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

Summary

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.

Abstract

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 "cut and paste" 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 "leapfrog transplants" (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.

Introduction

How genotype encodes phenotype has been a major question in biology since the rediscovery of Mendel'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 species^2,3. The "frog community" 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 animal^{4,5,6,7,8,9,10,11}. 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—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 embryogenesis^14,15,16, 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 "carriers", 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 stages^18,19. For leapfrogging, Blackler's approach was modified by performing the transplantations at the blastula stage¹⁷, when PGCs are localized in the vegetal pole of the embryo^20,21. 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 ("leapfrogged") 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 "host transfer" 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.

Protocol

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

1. 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.
2. Generate sgRNAs using procedures previously described²⁴ .
   NOTE: Briefly, short DNA templates coding for sgRNAs are created using overlapping deoxyoligonucleotides containing 5' bacteriophage T7 promoters, which are annealed and "filled in" 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's instructions²⁴.
3. Shortly before performing microinjections, prepare Cas9-sgRNA complexes by first denaturing ~250 ng of sgRNA in a 3 µL total volume of RNAse-free (diethylpyrocarbonate-treated) H₂O at 60-65 °C for 5 min, followed by quick-cooling on ice for 5 min. Centrifuge the sgRNA for a few seconds, add 1 µL of 1 µg/µL Cas9 protein, and incubate for 10 min at 37 °C to promote complex formation.
   NOTE: Following this incubation, the tube can be kept on ice while preparing the embryos for microinjection.
4. Obtain X. tropicalis embryos by in vitro fertilization using standard methods, as previously described²⁵. De-jelly^25,26 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'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.
5. Immediately transfer the embryos to be injected to a 1% agarose-coated dish containing 1x MMR using a Pasteur pipette.
6. Inject at the 1-cell stage. See references^26,27 for detailed descriptions of Xenopus microinjection methods.
   1. 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 °C. Also create a dish of uninjected sibling embryos that will serve as graft recipients.
7. 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.
   1. 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.
   2. 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 µg of gentamycin sulfate/mL.

2. Transplantation of PGCs

1. 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 °C incubator and allow them to equilibrate to room temperature for an additional 0.5 h.
2. 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.
3. 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.
4. 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.
5. 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.
6. 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.
7. 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.
8. 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.
9. 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.
10. Once the graft has been placed, use a hairloop or the shaft of an eyebrow hair knife to gently slide the "carcass" 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.
11. 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.
12. 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).
13. Continue making transplants until the embryos reach approximately early gastrula stage 10, which is approximately 6.5 hpf in X. tropicalis (see Figure 2E).

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).

1. 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.
   1. 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.
2. Transfer the 24-well plate containing embryos to a 25 °C incubator for overnight culture.
   NOTE: The next day, the embryos will have reached tailbud stages.
3. Move the graft recipients to clean agarose-coated 6-well plates containing 1/9x MMR supplemented with 50 µg of gentamycin sulfate/mL, with 1 embryo per well. Maintain a clear record of donor carcasses that match these graft recipients.
4. To assess mutagenesis efficiency by sequencing PCR amplicons^5,17,24 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 µL lysis buffer²⁴ containing proteinase K (PK) by repeated up-and-down pipetting using a P200 pipette.
5. Perform embryo lysis²⁴ at 56 °C for 6 h to overnight to permit PK digestion. Inactivate PK by heating it to 90-95 °C for 10 min, followed by quick cooling to 4 °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 °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).
6. 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.

Results

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...

Discussion

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...

Materials

List of materials used in this article

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