Efficient Genome Editing of Mice by CRISPR Electroporation of Zygotes

Summary

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.

Abstract

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.

Introduction

Genome editing in live mice has been considerably simplified and has become accessible and more affordable since the emergence of CRISPR editing^1,2,3. Initial animal editing attempts used microinjection to deliver CRISPR Cas9 mRNA/sgRNA into pronuclear-stage embryos^4,5,6. 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'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 step^{7,10,11,12,13,14,15}, 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/6N^9,16. Independent groups have also developed electroporation-based procedures with efficiencies greater than or matching microinjection^{11,12,13,14,15,17}, with electroporation protocols functioning well in rat^18,19, pig^20,21,22 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 delivery^9,16,25. 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) locus^9,16 have been included in the Supplementary Table 1.

Protocol

All animal care and use throughout this protocol adhered to Animal Welfare Act policies the ILAR Guide for Care and Use of Laboratory Animals and followed guidelines from the AVMA for euthanasia and the University of Pennsylvania Institutional Animal Care and Use Committee (IACUC) guidelines and policies. The animal care and use protocol was reviewed and approved by the University of Pennsylvania IACUC for this project. As a matter of compliance and caution, please seek out all necessary authorizations prior to attempting this protocol.

1. sgRNA and optional donor oligo design

1. sgRNA design
   1. Select candidate sgRNAs from any widely used online algorithms, such as Sequence Scan for CRISPR (SSC; http://crispr.dfci.harvard.edu/SSC/)²⁸ or CRISPick (https://portals.broadinstitute.org/gppx/crispick/public)²⁹. As each platform is unique, please carefully read the user instructions in order to design ideal sgRNAs. For in/del experiments, select two to three sgRNAs to test. For deletions, sgRNAs that flank the target region are suggested, for example, two sgRNAs upstream of the target and two sgRNAs downstream of the target.
      NOTE: While various online sgRNA algorithms factor in off-targets to avoid flawed sgRNA designs, NCBI's BLAST tool is helpful in confirming the quality of the selected sgRNA sequences.
   2. Insert 19-20 nucleotides (nt) determined from the previous step (1.1.1) into the variable region of the sgRNA oligo (Supplementary Table 1) and purchase synthesized sgRNA as custom DNA oligos.
      NOTE: PAGE purification is not required.
2. (Optional) Donor oligo design for homology directed repair (HDR)-mediated knock-in.
   1. Design the appropriate single-stranded oligodeoxynucleotide (ssODN) based on the desired experimental outcome (precise point mutation, loxP insertion, small tag insertion, etc.), keeping the total length to 100-200 nt with homology arms of 50 nt or more flanking the central region.
   2. To ensure higher knock-in efficiency, design the sgRNA to be complementary to the ssODN³⁰, and replace the protospacer adjacent motif (PAM) sequence with a silent mutation to prevent recurring cutting by the Cas9 enzyme.
   3. Order ssODN using the custom oligo option.

2. sgRNA synthesis

1. Template generation for in vitro transcription (IVT)
   1. To generate the DNA template for sgRNA IVT through PCR, prepare an IVT reaction in a PCR strip tube that is RNAse-free. (see Supplementary Table 2).
   2. Use the following thermal cycler conditions:
      95 °C for 2 min; 30 cycles of 95 °C for 2 min, 57 °C for 10 s, and 72 °C for 10 s; then 72 °C for 2 min
   3. To verify the success of the sgRNA DNA template PCR reaction, electrophorese 5 µL of the product combined with 1 µL of 6x loading dye on a 2% (wt/vol) agarose gel.
      NOTE: The anticipated product size is 127 bp. Any remaining PCR reaction can be stored at −20 °C for up to 5 months.
2. In vitro transcription (IVT) of sgRNA
   1. To generate the sgRNA, prepare the reaction mixture (T7 IVT as per manufacturer instructions) in a PCR tube and flick to mix the reagents.
      NOTE: See Supplementary Table 3 for the reaction setup example. The IVT reaction is RNase contamination sensitive; therefore, make every effort to provide an RNase-free environment.
   2. To allow the reaction to initiate and proceed, incubate the IVT reaction mixture for >18 h at 37 °C in either a thermal cycler or heat block.
   3. To degrade the original DNA template (step 2.1.3), introduce 1 µL of DNase I (2 units per reaction) and incubate the reaction for 20 min at room temperature (RT).
   4. To promote RNA binding to the magnetic purification beads, combine 129 µL of 100% ethanol in the reaction, which will now be 150 µL.
   5. To resuspend the purification beads, which tend to settle during storage, vortex the aliquot for at least 10 s.
   6. To purify the sgRNAs, add 100 µL of the resuspended beads (step 2.2.5) to the IVT reaction (step 2.2.4) and gently mix by pipetting up and down 10x.
   7. To allow for binding, allow the mixture to rest at RT for 5 min.
   8. To isolate the sgRNA from the unincorporated reaction products, place the reaction on the magnetic stands for 5 min at RT and wait until a small pellet forms.
   9. To wash the sgRNAs, first carefully discard the supernatant and then add 200 µL of 80% ethanol away from the pellet.
      NOTE: During the 80% (vol/vol) ethanol wash step, it is critical to avoid disturbing the pellet by pipetting, as this will reduce the sgRNA concentrations considerably. Instead, gently pipette the 80% ethanol so the pellet is submerged, and gently remove the volume with a pipette.
   10. Repeat the previous step (step 2.2.9) and air dry the pellet for no more than 5-6 min or the sgRNA will be permanently associated with the beads.
   11. Elute the sgRNA with 20µL of nuclease-free water by pipetting the pellet 10x, incubating for 2 min at (RT), and then placing it back on the magnetic stand to separate the beads from the now purified sgRNA.
   12. To evaluate the quality and quantity of the sgRNA, use an instrument like a spectrophotometer or bioanalyzer. Alternatively, on a 2% agarose gel, run 2 µL of sgRNAs, similar to step 2.1.3.
       ​NOTE: Quality is confirmed by a single clear band. The remainder of the sgRNA can either be used immediately or stored in −80 °C conditions.

3. Superovulation

1. To provide enough embryos to test each sgRNA design, superovulate two to three C57BL/6J females that are between 3-5 weeks old, as previously described⁹. Electroporating 20-30 embryos per sgRNA is recommended to generate enough results to confirm effectiveness.
   NOTE: If fertilization is successful, expect 10-20 embryos per mating.
2. To induce ovulation, follow this hormone injection schedule:
   1. Day 1: Administer 5 IU of pregnant mare serum gonadotropin (PMSG) (100 µL) through an intraperitoneal (IP) injection using a 26 G syringe into 3-5-week-old female mice.
   2. Day 3: After 46-48 h of PMSG injection, inject 5 IU of human chorionic gonadotropin (hCG) (100 µL) through an IP injection to induce ovulation.
   3. To set up breeding, pair females 1:1 with a male of reliable breeding right after hCG injection.
      ​NOTE: To prevent hormones from degrading and losing activity, perform injections under 30 min of thawing.

4. Embryo collection and processing

1. Embryo collection
   1. To euthanize females and retrieve the necessary material, as previously described³¹, be sure to first confirm the appropriate method in accordance with institutional policies. For example, use CO₂ asphyxiation, cervical dislocation, and/or other approved methods.
      NOTE: Normally, >75% of hormone-stimulated females display copulatory plugs, indicating successful mating.
   2. To prevent hairs from making tissue collection difficult, place the euthanized females on their backs and spray 70% (vol/vol) ethanol on the abdomen region to be surgically opened.
      NOTE: From this point on, it is advised to perform the surgical steps in a ventilated hood in order to maintain an aseptic environment and reduce the potential for contamination.
   3. To open the abdominal cavity and remove the oviducts, cut the skin/hair (subcutaneous) layer with surgical scissors, and locate the ovaries (Figure 4), which are found near the kidney and share a section of a fat pad. The oviducts are located proximal to the ovaries (Figure 4). Surgically remove both oviducts from the female and place them into individual 50 µL droplets of M2+BSA (4 mg/mL BSA) medium (Figure 5A).
      NOTE: Detailed instructions for this section of the procedure can be viewed visually³¹ or followed stepwise⁹ using previously published protocols.
   4. To release the cumulus-oocyte complexes (CoCs) containing oocytes (Figure 4), use dissection forceps to nick the ampulla of the oviduct while viewing through a stereomicroscope.
   5. To reduce the amount of debris (blood, fat, tissue), transfer and combine each CoC to a single 50 µL droplet of M2+BSA media with a handheld pipette set to 20-30 µL to avoid the carryover of debris.
2. Removal of cumulus cells
   1. To begin the removal of cumulus cells from the zygotes (Figure 4), transfer CoCs with as little extra media as possible to a droplet of 100 µL M2+hyaluronidase (Figure 5B), and gently mix by pipetting the CoCs until the embryos are visibly separated from the much smaller cumulus cells. Be sure to keep the exposure of M2+hyaluronidase to the embryos to a minimum, as it might impact the viability.
      NOTE: One can either preheat (37 °C) or add an additional 50 µL of M2+hyaluronidase if the embryos are not separating from the cumulus cells within 2 min.
   2. To dilute hyaluronidase and clear away loose cumulus cells from the zygotes, use a mouth-operated pipette to move the zygotes to a fresh 50 µL M2+BSA droplet.
      NOTE: Most steps beyond this point require the use of a mouth-operated pipette unless otherwise noted. While the risk of contamination from the user is low, the use of an inline air filter is recommended to maintain an aseptic environment. Please refer to previously described methods to prepare and use this instrument^9,31.
   3. Using a mouth pipette, pass the embryos through at least four to five more 50 µL M2+BSA droplets to reduce the number of cumulus cells.
3. (OPTIONAL) Thinning of the zona pellucida with acid tyrode (AT) solution.
   1. To erode the zona pellucida of the embryo and reduce additional physical obstacles for reagent delivery, transfer the embryos to a prewarmed (37 °C) 100 µL droplet of AT solution on a 60 mm plate (Figure 5C). Continuously observe the zygotes using a stereomicroscope until approximately 30% of the zona pellucida is eroded⁹. The total AT exposure must be 60-90 s. Prolonged exposure to AT can entirely dissolve the zona pellucida and jeopardize embryo viability.
      NOTE: For detailed instructions and images of this section of the procedure, please refer to previously published methods^9,16.
   2. To dilute the AT, use a mouth pipette to pass the embryos through at least four 50 µL droplets of M2+BSA.
   3. To determine whether an appropriate number of embryos have been processed, use a stereomicroscope to count the embryos. Depending on the number of female mice used, one can expect approximately 10-20 viable zygotes per female.
      ​NOTE: At this stage, embryos should be relatively free from debris that would prevent the evaluation of quantity and quality. For example, if using five females, the expected zygote numbers would likely be between 50-100, and a mechanical cell counter would be appropriate to keep track of embryos. It is common to lose about 10%-20% of embryos due to failure to fertilize or low-quality oocytes being ovulated. During the next step, briefly store the embryos in 50 µL of M2+BSA for no more than 30 min in an incubator.

5. RNP assembly and electroporation

1. RNP assembly
   1. To assemble the RNP complex, use the attached example tables to combine the appropriate reaction mixtures based on the desired editing strategy in RNP buffer (100 mM HEPES pH 7.5, 750 mM KCl, 5 mM MgCl₂, 50% glycerol, and 100 mM Tris(2-carboxyethyl)phosphine hydrochloride [TCEP] in nuclease-free water)
      NOTE: TCEP is a convenient reducing agent but has a short half-life in aqueous solutions; therefore, add TCEP just before RNP complex assembly.
      1. For non-homologous end joining (NHEJ)-mediated indels, refer to Supplementary Table 4.
      2. For homology directed repair (HDR)-mediated editing, refer to Supplementary Table 5.
      3. For engineering deletions using paired sgRNAs, refer to Supplementary Table 6.
         1. Regardless of strategy, combine the desired reagents at RT in a PCR tube.
         2. To allow RNP complex formation, incubate the mixture for 10 min at 37 °C.
            NOTE: RNP complex can be stored at RT for up to 1 h.
2. Electroporation
   1. To dilute the BSA before electroporation, pass the embryos through at least two droplets of 50 µL of reduced serum media.
      NOTE: BSA increases resistance during electroporation and could damage the zygotes.
   2. To prepare and mix the zygotes (step 4.3.2) with the RNP complex (step 5.1.1.2) for electroporation, mouth pipette 25-30 embryos into a 10 µL droplet of reduced serum media, and then use a handheld pipette to add 10 µL of the RNP complex (step 5.1.1.2).
   3. To dilute the viscous glycerol from the RNP complex and homogenize the sample, mix by pipetting 10x or until the mixture no longer shows signs of viscosity (swirl pattern) by using a stereomicroscope.
   4. To prepare the sample for insertion into the electroporator, pipette this 20 µL of embryo + RNP mixture into a 0.1 cm electroporation cuvette while avoiding bubbles.
   5. To deliver the editing reagents into the zygote, electroporate the embryos using a square-wave protocol with these conditions: 30 V, 4-6 pulses, 3 ms pulse length, and 100 pulse intervals.
   6. To retrieve zygotes from the cuvette, use a hand pipette to deliver 50 µL of KSOM+BSA (1mg/mL BSA) into the top of the cuvette to flush the embryos that may have settled to the bottom. Gently pipette this mixture out and onto a 60 mm plate.
   7. Repeat step 5.2.6 at least 2x-3x and combine each flush on a 60 mm plate until most embryos are recovered.
      ​NOTE: A typical recovery is >90% of the originally loaded embryos. To ensure embryo survival, test the incubator and check the expiration dates for all reagents. Embryos are susceptible to non-ideal culture conditions such as temperature, CO₂ level, humidity, and pH.

6. Embryo culture and genotyping

1. Embryo culture
   1. To culture zygotes to a desired stage, use a mouth pipette to transfer 20-30 zygotes into a droplet of KSOM+BSA in a pre-equilibrated culture plate and move the embryos to the droplet (Figure 5D). Incubate the plate overnight in 5% CO₂, at 37 °C, with 95% humidity.
      NOTE: Pre-equilibrate by placing a prepared 35mm culture plate in an incubator either the day prior or at least 4 h before incubation (Figure 5D).
   2. To promote ideal growth conditions, transfer only two-cell embryos on the following day to a fresh droplet of KSOM+BSA mixture using a stereomicroscope and return to the incubator. Be sure to leave or discard the dead or unfertilized embryos.
      NOTE: Pronuclear-stage embryos typically grow into morulae after 72 h of culture and blastocysts after 84 h.
2. Genotyping
   1. To dilute away the components of the media that might interfere with a PCR reaction, wash the morula/blastocyst embryos using a mouth pipette by passing through two drops of DPBS.
   2. To load single embryos for lysis, transfer individual embryos to one well of an 8-well PCR strip by setting a pipette to 1 µL, and then add 10 µL of lysis buffer (50 mM KCl, 10 mM Tris-HCl ph 8.5, 2.5 mM MgCl₂, 0.1 mg/mL gelatin, 0.45% Nonidet P-40, 0.45% Tween 20, 0.2 mg/mL proteinase K in nuclease-free H₂O).
      NOTE: Add proteinase K just before preparing the lysis buffer.
   3. To lyse the embryos, program a thermal cycler to 55 °C for 4 h, and then inactivate the proteinase K with a 10 min incubation at 95 °C.
      NOTE: Particularly for first-time users, attempt an editing experiment at the Tyr loci. This workflow has been optimized and can serve as a positive control. If necessary, store the embryo lysates at 4 °C for 2-3 days or in the freezer for up to 2 weeks before PCR analysis. Prevent repeated freeze-thaw cycles.
   4. To increase the chances of successful genotyping analysis, perform a nested PCR strategy by amplifying slightly longer DNA fragments that flank the targeted site as well as regions that will be used to amplify a more precise DNA fragment. Follow the conditions in Supplementary Table 7.
   5. Follow the below-mentioned thermal cycler conditions:
      95 °C for 2 min
      30+ cycles: 95 °C for 2 min, 60 °C for 10 s, and 72 °C for 10 s
      ​72 °C for 2 min
   6. To prepare the second stage of a nested PCR strategy, make a 1:10 dilution of the first PCR product (step 6.2.5) and load 2 µL of this into the next PCR reaction (with primers designed to be inside the previous amplicon).
      NOTE: The first PCR reaction must be diluted to reduce the carryover of flanking primers in the second PCR reaction. Prepare the nested PCR by following the steps in Supplementary Table 8.
   7. Place the PCR reaction in a thermal cycler using the following cycling conditions:
      95 °C for 2 min
      30 cycles: 95 °C for 2 min, 60 °C for 10 s, and 72 °C for 10 s
      72 °C for 2 min
      NOTE: Normally, >90% of PCR reactions are successful when the amplicon size is 500 bp or less.
   8. (Optional control) For NHEJ-mediated in/del mutations using Tyr as a positive control, digest 10 µL of the nested PCR products from step 6.2.7 by incubating at 37 °C for 4 h using 10 U of HinfI in a 20 µL reaction (see Supplementary Table 9). On a 2% agarose gel, run the digested product to evaluate editing and use a nondigested PCR product as a loading control. Run the gel at 135 V for 30 min.
      NOTE: The use of HinfI as an appropriate restriction enzyme was selected due to a conveniently located HinfI site present within the sgRNA target region that is predicted to be ablated upon a successful NHEJ editing. A detailed method and results have been previously described^9,16.
   9. (Optional control) For HDR-mediated mutations using Tyr as a positive control, digest 10 µL of the nested PCR products from step 6.2.7 by incubating at 37 °C for 4 h using 10 U of EcoRI in a 20 µL reaction (see Supplementary Table 10). On a 2% agarose gel, run the digested product to evaluate editing and use a nondigested PCR product as a loading control. Run the gel at 135 V for 30 min.
      NOTE: The choice of EcoRI as a diagnostic restriction enzyme takes advantage of the original sequence found at the sgRNA target region, where, upon successful HDR, the naturally occurring HinfI site is replaced with an EcoRI site, and a frameshift mutation is forced that interferes with the Tyr gene. For HDR analysis, perform both NHEJ (step 6.2.8) and HDR (step 6.2.9) analyses on each sample as both outcomes can occur and can help determine mosaicism. A detailed method and results have been previously described^9,16.

Results

This method generates more than 100 µg of sgRNA (20 µL at >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 mo...

Discussion

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 RNPs^7,10,11,12,

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
0.1-cm-gap electroporation cuvette	Bio-Rad	cat. no. 1652089	Electroporation
26-G, 1/2-inch needle	BD	cat. no. 305111	Superovulation
3–8-month-old male mice and 3- to 5-week-old female mice	JAX	cat. no. 000664	Superovulation
35-mm Tissue culture dish	Greiner Bio-One,	cat. no. 627-160	Embryo Culture
60-mm Tissue culture dish	Greiner Bio-One,	cat. no. 628-160	Embryo Processing
6x loading dye	Thermo Fisher Scientific	cat. no. R0611	sgRNA Synthesis and Genotyping
Acidic Tyrode's (AT) solution, embryo culture grade	Sigma-Aldrich,	cat. no. T1788	Embryo Processing
BSA, embryo culture grade	Sigma-Aldrich	cat. no. A3311	Embryo Processing and Culture
Cas9 protein	Alt-R S.p. Cas9 nuclease 3NLS	cat. no. 1074181	Electroporation
DNase I, RNase-free	New England BioLabs,	cat. no. M0303	sgRNA Synthesis
DPBS(calcium and magnesium free)	Gibco	cat. no. 14190-144	Embryo Processing
EcoRI	NEB	cat. no. R3101S	Genotyping
EDTA, anhydrous	Sigma-Aldrich	cat. no. EDS-100G	RNP Buffer
Ethanol	Koptec	cat. no. V1016	sgRNA Synthesis
Gelatin (powder) type B, laboratory grade	Fisher,	cat. no. G7-500	Lysis Buffer
Glycerol, molecular-biology grade	Fisher	cat. no. BP229	RNP Buffer
Taq Polymerase	Promega	cat. no. M712	Genotyping
HEPES, cell culture grade	Sigma-Aldrich	cat. no. H4034	RNP Buffer
HinfI (10,000 U/mL)	NEB	cat. no. R0155S	Genotyping
HiScribe T7 High Yield RNA Synthesis Kit	New England BioLabs,	cat. no. E2040	sgRNA Synthesis
Human chorion gonadotropin, lyophilized (hCG)	Millipore	cat. no. 230734	Superovulation
Hyaluronidase/M2	Millipore	cat. no. MR-051-F	Embryo Processing
KSOMaa Evolve medium (potassium-supplemented simplex-optimized medium plus amino acids)	Zenith Biotech	cat. no. ZEKS-050	Embryo Culture
LE agarose, analytical grade	BioExpress	cat. no. E-3120-500	sgRNA Synthesis and Genotyping
M2 medium	Zenith Biotech	cat. no. ZFM2-050	Embryo Processing
Magnesium chloride, anhydrous (MgCl2)	Sigma-Aldrich	cat. no. M8266	RNP and Lysis Buffer
Mineral Oil	Millipore	cat. no. ES-005C	Embryo Culture
Nonidet P-40,substitute (NP-40)	Sigma-Aldrich	cat. no. 74385	Lysis Buffer
Nuclease-free water, molecular-biology grade	Ambion	cat. no. AM9937	sgRNA Synthesis and Genotyping
Oligos for sgRNA synthesis, donor oligo and PCR primers for genotyping	Integrated DNA Technologies	custom orders	sgRNA Design
Reduced serum medium	Thermo Fisher Scientific	cat. no. 31985062	Embryo Culture
High-fidelity DNA polymerase	New England BioLabs,	cat. no. M0530	sgRNA Synthesis
Potassium chloridemolecular-biology grade (KCl)	Sigma-Aldrich	cat. no. P9333	RNP and Lysis Buffer
Pregnant mare serum gonadotropin lyophilizd ((PMSG)	ProspecBio	cat. no. HOR-272	Superovulation
Proteinase K, molecular-biology grade	Fisher	cat. no. BP1700-100	Lysis Buffer
RNase-free 1.5-mL microcentrifuge tube	VWR	cat. no. 20170-333	sgRNA Synthesis and Genotyping
RNase-free eight-well PCR strip tubes	VWR	cat. no. 82006-606	sgRNA Synthesis and Genotyping
Magnetic purification beads	GE Healthcare	cat. no. 65152105050250	sgRNA Synthesis
Tris (2-carboxyethyl) phosphine hydrochloride (TCEP)	Sigma-Aldrich	cat. no. C4706	RNP Buffer
Tris-HCl solution, pH 8.5 molecular-biology grade	Teknova	cat. no. T1085	Lysis Buffer
Tween 20 molecular-biology grade	Sigma-Aldrich	cat. no. P7949-500	Lysis Buffer