This work was performed with the support of a grant, 5R21HD080684-02, from the National Institute of Child Health and Human Development. The author wishes to thank Ken Cho for his continuing enthusiasm and support. The author would also like to acknowledge Bruce Blumberg for use of his camera, Rebekah Charney, for the critical reading of the manuscript, and Sean McNamara and Marcin Wlizla at the National Xenopus Resource (RRID:SCR_013731), for the valuable conversations regarding X. tropicalis feeding and animal care regimens.

All methods described here have been approved by the Institutional Animal Care and Use Committee of the University of California, Irvine.
1. Preparations for PGC Transplantation
<ol>
	<li>Prepare dissection tools in advance, as previously described23. 
	NOTE: Eyebrow hair knives are used to make incisions with the assistance of a hair loop to stabilize the embryo while performing the surgeries. Eyebrow hairs and hair loops are glued into borosilicate glass Pasteur pipettes that have first been drawn in a flame and broken on the thinned portion (see Figure 1A, arrowhead) to create a shorter, narrower tip (Figure 1B) for greater dexterity when performing surgeries.</li>
	<li>Generate sgRNAs using procedures previously described24 . 
	NOTE: Briefly, short DNA templates coding for sgRNAs are created using overlapping deoxyoligonucleotides containing 5&#39; bacteriophage T7 promoters, which are annealed and &#34;filled in&#34; by an error-free, thermostable DNA polymerase. In vitro sgRNA synthesis reactions are incubated for several hours to overnight (depending on the kit used). Reactions are DNAse-treated to remove the template. The sgRNAs are purified by standard methods of phenol/chloroform extraction and ammonium acetate/isopropanol precipitation, according to the kit manufacturer&#39;s instructions24.</li>
	<li>Shortly before performing microinjections, prepare Cas9-sgRNA complexes by first denaturing ~250 ng of sgRNA in a 3 &#181;L total volume of RNAse-free (diethylpyrocarbonate-treated) H2O at 60-65 &#176;C for 5 min, followed by quick-cooling on ice for 5 min. Centrifuge the sgRNA for a few seconds, add 1 &#181;L of 1 &#181;g/&#181;L Cas9 protein, and incubate for 10 min at 37 &#176;C to promote complex formation. 
	NOTE: Following this incubation, the tube can be kept on ice while preparing the embryos for microinjection.</li>
	<li>Obtain X. tropicalis embryos by in vitro fertilization using standard methods, as previously described25. De-jelly25,26 the embryos at 10 min post-fertilization. 
	NOTE: Fertilization time is considered the time after the sperm suspension is added to the eggs and the dish is flooded with 1/9thX Marc&#39;s Modified Ringers (MMR)26. For X. tropicalis25, unlike X. laevis, perform the de-jellying in 1/9x MMR containing 3% cysteine free base (not the HCl salt), pH adjusted to 7.8-8.0, followed by multiple washes in 1/9x MMR. Transfer the embryos to an agarose (1%)-coated dish containing 1/9x MMR at room temperature.</li>
	<li>Immediately transfer the embryos to be injected to a 1% agarose-coated dish containing 1x MMR using a Pasteur pipette.</li>
	<li>Inject at the 1-cell stage. See references26,27 for detailed descriptions of Xenopus microinjection methods.
	<ol>
		<li>Inject each embryo at a single site in the animal pole, with 4 nL of Cas9-sgRNA complex (final amount = 1 ng of Cas9 and ~250 pg of sgRNA; see the Discussion)24. After 10-20 min of injection site healing, transfer the embryos to an agarose-coated dish containing 1/9x MMR and incubate at 25 &#176;C. Also create a dish of uninjected sibling embryos that will serve as graft recipients.</li>
	</ol>
	</li>
	<li>Prepare Petri plates (60 mm) that contain a ~5 mm-thick layer of 1% agarose made in 0.3x MMR, if doing transplantations in X. tropicalis, or 1x MMR for X. laevis.
	<ol>
		<li>Create depressions ~3-4 mm deep by inserting a 3- x 4-well mold (created by cutting a 96-well PCR plate) into the molten agarose when pouring the plates (see Figure 1C and D). Hold the mold several millimeters above the bottom of the Petri dish using a clamp attached to a ring stand until the agarose has hardened.</li>
		<li>Additionally, make a 24-well plate to house individual embryos by coating the wells with a thin layer of 1% agarose made in 1/9x MMR. 
		NOTE: The culture medium added to these wells is 1/9x MMR supplemented with 50 &#181;g of gentamycin sulfate/mL.</li>
	</ol>
	</li>
</ol>
2. Transplantation of PGCs
<ol>
	<li>When the embryos just reach Nieuwkoop and Faber28 blastula stage 9 (Figure 2A), ~4.5 h post-fertilization (hpf) for X. tropicalis, remove them from the 25 &#176;C incubator and allow them to equilibrate to room temperature for an additional 0.5 h.</li>
	<li>To begin the transplantation at 5 hpf (Figure 2B), use a Pasteur pipette to transfer one PGC donor embryo (injected with Cas9-sgRNA) to the 60 mm agarose dish containing depressions (created in step 1.7) in 0.3x MMR (or 1x MMR if using X. laevis). Also transfer one uninjected sibling embryo, the graft recipient, to this dish. 
	NOTE: It is critical that the identities of each of these embryos are not confused.</li>
	<li>Manually remove the vitelline envelopes from each embryo using forceps and rotate both embryos so that their vegetal poles are in view and accessible for surgery. 
	NOTE: A description of manual de-vitellination of embryos has previously been described26.</li>
	<li>Using the sharp tip of an eyebrow hair knife, make four shallow incisions in the shape of a square on the vegetal pole of the recipient embryo, inside the zone where future bottle cells that mark the blastopore will form. Make the incisions by inserting the tip of the eyebrow hair knife into the embryo, just below the surface, and make upward slicing movements while stabilizing the embryo with the hair loop. 
	NOTE: Figure 2 shows vegetal views of an embryo at half-hourly intervals from 4.5-7.0 hpf to show the cell size and to provide a guide for estimating where the bottles cells will form (dashed white circle). The aim is to make incisions where the dashed black box is indicated.</li>
	<li>Once the four sides of the square are delineated by incisions, deepen each incision with the eyebrow hair knife using similar cutting motions to reach a depth of approximately 1/3 to 1/2 of the distance to the blastocoel floor.</li>
	<li>Free the vegetal tissue explant from the recipient embryo (Figure 3A and B) by making a horizontal (parallel to the vegetal surface) cut(s) in the deep vegetal region. 
	NOTE: The size of the PGC-containing vegetal explant is approximately 0.4-0.45 mm per side and 0.25-0.3 mm in depth. This vegetal explant from the recipient embryo is no longer needed and should therefore be set aside to be discarded.</li>
	<li>Working quickly, repeat this procedure (steps 2.4-2.6) on the PGC donor embryo to create a similarly sized vegetal tissue fragment for transplantation. 
	NOTE: It is important to carry out this second dissection with little delay to minimize the time for the recipient embryo to heal its open wound.</li>
	<li>Once the tissue containing PGCs is removed from the donor embryo, use the eyebrow hair knife and hair loop to move this explant into position in the opening created in the recipient embryo. 
	NOTE: The interior surface of the donor graft must be facing the interior of the recipient embryo. It is not necessary to match the orientations of the dorsal-ventral and left-right axes of the graft with the recipient embryo.</li>
	<li>Use the long edge of the shaft of the eyebrow hair knife, held parallel to the surface of the graft, to gently press the graft into the opening in the vegetal surface of the recipient embryo.</li>
	<li>Once the graft has been placed, use a hairloop or the shaft of an eyebrow hair knife to gently slide the &#34;carcass&#34; of the donor embryo across the agarose surface and into an agarose depression. Make sure that the open wound of the carcass is facing the bulk liquid. If not, use the eyebrow hair knife to rotate the embryo to achieve this orientation.</li>
	<li>Slide the recipient embryo, with the graft healing in place, into an adjacent depression and likewise make sure that the grafted vegetal pole tissue is facing the bulk liquid.</li>
	<li>Repeat this procedure (steps 2.1-2.11) using another pair of donor and recipient embryos to create another transplant/carcass pair; transfer these to empty depressions. 
	NOTE: It is critical that one makes notations to keep track of matched pairs of carcasses and transplant-bearing embryos. The donor carcasses will be used as a proxy for the efficiency of Cas9-sgRNA-induced mutagenesis in the transplanted PGCs, which cannot be easily assayed until the transplant-bearing animals reach sexual maturity (see step 3.3, below, and the Discussion).</li>
	<li>Continue making transplants until the embryos reach approximately early gastrula stage 10, which is approximately 6.5 hpf in X. tropicalis (see Figure 2E).</li>
</ol>
3. Post-healing Care of Embryos
NOTE: Grafts heal into place within ~30 min, and it is normal to observe some yolky debris from cell lysis exuding from the embryo (Figure 3C).
<ol>
	<li>Once healed, use a Pasteur pipette to very gently transfer embryos from the depressions to individual wells of an agarose-coated 24-well plate. The exudate will be removed (Figure 3D) by fluid mixing during the transfer.
	<ol>
		<li>Rotate the embryos so that they are placed vegetal-pole up, facing the bulk solution. Again, place the donor carcasses and graft recipients in adjacent wells to assist in keeping track of embryo pairs; record this information.</li>
	</ol>
	</li>
	<li>Transfer the 24-well plate containing embryos to a 25 &#176;C incubator for overnight culture. 
	NOTE: The next day, the embryos will have reached tailbud stages.</li>
	<li>Move the graft recipients to clean agarose-coated 6-well plates containing 1/9x MMR supplemented with 50 &#181;g of gentamycin sulfate/mL, with 1 embryo per well. Maintain a clear record of donor carcasses that match these graft recipients.</li>
	<li>To assess mutagenesis efficiency by sequencing PCR amplicons5,17,24 or using other methods that rely on PCR amplicons (see the Discussion), move individual carcasses to 0.2 mL PCR strip tubes. Remove most of the medium and homogenize in 100 &#181;L lysis buffer24 containing proteinase K (PK) by repeated up-and-down pipetting using a P200 pipette.</li>
	<li>Perform embryo lysis24 at 56 &#176;C for 6 h to overnight to permit PK digestion. Inactivate PK by heating it to 90-95 &#176;C for 10 min, followed by quick cooling to 4 &#176;C. Use lysates without further cleanup steps to seed the PCR reactions to obtain short amplicons for direct Sanger DNA sequencing24. Store the lysates at -20 &#176;C. 
	NOTE: Within several days, the tadpoles will reach feeding stages (approximately stage 45-46) and can be fed a suspension of planktonic powder (sera Micron).</li>
	<li>After ~1 week, with daily changes of culture medium to maintain cleanliness and minimize microbial overgrowth, move tadpoles to small tanks in a circulating aquatic system with drip flow. Use the sequencing data to segregate tadpoles with highly efficiently mutagenized PGCs from tadpoles with lower mutagenesis efficiency. Once metamorphosis is complete, move the froglets to the adult aquatic system in the lab. 
	NOTE: Regimens developed by the National Xenopus Resource (Marine Biology Laboratory, Woods Hole, MA) provide a guide for tadpole feeding29 and adult frog maintenance30.</li>
</ol>

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K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=CRISPR/Cas9:+An+inexpensive,+efficient+loss+of+function+tool+to+screen+human+disease+genes+in+Xenopus.">CRISPR/Cas9: An inexpensive, efficient loss of function tool to screen human disease genes in Xenopus.</a> Dev Biol. 408 (2), 196-204 (2015).</li><li>Yu, C., Zhang, Y., Yao, S., Wei, Y. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+PCR+Based+Protocol+for+Detecting+Indel+Mutations+Induced+by+TALENs+and+CRISPR/Cas9+in+Zebrafish.">A PCR Based Protocol for Detecting Indel Mutations Induced by TALENs and CRISPR/Cas9 in Zebrafish.</a> PLoS ONE. 9 (6), e98282 (2014).</li><li>Mock, U., Hauber, I., Fehse, B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Digital+PCR+to+assess+gene-editing+frequencies+(GEF-dPCR)+mediated+by+designer+nucleases.">Digital PCR to assess gene-editing frequencies (GEF-dPCR) mediated by designer nucleases.</a> Nat Protoc. 11 (3), 598-615 (2016).</li><li>Varshney, G. K., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=High-throughput+gene+targeting+and+phenotyping+in+zebrafish+using+CRISPR/Cas9.">High-throughput gene targeting and phenotyping in zebrafish using CRISPR/Cas9.</a> Genome Res. 25 (7), 1030-1042 (2015).</li><li>Shi, J., Wang, E., Milazzo, J. P., Wang, Z., Kinney, J. B., Vakoc, C. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Discovery+of+cancer+drug+targets+by+CRISPR-Cas9+screening+of+protein+domains.">Discovery of cancer drug targets by CRISPR-Cas9 screening of protein domains.</a> Nat Biotechnol. 33 (6), 661-667 (2015).</li><li>Koebernick, K., Loeber, J., Arthur, P. K., Tarbashevich, K., Pieler, T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Elr-type+proteins+protect+Xenopus+Dead+end+mRNA+from+miR-18-mediated+clearance+in+the+soma.">Elr-type proteins protect Xenopus Dead end mRNA from miR-18-mediated clearance in the soma.</a> Proc Natl Acad Sci USA. 107, 16148-16153 (2010).</li><li>Yang, J., Aguero, T., King, M. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+Xenopus+maternal-to-zygotic+transition+from+the+perspective+of+the+germline.">The Xenopus maternal-to-zygotic transition from the perspective of the germline.</a> Curr Top Dev Biol. 113, 271-303 (2015).</li></ol>

The author has nothing to disclose.

This report provides a detailed protocol for the transplantation of vegetal tissue containing PGCs. Transplantation of PGCs is used in conjunction with genome-editing technologies (e.g., CRISPR/Cas9) to modify the germline of an animal while maintaining nearly all of its somatic tissues as genetically wild type. For leapfrogging to be successful, there are a number of critical factors to consider prior to performing performing transplantations.
To ensure the complete replacement of th...

How genotype encodes phenotype has been a major question in biology since the rediscovery of Mendel&#39;s laws. An understanding of the roles of genes, their regulation and interactions within gene networks, and the functions of encoded products promises to provide tools for uncovering new biology and ameliorating disease states. For over half a century1, the African clawed frog, Xenopus, has been a leading model for studies on a wide variety of topics in basic biology and biomedicine, including the genetic control of development. Historically, most research on Xenopus has used the allotetraploid frog, X. laevis, but more recently, due to its diploidy, X. tropicalis has been developed as an amphibian genetic model. Complete genome sequences have been assembled from both Xenopus species2,3. The &#34;frog community&#34; is now at a turning point where basic genome modification technology permits the study of gene function, virtually at will. Programmable CRISPR/Cas9 endonucleases have made the mutagenesis of genes highly efficient, with biallelic mutation possible in most cells of the animal4,5,6,7,8,9,10,11. These studies, underscored by Bhattacharya et al.12 and Shigeta et al.13, have shown that the function of many genes can be studied by mutagenesis in F0 mosaic animals. This approach has many advantages; however, Cas9-sgRNA-microinjected embryos often display variable phenotypes due to incomplete loss of function (LOF). More significantly, the generation of mutant lines is highly advantageous for some applications&#8212;in particular when studying genes that have a maternal mRNA contribution. Maternal mRNAs and proteins, and their influences on epigenetics, persist for an extended period into embryogenesis14,15,16, rendering the early developmental contributions of many genes refractory to F0 analyses. Therefore, other LOF approaches are required.
When seeking to create mutant lines, the path to obtaining homozygous LOF embryos has several obstacles. First, efficient mutagenesis to produce biallelic mutations can be disadvantageous because loss of essential gene functions results in failure to survive to sexual maturity, interfering with the production of a viable line. A common solution is the careful titration of the amount of Cas9-sgRNA delivered. Here, the goal is to achieve a balance between reducing lethality while also maximizing germline mutagenesis efficiency. A second problem arises from the standard breeding scheme, where phenotypic analyses are deferred until the F2 generation. Using the standard approach, sexually mature F0 animals that transmit mutant alleles through the germline are outcrossed to produce F1 heterozygous &#34;carriers&#34;, which are then grown to sexual maturity. Two F1 heterozygotes are then intercrossed to produce F2 mutant embryos at an expected Mendelian frequency of 25%. Thus, two generations of breeding are necessary for analysis of mutant phenotypes. Mutant animals could be either homozygotes or compound heterozygotes (i.e., progeny containing two different mutant alleles, which depends on the genotypes of the parental animals used in the F1 intercross).
These obstacles can be overcome by confining programmable nuclease-mediated mutagenesis to germ cells, which underlies a new method called leapfrogging17. Leapfrogging has two main components: (1) the microinjection of Cas mRNA together with sgRNA, or nuclease-sgRNA complexes (or, in principle, TALENs or zinc finger nucleases) at the single-cell stage to efficiently mutagenize embryonic genomes, followed by (2) the transplantation of PGCs into wildtype sibling embryos, where the endogenous PGCs were removed. When both steps are efficient, complete germline replacement with mutant germ cells can be obtained. Blackler demonstrated in the early 1960s that Xenopus PGCs could be transplanted between embryos at the late neurula and early tailbud stages18,19. For leapfrogging, Blackler&#39;s approach was modified by performing the transplantations at the blastula stage17, when PGCs are localized in the vegetal pole of the embryo20,21. Transplantation before gastrulation has two main advantages. First, the engraftment of transplants and the subsequent normal development was found to be more efficient when the transplantation is performed at the blastula stage (unpublished observations). Second, by performing blastula-stage transplantations shortly after zygotic transcription has begun, one can avoid the lethality arising from developmental gene mutations that disrupt gastrulation or that otherwise lead to malformed late neurulae. PGC transplant-bearing (&#34;leapfrogged&#34;) embryos are grown to sexual maturity, and intercrosses between these F0 animals have demonstrated that, in many cases, 100% of the F1 progeny display the LOF phenotype (most being compound heterozygotes), indicating complete germline replacement with the targeted mutations.
It is expected that leapfrogging will accelerate genetic approaches in Xenopus. Leapfrogging also provides an alternative to the &#34;host transfer&#34; method22 for the LOF analysis of maternal-effect genes (unpublished observations). In the current publication, a detailed description of the method, especially focusing on PGC transplantation, is presented in X. tropicalis (with minor modifications for X. laevis). The transplantation of PGCs is demonstrated here to facilitate a more rapid transfer of this technology to other laboratories working with Xenopus. The principles of this method should be successful in other amphibians (e.g., urodeles), and organism-specific modifications in the methodology should allow for application to many other animals in which efficient mutagenesis can be accomplished and PGCs are readily transplantable.

<table><tbody><tr><td>Dumont #5 forceps</td><td>Fine Science Tools</td><td>11252-20 or 11252-30</td><td></td></tr><tr><td>Eyebrow hair knife</td><td></td><td></td><td>Homemade</td></tr><tr><td>Hair loop</td><td></td><td></td><td>Homemade</td></tr><tr><td>Pasteur pipettes, borosilicate glass</td><td>Fisher Scientific</td><td>13-678-20A</td><td></td></tr><tr><td>Krazy Glue (Cyanoacrylate-based)</td><td>Elmer&#039;s Products, Inc.</td><td>KG581</td><td>To affix eyebrow hair and hair loops into Pasteur pipettes, other similar glues can be used.</td></tr><tr><td>Oligodeoxynucleotides, custom ordered</td><td>Integrated DNA Technologies</td><td>Custom ordered</td><td>Template oligos for sgRNA synthesis, see Nakayama et al., 2014 for design details.</td></tr><tr><td>Megascript T7 kit</td><td>Ambion/ThermoFisher</td><td>AM1334</td><td>Or use Megashortscript T7 (AM1354) kit</td></tr><tr><td>Phenol, Tris buffered</td><td></td><td></td><td>High quality distilled phenol from any commercial supplier</td></tr><tr><td>Chloroform</td><td></td><td></td><td>High quality chloroform from any commercial supplier</td></tr><tr><td>Ethanol</td><td></td><td></td><td>High quality ethanol from any commercial supplier</td></tr><tr><td>Diethylpyrocarbonate (DEPC)</td><td>Sigma Chemical Co.</td><td>D5758-50ML</td><td></td></tr><tr><td>Cas9 protein, with nuclear localization signal</td><td>PNA Bio, Inc.</td><td>CP01</td><td>Reconstituted using DEPC-treated water according to manufacturer&#039;s recommendations</td></tr><tr><td>10X Marc&#039;s Modified Ringers solution</td><td></td><td></td><td>Homemade. Recipe (ref 26) for 1X MMR (lacking EDTA) is 100mM NaCl, 2 mM KCl, 1 mM MgSO4, 2 mM CaCl2, 5 mM HEPES. Solution is pH adjusted to 7.4.</td></tr><tr><td>Agarose</td><td></td><td></td><td>Any Molecular Biology grade agarose is sufficient</td></tr><tr><td>60 X15 mm Petri plates</td><td>Falcon</td><td>351007</td><td></td></tr><tr><td>24-well plates</td><td>Falcon</td><td>3047</td><td></td></tr><tr><td>L-Cysteine</td><td>Sigma Chemical Co.</td><td>C7352-100G</td><td>Free base, not HCl salt</td></tr><tr><td>Proteinase K</td><td>Roche</td><td>03 115 828 001</td><td>Typically ~20mg/ml from Roche</td></tr><tr><td>sera Micron</td><td>sera</td><td></td><td>Tadpole food. Resuspend in growth medium. Can be purchased from a variety of online retailers</td></tr></tbody></table>

primordial germ cell transplantation for crispr/cas9-based leapfrogging in xenopus

The creation of mutant lines by genome editing is accelerating genetic analysis in many organisms. CRISPR/Cas9 methods have been adapted for use in the African clawed frog, Xenopus, a longstanding model organism for biomedical research. Traditional breeding schemes for creating homozygous mutant lines with CRISPR/Cas9-targeted mutagenesis have several time-consuming and laborious steps. To facilitate the creation of mutant embryos, particularly to overcome the obstacles associated with knocking out genes that are essential for embryogenesis, a new method called leapfrogging was developed. This technique leverages the robustness of Xenopus embryos to &#34;cut and paste&#34; embryological methods. Leapfrogging utilizes the transfer of primordial germ cells (PGCs) from efficiently-mutagenized donor embryos into PGC-ablated wildtype siblings. This method allows for the efficient mutation of essential genes by creating chimeric animals with wildtype somatic cells that carry a mutant germline. When two F0 animals carrying &#34;leapfrog transplants&#34; (i.e., mutant germ cells) are intercrossed, they produce homozygous, or compound heterozygous, null F1 embryos, thus saving a full generation time to obtain phenotypic data. Leapfrogging also provides a new approach for analyzing maternal effect genes, which are refractory to F0 phenotypic analysis following CRISPR/Cas9 mutagenesis. This manuscript details the method of leapfrogging, with special emphasis on how to successfully perform PGC transplantation.

Following transplantation, the qualitative determination of the efficacy of CRISPR/Cas9 mutagenesis should be performed before expending the effort in animal husbandry to grow and maintain the animals to sexual maturity. Because animals carrying leapfrog transplants are somatically wildtype and the germline is difficult to access for direct measurements, saving the carcasses of donor embryos becomes important. DNA analysis from the carcasses serves as a proxy for the extent of mutagenesis...

Watch this Scientific Journal Video about Primordial Germ Cell Transplantation for CRISPR/Cas9-based Leapfrogging in Xenopus at JoVE.com

Primordial Germ Cell Transplantation for CRISPR/Cas9-based Leapfrogging in Xenopus

Genes essential for survival pose technical hurdles for creating mutant lines. Leapfrogging circumvents lethality by combining genome editing with primordial germ cell transplantation to create wild-type animals carrying germline mutations. Leapfrogging also permits the efficient generation of homozygous null mutants in the F1 generation. Here, the transplantation step is demonstrated.

Primordial Germ Cell Transplantation for CRISPR/Cas9-based Leapfrogging in Xenopus

The creation of mutant lines by genome editing is accelerating genetic analysis in many organisms. CRISPR/Cas9 methods have been adapted for use in the ...

primordial-germ-cell-transplantation-for-crisprcas9-based

Research

JoVE Journal

Genetics

8.3K Views.  University of California, Irvine. Genen essentieel voor overleving vormen technische hindernissen voor het maken van mutant lijnen. Sprongen omzeilt letaliteit door het combineren van genoom bewerken met primordiale geslachtscellen transplantatie wild-type dieren uitvoering germline mutaties maken. Sprongen maakt ook de efficiënte generatie homozygoot null mutanten in de F1 generatie. Hier, wordt de transplantatie stap aangetoond.

Video: Primordial Germ Cell Transplantation for CRISPR/Cas9-based Leapfrogging in Xenopus

Manipulation and In Vitro Maturation of Xenopus laevis Oocytes, Followed by Intracytoplasmic Sperm Injection, to Study Embryonic Development

Amphibian eggs have been widely used to study embryonic development. Early embryonic development is driven by maternally stored factors accumulated during oogenesis. In order to study roles of such maternal factors in early embryonic development, it is desirable to manipulate their functions from the very beginning of embryonic development. Conventional ways of gene interference are achieved by injection of antisense oligonucleotides (oligos) or mRNA into fertilized eggs, enabling under- or over-expression of specific proteins, respectively. However, these methods normally require more than several hours until protein expression is affected, and, hence, the interference of gene functions is not effective during early embryonic stages. Here, we introduce an experimental system in which expression levels of maternal proteins can be altered before fertilization. Xenopus laevis oocytes obtained from ovaries are defolliculated by incubating with enzymes. Antisense oligos or mRNAs are injected into defolliculated oocytes at the germinal vesicle (GV) stage. These oocytes are in vitro matured to eggs at the metaphase II (MII) stage, followed by intracytoplasmic sperm injection (ICSI). By this way, up to 10% of ICSI embryos can reach the swimming tadpole stage, thus allowing functional tests of specific gene knockdown or overexpression. This approach can be a useful way to study roles of maternally stored factors in early embryonic development.

Manipulation and In Vitro Maturation of Xenopus laevis Oocytes, Followed by Intracytoplasmic Sperm Injection, to Study Embryonic Development

We describe methods of manipulating Xenopus laevis immature oocytes, in vitro maturation of oocytes to eggs, and intracytoplasmic sperm injection. This protocol allows degradation of some maternal proteins and overexpression of genes of interest at fertilization, and hence is valuable to study roles of specific factors in early embryonic development.

Amphibian eggs have been widely used to study embryonic development. Early embryonic development is driven by maternally stored factors accumulated during ...

Developmental Biology

Stem cell-like Xenopus Embryonic Explants to Study Early Neural Developmental Features In Vitro and In Vivo

Understanding the genetic programs underlying neural development is an important goal of developmental and stem cell biology. In the amphibian blastula, cells from the roof of the blastocoel are pluripotent. These cells can be isolated, and programmed to generate various tissues through manipulation of genes expression or induction by morphogens. In this manuscript protocols are described for the use of Xenopus laevis blastocoel roof explants as an assay system to investigate key in vivo and in vitro features of early neural development. These protocols allow the investigation of fate acquisition, cell migration behaviors, and cell autonomous and non-autonomous properties. The blastocoel roof explants can be cultured in a serum-free defined medium and grafted into host embryos. This transplantation into an embryo allows the investigation of the long-term lineage commitment, the inductive properties, and the behavior of transplanted cells in vivo. These assays can be exploited to investigate molecular mechanisms, cellular processes and gene regulatory networks underlying neural development. In the context of regenerative medicine, these assays provide a means to generate neural-derived cell types in vitro that could be used in drug screening.

Stem cell-like Xenopus Embryonic Explants to Study Early Neural Developmental Features In Vitro and In Vivo

In Xenopus embryos, cells from the roof of the blastocoel are pluripotent and can be programmed to generate various tissues. Here, we describe protocols to use amphibian blastocoel roof explants as an assay system to investigate key in vivo and in vitro features of early neural development.

Understanding the genetic programs underlying neural development is an important goal of developmental and stem cell biology. In the amphibian blastula, ...

Functional Cloning Using a Xenopus Oocyte Expression System

Identification of genes responsible for embryonic induction poses a number of challenges; to name a few, secreted molecules of interest may be low in abundance, may not be secreted but tethered to the signaling cell(s), or may require the presence of binding partners or upstream regulatory molecules. Thus in a search for gene products capable of eliciting an early lens-inductive response in competent ectoderm, we utilized an expression cloning system that would allow identification of paracrine or juxtacrine factors as well as transcriptional or other regulatory proteins. Pools of mRNA were injected into Xenopus oocytes, and responding tissue placed directly on the oocytes and co-cultured. Following functional cloning of ldb1 from a neural plate stage cDNA library based on its ability to elicit the expression of the early lens placode marker foxe3 in lens-competent animal cap ectoderm, we characterized the mRNA expression pattern, and assayed developmental progression following overexpression or knockdown of ldb1. This system is suitable in a very wide variety of contexts where identification of an inducer or its upstream regulatory molecules is sought using a functional response in competent tissue.

Functional Cloning Using a Xenopus Oocyte Expression System

We describe a Xenopus oocyte and animal cap system for the expression cloning of genes capable of inducing a response in competent ectoderm, and discuss techniques for the subsequent analysis of such genes. This system is useful in the functional identification of a wide range of gene products.

Identification of genes responsible for embryonic induction poses a number of challenges; to name a few, secreted molecules of interest may be low in ...

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 ...

Generation of Chimeric Axolotls with Mutant Haploid Limbs Through Embryonic Grafting

A growing set of genetic techniques and resources enable researchers to probe the molecular origins of the ability of some species of salamanders, such as axolotls, to regenerate entire limbs as adults. Here, we outline techniques used to generate chimeric axolotls with Cas9-mutagenized haploid forelimbs that can be used for exploring gene function and the fidelity of limb regeneration. We combine several embryological and genetic techniques, including haploid generation via in vitro activation, CRISPR/Cas9 mutagenesis, and tissue grafting into one protocol to produce a unique system for haploid genetic screening in a model organism of regeneration. This strategy reduces the number of animals, space, and time required for the functional analysis of genes in limb regeneration. This also permits the investigation of regeneration-specific functions of genes that may be required for other essential processes, such as organogenesis, tissue morphogenesis, and other essential embryonic processes. The method described here is a unique platform for conducting haploid genetic screening in a vertebrate model system.

This goal of this protocol is to produce chimeric axolotls with haploid forelimbs derived from Cas9-mutagenized donor tissue using embryonic tissue grafting techniques.

A growing set of genetic techniques and resources enable researchers to probe the molecular origins of the ability of some species of salamanders, such as ...

Generating Chimeric Zebrafish Embryos by Transplantation

One of the most powerful tools used to gain insight into complex developmental processes is the analysis of chimeric embryos. A chimera is defined as an organism that contains cells from more than one animal; mosaics are one type of chimera in which cells from more than one genotype are mixed, usually wild-type and mutant. In the zebrafish, chimeras can be readily made by transplantation of cells from a donor embryo into a host embryo at the appropriate embryonic stage. Labeled donor cells are generated by injection of a lineage marker, such as a fluorescent dye, into the one-cell stage embryo. Labeled donor cells are removed from donor embryos and introduced into unlabeled host embryos using an oil-controlled glass pipette mounted on either a compound or dissecting microscope. Donor cells can in some cases be targeted to a specific region or tissue of the developing blastula or gastrula stage host embryo by choosing a transplantation site in the host embryo based on well-established fate maps.

A step-by-step guide to generating targeted chimeric zebrafish embryos by transplantation at the blastula or gastrula stage.

One of the most powerful tools used to gain insight into complex developmental processes is the analysis of chimeric embryos. A chimera is defined as an ...

Biology

Fertilization of Xenopus oocytes using the Host Transfer Method

Studying the contribution of maternally inherited molecules to vertebrate early development is often hampered by the time and expense necessary to generate maternal-effect mutant animals. Additionally, many of the techniques to overexpress or inhibit gene function in organisms such as Xenopus and zebrafish fail to sufficiently target critical maternal signaling pathways, such as Wnt signaling. In Xenopus, manipulating gene function in cultured oocytes and subsequently fertilizing them can ameliorate these problems to some extent. Oocytes are manually defolliculated from donor ovary tissue, injected or treated in culture as desired, and then stimulated with progesterone to induce maturation. Next, the oocytes are introduced into the body cavity of an ovulating host female frog, whereupon they will be translocated through the host's oviduct and acquire modifications and jelly coats necessary for fertilization. The resulting embryos can then be raised to the desired stage and analyzed for the effects of any experimental perturbations. This host-transfer method has been highly effective in uncovering basic mechanisms of early development and allows a wide range of experimental possibilities not available in any other vertebrate model organism.

Fertilization of Xenopus oocytes using the Host Transfer Method

Procedure for fertilizing Xenopus oocytes by the host transfer method.

Studying the contribution of maternally inherited molecules to vertebrate early development is often hampered by the time and expense necessary to ...

Production of Transgenic Xenopus laevis by Restriction Enzyme Mediated Integration and Nuclear Transplantation

Stable integration of cloned gene products into the Xenopus genome is necessary to control the time and place of expression, to express genes at later stages of embryonic development, and to define how enhancers and promoters regulate gene expression within the embryo. The protocol demonstrated here can be used to efficiently produce transgenic Xenopus laevis embryos. This transgenesis approach involves three parts: 1. Sperm nuclei are isolated from adult X. laevis testis by treatment with lysolecithin, which permeabilizes the sperm plasma membrane. 2. Egg extract is prepared by low speed centrifugation, addition of calcium to cause the extract to progress to interphase of the cell cycle, and a high-speed centrifugation to isolate interphase cytosol. 3. Nuclear transplantation: the nuclei and extract are combined with the linearized plasmid DNA to be introduced as the transgene and a small amount of restriction enzyme. During a short reaction, egg extract partially decondenses the sperm chromatin and the restriction enzyme generates chromosomal breaks that promote recombination of the transgene into the genome. The treated sperm nuclei are then transplanted into unfertilized eggs. Integration of the transgene usually occurs prior to the first embryonic cleavage such that the resulting embryos are not chimeric. These embryos can be analyzed without any need to breed to the next generation, allowing for efficient and rapid generation of transgenic embryos for analyses of promoter and gene function. Adult X. laevis resulting from this procedure also propagate the transgene through the germline and can be used to generate lines of transgenic animals for multiple purposes.

Production of Transgenic Xenopus laevis by Restriction Enzyme Mediated Integration and Nuclear Transplantation

This video protocol demonstrates a method for generating transgenic Xenopus laevis by introduction of transgenes into sperm nuclei followed by nuclear transplantation into unfertilized eggs.

Stable integration of cloned gene products into the Xenopus genome is necessary to control the time and place of expression, to express genes at later ...

Application of Mouse Parthenogenetic Haploid Embryonic Stem Cells as a Substitute of Sperm

In organisms with sexual reproduction, germ cells are the source of totipotent cells that develop into new individuals. In mice, fertilization of an oocyte by a spermatozoon creates a totipotent zygote. Recently, several publications have reported that haploid embryonic stem cells (haESCs) can be a substitute for gametic genomes and contribute to embryos, which develop into mice. Here, we present a protocol to apply parthenogenetic haESCs as a substitute of sperm to construct embryos by intracytoplasmic injection into oocytes. This protocol consists of steps for preparing haESCs as sperm replacement, for injection of haESC chromosomes into oocytes, and for culture of semi-cloned embryos. The embryos can yield fertile semi-cloned mice after embryo transfer. Using haESCs as sperm replacement facilitates genome editing in the germline, studies of embryonic development, and investigation of genomic imprinting.

This article aims to demonstrate the use of parthenogenetic haploid embryonic stem cells as a substitute for sperm for the construction of semi-cloned embryos.

In organisms with sexual reproduction, germ cells are the source of totipotent cells that develop into new individuals. In mice, fertilization of an ...

A Simple and Effective Transplantation Device for Zebrafish Embryos

Classical embryological manipulations, such as removing cells and transplanting cells within or between embryos, are powerful techniques to study complex developmental processes. Zebrafish embryos are ideally suited for these manipulations since they are easily accessible, relatively large in size, and transparent. However, previously developed devices for cell removal and transplantation are cumbersome to use or expensive to purchase. In contrast, the transplantation device presented here is economical, easy to assemble, and simple to use. In this protocol, we first introduce the handling of the transplantation device as well as its assembly from commercially and widely available parts. We then present three applications for its use: generation of ectopic clones to study signal dispersal from localized sources, extirpation of cells to produce size-reduced embryos, and germline transplantation to generate maternal-zygotic mutants. Finally, we show that the tool can also be used for embryological manipulations in other species such as the Japanese rice fish medaka.

Embryological manipulations such as extirpation and transplantation of cells are important tools to study early development. This protocol describes a simple and effective transplantation device to perform these manipulations in zebrafish embryos.

Classical embryological manipulations, such as removing cells and transplanting cells within or between embryos, are powerful techniques to study complex ...

Primordial Germ Cell Transplantation for CRISPR/Cas9-based Leapfrogging in Xenopus

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