This two-day symposium will showcase the best in developmental biology across the life course. From embryogenesis through to ageing, areas of current excitement in the field will be highlighted by plenary talks from 15 internationally renowned speakers, along with selected short talks from abstracts.
The symposium will honour the memory of Rosa Beddington, a leading UK embryologist who was Head of the Division of Mammalian Development at the MRC National Institute for Medical Research from 1993-2001. Many of Rosa’s colleagues and lab alumni will be attending the symposium.
We invite submissions for short talks (15 minutes) from early-career researchers (PhD students, postdocs and recently-established PIs). Please submit your abstracts to events@crick.ac.uk by 1 December 2018. Abstracts should be no more than 500 words including the title, authors and institution information. The presenter for short talks must have registered their attendance to the conference by 1 December 2018. Please ensure the presenter’s name is underlined on your submission.
We look forward to seeing you at the Crick in February,
The Organising Committee (James Briscoe, Alex Gould, Rita Sousa-Nunes and Jean-Paul Vincent)
Several MRC-funded positions are available in the Scholpp lab in the Living Systems Institute (LSI) at the University of Exeter to elucidate various aspects of cytoneme-mediated Wnt trafficking in vertebrates.
We are looking for
a Postdoc (3years, starting in Spring 2019), two PhD students (3.5years, starting in Autumn 2019, UK/EU only), and a Tech (2years, starting in Spring 2019)
Application deadlines: November / December 2018!
More information about the lab, about the LSI, and about our recent research in The Node.
Several postdoctoral positions are available in the group of Taija Mäkinen at Uppsala University. The lab studies fundamental mechanisms of tissue morphogenesis in the vascular system. The aim is to understand how endothelial cells communicate with each other and the tissue environment to co-ordinate vascular morphogenesis and formation of functionally specialised blood and lymphatic vessel types. To do so, the group utilises advanced mouse genetic tools and state-of-the-art cell and molecular biology techniques (including single cell RNA sequencing, flow cytometry, confocal, light-sheet and super-resolution microscopy). For more details about the group’s research please see: http://www.makinenlab.com/
Selected recent publications from the laboratory:
Zhang et al, Nat Commun 2018; Frye et al, Nat Commun 2018; Zhang et al, Development 2018; Wang et al, Development 2017; Martina-Almedina et al, J Clin Invest 2016; Martinez-Corral et al, Circ Res 2015; Stanczuk et al, Cell Rep 2015; Tatin et al, Dev Cell 2013; Lutter et al, J Cell Biol 2012; Bazigou et al, J Clin Invest 2011.
Work description:
The selected candidate(s) will work on one of the following topics: 1) functional characterisation of tissue-specific lymphatic endothelial progenitor cells (as part of an ERC-funded project), 2) identification and functional characterisation of vascular-bed specific genes, or 3) elucidation of disease mechanisms in vascular malformations. In addition, one position will be in collaboration with the group of Ingvar Ferby at Uppsala University (http://www.imbim.uu.se/forskargrupper/cancer/ferby-ingvar/), exploring how vesicular trafficking and compartmentalisation of growth factor receptors instruct behaviour of epithelial and endothelial cells using e.g. live-cell imaging approaches.
Qualifications:
We are looking for highly motivated individuals with a PhD and research background in molecular or cell biology, developmental biology or biochemistry, and a proven track record of successful scientific work. Strong background in molecular/cell biology, mouse genetics, flow cytometry and/or imaging is required.
How to apply:
To apply, please send your CV together with the names of three references and a short description of yourself and the motivation to join the group to: taija.makinen@igp.uu.se
The position is open until 28 December 2018, or until suitable candidate(s) is found.
Are you interested in applying mathematics, statistics or deep learning/machine learning to biomedical problems? Apply now for a MRC WIMM Prize Studentship, to start in October 2019. The studentship is fully-funded for four years, including a stipend of £18,000 p.a. and all University and College fees paid.
Applicants with a background in Physics, Mathematics, Engineering, Statistics or Computer Science are encouraged to apply. To be eligible for a full award, applicants must have no restrictions on how long they can stay in the UK and must have been ordinarily resident in the UK for at least 3 years prior to the start of the studentship. Further details about residence requirements may be obtained here.
For further information on how to apply can be found here.
All applications must be received by 12 noon (UK time) on Friday, 11 January 2019
Interviews will take place the week commencing 28 January 2019.
The Zebrafish Interest Group at the University of Utah held its first Utah Fish Conference (UFC) on October 8, 2018. The conference was organized by pre- and post-doctoral trainees to celebrate the University’s Zebrafish Interest Group (ZIG), as well as to unite the Mountain West fish community. This 1-day event hosted over 80 attendees from 6 institutes. UFC was sponsored by the University’s ZIG, as well as Tecniplast USA, Aquatic Enterprises, IDT, Aquaneering, wFluidx, NCI, NEB, Zeiss, ThermoFisher by Life Technologies, and Developmental Dynamics.
UFC was held at the Crocker Science Center at the University of Utah. The talks were well-attended by an active audience.
The event featured two excellent keynotes, Trista E. North at Boston Children’s Hospital, and Bruce Draper at University of California-Davis. Dr. North jump-started the day’s event with a stimulating talk on the hematopoietic signaling connectome. She was followed by two hour-long sessions of trainee talks, which represented labs across the University and trainees from outside institutes. There was an active poster session comprising 30 posters from 16 labs, with presenters from all career stages, from undergraduate to faculty researchers.
Members of the Organizing Committee (Left to Right: Chelsea Herdman, Alexis Fulbright, and Penny Lam) caught chatting with Dr. North during a coffee break.
An “Ask Me Anything” panel followed the poster session and featured both senior and junior faculty from the University, as well as Dr. North. A pre-doctoral trainee, Deeptha Vasudevan, moderated questions from the audience, and the panel covered topics from how to settle on a career path to advice on how to start your lab. The AMA was followed by an exciting evening talk from Dr. Bruce Draper about sex determination.
The poster session featured 30 posters from a diverse group of labs.
During dinner, the awards for best talk and best posters were announced. Robert Mackin (U of Idaho) won the Outstanding Young Investigator award for his excellent talk. Poster awards were given to 2 trainees in each category: undergraduate, graduate, or post-doctoral trainee. Awards went to: Jeffrey Dunn (BYU) and Samuel Caton (BYU) in the undergraduate category; Dana Klatt Shaw (U of Utah) and Srishti Kotiyal (U of Utah) in the graduate category; and Chelsea Herdman (U of Utah) and Angie Serrano (U of Utah) in the post-doctoral category. Awards were cash prizes sponsored by Developmental Dynamics. Following dinner, an after-party was hosted at The Porcupine Pub, sponsored by ThermoFisher by Life Technologies. The conversation was lively and offered more opportunities for trainees to intermix and mingle with faculty, Dr. North, and Dr. Draper.
The UFC was an invigorating moment for the University’s ZIG community, as well as for the Mountain West fish community. It was an excellent way to highlight the working happening within the region, and unite different institutes.
UFC2018 Organizing Committee:
Gabriel Bossé, PhD, @GabrielBosse1 (Randall Peterson Lab)
Join us for the Conference on Cell Competition in Development & Disease, in Tahoe, California!
February 24–28, 2019
Granlibakken Tahoe – Lake Tahoe, California, USA
Cell competition is a highly conserved process that promotes the context-dependent elimination of less fit cells and stimulates growth of more fit cells during growth and homeostasis. It has long been known that the basis of competition is the ability of growing cells to monitor their fitness and that of their neighbors, but only recently have signaling and effector mechanisms been identified. New technologies have uncovered the prevalence of cell competition in humans, with surprising outcomes and implications for human disease. This conference aims to bring together, for the first time, researchers from diverse fields who study competitive and cooperative interactions between cells.
Plenary Session Topics:
• Evolution of Competition and Cooperation • Stem Cell Competition • The Germline • Mosaicism and Selection in Normal Tissues • Cell Selection in Human Disease • Aging and Pre-Malignancy • Competition in Cancer
Plus workshops on technologies for clonal tracking, oncogenes and tumor suppressors as drivers of competition, and computational modeling of cell competition
Scientific Organizers: Margaret A. Goodell, Baylor College of Medicine, USA Laura A. Johnston, Columbia University, USA Thomas P. Zwaka, Icahn School of Medicine at Mount Sinai, USA
Scholarship/Discounted Abstract Deadline: Oct 24, 2018; Abstract Deadline: Nov 28, 2018; Discounted Registration Deadline: Jan 8, 2019
A postdoctoral position is available in the Developmental Mechanobiology and Regeneration lab of Joel Boerckel at the University of Pennsylvania in Philadelphia, PA, USA (http://www.med.upenn.edu/orl/boerckellab/). Our laboratory is housed in the Departments of Orthopaedic Surgery and Bioengineering and studies how mechanical cues direct morphogenesis, repair, and regeneration. A major focus of the lab is understanding how the transcriptional co-activators YAP and TAZ mediate progenitor cell mechanosensation, motility, and differentiation during development and regeneration.
We are looking for postdoctoral fellows with backgrounds in either cell and developmental biology or bioengineering to join our multidisciplinary team. NIH-funded projects are available to define the roles of YAP and TAZ in osteoprogenitor cell mobilization, mechanical regulation of endochondral ossification, and development-mimetic tissue engineering. Projects feature a combination of cell and bioreactor culture, biomaterials, and animal modeling using transgenic mice and rats.
What we offer:
Highly collaborative and collegial environment in the McKay Orthopaedic Research Labs (http://www.med.upenn.edu/orl)
Regular interactions with world-class colleagues and visiting speakers through the Penn Center for Musculoskeletal Diseases (https://www.med.upenn.edu/pcmd) and the Center for Engineering Mechanobiology (https://cemb.upenn.edu/) at UPenn.
Supportive mentorship for multi-faceted career development and opportunities tailored towards individual career goals.
A stimulating environment with freedom to develop new research directions.
An NIH funded position at NRSA postdoctoral stipend levels (with potential for renewal up to four additional years).
A department located in a dynamic East-coast city with affordable cost of living.
What we’re looking for:
Enthusiastic and ambitious individuals with a strong interest in our research and a collaborative and collegial laboratory environment.
Fearlessness in learning new techniques and designing projects independently.
Willingness to apply for applicable postdoctoral fellowships and eagerness to take advantage of other career development opportunities.
Interest in working with junior lab members and summer undergraduates.
Strong verbal and written communication skills.
Start date immediately or upon mutual agreement.
Application materials (email to boerckel@pennmedicine.upenn.edu):
Cover letter outlining relevant expertise and scientific interests
A Postdoctoral position is available in the Charalambous lab in the Department of Medical and Molecular Genetics at King’s College, London. Our team investigates genetic and epigenetic determinants of mammalian body composition. The candidate will be part of an exciting collaboration with the University of Bath, investigating the role of imprinted genes in adipose tissue development and turnover.
There is considerable phenotypic variation of body composition in the human population. This is clinically important because while some people maintain their health when exposed to a ‘Western’ lifestyle others develop obesity and associated metabolic disorders such as Type II Diabetes and cardiovascular disease. There is strong evidence that a large component of the variation in body composition is genetic, and furthermore – many of the genes may be acting in developmental pathways to modify skeletal muscle mass and adipose plasticity for a lifetime. By using in-vivo models to manipulate lean:adipose distribution we hope to identify the genetically determined developmental pathways that determine body composition, and understand how their compromise predisposes to metabolic disease.
The candidate will be part of a team based at Guy’s Campus, King’s College London, embedded within a centre of excellence for developmental biology and stem cell research. Moreover, the Department of Medical and Molecular Genetics has recently recruited a critical mass of researchers in the field of epigenetics, providing a strong source of crossover opportunities with this project.
Qualifications:
Applicants should have a recent Ph.D. degree or M.D./Ph.D. degree. Candidates with experience in stem cell and/or developmental biology, confocal microscopy, image acquisition and analysis will be preferred, as will those with bioinformatics skills. This position seeks a highly motivated individual with a strong interest in developmental biology.
This Editorial by Sergey Prykhozhij andJason Berman originally appeared in Disease Models and Mechanisms, an online Open Access sister journal to Development focusing on the use of model systems to better understand, diagnose and treat human disease. The Editorial focuses on three new papers on point mutant knock ins in zebrafish, and will thus be of interest to many developmental biologists.
The zebrafish is an increasingly popular model organism for human genetic disease research. CRISPR/Cas9-based approaches are currently used for multiple gene-editing purposes in zebrafish, but few studies have developed reliable ways to introduce precise mutations. Point mutation knock-in using CRISPR/Cas9 and single-stranded oligodeoxynucleotides (ssODNs) is currently the most promising technology for this purpose. Despite some progress in applying this technique to zebrafish, there is still a great need for improvements in terms of its efficiency, optimal design of sgRNA and ssODNs and broader applicability. The papers discussed in this Editorial provide excellent case studies on identifying problems inherent in the mutation knock-in technique, quantifying these issues and proposing strategies to overcome them. These reports also illustrate how the procedures for introducing specific mutations can be straightforward, such that ssODNs with only the target mutation are sufficient for generating the intended knock-in animals. Two of the studies also develop interesting point mutant knock-in models for cardiac diseases, validating the translational relevance of generating knock-in mutations and opening the door to many possibilities for their further study.
Introduction
One of the great potentials of zebrafish (Danio rerio) is to generate accurate models of human genetic diseases to recapitulate their clinical features, to understand the molecular mechanisms that underpin them, and to model treatments and disease management approaches. Before the advent of precise genome editing, understanding a missense mutation within a protein-coding human gene in zebrafish necessitated designing an mRNA expression vector or a transgenic construct. These approaches did not actually replace the normal (wild-type) zebrafish gene with an altered human one. Additionally, expression of a human (trans)gene raises the question whether that gene is, in fact, functional in fish. Does the mutant gene function in fish? Are expression levels sufficient? Is expression in the right place and at the right time? These answers may be elusive.
Imagine instead the ability to map an amino acid in a protein from another species to a specific zebrafish protein residue and then mutate it. This is the essence of point mutation knock-in; namely, the replacement of wild-type nucleotides with mutant ones by inducing endogenous recombination with genome-editing reagents. An investigator can then confidently know that his or her gene of interest is modified precisely and expressed in the biologically relevant way – under the control of the endogenous promoter. With this refined approach, the mutation is the only variable under study. One potential disadvantage of current knock-in methodologies is that they cannot be used directly for introducing dominant and lethal mutations, which might be better studied using transgenic approaches that afford inducibility.
Generation of precise point mutations in zebrafish for basic research and disease modeling studies is an important contribution of the burgeoning CRISPR/Cas9 technology, which uses a single-guide RNA (sgRNA) molecule to guide the Cas9 endonuclease to the genomic site of choice. Single-stranded oligodeoxynucleotides (ssODNs), or oligos for short (as referred to herein), are currently the donor templates of choice for mutation knock-ins (Fig. 1A). These oligos can vary in length and be either ‘target’ (T) or ‘non-target’ (NT) in orientation, and have either symmetric or asymmetric homology arms. NT oligos correspond to the strand not bound by the sgRNA, which contains the protospacer-adjacent motif (PAM) sequence; hence, the oligos are sometimes referred to as ‘sense’. Conversely, T oligos are derived from the strand bound by the sgRNAs and are correspondingly often referred to as ‘anti-sense’.
Fig. 1. Comparison of different study results using oligos for point mutation knock-ins. The figure panels show the Cas9-sgRNA CRISPR complex cutting genomic DNA and subsequent homology-directed repair by resection and knock-in mutation insertion. Synthesis-dependent strand-annealing (SDSA) is the DNA repair process involved in generating knock-ins when an ssODN (oligo) is present. (A) The basic strategy of point mutation knock-ins. The first step includes the identification of a functional sgRNA to couple with the Cas9 nuclease and direct it to the genomic site of choice. Second, the donor oligo with the mutation of interest and mutation(s) in sgRNA site or PAM is designed. Mutating the sgRNA homology site and/or the PAM site prevents subsequent rounds of Cas9-induced cuts of the edited genomic site. Third, upon the Cas9-induced break in genomic DNA, homology-dependent repair using the provided oligo can occur and the mutation is inserted into the genome. (B) The results of studies employing a comparison of ‘NT 126 S’ (sense symmetric) and ‘T 126 A right’ (anti-sense asymmetric) oligo knock-in efficiencies in zebrafish and in vitro (Prykhozhij et al., 2018b; Richardson et al., 2016). (C) The results of a cell culture study demonstrating which types of asymmetric oligos are more efficient (Liang et al., 2016). (D) The results of the study in this issue (Boel et al., 2018) that shows that symmetric oligos such as ‘NT 120 S’ perform nearly as well as two of the asymmetric oligos (‘NT 120 A left’ and ‘T 120 A right’), which, in turn, perform much better than their counterparts ‘NT 120 A right’ and ‘T 120 A left’.
Our own group previously reviewed zebrafish studies aimed at developing precise point mutation techniques using CRISPR/Cas9 (Prykhozhij et al., 2018a), including the first successful zebrafish study on germline transmission of a point mutation knock-in (Armstrong et al., 2016). In this Editorial, we will explore the implications of three papers published in this issue of Disease Models & Mechanisms that demonstrate point mutation knock-ins in zebrafish, and the advantages and challenges of working with such precise mutants (Boel et al., 2018; Tessadori et al., 2018; Farr et al., 2018). These papers pave the way toward CRISPR-based oligo knock-ins becoming widely acceptable as the preferred approach to generate point mutations in zebrafish.
The focus on technology
The paper by Boel et al. focuses on the technical aspects of knock-in optimization (Boel et al., 2018). The first key observation in the Boel paper is that using oligos to guide homology-directed repair (HDR) is quite error prone. Although this has been previously shown in zebrafish (Burg et al., 2016; Hruscha et al., 2013; Hwang et al., 2013; Prykhozhij et al., 2018b), this report does so much more systematically. The authors identified typical ‘knock-in with indel’ events, insertions of partial and multiple oligos, inverse oligo insertions and other abnormal events. The prevalence of these abnormal events ranged from a few percent to >50% of the total knock-in events, thereby reducing the correct knock-in rates from 4-8% (total knock-ins) to 1-4% (correct). This result suggests that caution and routine use of next-generation sequencing (NGS) approaches are needed to assess the error rate of any new knock-in strategy. Given that each oligo in this study contained six nucleotide changes, the authors examined how much the distance between the Cas9-induced cut site and mutation influences the knock-in efficiency. Predictably, increasing this distance resulted in decreased knock-in efficiency, which was more pronounced for oligos with shorter homology arms, such as 60 nucleotides (nt), and for asymmetric (A) oligos, in which, for example, the left homology arm is 30 nt and the right is 90 nt long. This is consistent with previous studies (Paquet et al., 2016), and suggests that researchers use longer or symmetric (S) homology arms for engineering knock-ins in which the mutation site is at large distances from the Cas9-induced cut sites.
Boel and colleagues also investigated whether asymmetric homology arm design for 120 nt oligos can improve knock-in efficiency (Boel et al., 2018). No single type of oligo design performed best in their system, so they concluded that homology arm asymmetry is likely not a general strategy for improving knock-in rates. This somewhat contradicts previous studies in cell culture systems that found target asymmetric ‘T 126 A right’ oligos superior to non-target symmetric ‘NT 126 S’ oligos (Fig. 1B) (Richardson et al., 2016), and that ‘NT 97 A left’ or ‘T 97 A right’ were both superior to the symmetric ones (Fig. 1C) (Liang et al., 2017).
In zebrafish, our group recently confirmed the in vitro findings of Richardson et al. (2016) that target asymmetric oligos perform significantly better than the non-target symmetric oligos for two different knock-ins into the tp53 gene (Fig. 1B) (Prykhozhij et al., 2018b). Boel and colleagues also note, however, that for three out of the four sites in the zebrafish genome they targeted, the combined knock-in rates for 120 nt asymmetric (‘NT 120 A left’ and ‘T 120 A right’) oligos were higher than those of symmetric ones. These 120 nt asymmetric oligos also performed consistently better than the ones with opposite asymmetries (‘NT 120 A right’ and ‘T 120 A left’) (Fig. 1D), which is consistent with the known properties of homologous recombination-based DNA repair by resection and synthesis-dependent strand annealing (Paix et al., 2017). The authors make a reasonable recommendation to test these oligos in parallel with symmetric ones. Boel et al.’s main aim was technical knock-in optimization. The oligos they tested contained six nucleotide mismatches and were not intended to change the amino acid sequence of the protein product (Boel et al., 2018). The oligos used in our study contained non-silent disease-relevant mutations, which is likely a more common scenario for mutation knock-in studies. Targeted cells might have different tolerance for silent and non-silent mutations, which can result in different frequencies of successfully targeted clones. Future efforts will either resolve these efficiency discrepancies or perhaps render them irrelevant due to further improvement of genome-editing technologies.
Modeling diseases via knock-ins with short oligos
Large-scale sequencing technology has greatly facilitated the identification of novel potentially disease-causing genomic variants. We have yet to understand the functional implications of these variants, but zebrafish can assist us in clarifying the complex genetic and molecular underpinning of these variants. The zebrafish has become one of the most sought after organisms in which to generate mutants that can serve as models of diseases caused by specific genomic variants. Most of these variants cannot be accurately modeled by complete gene inactivation or knockout and require knock-in approaches to introduce a specific mutation. Despite recent technological advances, few disease-associated zebrafish point mutants have been generated to date. The papers by Farr et al. and Tessadori et al. in this issue of Disease Models & Mechanisms describe point mutation knock-ins of several genetic variants implicated in inherited cardiac diseases (Farr et al., 2018; Tessadori et al., 2018). Tessadori et al. focus on mutations found in Cantú syndrome, a rare disease characterized by multiple cardiac abnormalities, bony changes and hair thickening. Farr et al. describe a zebrafish model of a PBX3 A136V mutation, which is enriched in a subset of congenital heart defects.
Both studies employed in vitro-transcribed sgRNA and either nCas9n mRNA (Jao et al., 2013) or Cas9 protein (Tessadori et al., 2018), together with oligos encoding the modifications. The distances between the Cas9 cut site and the nucleotide(s) to be mutated were 0-5 nt in the paper by Tessadori et al. and 10 nt in the paper by Farr et al. Both distances were in the previously reported optimal range (Paquet et al., 2016). Importantly, the mutations overlapped the sgRNA binding sites. Tessadori et al. used non-target strand mutant oligos with 25 nt homology arms and Farr et al. tested both non-target and target oligos, which were asymmetric (35 nt and 15 nt homology arms) if counted from the cut site. Interestingly, Farr and colleagues found that oligos with 25 nt homology arms worked well, whereas oligos with 20 nt and 40 nt homology arms failed. It is not clear whether homology arm asymmetry accounts for the performance differences, but it is likely that the shorter 10 nt homology arms are generally too short and 50 nt homology arms oligos might be too long for HDR. Once knock-in reagents were injected and the fish were subsequently bred, the resulting knock-in alleles needed to be detected and verified. Both papers employed Sanger sequencing and restriction enzyme digestion of the targeted site to evaluate knock-ins. Alternative approaches, such as allele-specific PCR, could be a more broadly applicable option (Prykhozhij et al., 2018b).
Both groups leveraged cut site proximity to the mutation sites and undertook some oligonucleotide size optimization to achieve efficient knock-in generation and germline transmission. The approaches described in these papers might prove effective for other mutation knock-ins in which the desired change is proximal to the Cas9 cut site. However, longer homologies and asymmetric homology arm designs might be needed for targeting mutation sites that are more distant from the predicted Cas9 cut site. Boel et al. (2018) suggest that much of the increase in efficiency observed due to lengthening oligos from 60 nt to 120 nt comes from aberrant repair events. It is therefore possible that 50-60 nt is a ‘sweet spot’ oligo length for cut-site proximal target mutation. The Farr and Tessadori groups should be lauded for identifying the minimal effective oligo size and simple knock-in procedures.
The main aim of these two studies was to model heart disease in zebrafish. Therefore, the mutations the authors introduced needed to produce a tractable, disease-relevant phenotype. Farr et al. aimed to test whether the zebrafish pbx4 A131V variant (which is homologous to the human PBX3 A136V mutation) could function as a modifier allele, resulting in a congenital heart defect. The pbx4 A131V allele did not produce a clearly discernable phenotype, but it was also not completely functional because its presence increased the severity of heart defects when combined with a null pbx4 allele (Farr et al., 2018). Tessadori and colleagues identified a convincing phenotype for their heterozygous and homozygous kcnj8V65M Cantú syndrome mutants. Their abcc9 G989E knock-in had a phenotype similar to that of the kcnj8V65M knock-in mutants, confirming a more generalized genotype-phenotype correlation (Tessadori et al., 2018). These newly developed zebrafish models could be used to improve our understanding of genetic heart disease and to test therapeutic approaches.
Conclusions
In sum, these three papers highlight various technical optimizations that can achieve robust and reproducible knock-in mutations, leading to zebrafish models with greater fidelity to the human diseases they are modeling. Lessons from these papers will be instructive to other investigators by providing important factors to consider when designing CRISPR/Cas9-based knock-ins in zebrafish. Although the generation of knock-in mutations continues to pose challenges, its successful implementation promises to be of tremendous value to the broader model organism community to study complex genetics that contribute to disease, in genes and in non-coding regions of the genome. By incorporating these mutants into high-throughput drug screening pipelines, the zebrafish holds great potential to provide rapid, cost-effective preclinical therapeutic data in a uniquely whole-organism vertebrate context that can streamline confirmatory murine studies and ultimately inform future clinical trials for patients with genetic disorders.
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