Halobenzoquinone-Induced Developmental Toxicity, Oxidative Stress

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Halobenzoquinone-induced developmental toxicity, oxidative stress, and apoptosis in zebrafish embryos Chang Wang, Xue Yang, Qi Zheng, Birget Moe, and Xing-Fang Li Environ. Sci. Technol., Just Accepted Manuscript • DOI: 10.1021/acs.est.8b02831 • Publication Date (Web): 20 Aug 2018 Downloaded from http://pubs.acs.org on August 20, 2018

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Environmental Science & Technology

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Halobenzoquinone-induced developmental toxicity, oxidative stress, and apoptosis

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in zebrafish embryos

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Chang Wang,1# Xue Yang,1, 2# Qi Zheng,3* Birget Moe4,5 and Xing-Fang Li4*

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1. Institute of Environment and Health, Jianghan University, Wuhan 430056, China

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2. School of Environmental Ecology and Biological Engineering, Wuhan Institute of Technology,

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Wuhan 430025, China 3. Key Laboratory of Optoelectronic Chemical Materials and Devices, Ministry of Education, Institute

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of Environment and Health, Jianghan University, Wuhan 430056, China

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4. Division of Analytical and Environmental Toxicology, Department of Laboratory Medicine and

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Pathology, Faculty of Medicine and Dentistry, University of Alberta, Edmonton, Alberta, Canada

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T6G 2G3

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5. Alberta Centre for Toxicology, Department of Physiology and Pharmacology, Faculty of Medicine,

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University of Calgary, Calgary, Alberta, Canada T2N 4N1

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#

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* To whom correspondence should be addressed:

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Qi Zheng: [email protected].

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Xing-Fang Li: [email protected], 1-780-492-5094.

Co-first authors

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ACS Paragon Plus Environment


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Abstract

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The developmental toxicity of water disinfection byproducts remains unclear. Here we report the

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study of halobenzoquinone (HBQ)-induced in vivo developmental toxicity and oxidative stress using

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zebrafish embryos as a model. Embryos were exposed to 0.5 to 10 µM of individual HBQs and 0.5 to 5

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mM haloacetic acids for up to 120 hours post-fertilization (hpf). LC50 values of the HBQs at 24 hpf were

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4.6 to 9.8 µM, while those of three haloacetic acids were up to 200 times higher at 1900 to 2600 µM.

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HBQ exposure resulted in significant developmental malformations in larvae, including failed inflation

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of the gas bladder, heart malformations, and curved spines. An increase in reactive oxygen species was

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observed along with a decrease in superoxide dismutase activity and glutathione content. Additionally,

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the antioxidant N-acetyl-L-cysteine significantly mitigated all HBQ-induced effects, supporting that

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oxidative stress contributes to HBQ toxicity. Further experiments examined HBQ-induced effects on

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DNA and genes. HBQ exposure increased 8-hydroxydeoxyguanosine levels, DNA fragmentation, and

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apoptosis in larvae, with apoptosis induction related to changes in the gene expression of p53 and mdm2.

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These results suggest that HBQs are acutely toxic, causing oxidative damage and developmental toxicity

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to zebrafish larvae.

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Table of Contents (TOC) Graphic

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Introduction

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Drinking water disinfection is an effective public health measure necessary to kill pathogenic

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microorganisms in source water. Disinfection of reclaimed water dramatically decreases the morbidity

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and mortality of infectious diseases.1 At present, the widely used liquid chlorine, chloramine, ozone, and

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chlorine dioxide are known to react with natural organic matter or environmental pollutants present in

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source water to generate disinfection byproducts (DBPs).2 Many identified DBPs are cytotoxic,

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genotoxic, and mutagenic in vitro or in vivo,3 and epidemiological studies suggest that the consumption

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of chlorinated drinking water is potentially associated with adverse reproductive health effects and an

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increased risk of bladder cancer.4 As such, several DBPs and classes of DBPs are regulated in North

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America, including chlorites, bromate, trihalomethanes, and haloacetic acids (HAAs).5, 6

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An emerging class of DBPs are the halobenzoquinones (HBQs).7, 8 HBQ-DBPs have been detected

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in treated drinking water and swimming pool water across North America at ng/L levels, with

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2,6-DCBQ, the most prevalent HBQ-DBP, detected at an occurrence frequency of 100%.7,9-11

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Quantitative structure toxicity relationship (QSTR) analysis predicted HBQs to be 1000-fold more toxic

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in comparison to regulated DBPs, and are likely to be carcinogenic and mutagenic.8 In vitro toxicity

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studies have reported that HBQs, including 2,6-DCBQ and 2,6-DBBQ, are cytotoxic to T24 human

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bladder cancer cells. The supplementation of glutathione (GSH) mediated the detoxification of HBQs in

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T24 cells,12 suggesting that HBQ cytotoxicity was related to the generation of ROS.13,

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Quinone-induced oxidative stress has also resulted in oxidative damage to proteins and DNA, forming

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protein and DNA adducts.15-18 The 2,5-HBQ isomers, 2,5-DCBQ and 2,5-DBBQ, have been shown to be

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more cytotoxic than their corresponding 2,6-HBQ analogues, although the compounds share the same

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mechanism of toxicity.19,20 Additionally, 2,6-DCBQ and 2,6-DBBQ have been shown to induce cell

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cycle arrest of human neural stem cells in vitro, indicating the potential developmental neurotoxicity of

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this class of DBPs.21 Although inconsistent correlations between maternal exposure to treated water and

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developmental effects in fetuses have been shown in epidemiological studies,22 an increased risk of

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various adverse developmental outcomes have been reported.23-25 Thus, further experimental studies

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using in vivo models are required 1) to determine if oxidative stress contributes to HBQ-induced toxicity


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in vivo and 2) to assess the potential developmental toxicity of this emerging class of DBPs.

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Zebrafish embryos are an established model for evaluating the developmental toxicity of

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environmental pollutants,26 and for high throughput screening, due to the high fecundity, rapid

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embryogenesis, and continuous reproduction of the transparent larvae.27 Although there are increasing

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studies regarding the toxicity mechanisms of DBPs, information on their aquatic toxicity is limited. In

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previous studies, adult zebrafish were used to assess the aquatic toxicity of dichloroacetonitrile and

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2,2-dichloroacetamide, which exhibited bioaccumulation and neurotoxicity.28-30 Zebrafish embryos were

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employed to evaluate the acute and developmental toxicity of different halogenated DBPs, with

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chlorinated DBPs shown to be less toxic than their corresponding bromine- and iodine-substituted

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analogues.31 Dr. Zhang's team has also established a marine polychaete model for testing developmental

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toxicity of DBPs.32, 33 DBPs are commonly released and generated in the aquatic environment, although

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few have been identified.34 Furthermore, potential human or environmental exposures to HBQs are not

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limited to DBPs. TCBQ is a commonly used global fungicide, with an annual production in China over

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2000 tons.35 TCBQ is also an oxidative metabolite of pentachlorophenol, another widely-used pesticide,

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disinfectant, and preservative.35 Both chemicals have been identified as source water contaminants. Thus,

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because of the ease at which these contaminants may enter the aquatic environment, it is important to

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evaluate the toxicity of HBQs in aquatic organisms.

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Based on previous studies, our hypothesis was that developing zebrafish embryos would show

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developmental toxicity and oxidative damage upon exposure to five HBQs (2,6-DCBQ, 2,5-DCBQ,

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2,5-DBBQ, TCBQ, and TBBQ). To test this hypothesis, we examined embryo mortality, the rate of

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malformations, ROS production, changes in the activity and concentration of antioxidant proteins,

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oxidative DNA damage, and apoptosis in zebrafish larvae to provide both morphological observations

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and quantitative molecular biological endpoints. For comparison, two regulated HAA-DBPs and

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iodoacetic acid (IAA) were also examined. The findings of this study will illustrate a useful method for

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screening the developmental toxicity of emerging DBPs and evaluating their potential aquatic toxicity.

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Materials and Methods

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Chemicals

and

Reagents.

Standards

of

2,6-dichloro-1,4-benzoquinone


(2,6-DCBQ),


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2,5-dichloro-1,4-benzoquinone

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tetrachloro-1,4-benzoquinone (TCBQ), and tetrabromo-1,4-benzoquinone (TBBQ) were obtained from

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Tokyo Chemical Industry (TCI; Toshima, Kita-ku, Tokyo, Japan) (Table S1). HBQs were dissolved in

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dimethyl sulfoxide (DMSO), which was used as the vehicle control in each assay. Tricaine (MS-222,

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150 mg/L), used to anesthetize the zebrafish larvae, and 2′,7′-dichlorofluorescin diacetate (DCFDA)

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were purchased from Sigma-Aldrich (St. Louis, MO, USA). Acridine Orange (AO) was purchased from

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the Biosharp Company (Shanghai, China). N-acetyl-L-cysteine (NAC) was purchased from TCI and

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diluted in sterilized water. Dichloroacetic acid (DCA) was purchased from Macklin Biochemical

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(Shanghai, China), dibromoacetic acid (DBA) from ZZ Standard (Shanghai, China), and IAA from

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Aladdin Bio-Chem Technology (Shanghai, China) (Table S1).

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Zebrafish Maintenance and Experimental Design. All experiments were performed on zebrafish

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embryos hatched by the wild type AB strain (3-months old, from the Institute of Hydrobiology, Chinese

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Academy of Sciences), maintained in 28 ± 0.5°C aerated water with 0.25-0.75‰ (w/v) salinity. Adult

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fish were paired, two males and two females in each tank, and were separated the night before the

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collection of fertilized eggs. Approximately thirty embryos were randomly selected, placed into 6-wells

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plates, and exposed to HBQs (0-16 µM) from 4 hours post fertilization (hpf) to 120 hpf. For all

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experiments, three replicate wells were prepared for each exposure group. A larger number of embryos

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was used in high concentration groups in all assays except the mortality assay to ensure sufficient

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numbers of surviving larvae for each assay. Both control and exposure groups contained less than 0.007%

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(v/v) DMSO. Embryos were also treated in the presence or absence of 50 µM of the antioxidant NAC, as

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it has been demonstrated previously that concentrations of NAC less than 50 µM are non-toxic and

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effective in eliminating ROS in zebrafish embryos.36,37 The HAAs (DCA, DBA, and IAA; 0-5 mM)

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were studied in parallel for comparison with the HBQs.

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Morphological Observations. The morphological development of embryos after HBQ exposure (0, 1, 2,

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5, 8, 10, 16 µM) was observed using a stereoscopic microscope (ZEISS SteREO Discovery. V12,

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Germany) during three stages of development: pharyngula, hatching, and swimming larva. Mortality (24

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hpf), hatching rate (60 hpf and 72 hpf), heart rate (72 hpf), uninflated swim bladder (120 hpf), and

(2,5-DCBQ),

2,5-dibromo-1,4-benzoquinone


(2,5-DBBQ),

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various other malformations (72 hpf; tail injury, pericardial edema, shortened body length, shortened

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yolk sac extension, developmental delay) were recorded.

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Measurement of ROS and Antioxidant Molecules. ROS generation in embryos exposed to HBQs (0.5,

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1, 5, 10 µM), HAAs (0.5, 1 mM), or H2O2 (0.5%, 1%) was measured using DCFDA. Briefly, 40 live

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larvae (120 hpf) for each treatment group were washed with cold PBS (pH 7.4) and homogenized in cold

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buffer (0.32 mM sucrose, 20 mM HEPES, 1 mM MgCl2, and 0.5 mM phenylmethyl sulfonylfluoride;

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pH 7.4). The homogenate was centrifuged at 12,000 rpm at 4°C for 10 min, and the supernatants were

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transferred to clean tubes for analysis. Fluorescence intensity was measured using a microplate reader

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(Synergy™ 2 Multi-Mode Microplate Reader, Biotek Instruments, VT, USA), with excitation and

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emission at 485 and 530 nm, respectively. Results are expressed as a percentage (%) relative to the

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control. In addition, an in vivo zebrafish ROS assay was performed. Briefly, after exposure of the larvae

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to 2,5-DCBQ (120 hpf; 0, 0.5, 2.5, 5 µM), the larvae were incubated with DCFDA in the dark for 1 h at

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28 °C. For microscopic examination, the larvae were mounted onto glass slides with 3% methylcellulose,

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and in vivo ROS generation was assessed using a fluorescence microscope (ZEISS Axio Vert.A1,

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Germany). The fluorescence intensity of individual larva was quantified by area of integrated optical

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density using Image J software (National Institutes of Health, Bethesda, MD, USA). In both the in vitro

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and in vivo ROS assays, 2,5-DCBQ- and 2,5-DBBQ-treated larvae (120 hpf) were co-exposed to the

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antioxidant NAC to confirm that the generated ROS induce oxidative stress. More details regarding

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these methods are available in the Supporting Information.

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To maintain intracellular redox homeostasis, ROS and other free radicals are eliminated by

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antioxidant systems, including antioxidant enzymes (e.g. superoxide dismutase; SOD) and non-enzyme

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inhibitors (GSH). To assess the antioxidant response to HBQ exposure, SOD activity and GSH

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production were measured according to the protocols of commercial kits from Nanjing Jiancheng

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Bioengineering Institute (NJBI; Nanjing, China). Briefly, 40 live larvae were homogenized and prepared

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as described above for the in vitro ROS assay with the supernatants collected for detection of SOD

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activity and GSH level. The results of the SOD assay are expressed in units of SOD activity per

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milligram of protein (U/mg), wherein 1 U of SOD is defined as the amount of sample that causes a 50%



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inhibition of cytochrome C reduction. GSH content is expressed as a percentage (%) relative to the

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control.

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Determination of DNA Damage. For analysis of DNA damage, larvae were exposed to 2,5-DCBQ,

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2,5-DBBQ, and 2,6-DCBQ (120 hpf; 1, 5, 10 µM). Approximately 40 live larvae were homogenized and

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prepared as described above for the in vitro ROS assay, with the supernatants collected for the

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determination of DNA damage. 8-OHdG, a biomarker of oxidative DNA damage, was measured using

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an enzyme-linked immunosorbent assay (ELISA) according to the manufacturer’s instruction (8-OHdG

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ELISA Kit, NJBI). In addition, a DNA ladder assay was used to study DNA fragmentation and the

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degree of DNA damage after exposure to 2,5-DCBQ and 2,5-DBBQ (120 hpf; 1, 5, 10 µM), following

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the manufacturer’s instruction (Apoptosis DNA Ladder Extraction Kit, KeyGEN BioTECH, Nanjing,

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China). After DNA extraction, gel electrophoresis and imaging were performed with a gel imaging

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system (G-BOX, Syngene, Cambridge, UK).

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Acridine Orange (AO) Staining and Apoptosis Imaging. Larval cell apoptosis was identified by AO

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staining, a nucleic acid-selective metachromatic stain useful for studying apoptosis patterns.38 After 96 h

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exposure to 2,5-DCBQ or 2,5-DBBQ (0.5, 2.5, 5 µM), 15 larvae from each group (n = 3) were washed

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twice in 30% Danieau's solution (58 mM NaCl, 0.7 mM KCl, 0.4 mM MgSO4, 0.6 mM Ca(NO3)2, and 5

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mM HEPES, pH 7.4), transferred to 30% Danieau's solution containing 5 µg/mL AO, and incubated for

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20 min. The larvae were anesthetized with tricaine (MS-222) prior to microscopic observation of

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apoptosis (Carl Zeiss, SteREO Discovery.V12, Germany). Image J software was used to quantify the

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fluorescence intensity of individual larva by area of integrated optical density.

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Quantitative Real-time PCR. To examine HBQ-induced apoptosis, changes in expression of two genes

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associated with apoptosis, p53 and mdm2, were measured. About 100 embryos were cultured in 10-cm

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culture dishes for each exposure group. After treatment with 2,5-DCBQ or 2,5-DBBQ (0.5, 2.5, 5 µM)

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for 120 hpf, 30 surviving larvae were randomly selected for qRT-PCR. Details on sample preparation

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and analysis are described in the Supporting Information. The GenBank accession numbers and forward

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and reverse primer sequences are listed in Table S2, while the fold changes in expression level were

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calculated using the 2-∆∆Ct method.39


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Statistical Analysis. Statistical analysis was performed using GraphPad Prism 7 (Graphpad Software, La

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Jolla, CA, USA) and SPSS 22.0 (IBM Corp., Armonk, NY, USA). Experimental results were expressed

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as the mean ± standard deviation (SD) or as the mean ± the standard error of the mean (SEM). Statistical

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differences between treatment groups in the presence or absence of NAC were determined by Student’s

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t-test. Statistical comparisons of analyses in which multiple treatment groups were tested (including

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ROS production, SOD activity, GSH production, 8-OHdG, AO, mdm2 expression, p53 expression,

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hatching/heart rate) were assessed using a one-way analysis of variance (ANOVA) with a Dunnett's

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post-test.40 Differences were considered statistically significant at p < 0.05. LC50 values, defined as the

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concentration that induces half of the maximum mortality, were calculated from a nonlinear regression

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model. The four-parameter logistic curve regression analysis was performed according to the following

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formula:

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y = minimum + (maximum – minimum) ÷ [1 + (x/LC50)Hill slope]

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where y is mortality and x is the log concentration of the test compound.

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Results and Discussion

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Acute Toxicity (LC50) of HBQs and HAAs. Zebrafish embryo mortality significantly increased with

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HBQ and HAA concentration (Fig. 1A; Fig. S1); however, no significant effect on embryo hatching rate

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at 60 hpf or 72 hpf was observed (Table S3). Mortality rates were used to calculate LC50 values for the

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five HBQs and three HAAs at 24, 48, 72, 96, and 120 hpf (Table 1; Table S4), with the toxic potency of

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the compounds ranked as 2,5-DCBQ ˃ 2,5-DBBQ ˃ 2,6-DCBQ ˃ TBBQ ˃ TCBQ > IAA > DBA ˃

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DCA. The calculated LC50 values show that the acute toxicity of HBQs in zebrafish embryos is two to

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three orders of magnitude greater than the HAAs. These findings are consistent with a previous study

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that evaluated the developmental toxicity of 20 halogenated DBPs in the marine polychaete, Platynereis

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dumerilii. Of the 20 DBPs, 2,5-dibromohydroquinone had the highest developmental toxicity, with a

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reported EC50 value 100 to 1000 times lower than those of the THMs or HAAs.32, 33 To clarify that the

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observed HBQ toxicity is linked to oxidative stress, we examined the effects of HBQs in the presence of

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NAC, a thiol antioxidant and ROS scavenger.41 When NAC (50 µM) was added to the high

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concentration treatment group (16 µM) of each HBQ, a significant reduction in mortality was observed


(Equation 1)


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(t-test, p < 0.05; Fig. 1B), implicating oxidative stress as a mechanism of HBQ-induced embryotoxicity.

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Exposure-induced Oxidative Stress. All five HBQs significantly induced ROS in zebrafish embryos in a

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dose-dependent manner at 120 hpf (ANOVA with Dunnett’s post-test, p < 0.05; Fig. 2A1). In

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comparison to the HAA-DBPs (Fig. 2A2), the HBQs generated similar levels of ROS, but at much lower

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concentrations, as 10 µM of each HBQ produced as much ROS as 1000 µM of each HAA (Fig. 2). Thus,

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the more potent embryotoxicity of the HBQs in relation to the HAAs is likely influenced by their ability

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to induce ROS. Again, it was found that NAC co-exposure mitigated the HBQ-induced effects; 50 µM

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of the antioxidant was sufficient to eliminate excess ROS in the 2,5-DCBQ and 2,5-DBBQ treatment

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groups (Fig. 2A1). For the localization of ROS production in tissue, in vivo imaging of DCFH-DA in

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whole zebrafish larvae was performed after HBQ exposure. Background fluorescence from normal

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physiological processes was observed in both the control and DCFDA treatment groups (Fig. 3A, B).

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However, increased fluorescence over the background was clearly visible in each of the HBQ treatment

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groups (Fig.3 D-F). The fluorescence of the 5 µM 2,5-DCBQ treatment group (Fig. 3F) was greatly

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reduced in the presence of exogenous NAC (Fig. 3C), but the adverse effects were not fully attenuated.

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Fluorescence intensity was greatest in the eyes, head, digestive tract, and yolk sac of the larvae, but also

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visible in the muscle tissue of the tail. In the higher concentration 2,5-DCBQ treatment groups (Fig. 3E,

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F), pericardial edema is clearly distinguished under fluorescence (Fig. 3C, E, F). The fluorescence

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intensity of each of the treatment groups was quantified in Figure S2A.

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SOD activity and GSH depletion were also investigated as indicators of oxidative stress, as both

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antioxidants are critically important for the elimination of excess ROS to prevent oxidative damage.42

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With increasing concentration of each HBQ, SOD activity was concurrently inhibited (ANOVA with

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Dunnett’s post-test, p < 0.05; Fig. 2B). Likewise, GSH depletion was dose-dependent in the 2,5-DCBQ

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and 2,5-DBBQ treatment groups (ANOVA with Dunnett’s post-test; Fig. 2C), consistent with in vitro

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studies that found HBQs reduced GSH levels in quinone-treated PC12, T24, and HepG2 cells.12,43, 44

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Both SOD activity and GSH concentrations were significantly recovered with NAC co-exposure (t-test,

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p < 0.05; Fig. 2B, C). As a free radical scavenger, GSH responds rapidly to ROS produced by toxic

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chemicals, 45 while SOD effectively maintains the oxidant balance of the highly reactive superoxide


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anion. The resulting reduction in free GSH and SOD activity increases the vulnerability of zebrafish

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larvae to HBQ-induced oxidative damage. The production of ROS is dependent on the semi-quinone and

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hydroxyl radical structural features of HBQs.5, 16 Because of their electron deficient structure, HBQs are

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reactive electrophiles which can readily react with various bio-nucleophiles, such as proteins and nucleic

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acids. The nucleophilic attack of GSH by quinones and HBQs has been demonstrated.16, 46 Thus, the

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reduction in GSH is likely a result of both GSH oxidation and conjugation.46

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DNA Damage and Apoptosis. Oxidative DNA damage in zebrafish larvae was assessed via the

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measurement of the DNA lesion, 8-OHdG, at 120 hpf. The content of 8-OHdG significantly increased at

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120 hpf in the high dose treatment groups of 2,6-DCBQ, 2,5-DCBQ, and 2,5-DBBQ (1, 5, 10 µM)

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(ANOVA with Dunnett’s post-test, p < 0.05; Fig. 4A), indicating DNA oxidation. This is consistent with

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in vitro studies that found HBQs significantly induced 8-OHdG formation in T24 cells and Chinese

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hamster lung cells.15, 47 Simultaneously, measurements (Fig. 4B) of DNA fragmentation clearly showed

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HBQ-induced DNA damage, as the DNA smear extended throughout the gel lane in the high dose

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treatment groups of 2,5-DCBQ (Fig. 4B1) and 2,5-DBBQ (Fig. 4B2). No significant fragmentation was

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visible in the control group (Fig. 4B1, second lane from left). DNA fragmentation is a hallmark of

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apoptosis, which could be induced as a result of irreparable HBQ-induced oxidative DNA damage. Thus,

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we examined apoptotic cells in vivo in whole zebrafish larvae using AO staining. Treatment

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concentrations of 2,5-DCBQ (Fig. 5A2-4) and 2,5-DBBQ (Fig. 5B2-4) exhibited higher fluorescence

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intensity in the heart, brain, yolk sac, and mouth of the larvae in comparison to the control group (Fig.

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5A1). Furthermore, co-exposure of 2,5-DCBQ (5 µM) with 50 µM NAC resulted in fewer apoptotic

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cells, demonstrated by the lower fluorescence intensity, and prevented pericardial edema (Fig. 5B1). The

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quantified fluorescence intensity of each treatment group is available in the Supporting Information

250

(Figure S2B).

251

For further examination of HBQ-induced apoptosis, two genes (p53, mdm2) associated with

252

apoptosis were examined in larvae at 120 hpf after exposure to 0.5, 2.5, or 5 µM 2,5-DCBQ or

253

2,5-DBBQ. The pattern of p53 expression was similar in larvae exposed to the HBQs investigated, as

254

p53 was significantly up-regulated (1.73-fold, 1.42-fold) in the 0.5 µM 2,5-DCBQ or 2,5-DBBQ



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treatment groups, but was unchanged at higher concentrations (2.5, 5 µM) (ANOVA with Dunnett’s

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post-test, p < 0.05; Fig. S3). Alternatively, mdm2 expression differed between the 2,5-DCBQ and

257

2,5-DBBQ treatment groups. Compared to the control, mdm2 was significantly down-regulated in larvae

258

exposed to 2.5 or 5 µM 2,5-DCBQ (2.71-fold, 2.76-fold) or 5 µM 2,5-DBBQ (2.12-fold), but was

259

significantly up-regulated (1.46-fold) in the 2.5 µM 2,5-DBBQ treatment group (ANOVA with Dunnett’s

260

post-test, p < 0.05; Fig. S3).

261

Both DNA damage and oxidative stress elicit the activation of p53,48 which in turn activates

262

apoptotic gene expression, leading to apoptosis.49,

50

263

significantly up-regulated in the lowest exposure group of 2,5-DCBQ and 2,5-DBBQ (0.5 µM), but was

264

not statistically changed in the higher exposure groups (2.5 and 5 µM). This is most likely related to the

265

expression levels of mdm2. The p53 protein is negatively regulated by mdm2, which suppresses p53

266

transcription.51 Hence, in the higher 2,5-DCBQ exposure groups, where mdm2 mRNA expression was

267

significantly down-regulated, upregulation of p53 was not necessary to maintain levels of p53 due to the

268

lack of its inhibitor. However, in the 2,5-DBBQ treatment groups, the relationship between p53 and

269

mdm2 is not as clear. These findings illustrate the delicate balance between these two proteins and the

270

control of apoptosis, and may indicate differing mechanisms of action between 2,5-DCBQ and

271

2,5-DBBQ. Nevertheless, the perturbation of these apoptosis genes indicate that HBQ exposure can

272

affect apoptosis signaling pathways in developing zebrafish.

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HBQ-Induced Malformations in Zebrafish Larvae. 2,6-DCBQ, 2,5-DCBQ, and 2,5-DBBQ induced the

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highest levels of mortality in exposed zebrafish embryos. Because 2,6-DCBQ is the most prevalent

275

HBQ-DBP detected in treated tap water,8 larvae exposed to these three HBQs were examined

276

morphologically to assess abnormal development. At 72 hpf, various physical malformations were

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observed in the zebrafish larvae. As shown in Figure 6, the observed abnormalities included heart

278

malformations, curved spine and caudal fin, pericardial edema, and shortened body length and yolk sac

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extension. Malformations were observed for exposure doses as low as 2 µM (Fig. 6B), where 2,5-DCBQ

280

clearly affected yolk sac extension and body length. Higher doses of 2,5-DCBQ (Fig. 6C-E) resulted in

281

more lethal damages, including severe tail injuries and pericardial edema. This was also observed in

Interestingly, p53 mRNA expression was


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higher doses of 2,6-DCBQ (Fig. 6H-J), where caudal scoliosis and pericardial edema were identified

283

(Fig. S4). Lower doses of 2,6-DCBQ significantly inhibited the rate of larval development

284

(developmental delay; Table S5). Uninflated swim bladders were observed in up to 20% of the larvae

285

exposed to 2,6-DCBQ, 2,5-DCBQ, or 2,5-DBBQ (Table S5; Fig. S5). Consistent with our other findings,

286

the number of deformities was significantly reduced with NAC co-exposure (Fig. 6F), with the

287

malformation rate of the 10 µM treatment groups decreasing from nearly 80% to less than 15% (Table

288

S5).

289

Oxidative stress and apoptosis have both been implicated in developmental toxicity during zebrafish

290

embryogenesis.36, 52 Environmental stresses induce malformations and mortality in fish in both natural

291

and aquaculture environments, while stress-induced apoptosis is thought to contribute to abnormal

292

development during embryogenesis.53 Upon HBQ exposure, several developmental malformations were

293

observed in zebrafish embryos. Low doses of HBQs also prevented the expansion of the gas bladder (Fig.

294

6), which further contributed to limited motility and ingestion, and eventually increased mortality.54

295

Heart edema and the separation of the atrium-ventricle were observed (Fig. 6E, S4), suggesting that the

296

developing heart may be a sensitive target organ in zebrafish embryos. Malformations of the heart and

297

pericardium may impact cardiac function and lead to an abnormal heartbeat and circulation failure,

298

resulting in stunted development (Fig. 6I). However, no adverse effects of HBQs or HAAs on zebrafish

299

heart rate were observed at 72 hpf (Table S6). The high percentage of apoptotic cells around the heart

300

(Fig. 5) may partially account for the resulting heart malformations.

301

This study presents an efficient approach to evaluate the toxicity and potential mechanisms of DBPs

302

using zebrafish embryos as a model of developmental toxicity. The results of this study show that HBQs

303

induce ROS generation and inhibit the cells’ anti-oxidative response in developing zebrafish, resulting in

304

death, physical malformations, oxidative DNA damage, and apoptosis. These findings were consistent

305

with in vitro studies, which linked the cytotoxicity of HBQs to ROS generation, antioxidant enzyme

306

inhibition, and oxidative DNA damage in mammalian cells.55-57 The acute toxicity and ROS induction of

307

HBQs was up to 200 times more potent than the two regulated HAAs (DCA and DBA) and one of the

308

most toxic DBPs, IAA. Hence, ours and others results support that aromatic halogenated DBPs,



Page 14 of 29

309

including halogenated phenolic and halopyrrole DBPs, induce significantly greater developmental

310

toxicity than aliphatic DBPs.58-60 More research into the pharmacodynamics of aliphatic and aromatic

311

halogenated DBPs are required to understand the toxicity difference of these DBP classes in vivo.

312

Furthermore, long-term exposures with environmentally-relevant concentrations of HBQs are required

313

to fully understand their toxicological significance.

314


Page 15 of 29


315

ASSOCIATED CONTENT

316

Supporting Information

317

The Supporting Information is available free of charge on the ACS Publications website, including

318

additional details of the methods and results (six tables and five figures).

319

AUTHOR INFORMATION

320

Corresponding Authors

321

*E-mail: [email protected].

322

[email protected].

323

ORCID

324

Xing-Fang Li: 0000-0003-1844-7700 641

325

Funding

326

This work was financially supported by grants from the National Natural Science Foundation of China

327

(21677062, 21507155), and grants from the Natural Sciences and Engineering Research Council of

328

Canada (NSERC), Alberta Health, and Alberta Innovates-Energy and Environmental Solutions.

329

Acknowledgements

330

We thank Dr. Mengxi Cao from Jianghan University for her kind assistance with zebrafish feeding and

331

experiments.

332



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water. Environ. Sci. Tech. 2017, 51, 3435-3444.



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Page 22 of 29

Table 1. Calculated LC50 Values of the HBQs and HAAs at 24 hpf Compounds Halobenzoquinones (HBQs) 2,5-DCBQ 2,6-DCBQ 2,5-DBBQ TBBQ TCBQ Haloacetic acids (HAAs) IAA DBA DCA

LC50 (mean ± SD) 2,5-dichloro-1,4-benzoquinone 2,6-dichloro-1,4-benzoquinone 2,5-dibromo-1,4-benzoquinone tetrabromo-1,4-benzoquinone tetrachloro-1,4-benzoquinone

µM 4.6 ± 0.2 6.6 ± 0.2 5.6 ± 0.2 9.4 ± 0.3 9.8 ± 0.4

Iodoacetic acid Dibromoacetic acid Dichloroacetic acid

1900 ± 240 2200 ± 190 2600 ± 170

485 486 487


Page 23 of 29


488 489 490 491 492 493 494 495 496 497 498 499 500 501 502 503

Figure 1. Effect of HBQs (1, 2, 5, 8, 10, 16 µM) on the mortality of zebrafish embryos at 24 hpf (A);

504

Effect of NAC co-exposure on HBQ-induced mortality (B). Error bars indicate mean ± SD. Three

505

replicate experiments were performed for each concentration (n = 3). t-test: HBQ group vs. respective

506

HBQ + NAC group: ***p < 0.001.

507 508



Page 24 of 29

509

510 511

Figure 2. ROS formation in larvae treated with 2,5-DCBQ or 2,5-DBBQ (1, 5, 10 µM) with or without

512

NAC co-exposure (A1). Effects of different concentrations (0.5, 1 mM) of DCA, DBA, IAA, or H2O2 on

513

ROS production in comparison to 2,5-DCBQ (A2). Effects of different concentrations (1, 5, 10 µM) of

514

five HBQs on SOD activity with or without NAC co-exposure (B). Effect of different concentrations (1,

515

5, 10 µM) of 2,5-DCBQ and 2,5-DBBQ on GSH levels with or without NAC co-exposure (C). Error

516

bars indicate mean ± SEM. Three replicate experiments were performed for each group. ANOVA with

517

Dunnett’s post-test: *p < 0.05, **p < 0.01, ***p < 0.001, ****p < 0.0001.

518


Page 25 of 29


519 520 521 522 523 524 525 526 527 528 529 530 531 532 533 534

Figure 3. In vivo assessment of ROS production in larvae at 120 hpf in the negative control (A),

535

DCFDA control (B), and the 2,5-DCBQ treatment groups (0.5, 2.5, 5 µM) (D-F). Co-exposure of 5 µM

536

2,5-DCBQ with 50 µM NAC (C) attenuated the detected fluorescence. FE: fluorescence enhancement;

537

PE: pericardial edema; SYSE: shortened yolk sac extension; SAV: separation of the atrium-ventricle;

538

USB: uninflated swim bladder; ISB: inflated swim bladder. Scale bar, 450 µm.

539 540



Page 26 of 29

541 542 543 544 545 546 547 548 549 550 551 552 553 554 555 556 557 558

Figure 4. Increase in 8-OHdG lesions at 120 hpf in the genomic DNA of zebrafish larvae after HBQ (1,

559

5, 10 µM) exposure (A). DNA fragmentation is also observed in the 2,5-DCBQ (B1) and 2,5-DBBQ (B2)

560

treatment groups (1, 5, 10 µM). Error bars indicate the mean ± SEM. Three replicate experiments were

561

performed for each exposure group (n = 3). ANOVA with Dunnett’s post-test: *p

Halobenzoquinone-Induced Developmental Toxicity, Oxidative Stress

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