Influence of DNA-methylation on zinc homeostasis in myeloid cells: Regulation of zinc transporters and zinc binding proteins

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Influence of DNA-methylation on zinc homeostasis in myeloid cells: Regulation of zinc transporters and zinc binding proteins Jana Elena Kessels a , Inga Wessels a , Hajo Haase b , Lothar Rink a , Peter Uciechowski a,∗ a b

Institute of Immunology, Medical Faculty, RWTH Aachen University, Pauwelsstr. 30, D-52074 Aachen, Germany Department of Food Chemistry and Toxicology, Berlin Institute of Technology, Gustav-Meyer-Allee 25, D-13355 Berlin, Germany

a r t i c l e

i n f o

Article history: Received 10 November 2015 Received in revised form 4 February 2016 Accepted 10 February 2016 Keywords: Myeloid cells Zinc Znt ZIP Chromatin remodeling Methylation

a b s t r a c t The distribution of intracellular zinc, predominantly regulated through zinc transporters and zinc binding proteins, is required to support an efficient immune response. Epigenetic mechanisms such as DNA methylation are involved in the expression of these genes. In demethylation experiments using 5-Aza2 -deoxycytidine (AZA) increased intracellular (after 24 and 48 h) and total cellular zinc levels (after 48 h) were observed in the myeloid cell line HL-60. To uncover the mechanisms that cause the disturbed zinc homeostasis after DNA demethylation, the expression of human zinc transporters and zinc binding proteins were investigated. Real time PCR analyses of 14 ZIP (solute-linked carrier (SLC) SLC39A; Zrt/IRT-like protein), and 9 ZnT (SLC30A) zinc transporters revealed significantly enhanced mRNA expression of the zinc importer ZIP1 after AZA treatment. Because ZIP1 protein was also enhanced after AZA treatment, ZIP1 up-regulation might be the mediator of enhanced intracellular zinc levels. The mRNA expression of ZIP14 was decreased, whereas zinc exporter ZnT3 mRNA was also significantly increased; which might be a cellular reaction to compensate elevated zinc levels. An enhanced but not significant chromatin accessibility of ZIP1 promoter region I was detected by chromatin accessibility by real-time PCR (CHART) assays after demethylation. Additionally, DNA demethylation resulted in increased mRNA accumulation of zinc binding proteins metallothionein (MT) and S100A8/S100A9 after 48 h. MT mRNA was significantly enhanced after 24 h of AZA treatment also suggesting a reaction of the cell to restore zinc homeostasis. These data indicate that DNA methylation is an important epigenetic mechanism affecting zinc binding proteins and transporters, and, therefore, regulating zinc homeostasis in myeloid cells. © 2016 Elsevier GmbH. All rights reserved.

1. Introduction A balanced zinc homeostasis is essential for an effective immune system [1]. Zinc deficiency negatively affects differentiation, growth, wound healing, thymus development, and the immune response [2–4]. In contrast, elevated zinc serum concentrations are able to influence immune cell functions adversely [5,6]. Most of the cellular zinc of eukaryotic cells is localized in the cytoplasm or within organelles, associated with proteins or anions [7,8]. Therefore, the quantities of free, available zinc in cells are limited [8]. The intracellular distribution and concentration of free zinc depends on fourteen human ZIP (importer), ten ZnT (exporter) zinc transporters, and on zinc binding proteins [1,9–11]. MT, buffering excess zinc and supplying zinc under zinc deficiency [10,12,13], is involved in cellular responses to metal toxicity and oxidative stress [10,14]. S100A8/S100A9 proteins capturing zinc effectively,

∗ Corresponding author. Fax: +49 241 8082613. E-mail address: [email protected] (P. Uciechowski).

are associated with wound healing and markers for inflammatory processes [15,16]. The intracellular zinc concentration influences leukocyte differentiation and function [7,17–19]. Excess of zinc increases pro-inflammatory cytokine production, but also long time zinc deficiency results in up-regulation of these cytokines and in increased reactive oxygen species production [20–22]. Thus, tight regulation of free intracellular zinc in leukocytes by zinc transporters and zinc binding proteins is crucial [23,24]. Molecular mechanisms regulating ZnT1, ZnT5, ZnT10, and MT are zinc-dependent transcriptional activators/repressors binding to metal response elements (MRE) or to the zinc transcriptional regulatory element (ZTRE) [14,25]. DNA methylation, an epigenetic mechanism controlling gene transcription [26–29], of MREs and of ZTREs has been implicated to reduce the capacity to recruit their corresponding binding factors [25,30]. Additionally, methylation of CpG islands in the ZIP8 and ZnT5 promoter regions resulted in decreased gene transcription [31,32]. Thus, the effects of DNA demethylation on zinc transporters and zinc binding proteins and subsequently its influence on zinc homeostasis were investigated in myeloid HL-60 cells.

http://dx.doi.org/10.1016/j.jtemb.2016.02.003 0946-672X/© 2016 Elsevier GmbH. All rights reserved.

Please cite this article in press as: J.E. Kessels, et al., Influence of DNA-methylation on zinc homeostasis in myeloid cells: Regulation of zinc transporters and zinc binding proteins, J Trace Elem Med Biol (2016), http://dx.doi.org/10.1016/j.jtemb.2016.02.003

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Fig. 1. DNA demethylation is associated with a significant increase of intracellular free zinc and total cellular zinc levels. (A) HL-60 cells were incubated for 3, 6, 24 and 48 h with and without 5 ␮mol L−1 AZA. Intracellular free zinc concentrations in HL-60 cells were analyzed using the fluorescent zinc probe FluoZin-3 in flow cytometry. Independent experiments were repeated n = 5. Depicted are mean values ± SEM. Statistical significances were quantified by student’s t-test (*, p < 0.05; **, p < 0.01; ns, not significant) (B) The same experiments as in A were performed with the zinc detection probe ZinPyr-1. Indicated are mean values ±SEM for n = 5 independent experiments. Statistical significances were quantified by student’s t-test (**, p < 0.01; ns, not significant). (C) The graph represents the absolute changes in free intracellular zinc dependent on AZA treatment from (A) and (B). (D) Total cellular zinc levels of AZA treated HL-60 cells and untreated HL-60 cells were measured via atomic absorption (mean values, ±SEM for n = 9; student’s t-test; *, p ≥ 0.05). (E) HL-60 cells were incubated with and without AZA for 48 h, then MTT assays were performed. The quantity of formazan was measured by recording changes in absorbance at 570 nm using a plate reading spectrophotometer (mean values ± SD, n = 3).

2. Material and methods

started by seeding 2 × 106 cells/mL on 6-well plates if not stated otherwise.

2.1. Cell culture 2.2. Treatment with 5-Aza-2 -deoxycytodine HL-60 cells were cultured in RPMI 1640 medium, supplemented with 10% low- endotoxin fetal calf-serum (FCS), 1% penicillin, 1% streptomycin and 1% l-glutamine (all Lonza, Cologne, Germany) in a humidified 37 ◦ C atmosphere containing 5% CO2 . All experiments

To accomplish inhibition of DNA-methyltransferases, AZA (Sigma–Aldrich, Munich, Germany) was added to the culture medium in a concentration of 5 ␮mol L−1 for the indicated times.

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Fig. 2. DNA demethylation results in enhanced mRNA expression of ZIP1 and ZnT3 and reduced expression of ZIP14. (A) HL-60 cells were incubated with and without 5 ␮mol L−1 AZA for 48 h. mRNA from AZA treated and untreated cells was isolated, reverse transcribed and quantified with SYBR Green real-time PCR. The gene expression of different ZIP zinc transporters was normalized to the housekeeping gene PBGD by the 2− Ct method. Indicated are mean values ± SEM for n = 3 independent experiments. Statistical significances were quantified by student’s t-test (*, p < 0.05; **, p < 0.01). (B) The same experiments as in A were performed to analyze the expression of ZnT zinc transporters. The diagram indicates the modulation in ZIP/ZnT mRNA levels of AZA treated cells relative to untreated cells (marked by a dotted line). Independent experiments were n = 3. Error bars represent ± SEM. Statistical significances were quantified by student’s t-test (*, p < 0.05).

The medium was exchanged with fresh AZA supplemented medium after 24 h [28]. Propidium staining using flow cytometry shows only small amounts of dead cells treated with or without AZA after 24 and 48 h (data not shown).

absorbance at 570 nm using a Sunrise plate reading spectrophotometer (Tecan, Crailsberg, Germany).

2.5. Reverse transcription and quantitative real-time PCR 2.3. Zinc measurement using flow cytometry and atomic absorption spectroscopy Free zinc levels in HL-60 cells were measured as described [33] using 1 ␮mol L−1 FluoZin-3AM ester (FluoZin-3) (Life technologies, Darmstadt, Germany) or 10 ␮mol L−1 ZinPyr-1 (NeuroBioTex, Galveston, TX, USA) per sample. In brief, 1 × 106 cells (F) were loaded with FluoZin-3 or ZinPyr-1 for 30 min. Additionally, aliquots of the cells were treated with the zinc specific membrane permeant chelator 50 ␮mol L−1 N,N,N ,N -tetrakis-(2-pyridylmethyl) ethylenediamine (TPEN, Sigma–Aldrich, Taufkirchen, Germany) (Fmin ), or 50 ␮mol L−1 sodium pyrithione (Sigma–Aldrich, Taufkirchen, Germany) plus 100 ␮mol L−1 ZnSO4 (Merck, Darmstadt, Germany) (Fmax ) at 37 ◦ C, 5% CO2 for 30 min. Fluorescence of FluoZin-3 and ZinPyr-1 was analyzed using a FACSCalibur (BD Biosciences, Heidelberg, Germany). Free zinc concentrations were calculated as [Zn] = KD × [(F−Fmin )/(Fmax −F)] with KD = 8.9 nmol L−1 [22] for FluoZin-3 and KD = 0.7 nmol L−1 for ZinPyr-1 [34]. For determination of the total cellular zinc content, 5 × 106 HL60 cells were treated with and without AZA for 48 h. Next, cells were washed, centrifuged and the pellet was acid digested (100 ␮L 65% HNO3 (suprapur), 100 ␮L 33% H2 O2 ) and heated. Samples were dissolved in 0.2% HNO3 and the zinc concentration was determined by flame atomic absorption, using a PerkinElmer Aanalyst 800 atomic absorption spectrometer. 2.4. MTT assay HL-60 cells were incubated with and without AZA for 48 h. After counting, the cells were plated on a 96-well plate with a starting number of 0.2 × 106 cells/100 ␮L and serially diluted. Then, 20 ␮L of MTT (3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide) tetrazolium (5 mg/mL) were added and cells incubated for 1 h. The reaction was stopped by adding dimethyl sulfoxide. The quantity of formazan (presumably directly proportional to the number of viable cells) was measured by recording changes in

Lysates from 4 × 106 HL-60 cells were prepared using 1 mL of Tri reagent (Ambion Life Technologies, Karlsbad, Germany) to isolate total RNA. cDNA was prepared by reverse-transcribing 1 ␮g mRNA using the qScript cDNA Synthesis Kit (Quanta Biosciences, Darmstadt, Germany) according to manufacturer’s instructions. Real-time PCR samples consisted of 5 ␮L dH2 O, 12.5 ␮L Power SYBR Green PCR MasterMix (Thermo Fisher, Rockford, IL 61105, USA), 2 ␮mol L−1 forward and reverse primer each and 5 ␮L of cDNA in a final volume of 25 ␮L. Primers for ZIP1-14, ZnT1-9, hMT1,2 (detecting transcripts of the isoforms MT1A, MT1E, MT1F, MT1G, MT1H, MT1H-like, MT1L, MT1X, and MT2A), S100A8/A9, Porphobilinogen deaminase (PBGD) had been previously described [17,24]. PCR was performed using a Step-One plus RealTime PCR System (Thermo Fisher) with the following conditions: 15 min at 95 ◦ C, followed by 40 cycles of 95 ◦ C for 15 s and 60 ◦ C for 60 s. For ZIP13 an annealing temperature of 62 ◦ C was used. The PCR conditions used for MT1,2 and S100A8/A9 had been described previously [17]. To quantify PCR results the comparative cycle threshold (Ct) method (2−CT ) was applied using PBGD as housekeeping gene.

2.6. Western blotting The preparation of cell extracts from HL-60 cells as well as SDS-polyacrylamide gel electrophoresis, and Western Blot analysis were described previously [22]. Membranes were blocked, and then incubated with a polyclonal rabbit/IgG anti-human ZIP1 antibody (Thermo Fisher, Rockford, IL 61105, USA). After washing, the membranes were incubated with horseradish peroxidaselinked anti-rabbit IgG secondary antibody, followed by detection with LumiGlo reagent (New England Biolabs, Frankfurt a. Main, Germany) on a LAS-3000 (Fujifilm Lifescience, Duesseldorf, Germany). As described recently, ␤-actin was used as control [22]. Relative density of the bands was determined with ImageJ software (Luke Miller; lukemiller.org).

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[28]. Experiments, in which AZA treatment did not result in significant enhanced IL-6 production after 48 h, were excluded. 2.8. Chromatin accessibility by real-time (CHART)-PCR assay The method has been previously described in detail [28,35]. In brief, nuclei from 5 × 106 cells were treated with 200 U MNase (Fermentas, St. Leon-Rot, Germany) for 60 min at 37 ◦ C, after which the reaction was stopped by the addition of 80 ␮L MNase stop buffer (0.5 mmol L−1 EDTA, 10 mmol L−1 EGTA). 25 mg/mL Proteinase K (Fermentas) and 2% SDS were added for overnight incubation at 37 ◦ C to achieve protein digestion. Genomic DNA was extracted using the QIAamp DNA Mini kit (Qiagen, Hilden, Germany). Realtime PCR was performed in 25 ␮L reaction volumes in duplicates using Power SYBRGreen PCR MasterMix (Life technologies, Darmstadt, Deutschland) containing 100 ng of DNA. PCR samples were composed as described. Primers for ZIP1 promoter regions I and II were generated with reference to Makhov and colleagues [36] and gene bank accession number NT 00448 using the Primer Express software 2.0 (Thermo Fisher), verified by nucleotide blast search (NCBI). ZIP1 promoter region I primer, forward: TTTCCTAATATCCGCTCTTGCTC; reverse: GTACCCACGACCGACTGCC (product size 151 bp). ZIP1 promoter region I contains binding sites for SP1, GATA binding protein 1 (GATA1), Myeloblastosis proto-oncogene protein (C-MYB) and cAMP response element-binding protein (CREB) [36]. ZIP1 promoter region II primer, forward: TTGGCAGTCGGTCGTGGGTAC; reverse: GAGGGTCTCACTACATGGTCTCC (product size 114 bp). ZIP1 promoter region II possesses transcription factor bindings sites for specificity protein 1 (SP1), nuclear factor kappalight-chain-enhancer of activated B-cells (NF␬B) and transcription factor E2F (E2F) [36]. Primer pairs for IL-1 region VIII and for IL-6 region I had been previously described [28,35]. Real-time PCR was performed using the following conditions: 15 min at 95 ◦ C, followed by 40 cycles of 95 ◦ C for 30 s and 58 ◦ C for 60 s; only for IL-6 region I an annealing temperature of 57 ◦ C was used. Ct values were correlated to percent accessibility using a standard curve generated from serial dilutions of genomic DNA. MNase  accessibility was calculated  according to the formula ˛ = |100 −

quantity(MNase+) quantity(MNase−)

× 100 | %.

2.9. Statistical analyses

Fig. 3. ZIP1, ZIP14 and ZnT3 mRNA levels significantly increase after DNA demethylation treatment. HL-60 cells were incubated with and without 5 ␮mol L−1 AZA for 3, 6, 24 and 48 h. mRNA from AZA treated and untreated cells was isolated, reverse transcribed and quantified with SYBR Green real-time PCR. The gene expression of ZIP1 (A), ZIP14 (B), and ZnT3 (C) were normalized to the housekeeping gene PBGD by the 2− Ct method. Expression of ZIP1, ZIP14, and ZnT3 mRNA in AZA treated cells was compared to untreated controls at each time point (dotted line). Indicated are mean values ± SEM for n = 3 independent experiments. Statistical significances were quantified by student’s t-test (*, p < 0.05; **, p < 0.01).

2.7. Enzyme-linked immunosorbent assay (ELISA) To assess IL-6 expression as positive control for DNA demethylation supernatants of AZA treated cells were collected at the indicated time points and frozen at −20 ◦ C for later analysis. IL-6 production was quantified from the thawed supernatants with the OptEIA ELISA kit (BD Bioscience, Heidelberg, Germany) according to manufacturer’s instructions using a Sunrise ELISA-reader (Tecan)

Statistical significance of results was analyzed by Student’s t-test (for paired samples) using the GraphPad Prism software (GraphPad Software, La Jolla, CA, USA). Kolmogorov-Smirnov test (KS-test) was used for data normalization. Error bars represent the standard error of the mean (SEM) or standard deviation (SD), as indicated. * indicates a significance of p < 0.05, **p < 0.01 and ***p < 0.001; ns, non-significant. 3. Results 3.1. The intracellular zinc concentration in HL-60 cells are increased after incubation with the DNA methyltransferase inhibitor 5-Aza-2 -deoxycytidine To investigate the effect of DNA methylation on zinc homeostasis, the promyelocytic leukemia cell line HL-60 was used. Recently, it was shown that zinc deficiency as well as DNA demethylation result in significantly enhanced promoter accessibilities and expression of inflammatory cytokines such as IL-1␤, IL-6, and tumor necrosis factor (TNF)-␣ in this cell line [22,28,35]. HL-60 cells were treated with 5 ␮mol L−1 DNA methyltransferase inhibitor AZA, a concentration used in previous studies [28,35]. Viability assays using propidium iodide staining confirmed

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5 ␮mol L−1 AZA as the non-cytotoxic concentration being appropriate for the experiments (data not shown). First, it was examined whether AZA-dependent DNA demethylation was able to influence intracellular free zinc levels. Two different zinc sensitive fluorescent probes FluoZin-3 and ZinPyr1 were used [34,37]. The zinc sensor probes vary in their affinity to zinc as well as in their cellular locations after staining, and had been mainly investigated in T cells, less in myeloid cells. After incubation with AZA for 24 h and 48 h both zinc sensors showed a very similar significant increase of intracellular zinc compared to untreated cells (Fig. 1A–C). Interestingly, the use of FluoZin-3 probes revealed as twice as high basal free zinc levels as with ZinPyr-1 in HL-60 cells. Taken together, the intracellular zinc concentration in HL-60 cells was increased in response to AZA treatment after 24 h, and this effect continues for additional 24 h. To confirm our data obtained from fluorescent probes, the total zinc levels in HL-60 cells were determined in an atomic absorption spectrometer. The results revealed a significant higher zinc concentration in AZA treated HL-60 cells compared to untreated HL-60 cells (Fig. 1D). To check the metabolic reactions of HL-60, MTT assays were performed. In these experiments it could be shown that HL-60 treated with AZA for 48 h have very similar metabolic activities as untreated HL-60 cells (Fig. 1E). 3.2. DNA demethylation in HL-60 cells leads to the up-regulation of ZIP1 and ZnT3 mRNA but reduces ZIP14 expression Increased zinc levels after AZA treatment indicate that changes in zinc transporter expression may be involved. Next, the cellular zinc transporter mRNA expression with and without AZA incubation (48 h) was investigated by real time PCR. All human ZIP and ZnT zinc transporters with the exception of ZnT10, whose expression is limited to fetal liver and fetal brain cells [9], were analyzed (Fig. 2). HL-60 cells expressed all ZIP mRNAs except for ZIP12 and all ZnT mRNAs with the exception of ZnT2 and ZnT8 (Fig. 2A and B). These results are in accordance with previous data from experiments with HL-60 and THP-1 cells [17,24]. After 48 h of treatment with AZA, the expression of ZIP1 and ZnT3 mRNAs was significantly increased when compared to untreated cells (Fig. 2A and B). Only the expression of Zip14 mRNA was significantly reduced (Fig. 2A). To investigate if demethylation leads to earlier alterations in mRNA expression of zinc transporters, kinetic experiments were performed. As a result, ZIP1 mRNA expression steadily increased over time, which was in line with the constantly rising free zinc concentrations after AZA treatment (Figs. 1 and 3A). The downregulation of ZIP14 mRNA only became significant after 48 h. (Fig. 3B). A significant change in the expression of ZnT3 mRNA was also observed after 48 h incubation with AZA (Fig. 3C). The data suggest that the time-dependent regulation of ZIP1 mRNA corresponds to the increase of intracellular zinc, whereas a similar correlation could not be observed for ZIP14 and ZnT3 expression. Western blot (Fig. 4A) and densitometric (Fig. 4B) analyses could show that there was a significant enhancement of ZIP1 protein in HL-60 incubated with AZA for 48 h compared to untreated cells. Enhanced mRNA and protein expression of the zinc importer ZIP1 may represent an explanation for the increase in intracellular free zinc upon AZA treatment. Therefore, the regulation of ZIP1 gene expression was subjected to a more detailed analysis. 3.3. Effects of DNA demethylation on the accessibility of the human ZIP1 promoter DNA methylation can result in altered promoter accessibility, subsequently affecting gene expression. To address whether AZA induced ZIP1 mRNA up-regulation could be a consequence

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of an altered ZIP1 promoter conformation, CHART assays were performed [22,28,35]. The effect of AZA was verified through significantly increased IL-6 production after 48 h of AZA treatment (Fig. 5A) [28]. The ZIP1 gene located on chromosome 1 has two promoter regions containing CpG-rich areas and important binding sites for several transcriptions factors (material and methods section) [36]. The ZIP1 promoter regions I (−1028 to −879) and II (−898 to −785) were investigated. ZIP1 promoter region II was completely inaccessible in HL-60 cells (Fig. 5B), even after AZA treatment. Region I of the ZIP1 promoter was 66.3% accessible in HL-60 cells, and opened up to 80.4% after 48 h incubation with AZA. But this increase was not significant (Fig. 5B). The IL-1 promoter region VIII (−673 to −583) remained inaccessible as described [35]. IL-6 promoter region I (+13 to −139) revealed significant accessibility after AZA incubation (Fig. 5B), as described before [28]. The ZIP1 promoter region I is already opened in HL-60 cells. Although there was an enhancement of accessibility after AZA treatment, DNA demethylation does not appear to influence ZIP1 promoter I, II accessibility. 3.4. DNA demethylation by 5-Aza-2 -deoxycytidine increases the expression of MT and S100A8, S100A9 zinc binding proteins Besides zinc transporters zinc binding and storing proteins contribute to the cellular zinc homeostasis [10,17]. Therefore, the influence of DNA methylation on gene expression of the main zinc regulating proteins of myeloid cells was also investigated in HL-60 cells. For the detection of MT, primers covering nine different isoforms of MT1 and 2 were used [17]. After incubation of HL-60 cells with AZA for 3, 6, 24 and 48 h, a significant increase in mRNA expression of S100A8, S100A9 and MT was detected after 48 h (Fig. 6A–C). S100A8 and S100A9 showed the highest mRNA up-regulation after AZA treatment (Fig. 6). Interestingly, a significant amplification of MT mRNA could be shown after 24 h, in addition (Fig. 6C). The data reveal that zinc binding proteins contributing to zinc homeostasis are regulated epigenetically via DNA methylation. 4. Discussion The present study reports changes in zinc metabolism in HL-60 cells induced by the DNA-demethylating agent AZA. Alterations of mRNA as well as protein expression of zinc transporter ZIP1 and zinc binding proteins MT and S100A8/9 via demethylation might be the potential underlying mechanism. The values for zinc concentrations in different cell lines reported in the literature vary from at least 10−5 –10−12 mol L−1 [11]. The concentrations measured in our study using fluorescent zinc probes were ∼10−11 mol L−1 and are comparable to formerly published data for HL-60 cells [22]. Depending on the probe used slightly different baseline concentrations of intracellular free zinc were measured (Fig. 1A and B). AZA treatment for 24 h and 48 h resulted in approximately same rises of intracellular zinc compared to untreated cells (Fig. 1A–C). Additionally, the determination of total cellular zinc concentrations (Fig. 1D) supported these observations, showing significant higher zinc concentrations in AZA treated HL60 cells than untreated HL-60 cells. The total zinc concentration in untreated HL-60 cells (6.6 (mean) ng zinc/106 ) was similar to those reported for mononuclear cells (7.4 (mean) ng zinc/106 ) [38]. The intracellular zinc levels are controlled to a high extent by zinc transporters; consequently, the expression of zinc transporters was analyzed under DNA demethylating conditions. The data reveal a significant increase of ZIP1 and ZnT3 mRNA expression after 48 h of incubation with AZA, whereas ZIP14 expression declined (Fig. 2).

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Fig. 4. ZIP1 protein is enhanced in HL-60 cells after DNA demethylation treatment. (A) Human ZIP1 protein expression in AZA treated and untreated HL-60 cells (48 h) was analyzed by Western blotting with antibodies against ZIP1 and ß-actin (n = 3). The blot with three samples from untreated (−) and AZA treated (±) cells shown is representative of n = 11 independent experiments. (B) The relative density of the protein bands of ZIP1 and ␤-actin was determined from AZA treated cells (AZA) and untreated controls (Med) are shown. Mean values ± SEM of n = 11 independent experiments. Statistical significances were quantified by student’s t-test (*, p < 0.05).

Fig. 5. Effects of DNA demethylation on the accessibility of the ZIP1 promoter. (A) Significant production of IL-6 after AZA treatment by HL-60 cells was the positive read-out for effective DNA demethylation throughout this study. Supernatants of untreated and AZA treated HL-60 cells (24, 48, and 72 h) were measured by ELISA. Bars represent mean values of IL-6 secretion ± SEM of n = 5 experiments. Statistical significances were quantified using student’s t-test (***, p < 0.001; ns, not significant). (B) HL-60 cells were incubated with and without 5 ␮mol L−1 AZA for 48 h. CHART assays were performed with primer sets for region I and II of the ZIP1 promoter, for region VIII of the IL-1 promoter (closed region), and for region I of the IL-6 promoter (open region after AZA treatment). Accessibility for treated and untreated samples was determined as described in materials and methods. The diagram shows mean values ± SEM of n = 3 independent experiments. Statistical significance was determined using student’s t-test (*, p < 0.05; ns, not significant).

The fact that only 3 out of 23 tested zinc transporter genes in HL-60 cells were influenced by AZA treatment suggests a specific methylation-dependent control mechanism rather than a broad impact on a whole functional groups of genes [39,40]. ZIP1 is ubiquitously expressed in human tissues and has been regarded as an important regulator of zinc homeostasis [41]. Our mRNA and Western blotting data indicate that ZIP1 is presumably involved in AZA-induced increase in free cellular zinc. The continuous increase of ZIP1 expression during AZA treatment supports the statement, whereas ZnT3 seems to play a minor role as only detected after 48 h (Figs. 1and 3). Our results are in line with other reports showing a correlation between increased ZIP1 expression and intracellular zinc levels. In one study, healthy prostate gland cells accumulated high levels of zinc. This ability was lost during malignant transformation accompanied by silencing of the ZIP1 gene [42]. In zinc deficient cells an increased expression of ZIP1 and translocation to the plasma membrane has been reported [43,44]. Vice versa, zinc sufficient conditions can lead to a reduced expression of the gene and the internalization of ZIP1 [41,43–45]. ZIP14 and ZnT3 have mainly been detected in lymphocytes and neurons, respectively. However, recent data suggest their

expression in other leukocytes as well [9,11,46,47]. The expression of ZnT3 was also observed in a number of other tissues and recently in myeloid cells [48]. The reduced expression of the zinc importer ZIP14 and the increased expression of the zinc exporter ZnT3 would be expected to lead to a reduction in cytoplasmic zinc levels. Since high intracellular zinc levels have toxic effects on several cellular functions [49], it cannot be excluded that this expression pattern prevents increase of cytoplasmic zinc above a certain threshold. To answer whether methylation could affect the accessibility of the ZIP1 promoter, two separate regions (I, II) characterized by a clustering of CpGs [36] with transcription factor binding sites were investigated by CHART. The ZIP1 region II was found to be inaccessible in HL-60 cells. Basic accessibility of region I was as high as 66.3% in untreated cells. AZA increased the accessibility, but not significantly (Fig. 5), so that DNA methylation alone does not seem to alter chromatin structure in this region of the ZIP1 promoter. If DNA methylation modulates binding abilities of SP1, GATA1, C-MYB and CREB transcription factors [36] to the ZIP1 promoter remains also to be determined.

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Fig. 6. AZA treatment of HL-60 cells results in increased mRNA expression of the zinc binding proteins S100A8/A9 and MT. HL-60 cells were incubated with and without 5 ␮mol L−1 AZA for 3, 6, 24 and 48 h. mRNA from AZA treated and untreated cells was isolated, reverse transcribed and quantified with SYBR Green real-time PCR. The gene expression of S100A8 (A), S100A9 (B), and MT (C) were normalized by the 2− Ct method. Expression of S100A8, S100A9, and MT mRNA in AZA treated cells was compared to untreated controls at each time point (dotted line). Indicated are mean values ± SEM for n = 3 independent experiments. Statistical significances were quantified by student’s t-test (*, p < 0.05; **, p < 0.01).

The expression of MT in most cells is increased in response to zinc excess but is down-regulated under zinc deficient conditions, an effect that is mediated by the metal responsive transcription factor 1 (MTF-1) [23,50]. Its zinc-dependent binding to an MRE in the MT promoter is essential for transcription of the gene [51]. Interestingly, methylation of CpG islands within this MRE can interfere with binding of MTF-1 and leads to a suppression of MT expression in lymphoid-derived cancer cells [30,52]. DNA methylation is an

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important epigenetic regulator contributing to gene repression of MT1 gene cluster in estrogen receptor positive and negative breast tumors [53]. Thus, increased zinc levels and demethylation of the MT promoter might contribute to increased MT expression in AZAtreated HL-60 cells, which is indicated by the observed increase of MT mRNA expression after 24 and 48 h (Fig. 6C). There is one report revealing that S100A8 and S100A9 mRNA were found to be up-regulated after 72 h of AZA treatment [54]; but the authors did not perform quantitative real-time PCR and there were no detailed descriptions about PCR conditions the authors had used. Zinc deficiency led to enhanced expression of S100A8 and S100A9 in THP-1 cells [55]. In our study increased expression of S100A8/A9 mRNA upon DNA demethylation was observed in presence of elevated zinc levels. Although a cellular counteracting effect to reduce excess zinc cannot be ruled out, the observed mRNA enhancement might directly be linked to DNAmethylation. Hyper-methylation of these genes and corresponding down-regulation has previously been described in MLL rearrangement leukemias [56], which supports this hypothesis. Furthermore, it has been demonstrated that methylation of MREs and ZTREs results in reduced gene expression [25,30,53]. Therefore, methylation of corresponding zinc responsive elements in ZIP, ZnT and metallothionein promoters is suggested to inhibit the expression of these genes in HL-60 cells, explaining why expression is increased after AZA treatment. In summary, the present study reveals that DNA methylation influences zinc levels in a myeloid cancer cell line by altering the expression of genes involved in zinc homeostasis. The data indicate that alteration of mRNA expression of zinc transporter and zinc binding proteins might be the underlying mechanisms. Hypermethylation of the ZIP1 promoter in HL-60 cells may reduce the expression of this key zinc importer, leading to elevated zinc levels after AZA treatment. Since ZIP-1 mRNA is earlier up-regulated through demethylation, and the highest intracellular zinc levels are reached after 24 h, the elevated zinc concentrations may lead to increased MT expression after 24 h, and ZnT3 and S100A8/9 expression after further 24 h to restore zinc homeostasis. However, one cannot exclude that other genes activated after AZA-treatment are also involved. Zinc deficiency is frequently observed in the elderly, and current studies indicate that zinc deficiency/dysregulated zinc homeostasis is linked to neurodegenerative disorders such as Parkinson’s disease (PD), and Alzheimer’s disease (AD) [57–59]. Additionally, altered expression of zinc transporters such as ZIP-1 has been reported in AD [60,61]. Although the functional role of zinc in AD and PD pathogenesis is still not clarified in vivo, zinc supplementation may be beneficial in the treatment of AD and PD, as shown in a Drosophila Parkinson’s model or in AD mice [62,63]. DNA methylation is also enhanced in aging and has been proposed as an underlying cause of zinc deficiency [64]. Both conditions are involved in cancerogenesis and have been implicated in immune diseases such as systemic lupus erythematosus [65–68]. Our results might contribute to a better molecular understanding of human diseases where dysregulated zinc homeostasis and DNA methylation are involved. Conflict of interests None of the authors declared a conflict of interest and disclose any financial and personal relationships with other people or organizations that could inappropriately influence (bias) their work. References [1] L. Rink, H. Haase, Zinc homeostasis and immunity, Trends Immunol. 28 (2007) 1–4.

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