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Developmental Biology

Efficient Genome Editing of Mice by CRISPR Electroporation of Zygotes

Published: December 16, 2022 doi: 10.3791/64302


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 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.


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'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.

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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 instrument9,31.
    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 methods9,16.
    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 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
    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 described9,16.
    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 described9,16.

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Representative 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 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.

Figure 1
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. Please click here to view a larger version of this figure.

Figure 2
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. Please click here to view a larger version of this figure.

Figure 3
Figure 3: Editing strategies' 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. Please click here to view a larger version of this figure.

Figure 4
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. Please click here to view a larger version of this figure.

Figure 5
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. Please click here to view a larger version of this figure.

Figure 6
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 "in/del" 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. Please click here to view a larger version of this figure.

Table 1: Phenotype and viability of Tyr-edited zygotes and mice. This table has been adapted with permission from Modzelewski et al.9. Please click here to download this Table.

Table 2: Results of CRISPR-EZ and microinjection deletion experiments. This table has been adapted with permission from Modzelewski et al.9. Please click here to download this Table.

Supplementary Table 1: List of oligos and donor ssODN. This table has been adapted with permission from Modzelewski et al.9. Please click here to download this File.

Supplementary Table 2: DNA template for the sgRNA reaction setup. This table has been adapted with permission from Modzelewski et al.9. Please click here to download this File.

Supplementary Table 3: In vitro transcription setup. This table has been modified with permission from Modzelewski et al.9. Please click here to download this File.

Supplementary Table 4: RNP complex setup for NHEJ formation. Please click here to download this File.

Supplementary Table 5: RNP complex setup for HDR formation. This table has been adapted with permission from Modzelewski et al.9. Please click here to download this File.

Supplementary Table 6: RNP complex setup for deletion. This table has been modified with permission from Modzelewski et al.9. Please click here to download this File.

Supplementary Table 7: Genotyping PCR setup. This table has been modified with permission from Modzelewski et al.9. Please click here to download this File.

Supplementary Table 8: Nested genotyping PCR setup. This table has been modified with permission from Modzelewski et al.9. Please click here to download this File.

Supplementary Table 9: HinfI restriction digest setup. Please click here to download this File.

Supplementary Table 10: EcoRi restriction digest setup. Please click here to download this File.

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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 "the average mouse lab" 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.

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There are no relevant financial declarations by the authors.


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).


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



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Diallo, C. K., Modzelewski, A. J. Efficient Genome Editing of Mice by CRISPR Electroporation of Zygotes. J. Vis. Exp. (190), e64302, doi:10.3791/64302 (2022).More

Diallo, C. K., Modzelewski, A. J. Efficient Genome Editing of Mice by CRISPR Electroporation of Zygotes. J. Vis. Exp. (190), e64302, doi:10.3791/64302 (2022).

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