The NICHD Zebrafish Core
- Benjamin Feldman, PhD, Staff Scientist and Director of the NICHD Zebrafish Core
- ChonHwa Tsai-Morris, PhD, Staff Scientist and Assistant Director of the NICHD Zebrafish Core
The NICHD Zebrafish Core was established in May 2012. The goal of the Core is to provide its clients with consultation, access to equipment and reagents, and service in the area of zebrafish genetics. NICHD investigators as well as investigators from other NIH institutes and from outside the NIH are its clientele. The oversight committee for the Core comprises Thomas Sargent (Chair), Harold Burgess, Ajay Chitnis, Igor Dawid, and Brant Weinstein. The Core's activities consist of (1) oversight and support of client-specific projects; (2) maintenance and improvement of equipment and infrastructure; (3) core-initiated research, also in the field of developmental biology; and (4) service and educational outreach.
Oversight and support of client-specific projects
In the past year, we engaged in research projects with seven labs: four from NICHD, one from NCI, one from NHLBI, and one from the Children’s National Medical Center.
Porter Lab (NICHD): Zebrafish models of human pediatric diseases.
Smith-Lemli-Opitz syndrome (SLOS) is an autosomal recessive, multiple malformation syndrome with pediatric onset characterized by intellectual disability and aberrant behavior. Phenotypic characterization is ongoing of zebrafish carrying mutant alleles of dhcr7, the zebrafish ortholog to human DHCR7 gene, which were generated with support from the Core in previous years. This project will continue in 2917. The Core also created three novel lines for the Porter lab, carrying mutations in either the npc1, npc2, or cln3 gene. Phenotypic characterization by the Porter lab is ongoing.
Stratakis Lab (NICHD): Function of zebrafish orthologs to human genes implicated in disorders of the pituitary-adrenal axis.
Gigantism arises due to excess growth hormone secretion during childhood, before the growth plates close. Since 2012, the Core has supported this lab’s investigation of the zebrafish ortholog to a human gene implicated as a driver of gigantism. The lab published a paper in June 2016, with Feldman as co-author, which included a description of the gene’s developmental expression in zebrafish. This year, the Stratakis lab also began to test the effect on growth and development of zebrafish in which this gene is chronically overexpressed in tissue-specific or ubiquitous locations, using the Gal4/UAS transgene system. The studies will continue in 2017. This year, the Core used Crispr/Cas9 methods to generate for the Stratakis lab zebrafish carrying loss-of-function mutations in the above gene as well as in three other zebrafish orthologs to genes implicated human growth anomalies. Characterization of the resulting phenotypes will begin in 2017.
Since 2012, the Core has also supported this lab’s investigation of the function of two zebrafish orthologs to human adrenal hyperplasia genes. Over previous years, the Core helped the lab generate and acquire zebrafish carrying loss-of-function mutation for each of these orthologs. Phenotypic characterization has begun this year; notable effects on juvenile growth in the case of one gene and on early embryogenesis in the case of the other were found. This project will continue into 2017.
Kaler Lab (NICHD). Modeling copper deficiency-associated distal motor neuropathy.
The Menkes' gene encodes ATP7A, a copper-binding ATPase localized to the plasma membrane and the trans-Golgi network, which is critical for proper intracellular copper distribution. Two ATP7A missense mutations cause a milder syndrome than Menkes disease, a distal motoneuropathy that is nevertheless debilitating to children and young adults. Since 2013, the Core has supported a project to clarify the structure-function relationship of ATP7A and motor neuron defects from the perspective of these missense mutations. This project was awarded a Bench-to-Bedside grant, with $10,000 per annum funding to the NICHD Zebrafish Core through FY 2016–2017. Over previous years, the Core supported the Kaler lab’s work to visualize and compare motor neuron growth during embryogenesis of wild-type zebrafish embryos and embryos homozygous for null mutations in their ATP7A ortholog, atp7a. The comparison will continue into 2017. In parallel with these studies, Feldman and Tsai-Morris have made a concerted effort to establish genome-editing technology in the Core, with the initial goal of inducing formation of zebrafish atp7a point mutations cognate to one of the human ATP7a motoneuropathy alleles. As described above, this work will continue into 2017.
Tanner Lab (NCI). Assessing human metastatic cell behaviors in a whole-body (zebrafish embryo) microenvironment.
The dual goal of this project is to (1) determine the trophic range of certain metastatic melanoma and breast carcinoma cell lines and (2) document cellular dynamics during early tumor formation from metastatic cells that have seeded into new microenvironments. Feldman assisted the Tanner lab during the inception phase in 2013–2014 and to a lesser degree in 2014–2015. The Tanner lab worked mostly independently in 2015–2016 and the Core’s chief role has been to maintain and manage access to needed equipment, fish tank space, and reagents. The Tanner lab has developed independent resources and will not need any further core support 2017.
Tuchman Lab (Children’s National Medical Center). Finding neuroprotective drugs to mitigate hyperammonemia, a consequence of urea cycle defects and liver failure.
Exposure of the brain to high ammonia causes neurocognitive deficits, intellectual disabilities, coma, and death. Since 2012, the Core has helped this lab use zebrafish embryos to identify small molecules able to diminish the effects of hyperammonemia. Over previous years, a library of hundreds of small molecules with known safety profiles for humans was screened, and several promising candidates were identified for follow-up validation studies in zebrafish and other animal models. This year, the Core helped the Tuchman lab increase throughput of this screen, bolstered by additional personnel from the Tuchman lab and by the Core’s implementation of NICHD’s massive embryo production systems as a source of embryos.
Maintenance and improvement of services, equipment, and infrastructure
The ‘do-it-yourself and pay-as-you-go’ model of facilitated, supervised research is working well and continues for complex and ad hoc projects. Additional ad hoc services include participation in monthly project-specific meetings, RNA syntheses, zebrafish line preservation via sperm freezing, and line recovery via in vitro fertilization. The core now also offers to generate zebrafish carriers of novel mutant alleles as a fee-for-service.
The Core also acquired the following: a state-of-art automated whole-mount in situ hybridization machine; a new camera and microscope adaptor components to establish a new photomicroscopy station; a portable UV illumination system and microscope blackout hood to visualize fluorescent zebrafish lines in the lab and for educational outreach activities. A new generation of wild-type zebrafish was introduced and new generations were established for other mutant and transgenic lines frequently needed for Core projects and/or educational outreach. The Core also added a new line of fish to its collection: a ubiquitously expressed Gal4 line.
Adopting emerging reverse genetic tools
Crispr/Cas9 technology has made gene targeting in zebrafish straightforward and has opened the door to more elaborate genetic modifications, widely referred to as “precise gene editing.” Over the past year, the Core acquired, adapted, and optimized reagents and protocols for Crispr/Cas9 gene disruption. The project has been greatly bolstered by the addition of a second Staff Scientist to the NICHD Zebrafish Core: Chon-Hwa Tsai-Morris. Dr. Tsai-Morris’s main assignment has been to improve these genetic tools and generate mutants for NICHD Zebrafish Core customers, as a new service. The Core has thus created and provided zebrafish carrying novel mutations in 15 distinct genes for the Weinstein, Sargent, Stratakis, Porter, and Kaler labs. In the process of getting Crispr-Cas9 up and running, the Core targeted an additional nine genes, for an overall success rate of over 75% per locus-specific disruption construct, known as a gRNA. The Core also established an optional quality-control strategy for selecting maximally mutagenized lines, which will be particularly useful for zebrafish and Crispr/Cas9 newcomers. The above data were presented at the Spring 2016 Mid-Atlantic Regional Zebrafish Meeting, the Spring 2016 Mid-Atlantic Regional Society for Developmental Biology Meeting, and the 2016 Allied Genetics Conference.
As part of a Bench-to-Bedside-funded project with the Kaler lab, the Core most recently began to explore and compare strategies for precise genome editing, whereby Crispr/Cas9–based gene disruption reagents are combined with a DNA donor sequence in order to generate zebrafish carrying atp7a point mutations; these are homologous to human ATP7A point mutations that the Kaler lab is studying in a class of patients with distal motoneuropathy. Precise gene editing is not as efficient as gene disruption, and zebrafish labs have reported varying levels of success, with relatively labor-intensive strategies required throughout. Through literature review and discussions at conferences, Feldman and Tsai-Morris selected two methods as the most promising. Using an initial gRNA targeting the atp7a gene and a single-stranded DNA donor sequence, we found that pre-synthesized Cas9 protein is better suited than in vitro–synthesized Cas9 RNA, data that was presented by Feldman in a late-breaking data talk at the 2016 Allied Genetics Conference. We also found that new gRNA target sites were likely needed. Accordingly, we identified an additional six gRNA targets in the Atp7a gene and found that two could be disrupted at suitable levels.
Future work will utilize these targets to test two classes of DNA donor sequences, respectively. First, we will use a gRNA site in one of Atp7a’s exons in combination with a single-stranded DNA donor for homology-directed repair. Second, we will use a gRNA site in an adjacent intron in combination with linearized double-stranded DNA for homologous recombination.
Development of other tools to facilitate research with zebrafish at NICHD
Automating immunohistochemistry.
A great deal of researcher time and effort goes into the manual processing of zebrafish embryos and tissue samples. Automated liquid exchange instruments can reduce the labor, but some models, including the Core’s recently retired model, required a prohibitively large antibody volume, leaving researchers using valuable antibodies with no automated option. To remedy this, the Core acquired a smaller-volume liquid-exchange system in November 2015, which is also designed for whole-mount procedures (Intavis Insitu Pro). The machine can also be used for immunohistochemistry of tissues from other organisms. Protocols for visualizing and localizing gene expression at both the protein and RNA levels were adapted to this instrument. To permit further reduction in the volume of valuable liquid reagents, the Core designed and 3D printed an adaptor sleeve, with help from Harold Burgess’s lab. Protein detection works very well, but the sensitivity of RNA detection does not yet match manual protocols. Future work, therefore, will be to identify protocol modifications that increase the sensitivity of RNA detection. Future work will also test an alternate tissue-holding unit to further reduce the required volume of valuable liquid reagents.
Implementing mass embryo production.
As part of the Central Aquatics Facility, NICHD has two large and two small mass embryo-production systems (MEPS), distributed between two of our procedure rooms. Initial optimization by the Research Animal Management Branch (RAMB) and Charles River staff already had one of the two larger MEPS running on a continuous basis, with one embryo collection per week. However, NICHD research through the Core and otherwise requires embryos more frequently. For small-molecule screens, at least 1500 embryos per experimental day are required. For microinjection experiments, two morning waves of at least 500 synchronously fertilized embryos are required. To achieve these goals, the Core made the following changes, working together with RAMB and Charles River: (1) The MEPS light cycle was adjusted to 9 am on and 11 pm off. (2) A second large MEPS was brought on line and seeded with 3000 visually curated healthy embryos derived from a wild-type EK stock (EK: Ekkwill strain) that was maximized for robustness via genetic diversity (Weinstein lab). (3) Spawning platforms are presented to these fish on Tuesdays, Wednesdays, and Thursdays at 9:00 am followed by a 9:45 am collection, and again at 9:45 am followed by a 10:45 am collection. (4) To eliminate spill-through of older embryos, a second and fully rinsed spawning platform is introduced at 9:45 am rather than re-using the first spawning platform, which had been our initial and less successful approach. (5) To facilitate embryo collection, a larger-size petri dish was introduced. Collection of MEPS embryos according to this new regime started on 8/1/16, when the breeding population was 12 weeks old. Beginning with week 15 and continuing to week 17, benchmarks were largely being achieved. Specifically more than 1500 embryos were obtained on six out of the eight collection days and more than 500 synchronous embryos were obtained on 12 out of the 16 morning collections during this period. Future work will expand this regime and implement a re-population schedule to include both the large MEPS. The plan is to repopulate the MEPS biannually and in staggered fashion (i.e., one new generation of 3000 curated embryos per quarter), such that each MEPS breeder population is used from 3 to 9 months of age.
Automating fish size measurement.
An ever more common phenotypic characterization of genetic variants in zebrafish is the measurement of their sizes and weights and how these change over time. Key members of this class of genetic variants include zebrafish models for human diseases known to drive alterations in the size or weight of affected individuals, such as inborn errors of metabolism. Indeed, the use of the Core’s macro-photography lens and milligram balance to document various lines and models has risen strikingly since the Core’s creation in 2012. Recognizing this new trend and with financial assistance from the Stratakis lab, the Core contracted Viewpoint Life Sciences in August, 2014, to write software for their behavior-tracking systems designed to measure birds-eye view lengths and surface areas of free-swimming zebrafish embryos, larvae, juveniles, and adults. A promising software package was developed and installed on one of three behavioral tracking units owned by NHGRI and shared by NICHD (Figure 1). We found that up to five free-swimming juveniles or adults with a minimum length of about 1.2 cm can be reproducibly measured for snout-to-tail base length and birds-eye view surface area over the same section of their bodies. Future work will include longitudinal growth studies using R&D Aquatics’ DC-96 tanks, in which 24 zebrafish can be reared in isolated chambers. R&D Aquatics has allowed the Core to pre-test a new set of baffles designed to permit isolation of fish beginning at a younger (or smaller stage); we found that juveniles of 8–10 mm of length can be successfully isolated, corresponding to approximately four weeks post fertilization for wild-type fish. In future we will also work to develop holding systems that permit smaller and younger fish to be measured across all life stages and to develop software that leverages the two orthogonal cameras on Viewpoint Life Science’s 3D system to enable volumetric size measurements.
Service
Feldman joined the NICHD Animal Care and Use Committee (ACUC) and is participating in monthly meetings and other oversight activities.
Educational outreach
Feldman helped orchestrate 2016 ‘Take Your Child to Work Day’ events at the Central Aquatics Facility. He also acquired a highly portable NightSea system, which can convert a high-school level microscope into a fluorescent scope, and used this to assist a teacher and her students from Sidwell Friends School in Washington DC to visualize their own fluorescent zebrafish embryos.
Additional Funding
- NICHD Customers: $6,750 in fee-for-use charges
- Non-NICHD Customers: $8,200 in fee-for-use charges
- Bench-to-Bedside: $10,000 (second year of three)
Publications
- Gore A, Athans B, Iben J, Johnson K, Russanova V, Castranova D, Pham V, Butler M, Williams-Simons L, Nichols J, Bresciani E, Feldman B, Kimmel C, Liu P, Weinstein B. Epigenetic regulation of hematopoiesis by DNA methylation. eLife 2016;5:e11813.
- Trivellin G, Bjelobaba I, Daly AF, Larco DO, Palmeira L, Faucz FR, Thiry A, Leal LF, Rostomyan L, Quezado M, Schernthaner-Reiter MH, Janjic MM, Villa C, Wu TJ, Stojilkovic SS, Beckers A, Feldman B, Stratakis CA. Characterization of GPR101 transcript structure and expression patterns. J Mol Endocrinol 2016;57:97-111.
- Feldman B. Taking the middle road: vertebrate mesoderm formation and the blastula-gastrula transition. In: Moore S., ed. Principles of Developmental Genetics, 2nd Edition. Academic Press 2014;203-236.
- Horstick EJ, Jordan DC, Bergeron SA, Tabor KM, Serpe M, Feldman B, Burgess HA. Increased functional protein expression using nucleotide sequence features enriched in highly expressed genes in zebrafish. Nucleic Acids Res 2015;43(7):e48.
Collaborators
- Perry Blackshear, PhD, Signal Transduction Laboratory, NIEHS, Research Triangle Park, NC
- Harold Burgess, PhD, Section on Behavioral Neurogenetics, NICHD, Bethesda, MD
- Elena Casey, PhD, Georgetown University, Washington, DC
- Stephen Kaler, MD, Section on Translational Neuroscience, NICHD, Bethesda, MD
- Forbes D. Porter, MD, PhD, Section on Molecular Dysmorphology, NICHD, Bethesda, MD
- Thomas Sargent, PhD, Section on Vertebrate Development, NICHD, Bethesda, MD
- Constantine Stratakis, MD, D(med)Sci, Section on Endocrinology and Genetics, NICHD, Bethesda, MD
- Kandice Tanner, PhD, Laboratory of Cell Biology, NCI, Bethesda, MD
- Mendel Tuchman, MD, Children's National Medical Center, Washington, DC
- Brant Weinstein, PhD, Section on Vertebrate Organogenesis, NICHD, Bethesda, MD
Contact
For more information, email bfeldman@mail.nih.gov or visit http://zcore.nichd.nih.gov.
