CRISPR-Cas technologies have revolutionized the field of genome editing. However, finding and isolating the desired germline edit remains a major bottleneck. Therefore, this protocol describes a robust method for quickly screening F0 CRISPR-injected zebrafish sperm for germline edits using standard PCR, restriction digest, and gel electrophoresis techniques.
The advent of targeted CRISPR-Cas nuclease technologies has revolutionized the ability to perform precise genome editing in both established and emerging model systems. CRISPR-Cas genome editing systems use a synthetic guide RNA (sgRNA) to target a CRISPR-associated (Cas) endonuclease to specific genomic DNA loci, where the Cas endonuclease generates a double-strand break. The repair of double-strand breaks by intrinsic error-prone mechanisms leads to insertions and/or deletions, disrupting the locus. Alternatively, the inclusion of double-stranded DNA donors or single-stranded DNA oligonucleotides in this process can elicit the inclusion of precise genome edits ranging from single nucleotide polymorphisms to small immunological tags or even large fluorescent protein constructs. However, a major bottleneck in this procedure can be finding and isolating the desired edit in the germline. This protocol outlines a robust method for screening and isolating germline mutations at specific loci in Danio rerio (zebrafish); however, these principles may be adaptable in any model where in vivo sperm collection is possible.
The CRISPR (Clustered Regularly Interspaced Short Palindromic Repeats)/Cas system is a powerful tool to perform loci-specific mutagenesis and precise genome editing in the Danio rerio (zebrafish) model system1,2,3,4. The Cas-ribonucleoprotein (RNP) is comprised of two main components: a Cas endonuclease (commonly Cas9 or Cas12a) and a locus-specific synthetic guide RNA (sgRNA)5. Together, the Cas-RNP generates a double-stranded break (DSB) in the desired locus that can be repaired by one of two intrinsic repair mechanisms. The non-homologous end joining (NHEJ) repair mechanism is error-prone and often results in a variety of insertions or deletions (indels) around the DSB. These indels can be deleterious if they introduce a frameshift mutation or premature stop in the resultant protein sequence. Alternatively, the homology-directed repair (HDR) mechanism uses a donor template with regions of homology surrounding the DSB site to repair the damage. Researchers can take advantage of the HDR system to generate precise genomic edits. Specifically, they can co-inject a double-stranded DNA donor construct that contains the desired edits as well as regions of homology flanking the DSB site in the genome. The increased economy of scale for these commercially produced CRISPR components has greatly reduced the barriers to screening multiple loci and to setting up larger-scale efforts for precise genome editing. However, in sexually reproducing animal models, a major bottleneck is the identification and isolation of germline-stable mutant animals.
The zebrafish model system exhibits several key qualities that enhance its use in reverse genetic studies. They are easy to rear in large numbers with basic aquatic housing equipment, and females exhibit high fecundity all year round6. Moreover, their external egg-laying and fertilization make them amenable to the microinjection of CRISPR/Cas components. The Cas-RNP is commonly injected into one-cell stage zebrafish embryos to generate DSBs/repair that is, in theory, inherited by all the daughter cells. However, diploid genomes require two DSB/repair events to mutagenize both homologous chromosomes. Furthermore, although Cas-RNP is injected at the one-cell stage, the DSB/repair may not occur until later points in development. Together, these factors contribute to the mosaic nature of F0-injected fish. A common practice is to outcross F0-injected fish and screen the F1 progeny for indels/specific edits. However, since not all F0-injected fish possess germline mutations, this practice results in many unproductive crosses that do not generate the desired edit. Screening the F0 germline rather than F1 somatic tissue increases the probability of isolating the desired germline edit and reduces the number of animals required in this process.
Sperm can easily be collected from F0-injected zebrafish without the need for euthanasia. This feature allows for the cryopreservation and rederivation of frozen sperm stocks7 but can also be exploited to rapidly screen, identify, and isolate the germline carriers of desired genomic mutations8,9. Brocal et al. (2016) previously described a sequencing-based method for screening germline edits in F0-injected male zebrafish10. Although useful for identifying the mutated alleles present in the germline, this approach can become costly in high throughput and may not be accessible for all labs. In contrast, the current protocol offers an approachable and cost-effective electrophoresis-based strategy for identifying germline edits. Specifically, this protocol outlines a robust method for screening and isolating germline mutations at specific loci using high-resolution agarose gel electrophoresis. In addition, this protocol describes a similar strategy for identifying the successful integration of a donor construct containing specific edits. As always, if specific edits are desired, sequencing-based strategies can be performed in tandem with the protocol described below. Although this protocol is specific to the zebrafish model system, these principles should be adaptable to any model in which the collection of sperm is a routine procedure. Together, these strategies will allow for the identification of F0-injected males with germline indels/edits that can be resolved on a gel after standard polymerase chain reaction (PCR) and/or restriction digest.
This study was carried out in line with the guidelines in the Guide for the Care and Use of Laboratory Animals of the National Institutes of Health. The protocol was approved by the University of Texas at Austin Animal Care and Use Committee (AUP-2021-00254).
1. Designing the sgRNA for CRISPR mutagenesis
2. Setting up the breeding tanks
3. Preparing the materials for the sperm collection procedure
4. Anesthetizing the male fish and collecting the sperm
5. Extracting DNA from the sperm samples
6. PCR amplification (and/or restriction digest) of the desired locus
7. Performing gel electrophoresis to separate PCR amplicons of varying sizes
8. Isolating germline stable alleles
The experimental approaches described in this protocol allow for the more rapid identification of genome edits or putative deleterious alleles by focusing on the analysis of thousands of genomes derived from the collection of F0-injected male sperm. Figure 2 highlights how to interpret the results obtained using this protocol.
To generate mutations in the p2ry12 locus, one-cell stage zebrafish embryos were injected with Cas9 endonuclease and a p2ry12-specific sgRNA (Figure 2B, gray highlight). F0-injected fish were put in the system until sexual maturity, and sperm was collected from six individual males. DNA was extracted from the sperm samples using high heat and basic conditions (HotSHOT DNA extraction) and neutralized prior to the PCR amplification of the p2ry12 locus. The PCR products were separated on a high-percentage (4%) agarose gel for 1.5 h at a high voltage (150 V). In this example, the wild-type amplicon ran as a single bright band of around 250 base pairs in length (Figure 2C; WT). In contrast, the F0-injected male amplicons containing indels ran on the gel as multiple bands (Figure 2C; lanes #1-2, #5-8). Alternatively, the mutated F0-injected male amplicons could run as a single band with altered gel mobility (not shown in the p2ry12 example, but evident in the PCR product of the dnah10 F0-male #2; Figure 2E). It was less clear on the p2ry12 example gel whether sample #3 and sample #4 contained indels in comparison to the wild-type band, so these individuals may not be the best founder candidates. The isolation of alleles from F0 males with more distinctive amplicon alterations is best for propagating stable germline carriers as they are easily scored on 4% agarose gel. For example, the F0-male #6 sperm sample appeared to contain a large deletion in one of the alleles (Figure 2C; lane 6; bright band with increased gel mobility). If this large deletion allele was selected for in the F1 generation, it would be easily distinguishable from the WT allele on a 4% agarose gel. Alternatively, if a collection of different alleles is desired, the F0-male #7 could be an effective founder candidate since its PCR product appeared to contain several discrete alleles of various mobilities. Once a founder male is selected, the desired allele can be isolated by outcrossing the individual back to the original wild-type strain used for injections.
To generate a specific knock-in mutation in the dnah10 gene, one-cell stage zebrafish embryos were injected with Cas9 endonuclease, a dnah10-specific sgRNA (Figure 2D; gray highlight), and a DNA donor oligonucleotide containing the desired edit (Figure 2D; red) and a donor-specific BstNI restriction site (Figure 2D, brackets). This design allows for the easy identification of donor integration with a restriction digest after PCR. Furthermore, by altering base pairs within the sgRNA recognition site (Figure 2D; gray highlight), this design prevents the Cas9 digestion of the donor sequence. Once the F0-injected fish reached sexual maturity, sperm was collected, and DNA was extracted using the hot shot method. From these samples, the dnah10 locus was amplified using PCR, and the products were separated on a high-percentage (4%) agarose gel for 1 h at a high voltage (150 V). In this example, the wild-type amplicon ran as a single bright band of around 400 base pairs in length (Figure 2E; top panel: WT). In contrast, the F0-injected male amplicons containing indels ran as multiple bands (Figure 2E; top panel: lanes #1, #4, and #5) or as a single band with decreased gel mobility (Figure 2E; top panel: lane #2). To determine if any of the sperm samples contained donor-integrated alleles, the PCR products were digested with the BstNI restriction enzyme for 1 h, and the product was run on a 4% agarose gel for 1 h at high voltage (150 V). Comparing the undigested PCR product (Figure 2D; top panel) to the digested product (Figure 2D; bottom panels) reveals samples with likely integration of the donor construct. In this example, the F0-injected male #1 had an additional band in the digested product that was not present in the PCR product (Figure 2D; bottom panel: lane #1, circled). Therefore, this male represents the best founder candidate for establishing a mutant line with the desired knock-in.
Once the putative F0 carrier male is outcrossed, the allele must be sequence-verified in the F1 progeny. It is suggested to directly Sanger sequence the amplicons derived from the F1 heterozygous fish followed by performing heterozygous allele analysis methods such as Poly Peak Parser13 or using a variety of next-generation sequencing-based methods such as MiSeq14 or Hi-Tom15 sequencing. This is a necessary and complementary approach to ensure that the precise edit or deleterious indel mutation is actually going germline and is not just an artifact of the gel electrophoresis analysis. Indeed, the use of next-generation sequencing approaches to sequence sperm DNA from F0 carriers can easily be used in place of the gel electrophoresis analysis described in this protocol. However, having an easily scorable agarose gel genotyping method is a more economical and egalitarian approach for the global community of zebrafish researchers.
Figure 1: Setup for the sperm squeezing procedure. (A) The sponge is positioned underneath a stereomicroscope at low magnification with overhead lighting. (B) The moistened 1 in x 1 in sponge, cut with an oval divot (dashed line), is used to hold the anesthetized male fish ventral side up during the procedure. (C) Anatomy of the ventral side of the anesthetized male fish depicting the anal fins (af, pink), pelvic fins (pf, blue), and cloaca (arrow), where sperm will be expelled during the procedure. (C') Filter forceps are used to gently squeeze the anesthetized male fish from the gills to the cloaca, while the expelled sperm is drawn into a glass pipette by capillary action (white arrow). (D) A capillary tube containing sufficient expelled sperm (opaque liquid; black bracket) for downstream DNA extraction and analysis. Please click here to view a larger version of this figure.
Figure 2: Workflow and representative results. (A) General workflow of the protocol. (B) Representative design of the p2ry12 sgRNA site (grey highlight) with a Cas9-specific PAM site (underlined). (C) Representative results of the 4% gel electrophoresis after 1.5 h at 150 V. Wild-type (WT) control and F0-p2ry12 CRISPR-injected male sperm samples (1-8) after PCR amplification. (D) Representative design of the dnah10 sgRNA site (grey highlight) with a Cas9-specific PAM site (underlined) near the targeted codon (bold). DNA donor sequence with the desired codon edit (red bold) and BstNI restriction site (brackets with apostrophe marking the cut site). (E) Representative results of the 4% gel electrophoresis after 1 h at 150 V. Top: PCR product of the wild-type (WT) and F0 dnah10 CRISPR-injected male sperm samples (1-5). Bottom: BstNI restriction digest product of the above samples. Sample 1 demonstrates the successful integration of the donor construct based on the additional band after digestion (red circle). Please click here to view a larger version of this figure.
This protocol describes a method for rapidly characterizing putative genome edits or targeted mutations using CRISPR-Cas technology by focused analysis on F0 male sperm genomes. This protocol should be amenable to other animal models where sperm is readily available for sampling without euthanasia. This method will increase the throughput of screening for desired edits and is especially useful for identifying rare HDR-mediated knock-in events. This approach also serves to reduce the number of experimental animals used to find a stable germline edit by facilitating the rapid screening of potentially thousands of genomes in one sperm sample from a putative F0 carrier, which is in contrast with more traditional approaches that may require the screening of hundreds of embryos derived from crosses of putative F0 carriers.
This protocol builds on established protocols for sperm collection in zebrafish7,10,14,16,17,18 by including a reproducible technique for identifying germline edits using high-resolution gel electrophoresis. This approach can easily be incorporated into any standard CRISPR/Cas workflow to increase the throughput for the screening and isolation of target genome edits. Additionally, this protocol is appropriate for labs staffed with personnel with a range of training and experience, as well as teaching labs. However, our method of sperm collection is not sufficient for cryopreservation, which has been expertly described in previous publications10,17.
This protocol uses a high-resolution agarose gel electrophoresis to identify putative male carriers of desired indel and precise genomic mutations. However, sperm genomic DNA is amenable to a myriad of other approaches, including fluorescent fragment analysis or barcoded sequencing14, high-resolution melt analysis18, or simple amplicon detection of fluorescent tags or epitopes using standard gel electrophoresis approaches. The downstream validation of all alleles using approaches such as Sanger-based or next-generation sequencing must be done before starting experimental work on putative alleles. Indeed, existing methods for screening for mutations in embryos using the next-generation sequencing approach14 could circumvent the need for analysis on agarose gels, which is described in this protocol. However, having a gel-scorable genotyping method is a more cost-effective and egalitarian approach to consider given that not all labs have easy access to cheap next-generation sequencing methods during the isolation and experimental phases of working with each mutant strain.
In summary, this protocol provides step-by-step directions for reproducibly screening sperm genomes from CRISPR/Cas-edited males such that fewer crosses and less PCR screening of embryos are needed to screen for desired edits. The application of this method will decrease the number of fish that need to be created and analyzed to successfully identify the edited alleles of interest, which also reduces the time and cost of personnel for generating stable lines.
The authors have nothing to disclose.
We would like to thank Anna Hindes at Washington University School of Medicine for her initial efforts in obtaining good-quality sperm genomic DNA using the hot shot method. This work was funded by the National Institute of Arthritis and Musculoskeletal and Skin Diseases of the National Institutes of Health under Award (R01AR072009 to R.S.G.).
Agarose powder | Fisher BioReagents | BP1356-100 | |
Breeding tanks | Carolina Biological | 161937 | |
BstNI Restriction Enzyme | NEB | R0168S | |
Cas9 Endonuclease | IDT | 1081060 | |
DNA Ladder, 100 bp | Thermo Scientific | FERSM0241 | |
dnah10 donor construct | Sigma-Aldrich | DNA Oligo in Tube; 0.025 nM, standard desalt purification, dry. Phosphorothioate bond on the donor at the first three phosphate bonds on both the 5’ and 3’ ends (5'-CCTCTCTCCCTTTCAGAAGCTTC TGCTCATCCGCTGCTTCTGCCT GGACCGAGTGTACCGTGCCGTC AGTGATTACGTCACGC-3') |
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dnah10 forward primer | Sigma-Aldrich | DNA Oligo in Tube; 0.025 nM, standard desalt purification, dry (5'-CATGGAACTCTTTCCTAATGAGT TTGGC-3') |
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dnah10 reverse primer | Sigma-Aldrich | DNA Oligo in Tube; 0.025 nM, standard desalt purification, dry ('5-AGTAGAGATCACACATCAACAGA ATACAGC-3') |
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dnah10 synthetic sgRNA | Synthego | Synthetic sgRNA, target sequence: 5'-GCTCATCCGCTGCTTCAGGC-3' | |
Electrophoresis power supply | Thermo Scientific | 105ECA-115 | |
Filter forceps | Millipore | XX6200006P | |
Fish (system) water | Generic | n/a | |
Gel electrophoresis system (including casting frame, comb, and electrophoresis chamber) | Thermo Scientific | B2 | |
Gel imaging light box | Azure Biosystems | AZI200-01 | |
Gel stain, 10000X | Invitrogen | S33102 | |
Glass bowl, 250 mL | Generic | n/a | |
Isolation tanks, 0.8 L | Aquaneering | ZT080 | |
Microcap capillary tube with bulb, 20 µL | Drummond | 1-000-0020/CA | |
Minicentrifuge | Bio-Rad | 12011919EDU | |
Micropipettes, various with appropriate tips | Generic | n/a | |
Microwave | Generic | n/a | |
Nuclease free water | Promega | P119-C | |
Paper towels | Generic | n/a | |
PCR tubes, 0.2 mL | Bioexpress | T-3196-1 | |
Plastic spoon, with drilled holes/slots | Generic | n/a | |
KCl solution, 0.2 M RNAse Free | Sigma-Aldrich | P9333 | |
p2ry12 forward primer | Sigma-Aldrich | DNA Oligo in Tube; 0.025 nM, standard desalt purification, dry (5'-CCCAAATGTAATCCTGACCAGT -3') |
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p2ry12 reverse primer | Sigma-Aldrich | DNA Oligo in Tube; 0.025 nM, standard desalt purification, dry (5'-CCAGGAACACATTAACCTGGAT -3') |
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p2ry12 synthetic sgRNA | Synthego | Synthetic sgRNA, target sequence: 5'-GGCCGCACGAGGTCTCCGCG-3' | |
Restriction Enzyme 10X Buffer | NEB | B6003SVIAL | |
NaOH solution, 50 mM | Thermo Scientific | S318; 424330010 | |
Sponge, 1-inch x 1-inch cut with small oval divot | Generic | n/a | |
Stereomicroscope | Zeiss | Stemi 508 | |
Taq polymerase master mix, 2X | Promega | M7122 | |
TBE Buffer Concentrate, 10X | VWR | E442 | |
Thermal Cycler | Bio-Rad | 1861096 | |
Tissue paper | Fisher Scientific | 06-666 | |
Tricaine-methanesulfonate solution (Syncaine, MS-222), 0.016% in fish water (pH 7.0±0.2) | Syndel | 200-266 | |
Tris Base, 1M (Buffered with HCl to ph 8.0) | Promega | H5131 |