https://orcid.org/0009-0006-9809-1095
Figures
Abstract
Fibroblast growth factor receptor 4 (FGFR4) has a role in many biological processes, including lipid metabolism, tissue repair, and vertebrate development. In recent years, FGFR4 overexpression and activating mutations have been associated with numerous adult and pediatric cancers. As such, FGFR4 presents an opportunity for therapeutic targeting which is being pursued in clinical trials. To understand the role of FGFR4 signaling in disease and development, we generated and characterized three alleles of fgfr4 knockout zebrafish strains using CRISPR/Cas9. To generate fgfr4 knockout crispants, we injected single-cell wildtype zebrafish embryos with fgfr4 targeting guide RNA and Cas9 proteins, identified adult founders, and outcrossed to wildtype zebrafish to create an F1 generation. The generated mutations introduce a stop codon within the second Ig-like domain of Fgfr4, resulting in a truncated 215, 223, or 228 amino acid Fgfr4 protein compared to 922 amino acids in the full-length protein. All mutant strains exhibited significantly decreased fgfr4 mRNA expression during development, providing evidence for successful knockout of fgfr4 in mutant zebrafish. We found that, consistent with other Fgfr4 knockout animal models, the fgfr4 mutant fish developed normally; however, homozygous fgfr4 mutant zebrafish were significantly smaller than wildtype fish at three months post fertilization. These fgfr4 knockout zebrafish lines are a valuable tool to study the role of FGFR4 in vertebrate development and its viability as a potential therapeutic target in pediatric and adult cancers, as well as other diseases.
Citation: Harrison EN, Jay AN, Kent MR, Sukienik TP, LaVigne CA, Kendall GC (2024) Engineering an fgfr4 knockout zebrafish to study its role in development and disease. PLoS ONE 19(11): e0310100. https://doi.org/10.1371/journal.pone.0310100
Editor: Yevgenya Grinblat, University of Wisconsin-Madison, UNITED STATES OF AMERICA
Received: May 8, 2024; Accepted: August 20, 2024; Published: November 22, 2024
Copyright: © 2024 Harrison et al. This is an open access article distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original author and source are credited.
Data Availability: All data generated in this study are included in this article and supplementary files. We are depositing all three fgfr4 knockout zebrafish lines, nch4, nch5, and nch6 at the Zebrafish International Resource Center (ZIRC; https://zfin.org/ZDB-LAB-191120-1). Pending this, data availability requests for the fgfr4 knockout zebrafish lines, nch4, nch5, and nch6 should be directed to the lead contact, Genevieve Kendall (Genevieve.Kendall@NationwideChildrens.org), Lindsey Ferguson (Lindsey.Ferguson@NationwideChildrens.org), or Logan Fehrenbach (Logan.Fehrenbach@NationwideChildrens.org), and are available with a completed Materials Transfer Agreement.
Funding: A.N.J. is supported by the Undergraduate Research Apprenticeship Program at The Ohio State University. M.K. is supported by a T32CA269052 Training Program in Basic and Translational Pediatric Oncology Research postdoctoral fellowship. G.C.K. is supported by an Alex’s Lemonade Stand Foundation A Award, a V Foundation for Cancer Research V Scholar Grant, and an R01 CA272872 grant through the National Cancer Institute of the National Institutes of Health. The funders had no role in study design, data collection and analysis, decision to publish, or preparation of the manuscript. The content is solely the responsibility of the authors and does not necessarily represent the official views of the National Institutes of Health.
Competing interests: The authors have declared that no competing interests exist.
Introduction
Fibroblast growth factor 4 (FGFR4) is a member of the fibroblast growth factor receptor family, a group of receptor tyrosine kinases that bind promiscuously to fibroblast growth factors (FGFs) and play a role in a wide range of biological processes. FGFR family members regulate a wide range of processes including organogenesis, neural development, metabolism, and inflammation [1]. FGFR4 signaling is critical in chordate development, tissue repair, and lipid metabolism. Additionally, FGFR4 has been implicated as a crucial mediator of myogenesis and muscle regeneration in mouse and chick models [2, 3].
In recent years, FGFR4 has been implicated as a potential therapeutic target in rhabdomyosarcoma (RMS), a pediatric soft tissue sarcoma. RMS is most often associated with displaying molecular features of the skeletal muscle lineage and is characterized by its failure to terminally differentiate into mature muscle [4, 5]. Though caused by multiple distinct genetic drivers, the subtype in which FGFR4 has been most often implicated is fusion-positive rhabdomyosarcoma. This subtype also has poorer patient prognosis compared to fusion-negative RMS subtypes [6, 7]. Fusion-positive RMS is most often genetically driven by a single chimeric fusion oncogene PAX3/7::FOXO1 which is caused by a chromosomal translocation event wherein the DNA binding domain of PAX3/PAX7 is juxtaposed to the transactivation domain of FOXO1 [8–10]. FGFR4 is transcriptionally upregulated in fusion-positive RMS, and its overexpression is a predictor for reduced overall survival in patients [11, 12]. FGFR4 has also been established as a direct transcriptional target of the PAX3::FOXO1 fusion protein [13, 14].
Beyond rhabdomyosarcoma, FGFR4 has also garnered attention in recent decades for its dysregulation in other adult and pediatric cancers. FGFR4 is frequently overexpressed in cancers, and it is prone to hotspot activating mutations in breast cancer and hepatocellular carcinoma, among others [11, 15]. FGFR4 is an exciting therapeutic target because many pan-FGFR inhibitors and FGFR4 inhibitors have been developed, three of which are currently FDA approved for cancer treatment [16]. Multiple phase 1 and phase 2 clinical trials are currently underway using select small molecule inhibitors of FGFR4 in cancers such as hepatocellular carcinoma and acute myeloid leukemia [17]. In addition, CAR T-cell therapy targeting FGFR4 in rhabdomyosarcoma is in preclinical development stages and has had success in mouse allograft models to decrease tumor burden [18].
Though broadly regarded as a therapeutic target of interest, a mechanistic understanding of the role of FGFR4 in rhabdomyosarcoma and other cancers remains unclear. Here, we have developed and thoroughly characterized three alleles of fgfr4 knockout zebrafish to fully understand the phenotypic consequences of the loss of this gene. These knockout zebrafish lines can be used to study the role of fgfr4 in biological processes related to development and disease. This knockout model can also be utilized in existing zebrafish disease models to thoroughly understand the role of FGFR4 in various diseases, including rhabdomyosarcoma and other cancers.
Methods
Zebrafish husbandry and breeding
Danio rerio were housed in an AAALAC-accredited, USDA-registered, OLAW-assured Aquaneering facility in compliance with the Guide for the Care and Use of Laboratory Animals. Vertebrate animal work was overseen by the Abigail Wexner Research Institute at Nationwide Children’s Hospital IACUC committee and Animal Resources Core. Methods of anesthesia and euthanasia followed the Guide for the Care and Use of Laboratory Animals. Anesthesia was performed using tricaine methanesulfonate (Fisher Scientific, NC0342409) immersion. Euthanasia was performed in adult zebrafish using rapid cooling followed by a secondary method. WIK zebrafish (Zebrafish International Resource Center, ZL84) served as our wildtype fish. Each strain was separately reared to adulthood (~3 months in our facility) and bred in-house by mass breeding. Larval fish (5–30 days post-fertilization) were housed at a density of approximately 15 larvae/L. Adult fish were housed at a density of 5–10 fish/L in mixed-sex groups. Fish were housed in 1.8L and 2.8L tanks on a recirculating life support system (Aquaneering, San Diego, CA) supplied with reverse osmosis water. fgfr4 mutant lines were outcrossed every two generations to maintain genetic diversity. An exchange of 10–15% of the total system water volume with fresh reverse osmosis water occurs daily. The system water was maintained at 26–27.5°C, conductivity of 450–600 μS/cm, and a pH of 7.2–7.5 using continuous monitoring via probes in the sump and an automatic dosing system. Larval fish were fed three times a day with 30,000 L-type rotifers Brachionus plicatilus gut loaded with RGComplete rotifer feed (Reed Mariculture) per 1.8L tank. Adult fish were fed twice a day with Gemma Micro 300 (Skretting).
Generating fgfr4 knockout fish
CRISPR/Cas9 knockout protocol was adapted from Talbot and Amacher, 2014 [19]. fgfr4 gRNA (IDT, see S1 Table for primer sequences) and Cas9 protein (IDT, 1081060) were injected into the single cell of WIK embryos to generate potential founders. Once sexually mature, potential founders were outcrossed to wildtype zebrafish to separate mosaic CRISPR-mediated mutations. After three fgfr4 mutant strains were identified by sequencing, heterozygous fish of each strain were in-crossed to generate homozygous zebrafish colonies.
High-resolution melt analysis (HRMA)
HRMA was performed as described previously [20]. Genomic DNA was isolated from the caudal fins of potential mutant zebrafish, as well as a minimum of three wildtype (WIK) fins as a reference group. Fin clips were incubated in 10mM Tris HCl (Sigma, 10708976001) pH 8.3, 50mM KCl (Sigma, P5405-250G), 0.3% Triton X-100 (Fisher, BP151-100), and 0.3% NP40 (Fisher, NC9375914) at 98°C for 10 minutes, then cooled to 4°C for 10 minutes. One-tenth volume of 10mg/mL proteinase K (Fisher, BP1700-100) was added to samples, which were then incubated for at least 1.5 hours at 55°C, proteinase K heat inactivated at 95°C for 10 minutes and held at 4°C until ready for use. Genomic DNA was added to 9 μL of Precision Melt Supermix (Bio-Rad, BP1700-100) and 0.5 μL of each 10 μM forward and reverse primers targeting the mutated region in fgfr4 (IDT, see S1 Table for primer sequences) and diluted to 20 μL with nuclease free water in a 384-well plate (Bio-Rad, HSP3805). The plate was run on a Bio-Rad CFX384 Real Time PCR Detection System with the following program: 95°C for 3 min; (95°C for 15 s, 60°C for 20 s, 70°C for 20 s) x 45 cycles; 65°C for 30 s; melt 65°C–95°C, 0.2°C/step hold 5 s; 95°C for 15 s. Bio-Rad Maestro and Bio-Rad Precision Melt Analysis software were used for data analysis, with known wildtype samples as the reference group.
A-tail cloning
To identify the sequence of CRISPR/Cas9-mediated mutations in the fgfr4 gene, genomic DNA used in the HRMA was used in a Phusion based PCR with 10 μM forward and reverse primers (see S1 Table for primer sequences). PCR products were purified using the Monarch PCR Cleanup Kit (NEB, T1030L). PCR products were A-tailed, ligated into the PGEM-T Easy vector (Promega, A1360), and then transformed into DH5-α cells (Fisher Scientific, FEREC0111). DH5-α cells were plated on LB-Amp plates sprayed with X-Gal/IPTG (Fisher Scientific, 21530077) and incubated overnight at 37°C. Colonies were picked, cultured overnight, and mini-prepped with QIAGEN QIAprep Spin Miniprep Kit (27106), and DNA was sequenced by Sanger sequencing through Eurofins Genomics using the SP6 sequencing primer. For sequence alignment, the wildtype reference used was Danio rerio fgfr4 cDNA from genome assembly GRCz11, NM_131430.1, chr21:37183912–37194363.
Genotyping
Zebrafish fin clips were incubated in 10mM Tris HCL pH 8.3 (Sigma, 10708976001) pH 8.3, 50mM KCl (Sigma, P5405-250G), 0.3% Triton X-100 (Fisher, BP151-100), and 0.3% NP40 (Fisher, NC9375914) at 98°C for 10 minutes, then cooled to 4°C for 10 minutes. One-tenth volume of 10mg/mL proteinase K (Fisher, BP1700-100) was added to samples, which were then incubated overnight at 55°C, proteinase K heat inactivated at 95°C for 10 minutes, and held at 4°C. The PCR reaction mix was created with NEB Phusion GC Buffer (NEB, B0519S), Phusion DNA Polymerase (NEB, M0530S), 10mM dNTPs (Fisher Scientific, 10-297-018), 10 μM forward primer and 10 μM reverse primer (IDT) according to the recommended ratios (see S1 Table for primer sequences). The samples were then amplified using the following program: 98°C x 30 sec, 98°C x 10 sec, 60°C x 30 sec, 72°C x 30 sec, repeat the previous three steps for 34 cycles (35 total cycles), 72°C x 10 min, and HOLD at 12°C. PCR products were purified using the Monarch PCR Cleanup Kit (NEB, T1030L) and digested with the TfiI restriction enzyme (NEB, R0546S) digest. Final products were run in a 1.2% agarose gel (Fisher Scientific, BP160-500) and imaged on a Bio-Rad Molecular Imager Gel Doc XR+ with Image Lab Software. With this genotyping method, the expected product size(s) for a wildtype zebrafish are 89 and 72 base pairs (bp), for a heterozygous fish are 163, 89, and 72bp, and for a homozygous fish is 163bp.
Real time quantitative polymerase chain reaction (RT-qPCR)
Wildtype WIK and fgfr4 homozygous mutant adult zebrafish were set up in breeding chambers with dividers and left overnight. The following morning, dividers were pulled, and embryos were collected. At 24 hours post fertilization, embryos of each cross were dechorionated either manually or via pronase (Sigma-Aldrich, 11459643001), euthanized, and aliquoted into samples of 12 embryos each. Extra embryo media was eluted off the embryo pellet, and samples were snap frozen and stored at -80°C.
Total RNA was isolated from each sample using the QIAGEN RNeasy Mini Kit (74104) including on-column DNAse digestion. The RT2 HT First Strand Synthesis kit (Qiagen, 330411) was used to synthesize cDNA using approximately 825 ng of total RNA input per sample. cDNA for each sample was diluted to 80 μL with nuclease-free water. 4 μL of diluted cDNA, 5 μL Universal SYBR Green Supermix (Bio-Rad, 1725122), and 0.5 μL of forward and reverse primers (see S1 Table for primer sequences) for the genes of interest were combined in a 384-well plate (Bio-Rad, HSP3805). The plate was run on a CFX384 Real Time PCR Detection System using the following program: 95°C for 2 min; (95°C for 15 s, 60°C for 1 min, Plate Read) x 40 cycles; 65°C for 30 s; (65°C, 0.5°C/step, Plate Read) x 60 cycles. Five biological replicates were used per strain with three technical replicates each. Data was analyzed using the CFX Maestro program.
Histology
Male and female adult homozygous mutant zebrafish from each strain were fixed in 4% PFA (Fisher Scientific, 50-276-31) in PBS for a minimum of 24 hours at room temperature. Cassettes were then transferred into 0.5M EDTA pH 7.8 (Fisher Scientific, BP120-1) for five days, after which zebrafish were paraffin embedded, sagittally sectioned, and hematoxylin and eosin (H&E) stained as described previously [21].
Imaging of zebrafish embryos and adults
Images of 24 hours post-fertilization (hpf) and 48hpf zebrafish embryos were taken on a Leica M205FA dissecting microscope. Images of adult zebrafish were taken by anesthetizing zebrafish, placing them on laminated grid paper, and taking images with a smartphone camera. Standard length was measured from anterior-most part of the head to the posterior-most part of the tail in embryos, or to the caudal fork of the tail fin in adults.
Results
Knockout fgfr4 zebrafish were generated using a CRISPR/Cas9 strategy (Fig 1A). Guide RNA against zebrafish fgfr4 and Cas9 protein were injected into single-cell wildtype embryos. After outcrossing these founders to wildtype to isolate individual CRISPR-mediated germline mutations, genomic DNA from putative F1 mutant fish was used for high resolution melt analysis (HRMA) to identify differences in melting temperature compared to wildtype references, which is indicative of genetic mutations. HRM analysis identified three clusters with inflection curves distinct from the wildtype references (Fig 1B). Sub-cloning and Sanger sequencing revealed that each of the clusters possessed one of the following mutations in the fgfr4 locus when aligned to wildtype fgfr4 cDNA: a 7 base pair (bp) deletion at 604-610bp (GATTCAG/-) and 6bp deletion at 644-649bp (AGGCTC/-), two 1bp insertion after 605bp (-/G) and 606bp (-/A), one substitution at 609bp (A/T), 38bp deletion at 609-646bp (CAGGAGTATATGTGTGTATGCTACGTGGCACCAAAGAG/-), and a 1bp insertion after 669bp (-/G) (S1 Fig). Each of these genomic mutations conferred premature stop codons in the fgfr4 sequence, resulting in truncated protein sequences with predicted lengths of 223, 228, and 215 amino acids (Fig 1C). These alleles are referred to as fgfr4nch4, fgfr4nch5, and fgfr4nch6, respectively. Because the kinase domain of Fgfr4 is lost in the generated mutants, all downstream wildtype function should also be lost.
(A) Schematic of generation of fgfr4 knockout zebrafish. Single cell wildtype zebrafish embryos were injected with fgfr4 guide RNA and Cas9 protein to generate mutant founders. Potential founders were outcrossed to isolate CRISPR/Cas9-mediated mutations, and putative crispant F1s were subjected to high resolution melt analysis to identify mutation sequences. (B) High resolution melt analysis (HRMA) of potential F1 mutants revealed three distinct clusters of potential fgfr4 crispants. (C) Schematic of wildtype Fgfr4 and predicted knockout strain protein lengths. Premature stop codons produced truncated Fgfr4 proteins with predicted length of 223, 228, and 215 amino acids respectively. Solid blocks indicate regions of wildtype Fgfr4 amino acid sequence alignment. The protein sequence aligned to the wildtype Fgfr4 protein ends at the same amino acid for all three alleles, with various additions of amino acids that do not align to the wildtype protein afterwards. Hatching indicates these additional amino acids.
We used real time quantitative polymerase chain reaction (RT-qPCR) as a proxy to validate the fgfr4 genomic knockout in all strains. To assess fgfr4 mRNA abundance, we designed RT-qPCR primers to target regions both 5’ and 3’ of the mutation site in all strains (Fig 2A). In all putative knockout strains, relative expression of fgfr4 mRNA was significantly decreased in homozygous fgfr4 knockout embryos compared to wildtype embryos (Fig 2B and 2C), indicating that the fgfr4 mRNA transcript is less stable, indicative of a true knockout. Because it has the lowest fgfr4 mRNA expression of all strains, the fgfr4nch4 line was used for the remainder of the described experiments, though all other strains had consistent phenotypes with those described below (data not shown).
(A) Schematic of wildtype zebrafish fgfr4 mature mRNA transcript and regions targeted for RT-qPCR. Blue triangle indicates approximate fgfr4 crispant mutation site for all strains relative to primers. Orange box indicates region 5’ of mutation site targeted by RT-qPCR primers, and green box indicates region 3’ of mutation site targeted by RT-qPCR primers. Relative expression of fgfr4 mRNA as measured by RT-qPCR targeting the regions 5’ (B) and 3’ (C) of the sequence mutation. Expression is normalized to gapdh and rpl13a. Each point represents a pool of n = 12, 24 hours post-fertilization (hpf) zebrafish embryos derived from a maternal zygotic. Multiple points represent biological replicates, while three technical replicates were used to generate each biological replicate. Error bar is the mean ± standard deviation. P values were calculated using a one-way Brown-Forsythe and Welch ANOVA, correcting for multiple comparisons with a Dunnett T3 test.
Homozygous fgfr4 mutant zebrafish embryos did not appear phenotypically different from wildtype zebrafish at early developmental timepoints (Fig 3A). Homozygous fgfr4 mutant zebrafish are not significantly different in size than stage-matched wildtype embryos at 24- and 48-hours post-fertilization (hpf). This standard early embryonic development is consistent with other zebrafish fgfr4 knockout models [22]. Further, all three fgfr4 mutant zebrafish alleles were found to exhibit Mendelian ratios (S2 Table), and homozygous fgfr4 fish were viable and fertile. Sex distributions were also standard in fgfr4 homozygotes (S3 Table). However, a size discrepancy between fgfr4 homozygotes and wildtype fish became apparent at roughly three months post-fertilization, wherein fgfr4 homozygotes are significantly smaller than age-matched wildtype zebrafish (Fig 3B), though this size discrepancy becomes nonsignificant at six months post-fertilization (S2 Fig). However, the internal anatomical histology of adult fgfr4 knockout zebrafish appeared overall normal (Fig 3C).
(A) Representative images from a phenotypic analysis of embryonic zebrafish. Homozygous fgfr4 mutant zebrafish do not display an early embryonic phenotype. Each point represents an individual fish standard length, and the bar represents the mean. An unpaired two-tailed t-test with Welch’s correction was used to calculate the p value. Scale bar is 560.1 μm in 24 hpf images and 700.4 μm in 48 hpf images. Standard length quantification was performed with n = 15 embryos per group. (B) Phenotypic analysis of adult zebrafish. Homozygous knockout fgfr4 zebrafish are significantly smaller than wildtype zebrafish at three months post fertilization. Standard length quantification was performed with n = 27 WT fish and n = 34 fgfr4 KO adult fish. Each point represents an individual fish standard length, and the bar represents the mean. An unpaired two-tailed t-test with Welch’s correction was used to calculate p value. Scale bar is 1 cm. (C) Representative hematoxylin and eosin (H&E) staining of a sagittal section from female and male adult fgfr4 knockout zebrafish. Internal anatomy in mutant zebrafish is normal in both male and female adult fgfr4 knockout zebrafish. This was assessed in 5 female and 3 male fgfr4 knockout fish total. Scale bar is 1mm.
Discussion
Here, we developed three fgfr4 knockout strains through a CRISPR/Cas9 single-cell injection strategy. Putative knockout strains were identified by HRMA, and the loss of fgfr4 expression was validated by RT-qPCR. Because all mutations confer premature stop codons that eliminate the kinase domain of Fgfr4, normal Fgfr4 activity should be removed in all mutants. Fgfr4 is not duplicated in zebrafish indicating that this single knockout should remove all Fgfr4 activity. The fgfr4 mutant strains were embryonically viable and were phenotypically normal at early developmental timepoints. However, at approximately three months post fertilization, juvenile fgfr4 knockout zebrafish are significantly smaller than age-matched wildtype fish. Despite their relatively small size at this timepoint, mutant fgfr4 zebrafish have normal internal anatomy and are fertile.
The results seen in this study recapitulate what has been seen in existing fgfr4 animal models. Similarly to zebrafish, Fgfr4 knockout mice are embryonically viable, develop normally, and experienced no impacts on fertility [23]. Fgfr4 knockout mice also metabolize high-fat diets differently than wildtype mice [24]. An fgfr4 zebrafish knockout model was similarly viable and developed normally in early stages of development [22], though adult phenotypes were not fully characterized. In zebrafish, the viability and lack of embryonic phenotype is likely a result of compensation from other FGFR family members [22].
The smaller size of fgfr4 mutant zebrafish in the juvenile stage has also been noted in Fgfr4 knockout mice, which weigh 10% less than their wildtype siblings at the time of weaning [23]. This difference in size may be due to the established and conserved role of fgfr4 in cholesterol metabolism. Because zebrafish embryos obtain all nutrition from the yolk until about five days post-fertilization, the metabolism-related phenotypic outcomes of an fgfr4 knockout may not appear until larvae are able to feed freely.
In the future, we plan to use these knockout zebrafish to understand the genetic cooperation between FGFR4 and PAX3::FOXO1, the predominant driver of fusion-positive rhabdomyosarcoma. We have found that when human PAX3::FOXO1 is incorporated into the zebrafish genome, we obtain tumors that histologically and molecularly recapitulate human RMS [25]. We hypothesize that incorporating PAX3::FOXO1 into the genome of fgfr4 knockout zebrafish, tumor occurrence and tumor volume will be significantly reduced. The fgfr4 knockout zebrafish strains described here are a valuable tool in the study of the role of FGFR4 in vertebrate development and disease.
Supporting information
S2 Table. Genotypes of fgfr4 knockout strains.
Genotypes obtained from heterozygous in-crosses of each fgfr4 knockout strain. Differences between observed and expected wildtype, heterozygous, and homozygous genotype distribution were nonsignificant by chi-square statistical tests. P values for actual and expected genotypes were 0.2950, 0.2706, and 0.3917 for fgfr4nch4, fgfr4nch5, and fgfr4nch6 respectively.
https://doi.org/10.1371/journal.pone.0310100.s002
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S3 Table. Sex split of homozygous fgfr4 knockout zebrafish.
Sex split data obtained from homozygous fgfr4 mutant zebrafish colonies. Differences between male and female observed and expected sex distributions were nonsignificant by chi-square statistical tests. P values for actual and expected sexes were 0.6184, 0.7891, and 0.5623 for fgfr4nch4, fgfr4nch5, and fgfr4nch6 respectively.
https://doi.org/10.1371/journal.pone.0310100.s003
(PDF)
S1 Fig. Mutation sequences identified by Sanger sequencing.
(A) Mutation sequence alignments to wildtype fgfr4 cDNA. Wildtype reference is Danio rerio fgfr4 cDNA from genome assembly GRCz11, NM_131430.1, chr21:37183912–37194363. Strain fgfr4nch4 contains a 7 base pair (bp) deletion at 604-610bp (GATTCAG/-) and 6bp deletion at 644-649bp (AGGCTC/-), strain fgfr4nch5 has two 1bp insertion after 605bp (-/G) and 606bp (-/A), and strain fgfr4nch6 has one substitution at 609bp (A/T), 38bp deletion at 609-646bp (CAGGAGTATATGTGTGTATGCTACGTGGCACCAAAGAG/-), and a 1bp insertion after 669bp (-/G). Highlighting indicates inserted and mismatched base pairs. (B) Trace sequencing files for all fgfr4 mutants.
https://doi.org/10.1371/journal.pone.0310100.s004
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S2 Fig. Homozygous fgfr4 knockout zebrafish are not significantly smaller than wildtype zebrafish at six months post-fertilization.
Standard length quantification was performed with n = 14 Wildtype (WT) fish and n = 12 fgfr4 KO fish. Each point represents an individual fish standard length, and the bar represents the mean. An unpaired two-tailed t-test with Welch’s correction was used to calculate the p value. Scale bar is 1 cm.
https://doi.org/10.1371/journal.pone.0310100.s005
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Acknowledgments
We would like to acknowledge the Animal Resources Core at Nationwide Children’s Hospital for phenomenal caretaking of our zebrafish colonies, in particular Dr. Laurie Goodchild, Dr. Carmen Arsuaga, Dr. Lindsey Ferguson, Logan Fehrenbach, Logan Bern, and Alexander Kramer. We also thank the Histopathology Core and the Processing and Banking Core at Nationwide Children’s Hospital for the processing and imaging of our histology samples.
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