Aquatic Toxicology 148 (2014) 195–203

Contents lists available at ScienceDirect

Aquatic Toxicology journal homepage: www.elsevier.com/locate/aquatox

Developmental exposure of zebrafish (Danio rerio) to bisphenol-S impairs subsequent reproduction potential and hormonal balance in adults Mohammad Naderi a,∗ , Marian Y.L. Wong b , Fatemeh Gholami c,1 a

Department of Marine Biology, Faculty of Marine Science, Khorramshahr University of Marine Science and Technology, Khorramshahr, Iran School of Biological Sciences, University of Wollongong, Wollongong, NSW, Australia c Department of Fundamental Science, Faculty of Biology, Yasuj Branch, Islamic Azad University, Yasuj, Iran b

a r t i c l e

i n f o

Article history: Received 2 November 2013 Received in revised form 7 January 2014 Accepted 10 January 2014 Keywords: Bisphenol-S Reproduction Sex steroid Thyroid hormone Vitellogenin Zebrafish (Danio rerio)

a b s t r a c t In the recent years, there has been a growing concern about the production and use of bisphenol-A substitute, namely bisphenol-S (BPS). Due to its novel nature, there have been few studies addressing the ability of BPS to disrupt the endocrine system of animals. In the present study, zebrafish (Danio rerio) embryos were exposed to and reared in various concentrations of BPS (0, 0.1, 1, 10 and 100 ␮g/l) for 75 days. Then adult males and females were paired in spawning tanks for 7 days in clean water and the consequent effects on fish development, reproduction, plasma vitellogenin (VTG), sex steroids and thyroid hormone levels were investigated as endpoints. After 75 days of exposure, there was a skewed sex ratio in favor of females. The results also showed that body length and weight significantly decreased in males exposed to 100 ␮g/l of BPS. Gonadosomatic index was significantly reduced in fish at ≥10 ␮g/l. Hepatosomatic index exhibited a significant increase in both male and female fish. At ≥1 ␮g/l of BPS, plasma 17␤-estradiol levels were significantly increased in both males and females. However, plasma testosterone showed a significant reduction in males exposed to 10 and 100 ␮g/l of BPS. A significant induction in plasma VTG level was observed in both males and females at ≥10 ␮g/l of BPS. Plasma thyroxine and triiodothyronine levels were significantly decreased at 10 and 100 ␮g/l of BPS in males, and at 100 ␮g/l in females. Egg production and sperm count were also significantly decreased in groups received 10 and 100 ␮g/l of BPS. Moreover, once time to hatching and hatching rates were calculated for fertilized eggs the postponed and decreased rates of hatching were observed. Taken together, these results suggest that developmental exposure to low concentrations of BPS has adverse effects on different parts of the endocrine system in zebrafish. © 2014 Elsevier B.V. All rights reserved.

1. Introduction The world is awash in a wide variety of man-made chemicals that have biological activity and that can impose adverse health consequences on wildlife and humans. In the past two decades, there has been an increasing concern over some classes of environmental contaminants known as endocrine-disrupting chemicals (EDCs), which have the ability to interfere with complex endocrine functions, often through mimicking or blocking endogenous hormones (reviewed by Diamanti-Kandarakis et al., 2009; Schug et al., 2011). Stemming from the production of various synthetic industrial chemicals and their by-products,

∗ Corresponding author at: No 13, Karoon lane, Second Golestan, Yasuj, Kohgiluyeh and Boyerahmad 75918-44349, Iran. Tel.: +98 9177416053(mob). E-mail address: [email protected] (M. Naderi). 1 Tel.: +98 9360346456. 0166-445X/$ – see front matter © 2014 Elsevier B.V. All rights reserved. http://dx.doi.org/10.1016/j.aquatox.2014.01.009

pharmaceuticals, and pesticides, these compounds have resulted in developmental deficiencies and reproductive impairments in a wide range of vertebrate and invertebrate species (Richter et al., 2007; LeBlanc, 2007; Crain et al., 2007; Zou, 2010; Rubin, 2011). Most notable among environmental EDCs are compounds that act as agonists or antagonists on vertebrate estrogen or androgen receptors. One such compound is xenoestrogen bisphenol-A (BPA, 2,2-bis-(4-hydroxyphenyl)-propane; CAS Registry No. 80-05-7). BPA is one of the key ingredients used to manufacture polycarbonate plastic and epoxy resins and a variety of other products including polyvinyl chloride (PVC), thermal printing paper used for cash register receipts, plastic containers for food and beverages, electronic components, and paper coatings (Staples et al., 1998; Biedermann et al., 2010; Vandenberg et al., 2009). Due to extensive rate of production and consumption, BPA ubiquitously presents in various environmental matrices (Tsai, 2006; Vandenberg et al., 2007, 2009; Flint et al., 2012). Owing to its adverse effects on humans (Bondesson et al., 2009; Vandenberg et al., 2010) and

196

M. Naderi et al. / Aquatic Toxicology 148 (2014) 195–203

animals (Goodman et al., 2006; Aluru et al., 2010; Li et al., 2011), several regulatory agencies including Health Canada (2009), the US Food and Drug Administration (2010), and European Union (European Commission, 2011) during recent years have banned use of BPA in infant formula bottles. Consequently, to comply with regulations on BPA, several functional alternatives have been introduced and developed into the global markets (Gallart-Ayala et al., 2011; Liao et al., 2012; Flint et al., 2012). Bisphenol-S (BPS, bis-(4-hydroxyphenyl)-sulfone; CAS Registry No. 80-09-1) another bisphenolic compound with more stability against high temperature and resistance to sunlight, is considered to be able to partially replace BPA in the industrial applications (Chen et al., 2002; Liao et al., 2012). This chemical is commonly used in epoxy glues, canned foodstuffs, thermal receipt papers, paper currencies, luggage tags, food cartons, flyers, newspapers, ˜ et al., 2010; Liao et al., 2012). Therefore, direct exposure etc. (Vinas of humans to BPS based on daily activities is inevitable. Similar to BPA, BPS has shown the ability to act as an estrogen mimic or an anti-androgen (Hashimoto et al., 2001; Kuruto-Niwa et al., 2005; Kitamura et al., 2005; Grignard et al., 2012). Correspondingly, it is highly likely that BPS has the potential to interfere with and disrupt the normal functions of endocrine system in organisms. Because of the novel nature of this compound, however, next to nothing is known about the effects of BPS on the endocrine system of animals in vivo. Only in a recent study, it has been demonstrated that BPS could disturb the balance of sex steroid hormones and impair normal reproduction of adult zebrafish (Danio rerio, Ji et al., 2013). The early life stages of fish is known to be the most vulnerable stage to effects of EDCs even at very low levels and have therefore frequently been used as the basis of toxicity tests (Kime, 1998). At this step, even minor concentrations of endogenous hormones (steroid and thyroid hormones in particular) play crucial roles in sex differentiation, metabolism, embryological growth and development, and osmoregulation (Power et al., 2001; McLachlan, 2001; Thomas, 2008). It is well documented that chronic exposure of developing fish to xentoestrogens, which can affect hormonal balance in animals, may lead to later consequences like intersex and sex reversal (Vajda and Norris, 2011). However, there is frequently little attempt to understand how exposure to EDCs during this narrow window of development affects later reproductive fitness in adults (but see van Aerle et al., 2002). To our knowledge, no study so far has assessed reproductive success of adult fish, which experienced developmental exposure to BPS. Zebrafish (D. rerio) was chosen as the test species for this study due to its well-known ecotoxicological and developmental advantages (Briggs, 2002; Stegeman et al., 2010). This study was conducted to evaluate the endocrine disrupting effects of BPS on reproduction and balance of steroid and thyroid hormones in adult zebrafish, which experienced early exposure to different concentrations of this chemical. We also examined other different endpoints such as survival rate, sex ratio, plasma vitellogenin (VTG), gonadosomatic index (GSI), hepatosomatic index (HSI) of zebrafish in F0 generation, as well as hatching rate and time to hatching in the offspring generation (F1 ).

2.2. Zebrafish maintenance and embryo exposure Wild-type adult zebrafish (D. rerio) of both sexes at an age of 4–5 months were purchased from a local supplier and acclimated in 30L glass tanks containing charcoal-filtered re-circulating water (pH 7.0–7.4) for 2 weeks prior to the experiment. Fish were kept at a temperature of 28 ± 0.5 ◦ C with a photoperiod of 14 h light: 10 h dark according to the protocols previously described by Shi et al. (2008) and Yu et al. (2010). Fish were fed with freshly hatched brine shrimp (Artemia nauplii) twice a day. Embryos obtained from group spawns were collected 2 h post-fertilization (hpf), washed and examined under a stereomicroscope. Those embryos that had developed normally and reached the blastula stage were selected and placed in several Petri dishes until 2 days post-fertilization (dpf) for subsequent experiments. Approximately 700 embryos (2 dpf) were randomly allocated into 5 l tanks containing 2.5 l of BPS exposure solution (0, 0.1, 1, 10 and 100 ␮g/l), thereby creating 5 experimental treatment conditions. Controls (i.e. 0 ␮g/l) only received 0.01% acetone (v/v). The experiment was run in three replicates. The fish were held in the exposure tanks for 75 dpf. During this period, any spawned eggs and dead embryos/larvae were removed and 50% of the exposure solution was renewed daily. 2.3. Sexing and pairing of adult fish After the exposure period, these fish that had been reared under the 5 treatments (i.e. the F0 generation) were sexed either by examining the microscopic features of the gonads or external examination of coloration (male zebrafish develop a conspicuously yellow/golden coloration on their underside, anal fin and caudal fin, whereas females develop a silver coloration, (Paull et al., 2008)). Males and females were distributed in separate tanks containing charcoal-filtered tap water and allowed to acclimate to the new condition for two weeks. Thereafter, males and females from the same treatments were randomly assigned to 3 l tanks (spawning tanks; 2 males × 2 females colonies) and acclimatized for 3 days. For the next 7 days, the numbers of eggs spawned in each treatment group were collected and counted daily, and hatching rate and time to hatching (h) were calculated. Three replicates were considered for each spawning tank. At the end of the experimental period, the fish were anesthetized with 2-phenoxyethanol, body weight and total length were measured, and then the gonads and livers were removed and weighed. GSI (100 × [gonad weight (g)/body weight (g)]) and HSI (100 × [liver weight (g)/body weight (g)]) were calculated. Sperm count was measured for both treated males and solvent controls as previously described in Larsen and Baatrup (2010). Blood was sampled from each fish by severing the caudal peduncle and collecting the blood into heparinized microcapillary tubes. Blood samples from 4 to 6 fish of the same sex from the same treatment were pooled to generate a total of three replicate pooled blood samples for each treatment. The blood samples were centrifuged at 3000 × g for 5 min at 4 ◦ C, and the plasma collected and stored at −80 ◦ C until assay. 2.4. Plasma hormones and vitellogenin measurement

2. Materials and methods 2.1. Chemicals Bisphenol-S (technical grade, purity >98%) was purchased from TCI Europe NV. (Zwijndrecht, Belgium). Stock solutions of BPS were prepared in HPLC-grade acetone and stored in the dark at 4 ◦ C. Solvent concentration was kept at 0.01% acetone (v/v) throughout the experiment. 2-Phenoxyethanol (ethylene glycol monophenyl ether) was obtained from Merck Schuchardt (Hohenbrunn, Germany). All other chemicals used were analytical grade.

Plasma concentrations of 17␤-estradiol (E2), testosterone (T), VTG, thyroxine (T4) and triiodothyronine (T3) were measured using enzyme-linked immunosorbent assay (ELISA) kits (Uscnlife, Wuhan, China; RD Chemical, Mountain View, CA, USA), following the manufacturer’s instructions. 2.5. Statistical analysis All values are represented as mean ± S.E.M. Kolmogorov– Smirnov, and Levene’s tests were applied in order to verify the

M. Naderi et al. / Aquatic Toxicology 148 (2014) 195–203

197

Fig. 1. Survival rate of zebrafish exposed to BPS for 75 days. Values represent mean ± SEM (n = 3 samples). * Significant difference compared to the solvent control (p < 0.05).

data distribution and the homogeneity of variances, respectively. Two-way ANOVA was used to calculate statistical significant effects of treatment, sex and their interaction on each response variable. Once a significant difference was observed Dunnett’s post hoc test was performed to compare differences among treatment groups using SPSS 16.0 (SPSS, Chicago, IL, USA). Sex ratio, survival and hatching rates were tested with the Chi-Square test (X2 -test). For all statistical results, a probability of p < 0.05 was considered significant. 3. Results 3.1. Sex ratio and mortality rates Exposure of zebrafish embryos to different doses of BPS resulted in a skew in the sex ratio towards females in adults. The proportion of females in groups treated with 10 and 100 ␮g/l of BPS were 58.8% and 66.7% respectively, compared with 46.3% in the solvent control group (X2 -test, X2 = 12.14, p = 0.016). Fish reared in the treatment with the highest dosage of BPS (100 ␮g/l) showed significantly higher mortality rates relative to those from the solvent control group (X2 -test, X2 = 27.50, p = 0.001; Fig. 1). 3.2. Morphological traits BPS affected body length, weight, GSI and HSI of adult zebrafish (Table 1). For body length, there was a significant effect of treatment (Two-way ANOVA: F4,4 = 7.7, p = 0.036) and sex (F1,4 = 151.2, p = 0.001) but no significant interaction between treatment and sex (F4,110 = 1.26, p = 0.28). Specifically, body length was significantly lower in male fish than female fish (F1,4 = 151.2, p = 0.001) and significantly lower in fish treated with 100 ␮g/l of BPS compared to the solvent controls (Dunnett’s test: F4 = 7.85, p = 0.001; Table 1). For weight, there was no significant effect of treatment (Two-way ANOVA: F4,4 = 0.842, p = 0.564), but a significant effect of sex (F1,4 = 13.92, p = 0.020) and an interaction between treatment and sex (F4,110 = 6.27, p = 0.001). Specifically, males were significantly lighter than females (F1,4 = 13.92, p = 0.020) and this difference was most pronounced in the 100 ␮g/l BPS treatment (F4 = 15.66, p = 0.002; Table 1). For GSI, there was no significant effect of treatment (Two-way ANOVA: F4,4 = 1.71, p = 0.306), but a significant effect of sex (F1,4 = 704.31, p = 0.001) and an interaction between treatment and sex (F4,110 = 10.81, p = 0.001). Specifically, GSI of males was significantly lower in groups treated with 10 and 100 ␮g/l of BPS compared to the solvent controls (F4 = 34.72, p = 0.001), whereas the GSI of females was significantly lower in groups treated with 100 ␮g/l BPS, compared to the solvent controls (F4 = 14.42, p = 0.001). Finally, for HSI, there was a significant effect of treatment (Two-way ANOVA: F4,4 = 6.33, p = 0.05) and an

Fig. 2. Plasma levels of (A) T, (B) E2 and (C) VTG in zebrafish after 75 days of exposure to BPS. Data are expressed as the mean ± SEM (n = 3 samples). * Significant difference compared to the solvent control (p < 0.05).

interaction between treatment and sex (F4,110 = 7.38, p = 0.001), but no significant effect of sex (F1,4 = 0.736, p = 0.439). Post-hoc tests revealed that female fish treated with 10 and 100 ␮g/l of BPS had significantly higher HSI when compared with the control fish (Dunnett’s test: F4 = 25.22, p = 0.001). However, only male fish in the 100 ␮g/l exposure group exhibited significant differences in HSI relative to the solvent controls (Dunnett’s test: F4 = 29.01, p = 0.001; Table 1). 3.3. Endocrinological traits There was a significant interaction between treatment and sex on plasma T levels (Two-way ANOVA: F4,20 = 5.74, p = 0.003). Specifically, plasma T levels were significantly lower in males exposed to 10 and 100 ␮g/l of BPS compared to the solvent controls (Dunnett’s test: F4 = 9.85, p = 0.02), whereas in females no significant change was found (Dunnett’s test: F4 = 2.08, p = 0.158; Fig. 2a). In addition, there was a significant interaction between treatment and sex on plasma E2 levels (Two-way ANOVA: F4,20 = 5.74, p = 0.003) and a significant sex effect (F1,4 = 17.29, p = 0.014; Fig. 2b). Specifically in males, plasma E2 levels were higher following exposure to 1, 10 and 100 ␮g/l of BPS relative to the solvent controls (Dunnett’s test: F4 = 16.54, p = 0.001), while in females, individuals in groups exposed to 10 and 100 ␮g/l of BPS had significantly higher plasma E2 levels compared to the solvent controls (Dunnett’s test: F4 = 13.16, p = 0.001; Fig. 2b). BPS treatment also had a significant effect on plasma VTG levels although this differed for males and

198

M. Naderi et al. / Aquatic Toxicology 148 (2014) 195–203

Table 1 Effects of bisphenol-S on somatic indices of male and female zebrafish.

Solvent control 0.1 1 10 100

Female

Male

BPS concentrations (␮g/l) Body weight (mg)

Body length (mm)

HSI (%)

GSI (%)

Body weight (mg)

Body length (mm)

HSI (%)

GSI (%)

238.33 ± 9.09 239.17 ± 7.39 240 ± 7.56 235.58 ± 6.36 176.83 ± 6.36*

24.28 ± 1.024 22.68 ± 0.617 23.48 ± 0.774 23.45 ± 1.163 19.41 ± 0.548*

1.20 ± 0.018 1.21 ± 0.016 1.25 ± 0.032 1.23 ± 0.012 1.53 ± 0.037*

1.08 ± 0.024 1.07 ± 0.028 1.02 ± 0.033 0.82 ± 0.022* 0.74 ±0.02*

271.33 ± 7.41 269.67 ± 7.13 273.58 ± 6.16 282 ±5.32 276.33 ± 5.19

28.84 ± 0.565 28.15 ± 0.706 28.77 ± 0.64 28.67 ± 0.409 26.67 ± 0.578

1.23 ± 0.024 1.26 ± 0.036 1.20 ± 0.015 1.43 ± 0.021* 1.49 ± 0.044*

11.49 ± 0.257 10.89 ± 0.258 10.87 ± 0.282 10.82 ± 0.26 9.02 ± 0.138*

The values are represented as mean ± SEM (n = 3; each sample included 12 fish). * Indicates significant difference between exposure and solvent control group (p < 0.05).

females (Two-way ANOVA: F4,20 = 18.9, p = 0.001; Fig. 2c). Specifically, VTG induction was significantly higher for females exposed to 10 and 100 ␮g/l of BPS relative to the solvent controls (Dunnett’s test: F4 = 28.99, p = 0.001). For males, 100 ␮g/l of BPS was the only concentration that caused significant increase in plasma VTG level relative to the solvent controls (Dunnett’s test: F4 = 12.20, p = 0.001). Analysis of plasma thyroid hormones showed a significant effect of treatment on T3 levels (Two-way ANOVA: F4,4 = 9.77, p = 0.024) as well as a significant interaction between treatment and sex on T4 levels (Two-way ANOVA: F4,20 = 3.57, p = 0.024). Specifically, there were significantly lower levels of plasma T3 and T4 in males in groups exposed to 10 and 100 ␮g/l of BPS relative to the solvent controls (Dunnett’s test: F4 = 4.48 and 27.85, p = 0.025, 0.001; Fig. 3a and b). In females, plasma T3 and T4 concentrations were significantly lower in groups exposed to100 ␮g/l BPS compared to the solvent controls (Dunnett’s test: F4 = 5.52 and 5.53, p = 0.013).

experienced 75 days exposure to 10 and 100 ␮g/l of BPS compared to the solvent controls (Dunnett’s test: F4 = 28.16, p = 0.001). In addition, hatching rate and time to hatching of embryos in the 10 and 100 ␮g/l of BPS treatments were adversely affected, even after fish

3.4. Reproductive traits The mean numbers of eggs produced per-colonies per-day in each treatment differed significantly depending on treatment (Fig. 4a). Colonies of zebrafish produced significantly fewer eggs during the 7 day spawning period in the groups that had

Fig. 3. Plasma levels of (A) T3 and (B) T4 in zebrafish after 75 days of exposure to BPS. Data are expressed as the mean ± SEM (n = 3 samples). * Significant difference compared to the solvent control (p < 0.05).

Fig. 4. Reproductive parameters of zebrafish colonies, which experienced exposure to various concentrations of BPS during 7 days. (A) Eggs per colonies per day, (B) hatching rate, (C) time to hatching, and (D) sperm count. Data are expressed as the mean ± SEM (n = 3 samples). * Significant difference compared to the solvent control (p < 0.05).

M. Naderi et al. / Aquatic Toxicology 148 (2014) 195–203

were transferred to clean water (X2 -test, X2 = 9.8 and F4 = 64.98, p = 0.001; Fig. 4b and c). Sperm count in the groups that received two highest concentrations of BPS was significantly lower than the solvent control group (Dunnett’s test: F4 = 7.46, p = 0.005; Fig. 4d), while no significant differences between fish exposed to 0.1 and 1 ␮g/l of BPS were found relative to the solvent controls (p = 0.555 and 0.436). 4. Discussion Increasing concerns and passing subsequent restrictions and regulations on BPA production and use has led to the development of several potential chemical replacements for BPA, such as BPS. Although the estrogenic potential of this compound has been demonstrated in vitro (Kuruto-Niwa et al., 2005; Grignard et al., 2012), given the few years of its introduction to global markets, the impact of BPS on fish reproduction has remained largely unknown. Besides, it has been widely indicated that early life stages of fish development are highly sensitive to toxicants in comparison with other life stages (Kime, 1998; Hutchinson et al., 1998; Luckenbach et al., 2001; Voelker et al., 2007). Therefore, exposure of fish embryos and larvae to even very low concentrations of weak estrogen mimics like BPS may lead to a variety of physiological disorders much later in life. The results of present study clearly showed that embryonic exposure to BPS can contribute to developmental toxicity, hormonal imbalance and reproduction failure in adult zebrafish. Developmental exposure of zebrafish to different concentrations of BPS led to decreased survival rate in the group exposed to 100 ␮g/l. In a recent study, however, Ji et al. (2013) have reported no mortality for adult zebrafish exposed to different concentrations of BPS (0.5–50 ␮g/l). Therefore, our result clearly confirmed higher sensitivity of embryos and larvae to toxic effects of xenobiotic compounds. The other conspicuous finding in zebrafish exposed to BPS (F0 ) was a skewed sex ratio towards more females. Several studies have shown that exposure of teleost to estrogenic chemicals can lead to shifting of the sex ratio in favor of females (Länge et al., 2001; Hill and Janz, 2003; Nash et al., 2004; Zhong et al., 2005; Zha et al., 2008a). In addition, female-skewed sex ratios in zebrafish and/or intersex condition in medaka (Oryzias latipes) and carp (Cyprinus carpio) have been reported following exposure to BPA (Kang et al., 2002; Mandich et al., 2007; Crain et al., 2007). It is well known that sex steroid hormones (T and E2) and their balance play important roles in sex differentiation, determination and maturation of teleost (Baroiller et al., 1999; Devlin and Nagahama, 2002; Fenske and Segner, 2004). Therefore, altered steroid levels obviously influence sexual differentiation in fish. Zebrafish is considered as a hermaphrodite species with protogynic gonads, which later develop into ovaries or testes approximately 5–7 weeks post hatching (Maack and Segner, 2003; Spence et al., 2008). During this crucial period, disruption of sex steroid hormonal balance may impair or change the gonadal sex differentiation. In our experiment, increased levels of E2 was observed for both BPS treated males and females, while plasma T levels showed a significant reduction in males. This could be a possible explanation for altered sex ratio in favor of females. Furthermore, BPS has shown the ability to act as an estrogen mimic that binds to estrogen receptors and subsequently exerts estrogenic activity. Therefore, this chemical may be potent enough to influence the sexual differentiation of zebrafish. The findings of our study also suggest that long-term BPS exposure contributes to decreased growth and GSI, and increased HSI in zebrafish. There were significant differences in both total length and body weight of male zebrafish at 100 ␮g/l of BPS compared to those of the solvent controls. In this regard, there are conclusive evidences that natural and synthetic estrogens affect the growth of fish. Yokota et al. (2000) reported the growth

199

suppression in Japanese medaka (O. latipes) exposed to BPA during the early life stage from fertilized eggs to 60 dpf. In another study, Aluru et al. (2010) reported that exposure of rainbow trout (Oncorhynchus mykiss) oocytes to BPA resulted in growth suppression in adults. The authors suggested that BPA through interfering with somatotropic axis (growth hormone and insulin-like growth factors) culminated in the growth impairment. Furthermore, there are other pathways by which xenoestrogens like BPS may be involved in growth suppression. Disruption of thyroid hormones, as clearly observed in our study, could be a possible reason. Several estrogenic compounds like BPA have been shown to interfere with secretion of T3 and T4, which play key roles in the growth enhancement of fish (Yokota et al., 2000; Davis et al., 2007; Aluru et al., 2010). Vitellogenesis may be another causative factor in the growth deficiency. This process requires lots of energy resources that may lead to a shift in energy utilization from somatic growth to VTG production. Therefore, the growth suppression of male zebrafish might be due to estrogenic activity of BPS. Both GSI and HSI are considered as common indicators of exposure to environmental contaminants, which are widely used as unspecific biomarkers to examine possible adverse effects of EDCs accompanied by other more specific biomarkers (Verslycke et al., 2002; Kleinkauf et al., 2004; Hutchinson et al., 2006; Du et al., 2009). GSI and HSI alterations have been reported in several fish species exposed to xenoestrogens such as E2, 17␣-ethynylestradiol, nonylphenol, and octylphenol (Mills et al., 2001; Van den Belt et al., 2004; Brion et al., 2004). In a recent study conducted in zebrafish, no significant effect of BPS on HSI in males or females has been found, but GSI value exhibited a significant reduction after 21 days of exposure (Ji et al., 2013). Previously, it has been documented that estrogenic EDCs caused reductions in GSI by alternation in the numbers and sizes of Sertoli cells and germ cells (Miles-Richardson et al., 1999; Kinnberg et al., 2000; Yang et al., 2006). Moreover, it has been proposed that VTG production due to estrogenic effect of xenobiotics could inhibit or disturb gonadal development in female fish (Scholz and Gutzeit, 2000; Versonnen et al., 2003). VTG production can also lead to hyperplasia or hypertrophy of hepatocyts (Arukwe and Goksøyr, 1998; Zha et al., 2008b), which may be responsible for elevated values of HSI in this study. In the present study, an evaluation of actual concentrations in the exposure medium limits our ability to draw conclusions about the effects of exact concentrations of BPS, nevertheless our results still provide support for the fact that relative concentration differences do have an impact. Further, the precise mechanism(s) which may be involved in alterations of plasma sex steroid hormones in zebrafish was not elucidated. However, previous studies have suggested several different direct and indirect mechanisms of action. Xenoestrogens have shown to affect the activity of steroidogenic and steroid metabolizing enzymes both in vivo and in vitro (Kishida et al., 2001; Vaccaro et al., 2005; Cionna et al., 2006; Kortner and Arukwe, 2007). There also several reports regarding indirect effects of estrogenic chemicals associated with altered HPG feedback loops (Harris et al., 2001; Mills and Chichester, 2005). It has been demonstrated that BPA have an up-regulatory effect on expression of aromatase (cyp19b) mRNA in early zebrafish embryos (Chung et al., 2011). Zebrafish has two distinct types of aromatase genes, enzyme responsible for conversation of T to E2, which are expressed in its gonad (CYP19A) and brain (cyp19b). Ji et al. (2013) have reported that short-term exposure of zebrafish to BPS caused significant increased and decreased in plasma levels of E2 and T, respectively. The observed alterations were along with up-regulation of cyp19a, and down-regulation of cytochrome P450 17A and 17␤Hydroxysteroid dehydrogenases transcripts. Two later enzymes are involved in biosynthesis of T. As a result, it is highly likely that in our study BPS affected the synthesis of steroid hormones in a similar manner.

200

M. Naderi et al. / Aquatic Toxicology 148 (2014) 195–203

VTG, a precursor to egg yolk protein, is normally produced in the liver of female oviparous vertebrates under the influence of circulating endogenous estrogen, released to the blood stream, taken up and modified by developing eggs. This protein is normally undetectable in the plasma of immature and male oviparous animals. However, since males and juveniles also possess the VTG gene their exposure to environmental estrogens or estrogen mimics can trigger expression of the gene, subsequently leading to VTG production (Kime, 1999; Ankley and Johnson, 2004). Measurement of VTG levels in males or immature fish is one of the most widely used biomarkers for exposure to estrogenic chemicals in the aquatic environment (Jin et al., 2008; Oehlmann et al., 2009). In addition, it has been confirmed that induction of VTG in males and juveniles can act as the indicator of reproductive disruption (Kime et al., 1999: Mills et al., 2003; Lin and Janz, 2006). BPA, owing to its estrogenic properties and via binding to specific estrogen receptors has shown to induce synthesis of VTG in different fish species (Ishibashi et al., 2005; Sohoni et al., 2001; Brian et al., 2007; Mandich et al., 2007; Correia et al., 2007; Hatef et al., 2012). BPS is also known to have estrogenic potential; therefore it is possible that BPS through exerting estrogenic effects triggered vitellogenic response in zebrafish. Besides, increased level of E2 after exposure to BPS could another explanation for enhanced concentrations of plasma VTG in the present study. This mechanism of action has previously been reported for other xenoestrogens like nonylphenol (Harris et al., 2001). Thyroid hormones are known to be involved in a wide range of essential physiological functions including embryonic growth and development, metamorphosis, osmoregulation, sexual differentiation, reproduction, and behavior (Cyr and Eales, 1996; Yen, 2001; Liu and Chan, 2002; Thomas, 2008; Jugan et al., 2010). In fish, it has been noted that functions of thyroidal states are susceptible to disruption by different EDCs (Lema et al., 2008; Shi et al., 2009; Li et al., 2009; Liu et al., 2011; Naderi et al., 2013; Wang et al., 2012; Yu et al., 2013). Estrogenic chemicals have shown the ability to interfere with the thyroid system by a direct impact on the synthesis of thyroid hormones. Inhibited thyroid peroxidase (TPO) activity has been reported in vitro, following exposure to BPA and nonylphenol (Schmutzler et al., 2004; Ghisari and Bonefeld-Jorgensen, 2005; Schmutzler et al., 2007). TPO is a key enzyme in the synthesis of both T4 and T3 (Dunn and Dunn, 2001; Thomas, 2008). Correspondingly, impaired TPO activity can lead to reduction of thyroid hormones, T4 in particular. Since in fish, thyroid follicles secrete primarily T4, which is then converted to biologically active T3 almost entirely in peripheral tissues, the reduction in plasma T4 levels is contributed to decrease in T3 formation. Deiodinases (Dio) are responsible enzymes for conversion of T4 to T3 by removing iodine from the outer ring of T4. There are three types of deiodinase enzymes (Dio1, Dio2, and Dio3) found in fish (Orozco and Valverde, 2005). Dio1 and Dio2 are known to be involved in converting T4 into T3. It has been demonstrated that Dio1 had a considerable influence on iodine recovery and thyroid hormones degradation (Van der Geyten et al., 2005). Dio2 plays a pivotal role in producing active T3 (Van der Geyten et al., 2005; Yu et al., 2010), and Dio3 is a purely inactivating enzyme (Orozco and Valverde, 2005). Previous studies have revealed that Dio activity in fish is sensitive to environmental contaminants. For example, Wei et al. (2008) have reported that PFOA (Perfluorooctane sulfonate) exposure significantly suppressed the expression of Dio2 in rare minnows (Gobiocypris rarus).Yu et al. (2010) also revealed significant up-regulation of Dio1 and Dio2 gene transcription in zebrafish larvae after exposure to DE-71. Up-regulated Dio1 and Dio2 genes expression associated with decreased T4 and increased T3 was also found in zebrafish embryos exposed to xenobiotic chemicals including BDE-209 (decabromodiphenyl ether); TDCPP (Tris(1,3dichloro-2-propyl) phosphate), hexaconazole and tebuconazole

(Chen et al., 2012; Yu et al., 2013; Wang et al., 2012). Accordingly, another plausible explanation for thyroid hormone alterations in our study would at least partly be associated with a decrease in the transcription of Dio2 (responsible for the reduction in deiodination of T4 into T3), and increased Dio1 activity (responsible for the decreased plasma T3 and T4). However, further studies are required to clarify the mechanisms involved in decreased plasma T3 and T4 levels observed in this study. Reproduction is a complex process that relies upon the appropriate interaction of multiple factors at the levels of the hypothalamus–pituitary–gonad (HPG) axis and the lower genital tract. As a result, any external factor that interacts with and interrupt this axis can have far-reaching and adverse consequences for reproduction (Hutchinson et al., 2006; Brian et al., 2007; Thomas, 2008). Despite the fact that zebrafish colonies spawned in clean water, the results indicated that the egg production in the groups exposed to 10 and 100 ␮g/l of BPS were significantly decreased. This reduction in the egg production was accompanied with the depression in sperm count of males treated with highest doses of BPS. Reduced egg production was also observed in adult zebrafish, when exposed to BPS for a period of 21 days (Ji et al., 2013). BPA has also shown to reduce egg production, sperm quality and count in several fish species (Haubruge et al., 2000; Sohoni et al., 2001; Lahnsteiner et al., 2005). The inhibitory effects on egg production and sperm count in fish have also reported associated with different estrogenic compounds (Vom Saal et al., 1998; Van den Belt et al., 2004; Lin and Janz, 2006; Santos et al., 2007; Robinson et al., 2007; Coe et al., 2008; Johnson et al., 2008; Liu et al., 2010). In addition to sex differentiation and sexual maturation, sex steroids influence quality and quantity of both eggs and sperm (Liley and Stacey, 1983; Devlin and Nagahama, 2002; Bobe and Labbé, 2010; Lubzens et al., 2010). Therefore, any disturbance in balance of sex steroids by which other parts of the HPG axis may be affected, subsequently leads to reproductive dysfunction (Brian et al., 2007; Xi et al., 2011; Naderi et al., 2013). Our results that showed the altered plasma levels of E2 and T were accompanied by reductions in normal growth of gonads, egg production and sperm counts can support this hypothesis. In our study, the delayed and reduced hatching rates were also observed in F1 generation. Similar results have been observed in zebrafish, rainbow trout (O. mykiss), and medaka (O. latipes) treated with BPA (Ishibashi et al., 2005; Aluru et al., 2010; McCormick et al., 2010). Moreover, the reduced hatchability and time to hatching were reported in zebrafish that have been undergoing parental exposure to environmentally relevant concentrations of BPS (Ji et al., 2013). Our results are well-consistent with these findings, further suggesting that BPS would impair the reproductive success in adult fish, which have experienced developmental exposure to this chemical. In summary, the data presented here demonstrated that developmental exposure of zebrafish to BPS resulted in developmental toxicity, reproduction impairments, imbalance of steroid and thyroid hormones, and induction of VTG in males, which obviously highlighted estrogenic activity of this compound. Taking into account the growing interest in application of BPS in a variety of products, further studies are required to clarify the precise mechanisms of BPS by which the endocrine system in animals might be affected. Acknowledgement The authors thank Mr. Pasha Zanoosi for valuable comments and suggestion on the manuscript. References Aluru, N., Leatherland, J.F., Vijayan, M.M., 2010. Bisphenol A in oocytes leads to growth suppression and altered stress performance in juvenile rainbow trout. PloS One 5, e10741.

M. Naderi et al. / Aquatic Toxicology 148 (2014) 195–203 Ankley, G.T., Johnson, R.D., 2004. Small fish models for identifying and assessing the effects of endocrine-disrupting chemicals. Ilar Journal 45, 469–483. Arukwe, A., Goksøyr, A., 1998. Xenobiotics, xenoestrogens and reproduction disturbances in fish. Sarsia 83, 225–241. Baroiller, J.-F., Guiguen, Y., Fostier, A., 1999. Endocrine and environmental aspects of sex differentiation in fish. Cellular and Molecular Life Sciences—CMLS 55, 910–931. Biedermann, S., Tschudin, P., Grob, K., 2010. Transfer of bisphenol A from thermal printer paper to the skin. Analytical and Bioanalytical Chemistry 398, 571–576. Bobe, J., Labbé, C., 2010. Egg and sperm quality in fish. General and Comparative Endocrinology 165, 535–548. Bondesson, M., Jönsson, J., Pongratz, I., Olea, N., Cravedi, J.-P., Zalko, D., Håkansson, H., Halldin, K., Di Lorenzo, D., Behl, C., 2009. A CASCADE of effects of bisphenol A. Reproductive Toxicology 28, 563–567. Brian, J.V., Harris, C.A., Scholze, M., Kortenkamp, A., Booy, P., Lamoree, M., Pojana, G., Jonkers, N., Marcomini, A., Sumpter, J.P., 2007. Evidence of estrogenic mixture effects on the reproductive performance of fish. Environmental Science and Technology 41, 337–344. Briggs, J.P., 2002. The zebrafish: a new model organism for integrative physiology. American Journal of Physiology—Regulatory, Integrative and Comparative Physiology 282, R3–R9. Brion, F., Tyler, C., Palazzi, X., Laillet, B., Porcher, J., Garric, J., Flammarion, P., 2004. Impacts of 17␤-estradiol, including environmentally relevant concentrations, on reproduction after exposure during embryo-larval-, juvenile-and adult-life stages in zebrafish (Danio rerio). Aquatic Toxicology 68, 193–217. Chen, M.Y., Ike, M., Fujita, M., 2002. Acute toxicity, mutagenicity, and estrogenicity of bisphenol-A and other bisphenols. Environmental Toxicology 17, 80–86. Chen, Q., Yu, L., Yang, L., Zhou, B., 2012. Bioconcentration and metabolism of decabromodiphenyl ether (BDE-209) result in thyroid endocrine disruption in zebrafish larvae. Aquatic Toxicology 110, 141–148. Chung, E., Genco, M.C., Megrelis, L., Ruderman, J.V., 2011. Effects of bisphenol A and triclocarban on brain-specific expression of aromatase in early zebrafish embryos. Proceedings of the National Academy of Sciences 108, 17732–17737. Cionna, C., Maradonna, F., Olivotto, I., Pizzonia, G., Carnevali, O., 2006. Effects of nonylphenol on juveniles and adults in the grey mullet, Liza aurata. Reproductive Toxicology 22, 449–454. Coe, T.S., Hamilton, P.B., Hodgson, D., Paull, G.C., Stevens, J.R., Sumner, K., Tyler, C.R., 2008. An environmental estrogen alters reproductive hierarchies, disrupting sexual selection in group-spawning fish. Environmental Science and Technology 42, 5020–5025. Correia, A.D., Freitas, S., Scholze, M., Goncalves, J.F., Booij, P., Lamoree, M.H., Mananos, E., Reis-Henriques, M.A., 2007. Mixtures of estrogenic chemicals enhance vitellogenic response in sea bass. Environmental Health Perspectives 115, 115–121. Crain, D.A., Eriksen, M., Iguchi, T., Jobling, S., Laufer, H., LeBlanc, G.A., Guillette, L.J., 2007. An ecological assessment of bisphenol-A: evidence from comparative biology. Reproductive Toxicology 24, 225–239. Cyr, D.G., Eales, J., 1996. Interrelationships between thyroidal and reproductive endocrine systems in fish. Reviews in Fish Biology and Fisheries 6, 165–200. Davis, L.K., Hiramatsu, N., Hiramatsu, K., Reading, B.J., Matsubara, T., Hara, A., Sullivan, C.V., Pierce, A.L., Hirano, T., Grau, E.G., 2007. Induction of three vitellogenins by 17␤-estradiol with concurrent inhibition of the growth hormone-insulin-like growth factor 1 axis in a euryhaline teleost, the tilapia (Oreochromis mossambicus). Biology of Reproduction 77, 614–625. Devlin, R.H., Nagahama, Y., 2002. Sex determination and sex differentiation in fish: an overview of genetic, physiological, and environmental influences. Aquaculture 208, 191–364. Diamanti-Kandarakis, E., Bourguignon, J.-P., Giudice, L.C., Hauser, R., Prins, G.S., Soto, A.M., Zoeller, R.T., Gore, A.C., 2009. Endocrine-disrupting chemicals: an Endocrine Society scientific statement. Endocrine Reviews 30, 293–342. Du, Y., Shi, X., Liu, C., Yu, K., Zhou, B., 2009. Chronic effects of water-borne PFOS exposure on growth, survival and hepatotoxicity in zebrafish: a partial life-cycle test. Chemosphere 74, 723–729. Dunn, J.T., Dunn, A.D., 2001. Update on intrathyroidal iodine metabolism. Thyroid 11, 407–414. European Commission., 2011. Bisphenol A: EU ban on baby bottles to enter into force tomorrow. European Union, Brussels, Available at http://europa.eu/rapid/pressrelease IP-11-664 en.htm. Fenske, M., Segner, H., 2004. Fenske, M., and Segner, H., Aromatase modulation alters gonadal differentiation in developing zebrafish (Danio rerio). Aquatic Toxicology 67, 105–126. Flint, S., Markle, T., Thompson, S., Wallace, E., 2012. Bisphenol A exposure, effects, and policy: a wildlife perspective. Journal of Environmental Management 104, 19–34. Gallart-Ayala, H., Moyano, E., Galceran, M., 2011. Analysis of bisphenols in soft drinks by on-line solid phase extraction fast liquid chromatography–tandem mass spectrometry. Analytica Chimica Acta 683, 227–233. Ghisari, M., Bonefeld-Jorgensen, E.C., 2005. Impact of environmental chemicals on the thyroid hormone function in pituitary rat GH3 cells. Molecular and Cellular Endocrinology 244, 31–41. Goodman, J.E., McConnell, E.E., Sipes, I.G., Witorsch, R.J., Slayton, T.M., Yu, C.J., Lewis, A.S., Rhomberg, L.R., 2006. An updated weight of the evidence evaluation of reproductive and developmental effects of low doses of bisphenol A. CRC Critical Reviews in Toxicology 36, 387–457. Grignard, E., Lapenna, S., Bremer, S., 2012. Weak estrogenic transcriptional activities of bisphenol A and bisphenol S. Toxicology in Vitro 26, 727–731.

201

Harris, C.A., Santos, E.M., Janbakhsh, A., Pottinger, T.G., Tyler, C.R., Sumpter, J.P., 2001. Nonylphenol affects gonadotropin levels in the pituitary gland and plasma of female rainbow trout. Environmental Science and Technology 35, 2909–2916. Hashimoto, Y., Moriguchi, Y., Oshima, H., Kawaguchi, M., Miyazaki, K., Nakamura, M., 2001. Measurement of estrogenic activity of chemicals for the development of new dental polymers. Toxicology in Vitro 15, 421–425. Hatef, A., Alavi, S.M.H., Abdulfatah, A., Fontaine, P., Rodina, M., Linhart, O., 2012. Adverse effects of bisphenol A on reproductive physiology in male goldfish at environmentally relevant concentrations. Ecotoxicology and Environmental Safety 76, 56–62. Haubruge, E., Petit, F., Gage, M.J., 2000. Reduced sperm counts in guppies (Poecilia reticulata) following exposure to low levels of tributyltin and bisphenol A. Proceedings of the Royal Society of London. Series B: Biological Sciences 267, 2333–2337. Health Canada, 2009. Government of Canada Acts to Protect Newborns and Infants from Bisphenol A in Polycarbonate Plastic Baby Bottles. Health Canada, Ottawa, ON, Available at http://hc-sc.gc.ca/ahc-asc/media/nrcp/ 2009/2009 106-eng.php. Hill, R.L., Janz, D.M., 2003. Developmental estrogenic exposure in zebrafish (Danio rerio): I. Effects on sex ratio and breeding success. Aquatic Toxicology 63, 417–429. Hutchinson, T.H., Ankley, G.T., Segner, H., Tyler, C.R., 2006. Screening and testing for endocrine disruption in fish—biomarkers as signposts, not traffic lights, in risk assessment. Environmental Health Perspectives 114, 106–114. Hutchinson, T.H., Solbe, J., Kloepper-Sams, P.J., 1998. Analysis of the ecetoc aquatic toxicity (EAT) database III—comparative toxicity of chemical substances to different life stages of aquatic organisms. Chemosphere 36, 129–142. Ishibashi, H., Watanabe, N., Matsumura, N., Hirano, M., Nagao, Y., Shiratsuchi, H., Kohra, S., Yoshihara S.-i. Arizono, K., 2005. Toxicity to early life stages and an estrogenic effect of a bisphenol A metabolite, 4-methyl-2, 4-bis (4hydroxyphenyl) pent-1-ene on the medaka (Oryzias latipes). Life Sciences 77, 2643–2655. Ji, K., Hong, S., Kho, Y., Choi, K., 2013. Effects of bisphenol S exposure on endocrine functions and reproduction of zebrafish. Environmental Science and Technology 47, 8793–8800. Jin, Y., Wang, W., Xu, C., Fu, Z., Liu, W., 2008. Induction of hepatic estrogen-responsive gene transcription by permethrin enantiomers in male adult zebrafish. Aquatic Toxicology 88, 146–152. Johnson, L.L., Lomax, D.P., Myers, M.S., Olson, O.P., Sol, S.Y., O’Neill, S.M., West, J., Collier, T.K., 2008. Xenoestrogen exposure and effects in English sole (Parophrys vetulus) from Puget Sound, WA. Aquatic Toxicology 88, 29–38. Jugan, M.-L., Levi, Y., Blondeau, J.-P., 2010. Endocrine disruptors and thyroid hormone physiology. Biochemical Pharmacology 79, 939–947. Kang, I., Yokota, H., Oshima, Y., Tsuruda, Y., Oe, T., Imada, N., Tadokoro, H., Honjo, T., 2002. Effects of bisphenol A on the reproduction of Japanese medaka (Oryzias latipes). Environmental Toxicology and Chemistry 21, 2394–2400. Kime, D., Nash, J., Scott, A., 1999. Vitellogenesis as a biomarker of reproductive disruption by xenobiotics. Aquaculture 177, 345–352. Kime, D.E., 1998. Disruption in eggs, embryos, larvae and juvenile fish. In: Kime, D.E. (Ed.), Endocrine Disruption in Fish. Kluwer Academic Publishers, Boston, MA, pp. 187–200. Kime, D.E., 1999. A strategy for assessing the effects of xenobiotics on fish reproduction. Science of the Total Environment 225, 3–11. Kinnberg, K., Korsgaard, B., Bjerregaard, P., Jespersen, A.S., 2000. Effects of nonylphenol and 17␤-estradiol on vitellogenin synthesis and testis morphology in male platyfish Xiphophorus maculatus. Journal of Experimental Biology 203, 171–181. Kishida, M., McLellan, M., Miranda, J.A., Callard, G.V., 2001. Estrogen and xenoestrogens upregulate the brain aromatase isoform (P450aromB) and perturb markers of early development in zebrafish (Danio rerio). Comparative Biochemistry and Physiology Part B: Biochemistry and Molecular Biology 129, 261–268. Kitamura, S., Suzuki, T., Sanoh, S., Kohta, R., Jinno, N., Sugihara, K., Yoshihara, S., Fujimoto, N., Watanabe, H., Ohta, S., 2005. Comparative study of the endocrinedisrupting activity of bisphenol A and 19 related compounds. Toxicological Sciences 84, 249–259. Kleinkauf, A., Connor, L., Swarbreck, D., Levene, C., Walker, P., Johnson, P.J., Leah, R.T., 2004. General condition biomarkers in relation to contaminant burden in European flounder (Platichthys flesus). Ecotoxicology and Environmental Safety 58, 335–355. Kortner, T.M., Arukwe, A., 2007. The xenoestrogen, 4-nonylphenol, impaired steroidogenesis in previtellogenic oocyte culture of Atlantic cod (Gadus morhua) by targeting the StAR protein and P450scc expressions. General and Comparative Endocrinology 150, 419–429. Kuruto-Niwa, R., Nozawa, R., Miyakoshi, T., Shiozawa, T., Terao, Y., 2005. Estrogenic activity of alkylphenols, bisphenol S, and their chlorinated derivatives using a GFP expression system. Environmental Toxicology and Pharmacology 19, 121–130. Lahnsteiner, F., Berger, B., Kletzl, M., Weismann, T., 2005. Effect of bisphenol A on maturation and quality of semen and eggs in the brown trout, Salmo trutta f. fario. Aquatic Toxicology 75, 213–224. Länge, R., Hutchinson, T.H., Croudace, C.P., Siegmund, F., Schweinfurth, H., Hampe, P., Panter, G.H., Sumpter, J.P., 2001. Effects of the synthetic estrogen 17␣ethinylestradiol on the life-cycle of the fathead minnow (Pimephales promelas). Environmental Toxicology and Chemistry 20, 1216–1227. Larsen, M.G., Baatrup, E., 2010. Functional behavior and reproduction in androgenic sex reversed zebrafish (Danio rerio). Environmental Toxicology and Chemistry 29, 1828–1833.

202

M. Naderi et al. / Aquatic Toxicology 148 (2014) 195–203

LeBlanc, G.A., 2007. Crustacean endocrine toxicology: a review. Ecotoxicology 16, 61–81. Lema, S.C., Dickey, J.T., Schultz, I.R., Swanson, P., 2008. Dietary exposure to 2,2 ,4,4 -tetrabromodiphenyl ether (PBDE-47) alters thyroid status and thyroid hormone—regulated gene transcription in the pituitary and brain. Environmental Health Perspectives 116, 1694. Li, D.-K., Zhou, Z., Miao, M., He, Y., Wang, J., Ferber, J., Herrinton, L.J., Gao, E., Yuan, W., 2011. Urine bisphenol-A (BPA) level in relation to semen quality. Fertility and Sterility 95 (625-630), e624. Li, W., Zha, J., Spear, P.A., Li, Z., Yang, L., Wang, Z., 2009. Changes of thyroid hormone levels and related gene expression in Chinese rare minnow (Gobiocypris rarus) during 3-amino-1,2,4-triazole exposure and recovery. Aquatic Toxicology 92, 50–57. Liao, C., Liu, F., Kannan, K., 2012. Bisphenol S, a new bisphenol analogue, in paper products and currency bills and its association with bisphenol a residues. Environmental Science and Technology 46, 6515–6522. Liley, N., Stacey, N., 1983. Hormones, pheromones, and reproductive behavior in fish. In: Hoar, W.S., Randall, D.J., Donaldson, E.M. (Eds.), Fish Physiology, 4B. Academic Press, New York, NY, pp. 1–63. Lin, L.L., Janz, D.M., 2006. Effects of binary mixtures of xenoestrogens on gonadal development and reproduction in zebrafish. Aquatic Toxicology 80, 382–395. Liu, Y.W., Chan, W.K., 2002. Thyroid hormones are important for embryonic to larval transitory phase in zebrafish. Differentiation 70, 36–45. Liu, C., Deng, J., Yu, L., Ramesh, M., Zhou, B., 2010. Endocrine disruption and reproductive impairment in zebrafish by exposure to 8:2 fluorotelomer alcohol. Aquatic Toxicology 96, 70–76. Liu, S., Chang, J., Zhao, Y., Zhu, G., 2011. Changes of thyroid hormone levels and related gene expression in zebrafish on early life stage exposure to triadimefon. Environmental Toxicology and Pharmacology 32, 472–477. Lubzens, E., Young, G., Bobe, J., Cerdà, J., 2010. Oogenesis in teleosts: how fish eggs are formed. General and Comparative Endocrinology 165, 367–389. Luckenbach, T., Triebskorn, R., Müller, E., Oberemm, A., 2001. Toxicity of waters from two streams to early life stages of brown trout (Salmo trutta f.fario L.), tested under semi-field conditions. Chemosphere 45, 571–579. Maack, G., Segner, H., 2003. Morphological development of the gonads in zebrafish. Journal of Fish Biology 62, 895–906. Mandich, A., Bottero, S., Benfenati, E., Cevasco, A., Erratico, C., Maggioni, S., Massari, A., Pedemonte, F., Vigano, L., 2007. In vivo exposure of carp to graded concentrations of bisphenol A. General and Comparative Endocrinology 153, 15–24. McCormick, J.M., Paiva, M.S., Häggblom, M.M., Cooper, K.R., White, L.A., 2010. Embryonic exposure to tetrabromobisphenol A and its metabolites, bisphenol A and tetrabromobisphenol A dimethyl ether disrupts normal zebrafish (Danio rerio) development and matrix metalloproteinase expression. Aquatic Toxicology 100, 255–262. McLachlan, J.A., 2001. Environmental signaling: what embryos and evolution teach us about endocrine disrupting chemicals. Endocrine Reviews 22, 319–341. Miles-Richardson, S., Pierens, S., Nichols, K., Kramer, V., Snyder, E., Snyder, S., Render, J., Fitzgerald, S., Giesy, J., 1999. Effects of waterborne exposure to 4-nonylphenol and nonylphenol ethoxylate on secondary sex characteristics and gonads of fathead minnows (Pimephales promelas). Environmental Research 80, S122–S137. Mills, L.J., Gutjahr-Gobell, R.E., Haebler, R.A., Borsay Horowitz, D.J., Jayaraman, S., Pruell, R.J., McKinney, R.A., Gardner, G.R., Zaroogian, G.E., 2001. Effects of estrogenic (o,p-DDT; octylphenol) and anti-androgenic (p,p -DDE) chemicals on indicators of endocrine status in juvenile male summer flounder (Paralichthys dentatus). Aquatic Toxicology 52, 157–176. Mills, L.J., Gutjahr-Gobell, R.E., Horowitz, D.B., Denslow, N.D., Chow, M.C., Zaroogian, G.E., 2003. Relationship between reproductive success and male plasma vitellogenin concentrations in cunner, Tautogolabrus adspersus. Environmental Health Perspectives 111, 93–100. Mills, L.J., Chichester, C., 2005. Review of evidenCE: are endocrine-disrupting chemicals in the aquatic environment impacting fish populations? Science of the Total Environment 343, 1–34. Naderi, M., Zargham, D., Asadi, A., Bashti, T., Kamayi, K., 2013. Short-term responses of selected endocrine parameters in juvenile rainbow trout (Oncorhynchus mykiss) exposed to 4-nonylphenol. Toxicology and Industrial Health, http://dx.doi.org/10.1177/0748233713491806, Epub ahead of print 14 June 2013. Nash, J.P., Kime, D.E., Van der Ven, L.T., Wester, P.W., Brion, F., Maack, G., Stahlschmidt-Allner, P., Tyler, C.R., 2004. Long-term exposure to environmental concentrations of the pharmaceutical ethynylestradiol causes reproductive failure in fish. Environmental Health Perspectives 112, 1725–1733. Oehlmann, J., Schulte-Oehlmann, U., Kloas, W., Jagnytsch, O., Lutz, I., Kusk, K.O., Wollenberger, L., Santos, E.M., Paull, G.C., Van Look, K.J.W., Tyler, C.R., 2009. A critical analysis of the biological impacts of plasticizers on wildlife. Philosophical Transactions of the Royal Society B: Biological Sciences 364, 2047–2062. Orozco, A., Valverde-R, C., 2005. Thyroid hormone deiodination in fish. Thyroid 15, 799–813. Paull, G.C., Van Look, K.J., Santos, E.M., Filby, A.L., Gray, D.M., Nash, J.P., Tyler, C.R., 2008. Variability in measures of reproductive success in laboratory-kept colonies of zebrafish and implications for studies addressing population-level effects of environmental chemicals. Aquatic Toxicology 87, 115–126. Power, D., Llewellyn, L., Faustino, M., Nowell, M., Björnsson, B.T., Einarsdottir, I., Canario, A., Sweeney, G., 2001. Thyroid hormones in growth and development of fish. Comparative Biochemistry and Physiology Part C: Toxicology and Pharmacology 130, 447–459.

Richter, C.A., Birnbaum, L.S., Farabollini, F., Newbold, R.R., Rubin, B.S., Talsness, C.E., Vandenbergh, J.G., Walser-Kuntz, D.R., vom Saal, F.S., 2007. In vivo effects of bisphenol A in laboratory rodent studies. Reproductive Toxicology 24, 199–224. Robinson, C.D., Brown, E., Craft, J.A., Davies, I.M., Megginson, C., Miller, C., Moffat, C.F., 2007. Bioindicators and reproductive effects of prolonged 17␤-oestradiol exposure in a marine fish, the sand goby (Pomatoschistus minutus). Aquatic Toxicology 81, 397–408. Rubin, B.S., 2011. Bisphenol A: an endocrine disruptor with widespread exposure and multiple effects. Journal of Steroid Biochemistry and Molecular Biology 127, 27–34. Santos, E.M., Paull, G.C., Van Look, K.J., Workman, V.L., Holt, W.V., Van Aerle, R., Kille, P., Tyler, C.R., 2007. Gonadal transcriptome responses and physiological consequences of exposure to oestrogen in breeding zebrafish (Danio rerio). Aquatic Toxicology 83, 134–142. Schmutzler, C., Bacinski, A., Gotthardt, I., Huhne, K., Ambrugger, P., Klammer, H., Schlecht, C., Hoang-Vu, C., Grüters, A., Wuttke, W., 2007. The ultraviolet filter benzophenone 2 interferes with the thyroid hormone axis in rats and is a potent in vitro inhibitor of human recombinant thyroid peroxidase. Endocrinology 148, 2835–2844. Schmutzler, C., Hamann, I., Hofmann, P.J., Kovacs, G., Stemmler, L., Mentrup, B., Schomburg, L., Ambrugger, P., Grüters, A., Seidlova-Wuttke, D., 2004. Endocrine active compounds affect thyrotropin and thyroid hormone levels in serum as well as endpoints of thyroid hormone action in liver, heart and kidney. Toxicology 205, 95–102. Scholz, S., Gutzeit, H., 2000. 17␣-ethinylestradiol affects reproduction, sexual differentiation and aromatase gene expression of the medaka (Oryzias latipes). Aquatic Toxicology 50, 363–373. Schug, T.T., Janesick, A., Blumberg, B., Heindel, J.J., 2011. Endocrine disrupting chemicals and disease susceptibility. Journal of Steroid Biochemistry and Molecular Biology 127, 204–215. Shi, X., Du, Y., Lam, P.K., Wu, R.S., Zhou, B., 2008. Developmental toxicity and alteration of gene expression in zebrafish embryos exposed to PFOS. Toxicology and Applied Pharmacology 230, 23–32. Shi, X., Liu, C., Wu, G., Zhou, B., 2009. Waterborne exposure to PFOS causes disruption of the hypothalamus–pituitary–thyroid axis in zebrafish larvae. Chemosphere 77, 1010–1018. Sohoni, P., Tyler, C., Hurd, K., Caunter, J., Hetheridge, M., Williams, T., Woods, C., Evans, M., Toy, R., Gargas, M., 2001. Reproductive effects of long-term exposure to bisphenol A in the fathead minnow (Pimephales promelas). Environmental Science and Technology 35, 2917–2925. Spence, R., Gerlach, G., Lawrence, C., Smith, C., 2008. The behaviour and ecology of the zebrafish, Danio rerio. Biological Reviews 83, 13–34. Staples, C.A., Dome, P.B., Klecka, G.M., Oblock, S.T., Harris, L.R., 1998. A review of the environmental fate, effects, and exposures of bisphenol A. Chemosphere 36, 2149–2173. Stegeman, J.J., Goldstone, J.V., Hahn, E., 2010. Perspectives on zebrafish as a model in environmental toxicology. In: Perry, S.F., Farrell, A.P., Brauner, C.J. (Eds.), Fish Physiology, 28. Academic Press, New York, NY, pp. 367–439. Thomas, P., 2008. The endocrine system. In: Di Giulio, R.T., Hinton, D.E. (Eds.), The Toxicology of Fishes. Taylor and Francis, New York, NY, pp. 457–478. Tsai, W.-T., 2006. Human health risk on environmental exposure to bisphenol-A: a review. Journal of Environmental Science and Health Part C: Environmental Carcinogenesis Reviews 24, 225–255. US Food and Drug Administration, 2010. Update on Bisphenol A for Use in Food Contact Applications. US Food and Drug Administration, Washington DC, Available at http://www.fda.gov/newsevents/publichealthfocus/ucm064437.htm. Vajda, A.M., Norris, D.O., 2011. Endocrine-active Chemicals (EACs) in Fishes. In: David Norris, D.O., Lopez, K.H. (Eds.), Hormones and Reproduction of Vertebrates, 1. Academic Press, San Diego, pp. 245–264. Vaccaro, E., Meucci, V., Intorre, L., Soldani, G., Di Bello, D., Longo, V., Gervasi, P.G., Pretti, C., 2005. Effects of 17␤-estradiol, 4-nonylphenol and PCB 126 on the estrogenic activity and phase 1 and 2 biotransformation enzymes in male sea bass (Dicentrarchus labrax). Aquatic Toxicology 75, 293–305. van Aerle, R., Pounds, N., Hutchinson, T.H., Maddix, S., Tyler, C.R., 2002. Window of sensitivity for the estrogenic effects of ethinylestradiol in early life-stages of fathead minnow, Pimephales promelas. Ecotoxicology 11, 423–434. Van den Belt, K., Berckmans, P., Vangenechten, C., Verheyen, R., Witters, H., 2004. Comparative study on the in vitro/in vivo estrogenic potencies of 17␤estradiol, estrone, 17␣-ethynylestradiol and nonylphenol. Aquatic Toxicology 66, 183–195. Van der Geyten, S., Byamungu, N., Reyns, G., Kühn, E., Darras, V., 2005. Iodothyronine deiodinases and the control of plasma and tissue thyroid hormone levels in hyperthyroid tilapia (Oreochromis niloticus). Journal of Endocrinology 184, 467–479. Vandenberg, L.N., Chahoud, I., Heindel, J.J., Padmanabhan, V., Paumgartten, F.J., Schoenfelder, G., 2010. Urinary, circulating, and tissue biomonitoring studies indicate widespread exposure to bisphenol A. Environmental Health Perspectives 118, 1055. Vandenberg, L.N., Hauser, R., Marcus, M., Olea, N., Welshons, W.V., 2007. Human exposure to bisphenol A (BPA). Reproductive Toxicology 24, 139–177. Vandenberg, L.N., Maffini, M.V., Sonnenschein, C., Rubin, B.S., Soto, A.M., 2009. Bisphenol-A and the great divide: a review of controversies in the field of endocrine disruption. Endocrine Reviews 30, 75–95. Verslycke, T., Vandenbergh, G.F., Versonnen, B., Arijs, K., Janssen, C.R., 2002. Induction of vitellogenesis in 17␣-ethinylestradiol-exposed rainbow trout

M. Naderi et al. / Aquatic Toxicology 148 (2014) 195–203 (Oncorhynchus mykiss): a method comparison. Comparative Biochemistry and Physiology Part C: Toxicology and Pharmacology 132, 483–492. Versonnen, B.J., Arijs, K., Verslycke, T., Lema, W., Janssen, C.R., 2003. In vitro and in vivo estrogenicity and toxicity of o-, m-, and p-dichlorobenzene. Environmental Toxicology and Chemistry 22, 329–335. ˜ Vinas, P., Campillo, N., Martínez-Castillo, N., Hernández-Córdoba, M., 2010. Comparison of two derivatization-based methods for solid-phase microextraction–gas chromatography–mass spectrometric determination of bisphenol A, bisphenol S and biphenol migrated from food cans. Analytical and Bioanalytical Chemistry 397, 115–125. Voelker, D., Vess, C., Tillmann, M., Nagel, R., Otto, G.W., Geisler, R., Schirmer, K., Scholz, S., 2007. Differential gene expression as a toxicant-sensitive endpoint in zebrafish embryos and larvae. Aquatic Toxicology 81, 355–364. Vom Saal, F.S., Cooke, P.S., Buchanan, D.L., Palanza, P., Thayer, K.A., Nagel, S.C., Parmigiani, S., Welshons, W.V., 1998. A physiologically based approach to the study of bisphenol A and other estrogenic chemicals on the size of reproductive organs, daily sperm production, and behavior. Toxicology and Industrial Health 14, 239–260. Wang, Q., Liang, K., Liu, J., Yang, L., Guo, Y., Liu, C., Zhou, B., 2012. Exposure of zebrafish embryos/larvae to TDCPP alters concentrations of thyroid hormones and transcriptions of genes involved in the hypothalamic-pituitary-thyroid axis. Aquatic Toxicology, http://dx.doi.org/10.1016/j.aquatox.2012.11.009, Epub ahead of print. 19 Nov 2012. Wei, Y., Liu, Y., Wang, J., Tao, Y., Dai, J., 2008. Toxicogenomic analysis of the hepatic effects of perfluorooctanoic acid on rare minnows (Gobiocypris rarus). Toxicology and Applied Pharmacology 226, 285–297. Xi, W., Lee, C., Yeung, W., Giesy, J.P., Wong, M., Zhang, X., Hecker, M., Wong, C.K., 2011. Effect of perinatal and postnatal bisphenol A exposure to the regulatory circuits at the hypothalamus–pituitary–gonadal axis of CD-1 mice. Reproductive Toxicology 31, 409–417.

203

Yang, F.X., Xu, Y., Hui, Y., 2006. Reproductive effects of prenatal exposure to nonylphenol on zebrafish (Danio rerio). Comparative Biochemistry and Physiology Part C: Toxicology and Pharmacology 142, 77–84. Yen, P.M., 2001. Physiological and molecular basis of thyroid hormone action. Physiological Reviews 81, 1097–1142. Yokota, H., Tsuruda, Y., Maeda, M., Oshima, Y., Tadokoro, H., Nakazono, A., Honjo, T., Kobayashi, K., 2000. Effect of bisphenol A on the early life stage in Japanese medaka (Oryzias latipes). Environmental Toxicology and Chemistry 19, 1925–1930. Yu, L., Chen, M., Liu, Y., Gui, W., Zhu, G., 2013. Thyroid endocrine disruption in zebrafish larvae following exposure to hexaconazole and tebuconazole. Aquatic Toxicology 139, 35–42. Yu, L., Deng, J., Shi, X., Liu, C., Yu, K., Zhou, B., 2010. Exposure to DE-71 alters thyroid hormone levels and gene transcription in the hypothalamic–pituitary–thyroid axis of zebrafish larvae. Aquatic Toxicology 97, 226–233. Zha, J., Sun, L., Spear, P.A., Wang, Z., 2008a. Comparison of ethinylestradiol and nonylphenol effects on reproduction of Chinese rare minnows (Gobiocypris rarus). Ecotoxicology and Environmental Safety 71, 390–399. Zha, J., Sun, L., Zhou, Y., Spear, P.A., Ma, M., Wang, Z., 2008b. Assessment of 17␣-ethinylestradiol effects and underlying mechanisms in a continuous, multigeneration exposure of the Chinese rare minnow (Gobiocypris rarus). Toxicology and Applied Pharmacology 226, 298–308. Zhong, X., Xu, Y., Liang, Y., Liao, T., Wang, J., 2005. The Chinese rare minnow (Gobiocypris rarus) as an in vivo model for endocrine disruption in freshwater teleosts: a full life-cycle test with diethylstilbestrol. Aquatic Toxicology 71, 85–95. Zou, E., 2010. Aquatic invertebrate endocrine disruption. In: Breed, M.D., Moore, J. (Eds.), Encyclopedia of Animal Behavior, 1. Academic Press, London, pp. 112–123.