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. Author manuscript; available in PMC: 2021 Jun 1.
Published in final edited form as: Aquat Toxicol. 2021 Mar 26;235:105815. doi: 10.1016/j.aquatox.2021.105815

The ecotoxicological contaminant Tris(4-chlorophenyl)methanol (TCPMOH) impacts embryonic development in zebrafish (Danio rerio)

Julian Navarrete 1, Peyton Wilson 1, Nicholas Allsing 2, Chandi Gordon 1, Rachel Margolis 1, Ashley V Schwartz 3, Christine Cho 1, Brynn Rogowski 1, Jennifer Topps 1, Uduak Z George 3, Karilyn E Sant 1
PMCID: PMC8113121  NIHMSID: NIHMS1697255  PMID: 33838494

Abstract

Tris(4-chlorophenyl)methanol (TCPMOH) is a water contaminant with unknown etiology, but is believed to be a byproduct of DDT manufacturing. It is highly persistent in the environment, and bioaccumulates in marine species. TCPMOH has also been measured in human breast milk, which poses a risk for developing infants. However, almost no toxicity data is currently available. In this study, we investigate the hazard posed by developmental TCPMOH exposures using the zebrafish model (Danio rerio). Zebrafish (Danio rerio) embryos were exposed to 0, 0.1, 0.5, 1, or 5 μM TCPMOH beginning at 24 hours post fertilization (hpf). Embryonic mortality and incidence of morphological deformities increased in a concentration-dependent manner with TCPMOH exposure. RNA sequencing assessed changes in gene expression associated with acute (4 hour) exposures to 50nM TCPMOH. Developmental exposure to TCPMOH decreased expression of ahr2, as well as metabolic enzymes cyp1a1, cyp1b1, cyp1c1, cyp1c2, and cyp2y3 (p<0.05). These findings were concordant with decreased Cyp1a1 induction measured by the ethoxyresorufin-O-deethylase (EROD) assay (p<0.05). Pathways associated with xenobiotic metabolism, lipid metabolism, and transcriptional and translational regulation were decreased. Pathways involved in DNA replication and repair, carbohydrate metabolism, and endocrine function were upregulated. Overall, this study demonstrates that TCPMOH is acutely toxic to zebrafish embryos at elevated concentrations.

Keywords: Tris(4-chlorophenyl)methanol, TCPMOH, zebrafish, embryo

1. INTRODUCTION

Tris(4-chlorophenyl)methanol (TCPMOH) is an aquatic pollutant that has been increasingly detected in global environments. TCPMOH is thought to be a metabolite or degradation product of another emerging and persistent environmental contaminant, tris(4-chlorophenyl)methane (TCPM) (Trego et al., 2019). The origin of TCPMOH is unknown, but evidence suggests that TCPM is anthropogenic, and that it is a manufacturing by-product of the organochlorine pesticide (dichlorodiphenyltrichloroethane) DDT (Buser, 1995). Recently, TCPM and TCPMOH were both detected in water near a dumping site for DDT (Kivenson et al., 2019). TCPMOH is also cited in several chemical patents for the production of synthetic polymers and the process to formulate lightfast dyes (Jarman et al., 1992).

Almost no empirical toxicokinetic or environmental fate and transport data are available for TCPMOH, so the current knowledge is based largely on quantitative structure-activity relationships models (QSARS). TCPMOH is lipophilic (LogKOW=6.372) and has a predicted bioconcentration factor of 1,692, suggesting bioaccumulation and potentially biomagnification in wildlife. The predicted half-life of TCPMOH in fish is 3.88 days, and the biodegradation half-life is 56–180 days, though additional experimentation is needed to better quantify environmental persistence. A summary of predicted physiochemical properties and kinetics is provided in Table S1.

Very few studies have investigated TCPMOH as a potential toxicant or environmental contaminant. TCPMOH has not been quantified in any water sources, drawing attention to a major gap in the existing data. TCPMOH has been readily detected in marine species (including cetaceans and mussels) and sea birds at elevated concentrations (complete list and details in Table S2). It frequently co-occurs in humans and ecological samples with the legacy pollutant dichlorodiphenyltrichloroethane (DDT) and its metabolites (Kunisue et al., 2004; Millow et al., 2015; Shaul et al., 2015; Trego et al., 2018), further suggesting that the source of TCPMOH may be related to DDT manufacture. In several studies, TCPMOH has been detected at the same or greater concentrations that DDT and metabolites (De Boer, 2000; Mackintosh et al., 2016; Shaul et al., 2015; Trego et al., 2018; Trego et al., 2019). Though the target of very few studies, TCPMOH has been detected in human adipose tissue, suggesting risk for human exposure (Minh et al., 2000). TCPMOH has also been found in human breast milk, concentrated to 23 ng/g lipid weight, suggesting the potential for developmental exposures through breastfeeding or potentially via maternal-fetal transfer during gestation (Kunisue et al., 2004). The maternal transfer of TCPMOH to offspring has been supported in several cetacean species, by as much as 84% transfer during seal gestation and lactation (Kajiwara et al., 2008b) and confirmed transplacental transfer of 3.3–5.3% TCPMOH in whales (Kajiwara et al., 2008a). Despite these known developmental exposures, no developmental toxicity data is currently available and this lack of data and risk for exposure demand the need for toxicological inquiry.

The mechanisms of toxicity for TCPMOH have not been well-characterized, especially in vivo. The limited number of published toxicology studies on TCPMOH toxicity suggest that it modulates endocrine activity—namely via androgen receptor binding, changes in androgenic function, or hydroxylation of estradiol (Foster et al., 1999; Körner et al., 2004; Schrader and Cooke, 2002; Segura-Aguilar et al., 1997). It is also found to induce cytochrome P450 activation (namely Cyp1A1 and Cyp1B1) and other phase I biotransformation enzymes, which together are common indicators of polycyclic aromatic hydrocarbon (PAH) exposures or activation of the aryl hydrocarbon receptor (AHR) pathway (Poon et al., 1997; Trego et al. 2019). Phase II biotransformation enzymes, primarily those catalyzing conjugation and biosynthesis or the endogenous antioxidant glutathione, were also increased by TCPMOH exposures (Poon et al., 1997). These studies were all conducted in vitro or in adult rodents, and the aquatic and developmental toxicity of TCPMOH remains uncharacterized.

The goal of this study is to assess the aquatic toxicity of TCPMOH and its impact on embryonic development in the zebrafish model (Danio rerio). We use non-targeted methods to assess mechanisms of aquatic toxicity, as well as a priori methods such as targeted gene expression and enzymatic assays based upon structural similarity to other halogenated organic compounds (HOCs). We hypothesized that TCPMOH would impair embryonic development, and that activation of common phase I and II detoxification pathways would occur due to exposures.

2. METHODS

2.1. Chemicals

Tris(4-chlorophenyl)methanol (TCPMOH; CAS #3010-80-8) was purchased from Sigma Aldrich (St. Louis, MO). Dimethyl sulfoxide (DMSO) and 7-Ethoxyresorufin were purchased from Fisher Scientific (Pittsburgh, PA). Stock solutions of TCPMOH [1–50 mM] for embryonic exposures were prepared in DMSO and stored at room temperature in amber glass vials away from light until use. All experimental procedures involving TCPMOH were performed using appropriate safety precautions.

2.2. Zebrafish Husbandry & Care

Wild-type (AB) strain zebrafish were originally given to San Diego State University from the University of California San Diego. Prior to the study, adult zebrafish were housed in an Aquaneering zebrafish system at San Diego State University in the Environmental Health Laboratory. Fish were provided the recommended amount of GEMMA Micro 300 powdered diet once daily (Skretting; Westbrook, ME). Temperature was maintained at 28°C, pH 7.2–7.3, conductivity 620–700, and light cycling was maintained at a 12:12 light:dark cycle. Nitrates, nitrites, ammonia, and chlorine were measured weekly. Breeding tank populations contained 15–20 adult fish (2:3 male:female ratio). All animal use protocols have been approved by the San Diego State University Institutional Animal Care & Use Committee and meet or exceed all recommended practices for zebrafish care (PHS Assurance Number 16–00430).

Embryos were collected from breeding tanks between 0–1 hours post-fertilization (hpf), washed, and housed in clean polystyrene dishes containing 0.3X Danieau’s medium (17 mM NaCl, 2 mM KCl, 0.12 mM MgSO4, 1.8 mM Ca(NO3)2, 1.5 mM HEPES, pH 7.6). At 6–8 hpf, embryos were sorted for viability and quality, and embryos from different breeding tanks were consolidated and then randomized into clean petri dishes with fresh 0.3X Danieau’s medium and incubated at 28.5°C overnight on a 12:12 hour light:dark cycle.

2.3. Exposures

At 24 hpf, embryos were manually dechorionated using watchmakers’ forceps prior to the initiation of all experiments. Embryos were incubated at 28.5°C throughout the duration of experiments on a 12:12 hour light:dark cycle.

For all microscopy experiments, embryos were individually transferred to untreated 24-well polystyrene plates, with one embryo per well. Each well contained 1 mL of 0.3X Danieau’s medium that had been previously constituted with 0.01% v/v DMSO (vehicle control), 0.1 μM, 0.5 μM, 1 μM, or 5 μM TCPMOH. These concentrations are supra-environmental and were selected to discover the range of acute developmental toxicity and mortality and based on preliminary studies. Exposure media were 95% refreshed daily from 1–7 days post fertilization (dpf) to mimic subchronic developmental exposure. Experiments were replicated 5 times, each with biological replicates of 10–12 embryos per exposure group.

For Ethyoxyresorufin-O-deethylase (EROD) assays, dechorionated 24 hpf embryos were transferred into borosilicate glass scintillation vials. Each vial contained 5 embryos in 5 mL of 0.3X Danieau’s concentrated with DMSO (0.05% v/v) or TCPMOH (0.5 μM or 1 μM). 7-ethoxyresorufin (ER-7) was added to the media at a concentration of 0.25 mg/mL. Vials were covered and static exposures were maintained until 4 dpf. Experiments were replicated 3 times, each with 3–5 embryos per exposure group.

For RNA sequencing experiments following acute exposures, embryos were maintained in 0.3X Danieau’s medium in clean 100 mm polystyrene dishes until 96 hpf, refreshing the water daily. At 96 hpf, embryos (all hatched) were transferred in groups of 10 embryos into clean 100 mm polystyrene dishes containing 30 ml of 0.3X Danieau’s medium. Concentrated stock solutions were added to the dishes to produce final exposure concentrations of 0.01% v/v DMSO (control) or 50 nM TCPMOH. Embryos were placed into a 28.5°C incubator for 4 hours, and embryos were collected for RNA isolation at 100 hpf. Experiments were replicated 5 times, each with 10–12 embryos per exposure group.

2.4. Microscopy

Wild-type (AB) embryos were imaged daily from 1–7 dpf to observe mortality, growth, swim bladder inflation, and incidence of structural deformities such as pericardial and yolk sac edema, craniofacial deformities, spinal deformities, and intestinal effusion. For all morphology experiments, the 5 μM exposures to TCPMOH were excluded due to the high mortality. All imaging was performed using a Nikon Ti-2 inverted microscope. Embryos were washed thoroughly and momentarily anaesthetized in 2% v/v MS-222 solution (prepared as 4 mg/mL Finquel tricaine powder in water, pH buffered, and stored at −20°C until use). Embryos were mounted in drops of 3% methylcellulose for imaging and oriented lengthwise. Brightfield images were acquired at 20X and 40X magnification for experiments examining morphology and growth. All measurements were quantified using the Nikon NIS Elements Advanced Research software.

2.5. EROD Assay

EROD activity in zebrafish embryos was used as an in vivo biomarker to measure the catalytic induction of cytochrome P4501A (Cyp1a). At 4 dpf, embryos were washed thoroughly and momentarily anaesthetized in 2% v/v MS-222 solution. Embryos were mounted in drops of 3% methylcellulose for imaging and oriented lengthwise. Images were acquired using 2X brightfield, 10X brightfield, and 10X RFP filter using the Nikon Ti-2 inverted epifluorescent microscope. RFP laser intensity, exposure time, and gain parameters were maintained the same across all fish, and fluorescence was normalized to background intensity.

2.6. RNA isolation

RNA was extracted from 4 dpf embryos (pools of 8–10 embryos) for RNA sequencing and qPCR. Embryos were collected into RNAlater and stored at −20 °C until RNA isolation. A total of 5 samples per exposure group was collected for RNA sequencing and 3–4 samples per exposure group was collected for qPCR, and each sample was obtained from a separate experimental replicate. RNAlater was removed from each sample and kept on ice prior to processing. Samples were briefly pulse sonicated using a Branson SFX250 Sonifier at 10% amplification, and RNA was processed with the GeneJET RNA Purification Kit (Fisher Scientific) according to manufacturer protocols. Extracted RNA for RNA sequencing was stored at −80°C until use. Extracted RNA for qPCR was first converted to cDNA then amplified using qPCR (detailed in Supplemental Information, Table S3).

2.7. Library preparation and RNA sequencing

RNA quality, library preparation, and sequencing were all performed at the University of California San Diego Institute for Genomic Medicine Genomics Center (San Diego, CA). RNA quantity and quality were assessed using Agilent 4200 TapeStation (Agilent Technologies, Santa Clara, California). RNA Integrity Numbers (RINs) ranged from 7.8 to 9.8, and RNA concentrations ranged from 130–270 ng/μl (Table S4). Stranded libraries were prepared using the Illumina Stranded mRNA Prep Kit following manufacturer instructions with Poly(A) enrichment (Illumina, San Diego, CA). Libraries were sequenced using the Illumina NovaSeq 6000 platform using paired end 100 bp (PE100; 2×100bp) reads to a sequencing depth of 25M reads.

2.8. Bioinformatics

RNAseq FASTQ files were imported on a local instance of Galaxy and labeled according to treatment (Afgan et al., 2018). FastQC was used to generate a reading of the quality of the sequences, and Cutadapt was used to trim and filter out any low-quality sequences (Andrews, 2010; Martin, 2011). The parameters for Cutadapt included settings for paired-end reads, a minimum filter length of 20, and a quality cutoff of 20. A zebrafish genome file and a transcriptome annotation file were then obtained from the Lawson Lab at UMASS Medical School (v. 4.3.2) (Lawson et al., 2020). This annotation file improves upon the zebrafish reference genome assembly (GRCz11) from 2017, adding more comprehensive transcript annotation. These files were aligned and annotated using the Spliced Transcripts Alignment to a Reference (STAR) software (Dobin et al., 2012). In order to count the number of reads per annotated gene, the featureCounts program was executed using the output from STAR (Liao et al., 2013). DESeq2 software was then used to find the differentially expressed features between the treatments (Love et al., 2014). Significantly changed genes were defined as those with DEseq2 FDR values <0.05 and also have expression changes of >15%. The DESeq2 output was then uploaded to the RNA-Enrich version of LRpath in order to test for biologically-relevant gene sets, specifically using the KEGG Pathway database (Kim et al., 2012; Lee et al., 2016). Gene Ontology enrichment analysis was performed using FishEnrich (Chen et al., 2013; Kuleshov et al., 2016). Alterations to gene expression and pathway enrichment by exposure were determined by using the Benjamini-Hochberg adjusted p-value (p<0.05). Data has been deposited into the Gene Expression Omnibus (GEO) and can be viewed using the accession number GSE165920.

2.9. Statistical analysis

Levene’s tests and Welch’s tests were performed to inform the appropriate statistical test for means comparisons. All normally distributed data is presented as the mean ± standard error, and assessed using ANOVAs with Tukey post-hoc test or independent t-tests. For non-parametric data in the EROD experiment, Kruskall-Wallis nonparametric test with Games-Howell post-hoc test were used to compare exposure groups and data is presented as quantiles. For percent survival and incidence of structural defects, Fisher’s exact tests were used. Probit was used to estimate LC50 concentrations based on empirical data. A confidence level of 95% (α=0.05) was utilized. IBM SPSS v26 was used for data analysis.

3. RESULTS

3.1. Survival

We aimed to assess the total survival and the presence of morphological defects in zebrafish exposed to 0.01% v/v DMSO (control), 0.1, 0.5, 1, or 5 μM TCPMPOH in order to gain preliminary understanding of toxicity (Figure 1). Embryo survival was not impacted in exposures to DMSO nor 0.1 μM TCPMOH. Total survival decreased as early as 2 dpf for embryos most highly exposed (5 μM), and a significant reduction was observed beginning at 4 dpf (p<0.05). No other mortality was observed prior to 4 dpf. By 5 dpf, there was significantly reduced total survival of embryos exposed to 1 μM TCPMOH, and the same was true at 6 dpf for those exposed to 0.5 μM TCPMOH (p<0.05).

Figure 1.

Figure 1.

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Survival (%) of zebrafish embryos exposed to 0 (DMSO), 0.1, 0.5, 1, or 5 μM TCPMOH daily until 7 dpf. Total survival decreases temporally and in a concentration-dependent manner throughout the 7 day period (p<0.05). Fisher’s exact tests were used to assess differences in frequencies between groups. Asterisks (*) indicate statistically significant changes from controls (p<0.05). n=20–38 embryos per exposure group

The median lethal concentrations (LC50) were estimated for each day of exposure starting on 2–7 dpf (Table S5) using probit regression. Though estimated LC50 values were above the empirical range from 1–4 dpf, there was a significant decrease of LC50 beginning at 4 dpf (p<0.001). The LC50 concentrations were 3.99, 2.47, and 1.02 μM TCPMOH for 5, 6, and 7 dpf, respectively (p<0.001).

3.2. Fish Length

To assess the impact of TCPMOH exposures on embryonic growth, total anterior-posterior fish length was measured daily from 2–5 dpf (Figure 2). Because measurements from 6–7 dpf were largely influenced by mortality, these measurements were excluded from the analysis. Overall, embryos exposed to the highest concentration of TCPMOH (5 μM) displayed stunted growth compared to controls (p<0.05). This growth deficiency was particularly apparent at 5 dpf, when embryos exposed to 5 μM TCPMOH were 7.2% smaller on average than controls. No other statistically significant trends in growth and exposure were observed over time (p>0.05).

Figure 2.

Figure 2.

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Total anterior-posterior fish length in embryos exposed to 0.01% DMSO, 0.1, 0.5, 1, or 5 μM TCPMOH. Embryos exposed to 5 μM TCPMOH displayed stunted growth compared to controls (p<0.05). No other statistically significant trends were observed (p>0.05). ANOVA with Tukey post-hoc tests were used. Asterisks (*) indicate statistically significant changes from controls (p<0.05). n=20–38 embryos per exposure group

3.3. Embryonic morphology

Embryos were individually examined daily for pericardial edema, yolk edema, swim bladder inflation, and craniofacial malformations in exposure groups of DMSO (control), 0.1, 0.5, or 1 μM TCPMOH (Figure 3). Incidence of pericardial edema and yolk edema was observed for embryos as early as 2 dpf, and subsequent exposures increased the percentage of these structural defects. At 5 dpf, embryos exposed to 1 μM had a significantly higher incidence of pericardial edema, yolk edema, craniofacial malformations, and decreased swim bladder inflation as compared to controls (p<0.01 for all deformities; Figures 3A–D). Swim bladder inflation decreased in a concentration-dependent manner beginning at 4 dpf, with reduced inflation in embryos exposed to 1 μM TCPMOH at 4 dpf (p<0.001; Figure 3C). At 5 dpf, all TCPMOH-exposed embryo groups had reduced swim bladder inflation compared to controls (p<0.022 for all groups). Yolk and pericardial edema frequently co-occurred in embryos (11.3% of total embryos), which are classic indicators of blue sac disease.

Figure 3.

Figure 3.

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TCPMOH exposures increase the incidence of embryonic defects and decrease swim bladder inflation. Embryos were individually examined microscopically, daily from 1–7 dpf. The most common deformities presented were pericardial edema (A) and yolk edema (impaired yolk utilization) (B) manifesting as an inflated pericardial sac due to fluid retention and inflated, malformed yolk sac, respectively. Decreased (C) swim bladder inflation occurred in a concentration-dependent trend. Increased incidence of craniofacial malformations (D), presenting mostly as lower jaw deformities, were increased in the 1 μM exposure group. Fisher’s exact tests were used to assess differences in frequencies between groups. Asterisks (*) indicate statistically significant changes from controls (p<0.05). n=20–38 embryos per exposure group

3.4. EROD activity

EROD activity was quantified in 4 dpf zebrafish embryos at 96 hpf following static exposures to 0 (DMSO), 0.5 or 1 μM TCPMOH (Figure 4). EROD activity is a useful in vivo biomarker of cyp1a1 activity through the rate of cyp1a1 mediated deethylation of 7-ER (Jönsson et al., 2009). Induction of cyp1a1 can be proportionally quantified by 7-ethoxy-resorufin-O-deethylase (EROD) activity in the presence of 7-Ethoxyresorufin (7-ER), and activity is directly proportional to RFP fluorescence. Data was normalized to background fluorescence and intensity. There was a concentration-dependent decrease of in vivo CYP1A1 activity with significant attenuation in embryos exposed to 1 μM (p=0.019). Median fluorescence was decreased by 31% and 38% in embryos exposed to 0.5 and 1 μM TCPMOH, respectively.

Figure 4.

Figure 4.

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Embryos were exposed to 0, 0.5, or 1 μM TCPMOH once at 24 hpf in the presence of 0.25 mg/mL 7-ethoxyresorufin (7-ER) in DMSO. At 96 hpf, embryos were screened for EROD activity, as quantified by red fluorescent protein (RFP) intensity. Boxplots represent the median (quartile) relative RFP intensity to the background intensity and control (DMSO) group. Boxes represent true quartiles (25th, 50th, 75th percentile). Kruskal-Wallis test with Games-Howell post-hoc tests were used for nonparametric data. Images are shown for (A) DMSO and (B) 0.5 μM TCPMOH exposed fish. Asterisks (*) indicate statistically significant changes from controls (p<0.05). n=9–11 embryos per exposure group.

3.5. Gene expression following acute exposures (RNA sequencing)

We examined comparative gene expression to assess the most significantly impacted genes by 4-hour 50 nM TCPMOH exposure. The 10 most significant upregulated and 10 most significantly downregulated genes are shown (Table S6). The complete DEseq2 results can be found in GEO (accession number GSE165920). In total, 516 genes were significant changed by TCPMOH exposure (FDR<0.05), 370 of which were downregulated and 146 upregulated (Figure 5). Processes related to ribosomal function and protein folding, RNA processing, cholesterol transport, and responses to xenobiotic and light stimuli were enriched due to TCPMOH exposures (Table 1). Pathways significantly impacted by acute TCPMOH exposure were also examined using the Kyoto Encyclopedia of Genes and Genomes (KEGG; https://www.genome.jp/kegg/). A complete list of pathways significantly affected by exposure (p<0.05) and the direction and magnitude of their association are presented (Table S7).

Figure 5.

Figure 5.

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Overall distribution and altered expression of transcripts within the AHR Xenobiotic Response and Nrf2 Antioxidant Response pathways. Log2 fold change values and p-values (FDR) were graphed as a volcano plot, showing that the majority of significantly impacted genes were downregulated. Within the AHR and Nrf2 pathways, transcription factors ahr2 and nfe2l2b were both decreased, as were their enzymatic targets. FDR<0.05

Table 1.

Gene Ontologies significantly enriched due to TCPMOH exposure (50 nM) from 96–100 hpf.

Gene Ontology Term Adjusted P-value Combined Score
GO Biological Process 2018 response to light stimulus (GO:0009416) 3.13E-09 34.93
ribosome biogenesis (GO:0042254) 0.0002 22.40
photoperiodism (GO:0009648) 0.0005 37.23
entrainment of circadian clock by photoperiod (GO:0043153) 0.0059 35.61
maturation of SSU-rRNA (GO:0030490) 0.0064 20.36
negative regulation of cellular macromolecule biosynthetic process (GO:2000113) 0.0066 13.68
negative regulation of nucleic acid-templated transcription (GO:1903507) 0.0066 10.19
maturation of SSU-rRNA from tricistronic rRNA transcript (SSU-rRNA, 5.8S rRNA, LSU-rRNA) (GO:0000462) 0.0076 21.80
rRNA metabolic process (GO:0016072) 0.0076 19.83
ncRNA processing (GO:0034470) 0.0079 18.10
rRNA processing (GO:0006364) 0.0079 15.42
maturation of LSU-rRNA from tricistronic rRNA transcript (SSU-rRNA, 5.8S rRNA, LSU-rRNA) (GO:0000463) 0.0087 27.61
cellular response to radiation (GO:0071478) 0.0104 14.80
cellular response to light stimulus (GO:0071482) 0.0104 12.46
negative regulation of gene expression (GO:0010629) 0.0104 9.42
negative regulation of transcription, DNA-templated (GO:0045892) 0.0139 9.44
pyrimidine dimer repair (GO:0006290) 0.0157 39.35
cholesterol efflux (GO:0033344) 0.0368 15.34
response to hydrogen peroxide (GO:0042542) 0.0379 29.60
chaperone mediated protein folding requiring cofactor (GO:0051085) 0.0417 17.71
negative regulation of lymphocyte activation (GO:0051250) 0.0424 23.29
thymus development (GO:0048538) 0.0424 20.23
‘de novo’ posttranslational protein folding (GO:0051084) 0.0424 14.23
response to reactive oxygen species (GO:0000302) 0.0424 14.12
cellular response to xenobiotic stimulus (GO:0071466) 0.0424 14.03
GO Cellular Component 2018 preribosome (GO:0030684) 0.0004 20.91
nucleolus (GO:0005730) 0.0004 14.67
mitochondrial intermembrane space (GO:0005758) 0.0112 21.42
mitochondrial envelope (GO:0005740) 0.0112 20.06
preribosome, large subunit precursor (GO:0030687) 0.0281 16.21
small-subunit processome (GO:0032040) 0.0281 8.07
nuclear lumen (GO:0031981) 0.0281 6.96
GO Molecular Function 2018 snoRNA binding (GO:0030515) 0.0002 37.03
hydrolase activity, acting on acid anhydrides, catalyzing transmembrane movement of substances (GO:0016820) 0.0067 19.21
P-P-bond-hydrolysis-driven transmembrane transporter activity (GO:0015405) 0.0067 18.62
ATPase activity, coupled to transmembrane movement of substances (GO:0042626) 0.0067 14.78
G-protein coupled photoreceptor activity (GO:0008020) 0.0311 14.76
ATPase activity, coupled to movement of substances (GO:0043492) 0.0314 12.85
cholesterol transporter activity (GO:0017127) 0.0340 15.95
transcription cofactor binding (GO:0001221) 0.0476 21.55
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Two key pathways involved in the metabolic response to common HOCs were significantly downregulated: “Metabolism of xenobiotics by cytochrome P450” and “Glutathione metabolism”. Because these pathways are often highly inducible, we had expected gene expression within these pathways to be increased a priori. To more closely examine this relationship, gene expression within the AHR Xenobiotic Response pathway and Nrf2 Antioxidant Response pathway was assessed (Tables S8–S10). Expression of the ahr2 transcription factor was significantly decreased, as well as expression of AHR targets and cytochrome P450 isoforms cyp1a1, cyp1b1, cyp1c1, cyp1c2, cyp2y3. Only expression of arntl1a and arntl1b, two nuclear factors involved in the coordination of circadian rhythm in response to lipids and xenobiotics, were upregulated. No genes were significantly upregulated within the Nrf2 Antioxidant Response pathway. Decreased gene expression of transcription factor nfe2l2b (“Nrf2b”), rate-limiting glutathione biosynthesis enzyme gclc, and two glutathione-S-transferases gsta.1 and gstcd occurred. Additionally, “GnRH signaling pathway,” “Melanogenesis,” and “Progesterone-mediated oocyte maturation” were increased due to TCPMOH exposure, so we examined gene expression within commonly modulated endocrine activity pathways more closely (Table S10). No genes commonly associated with modulated endocrine activity, such as estrogenic or androgenic function, were significantly impacted by exposures.

KEGG Pathway hierarchies were then annotated to examine overall patterns due to TCPMOH exposures (Figure 6). All Metabolism pathways were decreased by exposure, except for those related to carbohydrate metabolism. All Environmental Information Processing pathways, related to signal transduction processes such as Wnt and Hedgehog signaling, were upregulated. All Cellular Processes pathways, related to cell cycle regulation and cell-cell interactions, were also significantly upregulated. Though variable responses were observed for the Organismal Systems pathways, a cluster of those related to endocrine function were upregulated. For Genetic Information Processing pathways, processes related to DNA replication and repair were all upregulated, while pathways related to overall transcriptional and translational regulation were decreased.

Figure 6.

Figure 6.

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KEGG Pathways most significantly changed due to acute TCPMOH exposure during development. The ten most significantly up- and downregulated pathways are shown. The KEGG pathway hierarchies were used to group pathways by overarching processes, including cellular processes, environmental information processing, genetic information processing, metabolism, and organismal systems. Regression coefficients and p-values were plotted, with coefficients >0 indicating upregulated pathways and coefficients <0 indicating downregulated pathways. n=5 samples per treatment group, each containing 8–10 embryos

4. DISCUSSION

The goal of this study was to report the aquatic toxicity of TCPMOH using zebrafish embryos as a model. Because of its structural similarity to other highly toxic HOCs, we had hypothesized that TCPMOH would impair embryonic development, activating phase I and phase II metabolic enzymes such as Cyp1a. Our results demonstrate that TCPMOH is acutely toxic at elevated concentrations and provide concentration-response data for aquatic developmental toxicity. However, gene expression and EROD activity assays suggest that Cyp1a may be impaired by TCPMOH exposures.

TCPMOH significantly increased mortality and structural defects in embryos throughout the developmental period (Figure 1), and embryonic growth was significantly impacted only by the highest TCPMOH concentration (Figure 2). The magnitude and timing of mortality was increased in a concentration-dependent manner with increasing exposures. Beginning at 4 dpf, survival was reduced in the most highly exposed groups, and each subsequent day increased incidence in a concentration-dependent manner. This trend was also confirmed for several structural deformities (yolk sac edema, pericardial edema) as well as for impaired swim bladder inflation (Figure 3). This data correlates increasing exposures with decreased embryonic success, but also suggests a temporal contribution to the observed pathologies. Most incidence rates were not significantly increased until 4 dpf, the timing at which several major organs become physiologically and metabolically active—including the liver. The liver is a major metabolic target of HOCs, coordinating a number of biotransformation pathways for these highly lipophilic compounds. To better understand the temporal effects observed in the current study, information about the adsorption, distribution, metabolism, and excretion of TCPMOH is needed to infer target organ toxicity.

The occurrence of pericardial edema and yolk sac edema in fish embryos are hallmarks of blue sac disease. Blue sac disease is a somewhat common fry pathology, frequently attributed to a number of environmental conditions including water quality (Wolf, 1957). In this study, incidence of pericardial edema and yolk sac edema were both elevated (Figure 3), but additionally, we showed co-occurrence of these pathologies in more than 11% of exposed embryos. Pericardial edema and yolk sac edema are frequently observed following embryonic exposures to other HOCs, including TCDD and PCBs, and often these hallmarks manifest with other pathologies, including reduced swim bladder inflation as shown (Figure 3). One of the best predictors of blue sac disease in fish is the continuous induction of Cyp1a (Brinkworth et al., 2003; Brown et al., 2015; Hornung et al., 1999; Rousseau et al., 2015). Interestingly, our data show decreased gene expression of cyp1a1 following both acute (4-hour) and developmental exposures, and these changes also correlated with decreased EROD activity (Table S8, Figures 4 and S1). The presence of these pathologies without the signature Cyp1a activation was surprising, and it raises questions about the mechanism of TCPMOH toxicity.

Based upon the xenobiotic responses of similar PAHs (namely HOCs), we more closely examined the gene expression of phase I detoxification enzymes as well as Cyp1a activity (Table S7, Figures 4 and S1). Our data suggests that TCPMOH may actually antagonize or inhibit Cyp1a. Other studies have also shown that other HOCs, such as ortho-substituted polychlorinated biphenyls, may inhibit AHR signaling and/or Cyp1a function (Schlezinger et al., 2006; Suh et al., 2003). Metabolic compensation by other metabolic (Phase I) enzymes was not observed, as expression of other CYP enzymes was alse decreased after acute exposure (Table S8). However, cyp2e1 expression was increased following prolonged developmental exposures but not acute exposures (Figure S1), suggesting that this is not likely to be a direct effect of acute TCPMOH toxicity. Our data conflicts with a published study using a rat model, showing increased Cyp1a expression in the liver with increasing TCPMOH exposures (Poon et al., 1997). Interspecies physiologic differences including different sites of xenobiotic absorption, tissue lipid concentrations, and species-specific metabolic pathways may explain these differing Cyp1a responses. For example, hepatocyte differentiation and physiologic function are established between 3–5 dpf in zebrafish, so gene expression analysis at 4 dpf may not fully capture liver metabolic response and function (reviewed in Wilkins and Pack, 2013). Additionally, gene expression of the transcription factor ahr2 was also decreased in our study, which is the AHR ortholog most like the mammalian AHR (Andreasen et al., 2002; Christen and Fent, 2014). Expression of ahr1a and ahr1b were unaffected by exposures, which was expected since these homologs are not expected to play a major role in the xenobiotic response to PAHs (Andreasen et al., 2002; Garner et al., 2013; Karchner et al., 2005; Souder and Gorelick, 2019). Together, our data suggests that the metabolic response to TCPMOH may not be directly coordinated by the AHR signaling pathway. There is need to assess whether TCPMOH binds to the AHR in future studies, or whether other independent pathways coordinate the response to TCPMOH.

We also examined the contributions of several phase II detoxification genes in response to embryonic TCPMOH exposures (Table S9, Figure S1). Gene expression of several genes involved in the antioxidant response was significantly decreased, including several heat-shock proteins and glutathione-S-transferase genes. The rate-limiting enzyme for glutathione biosynthesis, gclc, was decreased by exposure, as was nfe2l2b (“Nrf2b”), the teleost-specific duplicate of the mammalian Nrf2—a transcription factor coordinating the adaptive antioxidant response. Decreased expression across all of these antioxidant response targets suggests that the Nrf2 pathway may not play an important role in the immediate response to TCPMOH in embryos. Gene expression of nrf2a, the Nrf2 homolog most similar to the mammalian Nrf2 (Timme-Laragy et al., 2012), was unaffected by acute TCPMOH exposures (Table S9) though expression was increased in embryos following daily developmental exposures (Figure S1). Interestingly, we also observed increased expression of Nrf2a target hmox1 following these longer developmental exposures. This suggests that the antioxidant response may not be immediately or directly involved in the response to TCPMOH, but that these processes may become indirectly activated following more chronic exposures. Additional work examining the temporal response to TCPMOH is required to understand the coordination of these mechanisms.

Several studies have shown that TCPMOH exposures may produce endocrine activity in mammals (Foster et al., 1999; Körner et al., 2004; Schrader and Cooke, 2002; Segura-Aguilar et al., 1997). Here, we found that several endocrine pathways were upregulated by TCPMOH exposures, including GnRH signaling pathway, Melanogenesis, and Progesterone-mediated oocyte maturation (Table S7). Most of the significantly impacted genes in these pathways were calmodulins, kinases, or phosphatases specific to regulation of circadian rhythm and photoperiod. When common hormone targets associated with estrogenic, androgenic, or other classic endocrine pathways (such as thyroid and pancreatic function) were examined, there were no significant changes in gene expression (Table S10). However, swim bladder inflation was particularly impaired due to TCPMOH exposures (Figure 3). Swim bladder inflation is a sensitive indicator of xenobiotic exposures and can also be an indicator of endocrine activity, namely disruption of thyroid hormone function (Godfrey et al., 2017; Stinckens et al., 2020; reviewed in Price and Mager, 2020). Here, genes associated with thyroid function were not significantly impacted by acute TCPMOH exposure (Table S10), and therefore failed swim bladder inflation is not likely to be the result of thyroid hormone disruption.

The concentrations of TCPMOH used in this study were likely supra-environmental and selected to be able to assess acute toxicity in this sentinel study. However, there is very limited information about the prevalence and persistence of TCPMOH or TCPM in environmental matrices. A study conducted by Falandysz et al. quantified TCPM and TCPMOH in Baltic coastal fish and sediments, finding elevated concentrations in marine species but nondetectable levels in marine sediments (Falandysz et al., 1999). The authors attributed this to a high TCPMOH bioaccumulation factor, though the bioconcentration kinetics require empirical elucidation. This is supported by the growing evidence of high TCPMOH concentrations found in marine species globally (detailed in Table S2). Kivenson et al (2019) investigated ocean contaminants co-occurring near DDT dumping sites off of the California coast. The authors detected elevated TCPM and TCPMOH concentrations near these sites using relative quantification methods (non-targeted GC × GC–HR–ToF–MS), suggesting that TCPM and TCPMOH may be contaminants of emerging concern along the Pacific coast. However, these measurements were relative, and additional research is needed to provide absolute quantification of TCPM and TCPMOH in coastal waters. This information is crucial to conduct risk assessments for these compounds, but also to help scientists uncover how species are exposed and to identify their sources globally.

A limitation of this work is the lack of data related to TCPMOH kinetics. Here, we did not absolutely quantify TCPMOH concentrations in our samples, nor the variance between exposures. Instead, we spiked known (but unverified) concentrations of TCPMOH into our media using controlled, laboratory-created solutions that were freshly prepared on a regular basis. Because of the low water solubility, high bioconcentration factor, and half-life longer than the study duration, this study assumed near-complete uptake of TCPMOH from the media. Without absolute quantification, the actual exposures to TCPMOH in embryos and related toxicity and risk are uncertain or relative. Analytical research is necessary for future studies looking to characterize the risks posed by TCPMOH.

In conclusion, this study characterizes the developmental toxicity of a marine contaminant, TCPMOH, using the zebrafish model. To our knowledge, this is the first study to assess the aquatic toxicity of these compounds in embryos. Elevated exposures to TCPMOH increased embryonic mortality, incidence of structural defects, and modulated gene expression and activity of enzymes involved in detoxification. We provide empirical concentration-response data, providing a foundation for future toxicological research into the mechanisms and physiological consequences of TCPMOH exposures.

Supplementary Material

supplemental data

HIGHLIGHTS.

  • Tris(4-chlorophenyl)methanol (TCPMOH) is an aquatic pollutant, found globally in marine species.

  • TCPMOH increased embryonic mortality and the incidence of pericardial edema and yolk sac edema in zebrafish embryos from 1–7 days post fertilization (dpf).

  • Exposures to TCPMOH reduced Cyp1a activity in 4 dpf embryos, which was concordant with cyp1a1 and ahr2 gene expression.

  • Pathways associated with carbohydrate metabolism, endocrine function, and DNA replication and repair were upregulated, while pathways related to xenobiotic and lipid metabolism, transcriptional and translational regulation were decreased.

ACKNOWLEDGEMENTS

This research was supported by the National Institutes of Health (K01ES031640), the San Diego State University Grants Program, and student support was provided in part by the CSUPERB-Howell Scholars program (to JN and AVS). The authors would like to acknowledge the outstanding animal husbandry provided by Pria Bose, Mia Bowers, Alex Fox, Alexa Garcia, Shawnee Huang, Sarah McHenry, and Sydney Stegman. This publication includes data generated at the UC San Diego IGM Genomics Center utilizing an Illumina NovaSeq 6000 that was purchased with funding from a National Institutes of Health SIG grant (#S10 OD026929)

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