type Culture Collection (Shanghai, China). The cells were cultured in RPMI-1640 medium (Gibco Life Technologies, Carlsbad, CA, USA) supplemented with 10% fetal bovine serum (Hyclone Laboratories, Inc., Logan, UT, USA) and 100 U/ml penicillin/streptomycin (Hyclone Laboratories, Inc.). Activation of the THP-1 cells was initiated in complete medium with 1 μg/ml PMA (Promega Corporation, Madison, WI, USA) for 18 h (21).

AlamarBlue® assay

Cell viability was measured using the alamarBlue assay, according to the manufacturer’s instructions (Invitrogen Life Technologies, Carlsbad, CA, USA). Briefly, the THP-1 and J774A.1 cells were plated in 96-well plates, at a concentration of 5.0×103 cells/well. The cells were then treated with various concentrations of CdCl2 (0–64 μM; Sigma-Aldrich, St. Louis, MO, USA) for 24 h, followed by reading with a microplate reader (Thermo Electron Corporation, Waltham, MA, USA) at 590 nm with excitation at 530 nm.

Western blot analysis

The THP-1 cells were collected after washing with cold phosphate-buffered saline (PBS; Solarbio Science & Technology Co., Ltd., Beijing, China), and total proteins were extracted using lysis buffer (Solarbio Science & Technology Co., Ltd., Beijing, China) that was pre-mixed with a protease inhibitor cocktail (Roche Diagnostics, Basel, Switzerland). Protein lysates (30–50 μg total proteins) were then separated by 10% SDS-PAGE and further analyzed by western blotting, as previously described (22). Briefly, proteins were transferred onto pure nitrocellulose membranes, followed by primary antibody (in 1% milk) incubation overnight at 4°C. The primary antibodies used in the present study were as follows: Anti-GAPDH (1:1,000; Santa Cruz Biotechnology, Inc., Dallas, TX, USA) and anti-ferroportin (1:500; Sigma-Aldrich). Subsequently, the secondary antibodies (in 1% milk) were applied for detection of the target proteins (for 1 h at 37°C). The secondary antibodies used in the present study were as follows: Goat anti-rabbit-HRP (for GAPDH; 1:10,000; ComWin Biotech Co., Ltd., Beijing, China) and goat anti-rat-HRP (for FPN; 1:5,000; ComWin Biotech Co., Ltd.). Bands were analyzed by Image J software (version 1.48; National Institutes of Health, Bethesda, MD, USA) following coloration. GAPDH was used as an internal control.

For the inhibition of transcription or translation, THP-1 cells were treated with CdCl2 in the presence or absence of 1 μg/ml actinomycin D (a transcriptional inhibitor; ComWin Biotech Co., Ltd.) or cycloheximide (an inhibitor of protein biosynthesis; ComWin Biotech Co., Ltd.) for 12 h. Subsequently, cells were cultured with new medium containing CdCl2 for another 12 h.

RNA extraction and reverse transcription-quantitative polymerase chain reaction (RT-qPCR)

Following treatment with 4 and 8 μM CdCl2 for 24 h (where 100 μM ferric ammonium citrate was used as a positive control), total RNAs were extracted from the THP-1 cells using TRIzol®, according to the manufacturer’s instructions (Invitrogen Life Technologies). For the reverse transcription reaction, 2 μg total RNAs were used to synthesize first strand cDNA, using oligo (dT) primers. qPCR was conducted to determine gene expression levels using SYBR® Green qPCR Master mix (Qiagen, Hilden, Germany) on a Mx3005P qPCR system (Agilent Technologies, Inc., Santa Clara, CA, USA). The primer sequences used in the present study were as follows: Forward: 5′-CGGTGTCTGTGTTTCTGGTAGA-3′ and reverse: 5′-CTGGGCCACTTTAAGTCTAGC-3′ for Ferroportin; and forward: 5′-GAAGGTGAAGGTCGGAGT-3′ and reverse: 5′-GAAGATGGTGATGGGATTTC-3′ for GAPDH. GAPDH was used as a housekeeping gene for normalization. The qRT-PCR reaction was run at 95°C for 5 min (pre-denaturation) followed by 45 cycles at 95°C for 15 sec, at 55°C for 30 sec, and at 72°C for 30 sec. Following the reaction, a melting curve analysis from 60–95°C was applied to all reactions to ensure consistency and specificity of the amplified products. Bands formed during agarose gel electrophoresis were quantified using the Image J software.

Luciferase reporter assay

A DNA fragment of the IRE sequence encoding the 5′-UTR of human ferroportin mRNA was cloned and then subcloned into the 5′-UTR of the luciferase gene, within the pGL3-Promoter luciferase reporter vector (Promega Corporation). The HindIII (Invitrogen Life Technologies) and NCOI (Invitrogen Life Technologies) enzymes were used in the construction of the plasmid. The constructed plasmid was validated by DNA sequencing using RVprimer3 primer (Invitrogen Life Technologies) and sequence alignment was performed with ClustalX software (version 2.1; Conway Institute University College Dublin, Dublin, Ireland). In the transfection experiments, 0.8 μg target plasmid and 80 ng Renilla luciferase plasmid were co-transfected into HEK293T cells using Lipofectamine® 2000 (Invitrogen Life Technologies) in 24-well plates. Following a 24 h incubation, the cells were washed with cold PBS, and then subjected to luciferase activity determination, using a Dual-Luciferase Reporter Assay system (Promega Corporation). Relative firefly luciferase activities were calculated by normalization to those of Renilla luciferase.

Labile iron pool (LIP) measurement

Following treatment of THP-1 and J774A.1 cells with 8 μM CdCl2 for 24 h, intracellular LIP levels were evaluated according to the standard calcein acetoxymethyl ester staining method (Sigma-Aldrich), as described previously (23). Briefly, cells were washed twice with PBS and treated with 0.5 μM calcein (Sigma-Aldrich) for 15 min at 37°C. Subsequently, cells were washed twice with PBS and divided into two parts. An aliquot was treated with 100 μM desferoxamine (Sigma-Aldrich) for 1 h at 37°C and the other was left untreated. The intracellular fluorescence was then measured by FACS analysis with excitation at 488 nm and reading at 525 nm with a flow cytometer (FACS Calibur®; Becton Dickinson, Franklin Lakes, NJ, USA). The LIP levels were determined from deduction of the cellular fluorescence of deferoxamine-treated cells, compared with the fluorescence of untreated cells.

Statistical analysis

Two-tailed Student’s t-test and one-way analysis of variance were used to analyze the experimental data. The SPSS Statistics 17.0 package (SPSS, Inc., Chicago, IL, USA) was utilized to analyze the data. The data are represented as the mean ± standard error. P<0.05 was considered to indicate a statistically significant difference.

Results and Discussion

Ferroportin is the only known iron exporter in mammalian cells, and it is mainly expressed on macrophages and duodenal enterocytes (24–26). Macrophages have a key role in maintaining iron homeostasis, through governing iron egress into plasma (27,28). The present study focused on the biological effects of Cd on the concentration of ferroportin in macrophages. To accurately determine the potential effects of Cd on ferroportin, sublethal concentrations of CdCl2 were identified that did not induce significant toxicity in macrophages, in order to determine the effects of relatively low non-toxic concentrations of Cd. Cell viability was evaluated following treatment with various concentrations of CdCl2 in THP-1 and J774A.1 cells. Cell viability of THP-1 cells was significantly reduced in response to treatment with CdCl2 at concentrations ≥32 μM for 24 h, as determined by an alamarBlue assay (Fig. 1A; P<0.05). The cell viability of J774A.1 cells was significantly decreased in response to treatment with CdCl2 at concentrations ≥20 μM for 24 h (Fig. 1B; P<0.05). Therefore, two non-toxic concentrations, 4 and 8 μM, were selected for use in the following experiments.

Figure 1 Cell viability assay of THP-1 human and J774A.1 murine macrophage cell lines exposed to various concentrations of CdCl2. Cell viability was assessed by the alamarBlue® assay in (A) THP-1 and (B) J774A.1 cells following treatment with CdCl2 (0–64 μM) for 24 h (4–5 wells in a 96-well plate containing each concentration). The data are represented as the mean ± standard error.

The possible influence of CdCl2 treatment was then determined on the expression of ferroportin. Since, to the best of our knowledge, only a human ferroportin antibody has been reported to function effectively, and no efficient mouse ferroportin antibody is commercially available, the protein expression of ferroportin was only examined in the THP-1 cells. Following treatment of the cells with 4 and 8 μM CdCl2 for 24 h, the cells were collected for western blot analysis. Ferroportin protein expression was markedly increased, by >2-fold, in the 4 and 8 μM Cd-treated cells, as compared with the untreated cells (Fig. 2A). To investigate the mechanisms responsible for the Cd-induced increase in ferroportin protein, THP-1 cells were treated with CdCl2 in the presence or absence of 1 μg/ml actinomycin D (a transcriptional inhibitor) and 10 μg/ml cycloheximide (an inhibitor of protein biosynthesis) for 12 h. The protein expression of ferroportin in the cells simultaneously treated with Cd and actinomycin D was reduced by ~40%, as compared with the cells treated with Cd alone; however, the protein expression was still greater (~20%) in the Cd-treated cells, as compared with the untreated cells (Fig. 2B). Furthermore, the protein expression of ferroportin was markedly reduced, by ~90%, in the cells treated with Cd and cycloheximide, as compared with the cells treated with Cd alone (Fig. 2B). These results indicate that the regulation of ferroportin by Cd may occur at the transcriptional and post-translational levels, but appears more likely to occur at the post-translational level. Ferric ammonium citrate (100 μM) was used as a positive control to promote ferroportin concentration (Fig. 2B). In addition, the mRNA expression levels of ferroportin were determined in the cells treated with 8 μM CdCl2 by RT-qPCR. There were no significant differences in the ferroportin mRNA expression levels between the Cd-treated and untreated cells (P>0.05, data not shown). These results suggest that the regulation of ferroportin by Cd primarily occurs at the post-transcriptional level.

Figure 2 Ferroportin protein expression in THP-1 human macrophage cells upon treatment with CdCl2, as determined by western blotting. Cells were treated with (A) 4 or 8 μM CdCl2 for 24 h, (B) with or without 1 μg/ml actinomycin D or 10 μg/ml cycloheximide for 12 h. FAC, ferric ammonium citrate.

In regards to the post-transcriptional regulation of ferroportin, an IRE within its 5′-UTR has been recognized as being under the control of IRPs (15,17). As a divalent metal ion, Cd possesses similar properties to iron (29), and it has long been postulated that Cd may interfere with the transport and metabolism of numerous essential metals, including iron, copper and zinc, presumably through competition between Cd and the other metals (30,31). Therefore, the present study investigated the possible interruption of IRP-IRE actions on the 5′-UTR of ferroportin, by Cd. A construct was generated by inserting a fragment of the IRE DNA sequence ahead of the 5′-UTR of luciferase, within a pGL3-promoter luciferase reporter vector (Fig. 3A). Treatment with CdCl2 significantly enhanced luciferase activity by >27% in the cells transfected with vector+IRE, as compared with the untreated cells (Fig. 3B; P<0.05). Conversely, there were no significant differences in luciferase activity observed in the cells transfected with the construct devoid of IRE, with or without CdCl2 treatment (Fig. 3B). Notably, the luciferase activity of the cells transfected with vector+IRE was increased by 33%, as compared with the cells transfected with vector only (Fig. 3B, P<0.05), thus implying that IRE is required for maintaining ferroportin levels. These results collectively demonstrate that Cd regulates ferroportin protein expression predominantly through the IRE-IRP regulatory system, and Cd presumably replaced iron in driving IRP removal from IRE in the 5′-UTR of ferroportin.

Figure 3 Effects of CdCl2 in driving the iron responsive element-iron regulatory protein (IRE-IRP) regulatory system. (A) Schematic diagram delineating the construction of the IRE-containing vector, known as vector+IRE. The IRE region was inserted into the 5′-untranslated region of the luciferase gene within a pGL3-promoter luciferase reporter vector, using restriction endonuclease, HindIII and NcoI. (B) Analysis of relative luciferase activities. pGL3-based constructs with IRE (vector+IRE) or without IRE (vector) were co-transfected with Renilla luciferase plasmid into HEK293T human embryonic kidney cells, and treated with or without 8 μM CdCl2 for 24 h. Relative luciferase activities were normalized to those of Renilla luciferase (n=4). The data are represented as the mean ± standard error.

LIP is a sensitive marker of iron storage and bioavailability, which dynamically binds to low-affinity ligands depending on different physiological settings (32,33). LIP levels are concertedly regulated and maintained within a strict range that meets cellular demand for iron, but prevents excess iron-triggered damage (23). To assess the effects of Cd-induced increased ferroportin on cellular iron storage and bioavailability, LIP levels were measured in the THP-1 and J774A.1 cells treated with 8 μM CdCl2. Treatment with Cd significantly decreased LIP levels by >3-fold in the THP-1 cells, as compared with the control cells (Fig. 4A; P<0.05). A similar decrease in LIP levels was observed in the J774A.1 cells treated with Cd, as compared with the untreated cells (Fig. 4B, P<0.05). These results suggest that increased LIP levels were correlated with increased ferroportin expression, in response to treatment with Cd.

Figure 4 Relative labile iron pool (LIP) levels in THP-1 human and J774A.1 murine macrophage cells exposed to CdCl2. Calcein acetoxymethyl ester-based intracellular LIP level assay was performed in (A) THP-1 and (B) J774A.1 cells following treatment with 8 μM CdCl2 for 24 h (n=4). Cadmium reduces intracellular iron by enforcing ferroportin translation via iron responsive element. The data are represented as the mean ± standard error.

Numerous previous studies have revealed deleterious actions of Cd in diverse systems and models (2–5); however, relatively few studies have attempted to determine the potential biological effects of exposure to chronic sublethal Cd levels. Increasing evidence has suggested that Cd may affect the homeostasis of essential metals, through competition or other unknown mechanisms (34,35). Cd burden was previously shown to stimulate the expression of divalent metal transporter 1 in enterocytes (36). Furthermore, a previous study demonstrated that Cd could attenuate erythropoietin production in the kidney, leading to an increase of hepcidin, which is often coupled with iron disorders (37). Park and Chung (38) also reported that Cd could disturb iron homeostasis by producing reactive oxygen species (38). However, research aiming to fully elucidate the effects of Cd on iron-associated genes and proteins remains limited, and further studies are required, in order to improve knowledge.

In conclusion, the present study identified a novel regulation of ferroportin by Cd. Ferroportin protein expression was upregulated in response to sublethal treatment of Cd in macrophages, resulting in attenuation of LIP. Notably, the results suggest that Cd may regulate ferroportin translation through the 5′-UTR IRE of ferroportin. These results therefore may have identified the molecular basis by which Cd impairs cellular iron storage and bioavailability, under non-toxic concentrations in macrophages.

Acknowledgements

The present study was supported by a grant from the National ‘973’ program (grant no. 2014CB932000) and the National Natural Science Foundation of China (grant no. 21377159). The authors of the present study would also like to thank the laboratory members for their assistance with the experiments and reagents.

References