Inhibition of red blood cell development by arsenic-induced disruption of GATA-1

Anemia is a hematological disorder that adversely affects the health of millions of people worldwide. Although many variables influence the development and exacerbation of anemia, one major contributing factor is the impairment of erythropoiesis. Normal erythropoiesis is highly regulated by the zinc finger transcription factor GATA-1. Disruption of the zinc finger motifs in GATA-1, such as produced by germline mutations, compromises the function of this critical transcription factor and causes dyserythropoietic anemia. Herein, we utilize a combination of in vitro and in vivo studies to provide evidence that arsenic, a widespread environmental toxicant, inhibits erythropoiesis likely through replacing zinc within the zinc fingers of the critical transcription factor GATA-1. We found that arsenic interacts with the N- and C-terminal zinc finger motifs of GATA-1, causing zinc loss and inhibition of DNA and protein binding activities, leading to dyserythropoiesis and an imbalance of hematopoietic differentiation. For the first time, we show that exposures to a prevalent environmental contaminant compromises the function of a key regulatory factor in erythropoiesis, producing effects functionally similar to inherited GATA-1 mutations. These findings highlight a novel molecular mechanism by which arsenic exposure may cause anemia and provide critical insights into potential prevention and intervention for arsenic-related anemias.


AsIII inhibits erythropoiesis in vivo.
To demonstrate that AsIII exposure inhibits erythropoiesis in vivo, we exposed male C57BL/6J mice to 20, 100, and 500 ppb AsIII (0.3, 1.3, and 6.7 µM, respectively) via drinking water for 30 days. Early erythroid cells (CD71 + /Ter119 − ; late BFU-E and CFU-E) were assessed in bone marrow by CD71 and Ter119 surface marker expression using flow cytometry 49,50 . AsIII exposure reduced the total number and percentage of early erythroid cells in a dose-dependent manner, starting from 0.02 µM (Fig. 1a-c). To further demonstrate that AsIII compromises the differentiation of early erythroid progenitors, we assessed the colony-forming ability of CFU-E from bone marrow of AsIII exposed mice. CFU-E colony formation was reduced by AsIII, starting at 20 ppb and was suppressed by over 60% with 500 ppb AsIII (Fig. 1d). These results provide evidence that in vivo drinking water exposure to environmentally relevant levels of AsIII suppress the differentiation of early erythroid progenitors in the bone marrow.
AsIII inhibits erythropoiesis, but not myelopoiesis. As an effort to characterize the inhibitory effects of AsIII on erythropoiesis and to investigate whether AsIII impairs other hematopoietic lineages, we developed an in vitro model of erythropoiesis using primary mouse bone marrow HPC stimulated with erythropoietin (EPO) and stem cell factor (SCF) 51 . The progression of HPC through the stages of hematopoiesis were assessed based on surface marker phenotype using flow cytometry 52,53 . We identified a significant reduction in the numbers of erythro-megakaryocytic progenitors (MEP, p = 0.0114; BFU-E, p = 0.00063; and CFU-E, p = 0.0056) after 24 h exposure to 0.5 μM AsIII (Fig. 2a,b and Supplementary Fig. S1), but no significant changes in CMPs were observed with either AsIII dose (Fig. 2b). Additionally, the number of early myeloid progenitors (i.e. pre-granulocyte macrophage (pre-GM)) was significantly increased (p = 0.0192) with AsIII exposure (Fig. 2c and Supplementary Fig. S1).
These results show that the AsIII-induced inhibition of erythropoiesis starts from very early stages of erythroid differentiation. AsIII likely disrupts the transition of CMP to MEP and subsequent differentiation of early stages of erythroid progenitors (i.e. processes dependent on GATA-1), rather than reducing the CMP population directly. Interestingly, the suppression of erythropoiesis skews hematopoietic differentiation in favor of myelopoiesis, despite EPO stimulation and the lack of myeloid supportive growth factors.
To determine whether the suppressive effects of AsIII on early erythroid progenitor differentiation persist to later-stages of maturation, we evaluated the progression of erythroblast differentiation every 24 h for 72 h based on CD71 and Ter119 surface marker expression and cell size 49,50 . A significant suppression of early erythroblast subsets (late BFU-E, CFU-E, and proerythroblasts, p = 0.0042) and basophilic erythroblasts (EryA, p = 0.0162) was observed after 24 h exposure to 0.5 μM AsIII (Fig. 2d,e and Supplementary Fig. S1). These effects were persistent up to 72 h, with substantially fewer AsIII exposed erythroblasts properly transitioning to late stages of  S1). Collectively, these results suggest that AsIII inhibits early stages of erythropoiesis and these effects are persistent throughout erythroblast differentiation, resulting in decreased production of mature erythroblast subsets. However, the suppressive effects of AsIII were selective to erythro-megakaryocytic progenitors, as myeloid progenitors were not reduced by AsIII exposure.
To further demonstrate that AsIII selectively inhibits erythropoiesis, we tested the effects of AsIII on the hemin or PMA-induced erythroid 54 , megakaryocytic, myelocytic, and monocytic differentiation, respectively [54][55][56][57] . In hemin stimulated K562 cells, erythropoietic differentiation was measured using surface marker expression of CD71 and CD235a by flow cytometry. Since the K562 cell line has some resistance to arsenic 58 , slightly higher concentrations of AsIII were used in these experiments. AsIII at 1 µM significantly reduced the percentage of CD71 + and CD235a + erythroid differentiated K562 cells (p = 0.0221 and 0.0469, respectively; Fig. 3a,b). Megakaryocytic differentiation of K562 cells was determined using CD41 surface marker expression by flow cytometry after exposure to 1 μM AsIII. AsIII significantly reduced the percentage of PMA differentiated megakaryocytic (CD41 high ) K562 cells (p = 0.0384; Fig. 3c). Further evaluation of forward scatter and CD11b surface marker expression (i.e. characteristics of myelopoiesis and monopoiesis, respectively), showed no significant differences with 1 μM AsIII (Fig. 3d,e).
These results showed that in the K562 cell model, AsIII also inhibits erythropoiesis and megakaryopoiesis, but not myelopoiesis or monopoiesis. The selective inhibition of erythroid and megakaryocytic differentiation and the lack of effects on myelocytic progenitors in two independent cell models (primary mouse bone marrow HPC and K562 cells) suggest that AsIII may be disrupting the function of GATA-1. These findings motivated us to follow-up with mechanistic studies focused on understanding the effects of AsIII on a prominent regulatory ZF transcription factor in erythropoiesis and a critical non-ZF regulator of myelopoiesis, GATA-1 and PU.1, respectively.

Figure 1.
Inhibition of bone marrow erythropoiesis in vivo by AsIII exposure. Male C57BL/6 J mice were exposed to 0, 20, 100, and 500 ppb AsIII (0, 0.3, 1.3, and 6.7 µM, respectively) in drinking water for 30 days. Bone marrow cells were isolated from the femurs of each mouse and utilized for flow cytometry and CFU-E colony forming assays. (a) Representative flow cytometry dot plot depicting effects of 0 and 500 ppb AsIII on early erythroid progenitor cells (EryP; CD71 + , Ter119 − ; late BFU-E and CFU-E) defined by CD71 and TER119 surface marker expression. (b) Total numbers or (c) percentages of early erythroid progenitor cells measured by CD71 and TER119 surface marker expression using flow cytometry. (d) Inhibition of CFU-E colony formation (shown as number of colonies/10 6 bone marrow cells). Data are expressed as mean ± SD, n = 5 mice/group, *p < 0.05, ** p < 0.001 in one-way ANOVA, followed by Tukey's post hoc test compared to no treatment group.
Using the RayBiotech Human GATA-1 Transcription Factor Activity Assay Kit, we observed the same trend of decline of GATA-1 activity (Fig. 4b). The DNA binding activity of GATA-1 was even more sensitive in primary mouse bone marrow erythroid cells with an inhibition of GATA-1 DNA binding found with very low concentrations of 0.1 and 0.5 μM AsIII (30% and 50%, respectively) (Fig. 4c). Further confirmation of the AsIII-induced impairment of GATA-1 DNA binding activity was found using chromatin immunoprecipitation (ChIP) coupled with quantitative real-time PCR (qPCR) in AsIII treated primary mouse bone marrow erythroid cells (Fig. 4d).  www.nature.com/scientificreports/ In addition to DNA binding activity, GATA-1 interaction with FOG-1 is also critical for its function as a transcriptional regulator 60 . We tested AsIII effects on the interaction between GATA-1 and FOG-1 using coimmunoprecipitation followed by western blotting. A significant inhibition of GATA-1 and FOG-1 interaction was found in primary mouse bone marrow erythroid progenitors starting after 24 h exposure to 0.1 and 0.5 µM AsIII (p = 0.0298 and 0.0211, respectively; Fig. 4e,f and Supplementary Fig. S4). Consistently, a suppression of GATA-1 and FOG-1 interaction was also found in K562 cells starting from 0.5 μM AsIII (Fig. 4g). At 2 μM concentration, AsIII reduced the interaction between GATA-1 and FOG-1 by ~ 75% (Fig. 4h). To demonstrate that AsIII inhibition of GATA-1 is through a zinc dependent mechanism, we used the specific zinc chelator, TPEN to demonstrate the necessity of zinc for GATA-1/FOG-1 interaction. We found that treating K562 cells with TPEN markedly decreased GATA-1/FOG-1 interaction (Fig. 4g,h and Supplementary Fig. S5), suggesting that AsIII treatment is functionally equivalent to zinc chelation in terms of removing zinc from GATA-1 protein and inhibiting GATA-1/FOG-1 interaction.
Since AsIII treatment decreased GATA-1 DNA binding and interaction with FOG-1, these findings suggest that AsIII disrupts the function of both the C-and N-terminal ZF domains of GATA. Our results show that AsIII inhibits GATA-1 DNA binding and protein-protein interaction activities, demonstrating GATA-1 as a molecular target of AsIII. To demonstrate that the AsIII-induced inhibition of GATA-1 is through ZF disruption, we treated K562 cells and primary mouse bone marrow erythroid progenitor cells with AsIII for 48 and 24 h, respectively. GATA-1 and PU.1 protein was purified from cell lysates using immunoprecipitation and the zinc and arsenic contents were measured by inductively coupled plasma-mass spectrometry (ICP-MS). In K562 cells treated with 1 or 2 μM AsIII, zinc contents were decreased by over 50% (Fig. 5a) and arsenic content was considerably increased (Fig. 5b). GATA-1 from primary mouse bone marrow erythroid progenitor cells showed greater sensitivity to AsIII. Zinc content in GATA-1 was decreased at 0.1 and 0.5 μM AsIII, with 0.5 μM reducing GATA-1 zinc content by approximately 50% (Fig. 5e). Similar to K562 cells, a drastic increase in GATA-1 arsenic content was observed (Fig. 5f). In contrast, AsIII did not modulate zinc content in PU.1, nor bind to PU.1 in either K562 or primary mouse bone marrow erythroid progenitor cells (Fig. 5c,d,g,h), indicating that PU.1 is not a molecular target of AsIII. Collectively, these results demonstrated that AsIII interacts with GATA-1, reducing zinc content and thus inhibiting DNA and protein binding activities (Fig. 4). More importantly, the interaction of AsIII with GATA-1 was ZF specific, as no changes to PU.1 were found.

AsIII causes GATA-1 zinc finger dysfunction in vivo.
To further demonstrate that GATA-1 is a sensitive molecular target of AsIII in vivo, we measured arsenic binding and zinc loss using GATA-1 protein collected from bone marrow of C57BL/6J mice exposed to 500 and 1000 ppb AsIII (6.7 and 13.3 µM) via drinking water for 2 weeks. GATA-1 protein was immunoprecipitated from bone marrow cells and ICP-MS was performed to measure zinc and arsenic content in GATA-1. Consistent with findings from our in vitro studies, zinc content in GATA-1 was significantly decreased with 1000 ppb AsIII (p = 0.0477; Fig. 6a). We also found a substantial increase of arsenic content in GATA-1 from mice exposed to 1000 ppb AsIII (Fig. 6b). These results corroborated our in vitro findings and provided additional evidence that AsIII interacts with GATA-1 to cause zinc loss and protein dysfunction in vivo.

Discussion
In this study, we found that AsIII selectively inhibits erythropoiesis, but not myelopoiesis. A prominent factor that contributes to this effect is GATA-1. GATA-1 is a ZF protein with two ZF domains, whereas PU.1 is not a ZF protein. The N-terminal ZF of GATA-1 is responsible primarily for binding with FOG-1 and the C-terminal ZF is responsible for DNA binding 18 . Our previous work in DNA repair inhibition by AsIII demonstrated that ZF proteins such as PARP-1 are sensitive molecular targets of AsIII 42 . We found that AsIII selectively interacts with ZF motifs with ≥ 3 cysteine residues 45 , which are a minority of ZF proteins. Both N-and C-terminal ZFs of GATA-1 contain 4 cysteine residues, making GATA-1 structurally favorable for arsenic binding. In contrast, PU.1 contains only two spatially separated cysteine residues and does not have zinc in its structure. As a result, AsIII does not interact with PU.1.
Our results showed that AsIII exposure not only impaired GATA-1 DNA binding, but also inhibited the interaction of GATA-1 with FOG-1, indicating that AsIII likely binds to both ZFs of GATA-1. Future studies will focus on demonstrating the precise interactions of AsIII with the zinc finger motifs in the GATA-1 protein.
Interaction with FOG-1 and DNA binding are both necessary functions of GATA-1 for commitment of HPC to the erythroid lineage and promotion of normal erythropoiesis 18,60 . Previous studies report that a single germline mutation on just one ZF of GATA-1 leads to dyserythropoietic anemia 25,26 . Additionally, a mutation in the N-terminal ZF of GATA-1 was found to result in X-linked thrombocytopenia and β-thalassemia 61 . These reports suggest that disruption of either ZF on GATA-1 can cause dysfunction resulting in hematological disorders. Our results suggest that AsIII impairs both ZF motifs on GATA-1, likely explaining why such low concentrations of AsIII cause an inhibition of erythropoiesis.
The interaction of AsIII with GATA-1 not only impaired erythropoiesis, but also produced a shift from erythropoiesis in favor of myelopoiesis (Fig. 7). The lineage commitment of CMPs is regulated by the functional antagonism of GATA-1 and PU.1 10,11,13,62 . Studies have shown that ectopic expression of GATA-1 or PU.1 blocks myeloid or erythroid differentiation, respectively through the functional repression of gene activation 11,62 . The  In this study we focused on GATA-1 because in addition to being essential for normal erythropoiesis, the peak expression and activity levels of GATA-1 occur during the stages of erythroid differentiation (i.e. BFU-E, CFU-E, proerythroblast) 15,17,18 which we have found in this and previous studies to be most impacted by AsIII exposure 41 . However, there are other zinc finger proteins which have important roles during hematopoiesis that may also be impacted by arsenic exposure 63 . In particular, GATA-2 is a zinc finger transcription factor, which is similar to GATA-1, contains two C4 zinc finger motifs 64,65 . GATA-2 is highly expressed in HSC and has essential regulatory functions during hematopoiesis [65][66][67][68][69][70] , in part through its activity in promoting the erythroid lineage commitment of HPC through the activation of GATA-1 expression 71,72 . The effects on GATA-1 observed in this study appear to be mediated through interactions of AsIII with the ZF motifs of GATA-1, as no effects on GATA-1 expression levels were found. It is plausible, however, that indirect effects mediated through other zinc finger proteins (e.g. GATA-2) may also contribute to the disruption of erythropoiesis, and as such will be a focus of future investigations.
In this work, the concentrations of AsIII that produced an inhibition of erythropoiesis started at 20 ppb (~ 0.3 µM), which is lower than 12% of the AsIII concentrations measured in water supplies in north central and western regions of the United States 33,34 . Some unregulated water sources in rural areas around the world contain much higher AsIII levels 33,34 . Many regions with high endemic AsIII are also reported to be disproportionately affected by anemia, even after statistical adjustment for common risk factors [35][36][37]39 . These studies emphasize the necessity for gaining a deeper mechanistic understanding of arsenic-caused anemias.
Findings from the present study showed that GATA-1 is a sensitive target of AsIII. GATA-1 ZF disruption was observed with environmentally relevant concentrations of AsIII in vitro and in vivo. We identified a molecular mechanism by which AsIII causes GATA-1 dysfunction producing effects functionally similar to germ-line mutations of GATA-1. Such mutations of GATA-1 ZF result in severe dyserythropoietic anemia 25,26 . Although inherited dyserythropoietic anemia is rare, millions of people worldwide are chronically exposed to AsIII through drinking water. This presents a very important environmental health issue that requires further attention from the scientific community.
Our work demonstrates for the first time that AsIII, a widespread environmental toxicant, inhibits erythropoiesis through disruption of the ZF transcription factor, GATA-1. AsIII disrupts the function of GATA-1 likely through interactions with its ZF motifs, resulting in dyserythropoiesis and an imbalance of HPC differentiation. Mice and in vivo drinking water exposures. Experiments involving mice were performed in accordance with protocols approved by the Institutional Animal Use and Care Committee at the University of New Mexico Health Sciences Center. All experiments were performed in accordance with relevant guidelines and regulations. Male C57BL/6J mice were purchased from Jackson Laboratory and acclimated for one week prior to in vivo and in vitro experiments. All mice used for in vitro and in vivo studies were approximately 12-13 weeks of age. Male mice were used in this study based on our reported observation of a decreased red blood cell counts in a group of men from rural Bangladesh who were exposed to AsIII in their drinking water 38 , to avoid any potential complication due to gender difference. Mice were maintained on a 12:12 reverse light:dark cycle and fed ad libitum 2920X Teklad rodent diet (Envigo). Mice were exposed via their drinking water to 0 (no treatment), 20 (0.3 μM), 100 (1.3 μM), and 500 ppb (6.7 μM) or to 0, 500 ppb, and 1000 ppb (13.3 μM) AsIII for 30 days (n = 5 mice/group) or 2 weeks (n = 6 mice/group), respectively.
Primary mouse bone marrow cell isolation. Bone marrow cells were isolated from femur bones as described by Ezeh et al. 73 51 . HPC were cultured in SF StemSpan hematopoietic progenitor expansion media (STEMCELL Technologies) containing AsIII and supplemented with 100 ng/mL murine stem cell factor (SCF) and 5 IU/mL (31.25 ng/mL) human recombinant EPO (Peprotech) to stimulate erythroid lineage commitment and differentiation. Erythroid progenitor cells were continuously exposed to AsIII throughout differentiation.
Flow cytometry. Erythroid and myeloid progenitors were evaluated based on surface marker phenotype as defined by Pronk et al. 52 and Grover et al. 53   For surface marker analysis, 0.5-1 × 10 6 cells were stained with 0.5 µg of monoclonal antibodies (all from BD Biosciences): cKit-APC-R700, CD34-PE-Cy7, SCA-1-BV605, CD16/32-BV510, CD150-BV421, CD105-BB515, CD71-PE and Ter119-FITC in 100 µL BD Horizon Brilliant Stain Buffer at room temperature in the dark for 30 min. Samples were washed twice and resuspended in 0.5 mL DPBS − with 2% FBS and 0.09% sodium azide prior to analysis using a BD Accuri C6 or BD LSRFortessa flow cytometer. Gates were set with the aid of fluorescence-minus-one controls. Flow cytometry data are reported as percentages or absolute cell numbers. Absolute cell numbers were generated by multiplying population subset percentages by total viable cell counts (for cell counts, see Supplementary Tables S1, S2).
Immunoprecipitation and co-immunoprecipitation. GATA-1 or PU.1 protein was isolated by immunoprecipitation and GATA-1/FOG-1 complex was co-immunoprecipitated as previously described 45 . Briefly, GATA-1, FOG-1, or PU.1 were purified using GATA-1 (D52H6) XP rabbit monoclonal antibody, PU.1 (9G7) rabbit monoclonal antibody (Cell Signaling Technologies), or FOG-1 rabbit polyclonal antibody (ab86281; Abcam), respectively using Dynabeads Protein-A Immunoprecipitation Kit (ThermoFisher Scientific). Immunoprecipitated GATA-1, FOG-1, and PU.1 were eluted from the beads and utilized for downstream experiments. GATA-1 and FOG-1 interaction was determined in co-immunoprecipitated GATA-1/FOG-1 complex by western blotting. Densitometry summaries were obtained from 3 independent blots after normalization to the density level of the whole image. To prepare samples for ICP-MS, a non-denaturing method was used to elute GATA-1 or PU.1 from beads, and the solution adjusted to pH > 7 using the neutralizing buffer provided in the immunoprecipitation kit.
GATA-1 DNA binding activity. GATA-1 DNA binding activity was determined using the fluorometric EpiQuik General Protein-DNA Binding Assay Kit (EpiGentek) according to the manufacturer's instructions. GATA-1 DNA binding activity was determined with immunoprecipitated GATA-1 by assessing the ability of GATA to bind a GATA consensus double-stranded DNA probe sequence 74 (GCC CCC GCT GAT TCC CTT ATC TAT GCC TTC CCAGC) or negative control double-stranded DNA probe sequence (ACA GGG ATG GGG GAG GGA ATG GGG TGA GGC CTGTC) using the EpiQuik General Protein-DNA Binding Assay Kit (Epigentek). All procedures strictly followed the instruction except 1:500 dilution of GATA-1 antibody was used. Nuclear extracts were prepared using the EpiQuik Nuclear Extraction Kit according to manufacturer's instructions (Epigentek). Fluorescence intensity was measured with a plate reader at Ex. 495 nm and Em. 520 nm. Human GATA-1 Transcription Factor Activity Assay (RayBiotech) was also used. Following erythroid differentiation, nuclear proteins were extracted from each treatment group (1.2 × 10 7 cells/group) using the Nuclear Extraction Kit according to instructions from the manufacturer (RayBiotech). Isolated nuclear proteins were then loaded into the wells of 96-well plate coated with double stranded oligonucleotides containing GATA-1 binding sequences. The Raybiotech Kit provides specific and non-specific competitor DNA oligos. Positive control (cell extract) were used with each of the competitor DNA oligos as control in the DNA binding assay. GATA-1 binding activity was determined using a primary antibody against GATA-1. Following primary antibody incubation, HRP-conjugated secondary antibody was added, and colorimetric signal was measured with a spectrophotometric plate reader at 450 nm.
ChIP assay and qPCR. GATA-1 DNA binding activity was determined using the ChIP-IT High Sensitivity and ChIP-IT Control qPCR Kits (Active Motif) according to manufacturer's instruction with some modifications. Briefly, ~ 2-3 × 10 6 mouse bone marrow erythroid progenitor cells were used for the ChIP assay. Crosslinking and cell lysis were performed in accordance with the ChIP-IT High Sensitivity Kit manual. Chromatin was sheared on ice using a micro ultrasonic cell disrupter (Kontes) to approximately 200-1000 bp. Chromatin fragment size was confirmed by analyzing size distribution in a 1.5% agarose gel. All immunoprecipitation, decrosslinking, and DNA purification steps were performed as described by the manufacturer. For immunoprecipitation, 7 µg of sheared chromatin was combined with ChIP grade GATA-1 antibody (4 µg; ab11852; Abcam), normal mouse IgG (2 µg; Active Motif) or RNA Polymerase II + bridging antibody (2 µg each; Active Motif). All assays exceeded the quality control criteria established by Active Motif for the ChIP-IT High Sensitivity Kit.
Following ChIP procedure, purified DNA was utilized to analyze the enrichment of GATA-1 binding by qPCR using the ChIP-IT qPCR Analysis Kit (Active Motif). Primers were designed to regions of the Klf-1 and Nfe2 genes as described by Suzuki et al. 72 The sequences used were cross referenced to available ChIP-sequencing data from the University of California Santa Cruz Genomics Institute Encyclopedia of DNA Elements to verify that they were valid genomic regions for evaluation of GATA-1 DNA binding activity (Supplementary Figs. S2, S3; https ://genom e.ucsc.edu/ENCOD E/index .html).
Primer sequences are listed in Supplementary Table S3. Primers efficiencies were tested and found to be ~ 100% for both Klf-1 and Nfe2 primer sets. In addition, universal negative control primers (Gapdh2) provided in the ChIP-IT qPCR Analysis Kit (Active Motif) were utilized for data analysis. Likewise, all other qPCR conditions were optimized. qPCR was performed with SsoAdvanced Universal SYBR Green Supermix (BioRad) according to manufacturer's instructions using a using a BioRad CFX384 Touch Real-Time PCR Detection System (BioRad). In an effort to minimize inter-sample variability, results from two independent experiments were expressed as fold enrichment over negative control primers, converted to percentage of untreated control, and merged for subsequent data analysis.

Scientific Reports
| (2020) 10:19055 | https://doi.org/10.1038/s41598-020-76118-x www.nature.com/scientificreports/ Inductively coupled plasma-mass spectrometry. Immunoprecipitated GATA-1 or PU.1 protein was collected, and protein content determined using the Pierce BCA Protein Assay Kit (ThermoFisher Scientific). Protein samples were diluted in trace metal grade nitric acid and zinc and arsenic contents in GATA-1 or PU.1 were measured by ICP-MS. Blank samples, internal standards, and standard curves were included with experimental samples as quality control for preparation and analyses. Data were calculated from the standard curve values generated during each experiment and values generated from negative quality control samples were subtracted as background prior to data analysis. Results were normalized to immunoprecipitated protein content.
Statistics. Flow cytometry data was processed using FlowJo version 10 (FlowJo LLC). Data were analyzed with GraphPad Prism 7. Differences between no treatment and treatment groups were determined using oneway ANOVA, followed by Tukey's post hoc test or two-tailed Student's t-test at a significance level of p < 0.05 or lower, as indicated in figure legends. All results from K562 cells are biological replicates and are representative of one of three independent experiments. Results from primary bone marrow HPC include at least three technical replicates and are representative of one of three independent experiments.

Data availability
The datasets generated/analyzed during the current study are available from the corresponding author (Ke Jian Liu: kliu@salud.unm.edu) on reasonable request.