Ciglitazone—a human PPARγ agonist—disrupts dorsoventral patterning in zebrafish

Animals

Adult wildtype (strain 5D) zebrafish were maintained and bred on a recirculating system using previously described procedures (Mitchell et al., 2018). All adult breeders were handled and treated in accordance with Institutional Animal Care and Use Committee-approved animal use protocols (#20150035 and #20180063) at the University of California, Riverside.

Chemicals

Ciglitazone (>99.4% purity) was purchased from Tocris Bioscience (Bristol, UK), and dorsomorphin (DMP) (99.7% purity) was purchased from Millipore Sigma (St. Louis, MO, USA). For both chemicals, stock solutions were prepared in high performance liquid chromatography-grade dimethyl sulfoxide (DMSO) and stored in two mL amber glass vials with polytetrafluoroethylene-lined caps. Working solutions were prepared by spiking stock solutions into particulate-free water from our recirculating system (pH and conductivity of ~7.2 and ~950 μS, respectively) immediately prior to each experiment, resulting in 0.2% DMSO within all vehicle control and treatment groups. Propylene glycol (>99.5% purity) and Oil Red O (ORO) (>75% dye content) were purchased from Fisher Scientific (Hampton, NH, USA) and Sigma–Aldrich (St. Louis, MO, USA), respectively.

Bioinformatics

Zebrafish-specific pparγ, pparαa, pparαb, pparδa, and pparδb transcript abundance (transcripts per million, or TPM) across developmental stages were obtained from White et al. (2017) (provided within White et al., 2017 as Supplementary File 3, RNA-seq TPM; .tsv file), and stages were converted into hpf per Kimmel et al. (1995). Five replicate TPM values per developmental stage were used to calculate the mean TPM ± standard deviation at each developmental stage. PPARα, PPARδ, and PPARγ amino acid sequences for Homo sapiens (NP_005027.2; NP_006229.1; NP_056953.2), Mus musculus (NP_035274.2; NP_035275.1; NP_035276.2), Rattus norvegicus (NP_037328.1; NP_037273.2; NP_001138838.1), and Danio rerio (NP_001154805.1 (αa); NP_001096037.1 (αb); XP_699900.6 (δa); NP_571543.1 (δb); NP_571542.1) were obtained from the National Center for Biotechnology Information (www.ncbi.nlm.nih.gov). Sequences were aligned using the Multiple Sequence Alignment Tool within Clustal Omega (https://www.ebi.ac.uk/Tools/msa/clustalo/), and the aligned file was used to generate a cladogram within Clustal Omega. Pairwise sequence alignments were also performed to obtain percent amino acid similarity using EMBOSS Matcher (https://www.ebi.ac.uk/Tools/psa/emboss_matcher/). The following default options were used for all pairwise alignments: Matrix = BLOSUM62; Gap Open = 1; Gap Extend = 4; and Alternatives = 1.

Embryo exposures and phenotyping

Embryos were sorted and exposed to either vehicle (0.2% DMSO) or ciglitazone (9.375, 12.5, 15, or 20 μM) from 4 to 24 hpf in glass petri dishes (20 embryos per replicate; three replicates per treatment). Ciglitazone concentrations were selected based on the maximum tolerated concentration (based on survival as an endpoint) in zebrafish embryos following a 4–24 hpf exposure. At 24 hpf, embryos were imaged under transmitted light at 2× magnification using a Leica MZ10 F stereomicroscope equipped with a DMC2900 camera and assessed for survival and dorsoventral patterning abnormalities (ventralization, dorsalization, or delayed development). Following previously described protocols (Dasgupta et al., 2017), ventralized embryos were defined as embryos with a swollen yolk sac extension; dorsalized embryos were defined as embryos with a tail deformity; and delayed embryos were defined as embryos that phenocopied embryos at a developmental stage prior to 24 hpf.

Morpholino injections

Morpholino antisense oligos were synthesized and obtained from Gene Tools, Inc. (Philomath, OR, USA). A fluorescein-tagged splice-blocking MO was designed to target the first exon-intron boundary (E1I1) of zebrafish pparγ-specific pre-mRNA (NCBI Gene ID: 557037), leading to insertion of intron 1 within pparγ mRNA (pparγ-MO sequence: 5′-TCAGCTCCTCTCTGACACTTACCAG-3′). We did not rely on a pparγ-specific translational MO due to the lack of a commercially available PPARγ-specific antibody that cross reacts with zebrafish PPARγ and, as such, inability to confirm knockdown of PPARγ protein. Gene Tools’ standard fluorescein-tagged negative control MO (nc-MO)—a MO that targets a human β-globin intron mutation—was used in order to account for potential non-target MO toxicity, and a zebrafish-specific, fluorescein-tagged chordin MO (chd-MO sequence: 5′-ATCCACAGCAGCCCCTCCATCATCC-3′) was used as a positive control for disruption of dorsoventral patterning (ventralization) at 24 hpf. Water injections were performed in order to account for potential toxicity associated with injection-related stress. MO stock solutions (1 mM) were prepared by resuspending lyophilized MOs in molecular biology-grade (MBG) water, and stocks were stored at room temperature in the dark.

Working solutions of nc-MOs and pparγ-MOs were diluted to 0.5 mM in MBG water and working solutions of chd-MOs were diluted to 0.125 mM in MBG water. Newly fertilized (1- to 8-cell stage, or before 1.25 hpf) zebrafish embryos were microinjected with MOs (~three nL per embryo) using a motorized Eppendorf Injectman NI2 and FemtoJet 4x similar to previously described protocols (McGee et al., 2013; Dasgupta et al., 2017). At 3 hpf, MO delivery in embryos was confirmed using a Leica MZ10 F stereomicroscope equipped with a DMC2900 camera and a GFP filter cube; non-fluorescent and/or coagulated embryos were discarded. Fluorescent embryos were then exposed to either vehicle (0.2% DMSO) or 12.5 μM ciglitazone from 4 to 24 hpf and assessed for dorsoventral patterning abnormalities as described above.

To confirm pparγ knockdown, injected embryos (20 per pool; three replicate pools per group) were snap-frozen in liquid nitrogen at 24 hpf and stored at −80 °C. Total RNA was extracted using an SV Total RNA Isolation System (Promega, Madison, WI, USA) and eluted in 30 µL of nuclease-free water. RNA quality and quantity were confirmed using an Agilent 2100 Bioanalyzer system and Qubit 4.0 Fluorometer (Thermo Fisher Scientific, Waltman, MA, USA), respectively. A total of ~140 ng RNA per replicate sample was reverse-transcribed into cDNA using a GoScript Reverse Transcription System (Promega, Madison, WI, USA). An E1E2 pparγ fragment (~228 bp) was then amplified (forward primer: 5′-CACATCTACAGTAGTGCAGTCAT-3′; reverse primer: 5′-TGTTGGGTTGTTCTCGTAGTC-3′) using approximately 50 ng of cDNA per sample, ZymoTaq PreMix (Zymo Research, Irvine, CA, USA), and an Eppendorf Mastercycler Nexus Thermocycler with the following conditions: 2 min at 95 °C followed by 45 cycles of 95 °C for 30 s, 49.5 °C for 1 min, and 72 °C for 30 s. PCR products were visualized using an Agilent 2100 Bioanalyzer system.

Oil Red O staining

To determine whether pparγ knockdown affected lipid homeostasis, embryos were injected with either water, nc-MOs, or pparγ-MOs, reared in particulate-free system water, and imaged under transmitted light at 6, 24, 48, 72, and 96 hpf. At each stage, a subset of embryos (seven per stage) were fixed in 4% paraformaldehyde (PFA)/1× phosphate-buffered saline (PBS) for 24 h and then transferred to 1× PBS. Fixed embryos were stained with ORO using previously described protocols (Passeri et al., 2009). Stained embryos were then imaged under transmitted light at 4× magnification using a Leica MZ10 F stereomicroscope equipped with a DMC2900 camera.

Ciglitazone and DMP co-exposures

Embryos were exposed to either vehicle (0.2% DMSO), 12.5 μM ciglitazone, 0.078 μM DMP, or 12 μM ciglitazone + 0.078 μM DMP from 4 to 24 hpf in glass petri dishes (20 embryos per replicate per timepoint; three replicates per treatment). Maximum tolerated concentrations for DMP and ciglitazone co-exposures were optimized based on preliminary experiments that tested combinations of 0.078 or 1.56 μM DMP in the presence or absence of 9.375, 12.5, or 15 μM ciglitazone. Although 0.625 µM DMP was used in our prior studies (Dasgupta et al., 2017; Dasgupta et al., 2018), we relied on 0.078 µM DMP since co-exposure with 0.625 μM DMP and 12.5 μM ciglitazone resulted in a significant increase in mortality. At 8 hpf, embryos were fixed overnight in 4% PFA/1× PBS. Fixed embryos were manually dechorionated and then incubated overnight with anti-phosphoSMAD 1/5/9 IgG antibody (1:100 dilution; Cell Signaling Technology, Danvers, MA, USA) using previously described protocols (Yozzo, McGee & Volz, 2013; Dasgupta et al., 2018). Embryos were then incubated overnight with an IgG-specific Alexa Fluor-conjugated secondary antibody (1:500 dilution; Millipore Sigma, St. Louis, MO, USA). Embryos were imaged at 8× magnification using a Leica MZ10 F stereomicroscope equipped with a GFP filter and DMC2900 camera. At 24 hpf, embryos were imaged under transmitted light at 2× magnification using a Leica MZ10 stereomicroscope equipped with a DMC2900 camera and assessed for survival and dorsoventral patterning abnormalities as described above.

mRNA-sequencing

To quantify potential effects of ciglitazone on the transcriptome, embryos were exposed to vehicle (0.2% DMSO), 12.5 μM ciglitazone, 0.078 μM DMP, or 12.5 μM ciglitazone + 0.078 μM DMP (20 embryos per dish; two dishes per replicate; four replicates per treatment) from 4 to 24 hpf and then immediately snap-frozen in liquid nitrogen at 24 hpf and stored at −80 °C. All embryos were homogenized in two mL cryovials using a PowerGen Homogenizer (Thermo Fisher Scientific, Waltman, MA, USA). Following homogenization, an SV Total RNA Isolation System (Promega, Madison, WI, USA) was used to extract total RNA from each replicate sample per the manufacturer’s instructions. RNA quantity and quality were confirmed using a Qubit 4.0 Fluorometer and 2100 Bioanalyzer system, respectively. Based on sample-specific Bioanalyzer traces, the RNA Integrity Number was >8 for all RNA samples used for library preparations.

Libraries were prepared using a QuantSeq 3′ mRNA-Seq Library Prep Kit FWD (Lexogen, Vienna, Austria) and indexed by treatment replicate per manufacturer’s instructions. Library quantity and quality were confirmed using a Qubit 4.0 Fluorometer and 2100 Bioanalyzer system, respectively. Raw Illumina (fastq.gz) sequencing files (16 files) are available via NCBI’s BioProject database under BioProject ID PRJNA544341, and a summary of sequencing run metrics are provided in Table S1 (>87.17% of reads were ≥Q30 across all runs). All 16 raw and indexed Illumina (fastq.gz) sequencing files were downloaded from Illumina’s BaseSpace and uploaded to Bluebee’s genomics analysis platform to align reads against zebrafish genome assembly GRCz10. After combining treatment replicate files, a DESeq2 application within Bluebee (Lexogen Quantseq DE1.2) was used to identify significant treatment-related effects on transcript abundance (relative to control) based on a false discovery rate p-adjusted value ≤ 0.05. Significantly affected transcripts were imported into the Database for Annotation, Visualization, and Integrated Discovery (DAVID) v6.8 for Gene Ontology (GO) enrichment analysis. Individual transcripts from significant GO terms (Benjamini score ≤ 0.05) were consolidated into a list of unique transcripts.

Statistical analysis

For data derived from exposures and MO injections, a general linear model (GLM) analysis of variance (ANOVA) (α = 0.05) was performed using SPSS Statistics 24, as these data did not meet the equal variance assumption for non-GLM ANOVAs. Treatment groups were compared with vehicle controls using pair-wise Tukey based multiple comparisons of least square means to identify significant treatment-related differences.

Zebrafish and mammalian PPARs are highly conserved

As expected, when comparing protein sequences of PPARs from human, mouse, rat, and zebrafish, we found that zebrafish PPARα (a/b), PPARδ (a/b), and PPARγ are closely related to mammalian PPARα, PPARδ, and PPARγ, respectively (Fig. 1A). For each PPAR, the DNA binding domain (DBD) and ligand binding domain (LBD) were highly conserved across all four species (Fig. 1B; Fig. S1). For example, the percent similarity between full-length zebrafish and human PPARγ was 74.9%, whereas the similarity between the DBD and LBD was 97.6% and 80.3%, respectively (Fig. 1B)—a finding that was similar to PPARα and PPARδ (Fig. S1). When comparing similarity across zebrafish-specific PPARs, similarity in the LBD was highest (72.7%) between PPARγ and PPARδb (Fig. S2). Finally, when comparing ligand binding pocket amino acid residues involved with rosiglitazone binding to human PPARγ (Sheu et al., 2005), we found that eight out of 13 residues that interact with thiazolidinediones are conserved between humans and zebrafish (Fig. S3).

Knockdown of pparγ adversely affects development within the first 96 h

Although there are elevated levels of pparγ transcripts between 0.75 and 5 hpf (due to maternally-loaded pparγ mRNA), zygotic transcription of pparγ mRNA does not occur until ~48 hpf (Fig. 2A). Similarly, maternally-loaded pparαa, pparαb, pparδa, and pparδb are all present within the embryo until 5 hpf (Fig. S4). While zygotic transcription of pparαa, pparαb, and pparδa is not initiated until ~24–30 hpf (Fig. S4), zygotic transcription of pparδb is initiated at 5 hpf and peaks at approximately 24 hpf (Fig. S4).

Injection of pparγ-MO resulted in insertion of intronic sequence within zygotic pparγ transcripts by 24 hpf (Fig. 2B). Within the first 48 h of development, injection of nc-MOs and pparγ-MOs resulted in mild defects on dorsoventral patterning relative to water-injected embryos (Figs. 2C–2FF). However, unlike nc-MO-injected embryos, injection of pparγ-MOs resulted in more severe developmental abnormalities including pericardial edema, cardiac looping defects, and stunted growth at 72 and 96 hpf (Fig. 2C–2FF)—a stage that coincides with a sharp increase in transcription of zygotic pparγ mRNA (Fig. 2A). However, the abundance of neutral lipids (as determined by ORO staining) within pparγ-MO-injected embryos was not qualitatively different across all stages (Fig. 2C–2FF).

Knockdown of pparγ does not block ciglitazone-induced toxicity at 24 hpf

Although ciglitazone exposure from 4 to 6 hpf did not impact epiboly at 6 hpf (Fig. 3A), initiation of ciglitazone exposure at 4 hpf resulted in a concentration-dependent effect on survival and dorsoventral patterning by 24 hpf (Figs. 3B–3F). Embryos injected with water, nc-MOs, or pparγ-MOs, and then exposed to vehicle (0.2% DMSO) from 4 to 24 hpf, exhibited mild effects on survival and dorsoventral patterning (Figs. 4A–4G). However, pparγ knockdown did not block dorsoventral patterning defects following exposure to ciglitazone from 4 to 24 hpf (Figs. 4A–4G).

DMP reverses the ventralizing effects of ciglitazone

While exposure to 12.5 µM ciglitazone resulted in ventralized and delayed embryos, the majority of embryos following co-exposure with 12.5 µM ciglitazone + 0.078 µM DMP were dorsalized (Figs. 5A–5E). Although exposure to 0.625 µM DMP (a positive control) disrupted BMP signaling as expected (Fig. 5F–5J), exposure to 12.5 µM ciglitazone, 0.078 µM DMP, or 12.5 µM ciglitazone + 0.078 µM DMP did not result in disruption of BMP signaling at 8 hpf (Fig. 5F–5J).

Ciglitazone exposure impacts cholesterol- and lipid-related biological processes by 24 hpf

Exposure to 12.5 μM ciglitazone, 0.078 μM DMP, or 12.5 μM ciglitazone + 0.078 μM DMP resulted in a significant change in the abundance of 1,641, 1,031, and 1,924 transcripts, respectively, relative to vehicle controls (Figs. 6A–6C; Tables S2–S4). Although the magnitude of affected transcripts was similar across all three treatment groups, there was a total of 580, 289, and 724 significantly affected transcripts that were unique to embryos exposed to 12.5 μM ciglitazone, 0.078 μM DMP, or 12.5 μM ciglitazone + 0.078 μM DMP, respectively (Fig. 6D). Interestingly, the most significantly altered DAVID-based biological process across all treatment groups was translation (Fig. 7A; Tables S5–S7). While all three treatment groups shared certain biological processes that were affected (Fig. 7B; Fig. S5), exposure to ciglitazone alone primarily affected cholesterol- and lipid-related biological processes (Fig. 7C)—an effect that was driven by transcripts specific to apolipoprotein A-IV a (apoa4a), A-IV b (apoa4b.2), A-Ia (apoa1a), A-Ib (apoa1b), Eb (apoeb), antifreeze protein type IV (afp4), and an unnamed transcript (zgc: 162608) (Table S8).