This protocol describes an approach to facilitate precise knock-in edits in zebrafish embryos using CRISPR-Cas9 technology. A phenotyping pipeline is presented to demonstrate the applicability of these techniques to model a Long QT Syndrome-associated gene variant.
Clustered regularly interspaced short palindromic repeats (CRISPR) in animal models enable precise genetic manipulation for the study of physiological phenomena. Zebrafish have been used as an effective genetic model to study numerous questions related to heritable disease, development, and toxicology at the whole-organ and -organism level. Due to the well-annotated and mapped zebrafish genome, numerous tools for gene editing have been developed. However, the efficacy of generating and ease of detecting precise knock-in edits using CRISPR is a limiting factor. Described here is a CRISPR-Cas9-based knock-in approach with the simple detection of precise edits in a gene responsible for cardiac repolarization and associated with the electrical disorder, Long QT Syndrome (LQTS). This two-single-guide RNA (sgRNA) approach excises and replaces the target sequence and links a genetically encoded reporter gene. The utility of this approach is demonstrated by describing non-invasive phenotypic measurements of cardiac electrical function in wild-type and gene-edited zebrafish larvae. This approach enables the efficient study of disease-associated variants in a whole organism. Furthermore, this strategy offers possibilities for the insertion of exogenous sequences of choice, such as reporter genes, orthologs, or gene editors.
CRISPR-based gene editing strategies in animal models enable the study of genetically heritable disease, development, and toxicology at the whole-organism level1,2,3. Zebrafish provide a powerful model that is closer in numerous physiological aspects to humans than murine or human-derived cell models4. An extensive array of genetic tools and strategies have been used in zebrafish for both forward5 and reverse genetic screening6. Comprehensive genetic mapping and annotation in zebrafish have facilitated gene-editing approaches as a primary technique to engineer targeted gene knockouts (KOs) and precise knock-ins (KIs)7.
Despite this, generating precise KI edits in zebrafish is limited by low efficiencies and the difficulty of accurate detection. Although transcription factor-like effector nucleases (TALENs) have been successfully used and optimized for KIs8, CRISPR provides an improved gene-editing strategy with simpler sgRNA targeting. Numerous studies have used CRISPR to generate precise KIs in zebrafish9,10,11,12,13,14,15,16,17,18,19,20, although these edits generated through CRISPR-mediated homology-directed repair (HDR) tend to be inefficient with low intrinsic success rates that require genotyping as a primary screen9,10,14,21. This demonstrates the need for an efficient KI CRISPR system in zebrafish, as well as a reliable high-throughput system for detecting precise edits.
The goal of this study was to describe a platform for generating a precise cardiac gene KI in zebrafish hearts with simple and high-throughput detection of successful edits. A CRISPR-Cas9-based two-sgRNA exon replacement approach is described, which is based on a TALEN approach8. This approach involves excision of the target sequence using two-sgRNA guides and replacement with an exogenous template sequence that contains the KI of interest as well as a genetically encoded intronic reporter gene (Figure 1). The integration of a genetically encoded fluorescent reporter within the target gene intronic sequence enables the efficient detection of positive edits. A phenotyping platform is then described for assessing cardiac electrical function in zebrafish larvae for non-invasive characterization of the gene variants associated with inherited LQTS, a cardiac electrical disorder that predisposes individuals to sudden cardiac death.
These approaches will enhance the access to and use of zebrafish KI gene edits to model inherited diseases and address biological and physiological questions, such as mapping gene expression patterns, and developmental regulation. Since zebrafish hearts better parallel human cardiac electrophysiological characteristics than murine models, they may be particularly attractive as a genetically tractable system for cardiac disease modeling7,22,23.
Studies using zebrafish were conducted in agreement with the policies and procedures of the Simon Fraser University Animal Care Committee and the Canadian Council of Animal Care and were completed under protocol # 1264K-18.
1. Design of CRISPR components for precise edits
2. Preparation of CRISPR components for embryo microinjection
3. Breeding of zebrafish and embryo microinjection
NOTE: Protocols for zebrafish breeding and the microinjection of single-cell embryos have been described previously29,30,31.
4. Reporter gene screening of CRISPR-Cas9-edited larval zebrafish
5. Phenotyping of CRISPR-Cas9-edited larval zebrafish
6. Genotyping of CRISPR-Cas9-edited larval zebrafish
The successful use of this two-sgRNA exon replacement CRISPR approach is highlighted by the introduction and simple detection of a precise edit to engineer the LQTS-associated variant, R56Q, in the zkcnh6a gene in zebrafish. Figure 6 shows a representative 3 dpf larvae injected at the one-cell embryo stage with CRISPR components as described above. Figure 6A shows the presence of the YFP mVenus reporter gene expression in the eye lens as a positive reporter of successful template integration. Figure 6B,C show Sanger sequencing chromatograms obtained from genomic DNA isolated from tail clip samples of wild-type and reporter gene-positive fish, respectively. Reporter gene-positive fish were found to have the precise edit, G to A, which introduces the R56Q variant into zkcnh6a. Genotyping showed a 100% correlation between YFP reporter gene expression and the presence of the precise R56Q gene edit, validating this fluorescence screening tool.
Phenotyping of gene-edited zebrafish larvae was conducted at 3 dpf. Figure 7 shows representative results from wild-type and R56Q gene-edited larvae. Heart rate was detected by video capture as described above. An example of the measurement of pericardial dimensions as a ratio of eye area is shown (Figure 7A). Figure 7B plots heart rate against normalized pericardial dimensions, highlighting a trend of bradycardia with increasing pericardial edema, which is associated with disorders of cardiac repolarization in zebrafish8,38,39,40. Figure 7C shows a representative example of ECG recordings from 3 dpf larvae. Standard intervals (QT, QRS) were measured from averaged ECG signals.
Figure 1: Integration of HDR template into the zebrafish genome. Dark grey, homology arms; green, sgRNA guide targets with silent mutation to prevent Cas9 recutting; light grey, target exon of interest; red line, point mutation; yellow, mVenus YFP reporter gene under an α-crystallin promoter; dashed lines indicate homology. Here, the targeted precise edit was R56Q in exon 2 of the zkcnh6a gene. Abbreviations: HDR = homology-directed repair; sgRNA = single-guide RNA; YFP = yellow fluorescent protein; DSB = double-stranded break; WT = wild type. Please click here to view a larger version of this figure.
Figure 2: Summary of steps to engineer precise edits in zebrafish genes using the two-sgRNA CRISPR-Cas9 approach (related protocol step numbers are indicated in parentheses). Abbreviations: sgRNA = single-guide RNA; YFP = yellow fluorescent protein; gDNA = genomic DNA; ECG = electrocardiogram; dpf = days post fertilization; MS-222 = tricaine methane sulfonate. Please click here to view a larger version of this figure.
Figure 3: Preparation of exogenous template fragments and sgRNA guides. (A) Sequential digestion and ligation of template fragments upstream and downstream of the mVenus YFP reporter gene sequence in pKHR5. (B) Annealing of complementary sgRNA pairs with restriction overhang for ligation into DR274. Abbreviations: sgRNA = single-guide RNA; YFP = yellow fluorescent protein. Please click here to view a larger version of this figure.
Figure 4: Construction of HDR template. Dark grey, homology arms; green, sgRNA guide targets with silent mutation to prevent Cas9 recutting; light grey, target exon of interest; red line, point mutation; yellow, mVenus YFP reporter gene under an α-crystallin promoter; dark blue line, added restriction sites. The two template fragments are integrated into the pKHR5 plasmid donor. Abbreviations: HDR = homology-directed repair; sgRNA = single-guide RNA. Please click here to view a larger version of this figure.
Figure 5: Microinjection of single-cell zebrafish embryos with CRISPR-Cas9 components. Scale bar = 0.5 mm. Abbreviations: HDR = homology-directed repair; sgRNA = single-guide RNA. Please click here to view a larger version of this figure.
Figure 6: Easy detection of mVenus YFP reporter gene fluorescence indicates positive HDR exogenous template integration into the target gene. (A) Example of mVenus YFP expression in a zebrafish eye (arrow) in an edited zebrafish larva. (B) Successful edits are confirmed by sequencing chromatograms (left, WT; right, R56Q edit). Please click here to view a larger version of this figure.
Figure 7: Phenotypic analysis of cardiac consequences in 3 dpf zebrafish following the precise R56Q edit in the zkcnh6a target gene. (A) Image detection of pericardial dimensions relative to eye size using the polygon tool in ImageJ. The boundaries of the pericardial sac were marked by the user from a single recording frame based on changes in translucency and pigmentation. Examples of normal pericardial dimensions and pericardial effusion are shown. Scale bar = 0.5 mm. (B) Correlation between pericardial dimensions (relative to eye dimension) and heart rate, R2 = 0.33. (C) Example of ECG recording from a 3 dpf zebrafish larva heart (left) and averaged complexes (right). Heart rate, 131 bpm; heart rate-corrected QTc interval, 460 ms. Abbreviations: dpf = days post fertilization; ECG = electrocardiogram. Please click here to view a larger version of this figure.
The engineering of precise gene edits using CRISPR-Cas9 is challenged by the low efficiencies of HDR mechanisms and their efficient detection. Here, a CRISPR-Cas9-based two-sgRNA exon replacement approach is described that produces precise edits in zebrafish with straightforward visual detection of positive edits. The efficacy of this approach is demonstrated by generating precise edits in the zkcnh6a gene. This paper shows how cardiac function in gene-edited zebrafish larvae may be assessed using non-invasive phenotypic measures of heart rate, pericardial dimensions, and ECG morphology. This approach, from introducing a gene edit to phenotypic evaluation, can be completed from start to finish within approximately 1 week.
The benefits of the above editing and phenotyping approach are the ease of CRISPR modification design, the wide applicability in multiple physiological systems, the ability to insert large genes or gene fragments, and the ability to track variant effects longitudinally through development and generations. The success of precise edits in this approach may be related to the combination of the large template size (due to the reporter gene insert and long homology arms), which has been shown to increase the efficiency of edits in zebrafish14, and the two-sgRNA-guides strategy, which has been used effectively in zebrafish TALEN-induced edits8.
One particular strength of the described approach is the ability to insert large genes or gene fragments. This may be useful, for example, to insert human orthologs41, allowing for more clinically translatable characterization and comparison between orthologs. Alternatively, genes encoding Cas enzymes could also be inserted, allowing for a line of zebrafish with in vivo CRISPR editing mechanisms, providing an inducible system. Similarly, alternative CRISPR mechanisms, such as prime editing, could be integrated and result in a line of zebrafish that are readily edited precisely and efficiently.
Despite the advantages of this approach, there are some limitations. First, only a single gene and locus have been modified, and further testing at other sites or in other genes is necessary to evaluate how broadly applicable this approach is. Due to the long homology arms required, the template design costs are higher; however, this may be offset by efficient screening. Another limitation is that the screening approach requires fluorescence detection capability. However, optical requirements are relatively low and can be custom-built or commercially purchased at a reasonably low cost. Using a two-sgRNA approach increases the number of potential off-target events; however, this is likely mitigated by the lower probability that the two sgRNA guides will both anneal in a manner that facilitates the incorporation of the template to yield reporter gene expression. Finally, using Cas9 mRNA may lead to mosaicisms as the Cas9 is not active until later developmental stages. This could be accounted for by sequencing particular tissue types; however, given the size of the zebrafish larvae, this is technically challenging.
In summary, this CRISPR-Cas9 two-sgRNA precise editing approach in zebrafish enables the simple visual detection of positive edits and may be adapted to incorporate large genes of interest at any locus. Combined with phenotypic measures, this allows for a reliable and high throughput platform for studying clinically-relevant cardiac variants.
The authors have nothing to disclose.
This research was supported by a Canadian Institutes of Health Research Project grant (T.W.C.) and Natural Sciences and Engineering Research Council of Canada Discovery grants (T.W.C.).
Program | |||
CRISPOR | TEFOR Infrastructure | ||
ENSEMBL | European Bioinformatics Institute | ||
ImageJ | National Institutes of Health (NIH) | ||
Micro-Manager | Open Source (Github) | ||
NEBiocalculator | New England Biolabs (NEB) | ||
EQUIPMENT | |||
24-well Plate | VWR | ||
25 mm Petri Dish | VWR | ||
Blackfly USB3 Camera | Teledyne FLIR | ||
C1000 Thermal Cycler | Bio-Rad | ||
Centrifuge 5415C | Eppendorf | ||
EZNA Gel Extraction Kit | Omega Biotek | ||
MAXIscript T7 Transcription Kit | Invitrogen | ||
MaxQ 5000 Incubator | Barnstead Lab Line | ||
Miniprep Kit | Qiagen | ||
mMessage mMachine T7 Ultra Transcription Kit | Invitrogen | ||
ND1000 Spectrophotometer | Nanodrop | ||
PCR Purification Kit | Qiagen | ||
PLI 100A Picoinjector | Harvard Apparatus | ||
PowerPac Basic Power Supply | Bio-Rad | ||
Stemi 305 Steroscope | Zeiss | ||
Wide Mini Sub Cell GT Electrophoresis System | Bio-Rad | ||
ZebTec Zebrafish Housing System | Tecniplast | ||
SERVICES | |||
Gene Synthesis | Genewiz | ||
Sanger Sequencing | Genewiz | ||
REAGENTS | |||
10β Competent Cells | NEB | ||
10X PCR Buffer | Qiagen | ||
100 mM Nucleotide Mixture | ABM | ||
Ampicillin | Sigma | ||
BamHI Endonuclease w/ buffer | NEB | ||
BsaI Endonuclease w/ buffer | NEB | ||
DR274 Plasmid (XL1 Blue bacterial agar stab) | Addgene | ||
EcoRI Endonuclease w/ buffer | NEB | ||
Glycerol | |||
HEPES | Sigma | ||
HindIII Endonuclease w/ buffer | NEB | ||
Kanamycin | Sigma | ||
Methylene Blue | Sigma | ||
MLM3613 Plasmid (XL1 Blue bacterial agar stab) | Addgene | ||
MS-222 (Tricaine) | Sigma | ||
pKHR5 Plasmid (DH5α bacterial agar stab) | Addgene | ||
PmeI Endonuclease w/ buffer | NEB | ||
SalI Endonuclease w/ buffer | NEB | ||
Sodium Hydroxide | Sigma | ||
T4 Ligase w/ buffer | Sigma | ||
Taq Polymerase | Qiagen | ||
TE Buffer | Sigma | ||
Tris Hydrochloride | Sigma | ||
XhoI Endonuclease w/ buffer | NEB | ||
RECIPES | |||
Solution | Component | Supplier | |
Annealing Buffer (pH 7.5-8.0) | 10 mM Tris | Sigma | |
50 mM NaCl | Sigma | ||
1 mM EDTA | Sigma | ||
E3 Media (pH 7.2) | 5 mM NaCl | Sigma | |
0.17 mM KCl | Sigma | ||
0.33 mM CaCl2 | Sigma | ||
0.33 mM MgSO4 | Sigma | ||
Injection Buffer (pH 7.5) | 20 mM HEPES | Sigma | |
150 mM KCl | Sigma |