This work was supported by Scientific Research (B) (19H03142: to SM), Scientific Research (A) (20H00444: to FS), Scientific Research (A) (21H04838: to SM), and Scientific Research on Innovative Areas &#34;Platform of Advanced Animal Model Support&#34; (16H06276: to ST) from the Ministry of Education, Culture, Sports, Science, and Technology. The funders had no role in study design, data collection and analysis, decision to publish, or manuscript preparation. We are grateful to Ryoichi Mori for his helpful discussion about genome-editing design.

All animal experiments were conducted humanely with approval from the Institutional Animal Experiment Committee of the University of Tsukuba, according to the Regulations for Animal Experiments of the University of Tsukuba and the Fundamental Guidelines for Proper Conduct of Animal Experiments and Related Activities in Academic Research Institutions under the jurisdiction of the Ministry of Education, Culture, Sports, Science, and Technology of Japan. C57BL/6J mice of both sexes, 10-25 weeks old, were used as the zygote donor. ICR mice (older than 10 weeks) were used as the recipient mice. The mice were obtained from commercial sources (see Table of Materials).
1. Confirmation of crRNA cleavage activity in a cell-free system
<ol>
	<li>Dissolve 2 nmol of crRNA (dry) and 20 nmol of tracrRNA (dry) in 25 &#181;L and 250 &#181;L of RNase-free water, respectively (see Table of Materials). 
	NOTE: CRISPR target sequences that were predicted to have high cleavage activity and as few off-targets as possible are selected using CRISPOR (see Table of Materials). In the case of gene cassette knock-in, target the sequence across the insertion site. The exon(s) to be floxed were selected using the KOnezumi (see Table of Materials).</li>
	<li>Amplify the DNA fragment (approximately 0.5-1 kb) containing the target site by genomic PCR9. Purify this PCR amplicon with the common PCR product extraction kit (see Table of Materials).</li>
	<li>Prepare 10 &#181;L of CRISPR mixture (100 ng/&#181;L of crRNA, 150 ng/&#181;L of tracrRNA, and 1 ng/&#181;L of Cas9 in the Cas9 Nuclease Reaction Buffer, see Table of Materials). To construct the CRISPR-Cas9 complex, place the CRISPR mixture in a heat block at 37 &#176;C for 30 min.</li>
	<li>Add the total volume (10 &#181;L) of CRISPR mixture to the PCR product (10 &#181;L, 20 ng/&#181;L) described in step 1.2 and incubate at 37 &#176;C for 1 h. Add 1 &#181;L of RNase (500 ng/&#181;L) to degrade the crRNA and tracrRNA at 37 &#176;C for 30 min, as RNAs must be absent during electrophoresis. Perform electrophoresis of the above samples using loading dye containing sodium dodecyl sulfate (2% w/v). 
	&#8203;NOTE: In rare cases, crRNAs with no cleavage activity are observed. In such cases, it is recommended to redesign the genome editing design.</li>
</ol>
2. Preparation of the mixture of crRNA, tracrRNA, Cas9 protein, and donor DNA for zygote microinjection
<ol>
	<li>Prepare CRISPR solution (50 ng/&#181;L of crRNA, 200 ng/&#181;L of tracrRNA, and 200 ng/&#181;L of Cas9 in RNase-free water). Prepare a donor DNA solution (20 ng/&#181;L of self-constructed plasmid DNA vector).
	<ol>
		<li>Filter the donor DNA solution through a sterile syringe filter with a 0.22 &#181;m pore size. Mix equal volumes of CRISPR solution and filtered donor DNA solution. Ensure that the final concentration of each is 25 ng/&#181;L of crRNA, 100 ng/&#181;L of tracrRNA, 100 ng/&#181;L of Cas9, and 10 ng/&#181;L of donor DNA. This mixture will hereafter be referred to as RNPD. 
		NOTE: When cutting two sites, such as during floxed mice production, the concentration of each crRNA must be 25 ng/&#181;L. The donor DNA is a circular plasmid DNA and can be purified with a common mini prep spin column kit (see Table of Materials). The prepared solution must be placed on ice and used for micro-injection as soon as possible.</li>
	</ol>
	</li>
</ol>
3. Obtaining mice zygotes by natural mating
<ol>
	<li>Inject 5 IU of pregnant mare&#39;s serum gonadotropin (PMSG, see Table of materials) subcutaneously into C57BL/6J female mice (10-25 weeks old) 3 days before microinjection. Approximately 46-48 h later, administer 5 IU of human chorionic gonadotropin (hCG, see Table of materials) intraperitoneally. Co-house the PMSG- and hCG-treated female mice with male C57BL/6J mice and check the vaginal plugs the following morning. 
	NOTE: Approximately 15-25 zygotes can be obtained from one female C57BL/6J mouse. The timeline is shown in Figure 1.</li>
	<li>Prepare two 35 mm Petri dishes for zygote culture, as shown in Figure 2. Place them in the incubator (37 &#176;C, 5% CO2) until use. These can be stored overnight.</li>
	<li>Euthanize the female mice with confirmed vaginal plugs by cervical dislocation and, using small scissors, incise through the abdominal skin and muscle layer from the midline of the lower abdomen to below the ribs. 
	NOTE: Follow local animal ethics committee recommendations for euthanasia.</li>
	<li>Collect the oviducts and place them in a 20 &#181;L drop of M2 medium containing 0.75 mg/mL of hyaluronidase (see Table of Materials) on the lid of the Petri dish. Pick up one oviduct with fine tip forceps and perfuse about 50 &#181;L of M2 medium with hyaluronidase through the fimbriae. 
	NOTE: At this time, the zygotes are loosely surrounded by numerous cumulus cells.</li>
	<li>After 30 s, pick up morphologically normal zygotes with a small amount of medium using a glass capillary and wash them by transferring them to fresh M16 medium droplets. Transfer the washed zygotes into a Petri dish (Figure 2) for pooling. 
	NOTE: The timeline is shown in Figure 1. Morphologically normal zygotes have a male and female pronucleus and two polar bodies that have not severely compromised the spherical shape15. The morphology of the zygote varies considerably among strains and species.</li>
	<li>Repeat steps 3.3 and 3.4 until all zygotes are retrieved. About 100 zygotes can be pooled in one M16 drop in the culture dish. Store the dish in the incubator (37 &#176;C, 5% CO2) until microinjection.</li>
</ol>
4. Preparation of the microinjection needle
<ol>
	<li>Pull some glass pipettes using the programmable pipette puller (see Table of Materials). 
	NOTE: Microinjection needles must have a thin tip and long taper. If the puller has a program for making needles for intracytoplasmic sperm injection, the same program can be used to make needles for microinjection.</li>
	<li>Break the tip of the microinjection needle by hitting the glass ball attached to the microforge tip. Ensure that the tip size of the microinjection needle is around 1 &#181;m in diameter. Check the tip size under a microscope at 1000x magnification.</li>
	<li>Bend the microinjection needle at about 2-3 mm from the tip using a microforge. 
	&#8203;NOTE: The angle of the bend depends on the setup of the micromanipulator. Too much bending will reduce the effectiveness of the piezo pulse.</li>
</ol>
5. Zygote microinjection
<ol>
	<li>Inject 1-1.5 cm of the operation liquid (inertia body that is needed for the piezo effect to make holes as an alternative to mercury, see Table of Materials) into the center of the microinjection needle and attach it to the manual microinjector holder with the piezo actuator.
	<ol>
		<li>Further, attach the holding needle to the opposite manual microinjector holder. Move the operation liquid to the microinjection needle tip with gradual pressure. Fix both the manual microinjector holders on the micromanipulator. 
		NOTE: The operation liquid coming out of the tip of the microinjection needle can be observed under a microscope at 50x magnification. After adjusting the amount of liquid emitted, the splashing of liquid can be observed when the piezo pulses are applied, confirming that the piezo pulses are effective. If this cannot be confirmed, re-prepare the microinjection needle.</li>
	</ol>
	</li>
	<li>Prepare an injection chamber with three droplets: (1) 10 &#181;L of M2 medium; (2) 5 &#181;L of 12% polyvinylpyrrolidone (PVP) in M2 medium; and (3) a 5 &#181;L mixture of crRNA, tracrRNA, Cas9 protein, and the Donor DNA (RNPD) solution. Cover the droplets with mineral oil in the same manner as step 3.2 and set the injection chamber on the inverted microscope stage.
	<ol>
		<li>Under the inverted microscope at a 50x magnification, lower the microinjection pipette into the 12% PVP drop. Aspirate and dispense 12% PVP to clean the inside of the microinjection needle tip and transfer it to the RNPD droplet.</li>
	</ol>
	</li>
	<li>Put the manual microinjector under negative pressure and suck up the RNPD solution into the microinjection needle. Wait for a few minutes until a sufficient volume is aspirated; under a microscope magnified 50x, fill the entire inside of the microinjection needle with liquid.
	<ol>
		<li>Stop the intake and allow the RNPD solution to be expelled gradually by setting the manual microinjector to positive pressure. Move the tip of the microinjection needle into the oil. 
		NOTE: Turn the knob of the manual microinjector counterclockwise for inhalation. To stop inhalation, rotate the knob clockwise and gradually increase the pressure while observing under a microscope. The RNPD solution will be gradually drained. Liquid level movement in the microinjection needle allows confirmation of the outflow.</li>
	</ol>
	</li>
	<li>Put 50 zygotes in the M2 droplet and gently lower the holding pipette into the same droplet. Transfer the microinjection needle to the M2 drop containing zygotes. Switch to a 20x objective and focus on the tip of the microinjection pipette. 
	NOTE: Put as many zygotes that can be injected in 10 min.</li>
	<li>Hold a zygote and focus on the pronucleus by moving the micromanipulator. Insert the microinjection needle into the zygote and bring the tip close to the pronucleus. Once the tip reaches the pronucleus membrane, apply the piezo pulse (intensity: 6-10, speed: 1) to pierce.
	<ol>
		<li>When the microinjection needle punctures the pronucleus, inject the RNPD solution, and observe the swelling of the pronucleus. Once the pronucleus is fully inflated, quickly pull out the needle. Perform the same procedure on both the female and male pronuclei. 
		NOTE: The inflation of the pronucleus by injection can be clearly seen under the microscope. The degree of this inflation is difficult to verbalize but can be confirmed by referring to previous reports15.</li>
	</ol>
	</li>
	<li>Move the injected zygote to another location within the M2 droplet to identify the zygotes before and after injection.</li>
	<li>Repeat steps 5.5-5.6 until all the zygotes in the M2 droplet have been injected. After the injection, transfer the zygotes into a fresh M16 dish.</li>
	<li>Select the survived zygotes 10 min after injection. Remove the lysed zygotes which are damaged by the injection and distribute survived zygotes to each droplet in the M16 dish. Each droplet must contain 20-24 zygotes for one pseudopregnant female. This reduces the time the M16 dish is exposed to the environment outside the incubator during embryo transfer. 
	&#8203;NOTE: Under optimal injection conditions, the survival rate is 90%-95%. This depends on the size of the needle tip, the strength of the piezo pulse, and the outflow pressure of the RNPD solution.</li>
</ol>
6. Embryo transfer into the oviduct
<ol>
	<li>For obtaining pseudopregnant mice, mate each female ICR mouse in the proestrus phase with a vasoligated male ICR mouse. Check the plugs the following day. 
	NOTE: Approximately 15 pseudopregnant mice are obtained from 20 mating pairs.</li>
	<li>Administer three types of mixed anesthetic agents (0.2 mg/kg of medetomidine, 4.0 mg/kg of midazolam, and 2.5 mg/kg of butorphanol, see Table of Materials) subcutaneously to the pseudopregnant female mice. Confirm appropriate anesthesia by a decrease in respiratory rate and disappearance of the tail-pinch and hindlimb pedal withdrawal reflexes. Apply ophthalmic ointment to lubricate the eyes as per local animal ethics committee recommendations.</li>
	<li>After razor-shaving the dorsal region of the mouse in the prone position and disinfecting the surgical area three times, alternating between an iodine or chlorhexidine-based scrub and alcohol, incise the dorsal skin approximately 1 cm along the spine with small dissecting scissors.</li>
	<li>Shift the dorsal incision wound to one side ventrally, incise the muscle layer through which the ovary can be seen, grasp the fat above the ovary with tweezers, and withdraw the ovary, oviduct, and uterus.
	<ol>
		<li>Secure the fat with a serrefine clamp (see Table of Materials). Place the reproductive tissues on sterile gauze to avoid direct contact with the coat.</li>
	</ol>
	</li>
	<li>In the following order, place a small amount of culture medium (M2 or M16), some air bubbles as a marker, and the zygotes in a pipette for transfer. 
	NOTE: Transfer approximately 10-12 zygotes per oviduct.</li>
	<li>Make an incision using a micro shear (see Table of Materials) between the ampulla and the fimbriae of the oviduct (just long enough to allow the glass pipette to enter), introduce the zygotes into the incision, and simultaneously confirm that the air bubble has been introduced.</li>
	<li>Perform the implantation into the other oviduct similarly (step 6.6).</li>
	<li>Suture the outer skin with autoclips (see Table of Materials). 
	&#8203;NOTE: Do not suture the incised muscle layer unless the incision wound is unavoidably large. The ideal size of the incised muscle layer should be less than 2-3 mm. If the incision is greater than 5 mm, the muscle layer should be sutured using a sterilized suture needle and thread (see Table of Materials).</li>
</ol>
7. Animal recovery
<ol>
	<li>Administer atipamezole hydrochloride (0.3 mg/kg) subcutaneously to mice. 
	NOTE: Follow local animal ethics committee recommendations for post-operative analgesia.</li>
	<li>Then, place the mice in cages and keep them warm on a hot plate at 37 &#176;C.</li>
	<li>After the mice are awakened, keep them warm for about 2 h. After confirming no abnormal behavior of the mice, return the cages to the breeding rack.</li>
</ol>

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The authors have declared no competing interests.

In the present study, fresh (not frozen-thawed) C57BL/6J mouse zygotes, obtained from natural mating, were used for genome editing mouse production. By injecting crRNA-tracrRNA-Cas9 and donor DNA (RNPD) into both pronuclei of these zygotes in a cell cycle phase where knock-in is most likely to occur, gene cassette knock-in and floxed mice were generated with a high birth rate and sufficient genome editing efficiency. The current study strengthens the extensibility of the SPRINT-CRISPR method15, where it ensures sufficient knock-in efficiency even in the C57BL/6J genetic background, and can be applied to the production of floxed mice.
Electroporation is a highly generalized method of producing genome-edited mice because of the low cost of the experimental devices and the ease of learning the technique8. Furthermore, the i-GONAD method19, in which genome editing is done with preimplantation embryos existing in the oviduct, does not require a recipient mouse. Compared to these methods, the present method presented here is costly and time-consuming when preparing and maintaining an experimental environment in terms of both hardware and software. However, once the experimental facilities are set up, complex gene cassette knock-in and floxed mice can be produced with only commercially available CRISPR elements (crRNA, tracrRNA, and Cas9 protein) and a simple, small amount of circular plasmid vector to be used as donor DNA. This means that many complex genome-edited mouse lines can be produced without the need for time-consuming (or relatively expensive) lsODN5,6 or adeno-associated virus vectors20 to prepare.
Unlike the original SPRINT-CRISPR method15, fresh (frozen-thawed) zygotes were used. When obtaining fresh zygotes by natural mating, the technical errors can be ignored that may occur during in vitro fertilization or freezing and thawing. Also, the embryo manipulation process can be completed in approximately half a day (Figure 1) following the current approach. On the other hand, some limitations of the present method include the requirement of many male mice, the loose control of the fertilization timing, and the limited ability to completely predict the number of zygotes for microinjection. Compared to the original SPRINT-CRISPR method, this method may have a higher birth rate (Table 1). Despite this, it cannot be concluded that the present method is better in terms of birth rate because of the differences in the experiment conductors, target genes, genetic backgrounds, and timing of embryo confirmation.
As shown in Table 1, a large number of mice in most of the gene cassette knock-in projects had random integration alleles. Random integration must be eliminated because it can lead to unintended disruption of endogenous genes and ectopic expression of transgenes. The challenge for the future is to find experimental conditions that reduce the number of random integration events while maintaining sufficient knock-in efficiency.
Almost all mouse zygote genome editing using genome editing effectors that result in DNA double-strand breaks is likely to induce unintended mutations at on-target sites9,21. The results of these unintended on-target mutations are not described here because we are not in a situation in which the next generation of mice can be managed in order to present clear evidence of them. The best way to rule out the concern of confusion caused by unintended on-target mutagenesis is to obtain the next generation of mice by mating founders with wild-type mice rather than intercrossing founders. In principle, the N1 mice resulting from this cross are wild-type (WT) on one side of the allele, so if the intended knock-in (KI) allele is detected, they are WT/KI heterozygous. Therefore, the on-target mutagenesis is not a critical problem in the laboratory mouse, which has a relatively fast life cycle and is a prolific animal. However, further improvement will be necessary when this technology is applied to relatively large mammals.
It has been confirmed that this technology can produce gene cassette knock-in mice in inbred strains other than C57BL/6J, but a sufficient number of projects are not secured, and the data are highly variable. For this reason, this information is not included in the present study. It is believed that further studies on this subject will be necessary to advance mouse genetics and disease model research.

Knock-in mice, in which the intended exogenous genes are introduced at the target loci, are widely used in many in vivo studies as gene humanized mice, fluorescent reporter mice, and Cre driver mice1,2. When knock-in mutations are induced by genome editing in mouse zygotes, single-stranded DNA (single-stranded oligo DNA donors, ssODN) or double-stranded DNA (dsDNA) is used as donor DNA2,3. ssODN is mainly used to knock-in relatively short gene fragments of less than 200 bp4. It is possible to knock-in fragments longer than 1 kb DNA using long ssODNs (lsODN)5,6, however their preparation is time-consuming. When dsDNA donors are used, gene cassette (&#62;1 kb) knock-in mice can be generated without laborious donor DNA preparation7.
The principal advantage of using ssODNs is that electroporation8 can generate knock-in mice. However, dsDNA donors must be introduced directly into the nucleus by zygote microinjection. Simultaneous knock-in at two sites is required to create floxed mice, in which each loxP sequence is knocked-in upstream and downstream of the target gene. There are two ways to generate floxed mice by genome editing in mouse zygotes-using two independent ssODNs, each carrying a single loxP site, or using a single dsDNA (or lsDNA) with a floxed sequence; the former is very inefficient9,10,11. Genome editing using dsDNA donors is the simplest approach to produce gene cassette knock-in and floxed mice if the environment is conducive to zygote microinjection.
Initially, mixtures of Cas9 mRNA and sgRNA or DNA vectors encoding sgRNA and Cas9 are used for genome editing in mouse zygotes12,13. Since high-quality and stable Cas9 proteins are now available at a low cost, the introduction of crRNA-tracrRNA-Cas9 ribonucleoprotein (RNP) into mouse zygotes14 has become popular. Recently, knock-in mice have been generated with high efficiency by introducing the crRNA-tracrRNA-Cas9 RNP and donor DNA into both pronuclei of mouse zygotes in a cell cycle phase where knock-in is most likely to occur15. Therefore, the present protocol describes techniques for producing various types of knock-in mice by this method.
There are many useful inbred strains of laboratory mice16. Genetic modification within the inbred background can disregard the influence of genetic background on the phenotype. This article describes a method for inducing gene cassette knock-in and floxed mutations in the zygote of the most commonly used inbred mouse line, the C57BL/6J17. Further, the timeline for the generation of genetically modified mice and the peripheral techniques used are also discussed, including the induction of superovulation and embryo transfer.

<table><tbody><tr><td>Atipamezole Hydrochloride</td><td>ZENOAQ</td><td>-</td><td></td></tr><tr><td>Autoclip Wound Clip</td><td>BD</td><td>427631</td><td></td></tr><tr><td>Autoclip Wound Clip Applier</td><td>BD</td><td>427630</td><td></td></tr><tr><td>Butorphanol</td><td>Meiji Seika Pharma</td><td>-</td><td></td></tr><tr><td>C57BL/6J mice</td><td>Jackson Laboratory Japan</td><td>-</td><td>older than 10 weeks old</td></tr><tr><td>Calibrated Pipet, 100ul</td><td>Drummond Scientific Company</td><td>2-000-100</td><td>for collecting zygote and embryo transfer</td></tr><tr><td>Cas9</td><td>Thermo Fisher Scientific</td><td>A36499</td><td></td></tr><tr><td>Cas9 Nuclease Reaction Buffer</td><td>NEB</td><td>B7203</td><td></td></tr><tr><td>CellTram 4r&amp;nbsp; oil</td><td>Eppendorf</td><td>5196000030</td><td></td></tr><tr><td>CRISPOR</td><td></td><td>http://crispor.tefor.net/</td><td>web tool for genome editing experiments with the CRISPR-Cas9 system</td></tr><tr><td>crRNA</td><td>IDT</td><td>-</td><td></td></tr><tr><td>Gel/PCR Extraction Kit</td><td>FastGene</td><td>FG-91302</td><td></td></tr><tr><td>hCG</td><td>ASKA Animal Health</td><td>-</td><td></td></tr><tr><td>Hyaluronidase</td><td>Merck Sigma-Aldrich</td><td>H3884</td><td></td></tr><tr><td>ICR mice</td><td>Jackson Laboratory Japan</td><td>-</td><td>older than 10 weeks old, weight 28 G or more</td></tr><tr><td>Inverted microscope</td><td>Leica</td><td></td><td></td></tr><tr><td>KOnezumi</td><td></td><td>https://www.md.tsukuba.ac.jp/LabAnimalResCNT/KOanimals/konezumi.html</td><td>a web application for automating gene disruption strategies to generate knockout mice</td></tr><tr><td>M16 medium</td><td>Merck Sigma-Aldrich</td><td>M7292</td><td></td></tr><tr><td>M2 medium</td><td>Merck Sigma-Aldrich</td><td>M7167</td><td></td></tr><tr><td>Medetomidine</td><td>ZENOAQ</td><td>-</td><td></td></tr><tr><td>Micro shears</td><td>Natsume Seisakusho</td><td>MB-54-1</td><td></td></tr><tr><td>Microforge</td><td>Narishige</td><td>MF-900</td><td>for fire polishing of holding pipette and bending the microinjection needle</td></tr><tr><td>Micromanipulator units</td><td>Narishige</td><td></td><td></td></tr><tr><td>Micropipette puller</td><td>Sutter Instrument</td><td>P-1000</td><td>programmable pipette puller</td></tr><tr><td>Midazolam</td><td>Maruishi Pharmaceutical</td><td>-</td><td></td></tr><tr><td>MILLEX-GV 0.22 &amp;micro;m filter</td><td>Merck Millipore</td><td>SLGV033R</td><td></td></tr><tr><td>Mineral oil</td><td>Nacalai</td><td>26114-75</td><td>for zygote culture and injection chamber</td></tr><tr><td>Petri dish (35mm, untreated)</td><td>Iwaki</td><td>1000-035</td><td>for zygote culture</td></tr><tr><td>PIEZO micromanipulator</td><td>PRIME TECH</td><td>PMM-150</td><td></td></tr><tr><td>Plasmid Mini Kit</td><td>FastGene</td><td>FG-90502</td><td>mini prep spin column kit</td></tr><tr><td>PMM Operation Liquid</td><td>PRIME TECH</td><td>KIT-A</td><td>operation liquid for microinjection</td></tr><tr><td>PMSG</td><td>ASKA Animal Health</td><td>-</td><td></td></tr><tr><td>Polyvinylpyrrolidone</td><td>Merck Sigma-Aldrich</td><td>P5288</td><td></td></tr><tr><td>RNase</td><td>QIAGEN</td><td>19101</td><td></td></tr><tr><td>RNase Free Water</td><td>IDT</td><td>-</td><td>tracrRNA Accessory Reagents</td></tr><tr><td>Serrefine clamp</td><td>Natsume Seisakusho</td><td>C-18</td><td></td></tr><tr><td>Sodium dodecyl sulfate</td><td>SIGMA</td><td>151-21-3</td><td></td></tr><tr><td>Suture needle with thread</td><td>Natsume Seisakusho</td><td>C11-60B2</td><td></td></tr><tr><td>Thin wall borosilicate glass&lt;br/&gt; without filament</td><td>Sutter Instrument</td><td>B100-75-10</td><td>for microinjection needle and holding pipette</td></tr><tr><td>tracrRNA</td><td>IDT</td><td>-</td><td></td></tr></tbody></table>

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 production outcomes of gene cassette knock-in and floxed mice using this protocol are shown in Table 1 and Table 2. The target genes were not stated because each genome-edited mouse line is currently being used in an independent, unpublished project. Instead, the target chromosomal regions were described.
The genotyping analyses were performed using a previously reported method18. In this genotyping method, mice in which the donor DNA is inserted into unintended chromosomal sites are not counted as positive individuals, even if the desired genome editing is induced. Hence, the production efficiency of the founder mice that are truly useful for actual purposes was shown, rather than the efficiency of the genome editing itself.
The median production efficiency of 13 independent gene cassette knock-in mice was 20.8%, with a maximum of 39.5% and a minimum of 7.9% (Table 1). This efficiency was considered to be sufficiently practical. Any lethal embryonic genes were not targeted during the gene cassette knock-in. The median birth rate under this non-lethal gene targeting condition was 34.0% (maximum 43.3% and minimum 15.9%). Since this is comparable to the birth rate in embryo transfer without genetic manipulation, we considered that the toxicity of the introduced genome editing elements and the physical damage of zygote microinjection was unlikely to affect embryonic development.
The median production efficiency of the 10 independent floxed mice was 7.7%, with a maximum of 20.7% and a minimum of 2.1% (Table 2). Although the functions of several target genes were unknown, the median birth rate was 30.2%, with a maximum of 43.8% and a minimum of 17.3%. This rate was comparable to that of gene cassette knock-in mice, suggesting that the micro-injection operation has a negligible effect on the embryonic development of floxed mice.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/64161/64161fig01.jpg" /> 
Figure 1: Timeline of zygote microinjection. White and dark color shows the light and dark cycle, respectively. The mice were maintained under a 14 h light/10 h dark cycle. PMSG: pregnant mare&#39;s serum gonadotropin; hCG: human chorionic gonadotropin <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/64161/64161fig01large.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/64161/64161fig02.jpg" /> 
Figure 2: Preparation of culture dishes. (A) Culture dish for the zygotes before the injection. Place two drops of M16 medium (20-25 &#181;L for each) in a 35 mm plastic Petri dish. (B) Culture dish for the zygotes after the injection. Place 10 drops (10-20 &#181;L for each drop) of M16 in a 35 mm plastic Petri dish. Drops are covered with mineral oil (3 mL). <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/64161/64161fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<table fo:font-size="8px" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td rowspan="3">Targeted Chromosomal Region</td>
			<td colspan="6">Number of</td>
			<td rowspan="3">Knock-in insertion length (kb)</td>
			<td colspan="2" rowspan="2">Homology arm length (kb)</td>
		</tr>
		<tr>
			<td colspan="2">Embryos</td>
			<td rowspan="2">Newborns</td>
			<td rowspan="2">Examined (weanings)</td>
			<td rowspan="2">Knock-in W/O rTG</td>
			<td rowspan="2">rTGc</td>
		</tr>
		<tr>
			<td>injected</td>
			<td>transferred</td>
			<td>5&#39; arm</td>
			<td>3&#39; arm</td>
		</tr>
		<tr>
			<td>2qE2</td>
			<td>349</td>
			<td>331</td>
			<td>114 (34.4%)a</td>
			<td>114</td>
			<td>9 (7.9%)b</td>
			<td>5 (4.4%)d</td>
			<td>1.0</td>
			<td>1.0</td>
			<td>1.0</td>
		</tr>
		<tr>
			<td>2qE2</td>
			<td>231</td>
			<td>215</td>
			<td>78 (36.3%)a</td>
			<td>74</td>
			<td>15 (20.3%)b</td>
			<td>23 (31.1%)d</td>
			<td>2.2</td>
			<td>1.0</td>
			<td>1.1</td>
		</tr>
		<tr>
			<td>5qG2</td>
			<td>288</td>
			<td>262</td>
			<td>90 (34.4%)a</td>
			<td>81</td>
			<td>19 (23.5%)b</td>
			<td>18 (22.2%)d</td>
			<td>1.6</td>
			<td>2.0</td>
			<td>2.0</td>
		</tr>
		<tr>
			<td>6qB1</td>
			<td>278</td>
			<td>259</td>
			<td>58 (22.4%)a</td>
			<td>46</td>
			<td>11 (23.9%)b</td>
			<td>11 (23.9%)d</td>
			<td>4.2</td>
			<td>1.2</td>
			<td>1.0</td>
		</tr>
		<tr>
			<td>6qE3 [ROSA26]</td>
			<td>295</td>
			<td>277</td>
			<td>97 (35.0%)a</td>
			<td>23</td>
			<td>2 (8.7%)b</td>
			<td>4 (17.4%)d</td>
			<td>6.5</td>
			<td>1.1</td>
			<td>3.0</td>
		</tr>
		<tr>
			<td>6qE3 [ROSA26]</td>
			<td>246</td>
			<td>219</td>
			<td>87 (39.7%)a</td>
			<td>83</td>
			<td>22 (26.5%)b</td>
			<td>18 (21.7%)d</td>
			<td>5.8</td>
			<td>1.1</td>
			<td>3.0</td>
		</tr>
		<tr>
			<td>7qF5</td>
			<td>279</td>
			<td>268</td>
			<td>116 (43.3%)a</td>
			<td>114</td>
			<td>45 (39.5%)b</td>
			<td>44 (38.6%)d</td>
			<td>1.4</td>
			<td>3.1</td>
			<td>1.0</td>
		</tr>
		<tr>
			<td>11qB4</td>
			<td>405</td>
			<td>353</td>
			<td>56 (15.9%)a</td>
			<td>47</td>
			<td>8 (17.0%)b</td>
			<td>12 (25.5%)d</td>
			<td>1.7</td>
			<td>1.0</td>
			<td>0.8</td>
		</tr>
		<tr>
			<td>11qD</td>
			<td>310</td>
			<td>294</td>
			<td>100 (34.4%)a</td>
			<td>98</td>
			<td>15 (15.3%)b</td>
			<td>76 (77.6%)d</td>
			<td>1.0</td>
			<td>1.8</td>
			<td>2.4</td>
		</tr>
		<tr>
			<td>12qE</td>
			<td>368</td>
			<td>320</td>
			<td>86 (26.9%)a</td>
			<td>84</td>
			<td>12 (14.3%)b</td>
			<td>19 (22.6%)d</td>
			<td>3.4</td>
			<td>1.1</td>
			<td>1.3</td>
		</tr>
		<tr>
			<td>12qF1</td>
			<td>315</td>
			<td>265</td>
			<td>76 (28.7%)a</td>
			<td>72</td>
			<td>17 (23.6%)b</td>
			<td>28 (38.9%)d</td>
			<td>1.0</td>
			<td>1.1</td>
			<td>1.1</td>
		</tr>
		<tr>
			<td>14qE5</td>
			<td>256</td>
			<td>215</td>
			<td>48 (22.3%)a</td>
			<td>48</td>
			<td>11 (22.9%)b</td>
			<td>21 (43.8%)d</td>
			<td>3.1</td>
			<td>1.0</td>
			<td>0.9</td>
		</tr>
		<tr>
			<td>14qE5</td>
			<td>287</td>
			<td>285</td>
			<td>85 (29.8%)a</td>
			<td>72</td>
			<td>15 (20.8%)b</td>
			<td>19 (26.4%)d</td>
			<td>2.6</td>
			<td>1.0</td>
			<td>0.9</td>
		</tr>
	</tbody>
</table>
Table 1: Knock-in mice production by micro-injection. The table shows the birth rate and the efficiency of producing mice that carry the intended knock-in allele without (W/O) random integration (rTG) of the donor DNA. It was possible to obtain mice with the intended knock-in allele in all projects. a: Number of Newborns/Number of transferred embryos; b: Number of knock-in alleles carried mice/number of examined mice; c: it is not certain if there is an intended allele; d: Number of rTG allele carried mice/number of examined mice; rTG: random integration of donor vector. W/O: without.
<table fo:font-size="8px" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td rowspan="3">Targeted Chromosomal Region</td>
			<td colspan="5">Number of</td>
			<td rowspan="3">floxed length (kb)</td>
			<td colspan="2" rowspan="2">Homology arm length (kb)</td>
		</tr>
		<tr>
			<td colspan="2">Embryos</td>
			<td rowspan="2">Newborns</td>
			<td rowspan="2">Examined (weanings)</td>
			<td rowspan="2">floxed W/O rTG</td>
		</tr>
		<tr>
			<td>injected</td>
			<td>transferred</td>
			<td>5&#39; arm</td>
			<td>3&#39; arm</td>
		</tr>
		<tr>
			<td>3qC</td>
			<td>325</td>
			<td>313</td>
			<td>93 (29.7%)a</td>
			<td>87</td>
			<td>9 (10.3%)b</td>
			<td>1.3</td>
			<td>1.2</td>
			<td>1.1</td>
		</tr>
		<tr>
			<td>4qB1</td>
			<td>210</td>
			<td>200</td>
			<td>36 (18.0%)a</td>
			<td>29</td>
			<td>6 (20.7%)b</td>
			<td>1.6</td>
			<td>1.2</td>
			<td>0.9</td>
		</tr>
		<tr>
			<td>4qC4</td>
			<td>272</td>
			<td>256</td>
			<td>112 (43.8%)a</td>
			<td>100</td>
			<td>5 (5.0%)b</td>
			<td>2.4</td>
			<td>1.0</td>
			<td>1.4</td>
		</tr>
		<tr>
			<td>5qF</td>
			<td>143</td>
			<td>137</td>
			<td>40 (29.2%)a</td>
			<td>40</td>
			<td>6 (15.0%)b</td>
			<td>1.8</td>
			<td>1.0</td>
			<td>1.1</td>
		</tr>
		<tr>
			<td>5qF</td>
			<td>255</td>
			<td>227</td>
			<td>80 (35.2%)a</td>
			<td>76</td>
			<td>2 (2.6%)b</td>
			<td>1.3</td>
			<td>1.1</td>
			<td>0.9</td>
		</tr>
		<tr>
			<td>9qE3.1</td>
			<td>289</td>
			<td>261</td>
			<td>80 (30.7%)a</td>
			<td>77</td>
			<td>9 (11.7%)b</td>
			<td>1.2</td>
			<td>1.2</td>
			<td>1.2</td>
		</tr>
		<tr>
			<td>10qB3</td>
			<td>284</td>
			<td>269</td>
			<td>88 (32.7%)a</td>
			<td>78</td>
			<td>3 (3.8%)b</td>
			<td>1.3</td>
			<td>1.1</td>
			<td>0.9</td>
		</tr>
		<tr>
			<td>10qC1</td>
			<td>243</td>
			<td>231</td>
			<td>40 (17.3%)a</td>
			<td>38</td>
			<td>4 (10.5%)b</td>
			<td>2.1</td>
			<td>1.0</td>
			<td>1.3</td>
		</tr>
		<tr>
			<td>14qE5</td>
			<td>390</td>
			<td>372</td>
			<td>139 (37.4%)a</td>
			<td>135</td>
			<td>5 (3.7%)b</td>
			<td>1.1</td>
			<td>0.8</td>
			<td>0.9</td>
		</tr>
		<tr>
			<td>17qA3.3</td>
			<td>383</td>
			<td>374</td>
			<td>99 (26.5%)a</td>
			<td>95</td>
			<td>2 (2.1%)b</td>
			<td>2.1</td>
			<td>0.6</td>
			<td>0.8</td>
		</tr>
	</tbody>
</table>
Table 2: Floxed mice production by micro-injection. The table shows the birth rate and the efficiency of producing mice that carry the intended flox allele without (W/O) random integration (rTG) of the donor DNA. It was possible to obtain mice with the intended flox allele in all projects. a: Number of newborns/number of transferred embryos; b: number of flox allele carried mice/number of examined mice. rTG: random integration of or donor vector; W/O: without.

Watch this Scientific Journal Video about Zygote Microinjection for Creating Gene Cassette Knock-in and Flox Alleles in Mice at JoVE.com

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

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 ...

zygote-microinjection-for-creating-gene-cassette-knock-flox-alleles

Nanyang Technological University

Research

JoVE Journal

Developmental Biology

4.4K Views.  University of Tsukuba. The present protocol describes zygote microinjection of CRISPR-Cas9 and donor DNA to efficiently produce gene cassette knock-in and floxed mice.

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

Imaging Subcellular Structures in the Living Zebrafish Embryo

In vivo imaging provides unprecedented access to the dynamic behavior of cellular and subcellular structures in their natural context. Performing such imaging experiments in higher vertebrates such as mammals generally requires surgical access to the system under study. The optical accessibility of embryonic and larval zebrafish allows such invasive procedures to be circumvented and permits imaging in the intact organism. Indeed the zebrafish is now a well-established model to visualize dynamic cellular behaviors using in vivo microscopy in a wide range of developmental contexts from proliferation to migration and differentiation. A more recent development is the increasing use of zebrafish to study subcellular events including mitochondrial trafficking and centrosome dynamics. The relative ease with which these subcellular structures can be genetically labeled by fluorescent proteins and the use of light microscopy techniques to image them is transforming the zebrafish into an in vivo model of cell biology. Here we describe methods to generate genetic constructs that fluorescently label organelles, highlighting mitochondria and centrosomes as specific examples. We use the bipartite Gal4-UAS system in multiple configurations to restrict expression to specific cell-types and provide protocols to generate transiently expressing and stable transgenic fish. Finally, we provide guidelines for choosing light microscopy methods that are most suitable for imaging subcellular dynamics.

Imaging the dynamic behavior of organelles and other subcellular structures in vivo can shed light on their function in physiological and disease conditions. Here, we present methods for genetically tagging two organelles, centrosomes and mitochondria, and imaging their dynamics in living zebrafish embryos using wide-field and confocal microscopy.

In vivo imaging provides unprecedented access to the dynamic behavior of cellular and subcellular structures in their natural context. Performing such ...

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

In Vivo Imaging of Transgenic Gene Expression in Individual Retinal Progenitors in Chimeric Zebrafish Embryos to Study Cell Nonautonomous Influences

The genetic and technical strengths have made the zebrafish vertebrate a key model organism in which the consequences of gene manipulations can be traced in vivo throughout the rapid developmental period. Multiple processes can be studied including cell proliferation, gene expression, cell migration and morphogenesis. Importantly, the generation of chimeras through transplantations can be easily performed, allowing mosaic labeling and tracking of individual cells under the influence of the host environment. For example, by combining functional gene manipulations of the host embryo (e.g., through morpholino microinjection) and live imaging, the effects of extrinsic, cell nonautonomous signals (provided by the genetically modified environment) on individual transplanted donor cells can be assessed. Here we demonstrate how this approach is used to compare the onset of fluorescent transgene expression as a proxy for the timing of cell fate determination in different genetic host environments.
In this article, we provide the protocol for microinjecting zebrafish embryos to mark donor cells and to cause gene knockdown in host embryos, a description of the transplantation technique used to generate chimeric embryos, and the protocol for preparing and running in vivo time-lapse confocal imaging of multiple embryos. In particular, performing multiposition imaging is crucial when comparing timing of events such as the onset of gene expression. This requires data collection from multiple control and experimental embryos processed simultaneously. Such an approach can easily be extended for studies of extrinsic influences in any organ or tissue of choice accessible to live imaging, provided that transplantations can be targeted easily according to established embryonic fate maps.

In Vivo Imaging of Transgenic Gene Expression in Individual Retinal Progenitors in Chimeric Zebrafish Embryos to Study Cell Nonautonomous Influences

Live tracking of individual WT retinal progenitors in distinct genetic backgrounds allows for the assessment of the contribution of cell non-autonomous signaling during neurogenesis. Here, a combination of gene knockdown, chimera generation via embryo transplantation and in vivo time-lapse confocal imaging was utilized for this purpose.

The genetic and technical strengths have made the zebrafish vertebrate a key model organism in which the consequences of gene manipulations can be traced ...

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 ...

Microinjection of mRNA and Morpholino Antisense Oligonucleotides in Zebrafish Embryos.

An essential tool for investigating the role of a gene during development is the ability to perform gene knockdown, overexpression, and misexpression studies. In zebrafish (Danio rerio), microinjection of RNA, DNA, proteins, antisense oligonucleotides and other small molecules into the developing embryo provides researchers a quick and robust assay for exploring gene function in vivo. In this video-article, we will demonstrate how to prepare and microinject in vitro synthesized EGFP mRNA and a translational-blocking morpholino oligo against pkd2, a gene associated with autosomal dominant polycystic kidney disease (ADPKD), into 1-cell stage zebrafish embryos. We will then analyze the success of the mRNA and morpholino microinjections by verifying GFP expression and phenotype analysis. Broad applications of this technique include generating transgenic animals and germ-line chimeras, cell-fate mapping and gene screening. Herein we describe a protocol for overexpression of EGFP and knockdown of pkd2 by mRNA and morpholino oligonucleotide injection.

Microinjection is a well-established and effective method for introducing foreign substances into fertilized zebrafish embryos. Here, we demonstrate a robust microinjection technique for performing mRNA overexpression, and morpholino oligonucleotide gene knockdown studies in zebrafish.

An essential tool for investigating the role of a gene during development is the ability to perform gene knockdown, overexpression, and misexpression ...

Biology

Microinjection of Zebrafish Embryos to Analyze Gene Function

One of the advantages of studying zebrafish is the ease and speed of manipulating protein levels in the embryo.  Morpholinos, which are synthetic oligonucleotides with antisense complementarity to target RNAs, can be added to the embryo to reduce the expression of a particular gene product.  Conversely, processed mRNA can be added to the embryo to increase levels of a gene product.  The vehicle for adding either mRNA or morpholino to an embryo is microinjection.  Microinjection is efficient and rapid, allowing for the injection of hundreds of embryos per hour.  This video shows all the steps involved in microinjection.  Briefly, eggs are collected immediately after being laid and lined up against a microscope slide in a Petri dish.  Next, a fine-tipped needle loaded with injection material is connected to a microinjector and an air source, and the microinjector controls are adjusted to produce a desirable injection volume.  Finally, the needle is plunged into the embryo's yolk and the morpholino or mRNA is expelled.

This video shows how morpholino or mRNA can be injected into zebrafish embryos at the one-cell stage to decrease or increase the level of specific gene products during subsequent development.

One of the advantages of studying zebrafish is the ease and speed of manipulating protein levels in the embryo.  Morpholinos, which are synthetic ...

Embryo Microinjection Techniques for Efficient Site-Specific Mutagenesis in Culex quinquefasciatus

Culex quinquefasciatus is a vector of a diverse range of vector-borne diseases such as avian malaria, West Nile virus (WNV), Japanese encephalitis, Eastern equine encephalitis, lymphatic filariasis, and Saint Louis encephalitis. Notably, avian malaria has played a major role in the extinction of numerous endemic island bird species, while WNV has become an important vector-borne disease in the United States. To gain further insight into C. quinquefasciatus biology and expand their genetic control toolbox, we need to develop more efficient and affordable methods for genome engineering in this species. However, some biological traits unique to Culex mosquitoes, particularly their egg rafts, have made it difficult to perform microinjection procedures required for genome engineering. To address these challenges, we have developed an optimized embryo microinjection protocol that focuses on mitigating the technical obstacles associated with the unique characteristics of Culex mosquitoes. These procedures demonstrate optimized methods for egg collection, egg raft separation and other handling procedures essential for successful microinjection in C. quinquefasciatus. When coupled with the CRISPR/Cas9 genome editing technology, these procedures allow us to achieve site-specific, efficient and heritable germline mutations, which are required to perform advanced genome engineering and develop genetic control technologies in this important, but currently understudied, disease vector.

Embryo Microinjection Techniques for Efficient Site-Specific Mutagenesis in Culex quinquefasciatus

This protocol describes microinjection procedures for Culex quinquefasciatus embryos that are optimized to work with CRISPR/Cas9 gene editing tools. This technique can efficiently generate site-specific, heritable, germline mutations that can be used for building genetic technologies in this understudied disease vector.

Culex quinquefasciatus is a vector of a diverse range of vector-borne diseases such as avian malaria, West Nile virus (WNV), Japanese encephalitis, ...

Gene Trapping Using Gal4 in Zebrafish

Large clutch size and external development of optically transparent embryos make zebrafish an exceptional vertebrate model system for in vivo insertional mutagenesis using fluorescent reporters to tag expression of mutated genes. Several laboratories have constructed and tested enhancer- and gene-trap vectors in zebrafish, using fluorescent proteins, Gal4- and lexA- based transcriptional activators as reporters 1-7. These vectors had two potential drawbacks: suboptimal stringency (e.g. lack of ability to differentiate between enhancer- and gene-trap events) and low mutagenicity (e.g. integrations into genes rarely produced null alleles). Gene Breaking Transposon (GBTs) were developed to address these drawbacks 8-10. We have modified one of the first GBT vectors, GBT-R15, for use with Gal4-VP16 as the primary gene trap reporter and added UAS:eGFP as the secondary reporter for direct detection of gene trap events. Application of Gal4-VP16 as the primary gene trap reporter provides two main advantages. First, it increases sensitivity for genes expressed at low expression levels. Second, it enables researchers to use gene trap lines as Gal4 drivers to direct expression of other transgenes in very specific tissues. This is especially pertinent for genes with non-essential or redundant functions, where gene trap integration may not result in overt phenotypes. The disadvantage of using Gal4-VP16 as the primary gene trap reporter is that genes coding for proteins with N-terminal signal sequences are not amenable to trapping, as the resulting Gal4-VP16 fusion proteins are unlikely to be able to enter the nucleus and activate transcription. Importantly, the use of Gal4-VP16 does not pre-select for nuclear proteins: we recovered gene trap mutations in genes encoding proteins which function in the nucleus, the cytoplasm and the plasma membrane.

This protocol describes the method of gene trap insertional mutagenesis using Gal4-VP16 as the primary reporter and GFP/RFP as secondary reporters in zebrafish. Approximately one in ten high-expressing F0 fish yield gene trap progeny co-expressing GFP and RFP. The screening procedure can be readily scaled to adapt to the size of the laboratory performing the insertional mutagenesis screen.

Large clutch size and external development of optically transparent embryos make zebrafish an exceptional vertebrate model system for in vivo insertional ...

Zebrafish Microinjection Techniques

One of the major advantages to working with zebrafish (Danio rerio) is that their genetics can be easily manipulated by microinjection of early stage embryos. Using this technique, solutions containing genetic material or knockdown constructs are delivered into the blastomeres: the embryonic cells sitting atop the yolk of the newly fertilized egg. Delivery into the cytoplasm is achieved either through direct injection into the blastomere, or via natural cytoplasmic movements that occur after a solution is injected into the yolk. Successful genetic manipulations are usually followed by quantification of embryonic phenotypes in order to elucidate the genetic mechanisms of development.
This video will provide an introduction to carrying out microinjections in zebrafish embryos. The discussion begins with a review of the essential tools for the technique, including the injection apparatus and the microinjector, which controls fluid movement with pressure pulses of air. Next, important preparatory steps are demonstrated, such as the pouring of agar plates to stabilize embryos during injection and calibration of the microinjection apparatus. The injection procedure is then presented along with tips on when and where injections should be performed. Finally, applications of the microinjection technique are discussed, including gene overexpression via mRNA injection, gene silencing by delivery of antisense morpholino oligonucleotides, and the generation of transgenic zebrafish using specially engineered plasmid DNA.

Introduction to Microinjection of Early Zebrafish Embryos

One of the major advantages to working with zebrafish (Danio rerio) is that their genetics can be easily manipulated by microinjection of early stage ...

Education

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Basic Biology

Biology II: Mouse, Zebrafish, and Chick

Gonad Microinjection: A Method of Compound Delivery Directly into the Germline of C. elegans

Source: Farboud, B. et al, <a href="https://www-jove-com.remotexs.ntu.edu.sg/video/57350/enhanced-genome-editing-with-cas9-ribonucleoprotein-diverse-cells">Enhanced Genome Editing with Cas9 Ribonucleoprotein in Diverse Cells and Organisms.</a>&#160;J. Vis. Exp. (2018).
This video describes microinjection, a common method of creating transgenic C. elegans strains.&#160; In the example, we will see microinjection used to introduce a ribonucleoprotein (RNP) complex for CRISPR-bases gene editing.

Source: Farboud, B. et al, Enhanced Genome Editing with Cas9 Ribonucleoprotein in Diverse Cells and Organisms.&#160;J. Vis. Exp. (2018).
This video ...

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

Summary

Explore More Videos

Imaging Subcellular Structures in the Living Zebrafish Embryo

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

In Vivo Imaging of Transgenic Gene Expression in Individual Retinal Progenitors in Chimeric Zebrafish Embryos to Study Cell Nonautonomous Influences

Generation of Genetically Modified Mice through the Microinjection of Oocytes

Microinjection of mRNA and Morpholino Antisense Oligonucleotides in Zebrafish Embryos.

Microinjection of Zebrafish Embryos to Analyze Gene Function

Embryo Microinjection Techniques for Efficient Site-Specific Mutagenesis in Culex quinquefasciatus

Gene Trapping Using Gal4 in Zebrafish

Zebrafish Microinjection Techniques

Gonad Microinjection: A Method of Compound Delivery Directly into the Germline of C. elegans