Name: efficient genome editing of mice by crispr electroporation of zygotes
Uploaded: 2022-12-16T13:20:00.000+00:00
Duration: 7 min 17 s
Description: Watch this Scientific Journal Video about Efficient Genome Editing of Mice by CRISPR Electroporation of Zygotes at JoVE.com

A.J.M. created the original concept that led to the development of CRISPR-EZ and produced the figures. C.K.D. compiled and adapted the internal and published protocols for this current manuscript. A.J.M. is supported by NIH (R00HD096108).

<ol>
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Presented here is a straightforward and highly efficient mouse genome editing technology. Electroporation can be used to generate modified embryos in 1-2 weeks (Figure 1) and can produce edited mice within 6 weeks9. Compared to contemporaneously developed electroporation-based protocols that deliver RNPs7,10,11,12,13,14,15,17,32, the method as described here is conceptually similar and offers efficiencies in the same range, with only minor differences in reagent development and parameters. Therefore, we suggest that readers compare and contrast based on needs and access to equipment. As the name implies, this protocol was developed with &#34;the average mouse lab&#34; in mind, with only common methods (IVT, PCR, gel electrophoresis), reagents (standard embryo and tissue culture consumable), or equipment (electroporator and cuvettes commonly used for bacterial transfection), which are likely accessible to most labs who are interested and able to collect embryos (or by kindly asking neighboring labs). Graduate student-level researchers with basic embryo handling skills can test sgRNAs for efficiency or conduct data-producing experiments multiple times within the timeframe of preimplantation development without the need to schedule with a core facility. However, if the desired goal is to generate animal models without the need for either microinjection or embryo manipulation, less intrusive methods are available10,32.
Many factors that impact Cas9 efficiency are actively being researched, such as the specificity of a genomic target, sgRNA sequence parameters, genome accessibility and topology, and, especially, cell type. Therefore, designing and testing sgRNAs is crucial for success. This protocol and other independently developed electroporation methods have been optimized to deliver Cas9 protein/sgRNA RNPs quickly and transiently into zygote-stage embryos, enhancing efficiency while minimizing mosaicism due to delivery at an early developmental stage and the short half-life of the RNP7,9,10,11,12,13,14,15,16,33,34.
For the most common genome editing strategies and various strains of mice, this method can outperform microinjections in terms of cost, efficiency, small insertions, in/del mutations, exon deletions, insertions, and point mutations9. With the current protocol, this method is ideal for the creation of in/del mutations and deletions up to 2.6 kb, which is much longer than the typical length of a protein-coding exon of 170 bp35. Long deletions are less efficient; however, this limitation is not unique to this protocol. Therefore, as Cas9-mediated editing is improved and new Cas protein variants are developed, the modular nature of this method allows for these improvements to directly augment the capabilities of our protocol.
While this method can perform a variety of simple edits, its potential for larger and more complex strategies, such as inserting conditional alleles or fluorescent tags, is currently being tested in our lab. Longer ssODNs could provide a viable approach for complicated genome editing and have shown success in microinjection-based embryo editing36; however, longer ssODNs are expensive to synthesize and have yet to be tested extensively with electroporation37. The use of adeno-associated viruses (AAV) to introduce up to 3.3 kb donor sequences has shown success as well but may not be accessible to most labs25. For the time being, we recommend standard embryonic stem cell genome editing and microinjection for complex mouse genome engineering.
Worries about Cas9 off-target effects remain8,38,39. Engineered Cas9 variations might increase target specificity, although this has not been fully investigated in RNP embryo electroporation studies. CRISPR-EZ could be a useful method for creating compound mutation models to explore complex genetics. While microinjections have made simultaneously editing multiple loci possible40, electroporation methods like the one described here make this more convenient and effective.

Genome editing in live mice has been considerably simplified and has become accessible and more affordable since the emergence of CRISPR editing1,2,3. Initial animal editing attempts used microinjection to deliver CRISPR Cas9 mRNA/sgRNA into pronuclear-stage embryos4,5,6. While microinjection is quite effective, the amount of practice required to fully master it might not be appropriate for trainees and students and also requires expensive equipment that a modestly funded lab is unable to afford. Microinjection is normally performed by expert technicians at transgenic facilities with schedules and service prices that are rate-limiting for many researchers. A more accessible approach is that of electroporation, which has been demonstrated to be quite effective for the delivery of CRISPR Cas9 mRNA/sgRNA into pronuclear-stage embryos7. Further improvements in CRISPR genome editing and delivery strategies suggested that pre-assembled RNPs already engaged with sgRNAs may be an effective means to reduce mosaicism8.
The rationale behind the development and use of this protocol was to bypass many of the limitations and obstacles associated with microinjection. As the name implies, an easy, in-house, and cost-effective method that could quickly determine whether untested sgRNA designs would be worthwhile using during a microinjection experiment would be a very convenient first pass quality control step (Figure 1). While this method cannot replace microinjection for more complex strategies, like introducing long donor DNA sequences for recombination-based outcomes, it is ideal for less complex strategies like small deletions or insertions and tagging genes. This method is appropriate for researchers with basic embryo manipulation skills who have simple editing needs, would like to test their hypothesis within the timeframe of preimplantation development, or prefer to test sgRNAs in embryos before scheduling an appointment with a microinjection specialist. Here, editing reagents are transiently delivered into pronuclear-stage embryos as Cas9/sgRNA RNPs via electroporation (a series of electrical pulses) to maximize efficiency while decreasing mosaicism8. Using an embryo genotyping method, editing results are available in approximately 1 week9, thus reducing the need for various microinjection applications at a significantly reduced cost.
This method&#39;s effectiveness peaks at the pronuclear embryo stage, when the embryo has not yet fused the maternal and paternal pronuclei or entered S-phase (Figure 2). Superovulation is used to maximize the number of zygotes but produces both pronuclear zygotes and unfertilized eggs. Healthy zygotes can also be pre-selected before electroporation to increase the overall efficiency. As other electroporation protocols have efficiently edited zygotes without the need to include a similar step7,10,11,12,13,14,15, an optional step of this protocol is the slight erosion of the zona pellucida (ZP). The ZP is a glycoprotein layer that aids spermatozoa binding, acrosome response, and fertilization surrounding pronuclear-stage embryos. In our experience, we found that a gentle acid-based erosion of the ZP provides reliable Cas9 RNP electroporation delivery with only a marginal impact on viability.
We have observed RNP delivery rates of up to 100% efficiency via electroporation in mouse strains that are commonly used in research like C57BL/6J and C57BL/6N9,16. Independent groups have also developed electroporation-based procedures with efficiencies greater than or matching microinjection11,12,13,14,15,17, with electroporation protocols functioning well in rat18,19, pig20,21,22 and cow23, so we suggest that readers compare the protocols to find the conditions that best suit their experimental and equipment needs. The system described here uses common materials and equipment, requiring only basic embryo manipulation skills. This technique is effective for a range of editing strategies, making this method broadly accessible to the research community.
Designing ideal small guide RNAs (sgRNAs) is essential for efficient editing. We recommend screening two to three sgRNA strategies per target site directly in mouse embryos, especially if mouse line generation is desired. Once designed, cloning-free methods like in vitro transcription (IVT) to produce high-quality sgRNAs are recommended3. The RNPs and sgRNAs are mixed with 30-50 processed pronuclear-stage embryos and exposed to a series of electrical pulses to first temporarily permeabilize the ZP and cell membrane, with subsequent pulses to keep the pores open and electrophorese the RNPs through the zygote24. After optimization, we found that six 3 ms pulses at 30 V for bulk embryos (~50) were optimal for editing effectiveness and viability, providing highly efficient Cas9/sgRNA RNP delivery9,16,25. Editing events in individual mouse morula can be confirmed using a variety of validation strategies common for CRISPR editing, such as restriction fragment length polymorphism (RFLP), T7 endonuclease digestion, and Sanger sequencing of the region of interest26.
The current method is most appropriate for simple editing schemes (Figure 3), such as insertion/deletions (indels), exon-sized deletions on the order of 500-2000 bp, and the delivery of point mutations and small insertions such as C- or N-Terminal tags (e.g., FLAG, HA, or V5)9,16,27. The potential for complex genome editing, like large insertions of fluorescent tags or conditional alleles, remains uncertain and is the present focus of upcoming improvements.
This method is easily mastered and can be used to quickly test sgRNAs in cultured mouse embryos in 1 week9 (Figure 1). Presented in this work is a six-step protocol, which includes 1) sgRNA design; 2) sgRNA synthesis; 3) superovulation and mating; 4) embryo culture, collection, and processing; 5) RNP assembly and electroporation; 6) embryo culture and genotyping. Information about all the materials used is provided (Table of Materials). As a positive control, reagents to edit the Tyrosinase (Tyr) locus9,16 have been included in the Supplementary Table 1.

<table><tbody><tr><td>0.1-cm-gap electroporation cuvette</td><td>Bio-Rad</td><td>cat. no. 1652089</td><td>Electroporation</td></tr><tr><td>26-G, 1/2-inch needle</td><td>BD</td><td>cat. no. 305111</td><td>Superovulation</td></tr><tr><td>3&amp;ndash;8-month-old male mice and 3- to 5-week-old female mice</td><td>JAX</td><td>cat. no. 000664</td><td>Superovulation</td></tr><tr><td>35-mm Tissue culture dish</td><td>Greiner Bio-One,</td><td>cat. no. 627-160</td><td>Embryo Culture</td></tr><tr><td>60-mm Tissue culture dish</td><td>Greiner Bio-One,</td><td>cat. no. 628-160</td><td>Embryo Processing</td></tr><tr><td>6x loading dye</td><td>Thermo Fisher Scientific</td><td>cat. no. R0611</td><td>sgRNA Synthesis and Genotyping</td></tr><tr><td>Acidic Tyrode&#039;s (AT) solution, embryo culture grade</td><td>Sigma-Aldrich,</td><td>cat. no. T1788</td><td>Embryo Processing</td></tr><tr><td>BSA, embryo culture grade</td><td>Sigma-Aldrich</td><td>cat. no. A3311</td><td>Embryo Processing and Culture</td></tr><tr><td>Cas9 protein</td><td>Alt-R S.p. Cas9 nuclease 3NLS</td><td>cat. no. 1074181</td><td>Electroporation</td></tr><tr><td>DNase I, RNase-free</td><td>New England BioLabs,</td><td>cat. no. M0303</td><td>sgRNA Synthesis</td></tr><tr><td>DPBS(calcium and magnesium free)</td><td>Gibco</td><td>cat. no. 14190-144</td><td>Embryo Processing</td></tr><tr><td>EcoRI</td><td>NEB</td><td>cat. no. R3101S</td><td>Genotyping</td></tr><tr><td>EDTA, anhydrous</td><td>Sigma-Aldrich</td><td>cat. no. EDS-100G</td><td>RNP Buffer</td></tr><tr><td>Ethanol</td><td>Koptec</td><td>cat. no. V1016</td><td>sgRNA Synthesis</td></tr><tr><td>Gelatin (powder) type B, laboratory grade</td><td>Fisher,</td><td>cat. no. G7-500</td><td>Lysis Buffer</td></tr><tr><td>Glycerol,&amp;nbsp;molecular-biology grade</td><td>Fisher</td><td>cat. no. BP229</td><td>RNP Buffer</td></tr><tr><td>Taq Polymerase</td><td>Promega</td><td>cat. no. M712</td><td>Genotyping</td></tr><tr><td>HEPES, cell culture grade</td><td>Sigma-Aldrich</td><td>cat. no. H4034</td><td>RNP Buffer</td></tr><tr><td>HinfI (10,000 U/mL)</td><td>NEB</td><td>cat. no. R0155S</td><td>Genotyping</td></tr><tr><td>HiScribe T7 High Yield RNA Synthesis Kit</td><td>New England BioLabs,</td><td>cat. no. E2040</td><td>sgRNA Synthesis</td></tr><tr><td>Human chorion gonadotropin, lyophilized&amp;nbsp;(hCG)</td><td>Millipore</td><td>cat. no. 230734</td><td>Superovulation</td></tr><tr><td>Hyaluronidase/M2</td><td>Millipore</td><td>cat. no. MR-051-F</td><td>Embryo Processing</td></tr><tr><td>KSOMaa Evolve medium (potassium-supplemented simplex-optimized medium plus amino acids)</td><td>Zenith Biotech</td><td>cat. no. ZEKS-050</td><td>Embryo Culture</td></tr><tr><td>LE agarose, analytical grade</td><td>BioExpress</td><td>cat. no. E-3120-500</td><td>sgRNA Synthesis and Genotyping</td></tr><tr><td>M2 medium</td><td>Zenith Biotech</td><td>cat. no. ZFM2-050</td><td>Embryo Processing</td></tr><tr><td>Magnesium chloride, anhydrous (MgCl&lt;sub&gt;2&lt;/sub&gt;)</td><td>Sigma-Aldrich</td><td>cat. no. M8266</td><td>RNP and Lysis Buffer</td></tr><tr><td>Mineral Oil</td><td>Millipore</td><td>cat. no. ES-005C</td><td>Embryo Culture</td></tr><tr><td>Nonidet P-40,substitute (NP-40)</td><td>Sigma-Aldrich</td><td>cat. no. 74385</td><td>Lysis Buffer</td></tr><tr><td>Nuclease-free water, molecular-biology grade</td><td>Ambion</td><td>cat. no. AM9937</td><td>sgRNA Synthesis and Genotyping</td></tr><tr><td>Oligos for sgRNA synthesis, donor oligo and PCR primers for genotyping</td><td>Integrated DNA Technologies</td><td>custom orders</td><td>sgRNA Design</td></tr><tr><td>Reduced serum medium</td><td>Thermo Fisher Scientific</td><td>cat. no. 31985062</td><td>Embryo Culture</td></tr><tr><td>High-fidelity DNA polymerase</td><td>New England BioLabs,</td><td>cat. no. M0530</td><td>sgRNA Synthesis</td></tr><tr><td>Potassium chloridemolecular-biology grade (KCl)</td><td>Sigma-Aldrich</td><td>cat. no. P9333</td><td>RNP and Lysis Buffer</td></tr><tr><td>Pregnant mare serum gonadotropin lyophilizd ((PMSG)</td><td>ProspecBio</td><td>cat. no. HOR-272</td><td>Superovulation</td></tr><tr><td>Proteinase K, molecular-biology grade</td><td>Fisher</td><td>cat. no. BP1700-100</td><td>Lysis Buffer</td></tr><tr><td>RNase-free 1.5-mL microcentrifuge tube</td><td>VWR</td><td>cat. no. 20170-333</td><td>sgRNA Synthesis and Genotyping</td></tr><tr><td>RNase-free eight-well PCR strip tubes</td><td>VWR</td><td>cat. no. 82006-606</td><td>sgRNA Synthesis and Genotyping</td></tr><tr><td>Magnetic purification beads</td><td>GE Healthcare</td><td>cat. no. 65152105050250</td><td>sgRNA Synthesis</td></tr><tr><td>Tris (2-carboxyethyl) phosphine hydrochloride (TCEP)</td><td>Sigma-Aldrich</td><td>cat. no. C4706</td><td>RNP Buffer</td></tr><tr><td>Tris-HCl solution, pH 8.5 molecular-biology grade</td><td>Teknova</td><td>cat. no. T1085</td><td>Lysis Buffer</td></tr><tr><td>Tween 20 molecular-biology grade</td><td>Sigma-Aldrich</td><td>cat. no. P7949-500</td><td>Lysis Buffer</td></tr></tbody></table>

efficient genome editing of mice by crispr electroporation of zygotes

With exceptional efficiency, accuracy, and ease, the CRISPR/Cas9 system has significantly improved genome editing in cell culture and lab animal experiments. When generating animal models, the electroporation of zygotes offers higher efficiency, simplicity, cost, and throughput as an alternative to the gold standard method of microinjection. Electroporation is also gentler, with higher viability, and reliably delivers Cas9/single-guide RNA (sgRNA) ribonucleoproteins (RNPs) into the zygotes of common laboratory mouse strains (e.g., C57BL/6J and C57BL/6N) that approaches 100% delivery efficiency. This technique enables insertion/deletion (indels) mutations, point mutations, the deletion of whole genes or exons, and small insertions in the range of 100-200 bp to insert LoxP or short tags like FLAG, HA, or V5. While constantly being improved, here we present the current state of CRISPR-EZ in a protocol that includes sgRNA production through in vitro transcription, embryo processing, RNP assembly, electroporation, and the genotyping of preimplantation embryos. A graduate-level researcher with minimal experience manipulating embryos can obtain genetically edited embryos in less than 1 week using this protocol. Here, we offer a straightforward, low-cost, efficient, high-capacity method that could be used with mouse embryos.

This method generates more than 100 &#181;g of sgRNA (20 &#181;L at &#62;6,000 ng/L concentration) for efficient Cas9/sgRNA RNP assembly. The routine superovulation method described here typically produces 10-20 viable embryos per plugged female. Due to handling errors and typical losses associated with embryo manipulation, an expected 80% of embryos are fertilized, viable, and in excellent condition after electroporation. To aid researchers in executing a successful experiment, we have provided an example strategy to target the mouse Tyr locus as a positive control (Figure 6), including oligo designs (Figure 6A, Supplementary Table 1) and genotyping strategies for both NHEJ and HDR events (Figure 6B) (step 6.2). Detailed examples of experimental success and outcomes are available9,16.
In a previous effort to compare this protocol to microinjection, collectively, over 30 distinct editing attempts involving seven distinct labs to generate mouse models were all successful9,16. Additionally, this method was used to investigate and report on the first essential retrotransposon in mammalian development27. When using a simple strategy such as delivering a single sgRNA, the delivery of RNPs was up to and including 100% in the C57BL/6J and C57BL/6N mouse strains, with in/del formation occurring at 50-100% and small oligo-based replacements ranging from 14%-63%9,16 (Table 1). When engineering genomic deletions, the editing effectiveness can vary from 3% to 100%, where contributing factors include genomic location, sgRNA design, and the size of the deletion (Table 2). For example, deletions smaller than 1,000bp are more successful than those larger than 1,000 bp9,16.
When using 60 embryos for electroporation, this protocol surpasses microinjection in terms of editing effectiveness, resulting in 3-4 founder animals compared to one founder from microinjection9. For gene knockout experiments in the C57BL/6N mice strain using identical sgRNA, this method outperformed microinjection, yielding an average of four founder animals9 (Table 2). For small insertions, such as V5 or HA tagging, ssODNs up to 162 nt have been successfully tested, and efforts are currently aimed at scaling up to 1000-2000 nt. The success of these experiments largely depends on the effectiveness of the sgRNA used for HDR outcomes to be robust. Nearly all HDR outcomes are mosaic, but between 31%-64% of embryos display some evidence of insertions9,16.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/64302/64302fig01.jpg" /> 
Figure 1: Overview of CRISPR-EZ. A graphical overview of the workflow. Once a strategy and design have been made, edited embryos can be generated and tested in approximately 1 week. This figure has been modified with permission from Modzelewski et al.9. and Chen et al.16. <a href="https://www-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/64302/64302fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/64302/64302fig02.jpg" /> 
Figure 2: Ideal timing to perform the procedure. The diagram shows relevant features and time points during the first cellular division after fertilization. To carry out this approach, researchers are provided with some visible clues, such as morphological changes, cell cycle time estimations, and chemical markers frequently observed before the first cleavage. Since the exact timing of insemination and fertilization is unknown, a sensible recommendation would be to regard hour 0 as midnight. Before the zygote enters S-phase is the ideal moment to deliver editing machinery for either NHEJ or HDR, which translates to executing this protocol between the hours of 7 AM and 11 AM in the morning. This figure has been modified with permission from Modzelewski et al.9. <a href="https://www-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/64302/64302fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/64302/64302fig03.jpg" /> 
Figure 3: Editing strategies&#39; success. Simple strategies such as a single sgRNA result in an in/del via NHEJ repair or a precision mutation when combined with an ssODN template through HDR. NHEJ repair can also be utilized for deletions using multiple sgRNAs. In general, the simpler the strategy, the more effective CRISPR-EZ is without further optimization. This figure has been modified with permission from Modzelewski et al.9. <a href="https://www-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/64302/64302fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/64302/64302fig04.jpg" /> 
Figure 4: Locating the oviduct and ampulla. After surgically isolating the reproductive tissues, one should secure the region by applying gentle pressure to the fat pad with forceps. The fat pad is a relatively safe region to manipulate to not harm the ovaries/oviduct. The area in the dashed square should be removed and placed into a droplet of M2 for further dissection. A cartoon diagram shows the region of interest with the relevant anatomical structures labeled. This figure has been modified with permission from Modzelewski et al.9. <a href="https://www-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/64302/64302fig04large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/64302/64302fig05.jpg" /> 
Figure 5: Suggested processing and culture plate setup. Schematics showing various plate setups to aid researchers in conducting both embryo processing and culture. (A) Typical M2+BSA wash plate. (B) M2+hyaluronidase plate for cumulus cell removal. (C) Optional AT solution plate for Zona thinning. (D) KSOM+BSA plate for embryo culture, with mineral oil overlay necessary for maintaining pH, humidity, and temperature. This figure has been modified with permission from Modzelewski et al.9. <a href="https://www-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/64302/64302fig05large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 6" class="xfigimg" src="/files/ftp_upload/64302/64302fig06.jpg" /> 
Figure 6: Genotyping example for NHEJ and HDR outcomes. (A) A cartoon diagram of the region surrounding the Tyr gene in the mouse genome. Below are details of the unedited (WT) sequence where a naturally occurring HinfI restriction site is found, which is within the target recognition site of the sgRNA (red text). Below this is one of many possible outcomes after successful CRISPR/Cas9 targeting where an &#34;in/del&#34; is formed. The exact nature of this edit is difficult to predict but will almost certainly disrupt the HinfI site. Further below is the donor oligo sequence that changes two base pairs to turn the HinfI site into an EcoRI recognition site. (B) Representative restriction fragment length polymorphism (RFLP) results after editing. NHEJ (top) and HDR (bottom) editing examples are shown. This figure has been modified with permission from Modzelewski et al.9. This figure has been modified with permission from Modzelewski et al.9. <a href="https://www-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/64302/64302fig06large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Table 1: Phenotype and viability of Tyr-edited zygotes and mice. This table has been adapted with permission from Modzelewski et al.9. <a href="https://www-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/64302/Table 1.xlsx" target="_blank">Please click here to download this Table.</a>
Table 2: Results of CRISPR-EZ and microinjection deletion experiments. This table has been adapted with permission from Modzelewski et al.9. <a href="https://www-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/64302/Table 2.xlsx" target="_blank">Please click here to download this Table.</a>
Supplementary Table 1: List of oligos and donor ssODN. This table has been adapted with permission from Modzelewski et al.9. <a href="https://www-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/64302/Supplementary Table 1.xlsx" target="_blank">Please click here to download this File.</a>
Supplementary Table 2: DNA template for the sgRNA reaction setup. This table has been adapted with permission from Modzelewski et al.9. <a href="https://www-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/64302/Supplementary Table 2.xlsx" target="_blank">Please click here to download this File.</a>
Supplementary Table 3: In vitro transcription setup. This table has been modified with permission from Modzelewski et al.9. <a href="https://www-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/64302/Supplementary Table 3.xlsx" target="_blank">Please click here to download this File.</a>
Supplementary Table 4: RNP complex setup for NHEJ formation. <a href="https://www-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/64302/Supplementary Table 4.xlsx" target="_blank">Please click here to download this File.</a>
Supplementary Table 5: RNP complex setup for HDR formation. This table has been adapted with permission from Modzelewski et al.9. <a href="https://www-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/64302/Supplementary Table 5.xlsx" target="_blank">Please click here to download this File.</a>
Supplementary Table 6: RNP complex setup for deletion. This table has been modified with permission from Modzelewski et al.9. <a href="https://www-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/64302/Supplementary Table 6.xlsx" target="_blank">Please click here to download this File.</a>
Supplementary Table 7: Genotyping PCR setup. This table has been modified with permission from Modzelewski et al.9. <a href="https://www-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/64302/Supplementary Table 7.xlsx" target="_blank">Please click here to download this File.</a>
Supplementary Table 8: Nested genotyping PCR setup. This table has been modified with permission from Modzelewski et al.9. <a href="https://www-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/64302/Supplementary Table 8.xlsx" target="_blank">Please click here to download this File.</a>
Supplementary Table 9: HinfI restriction digest setup. <a href="https://www-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/64302/Supplementary Table 9.xlsx" target="_blank">Please click here to download this File.</a>
Supplementary Table 10: EcoRi restriction digest setup. <a href="https://www-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/64302/Supplementary Table 10.xlsx" target="_blank">Please click here to download this File.</a>

Watch this Scientific Journal Video about Efficient Genome Editing of Mice by CRISPR Electroporation of Zygotes at JoVE.com

Efficient Genome Editing of Mice by CRISPR Electroporation of Zygotes

Here, we describe a simple technique intended for the efficient generation of genetically modified mice called CRISPR RNP Electroporation of Zygotes (CRISPR-EZ). This method delivers editing reagents by electroporation into embryos at an efficiency approaching 100%. This protocol is effective for point mutations, small genomic insertions, and deletions in mammalian embryos.

With exceptional efficiency, accuracy, and ease, the CRISPR/Cas9 system has significantly improved genome editing in cell culture and lab animal ...

efficient-genome-editing-of-mice-by-crispr-electroporation-of-zygotes

Nanyang Technological University

Research

JoVE Journal

Developmental Biology

2.9K Views.  The University of Pennsylvania. Here, we describe a simple technique intended for the efficient generation of genetically modified mice called CRISPR RNP Electroporation of Zygotes (CRISPR-EZ). This method delivers editing reagents by electroporation into embryos at an efficiency approaching 100%. This protocol is effective for point mutations, small genomic insertions, and deletions in mammalian embryos.

Video: Efficient Genome Editing of Mice by CRISPR Electroporation of Zygotes

CRISPR/Cas9-mediated Targeted Integration In Vivo Using a Homology-mediated End Joining-based Strategy

As a promising genome editing platform, the CRISPR/Cas9 system has great potential for efficient genetic manipulation, especially for targeted integration of transgenes. However, due to the low efficiency of homologous recombination (HR) and various indel mutations of non-homologous end joining (NHEJ)-based strategies in non-dividing cells, in vivo genome editing remains a great challenge. Here, we describe a homology-mediated end joining (HMEJ)-based CRISPR/Cas9 system for efficient in vivo precise targeted integration. In this system, the targeted genome and the donor vector containing homology arms (~800 bp) flanked by single guide RNA (sgRNA) target sequences are cleaved by CRISPR/Cas9. This HMEJ-based strategy achieves efficient transgene integration in mouse zygotes, as well as in hepatocytes in vivo. Moreover, a HMEJ-based strategy offers an efficient approach for correction of fumarylacetoacetate hydrolase (Fah) mutation in the hepatocytes and rescues Fah-deficiency induced liver failure mice. Taken together, focusing on targeted integration, this HMEJ-based strategy provides a promising tool for a variety of applications, including generation of genetically modified animal models and targeted gene therapies.

CRISPR/Cas9-mediated Targeted Integration In Vivo Using a Homology-mediated End Joining-based Strategy

The clustered regularly interspaced short palindromic repeats/CRISPR associated protein 9 (CRISPR/Cas9) system provides a promising tool for genetic engineering, and opens up the possibility of targeted integration of transgenes. We describe a homology-mediated end joining (HMEJ)-based strategy for efficient DNA targeted integration in vivo and targeted gene therapies using CRISPR/Cas9.

As a promising genome editing platform, the CRISPR/Cas9 system has great potential for efficient genetic manipulation, especially for targeted integration ...

Genetics

Enhanced Genome Editing with Cas9 Ribonucleoprotein in Diverse Cells and Organisms

Site-specific eukaryotic genome editing with CRISPR (clustered regularly interspaced short palindromic repeats)-Cas (CRISPR-associated) systems has quickly become a commonplace amongst researchers pursuing a wide variety of biological questions. Users most often employ the Cas9 protein derived from Streptococcus pyogenes in a complex with an easily reprogrammed guide RNA (gRNA). These components are introduced into cells, and through a base pairing with a complementary region of the double-stranded DNA (dsDNA) genome, the enzyme cleaves both strands to generate a double-strand break (DSB). Subsequent repair leads to either random insertion or deletion events (indels) or the incorporation of experimenter-provided DNA at the site of the break.
The use of a purified single-guide RNA and Cas9 protein, preassembled to form an RNP and delivered directly to cells, is a potent approach for achieving highly efficient gene editing. RNP editing particularly enhances the rate of gene insertion, an outcome that is often challenging to achieve. Compared to the delivery via a plasmid, the shorter persistence of the Cas9 RNP within the cell leads to fewer off-target events. 
Despite its advantages, many casual users of CRISPR gene editing are less familiar with this technique. To lower the barrier to entry, we outline detailed protocols for implementing the RNP strategy in a range of contexts, highlighting its distinct benefits and diverse applications. We cover editing in two types of primary human cells, T cells and hematopoietic stem/progenitor cells (HSPCs). We also show how Cas9 RNP editing enables the facile genetic manipulation of entire organisms, including the classic model roundworm Caenorhabditis elegans and the more recently introduced model crustacean, Parhyale hawaiensis.

Utilizing a preassembled Cas9 ribonucleoprotein complex (RNP) is a powerful method for precise, efficient genome editing. Here, we highlight its utility across a broad range of cells and organisms, including primary human cells and both classic and emerging model organisms.

Site-specific eukaryotic genome editing with CRISPR (clustered regularly interspaced short palindromic repeats)-Cas (CRISPR-associated) systems has ...

A Rapid and Facile Pipeline for Generating Genomic Point Mutants in C. elegans Using CRISPR/Cas9 Ribonucleoproteins

The clustered regularly interspersed palindromic repeats (CRISPR)-CRISPR-associated protein 9 (Cas9) prokaryotic adaptive immune defense system has been co-opted as a powerful tool for precise eukaryotic genome engineering. Here, we present a rapid and simple method using chimeric single guide RNAs (sgRNA) and CRISPR-Cas9 Ribonucleoproteins (RNPs) for the efficient and precise generation of genomic point mutations in C. elegans. We describe a pipeline for sgRNA target selection, homology-directed repair (HDR) template design, CRISPR-Cas9-RNP complexing and delivery, and a genotyping strategy that enables the robust and rapid identification of correctly edited animals. Our approach not only permits the facile generation and identification of desired genomic point mutant animals, but also facilitates the detection of other complex indel alleles in approximately 4 - 5 days with high efficiency and a reduced screening workload.

A Rapid and Facile Pipeline for Generating Genomic Point Mutants in C. elegans Using CRISPR/Cas9 Ribonucleoproteins

Here, we present a method to engineer the genome of C. elegans using CRISPR-Cas9 ribonucleoproteins and homology dependent repair templates.

The clustered regularly interspersed palindromic repeats (CRISPR)-CRISPR-associated protein 9 (Cas9) prokaryotic adaptive immune defense system has been ...

Generation of Genetically Modified Mice through the Microinjection of Oocytes

The use of genetically modified mice has significantly contributed to studies on both physiological and pathological in vivo processes. The pronuclear injection of DNA expression constructs into fertilized oocytes remains the most commonly used technique to generate transgenic mice for overexpression. With the introduction of CRISPR technology for gene targeting, pronuclear injection into fertilized oocytes has been extended to the generation of both knockout and knockin mice. This work describes the preparation of DNA for injection and the generation of CRISPR guides for gene targeting, with a particular emphasis on quality control. The genotyping procedures required for the identification of potential founders are critical. Innovative genotyping strategies that take advantage of the "multiplexing" capabilities of CRISPR are presented herein. Surgical procedures are also outlined. Together, the steps of the protocol will allow for the generation of genetically modified mice and for the subsequent establishment of mouse colonies for a plethora of research fields, including immunology, neuroscience, cancer, physiology, development, and others.

The microinjection of mouse oocytes is commonly used for both classic transgenesis (i.e., the random integration of transgenes) and CRISPR-mediated gene targeting. This protocol reviews the latest developments in microinjection, with a particular emphasis on quality control and genotyping strategies.

The use of genetically modified mice has significantly contributed to studies on both physiological and pathological in vivo processes. The pronuclear ...

Use of Freeze-thawed Embryos for High-efficiency Production of Genetically Modified Mice

The use of genetically modified (GM) mice has become crucial for understanding gene function and deciphering the underlying mechanisms of human diseases. The CRISPR/Cas9 system allows researchers to modify the genome with unprecedented efficiency, fidelity, and simplicity. Harnessing this technology, researchers are seeking a rapid, efficient, and easy protocol for generating GM mice. Here we introduce an improved method for cryopreservation of one-cell embryos that leads to a higher developmental rate of the freeze-thawed embryos. By combining it with optimized electroporation conditions, this protocol allows for the generation of knockout and knock-in mice with high efficiency and low mosaic rates within a short time. Furthermore, we show a step-by-step explanation of our optimized protocol, covering CRISPR reagent preparation, in vitro fertilization, cryopreservation and thawing of one-cell embryos, electroporation of CRISPR reagents, mouse generation, and genotyping of the founders. Using this protocol, researchers should be able to prepare GM mice with unparalleled ease, speed, and efficiency.

Here, we present a modified method for cryopreservation of one-cell embryos as well as a protocol that couples the use of freeze-thawed embryos and electroporation for the efficient generation of genetically modified mice.

The use of genetically modified (GM) mice has become crucial for understanding gene function and deciphering the underlying mechanisms of human diseases. ...

CRISPR-mediated Loss of Function Analysis in Cerebellar Granule Cells Using In Utero Electroporation-based Gene Transfer

Brain malformation is often caused by genetic mutations. Deciphering the mutations in patient-derived tissues has identified potential causative factors of the diseases. To validate the contribution of a dysfunction of the mutated genes to disease development, the generation of animal models carrying the mutations is one obvious approach. While germline genetically engineered mouse models (GEMMs) are popular biological tools and exhibit reproducible results, it is restricted by time and costs. Meanwhile, non-germline GEMMs often enable exploring gene function in a more feasible manner. Since some brain diseases (e.g., brain tumors) appear to result from somatic but not germline mutations, non-germline chimeric mouse models, in which normal and abnormal cells coexist, could be helpful for disease-relevant analysis. In this study, we report a method for the induction of CRISPR-mediated somatic mutations in the cerebellum. Specifically, we utilized conditional knock-in mice, in which Cas9 and GFP are chronically activated by the CAG (CMV enhancer/chicken &#223;-actin) promoter after Cre-mediated recombination of the genome. The self-designed single-guide RNAs (sgRNAs) and the Cre recombinase sequence, both encoded in a single plasmid construct, were delivered into cerebellar stem/progenitor cells at an embryonic stage using in utero electroporation. Consequently, transfected cells and their daughter cells were labeled with green fluorescent protein (GFP), thus facilitating further phenotypic analyses. Hence, this method is not only showing electroporation-based gene delivery into embryonic cerebellar cells but also proposing a novel quantitative approach to assess CRISPR-mediated loss-of-function phenotypes.

CRISPR-mediated Loss of Function Analysis in Cerebellar Granule Cells Using In Utero Electroporation-based Gene Transfer

Conventional loss-of-function studies of genes using knockout animals have often been costly and time-consuming. Electroporation-based CRISPR-mediated somatic mutagenesis is a powerful tool to understand gene functions in vivo. Here, we report a method to analyze knockout phenotypes in proliferating cells of the cerebellum.

Brain malformation is often caused by genetic mutations. Deciphering the mutations in patient-derived tissues has identified potential causative factors ...

CRISPR-Cas9 Genome Editing of Rat Embryos using Adeno-Associated Virus (AAV) and 2-Cell Embryo Electroporation

Genome editing technology is widely used to produce genetically modified animals, including rats. Cytoplasmic or pronuclear injection of DNA repair templates and CRISPR-Cas reagents is the most common delivery method into embryos. However, this type of micromanipulation necessitates access to specialized equipment, is laborious, and requires a certain level of technical skill. Moreover, microinjection techniques often result in lower embryo survival due to the mechanical stress on the embryo. In this protocol, we developed an optimized method to deliver large DNA repair templates to work in conjunction with CRISPR-Cas9 genome editing without the need for microinjection. This protocol combines AAV-mediated DNA delivery of single-stranded DNA donor templates along with the delivery of CRISPR-Cas9 ribonucleoprotein (RNP) by electroporation to modify 2-cell embryos. Using this novel strategy, we have successfully produced targeted knock-in rat models carrying insertion of DNA sequences from 1.2 to 3.0 kb in size with efficiencies between 42% and 90%.

CRISPR-Cas9 Genome Editing of Rat Embryos using Adeno-Associated Virus AAV and 2-Cell Embryo Electroporation

This protocol presents an optimized approach for producing genetically modified rat models. Adeno-associated virus (AAV) is used to deliver a DNA repair template, and electroporation is used to deliver CRISPR-Cas9 reagents to complete the genome editing process in 2-cell embryo.

Genome editing technology is widely used to produce genetically modified animals, including rats. Cytoplasmic or pronuclear injection of DNA repair ...

Somatic Genome-Engineered Mouse Models Using In Vivo Microinjection and Electroporation

Germline genetically engineered mouse models (G-GEMMs) have provided valuable insight into in vivo gene function in development, homeostasis, and disease. However, the time and cost associated with colony creation and maintenance are high. Recent advances in CRISPR-mediated genome editing have allowed the generation of somatic GEMMs (S-GEMMs) by directly targeting the cell/tissue/organ of interest.
The oviduct, or fallopian tube in humans, is considered the tissue-of-origin of the most common ovarian cancer, high-grade serous ovarian carcinomas (HGSCs). HGSCs initiate in the region of the fallopian tube distal to the uterus, located adjacent to the ovary, but not the proximal fallopian tube. However, traditional mouse models of HGSC target the entire oviduct, and thus do not recapitulate the human condition. We present a method of DNA, RNA, or ribonucleoprotein (RNP) solution microinjection into the oviduct lumen and in vivo electroporation to target mucosal epithelial cells in restricted regions along the oviduct. There are several advantages of this method for cancer modeling, such as 1) high adaptability in targeting the area/tissue/organ and region of electroporation, 2) high flexibility in targeted cell types (cellular pliancy) when used in combination with specific promoters for Cas9 expression, 3) high flexibility in the number of electroporated cells (relatively low frequency), 4) no specific mouse line is required (immunocompetent disease modeling), 5) high flexibility in gene mutation combination, and 6) possibility of tracking electroporated cells when used in combination with a Cre reporter line. Thus, this cost-effective method recapitulates human cancer initiation.

Somatic Genome-Engineered Mouse Models Using In Vivo Microinjection and Electroporation

This protocol describes microinjection and in vivo electroporation for regionally restricted CRISPR-mediated genome editing in the mouse oviduct.

Germline genetically engineered mouse models (G-GEMMs) have provided valuable insight into in vivo gene function in development, homeostasis, and disease. ...

Cancer Research

Zygote Microinjection for Creating Gene Cassette Knock-in and Flox Alleles in Mice

CRISPR-Cas technology has enabled the rapid and effortless generation of genetically modified mice. Specifically, mice and point mutant mice are readily produced by electroporation of CRISPR factors (and single-stranded oligo DNA donors) into the zygote. In contrast, gene cassette (&#62;1 kb) knock-in and floxed mice are mainly generated by microinjection of CRISPR factors and double-stranded DNA donors into zygotes. Genome editing technologies have also increased the flexibility of genetically modified mice production. It is now possible to introduce the intended mutations in the target genomic regions in a number of beneficial inbred mouse strains. Our team has produced over 200 gene cassette knock-in mouse lines, and over 110 floxed mouse lines by zygote microinjection of CRISPR-Cas9 following requests from several countries, including Japan. Some of these genome editing used BALB/c, C3H/HeJ, and C57BL/6N inbred strains, however most used C57BL/6J. Unlike the electroporation method, genome editing by zygote microinjection in various inbred strains of mice is not that easy. However, gene cassette knock-in and floxed mice on single inbred genetic backgrounds are as critical as genetic humanized, fluorescent reporter, and conditional knockout mouse models. Therefore, this article presents the protocol for the zygote microinjection of CRISPR factors and double-stranded DNA donors in C57BL/6J mice for generating gene cassette knock-in and floxed mice. This article exclusively focuses on nuclear injection rather than cytoplasmic injection. In addition to zygote microinjection, we outline the timeline for the production process and peripheral techniques such as induction of superovulation and embryo transfer.

The present protocol describes zygote microinjection of CRISPR-Cas9 and donor DNA to efficiently produce gene cassette knock-in and floxed mice.

CRISPR-Cas technology has enabled the rapid and effortless generation of genetically modified mice. Specifically, mice and point mutant mice are readily ...

CRISPR/Cas9-Mediated Highly Efficient Gene Targeting in Embryonic Stem Cells for Developing Gene-Manipulated Mouse Models

The CRISPR/Cas9 system has made it possible to develop genetically modified mice by direct genome editing using fertilized zygotes. However, although the efficiency in developing gene-knockout mice by inducing small indel mutation would be sufficient enough, the efficiency of embryo genome editing for making large-size DNA knock-in (KI) is still low. Therefore, in contrast to the direct KI method in embryos, gene targeting using embryonic stem cells (ESCs) followed by embryo injection to develop chimera mice still has several advantages (e.g., high throughput targeting in vitro, multi-allele manipulation, and Cre and flox gene manipulation can be carried out in a short period). In addition, strains with difficult-to-handle embryos in vitro, such as BALB/c, can also be used for ESC targeting. This protocol describes the optimized method for large-size DNA (several kb) KI in ESCs by applying CRISPR/Cas9-mediated genome editing followed by chimera mice production to develop gene-manipulated mouse models.

Here we present a protocol for developing genetically modified mouse models using embryonic stem cells, especially for large DNA knock-in (KI). This protocol is tuned up using CRISPR/Cas9 genome editing, resulting in significantly improved KI efficiency compared with the conventional homologous recombination-mediated linearized DNA targeting method.

The CRISPR/Cas9 system has made it possible to develop genetically modified mice by direct genome editing using fertilized zygotes. However, although the ...

Efficient Genome Editing of Mice by CRISPR Electroporation of Zygotes

Summary

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Chapters in this video

CRISPR/Cas9-mediated Targeted Integration In Vivo Using a Homology-mediated End Joining-based Strategy

Enhanced Genome Editing with Cas9 Ribonucleoprotein in Diverse Cells and Organisms

A Rapid and Facile Pipeline for Generating Genomic Point Mutants in C. elegans Using CRISPR/Cas9 Ribonucleoproteins

Generation of Genetically Modified Mice through the Microinjection of Oocytes

Use of Freeze-thawed Embryos for High-efficiency Production of Genetically Modified Mice

CRISPR-mediated Loss of Function Analysis in Cerebellar Granule Cells Using In Utero Electroporation-based Gene Transfer

CRISPR-Cas9 Genome Editing of Rat Embryos using Adeno-Associated Virus (AAV) and 2-Cell Embryo Electroporation

Somatic Genome-Engineered Mouse Models Using In Vivo Microinjection and Electroporation

Zygote Microinjection for Creating Gene Cassette Knock-in and Flox Alleles in Mice

CRISPR/Cas9-Mediated Highly Efficient Gene Targeting in Embryonic Stem Cells for Developing Gene-Manipulated Mouse Models