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 border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<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&#160; 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 &#181;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 
			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/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/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 ...

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Developmental Biology

4.1K 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

Technique to Target Microinjection to the Developing Xenopus Kidney

The embryonic kidney of Xenopus&#160;laevis (frog), the pronephros, consists of a single nephron, and can be used as a model for kidney disease. Xenopus embryos are large, develop externally, and can be easily manipulated by microinjection or surgical procedures. In addition, fate maps have been established for early Xenopus embryos. Targeted microinjection into the individual blastomere that will eventually give rise to an organ or tissue of interest can be used to selectively overexpress or knock down gene expression within this restricted region, decreasing secondary effects in the rest of the developing embryo. In this protocol, we describe how to utilize established Xenopus fate maps to target the developing Xenopus kidney (the pronephros), through microinjection into specific blastomere of 4- and 8-cell embryos. Injection of lineage tracers allows verification of the specific targeting of the injection. After embryos have developed to stage 38 - 40, whole-mount immunostaining is used to visualize pronephric development, and the contribution by targeted cells to the pronephros can be assessed. The same technique can be adapted to target other tissue types in addition to the pronephros.

Technique to Target Microinjection to the Developing Xenopus Kidney

Here, we present a protocol to use fate maps and lineage tracers to target injections into individual blastomeres that give rise to the kidney of Xenopus laevis embryos.

The embryonic kidney of Xenopus&#160;laevis (frog), the pronephros, consists of a single nephron, and can be used as a model for kidney disease. Xenopus ...

Microinjection for Transgenesis and Genome Editing in Threespine Sticklebacks

The threespine stickleback fish has emerged as a powerful system to study the genetic basis of a wide variety of morphological, physiological, and behavioral phenotypes. The remarkably diverse phenotypes that have evolved as marine populations adapt to countless freshwater environments, combined with the ability to cross marine and freshwater forms, provide a rare vertebrate system in which genetics can be used to map genomic regions controlling evolved traits. Excellent genomic resources are now available, facilitating molecular genetic dissection of evolved changes. While mapping experiments generate lists of interesting candidate genes, functional genetic manipulations are required to test the roles of these genes. Gene regulation can be studied with transgenic reporter plasmids and BACs integrated into the genome using the Tol2 transposase system. Functions of specific candidate genes and cis-regulatory elements can be assessed by inducing targeted mutations with TALEN and CRISPR/Cas9 genome editing reagents. All methods require introducing nucleic acids into fertilized one-cell stickleback embryos, a task made challenging by the thick chorion of stickleback embryos and the relatively small and thin blastomere. Here, a detailed protocol for microinjection of nucleic acids into stickleback embryos is described for transgenic and genome editing applications to study gene expression and function, as well as techniques to assess the success of transgenesis and recover stable lines.

Transgenic manipulations and genome editing are critical for functionally testing the roles of genes and cis-regulatory elements. Here a detailed microinjection protocol for the generation of genomic modifications (including Tol2-mediated fluorescent reporter transgene constructs, TALENs, and CRISPRs) is presented for the emergent model fish, the threespine stickleback.

The threespine stickleback fish has emerged as a powerful system to study the genetic basis of a wide variety of morphological, physiological, and ...

Embryo Microinjection and Electroporation in the Chordate Ciona intestinalis

Simple model organisms are instrumental for in vivo studies of developmental and cellular differentiation processes. Currently, the evolutionary distance to man of conventional invertebrate model systems and the complexity of genomes in vertebrates are critical challenges to modeling human normal and pathological conditions. The chordate Ciona intestinalis is an invertebrate chordate that emerged from a common ancestor with the vertebrates and may represent features at the interface between invertebrates and vertebrates. A common body plan with much simpler cellular and genomic composition should unveil gene regulatory network (GRN) links and functional genomics readouts explaining phenomena in the vertebrate condition. The compact genome of Ciona, a fixed embryonic lineage with few divisions and large cells, combined with versatile community tools foster efficient gene functional analyses in this organism. Here, we present several crucial methods for this promising model organism, which belongs to the closest sister group to vertebrates. We present protocols for transient transgenesis by electroporation, along with microinjection-mediated gene knockdown, which together provide the means to study gene function and genomic regulatory elements. We extend our protocols to provide information on how community databases are utilized for in silico design of gene regulatory or gene functional experiments. An example study demonstrates how novel information can be gained on the interplay, and its quantification, of selected neural factors conserved between Ciona and man. Furthermore, we show examples of differential subcellular localization in embryonic cells, following DNA electroporation in Ciona zygotes. Finally, we discuss the potential of these protocols to be adapted for tissue specific gene interference with emerging gene editing methods. The in vivo approaches in Ciona overcome major shortcomings of classical model organisms in the quest of unraveling conserved mechanisms in the chordate developmental program, relevant to stem cell research, drug discovery, and subsequent clinical application.

Embryo Microinjection and Electroporation in the Chordate Ciona intestinalis

We present transient transgenesis and gene knockdown in Ciona intestinalis, a chordate sister group to vertebrates, using microinjection and electroporation techniques. Such methods facilitate functional genomics in this simple invertebrate that features rudimentary characteristics of vertebrates, including notochord and head sensory epithelia, and many orthologs of human disease associated genes.

Simple model organisms are instrumental for in vivo studies of developmental and cellular differentiation processes. Currently, the evolutionary distance ...

Structure-function Studies in Mouse Embryonic Stem Cells Using Recombinase-mediated Cassette Exchange

Gene engineering in mouse embryos or embryonic stem cells (mESCs) allows for the study of the function of a given protein. Proteins are the workhorses of the cell and often consist of multiple functional domains, which can be influenced by posttranslational modifications. The depletion of the entire protein in conditional or constitutive knock-out (KO) mice does not take into account this functional diversity and regulation. An mESC line and a derived mouse model, in which a docking site for FLPe recombination-mediated cassette exchange (RMCE) was inserted within the ROSA26 (R26) locus, was previously reported. Here, we report on a structure-function approach that allows for molecular dissection of the different functionalities of a multidomain protein. To this end, RMCE-compatible mice must be crossed with KO mice and then RMCE-compatible KO mESCs must be isolated. Next, a panel of putative rescue constructs can be introduced into the R26 locus via RMCE targeting. The candidate rescue cDNAs can be easily inserted between RMCE sites of the targeting vector using recombination cloning. Next, KO mESCs are transfected with the targeting vector in combination with an FLPe recombinase expression plasmid. RMCE reactivates the promoter-less neomycin-resistance gene in the ROSA26 docking sites and allows for the selection of the correct targeting event. In this way, high targeting efficiencies close to 100% are obtained, allowing for insertion of multiple putative rescue constructs in a semi-high throughput manner. Finally, a multitude of R26-driven rescue constructs can be tested for their ability to rescue the phenotype that was observed in parental KO mESCs. We present a proof-of-principle structure-function study in p120 catenin (p120ctn) KO mESCs using endoderm differentiation in embryoid bodies (EBs) as the phenotypic readout. This approach enables the identification of important domains, putative downstream pathways, and disease-relevant point mutations that underlie KO phenotypes for a given protein.

Proteins often contain multiple domains that can exert different cellular functions. Gene knock-outs (KO) do not consider this functional diversity. Here, we report a recombination-mediated cassette exchange (RMCE)-based structure-function approach in KO embryonic stem cells that allows for the molecular dissection of various functional domains or variants of a protein.

Gene engineering in mouse embryos or embryonic stem cells (mESCs) allows for the study of the function of a given protein. Proteins are the workhorses of ...

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 ...

A Method for Characterizing Embryogenesis in Arabidopsis

Given the highly predictable nature of their development, Arabidopsis embryos have been used as a model for studies of morphogenesis in plants. However, early stage plant embryos are small and contain few cells, making them difficult to observe and analyze. A method is described here for characterizing pattern formation in plant embryos under a microscope using the model organism Arabidopsis. Following the clearance of fresh ovules using Hoyer's solution, the cell number in and morphology of embryos could be observed, and their developmental stage could be determined by differential interference contrast microscopy using a 100X oil immersion lens. In addition, the expression of specific marker proteins tagged with Green Fluorescent Protein (GFP) was monitored to annotate cell identity specification during embryo patterning by confocal laser scanning microscopy. Thus, this method can be used to observe pattern formation in wild-type plant embryos at the cellular and molecular levels, and to characterize the role of specific genes in embryo patterning by comparing pattern formation in embryos from wild-type plants and embryo-lethal mutants. Therefore, the method can be used to characterize embryogenesis in Arabidopsis.

A Method for Characterizing Embryogenesis in Arabidopsis

This protocol outlines a method for observing embryogenesis in Arabidopsis via ovule clearance followed by the inspection of embryo pattern formation under a microscope.

Given the highly predictable nature of their development, Arabidopsis embryos have been used as a model for studies of morphogenesis in plants. However, ...

In Vitro Ovule Cultivation for Live-cell Imaging of Zygote Polarization and Embryo Patterning in Arabidopsis thaliana

In most flowering plants, the zygote and embryo are hidden deep in the mother tissue, and thus it has long been a mystery of how they develop dynamically; for example, how the zygote polarizes to establish the body axis and how the embryo specifies various cell fates during organ formation. This manuscript describes an in vitro ovule culture method to perform live-cell imaging of developing zygotes and embryos of Arabidopsis thaliana. The optimized cultivation medium allows zygotes or early embryos to grow into fertile plants. By combining it with a poly(dimethylsiloxane) (PDMS) micropillar array device, the ovule is held in the liquid medium in the same position. This fixation is crucial to observe the same ovule under a microscope for several days from the zygotic division to the late embryo stage. The resulting live-cell imaging can be used to monitor the real-time dynamics of zygote polarization, such as nuclear migration and cytoskeleton rearrangement, and also the cell division timing and cell fate specification during embryo patterning. Furthermore, this ovule cultivation system can be combined with inhibitor treatments to analyze the effects of various factors on embryo development, and with optical manipulations such as laser disruption to examine the role of cell-cell communication.

In Vitro Ovule Cultivation for Live-cell Imaging of Zygote Polarization and Embryo Patterning in Arabidopsis thaliana

This manuscript describes an in vitro ovule cultivation method that enables live-cell imaging of Arabidopsis zygotes and embryos. This method is utilized to visualize the intracellular dynamics during zygote polarization and the cell fate specification in developing embryos.

In most flowering plants, the zygote and embryo are hidden deep in the mother tissue, and thus it has long been a mystery of how they develop dynamically; ...

Microstructured Devices for Optimized Microinjection and Imaging of Zebrafish Larvae

Zebrafish have emerged as a powerful model of various human diseases and a useful tool for an increasing range of experimental studies, spanning fundamental developmental biology through to large-scale genetic and chemical screens. However, many experiments, especially those related to infection and xenograft models, rely on microinjection and imaging of embryos and larvae, which are laborious techniques that require skill and expertise. To improve the precision and throughput of current microinjection techniques, we developed a series of microstructured devices to orient and stabilize zebrafish embryos at 2 days post fertilization (dpf) in ventral, dorsal, or lateral orientation prior to the procedure. To aid in the imaging of embryos, we also designed a simple device with channels that orient 4 zebrafish laterally in parallel against a glass cover slip. Together, the tools that we present here demonstrate the effectiveness of photolithographic approaches to generate useful devices for the optimization of zebrafish techniques.

Microinjection of zebrafish embryos and larvae is a crucial but challenging technique used in many zebrafish models. Here, we present a range of microscale tools to aid in the stabilization and orientation of zebrafish for both microinjection and imaging.

Zebrafish have emerged as a powerful model of various human diseases and a useful tool for an increasing range of experimental studies, spanning ...

Live Cell Imaging of Chromosome Segregation During Mitosis

Chromosomes must be reliably and uniformly segregated into daughter cells during mitotic cell division. Fidelity of chromosomal segregation is controlled by multiple mechanisms that include the Spindle Assembly Checkpoint (SAC). The SAC is part of a complex feedback system that is responsible for prevention of a cell progress through mitosis unless all chromosomal kinetochores have attached to spindle microtubules. Chromosomal lagging and abnormal chromosome segregation is an indicator of dysfunctional cell cycle control checkpoints and can be used to measure the genomic stability of dividing cells. Deregulation of the SAC can result in the transformation of a normal cell into a malignant cell through the accumulation of errors during chromosomal segregation. Implementation of the SAC and the formation of the kinetochore complex are tightly regulated by interactions between kinases and phosphatase such as Protein Phosphatase 2A (PP2A). This protocol describes live cell imaging of lagging chromosomes in mouse embryonic fibroblasts isolated from mice that had a knockout of the PP2A-B56&#947; regulatory subunit. This method overcomes the shortcomings of other cell cycle control imaging techniques such as flow cytometry or immunocytochemistry that only provide a snapshot of a cell cytokinesis status, instead of a dynamic spatiotemporal visualization of chromosomes during mitosis.

This protocol describes an easy and convenient method to label and visualize live chromosomes in mitotic cells using Histone2B-GFP BacMam 2.0 label and a spinning disc confocal microscopy system.

Chromosomes must be reliably and uniformly segregated into daughter cells during mitotic cell division. Fidelity of chromosomal segregation is controlled ...

Generation of Scaffold-free, Three-dimensional Insulin Expressing Pancreatoids from Mouse Pancreatic Progenitors In Vitro

The pancreas is a complex organ composed of many different cell types that work together to regulate blood glucose homeostasis and digestion. These cell types include enzyme-secreting acinar cells, an arborized ductal system responsible for the transportation of enzymes to the gut, and hormone-producing endocrine cells. 
Endocrine beta-cells are the sole cell type in the body that produce insulin to lower blood glucose levels. Diabetes, a disease characterized by a loss or the dysfunction of beta-cells, is reaching epidemic proportions. Thus, it is essential to establish protocols to investigate beta-cell development that can be used for screening purposes to derive the drug and cell-based therapeutics. While the experimental investigation of mouse development is essential, in vivo studies are laborious and time-consuming. Cultured cells provide a more convenient platform for screening; however, they are unable to maintain the cellular diversity, architectural organization, and cellular interactions found in vivo. Thus, it is essential to develop new tools to investigate pancreatic organogenesis and physiology.
Pancreatic epithelial cells develop in the close association with mesenchyme from the onset of organogenesis as cells organize and differentiate into the complex, physiologically competent adult organ. The pancreatic mesenchyme provides important signals for the endocrine development, many of which are not well understood yet, thus difficult to recapitulate during the in vitro culture. Here, we describe a protocol to culture three-dimensional, cellular complex mouse organoids that retain mesenchyme, termed pancreatoids. The e10.5 murine pancreatic bud is dissected, dissociated, and cultured in a scaffold-free environment. These floating cells self-assemble with mesenchyme enveloping the developing pancreatoid and a robust number of endocrine beta-cells developing along with the acinar and the duct cells. This system can be used to study the cell fate determination, structural organization, and morphogenesis, cell-cell interactions during organogenesis, or for the drug, small molecule, or genetic screening.

Generation of Scaffold-free, Three-dimensional Insulin Expressing Pancreatoids from Mouse Pancreatic Progenitors In Vitro

Here, we present a protocol to generate insulin expressing 3D murine pancreatoids from free-floating e10.5 dissociated pancreatic progenitors and the associated mesenchyme.

The pancreas is a complex organ composed of many different cell types that work together to regulate blood glucose homeostasis and digestion. These cell ...