Toxic effects of fluridone on early developmental stages of Japanese Medaka (Oryzias latipes)

Science of the Total Environment (2020) 134495

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Science of the Total Environment journal homepage: www.elsevier.com/locate/scitotenv

Toxic effects of fluridone on early developmental stages of Japanese Medaka (Oryzias latipes) Jiali Jin a,b,⇑, Tomofumi Kurobe b, Bruce G. Hammock b, Chelsea H. Lam b, Li Lin c, Swee J. Teh b a

Yangtze River Fisheries Research Institute, Chinese Academy of Fishery Sciences, Wuhan 430223, China Aquatic Health Program, Department of Anatomy, Physiology, and Cell Biology, School of Veterinary Medicine, University of California, Davis, CA 95616, USA c Guangzhou Key Laboratory of Aquatic Animal Diseases and Waterfowl Breeding, College of Animal Sciences and Technology, Zhongkai University of Agriculture and Engineering, Guangzhou, Guangdong 510225, China b

h i g h l i g h t s

g r a p h i c a l a b s t r a c t

 Fluridone is intensively used for

controlling invasive aquatic plants.  Fluridone reduced hatching success of

Medaka embryo in a dose dependent manner.  High levels of fluridone lead to abnormal swimming behavior of larval Medaka.  RNA-seq detected gene expression and biological processes altered by fluridone.  Fluridone induced oxidative stress and endocrine disruption in the larval Medaka.

a r t i c l e

i n f o

Article history: Received 17 June 2019 Received in revised form 11 September 2019 Accepted 15 September 2019

Keywords: Hatching success Mechanical stimulation test RNA-seq RT-qPCR

a b s t r a c t The herbicide fluridone is intensively applied to control invasive aquatic plants globally, including in the Sacramento and San Joaquin Delta (the Delta), California, USA. Our previous study revealed that the adult stage of Delta Smelt showed acute and sub-lethal adverse effects following 6 h of exposure to environmentally relevant concentrations of fluridone. To further investigate mechanisms of toxicity of fluridone and to assess its toxicity to early life stages of fish, we performed additional exposures using the fish model Japanese Medaka (Oryzias latipes). Male and female Medaka embryos were exposed to concentrations of fluridone for 14 d and showed reduced hatching success in a dose dependent manner. The half maximal effective concentration for the hatching success was 2.3 mg L 1. In addition, male and female Medaka larvae were acute exposed to fluridone for 6 h to assess their swimming behavior and gene expression patterns. Fish exposed to fluridone at 4.2 mg L 1 or higher became lethargic and showed abnormal swimming behavior. The response to the stimuli was significantly impaired by fluridone at 21 mg L 1 and above in males, and at 104 mg L 1 in females. Transcriptome analysis identified a total of 799 genes that were significantly differentially expressed, comprising 555 up-regulated and 244 down-regulated genes in males exposed to 21 mg L 1 of fluridone. The gene set enrichment analysis indicated a number of biological processes altered by fluridone. Among the genes involved in those biological processes, the expression of the genes, acetylcholinesterase, retinoic acid receptor, insulin receptor substrate, glutathione reductase, and glutathione S transferase, exhibited dose- and sex-dependent

⇑ Corresponding author at: Yangtze River Fisheries Research Institute, Chinese Academy of Fishery Sciences, Wuhan 430223, China. E-mail address: [email protected] (J. Jin). https://doi.org/10.1016/j.scitotenv.2019.134495 0048-9697/Ó 2019 Elsevier B.V. All rights reserved.

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J. Jin et al. / Science of the Total Environment (2020) 134495

responses to fluridone. The study indicated that fluridone exposure led to detrimental toxic effects at early developmental stages of fish, by disturbing the biological processes of growth and development, and the nervous system, inducing oxidative stress and endocrine disruption. Ó 2019 Elsevier B.V. All rights reserved.

1. Introduction Excessive growth of nuisance plants in aquatic systems can directly or indirectly compromise local economies or ecosystems by hampering boat traffic, degrading water quality, and altering habitats and food webs (Bremigan et al., 2005; Jacob et al., 2016). Invasive aquatic plants are a serious ecological concern in the major lakes and rivers throughout much of North America. Previous studies reported that native plants have been displaced by invasive aquatic plants, such as Eurasian Watermilfoil (Myriophyllum spicatum), Hydrilla (Hydrilla verticillata L.f.), and Water Hyacinth (Eichhornia crassipes Mart.) (Bremigan et al., 2005; Cheruvelil, 2004; Yi et al., 2011). Herbicide application is a widely used method to control invasive plants worldwide, and one of the most heavily used herbicides is fluridone (Jacob et al., 2016; Kamarianos et al., 1989; Siemering et al., 2008). Fluridone is a slow-acting, systemic herbicide that has been used to control the growth of submerged and emergent aquatic vascular plants since the 1970s (Arnold, 1979; Cahoon et al., 2015; Mccowen et al., 1979; Netherland et al., 2002). Fluridone functions by inhibiting the biosynthesis of carotenoids, pigments that are essential for absorbing light, thus leading to photodegradation of chlorophyll and eventual death of plants (Hamelink et al., 1986; Jacob et al., 2016; Mccowen et al., 1979; Sprecher et al., 1998). The half-life of fluridone in water is relatively long, ranging from 5 to 60 days, with an average of 20 days in aquatic systems after application (Jacob et al., 2016; Paul et al., 1994; West et al., 1983). Nevertheless, fluridone is typically applied multiple times to maintain the target concentration and provide sufficient interaction time to achieve effective treatment. While the persistence of fluridone in aquatic ecosystems makes it an effective herbicide, its persistence may also increase adverse effects on non-target species (Boyd and Tucker, 1998; Cahoon et al., 2015). Fluridone and its metabolite can accumulate in fish body after exposure. For example, fluridone concentrations accumulated in channel catfish was 2 to 9 times greater than that in the water after 60 day’s exposure of different concentrations of fluridone with the highest concentration of 2.0 mg/L (Hamelink et al., 1986). After spraying fluridone at the rate of 0.042 mg/L in pond, fluridone residues in edible parts of carp (Cyprinus carpio L) reached a maximum of 484 lg/kg on the 13th day and 30.7 lg/kg even on the 84th day after application (Kamarianos et al., 1989). In addition, there are several studies reporting toxicity of fluridone to aquatic invertebrates and fishes. For instance, Archambault and Cope (2016) reported that the hatching of freshwater snail eggs (Somatogyrus virginicus) was significantly delayed and reduced by fluridone exposure. Other researchers reported EC50s or LC50s of the tested organisms such as daphnids (Daphnia magna Strauss) (48-h EC50 was 3.6–6.3 mg/L), Rainbow Trout (Saimogairdneri Richardson) (96-h LC50 was 4.2–11.7 mg/L), Walleye (Stizostedion vitreum) (24-h LC50 was 3.6 mg/L) and unionid mussel (Lampsilis siliquoidea) (96-h LC50 was 511 lg/L), or investigated the whole ecological effects (such as plankton biomass and community structure, fish survival and yield) of fluridone in fish ponds or lakes after application (Archambault et al., 2015; Hamelink et al., 1986; Jacob et al., 2016; Paul et al., 1994; Parsons et al., 2009; Yi et al., 2011). However, sublethal effects of

fluridone on non-target aquatic organisms have rarely been assessed. Our laboratory previously examined the effects of fluridone on adult Delta Smelt (Hypomesus transpacificus), an endemic fish species in the Sacramento and San Joaquin River Delta (the Delta). In the study, we monitored swimming behavior and assessed representative sublethal endpoints, such as estrogen and total glutathione concentrations in the liver and acetylcholinesterase activity in the brain. Delta Smelt exposed to high concentrations (4.2 mg L 1 and higher) of fluridone showed abnormal swimming performance, such as lethargy and a spiral swimming pattern. In addition, the data from the sublethal endpoints suggested that fluridone disrupted the endocrine system and induced neurotoxicity and oxidative stress by 6 h acute exposure at environmentally relevant concentrations (Jin et al., 2018). Here, we performed a follow-up study on the influence of fluridone on early fish development using the model species, Japanese Medaka (Oryzias latipes). Medaka (Quart strain) has two major advantages for use in exposure studies. It is sexually dimorphic at the embryonic stage at 2 to 4 days post fertilization (dpf), allowing sex specific exposure tests using embryos (Kinoshita et al., 2009; Wada et al., 1998; Wagner et al., 2017). In addition, the whole genome sequence and annotation are available in public databases (http://www.ensembl.org/index.html, https://www.ncbi.nlm.nih.gov), which is essential for transcriptome analysis by RNA-seq. RNA-seq was used for initial screening of DEGs (differentially expressed genes), followed by RT-qPCR for studying dose response for gene of our interest in both sexes. To investigate the mechanisms of toxicity of fluridone in this study, embryonic and larval stage of male and female Medaka were exposed to fluridone, and their hatching success as well as response to mechanical stimulation were assessed. In addition, gene expression analyses (RNA-seq and RT-qPCR) were performed using larval stage of Medaka.

2. Materials and methods 2.1. Fluridone solutions An aqueous suspension formulation of fluridone containing 41.7% active ingredient (SonarÒ AS) was obtained from SePRO Corporation (Carmel, IN, USA). The stock solutions were prepared based on the percent active ingredient in the commercial formulation. Then, two sets of experimental solutions were prepared: the first set of fluridone solutions (0, 0.03, 0.5, 1, 2, and 4 mg L 1) was prepared for the embryo exposure and the second set (0, 0.08, 0.8, 4.2, 21 and 104 mg L 1) was for the larval fish exposure. The experimental solutions were prepared by spiking stock solutions into reconstituted water (containing Ca2+ 13.9 mg L 1, Na+ 26.3 mg L 1, K+ 2.1 mg L 1, and Mg2+ 12.1 mg L 1) that is used for routine Medaka culture in our laboratory (Ramírez-Duarte et al., 2017a). Because of the insolubility of fluridone in water, precipitate existed at the higher concentrations. Fluridone could therefore adhere to the surface of the chorion layer of Medaka embryos and thus lead to unrealistically low hatching rates. Therefore, a relatively narrow range of fluridone concentrations which did not form precipitate (the first set) was used for the embryo test

J. Jin et al. / Science of the Total Environment (2020) 134495

in the present study. The second set of fluridone concentrations (the wider range) was used for acute exposure to larval Japanese Medaka, which were not expected to cause significant mortality of the fish based on our previous study (Jin et al., 2018). The water quality parameters measured before spiking of fluridone were as follows: pH 7.86, alkalinity 120 mg L 1, hardness 80 mg L 1, conductivity 298.2 lS, salinity 0.2 ppt. Water quality was also measured after spiking fluridone, and the measurements were identical to the measurements before spiking the chemical. Before the initiation of the exposure, water sample of each experimental solution was collected and stored in amber glass vials at 80 °C for chemical analysis. The concentration of fluridone was measured by enzyme-linked immunosorbent assay following the manufacturer’s instructions (Abraxis, Inc, Warminster, PA, USA), which had been described in our previous study (Jin et al., 2018).

2.2. Exposures tests 2.2.1. Embryo exposure test The embryo exposure test was performed in 96-well plates, allowing us to use less experimental solution, have better temperature control, and to allow us to make repeated observations of individual embryos (Truong et al., 2011; Wagner et al., 2017). Japanese Qurt strain Medaka embryos (<6h post fertilization, hpf) were collected from the culture facility at the Aquatic Health Program, University of California, Davis. And the embryos were cleaned and sorted for viability according to the procedure described in Kurobe et al. (2018). Then 50 healthy embryos were placed in each 50 mL Pyrex beakers containing 15 mL of fluridone solution for batch exposure (one beaker per concentration). A 50% (7.5 mL) experimental solution was changed at 2 d post exposure (dpe). At 4 dpe, embryos were separated by sex based on the presence of leucophores on males and then transferred into 96-well plates (Wada et al., 1998) (Costar 9016, Non-sterile, Polystyrene, Flat Bottom, Medium Binding). Embryos were not placed initially in 96well plates to ensure that equal numbers of each sex were exposed, so that unfertilized eggs could be removed, and to conclusively attribute hatching failures to experimental conditions (Wagner et al., 2017). Each well had one embryo placed inside and contained 200 mL of experimental solution. A total of 16 embryos for each sex were used per treatment. Fish embryos were kept in an environmental chamber (Percival Scientific, Perry, IA, USA, Model 136LLVL) at 25 °C on a 16:8h light:dark cycle with a light intensity of 600 Lux during the exposure. More than 80% of the volume in each well was changed every 2 days for the remainder of the experiment. Each embryo was observed daily using a dissecting microscope and recorded for mortality, hatching success, and signs of abnormal development. The experiment was terminated at 14 dpe (4 days in beaker, 10 days in plate), and fish embryos that failed to hatch were counted. A model comparison approach was used to test for an effect of fluridone and sex on hatching success (Burnham and Anderson, 2002, McElreath, 2016). Binomial models were fit to the hatching data in which ‘proportion hatching success’ was the response variable. The models included an intercept model (null), a fluridone model (mg L 1), and a fluridone and sex model. The models were fit using the ‘mle2’ command in the R package ‘bbmle’ (Bolker, 2010; R Core Team, 2018), and compared using Akaike Information Criterion corrected for small sample size (AICc) (Burnham and Anderson, 2002, McElreath, 2016). Variables were considered statistically significant if the 95% confidence interval of the parameter estimate did not overlap zero. The concentration at which 50% hatching success occurred and the 95% confidence interval was calculated based on the top-ranked model by AICc using the methods described in Hammock et al. (2016).

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2.2.2. Larval exposure test Fertilized eggs of Medaka were collected, cleaned, and separated by sex as described above, and then raised until they hatched. Embryos were maintained at 25 °C in the embryo rearing solution, containing 1 g L 1 NaCl, 0.030 g L 1 KCl, 0.040 g L 1 CaCl2
H2O, 80 mg L 1 MgSO4, and 1.0 mg L 1 methylene blue in distilled water (Kurobe et al., 2018). Upon hatching, larvae were transferred into 500 mL beakers and raised in reconstituted water for one week. Larvae were fed three times a day using purified casein-based diet (DeKoven et al., 1992). Larval fish were fasted 24 h prior to the exposures and fasting continued until the end of the tests. Two exposure tests were conducted with larval fish: one for a stimuli response test and the other for gene expression analyses by RNA-seq and RT-qPCR. The larval fish at 7 ± 1 days post hatch (dph) were used for the exposure tests. For the stimuli response test, the male and female larval Medaka were separately exposed to fluridone solution (0, 0.08, 0.8, 4.2, 21 and 104 mg L 1) using 500 mL beakers. Five fish in triplicate were placed in each beaker containing 400 mL of experimental solution. The beakers were continuously aerated and placed in a water bath to maintain the temperature at 25 °C throughout the 6 h exposure. At the end of the exposure, the response of Medaka larvae to a mechanical stimulus (touch) was assessed as described by Stehr et al. (2006). Briefly, fish were transferred to a petri dish with experimental solution and then gently touched on the head with forceps. Fish that swam away were scored as responders after one, two, or three repeated stimuli, while all others were scored as non-responders. The data were analyzed using one-way ANOVA followed by Dunnett’s tests. For gene expression analyses (RNA-seq and RT-qPCR), the exposure was performed as described immediately above, except that 10 larvae were placed in each beaker. The male larval Medaka exposed to the control and the lowest concentration that significantly impaired response to mechanical stimuli (21 mg L 1) were selected for RNA-seq analysis. Only male fish were used for RNAseq because a sign of endocrine disruption (elevated concentrations of hepatic estradiol) was observed in our previous study (Jin et al., 2018). For RT-qPCR, all the fish exposed to the different concentrations of fluridone were used in the analysis. At the end of the 6 h exposure, fish were sacrificed using an overdose of buffered tricaine methane sulfonate (MS222) solution. The larval fish in each beaker were pooled and preserved in TRIZOL reagent (Thermo Fisher Scientific, Waltham, MA, USA) at 80 °C for later sample processing. This study was approved by the Institutional Animal Care and Use Committee (IACUC), University of California, Davis (IACUC Protocol Number: 19604). 2.3. RNA-seq All the details for library preparation, sequencing reaction for RNA-seq, and data analysis are available in our previous publication (Wagner et al., 2017). Total RNA was isolated from control and fluridone exposed groups (21 mg L 1) using TRIZOL Reagent by following the manufacturer’s instructions (Thermo Fisher Scientific). The assessment of total RNA quality, library preparation and sequencing reaction were all performed by the DNA Technologies Core Facility at University of California, Davis (http://dnatech. genomecenter.ucdavis.edu). The samples were sequenced on a HiSeq 2500 instrument (Illumina, San Diego, CA, USA) with single-end 50 bp reads (Wagner et al., 2017). The raw data from sequencing were subjected to trimming by FASTX-Toolkit (http://hannonlab.Cshl.edu/fastx_toolkit/index. html) before mapping to Medaka reference genome sequence using TopHat ver. 2.0.13 (Kim et al., 2013). The Medaka reference genome sequence was obtained from the Ensembl database

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(downloaded on December 18th, 2014, http://www.ensembl.org/ index.html). The data were further processed using Cufflinks, Cuffmerge, and Cuffdiff within Cufflinks ver. 2.2.1 package for transcript assembly, merging, and statistical analysis (Trapnell et al., 2010, 2012). False discovery rate (FDR) lower than 0.05 was set as the threshold to identify the genes that differentially expressed by fluridone exposure. Then the differentially expressed genes (DEGs) were annotated by BLASTX similarity searches against a protein database (nr) obtained from the NCBI GenBank website (downloaded on July 3rd, 2015. Approximately 178 million protein sequences were available, https://www.ncbi.nlm.nih.gov). Gene set enrichment analysis was performed in order to identify biological processes altered by fluridone exposure. The Medaka DNA sequences for the DEGs by fluridone exposure were retrieved and then used to obtain Ensembl transcript IDs by running BLASTN similarity searches against the annotated Medaka cDNA sequences downloaded from the Ensembl database (downloaded on January 5th, 2015, http://www.ensembl.org/index.html). Only nonredundant Ensembl transcript IDs were used for the gene set enrichment analysis, by removing sequences showing low similarities with a cutoff value of 95%. To implements the hypergeometric calculation, the hyperGTest function was used for identifying biological pathways altered by fluridone exposure compared with the control (P value cutoff = 0.05).

2.4. Reverse transcriptase quantitative PCR (RT-qPCR) Differently expressed genes (n = 10) were randomly selected to validate the RNA-seq results by RT-qPCR. In addition, the expression of five genes (acetylcholinesterase, AChE; retinoic acid receptor, RAR; insulin receptor substrate, IRS; glutathione reductase, GR; glutathione S transferase, GSH), selected based on the results of differentially expressed genes (DEG) and gene set enrichment analysis, were also analyzed in both sexes at all the fluridone concentrations. RT-qPCR was performed as described in our previous publications (Rochman et al., 2014; Ramírez-Duarte et al., 2016; Wagner et al., 2017). Briefly, one mg of total RNA was subjected to DNase treatment (AMPD1, Sigma-Aldrich, St. Louis, MO, USA), then followed by cDNA synthesis using superscript II Reverse Transcriptase (Thermo Fisher Scientific). The primers for RT-qPCR were designed by Primer 3.0 ver. 4.1.0 available online (http://primer3. ut.ee/) (Table S1). The reaction cocktail was prepared with Maxima SYBR Green qPCR Master Mix (Thermo Fisher Scientific) and the reactions were performed using LightCyclerÒ 480Ⅱ (Roche, Switzerland). All sample were run in triplicate. The relative expression in fold change of each gene was calculated by the 2 DDCt method and the data were normalized with the geometric means of three housekeeping genes, glyceraldehyde-3-phosphate dehydrogenase (GADPH), b-actin, and 18S ribosomal RNA (Schmittgen and Livak, 2008; Ramírez-Duarte et al., 2016). The gene expression data were log2 transformed before the statistical test using one-

way ANOVA, followed by Dunnett’s tests if significant differences were detected. The linear relationship of gene expression data from RNA-seq and RT-qPCR was assessed by Pearson’s correlation coefficient test in the package ‘Hmisc’ written in R (R Core Team, 2018; Wagner et al., 2017). 3. Results 3.1. Chemical concentrations Nominal concentrations of fluridone and concentrations measured by ELISA of experimental solutions used in larval Medaka exposures are shown in Table 1. Precipitate was observed at the highest concentration of fluridone which would interfere with the accuracy of the chemical determination. Therefore, the highest nominal concentration was not measured by ELISA. The nominal and measured concentrations were nearly identical (Table 1), so the nominal concentrations are reported for the remainder of the paper. 3.2. Hatching success of embryos Hatching success declined in a dose dependent manner in both sexes with increasing fluridone concentration (Table 2, Fig. 1). The fluridone parameter estimate in Model No. 2 was statistically significant (95% CI: 1.31, 0.72) (Table 3). No significant difference in hatching success due to sex was detected (Model No. 3 sex parameter estimate 95% CI: 0.92, 0.61) (Table 3). Based on the top-ranked model, the half effect concentration (EC50) of the total hatching success occurred at 2.3 mg L 1 (95% CI: 1.88, 2.79). At the highest concentration (4 mg L 1), the hatching rate of female and male embryos decreased to 12.5 and 25%, respectively (Table 2). No mortality was observed during the 14 day exposure of the embryos exposed to the fluridone at all the tested concentrations. One hatchling showing a developmental deformity (tail curvature) at 4 mg L 1 of fluridone. 3.3. Abnormal swimming behavior of larval fish and response to mechanical stimulation Abnormal swimming behavior was observed in larval fish exposed to high concentrations of fluridone. In the control group, fish were evenly distributed in the glass beakers with regular and constant swimming behaviors while fish exposed to fluridone (4.2 mg L 1 and higher) became lethargic and remained at the top of the experimental solution with slow and irregular swimming pattern. The stimulation test results are summarized in Fig. 2. The response to the stimuli was significantly decreased by fluridone for both males (ANOVA, F [5,17] = 8.6, P < 0.01) and females (ANOVA, F [5,17] = 8.8, P < 0.01). Based on Dunnett’s tests, these reductions occurred at the highest concentration in females (104 mg L 1), and at the two highest concentrations in males (21

Table 1 Nominal and measured concentrations of fluridone in the larval exposure test. Treatment

Nominal concentration mg L

Control 1 2 3 4 5 a

0 0.08 0.8 4.2 21 104

1

Measured concentration ± standard deviation

lM

mg L

0 0.24 2.5 13 64 320

Not detected 0.08 ± 0.01 0.8 ± 0.06 4.1 ± 0.33 22.5 ± 1.81 NAa

1

lM Not detected 0.25 ± 0.02 2.5 ± 0.18 12.5 ± 1.00 68.2 ± 5.50 NA

The concentration of fluridone was not available (NA) because precipitation interfered the accuracy of the chemical determination in the highest concentration.

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J. Jin et al. / Science of the Total Environment (2020) 134495 Table 2 Hatching success of Japanese Medaka embryo exposed to fluridone for 14 days. A total of 16 embryos for each sex were used per treatment. Fluridone (mg L

1

)

0 0.03 0.5 1 2 4

Female

Male

Total

No. of hatching

Hatching rate (%)

No. of hatching

Hatching rate (%)

No. of hatching

Hatching rate (%)

14 16 15 12 9 2

87.5 100 93.8 75 60 12.5

15 16 12 11 8 4

93.8 100 75 68.8 50 25

29 32 27 23 17 6

90.6 100 84.4 71.9 54.8 18.8

Fig. 1. Hatching success of Medaka embryos (both sexes) as a function of fluridone (mg L 1). The solid line represents the top-ranked model in Table 3 and the dashed line represents its 95% confidence interval. The ‘jitter’ function was used to prevent points from overlapping (R Core Team, 2018).

and 104 mg L exposure.

1

). No mortality was observed during the 6 h

3.4. Transcriptomic analysis The sequencing reaction generated averages of 59.7 and 63.9 million (M) reads for the control and fluridone treatments, respectively (Table S2). Among them, 79–81% of the reads were successfully mapped to the Medaka reference genome sequence, providing over 47 M reads per sample for the analysis (Table S2). In total, 59,037 unique genes were obtained, among which 799 genes were differentially expressed by fluridone, including 555 up- and 244 down-regulated genes. This included genes involved in growth and development (IRS and RAR), nervous system (AChE), and antioxidant system (GR and GST). The fluridone exposure altered a number of biological processes, such as cellular component organization, nervous system development, response to oxidative

Fig. 2. Response of the larval Medaka to the mechanical stimulation after fluridone exposure. Each bar represents the mean ± SE of three replicates. The data were collected from 5 fish per replicate. Asterisks (*) indicate statistical differences from the control group by one-way ANOVA with a post hoc Dunnett’s test (P < 0.05).

stress, calcium ion transport, androgen biosynthetic processes and androgen metabolic processes (Table 4).

Table 3 Model comparison of the hatching success models. The response variable in each model is proportion hatching success after 14 days. Model #

Model

DAICc

df

AICc wt

2 3 1

Fluridone Fluridone + Sex Intercept

0.0 1.9 64.8

2 3 1

0.72 0.28 <0.001

DAICc is the difference between model of interest and top-ranked model in Akaike Information Criterion Units corrected for small sample size, df is the degrees of freedom, and AICc wt is the Akaike weight.

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Table 4 List of biological processes up-regulated (A) and down-regulated (B) by fluridone exposure. GOBPID

P-value

Odds Ratio

Exp Count

Count

Size

BP Term

(A) Up-regulated biological processes GO:0055114 <0.001 2.837 GO:0006979 <0.001 6.802 GO:0015671 0.004 12.721 GO:0006820 0.007 2.45 GO:0030308 0.019 14.101 GO:0071466 0.019 6.356 GO:0008203 0.027 10.575 GO:0045926 0.037 8.459 GO:0006749 0.037 8.459 GO:0016125 0.037 8.459 GO:0051124 0.045 Inf GO:0090259 0.045 Inf GO:0006702 0.045 Inf GO:0002072 0.045 Inf GO:0008209 0.045 Inf GO:0071340 0.045 Inf

23 1 0 5 0 1 0 0 0 0 0 0 0 0 0 0

56 7 3 12 2 3 2 2 2 2 1 1 1 1 1 1

514 29 8 117 5 13 6 7 7 7 1 1 1 1 1 1

Oxidation-reduction process Response to oxidative stress Oxygen transport Anion transport Negative regulation of cell growth Cellular response to xenobiotic stimulus Cholesterol metabolic process Negative regulation of growth Glutathione metabolic process Sterol metabolic process Synaptic growth at neuromuscular junction Regulation of retinal ganglion cell axon guidance Androgen biosynthetic process Optic cup morphogenesis involved in camera-type eye development Androgen metabolic process Skeletal muscle acetylcholine-gated channel clustering

(B) Down-regulated biological processes GO:0016043 <0.001 0.386 GO:0051716 <0.001 0.63 GO:0007399 0.001 0.216 GO:0007417 0.002 0 GO:0098660 0.005 0.34 GO:0048513 0.007 0.496 GO:0030036 0.007 0.139 GO:0060322 0.009 0 GO:0048583 0.009 0.42 GO:0032502 0.009 0.637 GO:0007420 0.011 0 GO:0007010 0.018 0.363 GO:0034220 0.025 0.572 GO:0006816 0.028 0 GO:0050790 0.028 0.539 GO:0048468 0.035 0.403 GO:0007015 0.037 0 GO:0070588 0.044 0 GO:0022008 0.048 0.376

38 86 13 6 14 23 7 5 16 44 4 11 22 3 18 10 3 3 8

16 60 3 0 5 12 1 0 7 30 0 4 13 0 10 4 0 0 3

836 1900 287 131 307 506 150 102 349 980 97 232 480 77 392 210 71 67 169

Cellular component organization Cellular response to stimulus Nervous system development Central nervous system development Inorganic ion transmembrane transport Animal organ development Actin cytoskeleton organization Head development Regulation of response to stimulus Developmental process Brain development Cytoskeleton organization Ion transmembrane transport Calcium ion transport Regulation of catalytic activity Cell development Actin filament organization Calcium ion transmembrane transport Neurogenesis

Abbreviations: GOBPID: gene ontology (GO) molecular function identification number; P-value: statistical confidence in P-value obtained by hypergeometric test; Odds Ratio: odds ratio of enrichment of GO ID; Exp Count: the expected count of genes with the given GO term; Count: the count of genes that are annotated to GO ID in the set of differently expressed genes; Size: the count of genes that are annotated to GO ID in the background; BP Term: description of biological process (BP) for GO ID.

3.5. Validation of RNA-seq data by RT-qPCR A Pearson’s correlation coefficient test indicated that there was a significant positive correlation between the RNA-seq and RTqPCR results (r = 0.96, n = 10, P < 0.001, Fig. S1). 3.6. Gene expression analysis by RT-qPCR The relative expressions of the genes of interest (IRS, RAR, AChE, GR, and GST) that were differentially expressed and involved in the biological processes altered by fluridone, were analyzed for larval Medaka exposed at different concentrations of fluridone (Fig. 3). After 6 h exposure, IRS expression increased with fluridone concentration up to 21 mg L 1 in females (Fig. 3A). In males, the expression of IRS was slightly down-regulated by fluridone at the two lowest concentrations (0.08 and 0.8 mg L 1), and then upregulated in a dose dependent manner as concentration increased (Fig. 3B). However, IRS expression at all the concentrations showed no significant difference among treatments (Female: ANOVA, F [5,17] = 1.21, P = 0.36; Male: ANOVA, F [5,17] = 0.83, P = 0.55). The expression of RAR was slightly induced in females at 0.8 mg L 1 and higher concentrations, while the expression was downregulated in males at all the concentrations tested in this study (Fig. 3C and D). No statistically significant difference was detected for RAR expression (Female: ANOVA, F [5,17] = 0.12, P = 0.99; Male: ANOVA, F [5,17] = 0.63, P = 0.68). Similarly, AChE expression was

down-regulated in males while up-regulation was observed in females with the largest change at 0.8 mg L 1 of fluridone (Fig. 3E and F). However, no statistically significant difference was found in both sexes (Female: ANOVA, F [5,17] = 0.12, P = 0.99; Male: ANOVA, F [5,17] = 0.91, P = 0.51). Statistical differences of GR expression were not detected in females nor males (Female: ANOVA, F [5,17] = 0.87, P = 0.53; Male: ANOVA, F [5,17] = 0.57, P = 0.72), however, a dose dependent response was observed in both sexes (Fig. 3G and H). The gene expression of GST in females was induced by fluridone exposure and significantly increased in the 4.2 mg L 1 treatment (ANOVA, F [5,17] = 3.40, P < 0.05) (Fig. 3I). In males, although significant differences of GST expression were not detected (ANOVA, F [5,17] = 1.63, P = 0.23) (Fig. 3J), the expression of GST was down-regulated and negatively correlated with fluridone concentration.

4. Discussion Although fluridone has been actively used for controlling invasive aquatic plants, our understanding on the adverse effects of fluridone on non-target organisms is still limited. In our previous study, we reported acute toxicity (abnormal swimming behavior) and sublethal effects (endocrine disruption, neurotoxicity, and oxidative stress) in adult Delta Smelt following 6 h fluridone exposure tests (Jin et al., 2018). To better understand the mechanisms of

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Fig. 3. The gene expression of IRS (A and B), RAR (C and D), AChE (E and F), GR (G and H) and GST (I and J) in female (A, C, E, G, and I) and male (B, D, F, H, and J) larval Medaka exposed to fluridone. Each bar represents the mean ± SE of three replicates. The data are from 10 fish per replicate. The asterisk (*) indicates statistical differences from the control group based on a post hoc Dunnett’s test (P < 0.05).

toxicity of fluridone and its adverse effects on early developmental stages of fish, we assessed the hatching success of the fish model

Japanese Medaka, its response to mechanical stimuli, and investigated gene expression patterns after fluridone exposure.

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The hatching success of embryos decreased with increasing fluridone exposure in a dose dependent manner, suggesting toxic effects of fluridone on fish embryonic development (Fig. 1). The impairment of hatching success seems to be a general response of the embryos to various contaminants (Viant et al., 2006; Wagner et al., 2017). Fish in embryonic stages are a particularly susceptible life-stage given that they cannot evade or detoxify contaminants like adult fish due to their immobility and the immature defense system. In addition, the embryonic stage is more susceptible to a cascade of failures during development that could impact the hatching success, growth rate, and organogenesis, eventually causing teratogenic effects (Schulte and Nagel, 1994, Tzima et al., 2017; Wiegand et al., 2000, 2001). The RNA-seq data from larval fish also indicated that biological processes related to growth and development were affected by fluridone. Besides, the related DEGs, IRS and RAR, were significantly affected by fluridone exposure. The dose-response relationship of IRS expression with fluridone concentration indicated the disturbance of fluridone on IRS expression in larval fish. IRS are a family of cytoplasmic adaptor proteins mediating insulin signaling and play a central role in maintaining basic cellular functions such as regulation of cell differentiation, growth, proliferation, survival, and metabolism (Machado-Neto et al., 2018; Sesti et al., 2001). Studies on IRS demonstrated that knockout mice lacking IRS resulted in series of defects including retardant embryonal and postnatal growth (Tamemoto et al., 1994; Burks et al., 2000; Fantin et al., 2000; Mardilovich et al., 2009). RAR is a nuclear receptor for retinoic acid (RA), which is an essential chemical for the development of embryonic structures including the nervous system and notochords of chordates (Carter et al., 2015; Lohnes et al., 1994; Ruberte et al., 1993). RAR conducts the signaling of RA through expression of subsets of target genes involved in cellular differentiation, proliferation and apoptosis (Samarut and Rochette-Egly, 2012). Research on mice demonstrated that RAR is essential for normal development of many structures, since RAR mutant mice exhibited antepartum or postpartum lethality and a number of congenital abnormalities such as malformed eyes, glandular defects, and skeletal abnormalities (Lohnes et al., 1993, 1994). A study on Zebrafish also suggested that RAR plays a crucial role in maintaining the neural crest and mesodermal stem/progenitor cells during normal embryonic development and tissue regeneration (Wai, 2015). The disturbance of IRS, RAR, and/or other genes involved in developmental processes may have resulted in delayed hatching and hatch failure of the Medaka embryo. The observation and data of fish swimming behavior as well as RNA-seq analysis all suggest that fluridone functions as a neurotoxicant in fish. The gene, AChE, which is related to the function of the nervous system, showed dose- and sex-specific expression (Fig. 3). AChE plays an important role in cholinergic neuronal function by hydrolysis and homeostasis regulating of the neurotransmitter acetylcholine (ACh) in the body. AChE is used as a classical biomarker of neurotoxicity in response to the exposure to a number of pollutants like pesticides and herbicides (Gholami-Seyedkolaei et al., 2013; Menéndez-Helman et al., 2012; Dos Santos Miron et al., 2005). There is increasing evidence that many herbicides are neurotoxic to fishes through inhibiting the activity of AChE (Cattaneo et al., 2011; Glusczak et al., 2006; Da Fonseca et al., 2008; Murussi et al., 2014; Topal et al., 2017a, b). Changes in AChE activity are highly correlated with abnormal behaviors of fishes (Baldissera et al., 2018; Labenia et al., 2007; Rao et al., 2005). The inhibition of AChE could lead to the over accumulation of acetylcholine in synapses that consequently interferes with the normal function of the nervous system, which may negatively affect swimming and reproductive behavior of fishes (Bhattacharya, 1993; Dutta and Arends, 2003; Saglio and Trijasse, 1998; Schmidel et al., 2014). In contrast, the over excitation of

AChE activity could lead to a rapid degradation of ACh and hyperactivated cholinergic system, highly associated with anxiety-like behaviors such as hyperlocomotion (Baldissera et al., 2018; McCloskey et al., 2017). In our previous study on Delta Smelt, adults exposed to the same concentrations of fluridone also showed the abnormal swimming behavior, as well as a decline in AChE activity (Jin et al., 2018). Thus, the inhibition of AChE may explain the neurotoxic effects of fluridone on larval Medaka observed in this study. The transcriptome analysis revealed oxidative stress of fluridone on larval Medaka. Numerous studies indicate that various environmental pollutants could cause oxidative stress in fish by inducing generation of reactive oxygen species (ROS), which could lead to the oxidative injury of the cell, such as lipid peroxidation, protein and nuclear damage (Ahmad et al., 2000; Hermes-Lima, 2004; Modesto and Martinez, 2010; Nwani et al., 2013). The data from RNA-seq suggest that glutathione reductase (GR) and glutathione S transferase (GST) were differentially expressed in larval Medaka in response to fluridone exposure. GR is responsible for the maintenance of the cellular level of glutathione (GSH), a primary non-enzymatic antioxidant that scavenges free radicals by catalyzing the oxidized glutathione (GSSG) to reduced glutathione (Livingstone, 2001; Lesser, 2011; Monserrat et al., 2011; Regoli and Principato, 1995). Therefore, GR plays an important role in the oxidative defense systems in fish (Parvez and Raisuddin, 2005, Tripathi et al., 2006, Dong et al., 2014; Ramírez-Duarte et al., 2017b). In addition, GST takes part in the detoxification of endogenous and exogenous chemicals in the fish body, via mediating the catalyzing of a serial reaction between GSH and xenobiotics with electrophilic substances (Regoli and Principato, 1995; Glisic et al., 2015). There are a number of publications reporting the adverse effects of contaminants to antioxidant systems via the disturbance of the GR and GST levels in fish (Atli and Canli, 2010; Lushchak et al., 2009; Modesto and Martinez, 2010; Nwani et al., 2013). The GR and GST expression in larval Medaka showed a dose-response relationship with the fluridone concentration in the present study, suggesting that fluridone compromised the antioxidant system and detoxification ability and caused oxidative stress in larval Medaka. An increasing literature has revealed that many herbicides have estrogenic activity that disturbs the endocrine systems of aquatic organisms (Gronen et al., 1999; Mnif et al., 2011; Sonnenschein and Soto, 1998; Zhu et al., 2017). The present study indicated that fluridone showed endocrine disruption in Medaka, which was revealed by transcriptome data. The results support our previous finding that adult Delta Smelt exposed to fluridone showed increased levels of hepatic E2 concentrations (Jin et al., 2018). It is noteworthy that the expression pattern of all the genes tested by RT-qPCR (IRS, RAR, AChE, GR and GST) exhibited different expression patterns between female and male in the current study (Fig. 3). Sex-dependent differences in the response of organisms to pollutants are widely reported (Murray et al., 2003; Ookubo et al., 2009; Wang et al., 2018). The sex-biased factors underlying sex differences in physiology and protective/damage mechanisms are possibly due to the difference of gonadal hormones and inherent sexual inequalities in effects of sex chromosome genes (Arnold, 2017; Arnold et al., 2017). The data from the present study and our previous work suggest that fluridone application in the field compromises fish health (Jin et al., 2018). The recommended maximum application rate of fluridone is 0.09 mg L 1 for a one-time treatment or 0.15 mg L 1 for the maximum cumulative application rate per growth cycle (Yi et al., 2011; United States Environmental Protection Agency, 2004; Product’s label). Although these application rates suggest that field concentrations are lower than the concentrations at which we observed significant effects on Medaka (20 mg L 1), observed

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concentrations can exceed recommended application rates. For example, Jacob et al. (2016) reported that a nominal application rate of 0.079 mg L 1 to ponds resulted in a fluridone concentration as high as 0.33 mg L 1 after 24 h. In addition, the pellet form of fluridone has been applied in the Delta, which may increase the local concentration of fluridone in the benthic environment where fish eggs and larvae occur beyond the label specifications. To achieve the maximum levels of efficacy for controlling unwanted aquatic plants, multiple applications of herbicide are applied to treat young plants in spring (Isaacs et al., 1989; Pratt et al., 1997; Poovey et al., 2010, USDA and CDBW 2012). For fluridone, it requires 45 days of contact time to be effective according to the information reviewed by the Environmental Protection Agency (EPA) and is typically prescribed for one to four months (Archambault and Cope, 2016; United States Environmental Protection Agency, 2004). This can be an issue in the Delta because spawning of fish such as Delta Smelt take place in February through May, overlapping temporally with herbicides applications in the aquatic environment. Although field data are still lacking and there are no official documents reporting the fluridone concentrations in the ambient water in the Delta, it is very likely that fish in the aquatic environment are exposed to fluridone. 5. Conclusion Fluridone application in the field is considered to be an effective approach for controlling invasive aquatic plants, however like any herbicide, it poses a risk to non-target organisms. In this study, we report that fluridone significantly suppressed the hatching rate of Medaka embryos and compromised response of larval Medaka against stimulation. In addition, transcriptome analysis revealed that various biological processes such as developmental processes, nervous system development, oxidation–reduction processes, and the endocrine system were disrupted by fluridone exposure, suggesting that the chemical causes developmental toxicity, neurotoxicity, oxidative stress, and endocrine disruption in fish. Application of fluridone may compromise fish health at early developmental stages in aquatic environments. Declaration of Competing Interest The authors declare that they have no competing interests. Acknowledgements This study was supported by the California Department of Boating and Waterways (contract# C1370030) and United States Fish and Wildlife Service (contract# F15AP00771). The sequencing reactions for RNA-seq were carried out at the DNA Technologies and Expression Analysis Core Laboratory at the UC Davis, supported by NIH Shared Instrumentation Grant 1S10OD010786-01. We are grateful to Teh C. and other members at the Aquatic Health Program for their help running the exposure tests and sample processing. Jiali Jin acknowledges the financial support from the China Scholarship Council. Appendix A. Supplementary data Supplementary data to this article can be found online at https://doi.org/10.1016/j.scitotenv.2019.134495. References Ahmad, I., Hamid, T., Fatima, M., Chand, H.S., Jain, S.K., Athar, M., Raisuddin, S., 2000. Induction of hepatic antioxidants in freshwater Catfish (Channa punctatus Bloch) is a biomarker of paper mill effluent exposure. BBA 1523, 37–48.

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Archambault, J.M., Bergeron, C.M., Cope, W.G., Richardson, R.J., Heilman, M.A., Corey, J.E., Netherland, M.D., Heise, R.J., 2015. Sensitivity of freshwater molluscs to hydrilla-targeting herbicides: providing context for invasive aquatic weed control in diverse ecosystems. J. Freshwater Ecol. 30, 335–348. Archambault, J.M., Cope, W.G., 2016. Life stage sensitivity of a freshwater snail to herbicides used in invasive aquatic weed control. Freshwater Mollusk Biol. Conserv. 19, 69–79. Arnold, A.P., 2017. A general theory of sexual differentiation. J. Neurosci. Res. 95, 291–300. Arnold, A.P., Cassis, L.A., Eghbali, M., Reue, K., Sandberg, K., 2017. Sex hormones and sex chromosomes cause sex differences in the development of cardiovascular diseases. Arterioscler. Thromb. Vasc. Biol. 37, 746–756. Arnold, W.R., 1979. Fluridone–a new aquatic herbicide. J. Aquat. Plant Manag. 17, 30–33. Atli, G., Canli, M., 2010. Response of antioxidant system of freshwater fish Oreochromis niloticus to acute and chronic metal (Cd, Cu, Cr, Zn, Fe) exposures. Ecotoxicol. Environ. Saf. 73, 1884–1889. Baldissera, M.D., Souza, C.F., Zeppenfeld, C.C., Descovi, S.N., Moreira, K.L.S., Da Rocha, M.I.U.M., Da Veiga, M.L., Da Silva, A.S., Baldisserotto, B., 2018. Aflatoxin B1contaminated diet disrupts the blood-brain barrier and affects fish behavior: Involvement of neurotransmitters in brain synaptosomes. Environ. Toxicol. Pharmacol. 60, 45–51. Bhattacharya, S., 1993. Target and non-target effects of anticholinesterase pesticides in fish. Sci. Total Environ. 134, 859–866. Bolker, B., 2010. bbmle: Tools for general maximum likelihood estimation. R package version 0.9 5. Boyd, C.E., Tucker, C.S., 1998. Pond Aquaculture Water Quality Management. Kluwer, Norwell, MA, USA. Bremigan, M.T., Hanson, S.M., Soranno, P.A., Cheruvelil, K.S., Valley, R.D., 2005. Aquatic vegetation, Largemouth bass, and water quality responses to low-dose fluridone two-years post treatment. J. Aquat. Plant Manage. 43, 65–75. Burks, D.J., De Mora, J.F., Schubert, M., Withers, D.J., Myers, M.G., Towery, H.H., Altamuro, S.L., Flint, C.L., White, M.F., 2000. IRS-2 pathways integrate female reproduction and energy homeostasis. Nature 407, 377–382. Burnham, K.P., Anderson, D.R., 2002. Model Selection and Multimodel Inference: A Practical Information-Theoretic Approach. Springer, New York. Cahoon, C.W., York, A.C., Jordan, D.L., Seagroves, R.W., Everman, W.J., Jennings, K.M., 2015. Fluridone carryover to rotational crops following application to cotton. J. Cotton. Sci. 19, 631–640. Carter, C.J., Rand, C., Mohammad, I., Lepp, A., Vesprini, N., Wiebe, O., Carlone, R., Spencer, G.E., 2015. Expression of a retinoic acid receptor (RAR)-like protein in the embryonic and adult nervous system of a protostome species. J. Exp. Zool. Part B: Mol. Dev. Evol. 324, 51–67. Cattaneo, R., Clasen, B., Lucia Loro, V., de Menezes, C.C., Moraes, B., Santi, A., Toni, C., de Avila, L.A., Zanella, R., 2011. Toxicological responses of Cyprinus carpio exposed to the herbicide penoxsulam in rice field conditions. J. Appl. Toxicol. 31, 626–632. Cheruvelil, K.S., 2004. Examining Lakes at Multiple Spatial Scales: Predicting Fish Growth, Macrophyte Cover and Lake Physio-Chemical Variables PhD dissertation. Michigan State University, East Lansing. Da Fonseca, M.B., Glusczak, L., Moraes, B.S., De Menezes, C.C., Pretto, A., Tierno, M.A., Zanella, R., Gonçalves, F.F., Loro, V.L., 2008. The 2,4-D herbicide effects on acetylcholinesterase activity and metabolic parameters of piava freshwater fish (Leporinus obtusidens). Ecotoxicol. Environ. Safe. 69, 416–420. DeKoven, D.L., Núñez, J.M., Lester, S.M., Conklin, D.E., Marty, G.D., Parker, L.M., Hinton, D.E., 1992. A purified diet for Medaka (Oryzias latipes): refining a fish model for toxicological research. Lab. Anim. Sci. 42, 180–189. Dong, X., Liu, H., Chen, F., Li, D., Zhao, Y., 2014. MiR-214 promotes the alcoholinduced oxidative stress via down-regulation of glutathione reductase and cytochrome P450 oxidoreductase in liver cells. Alcohol. Clin. Exp. Res. 38, 68– 77. Dos Santos Miron, D., Crestani, M., Rosa Shettinger, M., Maria Morsch, V., Baldisserotto, B., Angel Tierno, M., Moraes, G., Vieira, V.L., 2005. Effects of the herbicides clomazone, quinclorac, and metsulfuron methyl on acetylcholinesterase activity in the silver Catfish (Rhamdia quelen) (Heptapteridae). Ecotoxicol. Environ. Safe. 61, 398–403. Dutta, H.M., Arends, D.A., 2003. Effects of endosulfan on brain acetylcholinesterase activity in juvenile Bluegill sunfish. Environ. Res. 91, 157–162. Fantin, V.R., Wang, Q., Lienhard, G.E., Keller, S.R., 2000. Mice lacking insulin receptor substrate 4 exhibit mild defects in growth reproduction, and glucose homeostasis. Am. J. Physiol. Endocrinol. Metab. 278, E127–E133. Gholami-Seyedkolaei, S.J., Mirvaghefi, A., Farahmand, H., Kosari, A.A., 2013. Effect of a glyphosate-based herbicide in Cyprinus carpio: Assessment of acetylcholinesterase activity, hematological responses and serum biochemical parameters. Ecotoxicol. Environ. Safe. 98, 135–141. Glisic, B., Mihaljevic, I., Popovic, M., Zaja, R., Loncar, J., Fent, K., Kovacevic, R., Smital, T., 2015. Characterization of glutathione-S-transferases in Zebrafish (Danio rerio). Aquat. Toxicol. 158, 50–62. Glusczak, L., dos Santos Miron, D., Crestani, M., da Fonseca, M.B., de Araújo Pedron, F., Duarte, M.F., Vieira, V.L., 2006. Effect of glyphosate herbicide on acetylcholinesterase activity and metabolic and hematological parameters in piava (Leporinus obtusidens). Ecotoxicol. Environ. Safe. 65, 237–241. Gronen, S., Denslow, N., Manning, S., Barnes, S., Barnes, D., Brouwer, M., 1999. Serum vitellogenin levels and reproductive impairment of male Japanese Medaka (Oryzias latipes) exposed to 4-tert-octylphenol. Environ. Health Persp. 107, 385–390.

10

J. Jin et al. / Science of the Total Environment (2020) 134495

Hamelink, J.L., Buckler, D.R., Mayer, F.L., Palawski, D.U., Sanders, H.O., 1986. Toxicity of fluridone to aquatic invertebrates and fish. Environ. Toxicol. Chem. 5, 87–94. Hammock, B.G., Lesmeister, S., Flores, I., Bradburd, G.S., Hammock, F.H., Teh, S.J., 2016. Low food availability narrows the tolerance of the copepod Eurytemora affinis to salinity, but not to temperature. Estuar. Coast. 39, 189–200. Hermes-Lima, M., 2004. Oxygen in biology and biochemistry: Role of Free Radicals. In: Storey, K.B., (Ed.), Functional Metabolism: Regulation and Adaptation. Cap, pp. 319–368. Isaacs, M.A., Murdock, E.C., Toler, J.E., Wallace, S.U., 1989. Effects of late-season herbicide applications on Sicklepod (Cassia obtusifolia) seed production and viability. Weed Sci. 37, 761–765. Jacob, A.P., Culver, D.A., Lanno, R.P., Voigt, A., 2016. Ecological impacts of fluridone and copper sulphate in Catfish aquaculture ponds. Environ. Toxicol. Chem. 35, 1183–1194. Jin, J.L., Kurobe, T., Ramírez-Duarte, W.F., Melissa, B.B., Lam, C.H., Pandey, P.K., Hung, T.C., Stillway, M.E., Zweig, L., Caudill, J., Lin, L., Teh, S.J., 2018. Sublethal effects of herbicides penoxsulam, imazamox, fluridone and glyphosate on Delta Smelt (Hypomesus transpacificus). Aquat. Toxicol. 197, 79–88. Kamarianos, A., Altiparmakis, J., Karamanlis, X., Kufidis, D., Kousouris, T.H., Fotis, G., Kilikidis, S., 1989. Experimental evaluation of fluridone effectiveness on fish productive aquatic systems. J. Aquat. Plant Manag. 27, 24–26. Kim, D., Pertea, G., Trapnell, C., Pimentel, H., Kelley, R., Salzberg, S.L., 2013. TopHat2: accurate alignment of transcriptomes in the presence of insertions, deletions and gene fusions. Genome Biol. 14, R36. Kinoshita, M., Murata, K., Naruse, K., Tanaka, M., 2009. Medaka: Biology, Management, and Experimental Protocols. Wiley-Blackwell, Iowa. Kurobe, T., Lehman, P.W., Haque, M.E., Sedda, T., Lesmeister, S., Teh, S.J., 2018. Evaluation of water quality during successive severe drought years within Microcystis blooms using fish embryo toxicity tests for the San Francisco Estuary, California. Sci. Total. Environ. 610, 1029–1037. Labenia, J.S., Baldwin, D.H., French, B.L., Davis, J.W., Scholz, N.L., 2007. Behavioral impairment and increased predation mortality in Cutthroat Trout exposed to carbaryl. Mar. Ecol. Prog. Ser. 329, 1–11. Lesser, M.P., 2011. Oxidative stress in tropical marine ecosystems. In: Abele, D., Vázquez Medina, J.P., Zenteno Savín, T. (Eds.), Oxidative Stress in Aquatic Ecosystems. John Wiley & Sons Ltd, Chichester, UK, pp. 7–19. Livingstone, D.R., 2001. Contaminant-stimulated reactive oxygen species production and oxidative damage in aquatic organisms. Mar. Pollut. Bull. 42, 656–666. Lohnes, D., Kastner, P., Dierich, A., Mark, M., LeMeur, M., Chambon, P., 1993. Function of retinoic acid receptor c (RARc) in the mouse. Cell 73, 643–658. Lohnes, D., Mark, M., Mendelsohn, C., Dollé, P., Dierich, A., Gorry, P., Gansmuller, A., Chambon, P., 1994. Function of the retinoic acid receptors (RARs) during development (I) Craniofacial and skeletal abnormalities in RAR double mutants. Development 120, 2723–2748. Lushchak, O.V., Kubrak, O.I., Storey, J.M., Storey, K.B., Lushchak, V.I., 2009. Low toxic herbicide Roundup induces mild oxidative stress in Goldfish tissues. Chemosphere 76, 932–937. Machado-Neto, J.A., Fenerich, B.A., Alves, A.P.N.R., Fernandes, J.C., Scopim-Ribeiro, R., Coelho-Silva, J.L., Traina, F., 2018. Insulin Substrate Receptor (IRS) proteins in normal and malignant hematopoiesis. Clinics, 73. Mardilovich, K., Pankratz, S.L., Shaw, L.M., 2009. Expression and function of the insulin receptor substrate proteins in cancer. Cell Commun. Signal. 7, 1–14. McCloskey, M.C., Young, T.J., Anderson, S.M., 2017. The influence of acetylcholinesterase on anxiety- and depression-like behaviors in fluoxetinetreated male mice. Bios 88, 29–38. Mccowen, M.C., Young, C.L, West, S.D., Parka, S.J., Arnold, W.R., 1979. Fluridone, a new herbicide for aquatic plant management. McElreath, R., 2016. Statistical Rethinking: A Bayesian Course with Examples in R and Stan. CRC Press. Menéndez-Helman, R.J., Ferreyroa, G.V., Afonso, M.D.S., Salibián, A., 2012. Glyphosate as an acetylcholinesterase inhibitor in Cnesterodon decemmaculatus. Bull. Environ. Contam. Toxicol. 88, 6–9. Mnif, W., Hassine, A.I., Bouaziz, A., Bartegi, A., Thomas, O., Roig, B., 2011. Effect of endocrine disruptor pesticides: A review. Int. J. Environ. Res. Public Health 8, 2265–2303. Modesto, K.A., Martinez, C.B., 2010. Effects of Roundup Transorb on fish: hematology, antioxidant defenses and acetylcholinesterase activity. Chemosphere 81, 781–787. Monserrat, J., Letts, R.E., Ferreira, J.L.R., Ventura-Lima, J., Rocha, A.M., 2011. Biomarkers of oxidative stress: benefits and drawbacks for their application in biomonitoring of aquatic environments. In: Abele, D., Vázquez Medina, J.P., Zenteno Savín, T. (Eds.), Oxidative Stress in Aquatic Ecosystems. John Wiley & Sons Ltd, Chichester, UK, pp. 317–326. Murray, H.E., Pillai, A.V., McArthur, S.R., Razvi, N., Datla, K.P., Dexter, D.T., Gillies, G. E., 2003. Dose- and sex-dependent effects of the neurotoxin 6hydroxydopamine on the nigrostriatal dopaminergic pathway of adult rats: Differential actions of estrogen in males and females. Neuroscience 116, 213– 222. Murussi, C.R., Thorstenberg, M.L., Leitemperger, J., Costa, M., Clasen, B., Santi, A., Menezes, C., Engers, V.K., Loro, V.L., 2014. Toxic effects of penoxsulam herbicide in two fish species reared in southern Brazil. Bull. Environ. Contam. Toxicol. 92, 81–84. Netherland, M.D., Honnell, D.R., Staddon, A.G., Getsinger, K.D., 2002. Comparison of immunoassay and HPLC for analyzing fluridone concentrations: New applications for immunoassay techniques. Lake Reserv. Manage. 18, 75–80.

Nwani, C.D., Nagpure, N.S., Kumar, R., Kushwaha, B., Lakra, W.S., 2013. DNA damage and oxidative stress modulatory effects of glyphosate-based herbicide in freshwater fish, Channa punctatus. Environ. Toxicol. Pharmacol. 36, 539–547. Ookubo, M., Yokoyama, H., Kato, H., Araki, T., 2009. Gender differences on MPTP (1methyl-4-phenyl-1,2,3,6-tetrahydropyridine) neurotoxicity in C57BL/6 mice. Mol. Cell. Endocrinol. 311, 62–68. Parsons, J.K., Couto, A., Hamel, K.S., Marx, G.E., 2009. Effect of fluridone on macrophytes and fish in a coastal Washington lake. J. Aquat. Plant Manag. 47, 31–40. Parvez, S., Raisuddin, S., 2005. Protein carbonyls: novel biomarkers of exposure to oxidative stress-inducing pesticides in freshwater fish Channa punctata (Bloch). Environ. Toxicol. Pharmacol. 20, 112–117. Paul, E.A., Simonin, H.A., Symula, J., Bauer, R.W., 1994. The toxicity of diquat, endothall, and fluridone to the early life stages of fish. J. Freshwater Ecol. 9, 229–239. Poovey, A.G., Glomski, L.A.M., Netherland, M.D., Skogerboe, J.G., 2010. Early Season Applications of Fluridone for Control of Curlyleaf Pondweed. Aquatic Plant Control Research Program. Engineer Research and Development Center. ERDC/ EL-TR-10-22. Pratt, J.R., Melendez, A.E., Barreiro, R., Bowers, N.J., 1997. Predicting the ecological effects of herbicides. Ecol. Appl. 7, 1117–1124. R Core Team, 2018. R: A language and environment for statistical computing. R foundation for statistical computing. Vienna, Austria. https://www.R-project. org/. Ramírez-Duarte, W.F., Jin, J., Kurobe, T., Teh, S.J., 2016. Effects of prolonged exposure to low pH on enzymatic and non-enzymatic antioxidants in Japanese Medaka (Oryzias latipes). Sci. Total Environ. 568, 26–32. Ramírez-Duarte, W.F., Kurobe, T., Teh, S.J., 2017a. Effects of low levels of ultraviolet radiation on antioxidant mechanisms of Japanese Medaka (Oryzias latipes). Chemosphere 181, 304–312. Ramírez-Duarte, W.F., Kurobe, T., Teh, S.J., 2017b. Impairment of antioxidant mechanisms in Japanese medaka (Oryzias latipes) by acute exposure to aluminum. Comp. Biochem. Physiol. C: Toxicol. Pharmacol. 198, 37–44. Rao, J.V., Begum, G., Pallela, R., Usman, P.K., Rao, R.N., 2005. Changes in behavior and brain acetylcholinesterase activity in mosquito fish, Gambusia affinis in response to the sub-lethal exposure to chlorpyrifos. Int. J. Environ. Res. Public Health 2, 478–483. Regoli, F., Principato, G., 1995. Glutathione, glutathione-dependent and antioxidant enzymes in mussel, Mytilus galloprovincialis, exposed to metals under field and laboratory conditions: implications for the use of biochemical biomarkers. Aquat. Toxicol. 31, 143–164. Rochman, C.M., Kurobe, T., Flores, I., Teh, S.J., 2014. Early warning signs of endocrine disruption in adult fish from the ingestion of polyethylene with and without sorbed chemical pollutants from the marine environment. Sci. Total Environ. 493, 656–661. Ruberte, E., Friederich, V., Chambon, P., Morriss-Kay, G., 1993. Retinoic acid receptors and cellular retinoid binding proteins. III. Their differential transcript distribution during mouse nervous system development. Development 118, 267–282. Saglio, P., Trijasse, S., 1998. Behavioral responses to atrazine and diuron in Goldfish. Arch. Environ. Contam. Toxicol. 35, 484–491. Samarut, E., Rochette-Egly, C., 2012. Nuclear retinoic acid receptors: conductors of the retinoic acid symphony during development. Mol. Cell. Endocrinol. 348, 348–360. Schmidel, A.J., Assmann, K.L., Werlang, C.C., Bertoncello, K.T., Francescon, F., Rambo, C.L., Beltrame, G.M., Calegari, D., Batista, C.B., Blaser, R.E., Roman Júnior, W.A., Conterato, G.M., Piato, A.L., Zanatta, L., Magro, J.D., Rosemberg, D.B., 2014. Subchronic atrazine exposure changes defensive behaviour profile and disrupts brain acetylcholinesterase activity of Zebrafish. Neurotoxicol. Teratol. 44, 62– 69. Schmittgen, T.D., Livak, K.J., 2008. Analyzing real-time PCR data by the comparative CT method. Nat. Protoc. 3, 1101–1108. Schulte, C., Nagel, R., 1994. Testing acute toxicity in the embryo of Zebrafish, Brachydanio rerio, as an alternative to the acute fish test: preliminary results. Atla-Altern. Lab. Anim. 22, 12–19. Sesti, G., Federici, M., Hribal, M.L., Lauro, D., Sbraccia, P., Lauro, R., 2001. Defects of the insulin receptor substrate (IRS) system in human metabolic disorders. FASEB J. 15, 2099–2111. Siemering, G.S., Hayworth, J.D., Greenfield, B.K., 2008. Assessment of potential aquatic herbicide impacts to California aquatic ecosystems. Arch. Environ. Contam. Toxicol. 55, 415–431. Sonnenschein, C., Soto, A.M., 1998. An updated review of environmental estrogen and androgen mimics and antagonists. J. Steroid Biochem. Mol. Biol. 65, 143– 150. Sprecher, L., Netherland, M.D., Stewart, A.B., 1998. Fhytoene and carotene response of aquatic plants to fluridone under laboratory conditions. J. Aquat. Plant Manag. 36, 111–120. Stehr, C.M., Linbo, T.L., Incardona, J.P., Scholz, N.L., 2006. The developmental neurotoxicity of fipronil: Notochord degeneration and locomotor defects in Zebrafish embryos and larvae. Toxicol. Sci. 92, 270–278. Tamemoto, H., Kadowaki, T., Tobe, K., Yagi, T., Sakura, H., Hayakawa, T., Terauchi, Y., Ueki, K., Kaburagi, Y., Satoh, S., Sekihara, H., Yoshioka, S., Horikoshi, H., Furuta, Y., Ikawa, Y., Kasuga, M., Yazaki, Y., Aizawa, S., 1994. Insulin resistance and growth retardation in mice lacking insulin receptor substrate-1. Nature 372, 182–186.

J. Jin et al. / Science of the Total Environment (2020) 134495 Topal, A., Alak, G., Ozkaraca, M., Yeltekin, A.C., Comaklı, S., Acıl, G., Kokturk, M., Atamanalp, M., 2017a. Neurotoxic responses in brain tissues of Rainbow trout exposed to imidacloprid pesticide: Assessment of 8-hydroxy-2-deoxyguanosine activity, oxidative stress and acetylcholinesterase activity. Chemosphere 175, 186–191. Topal, A., Alak, G., Altun, S., Erol, H.S., Atamanalp, M., 2017b. Evaluation of 8hydroxy-2-deoxyguanosine and NFkB activation, oxidative stress response, acetylcholinesterase activity, and histopathological changes in Rainbow trout brain exposed to linuron. Environ. Toxicol. Pharmacol. 49, 14–20. Trapnell, C., Roberts, A., Goff, L., Pertea, G., Kim, D., Kelley, D.R., Pimentel, H., Salzberg, S.L., Rinn, J.L., Pachter, L., 2012. Differential gene and transcript expression analysis of RNA-Seq experiments with TopHat and Cufflinks. Nat. Protoc. 7, 562–578. Trapnell, C., Williams, B.A., Pertea, G., Mortazavi, A., Kwan, G., van Baren, M.J., Salzberg, S.L., Wold, B.J., Pachter, L., 2010. Transcript assembly and quantification by RNA-Seq reveals unannotated transcripts and isoform switching during cell differentiation. Nat. Biotechnol. 28, 511–515. Tripathi, B.N., Mehta, S.K., Amar, A., Gaur, J.P., 2006. Oxidative stress in Scenedesmus sp. during short- and long-term exposure to Cu2+ and Zn2+. Chemosphere 62, 538–544. Truong, L., Harper, S.L., Tanguay, R.L., 2011. Evaluation of embryo toxicity using the Zebrafish model. Method. Mol. Biol. 691, 271–279. Tzima, E., Serifi, I., Tsikari, I., Alzualde, A., Leonardos, I., Papamarcaki, T., 2017. Transcriptional and behavioral responses of Zebrafish larvae to Microcystin-LR exposure. Int. J. Mol. Sci. 18, 365. USDA and CDBW, 2012. Water Hyacinth Control Program, Biological Assessment accessed on May 11th, 2019 (https://matome.naver.jp/odai/ 2146208819302928701 . United States Environmental Protection Agency, 2004. Report of the Food Quality Protection Act (FPQA) Tolerance Reassessment Progress and Risk Management Decision (TRED) for Fluridone.

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Viant, M.R., Pincetich, C.A., Hinton, D.E., Tjeerdema, R.S., 2006. Toxic actions of dinoseb in Medaka (Oryzias latipes) embryos as determined by in vivo 31P NMR, HPLC-UV and 1 H NMR metabolomics. Aquat. Toxicol. 76, 329–342. Wada, H., Shimada, A., Fukamachi, S., Naruse, K., Shima, A., 1998. Sex-linked inheritance of the lf locus in the Medaka fish (Oryzias latipes). Zool. Sci. 15, 123– 126. Wagner, S.D., Kurobe, T., Hammock, B.G., Lam, C.H., Wu, G., Vasylieva, N., Gee, S.J., Hammock, B.D., The, S.J., 2017. Developmental effects of fipronil on Japanese Medaka (Oryzias latipes) embryos. Chemosphere 166, 511–520. Wai, H., 2015. A study of the regulatory role of retinoic acid receptor gamma in Zebrafish development. Doctoral thesis. Aston Univ. Wang, D., Zhu, W., Chen, L., Yan, J., Teng, M., Zhou, Z., 2018. Neonatal triphenyl phosphate and its metabolite diphenyl phosphate exposure induce sex- and dose-dependent metabolic disruptions in adult mice. Environ. Pollut. 237, 10– 17. West, S.D., Burger, R.O., Poole, G.M., Mowrey, D.H., 1983. Bioconcentration and field dissipation of the aquatic herbicide fluridone and its degradation products in aquatic environments. J. Agric. Food Chem. 31, 579–585. Wiegand, C., Krause, E., Steinberg, C., Pflugmacher, S., 2001. Toxicokinetics of atrazine in embryos of the Zebrafish (Danio rerio). Ecotoxicol. Environ. Saf. 49, 199–205. Wiegand, C., Pflugmacher, S., Giese, M., Frank, H., Steinberg, C., 2000. Uptake, toxicity, and effects on detoxication enzymes of atrazine and trifluoroacetate in embryos of Zebrafish. Ecotoxicol. Environ. Saf. 45, 122–131. Yi, S.A., Francis, B.M., Jarrell, W.M., Soucek, D.J., 2011. Toxicological effects of the aquatic herbicide, fluridone, on male water mites (Hydrachnidiae: Arrenurus: Megaluracarus). Ecotoxicology 20, 81–87. Zhu, L.Z., Qi, S.Z., Cao, F.J., Mu, X.Y., Yang, Y., Wang, C., 2017. Quizalofop-P-ethyl exposure increases estrogen axis activity in male and slightly decreases estrogen axis activity in female Zebrafish (Danio rerio). Aquat. Toxicol. 183, 76–84.