Modulation of PPAR signaling disrupts pancreas development in the zebrafish, Danio rerio

Abstract

Peroxisome Proliferator Activated Receptors (PPARs) are transcription factors that regulate processes such as lipid and glucose metabolism. Synthetic PPAR ligands, designed as therapeutics for metabolic disease, provide a tool to assess the relationship between PPAR activity and pancreas development in vivo, an area that remains poorly characterized. Here, we aim to assess the effects of PPAR agonists and antagonists on gene expression, embryonic morphology and pancreas development in transgenic zebrafish embryos. To evaluate developmental perturbations, we assessed gross body and pancreas morphology at 4 days post fertilization (dpf) in response to developmental exposures with PPARα, PPARγ, and PPARβ/δ agonists and antagonists at 0, 0.01, 0.1, 1, and 10 μM concentrations. All ligand exposures, with the exception of the PPARα agonist, resulted in significantly altered fish length and yolk sac area. PPARγ agonist and antagonist had higher incidence of darkened yolk sac and craniofacial deformities, whereas PPARα antagonist had higher incidence of pericardial edema and death. Significantly reduced endocrine pancreas area was observed in both PPARγ ligands and PPARα agonist exposed embryos, some of which also exhibited aberrant endocrine pancreas morphology. Both PPARβ/δ ligands caused reduced exocrine pancreas length and novel aberrant phenotype, and disrupted gene expression of pancreatic targets pdx1, gcga, and try. Lipid staining was performed at 8 dpf and revealed altered lipid accumulation consistent with isoform function. These data indicate chronic exposure to synthetic ligands may induce morphological and pancreatic defects in zebrafish embryos.

INTRODUCTION

Embryogenesis and organogenesis are tightly controlled processes, mediated by closely coordinated cell signaling pathways. However, exogenous cues such as chemical exposure and malnutrition can disrupt proliferation, differentiation and apoptosis, which are all crucial procedures during development. Though the most severe consequences of these miscues are structural deformities and embryonic mortality, more subtle changes may also occur such as behavioral or metabolic perturbations. One such signaling pathway heavily involved in metabolic regulation is the Peroxisome Proliferator-Activated Receptors (PPARs) signaling pathway, a family of nuclear hormone receptors essential for regulating lipid homeostasis during development and throughout adulthood (Rees et al., 2008). In response to ligand activation by endogenous fatty acids, PPAR heterodimerizes with retinoic acid receptor α (RXRα) to bind peroxisome proliferator responsive element (PPRE) promoter sequences for transcription of genes related to fatty acid metabolism, differentiation, and proliferation (Monsalve et al., 2013; Lee, 2017). Though disruption of PPAR signaling may impact these processes through immediate action, alterations to metabolic programing are shown to persist into adulthood and have been linked to pathologies such as obesity and glucose intolerance (Erhuma et al., 2007; Lillycrop et al., 2008; Rees et al., 2008).

There are three mammalian isoforms of PPAR: PPARα, PPARγ and PPARβ/δ. PPARα is expressed in cells with high rates of fatty acid catabolism, such as hepatocyte, cardiomyocytes, and cells of the intestinal mucosa (Miyachi et al., 2002). PPARγ is widely known for its expression in adipose tissue and its pivotal role in adipocyte differentiation (Braissant and Wahli, 1998). Unlike gamma and alpha, PPARβ/δ is ubiquitously expressed at lower levels, and not as widely researched, making its primary function more elusive. However, this isoform has been hypothesized to regulate expression of other PPAR isoforms (Shi et al., 2002) and is suspected to function in embryo implantation and organogenesis as it is expressed very early in embryonic development (Barak et al., 2002; Lee et al., 2009).

PPARs’ widespread roles in maintaining lipid homeostasis links their disrupted activity to forms of metabolic disease. Inefficient ability to properly store and metabolize fatty acids leads to fat accumulation in non-adipose tissue, which can eventually lead to lipotoxicity. This phenomenon, co-occurring with inflammation and oxidative stress, results in cellular apoptosis and impaired tissue function (Sharma and Alonso, 2014). The pancreas, a primary metabolic organ, is sensitive to fatty acid accumulation and oxidative stress (Sant et al., 2016). Lipid accumulation in pancreatic tissue will result in damage to beta cells, with disease implications for diabetes (Matsuda et al., 2014). Additional damage to acinar and ductal cells of the exocrine pancreas will further impair nutrient storage and breakdown (Yang and Li, 2012).

PPARs have become an increasingly prevalent pathway for understanding mechanisms of metabolic disorders. In adulthood, PPARα and PPARβ/δ knockout mice have impaired insulin secretion (Sugden and Holness, 2004; Iglesias et al., 2012) and in vitro PPARγ activation induces insulin secretion in rat insulinoma INS-1 cells (Kim et al., 2012). Much less is known about the roles of PPARs during embryonic and fetal development (Suwik et al., 2020; reviewed in Michalik et al., 2002; Lee et al., 2009; Kadam et al., 2015). Studies have shown poor maternal diet impairs PPARγ activity in utero, and that a maternal food restriction diet results in low birth-weight offspring that later develop obesity, hypertriglycemia and insulin resistance during adulthood (Desai et al., 2015; Belenchia et al., 2018). However, the mechanisms, timing, and causality of these consequences remain unknown.

Small molecule, pharmacological agonists and antagonists have been designed to modulate the three PPAR isoforms with great specificity, and have been explored for therapeutic potential for obesity, diabetes, and high cholesterol. These compounds provide a powerful tool for observing the effects of both upregulation and downregulation of each isoform in a more physiologically relevant manner, when compared to PPAR knockdown or knockout transgenic models. Use of these synthetic compounds has been characterized in vitro as well as in murine models. However, there is little reported use in the zebrafish model—a well-established model in the field of developmental toxicology (Sipes et al., 2011; Volz et al., 2015).

Zebrafish provide many advantages for developmental studies. Many developmental regulatory pathways are highly conserved between mammals and fish species, including humans and zebrafish (Sipes et al., 2011). Furthermore, zebrafish embryos are useful models for observing specific organ development due to their transparency during development and easy transgenic manipulation. These strategies allow for direct, real-time visualization of organ systems and adipocytes in a whole, live organism (Tiso et al., 2009). Our goal is to utilize the zebrafish model to characterize the role of all three PPAR isoforms during embryogenesis. Unlike mammals, zebrafish express two subtypes of PPARα (pparaa & pparab) and PPARβ/δ (pparda & ppardb), while similarly only expressing one PPARγ homolog (Zhao et al., 2015). Despite expression of the orthologs in zebrafish, all three PPAR isoforms are conserved in zebrafish, as are the downstream signaling processes (Den Broeder et al., 2015). We aim to perform pharmacological modulation of each PPAR isoform to elucidate the effects of altered PPAR activity during embryogenesis.

We hypothesized that pharmacological modulations of PPARs will impact zebrafish pancreas development and lipid accumulation, due to the organ’s known sensitivity to lipotoxicity and toxicant-induced perturbations of PPAR signaling. In this study, we present data on the roles of isoform specific PPAR modulation in the developing endocrine and exocrine pancreas using transgenic zebrafish. We further assess the effectiveness of these pharmaceuticals in the zebrafish model through gene expression analysis of isoform specific PPAR target genes in combination with morphological and lipid homeostasis. Our data indicates that altered PPAR activity during embryogenesis disrupt normal development, and we report PPAR isoform-specific effects on pancreas development.

MATERIALS & METHODS

Chemicals & Reagents

Rosiglitazone, T0070907, GW 6471, and GW 590735 were purchased from Cayman Chemical (Ann Arbor, MI; ≥98% purity). GSK3787 was purchased from Millipore Sigma (Temecula, CA; ≥99% purity). GW501516 was purchased from AdipoGen (San Diego, CA; ≥98% purity). Both PPARγ and PPARβ/δ ligands, as well as GW590735 bind the ligand binding domain (LBD) of their respective isoform (Lehmann et al., 1995; Lee et al., 2002; Sierra et al., 2007), whereas GW6471 acts through recruiting corepressors of PPARα (Xu et al., 2002). A list of PPAR isoform specific ligands and their functions is included in Table 1. All compounds were dissolved in dimethyl sulfoxide (DMSO) to prepare 100 mM working stocks (highest concentration), and lower concentration working stocks were prepared through serial dilutions. GSK3787 was stored at 4°C away from the light and all other compounds were stored at −20°C. Nile Red was purchased from ThermoFisher Scientific (Waltham, Massachusetts) and stored at −20°C away from light exposure. All experimental procedures were conducted using appropriate safety precautions.

Table 1.

List of chemicals used to modulate PPAR isoform.

	Chemicals	Mode of action
Vehicle	DMSO
PPARα	GW590735 (↑)	Binds ligand binding domain (LBD) to activate downstream gene expression of PPARα
	GW6471 (↓)	Recruits corepressor to repress downstream gene expression of PPARα
PPARγ	Rosiglitazone (↑)	Binds ligand binding domain (LBD) to activate downstream gene expression of PPARγ
	T0070907 (↓)	Binds ligand binding domain (LBD) to repress downstream gene expression of PPARγ
PPARβ/δ	GW501516 (↑)	Binds ligand binding domain (LBD) to activate downstream gene expression of PPARβ/δ
	GSK3787 (↓)	Binds ligand binding domain (LBD) to repress downstream gene expression of PPARβ/δ

↑ is used for agonists and ↓ is used for antagonists

Animal Husbandry

Adult fish were housed in an automated Aquaneering zebrafish system maintained at 28.5°C in accordance with the recommendations in the Guide for the Care and Use of Laboratory Animals of the National Institutes of Health. Institutional approvals were provided by the San Diego State University and University of Massachusetts Amherst Institutional Animal Care and Use Committees (PHS Assurance Number 16–00430; Animal Welfare Assurance Number A3551–01). In both facilities, all adult fish and embryos were maintained on a 14 h light:10 h dark daily cycle and provided the recommended amount of GEMMA Micro 300 (Skretting; Westbrook, ME) twice daily. Breeding populations were housed in tanks containing 2:1 female to male ratio. All embryos were collected within 1 hour post fertilization.

Zebrafish Strains

To study the effects of altered PPAR activity on gross and pancreatic morphology, transgenic Tg(insulin:GFP) and Tg(ptf1a:GFP) zebrafish were used (diIorio et al., 2002; Lin et al., 2004). They were originally obtained as heterozygous populations on the wildtype AB background strain from Dr Philip diIorio at the University of Massachusetts Medical School (Worcester, Massachusetts) and bred to homozygosity. Tg(insulin:GFP) zebrafish express GFP in β cells, while Tg(ptf1a:GFP) zebrafish express GFP throughout the exocrine pancreas, as well as in the retina and hindbrain (Lin et al., 2004; Godinho et al., 2005). All embryos were collected from homozygous genotyped tanks.

Wildtype AB strain zebrafish were used to observe the effects of altered PPAR on lipid deposition. Larvae at 5 days post fertilization (dpf) larvae were transferred to 250 mL beakers 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 7.2), each with no more than 20 larvae. Larvae were fed the recommended amount of GEMMA Micro 75 (Skretting; Westbrook, ME) and water was refreshed once each day throughout the experiment.

Animal Exposures

A diagram of the utilized exposure paradigms for qPCR, morphology, and adiposity experiments is presented in Figure 1. To assess changes in gene expression within the PPAR signaling pathway, embryos were exposed to 10 μM Rosiglitazone, T0070907, GW6471, GW590735, GSK3787, or GW501516 at 3 dpf for 24 hours prior to RNA isolations. This exposure paradigm was selected because most PPAR isoforms have relatively higher PPAR mRNA expression by 3 dpf (Cheng et al., 2019). Concentrations were selected based on previous studies in zebrafish (Ouaddah-Boussouf, et al. 2016; Tiefenbach, et al 2010), and optimized in our laboratory to ensure that no mortality was observed. These embryos were maintained in individual wells of a 96 well plate containing 200 μL of exposure solution (1 embryo per well). Experimental replicates of 3–5 groups with 15 embryos per treatment were performed for RNA isolation. To further examine effects of PPAR β/δ on pancreatic gene expression, embryos were exposed to 10 μM GW501516 or GSK3787 from 24–96 hpf following manual dechorionation at 24 hpf using watchmakers’ forceps. To capture the entire window of pancreatic organogenesis, embryos were exposed daily in 24-well plates, with each well containing 1 mL of exposure solution. Given that the half-lives of these compounds are <1 day in humans and the recommended use of related pharmacological products is daily administration, we employed daily water exposures.

Figure 1.

Exposure paradigms utilized in this study to assess gene expression, morphology, and adiposity. Embryos were exposed for 24 hours from 3–4 dpf prior to assessment of gene expression in 4 dpf embryos. PPAR gene expression in embryos greatly increases beginning at 3 dpf, and therefore the gene expression paradigm was fitted to this window of exposure. For morphological experiments, embryos were exposed from 1–4 dpf prior to microscopy. To assess adiposity, embryos were exposed from 1–5 dpf, and then transferred to clean water until larval imaging at 8 dpf.

For morphometric analysis, Tg(insulin:GFP) and Tg(ptf1a:GFP) embryos were manually removed from the chorion at 24 hpf and exposed to rosiglitazone, T0070907, GW6471, GW590735, or GSK3787 at concentrations of 0 (0.01% v/v DMSO), 0.1, 1, and 10 μM in 0.3x Danieau’s medium. Embryos were exposed to GW501516 at concentrations of 0, 0.01, 0.1, and 1 μM in 0.3x Danieau’s medium due to high incidence of mortality during prolonged exposures at 10 μM. Embryos were housed in individual wells of a 24-well plate containing 1 mL of exposure solution, replaced entirely daily until 4 dpf, when larvae were placed in fresh Danieau’s medium and imaged. Experiments were replicated 2–3 times on groups of 15–20 embryos.

RNA Isolation & Quantitative PCR

Following exposures, embryos were rinsed in 0.3x Danieau’s medium and collected into RNAlater solution (Fisher Scientific; Waltham, MA), and stored at −80 °C until RNA extraction. RNA was isolated using the GeneJET RNA Purification Kit purchased from Fisher Scientific (Waltham, MA, USA). Following isolation, RNA concentrations and sample purity were determined using a μLITE spectrophotometer purchased from BioDrop (Cambridge, UK). 1 μg of RNA was converted into cDNA using the iScript cDNA Synthesis Kit (Bio-Rad; Hercules, CA). Afterward, samples were diluted to 0.25 ng/μL cDNA in nuclease-free water and stored at −20 °C.

Using a Bio-Rad CFX Connect Real-Time PCR Detection System, Quantitative Real-Time PCR (qRT-PCR) was performed to assess the expression of pparaa, pparg, fabp1a, and fabp1b1. A 20 μL reaction mixture was prepared containing 5 μL nuclease-free water, 0.5 μL of (5 pM) each primer, and 10 μL 2x iQ SYBR Green Supermix (BioRad), and 4 μL (1 ng) cDNA template. Sequences and optimization temperature are described in Table S1. CFX Manager software (BioRad) was used to analyze the data and fold changes were calculated using the ΔΔCT method (Livak and Schmittgen, 2001). These primers have been previously published and validated. Beta-actin (ba) and beta-2-macroglobulin (b2m) were used as housekeeping genes and the arithmetic mean of the Cq values of both genes was used as the housekeeping reference. There was no significant difference in the expression of both housekeeping genes due to exposure.

Microscopy

Tg(insulin:GFP) and Tg(ptf1a:GFP) embryos were imaged at 4 dpf to examine morphogenesis of the primary islet at 100x magnification and posterior extension of the exocrine pancreas at 50x magnification, respectively. All embryos were additionally analyzed for morphometry, including overall fish length at 20x magnification and yolk sac utilization at 50x magnification. Embryos are larvae with yolk sac edema or dark yolk were excluded from these measurements to avoid bias, though exclusion did not impact measurements (Table S2). Fluorescence and brightfield microscopy were conducted using a Zeiss Axiozoom v16 upright dissecting microscopy containing a GFP filter. To conduct imaging, embryos were anesthetized in 2% v/v MS-222 (prepared as 4 mg/ml tricaine powder in water, pH buffered, and stored at −20°C until use) and mounted in a 3% methylcellulose solution. Embryos were positioned on their left lateral side for optimal pancreas imaging. Because of the potential for classification bias for aberrant phenotypes, images were blinded and randomized prior to assessment.

Visualization of lipid droplets

Nile Red staining was performed to quantify neutral lipid deposits in exposed and control larval zebrafish. Embryos were exposed each day until 5 dpf, then transferred to clean water. We have previously shown that these developmental exposures (1–5 dpf) are sufficient to increase adiposity at later timepoints (Sant et al., 2021a; Sant et al., 2021b). Here, we assessed whether developmental modulation of PPARs alone were also capable of these effects. At 8 dpf larvae were transferred to glass petri dishes containing 30 mL of 0.3x Danieau’s medium with 15 μL of 5 mg/mL Nile Red (2.5 μg/mL). Following 45 minute incubation at room temperature in the dark, larvae were immediately washed in 0.3x Danieau’s medium. Stained lipid deposits of 2–3 randomly selected larvae within each dosing group per experimental replicate were visualized through fluorescence microscopy utilizing a Zeiss Axiozoom v16 upright dissecting microscope containing a GFP filter. All larvae not used for microscopy were randomly pooled into groups of 3 larvae from the same exposure concentration, transferred to a 50% acetone and water solution, and sonicated. Nile Red fluorescence was immediately quantified using a Cytation analyzer plate reader with excitation at 488 nm and emission at 565 nm.

Putative PPAR Transcription Factor Binding Sites (TFBS) in Pancreatic Genes

Putative TFBS, namely potential PPAR-responsive elements (PPREs), within the promoter and gene region of target genes were identified in silico using the UCSC Genome Browser track hub JASPAR2020 TFBS danRer11. The track lists predicted binding sites for transcription factor binding profiles in the JASPAR CORE 2020 vertebrates collection database to the zebrafish genome, GRCz11/danRer11 assembly (http://jaspar.genereg.net/). The output of the track for the target gene region and 6000 bp upstream of the transcription start site was filtered for all profiles containing “PPAR” and whose p-values ≤ 10–4 (track score ≥ 400). Exons were identified using the annotations for GRCz11 provided by NCBI RefSeq, available at UCSC Genome Browser (https://genome.ucsc.edu/).

Data analysis and statistics

Data is reported as mean +/− SEM. Tests for normality and equal variances were first performed to determine appropriate statistical models. Log transformation of gcga gene expression data prior to analysis was required to assume a normal distribution. One-way ANOVA with Tukey post hoc tests or Games-Howell post hoc tests were used to assess differences between exposure groups. Fisher’s exact test was performed to assess correlation between exposure concentrations and deformities. All statistics were performed using JMP Pro 14 (SAS Institute Inc., Cary, NC, USA). A confidence level of 95% (α=0.05) was used for all tests.

RESULTS

PPAR Agonists and Antagonists and Impacts on PPAR Signaling

We exposed embryos to 10 μM concentrations of PPARα, PPARβ/δ, and PPARγ ligands at 3 dpf for 24 hours, and assessed gene expression at 4 dpf (Figure S1). Gene expression of pparaa was significantly reduced by exposure to the PPARβ/δ agonist, GW501516, compared to controls (p<0.001) as well as when compared to the PPARβ/δ antagonist GSK3787 (p=0.036). The same relationship was also observed for expression of pparg, with reduced expression for embryos treated with PPARβ/δ agonist GW501516 compared to controls (p=0.050) and to the PPARβ/δ antagonist GSK3787 (p=0.039). Expression of fabp1a was unaffected across treatment groups (p>0.05). Gene expression of fabp1b1 was significant downregulated and upregulated by treatment with PPARβ/δ agonist GW501516 (p=0.009) and PPARβ/δ antagonist GSK3787 (p=0.037), respectively, and these exposures also differed from each other (p<0.001). Overall, the only exposures resulting in statistically significant changes compared to controls were via agonism and antagonism of the PPARβ/δ.

Embryonic Morphometry

Morphology associated with modulation of PPAR activity was characterized at 4 dpf, including fish length, yolk sac utilization, and incidence of phenotypic deformities (Figure 2). Zebrafish exposed to 10 μM PPARγ antagonist T0070907 and agonist Rosiglitazone both displayed significantly decreased fish length compared to controls (p=0.002 and p<0.001, respectively). Likewise, both agonism and antagonism of PPARγ impaired yolk sac utilization (p<0.001) compared to controls. Embryos exposed to 10 μM PPARα antagonist GW6471 displayed a significant increase in body length (p<0.001) and yolk sac area (p=0.002). Conversely, exposures to PPARα agonist GW590735 did not significantly alter fish length or yolk sac area at 96 hpf (p>0.05). Similar to PPARα antagonism, treatment with the PPARβ/δ antagonist GSK3787 increased fish length (p=0.020) and increased yolk sac area (p<0.001). PPARβ/δ agonism with GW501516 decreased both fish length (p<0.001) and yolk sac utilization (p<0.001), which was the opposite effect of PPARβ/δ antagonism (expected) and also similar to PPARγ modulation.

Figure 2.

PPAR isoform modulation impacts fish growth and yolk utilization. Fish length and yolk sac area were measured at 4 dpf, following developmental (3 day) exposures to PPAR agonists and antagonist across a range of concentrations. PPARγ agonist and antagonist exposures decreased fish length and increased yolk sac area at 10 μM. PPARα antagonist, but not agonist, exposures increased fish length as well as yolk sac area. Additionally, PPARβ/δ antagonist exposure increased fish length as well as yolk sac area, while agonist exposure decreased fish length and increased yolk sac area. Asterisks (*) indicate a difference between designated treatment groups and DMSO control (p<0.05); n=75–90 eleutheroembryos per group

Structural Deformities & Aberrant Phenotypes

Embryos were examined for the occurrence of morphological deformities and aberrant phenotypes at 4 dpf. Aberrant phenotypes were categorized by the presence of craniofacial malformation, pericardial edema, darkened yolk sac, or “multiple deformities” indicating the presence of two or more of these abnormal phenotypes in combination (Figure 3A). All embryos deemed to have multiple deformities were excluded from the analysis of individual deformities (craniofacial malformation, pericardial edema, or darkened yolk sac) to prevent redundance. No other obvious morphologies or malformations were observed. Exposure to 10 μM PPARγ antagonist increased total deformity occurrence by 30% (p=0.007) (Figure 3B). This increase was mostly accounted for by a 10-fold increase in the recording of multiple deformities (p<0.001), specifically the co-occurrences of pericardial edema and dark yolk (19.5%) as well as craniofacial malformation and dark yolk (10%). PPARγ agonist exposure additionally increased the incidence of total deformity in 0.1 μM (p=0.035) and 10 μM (p=0.003) exposure groups. Notably, craniofacial malformation alone was significantly increased with 0.1 μM (p=0.007) and 1 μM (p=0.026) concentrations.

Figure 3.

Modulation of all PPAR isoforms increases the incidence of morphological deformities. A.) Embryos were examined microscopically for craniofacial malformations, pericardial edema, and darkened yolk sacs following exposure to PPAR agonists and antagonists. B.) PPARγ antagonist exposure displayed increased occurrence of comorbidities presenting as darkened yolk sacs accompanied by either pericardial edema or craniofacial malformation. PPARγ agonist exposure displayed an increased occurrence of craniofacial malformations alone and total deformity as well. Treatment with 10 μM PPARα antagonist resulted in a significant increase in total deformity, with a notable increase in multiple deformities that include pericardial edema. PPARβ/δ antagonist exposure increased pericardial edema and total deformities at 1 μM. Asterisks (*) indicate a difference between designated treatment group and the controls (p<0.05); n= 75–90 eleutheroembryos.

PPARα antagonist exposure increased total morphological deformities at 10 μM by 38% (p=0.005). This is represented by a significant increase in multiple deformities (p=0.018), accounted for by the co-occurrence of pericardial edema and craniofacial malformation (25%), and the co-occurrence of all three morphologies (13%). PPARα agonist exposure did not induce any morphological deformities (p>0.05). PPARβ/δ antagonist exposure additionally did not increase the incidence of any deformity (p>0.05). However, exposure to 10 μM agonist did increase the total deformity incidence (p<0.001), with a significant increase in pericardial edema alone (p=0.048). Finally, both PPARβ/δ compounds perturbed swim bladder inflation at all concentrations and in a concentration-dependent manner (p<0.01 for all agonist and antagonist exposures; Table S3). No significant changes in swim bladder inflation were observed for the PPARα or PPARγ agonists nor antagonists (p>0.05).

Endocrine Pancreas Development

Primary β-cell islet area was quantified in 4 dpf zebrafish in order to observe pancreas-specific effects of PPAR modulation during embryogenesis (Figure 4). Both PPARγ antagonism (10 μM exposure) and agonism (10 μM exposure) significantly decreased islet area when compared to controls (p=0.047 and p=0.042, respectively). Agonism and antagonism of both PPARα and PPARβ/δ did not significantly impact islet area (p>0.05; Figure S2).

Figure 4.

PPARγ modulation decreases islet area. Islet areas were approximated by quantifying the β cell area in 4 dpf Tg(insulin-GFP) transgenic zebrafish. At the highest exposure concentration (10 μM), both antagonist and agonist exposure significantly decreased islet area. Boxplots are shown to indicate data quartiles, and p-values are shown comparing the control and 10 μM groups above the plots. n= 36–42 antagonist; n=30–40 agonist eleutheroembryos.

Islet morphology was also assessed in response to PPAR modulation. We have previously characterized several aberrant islet morphologies resulting from chemical exposures, which were also observed in this study. Fragmentation of islets was characterized by sections of β-cells remaining intact to the primary structure but deviating from the main cluster, and ectopic islets were identified as having a singular cell or small cluster of cells entirely isolated from the primary β-cell cluster (Figure 5A). Hypomorphic islets were characterized as being below the 10th percentile of controls for islet area. Antagonism of PPAR isoforms did not increase the occurrence of these variant islet morphologies (p>0.05) while agonism of all isoforms did display an increase in the incidence of these variant morphologies (Figure 5B). For PPARγ, 1 μM and 10 μM agonist exposure increased the overall occurrence of aberrant islet morphologies (p=0.001 and p<0.001, respectively). Fragmented islet structure alone significantly increased with 10 μM exposure, when compared to controls (p=0.040), demonstrating a 22% increase. Hypomorphic islets were increased by 9% in all exposure groups but was not statistically significant. For PPARα, agonism using exposures of 0.1, 1, and 10 μM increased the observed incidence of altered islet architecture through promotion of all three previously described aberrant islet morphologies (p=0.009, 0.003, and 0.005 respectively), and all concentrations increased total incidence of islet variants by 30% compared to controls. Fragmented islets were 23% of all deformities seen in 0.1 μM, demonstrating a significant increase over controls (p=0.046). Lastly, hypomorphic islets doubled to 22% and 24% in 1 μM and 10 μM exposure, respectively, however this morphology alone was not considered to be statistically significant (p>0.05). PPARβ/δ agonist (10 μM) exposure increased incidence of ectopic β-cell formation by nearly 16% (p=0.015), though no other significant effects were observed.

Figure 5.

PPAR modulation increases the incidence of aberrant islet morphologies at 4 dpf. Images were blinded prior to classification, and the incidence of ectopic β cells, fragmented islets, and hypomorphic islets (falling below the 10^th percentile of controls) was recorded. For all PPAR isoforms, antagonism did not increase the incidence of aberrant islet morphologies (p>0.05), while agonism increased incidence of aberrant islet morphologies (p<0.05). Asterisks (*) above bars indicate a difference in total aberrant islet morphology between designated treatment group and the controls while asterisks within bars indicate a difference only in the associated morphology between designated treatment group and the controls (p<0.05); n= 30–42 for each exposure group.

Exocrine Pancreas Development

Pancreas length was quantified at 4 dpf following agonist and antagonist exposure. Briefly, exocrine pancreas length was measured from the islet center to the proximal (caudal) tip. Modulation of both PPARα and PPARγ (antagonist nor agonist) did not significant impact pancreas length (Figure S3). PPARβ/δ agonist exposure displayed significance at all 0.01 (p=0.01), 0.1 (p<0.001), and 1 μM (p<0.001), with a concentration dependent decrease in pancreas length (Figure 6A). PPARβ/δ antagonist exposure produced a U-shaped concentration-response curve, demonstrating a significant decrease at 0.1 μM (p<0.001).

Figure 6.

PPAR modulation impairs exocrine pancreas length and morphology. A) Exocrine pancreas length was measured in Tg(ptf1a-GFP) transgenic along the nasal-caudal axis. PPARβ/δ antagonist exposure significantly decreased pancreas length, displaying an inverted U-shaped curve, and agonist exposure significantly decreased pancreas length in a concentration dependent manner. Data is presented as boxplots and represents the quantiles for each set of data, and significant p-values are indicated above each plot (p<0.05). n=40–45 antagonist, n=38–47 agonist eleutheroembryos. B) Aberrant pancreatic morphologies observed due to PPAR modulation included curved, wispy, and ectopic exocrine variants. Incidence of aberrant morphologies is presented in Table 2.

While modulation of PPARβ/δ was found to solely impair exocrine pancreas length, deviant exocrine pancreas morphologies were observed as a result of both PPARβ/δ and PPARγ modulation (Figure 6B; Table 2). Curved pancreata were defined as those with a pancreatic head located more ventrally than expected, often colocalizing with the liver. “Wispy” pancreata were a novel phenotype observed in this study, characterized by a thin pancreatic body and small protrusions or nodule-like structures. Antagonism of all PPAR isoforms did not significantly impact pancreatic morphology (p>0.05). PPARγ agonist 10 μM exposure increased the occurrence of curved and “wispy” pancreas morphologies by 18% (p=0.008) and 21% (p=0.004), respectively. Furthermore, PPARβ/δ agonist exposure revealed the presence of ectopic exocrine tissue at 0.01 and 1 μM, which was a phenotype that has not been previously reported. This was characterized as ptf1a-positive pancreatic tissue developing outside of the pancreas. While the incidence of this ectopic tissue was not statistically significant, this novel phenotype may be directly related to PPARβ/δ agonism.

Table 2.

Incidence of observed aberrant exocrine pancreas morphology due to PPAR agonist exposures^†.

Concentration (μM)	Normal	Total variants	Curved	Wispy	Ectopic
PPARα agonism (GW590735)
0	81% (25/31)	19% (6/31)	3% (1/31)	3% (1/31)	0
0.1	89% (31/35)	11% (4/35)	3% (1/35)	0	0
1	75% (24/32)	25% (8/32)	6% (2/32)	6% (2/32)	0
10	69% (22/32)	31% (10/32)	6% (2/32)	0	0
PPARβ/δ agonism (GW501516)
0	100% (47/47)	0	0	0	0% (0/47)
0.01	92% (35/38)	8% (3/38)	0	0	8% (3/38)
0.1	100% (40/40)	0	0	0	0
1	97% (36/39)	3% (3/39)	0	0	3% (1/39)
PPARγ agonism (Rosiglitazone)
0	95% (40/42)	5% (2/42)	5% (2/42)	0	0
0.1	92% (44/48)	8% (4/48)	4% (2/48)	0	0
1	90% (43/48)	10% (5/48)	2% (1/48)	2% (1/48)	0
10	56% (27/48)	44% (21/48)*	23% (11/48)*	21% (10/48)*	0

^†Images of aberrant pancreas morphologies are shown in Figure 6.

^*indicates a significant increase in aberrant morphologies compared to controls (p<0.05)

Pancreatic gene expression & Putative TFBS

Because of the efficacy of PPARβ/δ modulation on PPAR signaling and the impact of treatment on endocrine and exocrine pancreatic morphologies, gene expression of sensitive pancreas target genes was examined in zebrafish exposed daily to PPARβ/δ agonist or antagonist (Figure 7A). The most robust effect was observed for glucagon a (gcga) expression, which increased by more than 800% due to PPARβ/δ agonism compared to controls (p<0.001) and PPARβ/δ agonism (p<0.001). Expression of trypsin (try) was decreased by 80% and 90% in agonists compared to controls (p=0.004) and antagonist (p=0.010), respectively. Expression of pancreatic and duodenal homeobox 1 (pdx1) was significantly reduced by approximately 60% due to PPARβ/δ agonism compared to antagonism (p=0.027). No significant changes of gene expression were observed for amylase 2A (amy2a) or insulin (insa) due to PPARβ/δ modulation (p>0.05). For these reasons, locations of putative PPRE transcription factor binding sites (TFBS) were identified within these genes or proximally upstream (−6,000bp) (Figure 7B). All of these gene targets had several putative PPREs, and most had at least one located near the transcription start site (TSS). Specifically, the 3 genes with significantly modified expression all have putative PPARγ::RXRa TFBS near the gene TSS. Detailed gene locations and TFBS locations are listed in Table S4 & Table S5.

Figure 7.

Modulation of PPAR signaling impacts pancreatic gene expression. A) Gene expression of endocrine (gcga, insa, pdx1) and exocrine (try, amy2a) pancreas transcripts was assessed following PPARβ/δ modulation using qPCR. Agonism of PPARβ/δ increases gcga expression by almost 10-fold, and decreases expression of pdx1 and try, indicative of potential endocrine and exocrine effects. B) Putative PPAR transcription factor binding sites (TFBS) were examined in the promoters of pancreatic gene targets. Each gene had several isoform-specific or multiple isoform binding sites in the promoter or gene body, including several directly adjacent to the transcription start site. Asterisks (*) indicate a significant difference between controls and a PPAR modulator, while octothorpes (#) indicate a significant difference between the agonist and antagonist. α=0.05, n=6–10 samples per group each containing 8–10 pooled embryos

Lipid Accumulation

Nile Red staining was performed at 8 dpf to quantify lipid accumulation in zebrafish larvae following developmental PPAR modulation. Larvae developmentally exposed to 10 μM PPARα antagonist were excluded from this assay due to increased mortality by 7 dpf (Figure S4). As expected, PPARγ antagonism significantly decreased total body lipid accumulation (p<0.001), while PPARγ activation significantly increased total body lipid accumulation (p<0.001)—both in a concentration dependent manner (Figure 8A,B). PPARα agonism decreased lipid accumulation (p<0.001), and no significant change in lipid staining was observed due to PPARα antagonism (Figure 8C). Interestingly, both PPARβ/δ activation and inhibition resulted in a concentration dependent decrease in lipid accumulation (p<0.001) (Figure 8D).

Figure 8.

PPAR modulation alters lipid accumulation at 8 dpf. Nile Red plate reader assay was performed to quantify larval lipid accumulation 3 days after discontinuing exposure. All values are normalized to control. (A,B) Larvae exposed to the PPARγ antagonist had decreased neutral lipid accumulation, while those exposed to the agonist were increased. (C) PPARα antagonist exposure displayed an increasing trend in lipid accumulation, however not significant. Conversely, agonist exposure decreased lipid accumulation. (D) Both PPARβ/δ antagonist and agonist exposure significantly decreased lipid accumulation in a concentration dependent manner. Asterisks (*) indicate a difference between designated treatment group and the control (p<0.05). n=13–16 biological replicates for PPARα; n= 7–10 PPARβ/δ; n=10–13 for PPARγ

DISCUSSION

PPARs are essential regulators of nutrient homeostasis. The spatiotemporal expression patterns of PPARs are well-documented across vertebrates, including the zebrafish (Ibabe et al., 2002; Maradonna et al., 2013). Collectively, studies demonstrate the importance of properly regulated PPAR activity during fetal metabolic development. While few studies have observed the effects of PPAR modulation during development (Cheng et al., 2019; Sant et al., 2018; Hsieh et al., 2018), there is very little known about these consequences in the developing pancreas. The pancreas performs essential metabolic functions in lipid and glucose homeostasis and is known to be susceptible to lipotoxicity and oxidative stress (Yang and Li, 2012; Sant et al., 2016; Martinez-Useros et al., 2017; Wang and Wang, 2017). Additionally, toxicants with reported affinity for one or more PPAR isoforms have elicited adverse effects on pancreatic development following embryonic exposure in zebrafish (Sant et al., 2016; Brown et al., 2018; Jacobs et al., 2018), making it an important endpoint to observe in predisposition to metabolic disease. In this study, the use of pharmacological agonists and antagonists for PPAR allow for the modulation of specific isoforms throughout embryonic development. Our goal was to characterize the role of specific PPAR isoforms during embryonic development, namely during pancreatic organogenesis.

Zebrafish are a unique tool to study PPAR function and organogenesis. First, we utilized Tg(insulin-GFP) and Tg(ptf1a-GFP) zebrafish strains to directly visualize the developing endocrine and exocrine pancreas in live animals, respectively, and lipid stains to visualize adiposity. Because they develop externally, pharmacological treatments were also easy to administer and target specific developmental processes independent of any potential maternal metabolism. One challenge with the zebrafish model is the existence of multiple orthologs for some genes due to ancestral teleost whole-genome duplication events (Poslethwait et al., 2004). As a result of this, multiple orthologs of PPARs exist within the zebrafish genome—two PPARα orthologs (pparaa, pparab), two PPARβ/δ orthologs (pparda, ppardb), and one PPARγ ortholog (pparg). These duplications can be used as a tool to better understand the functionality of signaling processes, and can be explored through gene relationships such as complementation, subfunctionalization, and neofunctionalization. Works by Laprairie et al have utilized the zebrafish as a tool to better understand gene subfunctionalization within the PPAR family and its targets and have identified isoform-specific signaling processes related to fatty acid binding proteins (Laprairie et al., 2016a; 2016b; 2018).

We examined PPAR-related gene expression in response to selective PPAR ligands to gain isoform-specific insight. Exposure to selective PPARα and PPARγ agonists and antagonists for 24 hours failed to significantly impact regulation of PPAR signaling at 4 dpf. This was unexpected, since a prior study had demonstrated that genes fabp1a and fabp1b1 are specific targets of PPARα and PPARγ signaling, respectively (Laprairie et al., 2016a). This observation is consistent with a recent studying showing that exposures to ciglitazone (a PPARγ agonist) fail to activate the zebrafish PPARγ ligand-binding domain, despite over 80% sequence similarity between zebrafish and humans (Cheng et al., 2019). It is likely that, due to half-lives of <1 day in other vertebrate species, complete agonism and antagonism throughout the entire 24-h period was not maintained in this study. Likewise, many of the organs with the highest PPAR expression (such as the liver) were not fully functional at our 4 dpf timepoint, so it is also possible that modulation of PPARα and PPARγ may produce effects of greater magnitude once these metabolic processes and organs are fully developed. However, our data found a strong effect from treatment with PPARβ/δ agonist and antagonist, which was novel because little is known about the autoregulatory function of PPARβ/δ (Figure S1). Interestingly, treatment with PPARβ/δ agonist GW501516 decreased expression of pparaa, pparg, and PPARγ target fabp1b1. Inversely, treatment with PPARβ/δ antagonist GSK3787 increased expression of these genes. This suggests that PPARβ/δ may play a role in autoregulation of other PPAR isoforms or that compensatory function or shared functionalization may occur across isoforms, and may play a bigger role during development.

Exposure to PPAR modulators significantly impacted zebrafish growth (Figure 2). Fish length was decreased and yolk sac area was increased due to all PPARγ modulators, suggesting decreased nutrient utilization and growth by PPARγ agonism and antagonism. This was also observed for PPARβ/δ agonist treatment. Yolk sac area is representative of nutrient uptake by the embryos, as this is the sole nutrient source until onset of exogenous feeding around 5 dpf. As yolk sac area decreases, fish length is expected to increase due to increased nutrient uptake fueling growth (Sant and Timme-Laragy, 2018; Schwartz et al., 2021). Therefore, modulation of PPARγ and antagonism of PPARβ/δ both show canonical patterns of decreased embryonic nutrition leading to reduced growth. Deviations from this relationship could suggest increased fat accumulation or deposition, or enhanced metabolic activity without increased growth. Thus, when treated with PPARα antagonist, the increased fish growth coupled with increased remaining yolk area was surprising. However, the zebrafish yolk is largely bulk lipids as a nutrient source, and it is possible that antagonism of PPARα and the resulting impairment of lipid catabolism and fatty acid oxidation may introduce adaptive or even stochastic preference towards other energy-generating processes. Decreased survival to 7 dpf (Figure S4) suggests that these fish may be at a disadvantage once exogenous feeding begins at the onset of complete yolk depletion. Additional biochemical experiments are required in order to better understand this complex relationship.

Morphological deformities in the zebrafish larvae were detectable with overall PPARγ modulation and PPARα inhibition but did not correlate with alterations to PPARβ/δ activity (Figure 3). Darkened yolk sacs of embryos exposed to PPARγ regulators is a relatively novel observation. This phenomenon may be due to an inability of the embryo to properly break down and absorb the yolk, leaving these fats to become necrotic and dark in color (Fraher et al., 2016). Furthermore, pericardial edemas observed in isolation or with other deformities observed in this study are consistent with other reports of cardiotoxicity and hypertrophy, namely due to lipotoxicity in cardiomyocytes, following modulation of PPARα and PPARγ (Bonda et al., 2016; Czarnowska et al., 2016; Duan et al., 2005; Kar et al., 2018). Because of the elevated expression of PPARβ/δ in diverse tissues throughout development, the lack of malformations associated with agonism or antagonism was surprising. However, reduced swim bladder inflation was observed due to both agonism and antagonism of PPARβ/δ (Table S3), and our PPARβ/δ agonist exposures for morphological experiments were 10-fold lower than the other modulators due to increased mortality throughout the 4-day exposure period. Therefore, we were unable to observe the incidence of malformations due to the 10 μM GW501516 exposure. All three PPAR isoforms have known functions in β cell mass, insulin secretion and hormone regulation (Kim and Ahn, 2004; Sugden and Holness, 2004; Iglesias et al., 2012). Thus, we assessed the consequences of PPAR modulation on developmental of the endocrine pancreas (Figure 4). Islet β cells appeared to be most susceptible to modulation of PPARγ, as both agonism and antagonism decreased average islet area. One of the best characterized mechanisms of islet toxicity is due to lipotoxicity, namely due to saturated fatty acids and bulk lipid accumulation within the pancreas (Matsuda et al., 2014; Estadella et al., 2013). It is probable that PPARγ demonstrated the most significant effect on β-cell islet area due to its greater lipid accumulation, as seen through Nile Red staining (Figure 8). However, islet areas were not impacted by PPARα or PPAR β/δ modulation. Because PPARα is the most highly expressed isoform in pancreatic beta cells, it was expected to have a greater and more specific impact on the islet itself (Segerstolpe et al., 2016). Despite no effect on islet area, dysmorphogenesis of the islets following PPARα and PPARβ/δ activation suggests potential pancreatic toxicity beyond proliferation and growth. Likewise, all islet phenotypes were observed prior to exogenous feeding to capture the window of organogenesis. Therefore, it is possible that a dietary challenge (such as the addition of carbohydrates as a major nutrient source) is necessary to observe the effects of PPAR modulation on islet development.

We have previously shown that growth of the exocrine pancreatic tail is a highly sensitive process and susceptible to toxicological perturbation (Jacobs et al, 2018; Sant et al., 2017; 2018). Here, PPAR modulation was performed from 1–4 dpf throughout the dynamic windows of organogenesis. The exocrine pancreas originally forms as a tissue budding from the GI tract, and the period of elongation mostly occurs between 2–3 dpf in the zebrafish (Tiso and Argenton, 2009). Elongation of exocrine pancreatic tissue appears to be most sensitive to PPARβ/δ ligand exposure, as both PPARβ/δ agonist and antagonist significantly decrease pancreas length (Figure 6). We also show that PPARα and PPARγ modulation appeared to have no significant impact on pancreatic length (Figure S3). Other works have shown that PPARβ/δ is the most highly expressed isoform in ductal and acinar cells of the pancreas, providing potential insight into the sensitivity of the exocrine pancreas to exposure of these isoform ligands (Segerstolpe et al., 2016).

Perhaps the most novel finding of this work was the ability of PPAR modulation to impact the morphology of the pancreas (Figure 6; Table 2). The occurrence of “wispy” exocrine tissue has also not been reported with previous toxicant exposure experiments, and was observed due primarily to PPARγ agonism. Low concentration PPARβ/δ agonist (GW501516) treatment revealed the presence of ectopic exocrine pancreas cells, an occurrence that has not been previously observed. PPARβ/δ signaling is suggested to play an essential role in organogenesis, primarily reported in embryonic stem cells, the epidermal layer of the skin, and the colon (Higa et al., 2007; Rees et al., 2008). Because modulation of PPARβ/δ also significantly impacted pancreatic length, these data suggest that PPARβ/δ may be fundamentally important for the organogenesis and exocrine programming of the pancreas.

Because of the novel morphologies and decreased pancreatic length due to PPARβ/δ treatment, we assessed the expression and putative PPRE localization of genes related to pancreas development and function (Figure 7). Putative PPREs were located within each gene, with both PPARβ/δ and PPARγ having putative PPREs within Exon 1 of the genes gcga and pdx1. Agonism of PPARβ/δ resulted in decreased gene expression of pdx1, a transcription factor largely involved in insulin production and β cell function. Conversely, agonism of PPARβ/δ increased the expression of gcga by more than 800%. PDX1 expression is not commonly found in α cells (glucagon-positive), except when undergoing α to β cell transdifferentiation in islets—a process being widely explored as a therapy for diabetes and reduced β cell mass (Piran et al., 2014). For this reason, PDX1 expression is typically high in β cells and has an inverse relationship with glucagon expression (Wang et al. 2001). Here, we confirm this inverse relationship, with robustly decreased pdx1 expression and increased gcga expression. Though decreased islet areas, approximated by β cell fluorescence were not observed, it is possible that islet α cell mass is expanded due to PPARβ/δ agonism. Similarly to studies showing that a gain of PDX1 expression leads to α to β cell transdifferentiation, another study has shown that suppressed PDX1 expression may conversely favor α cell transition over β cell fates (Wang et al., 2001). Additional time-lapse imaging with transgenic expressions of both α and β cells could help to identify whether PPARβ/δ may play a larger role in islet transdiffentiation.

We also evaluated lipid accumulation in 8 dpf larvae following developmental PPAR ligand exposure (Figure 8). It is worth noting that these pharmaceutical regulators all have a half-life on the scale of hours, and lipid analysis was performed three days post-exposure (at 8 dpf). Nile Red staining results fit the known PPAR paradigms for PPARα and PPARγ, showing decreased and increased staining, respectively. Additionally, we found that both PPARβ/δ agonist and antagonist exposure significantly decreased lipid accumulation. Studies in mice and primates have revealed targeted activation of this isoform to produce a model “resistant to obesity” (Wagner and Wagner, 2010), supporting the decrease in lipid accumulation that we see in agonist exposure. However, decreased lipid accumulation was produced with antagonist exposure, which was unexpected. We observed that gene expression of fabp1b was increased by antagonism of PPARβ/δ (Figure S1). Animal studies using zebrafish and mice, as well as epidemiological studies, have shown positive associations between Fabp1 (FABP1 or fabp1b) and obesity, adiposity, or hepatic steatosis, which is contrary to our findings (Lu et al., 2020; Mukai et al., 2017; Newberry et al., 2008; Shi et al., 2012; Shimada et al., 2015; You et al., 2020). Though much remains unclear about the role of PPARβ/δ in adiposity, it is also possible that crosstalk between PPAR isoforms may occur. We showed that agonism of PPARβ/δ resulted in decreased expression of pparaa and pparg (Figure S1). There are reports of overlapping functions of this isoform for both PPARγ, in terms of proliferation and differentiation, as well as PPARα, in fatty acid transport. More spatiotemporal analysis is necessary to understand whether these effects are direct or indirect.

For all PPAR agonists and antagonists, indicators of potential systemic toxicity such as malformations, decreased fish growth, and modified yolk sac area were present at the highest assessed concentrations. For this reason, we used a concentration-response strategy throughout these morphology experiments (Figures 2 & 3). While the lower concentrations utilized appear to fall below the no-observed-adverse-effect levels for most of the endpoints assessed, it is important to note that systemic toxicity from the pharmacological agents may be possible at the higher concentrations. Therefore, we cannot conclude whether the morphological effects observed at the highest concentrations are strictly due to PPAR modulation or whether they are the result of systemic embryotoxicity. For this reason, we also employed a concentration-response study design throughout pancreatic experiments (Figures 4–8). Though many of these pancreatic consequences show a progressive concentration-dependent response, additional experiments are needed to confirm that these effects are due solely to PPAR modulation rather than toxicity. Future work using targeted PPAR knockout using PPAR mutant zebrafish strains will be able to illuminate whether this is a true effect of PPAR modulation or an artifact of pharmacological toxicity.

The implications of these findings go beyond pharmacological modulation of PPARs, since exposures to these agents during pregnancy are unlikely. However, it is important to note that ubiquitous environmental toxicants such as perfluorinated compounds and phthalates have reported affinity for one or more PPAR isoform (Hurst et al., 2003; Takacs and Abbott, 2007; Lau et al., 2010; Zhang et al., 2014; Behr et al., 2020; Khazee et al., 2021). We have previously demonstrated the ability of these toxicants to perturb pancreas development and adiposity in zebrafish (Sant et al., 2017; 2018; 2020; 2021a; 2021b; Jacobs et al., 2018). Because these chemicals may contaminate matrices such as drinking water, consumer products, and foods, exposures to these toxicants during organogenesis are far more likely. Therefore, elucidating the roles of PPAR signaling during embryogenesis and organogenesis, in an isoform-specific manner, may help to provide a stronger framework for an adverse outcome pathway or mechanism for adverse developmental toxicant outcomes.

In conclusion, we present data demonstrating that modulation of PPAR signaling during development can impair pancreatic organogenesis and increase adiposity. Specifically, we have found that perturbation of PPARβ/δ signaling results in reduced pancreas length, increased aberrant pancreas morphologies, and disrupted endocrine and exocrine gene expression in the zebrafish model. Although we do see morphological and expression indications of PPAR-specific modulation following agonist and antagonist exposures in zebrafish, the potential for crosstalk between isoforms and downstream signaling processes requires future exploration and causal analysis. Further research is needed to expand our understanding of how developmental PPAR function may physiologically impact early onset of metabolic diseases.