Di(2-ethylhexyl)phthalate Impairs Erythropoiesis via Inducing Klotho Expression and Not via Bioenergetic Reprogramming

doi:10.21203/rs.3.rs-95024/v1

Download PDF

Research

Di(2-ethylhexyl)phthalate Impairs Erythropoiesis via Inducing Klotho Expression and Not via Bioenergetic Reprogramming

https://doi.org/10.21203/rs.3.rs-95024/v1

This work is licensed under a CC BY 4.0 License

Version 1

posted

You are reading this latest preprint version

Background: Di(2-ethylhexyl)phthalate (DEHP) is one of the phthalates most widely used to manufacture various plastic products. However, the potential effects of DEHP on erythropoiesis have not been investigated comprehensively. Here, we aimed to investigate whether DEHP modulate the function of haematopoietic stem and progenitor cells (HSPCs) to influence erythropoiesis, and to explore the associated mechanisms.

Results: In the present study, human cell lines with a capacity to differentiate into erythroid cells and murine bone marrow cells were treated with DEHP. DEHP not only impaired HSPC function but also suppressed erythroid differentiation in a dose-dependent manner. In addition, DEHP removal restored HSPC activity. To explore how DEHP interferes with erythroid differentiation, we focused on energy metabolism and Klotho expression. DEHP suppressed erythroid differentiation via upregulation of Klotho expression in erythroblasts and not via cellular bioenergetic modulation.

Conclusions: Here, we have demonstrated a novel pathophysiological role of DEHP in erythroid differentiation. DEHP impaired erythroid differentiation by inducing Klotho expression and not by reprogramming cellular bioenergetic profiles during differentiation.

General Cell Biology & Physiology

Di(2-ethylhexyl)phthalate

phthalates

erythroid differentiation

Klotho

bioenergetics

Di(2-ethylhexyl)phthalate (DEHP) is one of the phthalates most frequently used as a plasticiser in plastic products such as personal care products and medical devices. It is non-covalently bound to the polymer; hence, DEHP easily leaches, migrates, or evaporates into the atmosphere, foods, or directly into body fluids [1−3]. Due to its ubiquity in the environment, concerns have been raised about the potential health impacts of continuous exposure to DEHP exposure. To date, the adverse effects of DEHP in the liver, the endocrine and reproductive systems, as well as in obesity and some types of cancers have been explored extensively [3, 4]. However, further investigations are required to determine if DEHP exposure influences other physiological processes. According to a recent study, DEHP affects the function of human haematopoietic stem and progenitor cells (HSPCs) isolated from human umbilical cord blood [5]. DEHP also impairs the migration and colony-forming capacity of human CD34⁺ HSPCs [6]. High-dosage and long-term dietary DEHP uptake in rodents caused chronic progressive nephropathy and significantly reduced haemoglobin (Hgb) production [7, 8]. However, DEHP had no effect in male cynomolgus monkeys treated with 500 mg/kg/day DEHP for 14 days [9]. It remains unclear whether DEHP exposure adversely affects erythropoiesis until now.

Previous studies have demonstrated that DEHP acts as an endocrine disruptor and reproductive toxicant via the inhibition of endocrine receptor signalling or the activation of peroxisome proliferator-activated receptors (PPARs) [3, 10]. In addition, DEHP causes epigenetic changes, especially DNA methylation [3, 11] and impairs both insulin signalling and glucose metabolism [11−14]. In liver and muscle cells, DEHP exposure can influence the levels of glucose metabolites, including glucose-6-phosphate, fructose-6-phosphate, pyruvate, lactate, glucose-1-phosphate, and glycogen [13]. In haematopoietic systems, long-term haematopoietic stem cells (HSC), which are at the top of the haematopoietic hierarchy, tend to generate energy via anaerobic glycolysis, whereas lineage-restricted progenitor cells generate adenosine triphosphate (ATP) primarily in the mitochondria via oxidative phosphorylation (OXPHOS) [15−18]. Erythroid progenitor cells also exhibit a similar glycolytic metabolism rearrangement when they differentiate into mature erythroid cells [19]. In the present study, we investigated whether DEHP influences erythroid differentiation through the modulation of glycolytic metabolism and mitochondrial respiration. In addition, we investigated potential DEHP-mediated molecular mechanisms.

DEHP suppresses erythropoiesis

To rapidly test whether DEHP influenced erythropoiesis, we used human chronic myelogenous leukaemia K562 and erythroleukemia HEL 92.1.7 (HEL) cell lines. The erythroid and megakaryocytic differentiation potentials of the two cell lines were comparable to those of human CD34⁺ HSPCs [20, 21]. Sodium butyrate (NaB) was used to induce erythroid differentiation in both cell lines [21]. The addition of DEHP in the culture medium did not affect K562 cell proliferation (Fig. 1a) and survival (Fig. 1b). Based on previous reports on DEHP plasma concentrations in chronic kidney disease (CKD) patients undergoing haemodialysis [22, 23], 1 µg/ml of DEHP was selected for application in the experiments in the present study.

DEHP treatment substantially decreased NaB-induced erythroid differentiation, which was evaluated based on Hgb production, in both K562 and HEL cells. Without NaB treatment, DEHP also decreased spontaneous erythroid differentiation, which was resulted from increasing cell numbers, in both cell lines (Fig. 1c). Erythropoietin (EPO) is the primary cytokine that regulates erythropoiesis [24, 25]. An increase in DEHP concentration gradually and significantly suppressed the formation of EPO-induced colony-forming unit-erythroids (CFU-Es) in bone marrow (BM) (Fig. 1d) and spleen (SP)HSPC cells (Fig. 1e). CFU-Es are differentiated form the earlier erythroid progenitor cells, burst-forming unit-erythroids (BFU-Es) [26]. DEHP also suppressed BFU-E formation (Fig. 1f). The results not only demonstrated that DEHP suppressed erythropoiesis but also indicated that the two human cell lines were comparable cell models that could be adopted in subsequent investigations.

DEHP suppresses the differentiation of HSPCs, whereas removal of DEHP restores HSPC self-renewal activity

To examine the effect of DEHP in the proliferation and differentiation of HSPCs, BM and SP cells were cultured in a methylcellulose-based medium. Consequently, HSPCs proliferate and differentiate to form discrete colonies, including the oligopotential progenitor CFU-granulocyte, erythrocyte, monocyte/macrophage, and megakaryocyte (CFU-GEMM), lineage-restricted progenitor CFU-granulocyte and monocyte/macrophage (CFU-GM) and the precursors of granulocytes (CFU-G) and monocytes/macrophages (CFU-M). DEHP decreased the formation of CFU-GEMM, CFU-GM, CFU-G and CFU-M in both BM and SP cells in a dose-dependent manner (Fig. 2). The results clearly indicated that DEHP influenced HSPC differentiation.

HSPC self-renewal can be examined by re-plating cells in methylcellulose-based medium. HSPCs retained a similar capacity to generate haematopoietic cells in the second plating, while the potential decreased gradually in the third and fourth iterations (Fig. 3a), which is consistent with our previous observations [27]. The experimental design used to examine whether DEHP removal restored HSPC potential is presented in Fig. 3b. In the first plating, DEHP inhibited total colony number growth significantly. Subsequently, a cell number similar to the number of the first-generation colony cells (2 × 10⁴ cells) was re-plated with or without DEHPs (1 µg/ml). Like normal HSPCs in BM cells retained similar ability to generate haematopoietic cells in the first and second plating, removal of DEHP in the second plating keep similar self-renewal capability. However, continuous exposure to DEHP significantly decreased HSPC function (Fig. 3c). Overall, the results suggested that DEHP interfered transiently with HSPC functions.

DEHP did not reprogram cellular bioenergetics during erythroid differentiation

Previous studies have indicated that DEHP treatment influences cellular glucose metabolism [13], and erythropoiesis is highly associated with reprogramming of glucose metabolism and energy consumption [19]. Mammalian cells generate ATP by mitochondrial (oxidative phosphorylation) and non-mitochondrial (glycolysis) metabolism. Cellular oxygen consumption rate (OCR) and extracellular acidification rate (ECAR) are key indicators of mitochondrial respiration and glycolysis, respectively. The indices could provide a systemic perspective of cellular metabolic function in live cells and can be evaluated using a Seahorse Extracellular Flux Analyzer [28, 29]. Here, we firstly examined whether DEHP influenced energy metabolism in undifferentiated K562 cells. K562 cells were treated with 1 µg/ml DEHP for 24 h, and then collected for OCR and ECAR assays. OCR is measured before and after the addition of various inhibitors to derive several mitochondrial respiration parameters. The cellular OCR baseline is measured before the addition of oligomycin, a complex V inhibitor. The ATP-linked respiration can be evaluated following treatment with oligomycin. Subsequently, FCCP, a protonophore, is added to collapse the inner membrane gradient, allowing the electron transport chain (ETC) to function at its maximum potential, and then maximal respiratory capacity is measured. Finally, antimycin A and rotenone, inhibitors of complex III and I, are added to shut down ETC function and reveal the non-mitochondrial respiration. Mitochondrial spare respiratory capacity is calculated by subtracting basal respiration from maximal respiratory capacity (Fig. 4a). DEHP treatment did not influence mitochondrial respiration parameters in undifferentiated K562 cells (Fig. 4b, c).

For the ECAR assay, cells were cultured in the assay buffer without glucose for 1 h. Glucose is used to elicit glycolytic activity. Afterward, oligomycin injection blocks mitochondrial respiration to allow glycolysis to function at its maximum rate. Finally, the addition of 2-DG, a glucose analogue, inhibits glycolysis, and, therefore, provides a baseline ECAR measurement. Glycolytic reserve capacity is calculated by subtracting basal ECAR from maximal ECAR (Fig. 4d). Similar to the OCR analysis case, DEHP treatment did not influence any glycolysis parameter in undifferentiated K562 cells (Fig. 4e, f). Subsequently, we analysed whether DEHP reprogramed cellular bioenergetics in the course of erythroid differentiation.

K562 cells were treated with NaB with or without DEHP (1 µg/ml) for 24 h, and then subjected to OCR and ECAR assays. Induction of erythroid differentiation increased basal and maximal mitochondrial respiration dramatically, in addition to ATP production (Fig. 5a, b). In addition, erythroid differentiation induction decreased glycolytic activity significantly (Fig. 5c, d). The results were consistent with previous findings on the rearrangement of glucose metabolism and energy consumption during erythropoiesis in chicken T2EC cells [19]. However, the addition of DEHP into differentiated cells did not influence cellular bioenergetics further (Fig. 5). The results indicated that the effects of DEHP on erythroid differentiation are not via modulating cellular bioenergetics.

DEHP induces Klotho expression, which affects erythroid differentiation

Previous studies have demonstrated that loss of Klotho, which was initially considered an anti-ageing gene, disrupted HSPC homing, and increased erythropoiesis [30]. Therefore, Klotho expression was analysed in K562 cells under different treatments. Compared to the wild-type (WT) K562 cells (Control, C), the expression of Klotho was decreased in NaB-treated K562 cells. DEHP addition in NaB-treated K562 cells induced Klotho expression markedly (Fig. 6a). The association between DEHP and Klotho expression was further examined in CFU-E cells. DEHP exposure induced Klotho expression in CFU-E cells significantly (Fig. 6b). To confirm the influence of Klotho in DEHP-mediated suppression of erythropoiesis, the knockdown approach was used. Three Klotho short-hair (shRNA) were expressed transiently in K562 cells for 3 days, and then endogenous Klotho protein expression was analysed using Western blot analysis. The C2 and E2 shRNAs exhibited better knockdown efficiency (Fig. 6c). Afterward, C2 and E2 shRNA-expressed K562 cells were treated with NaB with or without DEHP. Knockdown of Klotho increased erythroid differentiation significantly, which was consistent with the findings of a previous report [30]. Notably, Klotho knockdown abolished DEHP-mediated suppression of erythroid differentiation (Fig. 6d). Overall, the results indicated that DEHP downregulated erythroid differentiation via Klotho expression induction.

In the present study, our results demonstrated an unprecedented link between DEHP and Klotho in erythroid differentiation. DEHP seemed to transiently impair HSPC function and erythroid differentiation. We further revealed that DEHP suppressed erythroid differentiation through the upregulation of Klotho expression in haematopoietic cells and not via modulation of cellular bioenergetics (Fig. 7). Although our results indicated that DEHP does not interfere with cellular bioenergetics in haematopoietic cells, numerous studies have demonstrated that DEHP influences mitochondrial function, energy metabolism, and insulin resistance in other cell types [31−35].

Klotho proteins, Klotho and β-Klotho, are required for high-affinity binding of fibroblast growth factors (FGFs) including FGF19, FGF21, and FGF23 to their cognate FGF receptors (FGFR) [36]. In the present study, we focused on Klotho, because β-Klotho is expressed abundantly in metabolic tissues [37]. Klotho^−/− mice developed multiple phenotypes observed commonly in naturally aged WT mice in an accelerated manner, such as decreased spontaneous activity, hypogonadism, impaired bone mineralisation, thymic involution, B lymphocytopenia [38]. In contrast, β-Klotho loss protected against obesity by increasing energy expenditure, which is due to crosstalk among liver, microbiota, and brown adipose tissue [39]. Loss of Klotho not only interfered with HSC function but also impaired erythropoiesis [30]. Our results further demonstrated the critical role of Klotho in DEHP-mediated suppression of erythropoiesis.

In patients with CKD, increased DEHP concentrations were observed when they underwent haemodialysis [22, 23, 40]. Anaemia is mainly due to impaired EPO production in the kidney [41], and is highly associated with morbidity and mortality in CKD patients [41]. EPO is the primary cytokine regulating erythropoiesis; therefore, anaemia correction using recombinant human EPO or erythropoiesis-stimulating agents (ESAs) markedly improves the quality of life of CKD patients [42, 43]. Nevertheless, other factors could interfere with responses to EPO or ESAs [43, 44]. Unfortunately, some CKD patients still exhibit poor responses to EPO or ESAs, even when the therapies are combined with other therapeutic strategies to minimise inflammation, improve vitamin D status, and decrease hyperparathyroidism [44]. Conversely, increasing evidence indicates that Klotho, which is mainly expressed in the kidney, is correlated significantly with the development and progression of CKD and its complications [36, 45]. Here, our results suggest that DEHP is an uncharacterised factor involved in the progression of anaemia, or in the development of peripheral resistance or decreased response to EPO or ESAs in CKD patients. However, further studies are required to validate the findings above.

DEHP, the most common phthalate, leaches into the environment or directly into human tissue easily. Previous studies have demonstrated that DEHP interferes with some HSPC functions. In the present study, we demonstrated that exposure to DEHP transiently impairs HSPC function and erythroid differentiation. We also demonstrated that DEHP suppresses erythroid differentiation through the upregulation of Klotho expression in haematopoietic cells and not via modulation of cellular bioenergetics. Consequently, the present study provides a novel link between DEHP and Klotho in erythroid differentiation. In addition, our results could facilitate the assessment of novel DEHP-mediated effects in the environment and in human tissues.

Materials

DEHP (Acros Organics) was a kind gift from Dr. Tzong-Shyuan Lee (National Taiwan University, Taiwan), and was prepared using dimethylsulfoxide (DMSO) as the solvent at a concentration of 1 or 10 mg/ml [46, 47]. The reference sequence for Klotho shRNA design is NM 004795. The scramble shRNA (ASN0000000004) and Klotho shRNAs, including A2 (TRCN0000161418), D2 (TRCN0000419459), and E2 (TRCN0000433013), were obtained from National RNAi Core Facility (Academia Sinica, Taipei City, Taiwan). All plasmids were verified by DNA sequencing, which was carried out by Genomics (New Taipei City, Taiwan).

Cell culture, differentiation and transfection

Human K562 and HEL cells were obtained from the Bioresource Collection and Research Center (BCRC), Hsinchu City, Taiwan. K562 cells (4 × 10⁴ cells/ml) were cultivated in Iscove’s Modified Dulbecco’s Medium (IMDM) supplemented with 10% foetal bovine serum (FBS), 100 IU/ml penicillin, and 100 IU/ml streptomycin, and were passaged every two days. HEL cells (1 × 10⁵ cells/ml) were cultivated in Roswell Park Memorial Institute (RPMI) 1640 medium supplemented with 10% FBS, 1 mM sodium pyruvate, 100 IU/ml penicillin, and 100 IU/ml streptomycin, and were passaged every 2 to 3 days. For erythroid differentiation, K562 and HEL cells were treated with 2 mM NaB for K562 cells and at 0.1 mM NaB for HEL cells. Benzidine staining was performed to examine Hgb expression [21]. Transfection was performed using Lipofectamine™ 2000 reagent (Thermo Fisher Scientific Inc.) according to the manufacturer’s instructions. The K562 cells, which were used to express Klotho shRNAs, were cultured with 0.5 µg/ml puromycin for at least 12 days.

Mice and cell isolations

Pure C57BL/6 mice were purchased from the Laboratory Animal Center of National Yang-Ming University. BM cells were isolated carefully from tibia and femur. For SP cell preparation, a small piece of excised spleen (SP) was pressed through the strainer using the plunger end of a syringe. Subsequently, BM and SP cells were washed through the strainer and collected [27, 48].

Colony forming assay

The methylcellulose-based media MethoCult™ M3334 and M3434 were obtained from StemCell Technologies (Vancouver, Canada). Colony forming assays were performed as previously described [27, 49]. To detect BFU-E and other types of colonies (CFU-G, CFU-M, CFU-GM, and CFU-GEMM), 2 × 10⁴ BM cells or 1 × 10⁵ SP cells were plated in duplicate in MethoCult™ GF M3434 methylcellulose medium according to the manufacturer’s protocol. Colonies were counted after 10 days in culture. For re-plating analyses, 2 ⋅ 10⁴ BM cells were plated in duplicate in MethoCult™ GF M3434 methylcellulose medium. After 10 days, 2 × 10⁴ colony cells were re-plated. To detect CFU-E colonies, 2 × 10⁴ BM cells or 1 × 10⁵ SP cells were plated in duplicate in MethoCult™ M3334 methylcellulose medium according to the manufacturer’s protocol. CFUE colonies were counted after 2 days of culture.

Analysis of OCR and ECAR

OCR and ECAR were measured using a Seahorse XF24 Extracellular Flux Analyzer (Seahorse Bioscience, North Billerica, MA). Firstly, the Seahorse XF cell culture microplates were coated with 3.5 µg/cm² of Corning Cell-Tak™ adhesive reagent (Corning, NY, USA). For OCR, the constituents were prepared in sodium bicarbonate- and HEPES-free IMDM (Gibco™, Thermo Fisher Scientific) with 2% FBS (Gibco™, Thermo Fisher Scientific), 100 IU/ml penicillin (Sigma-Aldrich), and 100 IU/ml streptomycin (Sigma-Aldrich) (non-buffered IMDM). Ninety-thousand K562 cells were resuspended in 100 µl non-buffered IMDM for one well of the Seahorse XF cell culture microplates, transferred into microplates, and then incubated at room temperature for 30 min. Subsequently, 575 µl of non-buffered IMDM was added in each well. Oligomycin, FCCP, rotenone, antimycin, glucose, and 2-DG were obtained from Seahorse Bioscience (North Billerica, MA, US). To measure OCR, oligomycin and FCCP were diluted to 1 µM, and otenone and antimycin were diluted to 0.5 µM. For ECAR, culture medium was changed to assay medium (XF base medium DMEM supplemented with 2 mM glutamine), and cells were incubated in a non-CO₂ incubator at 37 °C for 1 h before ECAR assay. Glucose, oligomycin and 2-DG were diluted to 10 mM, 1 µM and 50 mM, respectively. The automatic measurement of OCR and ECAR in real time was performed according to the manufacturer’s protocol to detect live-cell bioenergetic profile.

Statistical analysis

Statistical analyses were performed with GraphPad Prism 6.0 (GraphPad Software Inc., San Diego, CA). Data are presented as means ± s.d. All experiments were performed at least three times. We compared the result of the treatment groups and the controls using Student’s t-test or two-way analysis of variance (ANOVA) followed by the Tukey's multiple-comparison post-hoc test. Differences between groups were considered significant at P < 0.05.

2-DG: 2-deoxyglucose; ATP: adenosine triphosphate; ANOVA: Analysis of variance; BFU-E: Burst-forming unit erythroid; BM: Bone marrow; CFU-E: Colony forming unit erythroid; CFU-GEMM: Colony forming unit-granulocyte, erythrocyte, monocyte/macrophage, and megakaryocyte; CFU-GM: Colony forming unit-granulocyte and monocyte/macrophage; CFU-G: Colony forming unit-granulocyte; CFU-M: Colony forming unit-monocyte/macrophage; CKD: Chronic kidney disease; ; DEHP: Di(2-ethylhexyl)phthalate; DMSO: Dimethylsulfoxide; ECAR: Extracellular acidification rate; EPO: Erythropoietin; ESA: Erythropoiesis-stimulating agent; ETC: electron transport chain; FBS: Foetal bovine serum; FCCP: Carbonyl cyanide-4 (trifluoromethoxy) phenylhydrazone; FGF: Fibroblast growth factor; Hgb: Haemoglobin; HSC: Haematopoietic stem cells; HSPC: Haematopoietic stem and progenitor cell; IACUC: Institutional Animal Care and Use Committee; IMDM: Iscove’s Modified Dulbecco’s Medium; NaB: Sodium butyrate; OCR: Oxygen consumption rate; OXPHOS: Oxidative phosphorylation; PPAR: Peroxisome proliferator-activated receptor; Short-hair RNA: shRNA; SP: Splenocytes; SCF: Stem cell factor; WT: Wild-type.

Ethics approval and consent to participate

All animal experiments were conducted in accordance with Guide for the Care and Use of Laboratory Animals and approved by the Institutional Animal Care and Use Committee (IACUC) of National Yang-Ming University (IACUC number: 1081012). No human data were used for this study, and thus consent was not required.

Consent for publication

No human data were used for this study, and thus consent was not required.

Availability of data and materials

The datasets used and/or analysed during the current study are available from the corresponding author on reasonable request.

Competing interests

The authors declare that they have no competing interests.

Funding

This research was supported by funding from Ministry of Science and Technology (MOST 105-2628-B-010-010-MY3 and MOST 108-2320-B-010 -043 -MY3) (Y.I.C.), a grant from Yen Tjing Ling Medical Foundation (CI-109-6). (Y.I.C.) This work is particularly supported by "Yin Yen-Liang Foundation Development and Construction Plan" of the School of Medicine, National Yang-Ming University (Y.I.C.).

Author’s contributions

CYT performed the experiments about DEHP in survival and differentiation in cell lines and identifying molecular mechanism, data analysis, and manuscript writing. TPF performed the analysis of OCR and ECAR and the identification of molecular mechanism, data analysis, and manuscript writing. YIC performed colony forming assay. SWC, HWC, ECYL and TAL participated in parts of experiments and supervised the animal harvest and care. TAL, DCT and YIC conceived and designed the study. YIC supervised the project, edited the manuscript and gave approval to its final version. All authors read and approved the final manuscript.

Acknowledgements

We are grateful to Dr. Tzong-Shyuan Lee (National Taiwan University, Taiwan) for providing DEHP. We thank the Instrumentation Research Center of National Yang-Ming University (Taiwan) for use of the shared services to complete this research. We also thank the National RNAi Core Facility at Academia Sinica in Taiwan for providing shRNA reagents and related services.

Author details

¹Department and Institute of Physiology, National Yang-Ming University, No. 155, Sec. 2, Linong St., Taipei City 112304, Taiwan. ²Division of Hematology and Oncology, Department of Medicine, Taipei Veterans General Hospital, No. 201, Sec. 2, Shipai Rd., Taipei 112201, Taiwan. ³Division of Nephrology, Department of Medicine, Taipei Veterans General Hospital, No. 201, Sec. 2, Shipai Rd., Taipei 112201, Taiwan. ⁴Institute of Clinical Medicine, National Yang-Ming University, No. 155, Sec. 2, Linong St., Taipei 112304, Taiwan. ⁵Center for Intelligent Drug Systems and Smart Bio-devices (IDS2B), No. 1001, University Rd., Hsinchu City 300093, Taiwan.

Serrano SE, Braun J, Trasande L, Dills R, Sathyanarayana S. Phthalates and diet: a review of the food monitoring and epidemiology data. Environ Health. 2014;13:43.
Buckley JP, Kim H, Wong E, Rebholz CM. Ultra-processed food consumption and exposure to phthalates and bisphenols in the US National Health and Nutrition Examination Survey, 2013-2014. Environ Int. 2019;131:105057.
Yang O, Kim HL, Weon JI, Seo YR. Endocrine-disrupting chemicals: Review of toxicological mechanisms using molecular pathway analysis. J Cancer Prev. 2015;20:12-24.
Rowdhwal SSS, Chen J. Toxic effects of di-2-ethylhexyl phthalate: An overview. Biomed Res Int. 2018;2018:1750368.
Gutierrez-Garcia AK, Flores-Kelly JM, Ortiz-Rodriguez T, Kalixto-Sanchez MA, De Leon-Rodriguez A. Phthalates affect the in vitro expansion of human hematopoietic stem cell. Cytotechnology. 2019;71:553-561.
Manz P, Cadeddu RP, Wilk M, Fischer JC, Fritz B, Haas R, et al. Influence of Di(2-ethylhexyl)phthalate on migration rate and differentiation of human hematopoietic stem and progenitor cells (CD34⁺). Clin Hemorheol Microcirc. 2015;61:111-118.
David RM, Moore MR, Finney DC, Guest D. Chronic toxicity of di(2-ethylhexyl)phthalate in rats. Toxicol Sci. 2000;55:433-443.
David RM, Moore MR, Finney DC, Guest D. Chronic toxicity of di(2-ethylhexyl)phthalate in mice. Toxicol Sci. 2000;58:377-385.
Pugh G, Jr., Isenberg JS, Kamendulis LM, Ackley DC, Clare LJ, Brown R, et al. Effects of di-isononyl phthalate, di-2-ethylhexyl phthalate, and clofibrate in cynomolgus monkeys. Toxicol Sci. 2000;56:181-188.
Desvergne B, Feige JN, Casals-Casas C. PPAR-mediated activity of phthalates: A link to the obesity epidemic? Mol Cell Endocrinol. 2009;304:43-48.
Rajesh P, Balasubramanian K. Phthalate exposure in utero causes epigenetic changes and impairs insulin signalling. J Endocrinol. 2014;223:47-66.
Mushtaq M, Srivastava SP, Seth PK. Effect of di-2-ethylhexyl phthalate (DEHP) on glycogen metabolism in rat liver. Toxicology. 1980;16:153-161.
Martinelli MI, Mocchiutti NO, Bernal CA. Dietary di(2-ethylhexyl)phthalate-impaired glucose metabolism in experimental animals. Hum Exp Toxicol. 2006;25:531-538.
Xu J, Zhou L, Wang S, Zhu J, Liu T, Jia Y, et al. Di-(2-ethylhexyl)-phthalate induces glucose metabolic disorder in adolescent rats. Environ Sci Pollut Res Int. 2018;25:3596-3607.
Vannini N, Girotra M, Naveiras O, Nikitin G, Campos V, Giger S, et al. Specification of haematopoietic stem cell fate via modulation of mitochondrial activity. Nat Commun. 2016;7:13125.
Suda T, Takubo K, Semenza GL. Metabolic regulation of hematopoietic stem cells in the hypoxic niche. Cell Stem Cell. 2011;9:298-310.
Takubo K, Nagamatsu G, Kobayashi CI, Nakamura-Ishizu A, Kobayashi H, Ikeda E, et al. Regulation of glycolysis by Pdk functions as a metabolic checkpoint for cell cycle quiescence in hematopoietic stem cells. Cell Stem Cell. 2013;12:49-61.
Papa L, Djedaini M, Hoffman R. Mitochondrial role in stemness and differentiation of hematopoietic stem cells. Stem Cells Int. 2019;2019:4067162.
Richard A, Vallin E, Romestaing C, Roussel D, Gandrillon O, Gonin-Giraud S. Erythroid differentiation displays a peak of energy consumption concomitant with glycolytic metabolism rearrangements. PloS One. 2019;14:e0221472.
Chang YI, Hua WK, Yao CL, Hwang SM, Hung YC, Kuan CJ, et al. Protein-arginine methyltransferase 1 suppresses megakaryocytic differentiation via modulation of the p38 MAPK pathway in K562 cells. J Biol Chem. 2010;285:20595-20606.
Hua WK, Chang YI, Yao CL, Hwang SM, Chang CY, Lin WJ. Protein arginine methyltransferase 1 interacts with and activates p38alpha to facilitate erythroid differentiation. PloS One. 2013;8:e56715.
Mettang T, Thomas S, Kiefer T, Fischer FP, Kuhlmann U, Wodarz R, et al. Uraemic pruritus and exposure to di(2-ethylhexyl) phthalate (DEHP) in haemodialysis patients. Nephrol Dial Transplant. 1996;11:2439-2443.
Faouzi MA, Dine T, Gressier B, Kambia K, Luyckx M, Pagniez D, et al. Exposure of hemodialysis patients to di-2-ethylhexyl phthalate. Int J Pharm. 1999;180:113-121.
Grover A, Mancini E, Moore S, Mead AJ, Atkinson D, Rasmussen KD, et al. Erythropoietin guides multipotent hematopoietic progenitor cells toward an erythroid fate. J Exp Med. 2014;211:181-188.
Wojchowski DM, Menon MP, Sathyanarayana P, Fang J, Karur V, Houde E, et al. Erythropoietin-dependent erythropoiesis: New insights and questions. Blood Cells Mol Dis. 2006;36:232-238.
Lodish H, Flygare J, Chou S. From stem cell to erythroblast: regulation of red cell production at multiple levels by multiple hormones. IUBMB life. 2010;62:492-496.
Chang YI, You X, Kong G, Ranheim EA, Wang J, Du J, et al. Loss of Dnmt3a and endogenous Kras cooperate to regulate hematopoietic stem and progenitor cell functions in leukemogenesis. Leukemia. 2015;29:1847-1856.
Vangapandu HV, Gandhi V. Extracellular flux assays to determine oxidative phosphorylation and glycolysis in chronic lymphocytic leukemia cells. Methods Mol Biol. 2019;1881:121-128.
van der Windt GJW, Chang CH, Pearce EL. Measuring bioenergetics in T cells using a Seahorse Extracellular Flux Analyzer. Curr Protoc Immunol. 2016;113:3.16B.1-3.16B.14.
Vadakke Madathil S, Coe LM, Casu C, Sitara D. Klotho deficiency disrupts hematopoietic stem cell development and erythropoiesis. Am J Pathol. 2014;184:827-841.
Chiang HC, Wang CH, Yeh SC, Lin YH, Kuo YT, Liao CW, et al. Comparative microarray analyses of mono(2-ethylhexyl)phthalate impacts on fat cell bioenergetics and adipokine network. Cell Biol Toxicol. 2017;33:511-526.
Inouye B, Ogino Y, Ishida T, Ogata M, Utsumi K. Effects of phthalate esters on mitochondrial oxidative phosphorylation in the rat. Toxicol Appl Pharmacol. 1978;43:189-198.
Kora S, Sado M, Terada H. Effect of the plasticizer di-(2-ethylhexyl)phthalate on oxidative phosphorylation in rat liver mitochondria: Modification of the function of the adenine nucleotide translocator. J Pharmacobiodyn. 1988;11:773-778.
Dales RE, Kauri LM, Cakmak S. The associations between phthalate exposure and insulin resistance, beta-cell function and blood glucose control in a population-based sample. Sci Total Environ. 2018;612:1287-1292.
Kataria A, Levine D, Wertenteil S, Vento S, Xue J, Rajendiran K, et al. Exposure to bisphenols and phthalates and association with oxidant stress, insulin resistance, and endothelial dysfunction in children. Pediatr Res. 2017;81:857-864.
Kuro OM. The Klotho proteins in health and disease. Nat Rev Nephrol. 2019;15:27-44.
Ito S, Kinoshita S, Shiraishi N, Nakagawa S, Sekine S, Fujimori T, et al. Molecular cloning and expression analyses of mouse betaklotho, which encodes a novel Klotho family protein. Mech Dev. 2000;98:115-119.
Kuro-o M. Klotho and aging. Biochim Biophys Acta. 2009;1790:1049-1058.
Somm E, Henry H, Bruce SJ, Aeby S, Rosikiewicz M, Sykiotis GP, et al. Beta-Klotho deficiency protects against obesity through a crosstalk between liver, microbiota, and brown adipose tissue. JCI insight. 2017;2:e91809.
Pollack GM, Buchanan JF, Slaughter RL, Kohli RK, Shen DD. Circulating concentrations of di(2-ethylhexyl) phthalate and its de-esterified phthalic acid products following plasticizer exposure in patients receiving hemodialysis. Toxicol Appl pharmacol. 1985;79:257-267.
Babitt JL, Lin HY. Mechanisms of anemia in CKD. J Am Soc Nephrol. 2012;23:1631-1634.
Hung SC, Tarng DC. ESA and iron therapy in chronic kidney disease: A balance between patient safety and hemoglobin target. Kidney Int. 2014;86:676-678.
de Oliveira Junior WV, Sabino Ade P, Figueiredo RC, Rios DR. Inflammation and poor response to treatment with erythropoietin in chronic kidney disease. J Bras Nefrol. 2015;37:255-263.
Icardi A, Paoletti E, De Nicola L, Mazzaferro S, Russo R, Cozzolino M. Renal anaemia and EPO hyporesponsiveness associated with vitamin D deficiency: The potential role of inflammation. Nephrol Dial Transplant. 2013;28:1672-1679.
Zou D, Wu W, He Y, Ma S, Gao J. The role of klotho in chronic kidney disease. BMC Nephrol. 2018;19:285.
Zhao JF, Hsiao SH, Hsu MH, Pao KC, Kou YR, Shyue SK, et al. Di-(2-ethylhexyl) phthalate accelerates atherosclerosis in apolipoprotein E-deficient mice. Arch Toxicol. 2016;90:181-190.
Lee KI, Chiang CW, Lin HC, Zhao JF, Li CT, Shyue SK, et al. Maternal exposure to di-(2-ethylhexyl) phthalate exposure deregulates blood pressure, adiposity, cholesterol metabolism and social interaction in mouse offspring. Arch Toxicol. 2016;90:1211-1224.
Chang YI, Kong G, Ranheim EA, Tu PS, Yu YS, Zhang J. Dnmt3a haploinsufficiency cooperates with oncogenic Kras to promote an early-onset T-cell acute lymphoblastic leukemia. Am J Transl Res. 2017;9:1326-1334.
Tu PS, Lin EC, Chen HW, Chen SW, Lin TA, Gau JP, Chang YI. The extracellular signal-regulated kinase 1/2 modulates the intracellular localization of DNA methyltransferase 3A to regulate erythrocytic differentiation. Am J Transl Res. 2020;12:1016-1030.

Download PDF

Version 1

posted

You are reading this latest preprint version

Di(2-ethylhexyl)phthalate Impairs Erythropoiesis via Inducing Klotho Expression and Not via Bioenergetic Reprogramming

Status:

Version 1

Abstract

Figures

Background

Results

DEHP suppresses erythropoiesis

DEHP suppresses the differentiation of HSPCs, whereas removal of DEHP restores HSPC self-renewal activity

DEHP did not reprogram cellular bioenergetics during erythroid differentiation

DEHP induces Klotho expression, which affects erythroid differentiation

Discussion

Conclusions

Methods

Materials

Cell culture, differentiation and transfection

Mice and cell isolations

Colony forming assay

Analysis of OCR and ECAR

Statistical analysis

Abbreviations

Declarations

Ethics approval and consent to participate

Consent for publication

Availability of data and materials

Competing interests

Funding

Author’s contributions

Acknowledgements

Author details

References

Status:

Version 1