Oocyte Host-Transfer and Maternal mRNA Depletion Experiments in Xenopus

Abstract

This protocol details the oocyte host-transfer method in Xenopus, using transplantation by intraperitoneal injection. This approach is suitable for the overexpression of mRNAs and for the use of antisense oligonucleotides to deplete maternal mRNAs, which are not replaced until zygotic genome activation in the mid-blastula transition. Xenopus oocyte host-transfer can also be used for highly efficient mutagenesis in the F₀ generation by prefertilization injection of genome editing reagents.

MATERIALS

It is essential that you consult the appropriate Material Safety Data Sheets and your institution’s Environmental Health and Safety Office for proper handling of equipment and hazardous material used in this protocol.

RECIPES: Please see the end of this protocol for recipes indicated by <R>. Additional recipes can be found online at http://cshprotocols.cshlp.org/site/recipes.

Reagents

Anesthetic solution (buffered MS222) <R>

Human chorionic gonadotropin (hCG; 10,000 IU/vial; Sigma-Aldrich CG10)

L-15 oocyte culture medium (OCM; 70%) <R>

Marc’s modified Ringer’s (MMR) solution–triple HEPES (10×) <R>

Nucleic acids for microinjection according to experimental goals

Details regarding oligonucleotide design, modification and use in oocytes for host-transfer are described elsewhere (Hulstrand et al. 2010; Olson et al. 2012).

Progesterone (10 mm in 100% ethanol, stored at 20˚C; Sigma-Aldrich P8783)

Tap water conditioned with AmQuel or other chloramine remover

Vital dye stocks <R>

Xenopus females for use as oocyte donors and as transfer hosts

Xenopus ovarian tissue (Nasco; optional, see Step 1)

Equipment

Containers for frogs (buckets or small tubs)

Dumont #4 or #5 forceps (Fine Science Tools)

Glass syringe with Luer lock adaptor (2 mL; Tomopal Inc. 140–1502) Incubator (18°C)

Kimwipes (large, sterilized)

Microcentrifuge tubes (1.5 mL)

Pasteur pipettes (fire-polished glass, sterile) Petri dishes (plastic 100 and 60 mm diameter) Rocking platform

Sterile syringe needles (16 gauge, 1 inch [16G1]; BD 305197) Teflon pestles for 1.5 mL microcentrifuge tubes (RPI 199228)

METHOD

The steps for the entire procedure are outlined in Figure 1.

FIGURE 1.

Steps for culturing and transferring oocytes by intraperitoneal injection. Up to several days before the day embryos are desired, isolate oocytes by manual defolliculation and inject with experimental antisense oligonucleotides and/or mRNAs. On the evening before host-transfer, stimulate the oocytes to mature by adding progesterone (Pg) to the medium. At the same time, inject presumptive host females with human chorionic gonadotropin (hCG) to induce ovulation. On the morning of the transfer day, sort healthy mature oocytes with visible “white spots” from germinal vesicle breakdown and color with desired vital dyes. The chosen host female is anesthetized and colored oocytes are transplanted using intraperitoneal (IP) injection. Three hours later, experimental oocytes are recovered and analyzed according to experimental goals.

Culture of Experimental Oocytes

• 1
  Prepare ovarian tissue surgically from anesthetized females as described in Schneider et al. (2010) and Olson et al. (2012) and place in OCM at 18°C.

  Alternatively, Xenopus ovarian tissue can be obtained commercially.

• 2
  Manually defolliculate 300–500+ oocytes using Dumont forceps. Dissect oocytes by tearing the follicle layer near the region where oocytes are attached to the ovary, using very light grip pressure on the forceps. Store oocytes in OCM at 18°C and begin experiments within 1–2 d.

  Frogs stimulated with hGC within 3 mo should not be used as oocyte donors. Healthy, high-quality oocytes are essential for successful host-transfer experiments. These should be fully grown and free of any damage to the membrane. Practice is essential for the rapid collection of many good quality oocytes and is necessary for good results with this method. For detailed descriptions of manual defolliculation as well as videos see: Smith et al. (1991), www.youtube.com/watch?v=us8rDNG69Sk(Manual defolliculation 2009), Protocol: Isolation of Xenopus Oocytes (Sive et al. 2010a), Hulstrand et al. (2010), Schneider et al. (2010), Olson et al. (2012), and Protocol: Isolation of Xenopus Oocytes (Newman et al. 2018).

• 3
  Microinject oocytes directly while in OCM; Ficoll is not needed. Transfer the oocytes to fresh OCM after injection and culture for up to 3 d at 18°C. Include appropriate controls, such as uninjected oocytes and mRNA rescued oocytes, in the case of oligonucleotide depletions, or control mRNAs for overexpression experiments.

  For a detailed description of microinjection see: Protocol: Microinjection of Xenopus Oocytes (Sive et al. 2010b). Antisense oligonucleotides are typically injected 36–48 h before fertilization, whereas mRNAs can be injected the day before host transfer and fertilization. Typical doses for antisense oligonucleotides are 2–6 ng for modified DNA-based oligonucleotides and 10–50+ ng for morpholino oligonucleotides. The optimal dose should be empirically determined in pilot studies. For overexpression/gain-of-function studies, high doses of mRNAs of 1 ng or more can be used. When rescuing a mRNA knockdown effect, minimal doses are used (20–200 pg), ideally below the phenotypic threshold, so that rescue of the depletion can be accurately assessed as opposed to an ectopic overexpression effect. For genome editing, equimolar amounts ( 2 µM each) of Cas9 protein and appropriate guide RNAs are assembled for 10 min at 37°C and 300 pg sgRNA/1.5 ng Cas9 protein are injected.

In Vitro Oocyte Maturation and Preparation of Host Female

• 4At the end of the day before transfer, add progesterone to each dish of oocytes ( ~75–200 in 8 mL OCM per dish). Use 1.6 µL of 10 mM progesterone stock per 8 mL OCM (2 µM final concentration). Swirl to mix and incubate overnight at 18°C. Also, place the anesthetic solution to warm overnight at 18°C.
• 5
  Prime two-to-three prospective host females by injecting human chorionic gonadotropin (hCG, ~1000 U) into the dorsal lymph sac to induce ovulation. This is best done 10–12 h before oocyte implantation.

  Albino females may be used as hosts to facilitate identification of transferred oocytes. Albinos may also be used as oocyte donors if desired, and if identifying animal-vegetal poles is not critical.

• 6
  Check the oocytes for maturation, white spots indicating germinal vesicle breakdown.

  If oocytes are stuck together, gently tease them apart otherwise this will cause blockage during the transfer. See Troubleshooting.

• 7
  Thaw the vital dye stocks and centrifuge briefly when thawed. Add 80 µL of dye stock to each group of oocytes in 8 mL OCM and swirl gently to mix.

  Only six different colors can be reliably made using standard vital dyes. For up to three groups, use the single dyes, Brown, Blue, and Red. Dyes can be combined to generate additional colors (Brown + Blue = Green [good for uninjected controls], Blue + Red = Mauve, Brown + Red = Orange [often hard to distinguish from either plain Brown or Red]).

• 8
  Incubate the oocytes in vital dyes with rocking for 10+ min at room temperature and then wash all oocytes together in a larger volume of OCM. However, make sure the colors are easily distinguishable before doing this.

  If oocyte colors are not easily distinguished, extend the staining time or wash separately and transfer into different host females.

• 9
  Place a host female into anesthetic solution for 5–10 min until she is nonresponsive to a toe-pinch or to being held upside-down.

  This can be done while oocytes are coloring. Choose a host that is laying healthy eggs and that has not yet ovulated many of her own eggs. Avoid females with stringy eggs or that are not obviously extruding eggs when squeezed.

Intraperitoneal Injection of Donor Oocytes

• 10Place the anesthetized female in dorsal recumbency on a damp Kimwipe.
• 11Prepare the injection syringe by fitting a new 16G1 needle to the syringe. Set aside the plunger. Wash the inside of the syringe and needle with OCM to coat the surface. This reduces the potential for oocyte sticking.
• 12Hold the empty syringe with needle next to the abdomen of the frog. With the beveled edge up, insert the needle tip at a 45° angle, making sure to penetrate skin and muscle. The skin surface can be braced with a gloved finger adjacent to the injection site if the needle does not pass smoothly.
• 13Hold the inserted syringe with the nondominant hand. Collect colored oocytes using a sterile, fire-polished glass Pasteur pipette in the dominant hand and introduce into the open end of the syringe. Oocytes will drain down the side and pool at the bottom but should not flow into the needle right away.
• 14Elevate the end of the syringe so it is perpendicular to the frog surface, keeping the needle in the frog. Oocytes should immediately flow by gravity into the needle and body cavity. If not, tap the side of the syringe gently. If oocytes are trapped around the outlet port flush them with additional OCM. Keep the overall volume to a minimum because excess OCM (>3–4 mL) can cause the frog to cease egg laying.
• 15Return the syringe to its original insertion angle and withdraw the needle. The oocytes should not spill out from the incision, although some OCM may leak out initially. The needle hole does not need to be sutured. Rinse the host female in water and allow recovery in a shallow volume of water. Once recovered, move to a larger container. Recovery is indicated by gulping and eye-bulging.
• 16Observe the host for resumption of egg-laying. This should occur within 20–30 min and the donor colored eggs should appear 2–3 h after implantation.

In Vitro Fertilization of Host-Transferred Oocytes

• 17Isolate testes from a male frog through nonsurvival surgery as described in Protocol: Isolating Xenopus laevis Testes (Sive et al. 2007a). Store the testes in a Petri dish of OCM at room temperature for use the same day or at 4°C for up to 1 wk.
• 18Prepare a sperm suspension by homogenizing ~¼ of a testis in 2–3 mL 0.3× MMR.
• 19
  Manually squeeze eggs from the female into a Petri dish or place the female in high salt MMR (1.2× MMR) solution to collect eggs. Rinse eggs collected in 1.2× MMR in 0.3× MMR. Blot off excess liquid and fertilize by adding the sperm suspension (from Step 18). Incubate fertilized eggs at the desired temperature and handle as normal embryos.

  See Troubleshooting.

• 20
  Dejelly cleaving eggs at the 2–4 cell stage using 2% cysteine as described in Protocol: Dejellying Xenopus laevis Embryos (Sive et al. 2007b), and sort when colors are easily distinguished.

  See Troubleshooting.

TROUBLESHOOTING

Problem (Step 6):

Oocytes fail to mature after progesterone treatment.

Solution:

Suspect poor oocyte quality (see Protocol: Isolation of Xenopus Oocytes [Newman et al. 2018]). Oocytes can be tested for maturation by treating a small batch with progesterone immediately after isolation before proceeding. If the ovary is deemed unsuitable, select another female and perform another surgery.

Problem (Step 19):

Experimental oocytes fail to be oviposited by the host female.

Solution:

This can occur if the female stops laying eggs for any reason. There is no good recourse for recovering eggs from the body cavity or oviducts; therefore, this situation is best prevented by selecting a host female that has just begun to oviposit eggs on her own (without squeezing) and that is laying high quality eggs. Rapid changes in water temperature and remaining under anesthesia for too long are also correlated with early cessation of egg-laying.

Problem (Step 20):

Experimental oocytes fail to fertilize and develop.

Solution:

This is the most common problem with this method and can generally be avoided by using only healthy, undamaged donor oocytes obtained by careful manual defolliculation. Also, avoid contamination of the OCM and be sure that MMR is correctly made and is at pH 7.6–7.8. Host-transferred oocytes also require higher amounts of sperm and/or longer incubation in the sperm suspension compared to a typical in vitro fertilization in Xenopus.

DISCUSSION

The early development of amphibians relies on maternal factors stored and localized in the egg during oogenesis (for review, see Houston 2013) and Xenopus is advantageous for studying the roles of these maternal gene products. Classic studies in other amphibians (Rugh 1935; Humphries 1956; Aplington 1957; Arnold and Shaver 1962; Lavin 1964; Smith et al. 1968) laid the foundation for Xenopus oocytes to be cultured, manipulated in vitro, and fertilized following transfer into the body cavity of a host female (Brun 1975). These host-transferred eggs are transported by peritoneal cilia to the ostium and oviduct, acquiring surface modifications and jelly coats needed for fertilization (Rugh 1935). This method was revived by Heasman and Wylie (Holwill et al. 1987; Heasman et al. 1991; Torpey et al. 1992) and coupled with antisense oligonucleotide injection into oocytes to elucidate basic mechanisms of axis formation, germ layer patterning and germline specification (for review, see Heasman 2006). Additionally, prefertilization injection of genome editing reagents has recently been used to generate highly efficient mutagenesis in the F₀ generation (Miyamoto et al. 2015a,b; Nakajima and Yaoita 2015; Ratzan et al. 2017; Aslan et al. 2017), providing a renewed rationale for performing the host-transfer method.

RECIPES

Anesthetic Solution (Buffered MS222)

Dissolve 2 g of MS222 (3-aminobenzoic acid ethyl ester; Sigma-Aldrich A5040) in 2 L of AmQuel-treated tap water. Add 1.4 g sodium bicarbonate; the resulting pH should be 7.0. Store in a foil-covered container at 4°C for up to 1 mo. Warm to room temperature before use.

L-15 Oocyte Culture Medium (OCM; 70%)

Using an autoclaved glass cylinder, add 214 mL of deionized distilled water (16–18 MΩ·cm at 25°C) to 500 mL of Leibovitz’s L-15 medium containing 2 mM l-glutamine (Gibco/Thermo-Fisher 11415064) in a sterile bottle or in the sterile L-15 bottle itself. Add 0.3 g of bovine serum albumin (BSA, fraction V) and 3.6 mL of 200× Penicillin-Streptomycin solution (10,000 U/mL, 200× stock; Gibco/Thermo-Fisher 15140122). Adjust the pH to 7.6–7.8 with 5 n NaOH (3–4 drops). (The phenol red indicator dye should be dark red/light purple. The pH is best checked by measuring a small sample in a separate beaker to avoid contaminating the OCM with the pH electrode.) Make fresh weekly and store at 16°C–18°C (or at 4°C for longer term storage).

This version of OCM with 70% L-15 results in more robust oocytes compared with other OCM recipes. The BSA in this recipe can be substituted with 0.05% poly(vinyl alcohol) (Sigma-Aldrich P8136) with equivalent results.

Marc’s Modified Ringer’s (MMR) Solution–Triple HEPES (10×)

Reagent	Quantity	Final concentration (10×)
NaCl	58.44 g	1 m
KCl	1.34 g	18 mm
CaCl2•2H2O	2.94 g	20 mm
MgCl2•6H2O	2.03 g	10 mm
HEPES (molecular grade acid, not the sodium salt)	35.75 g	150 mm

Combine the above solids in ~700 mL of deionized distilled water (16–18 MΩ·cm at 25°C). Adjust pH to 7.6–7.8 with NaOH, and bring the volume to 1 L. Filter-sterilize but do not autoclave. Store at 4°C. Readjust to pH 7.6–7.8 after dilution to 1× (or to 1.2× for high-salt MMR).

Vital Dye Stocks

Add the following amounts of dye powder to 50 mL of sterile deionized water in separate 50-mL Falcon tubes:

• 0.5 g/50 mL Bismarck Brown (Sigma-Aldrich B2759),

• 0.125 g/50 mL Neutral Red (Sigma-Aldrich 861251), and

• 0.05 g/50 mL Nile Blue A (Nile Blue sulfate; Sigma-Aldrich N0766).

Rock for 2 h to dissolve, and then centrifuge at ~3000g. (The final concentrations will be 1%, 0.25%, and 0.1% for Bismarck Brown, Neutral Red, and Nile Blue A, respectively.) Aliquot the supernatant in 1.5-mL microcentrifuge tubes (~1-mL aliquots) and store at −20°C.