Obtaining Xenopus laevis embryos

Abstract

The embryos of the African clawed frog, Xenopus laevis, are a powerful substrate for the study of complex fundamental biological and disease mechanisms in neurobiology, physiology, molecular biology, cell biology, and developmental biology. A simple and straightforward technique for generating a large number of developmentally synchronized embryos is In Vitro Fertilization (IVF). IVF permits simultaneous fertilization of thousands of eggs but requires the sacrifice of the parental male, which may not be feasible if the male comes from a stock of precious animals. An alternative to sacrificing a precious male is to use a natural mating which allows for the collection of many embryos with minimal preparation, but with the potential loss of the experimental advantage of developmental synchronization. Here we present both methods for obtaining X. laevis embryos.

MATERIALS

Reagents:

Ethyl 4-aminobenzoate, 98%; aka. Benzocaine (CAS# 94-09- 7)

• Make 10% stock in 95% Ethanol and store at room temperature.

Gentamicin Reagent Solution, 10 mg/mL in distilled water (e.g., ThermoFisher Scientific, Waltham, MA, USA).

Human Chorionic Gonadotropin (hCG) (available from several sources including: National Hormone and Peptide Program, Los Angeles, CA, USA; Sigma-Aldrich, St. Louis, MO, USA; Chorulon brand, Merck Animal Health, Madison, NJ, USA; BioVendor, Asheville, NC, USA)

• hCG should be resuspended to a concentration of 1000 U/mL dissolved in 1x PBS and stored at −20°C.

Ovine Luteinizing Hormone (oLH) (e.g., National Hormone and Peptide Program, Los Angeles, CA, USA)

• oLH should be resuspended to a concentration of 0.4 mg/mL dissolved in 1x PBS and stored at −20°C.

Pregnant Mare Serum Gonadotropin (PMSG) (e.g., BioVendor, Asheville, NC, USA)

• PMSG should be resuspended to a concentration of 100 U/mL dissolved in 1x PBS and stored at −20°C. The use of Pregnant Mare Serum Gonadotropin has been shown to improve follicular maturity.

Marc’s Modified Ringer Solution (MMR):

• Make 10x stock: 1 M NaCl, 20 mM KCl, 10 mM MgSO₄·7H₂O, 20 mM CaCl₂·2H₂O, 50 mM HEPES free acid; adjust the pH to 7.4–7.8 with NaOH. Sterilize in an autoclave. Dilute this stock with autoclaved Type 1 water (ASTM International 2018) as necessary. Following dilution, no further pH adjustment is required.

Modified Barth’s Saline (MBS):

• Make 10x stock: 800 mM NaCl, 10 mM KCl, 10 mM MgSO₄·7H₂O, 50 mM HEPES free acid, 25 mM NaHCO₃; adjust the pH to 7.8 with NaOH. Sterilize in an autoclave. Make a separate 0.1 M stock of CaCl₂, also sterilize in an autoclave. To make culture solutions dilute 10x MBS stock with Type 1 water and add CaCl₂ stock to a final concentration of CaCl₂ in 1x MBS at 0.7 mM.

Phosphate Buffered Saline (PBS) tablets, 100 g.

Equipment:

1ml syringe with PrecisionGlide Needle, 27 G x 3/8" (e.g., Becton, Dickinson and Company, Franklin Lakes, NJ, USA)

Pestle for 1.5ml microcentrifuge tubes (e.g., USA Scientific, Inc., Orlando, FL, USA)

Method:

Testes isolation for in vitro fertilization

1. Anesthetize the male frog via submersion in a bath of 0.15% benzocaine in system water for 15-30 minutes.

   • The male has reached proper anesthetic plane when it is no longer responsive to foot pinching and has lost the swallowing reflex when its throat is rubbed.

2. To ensure euthanasia, place the anesthetized male on a moist paper towel and using surgical scissors perform cervical transection severing the spine at the level of the atlas bone immediately posterior to the skull.

3. With the frog in the supine position (Figure 1A), cut through the skin (Figure 1B, C), fascia, and muscle with fine scissors to expose the lower abdominal cavity (Figure 1D).

   • After cutting through each layer, move it out of the way to allow access to the tissues below and eventually the abdominal cavity. This is a terminal procedure and since the goal is to gain easy access to the abdominal cavity the opening should be made large enough to permit that.
4. Locate and isolate the testes along the dorsal midline of the abdominal cavity (Figure 1E, F).

   • Due to the positioning of the testes on the dorsal side of the cavity, it is often necessary to adjust the position of fat bodies (Figure 1D) to reveal the testes. In a healthy, fully sexually mature male, each testis resembles in shape and size a grain of puffed rice, approximately 7mm long and 3mm in diameter. The testis may be closely associated with the fat bodies. One way to distinguish it is that the color of the testis is much paler then that of fat bodies. The testis is white to beige, not yellow, with a diffuse network of fine red vasculature on the surface.

5. Using scissors, cut the testes away from the surrounding organs.

6. Clean the testes on a dry paper towel or a Kimwipe by removing any attached remnants of fat and viscera and blood (Figure 1G).

   • Removing all tissues besides the testes should aid in fertilization rate and durability during extended storage. Be gentle when handling the testes with forceps, as they can be easily damaged. Keeping them intact before use should also increase their durability.
7. Store the testes in 1x MMR with 10 μg/mL of gentamicin antibiotic at 4°C.

   • High salt prevents the sperm from activating and when stored at 4°C in the presence of the antibiotic the testes will be viable for at least a week. If preferred, MBS can be substituted for MMR, here and wherever MMR is being used in the following steps. Use the same concentration of MBS as that indicated for MMR.

Figure 1. Isolation of testes from an X. laevis male.

(A) A ventral view of a euthanized X. laevis male, the dashed line indicates the general location of the opening that needs to be made to give access to the abdominal cavity. (B) Incision through the skin. (C) The generated flap of skin pulled back to provide access to the layer of ventral abdominal muscle underneath. (D) Both the skin and muscle layers pulled back giving access to the open abdominal cavity, the dashed line outlines the area of the abdominal cavity currently obscured by the fat bodies. (E) The fat bodies on the male’s left side have been pulled aside giving access to the testis indicated by the blue arrow. (F) Both testes, indicated by blue arrows, are visible after fat bodies have been moved aside. (G) Isolated and clean testes in a dish of 1x MMR and gentamicin.

Applying the sperm for in vitro fertilization

• 8.Collect eggs into a sterile petri dish (See obtaining eggs) and remove system or egg laying water that is incidentally transferred.
• 9.
  Cut approximately 1/8 of a testis and transfer it into a 1.5ml microcentrifuge tube with about 1ml of 1x MMR.

  • Alternatively, sperm suspension can be made in 1-2ml 0.1x MMR, which activates the sperm immediately. The remaining part of the testis, if not being used immediately, can be returned to storage as in step 7 above.
• 10.
  Use a microcentrifuge pestle to pulverize the testis piece and then add the sperm solution to the eggs using a transfer pipette.

  • Alternatively, the testis piece can be macerated directly in the dish with forceps.
• 11.With a fine tipped plastic pipette mix the sperm solution with the eggs, if possible spreading them into a monolayer.
• 12.
  After 5 minutes flood the dish with 0.1x MMR to activate the sperm.

  • Fill the dish when flooding.
• 13.After 15 minutes look for egg rotation and animal pole contraction which are good indicators of a successful fertilization.

Priming for natural mating

• 14.
  Perform this 1 to 7 days before planned ovulation

  • The priming injection on its own does not induce egg laying. Instead, it promotes consistent production of a higher number of mature eggs following the boosting injection.
• 15.Remove PMSG stocks from −20°C freezer and allow to thaw at room temperature or 37°C.
• 16.
  Fill a 1ml syringe with 30-50 U of 100 U/mL PMSG.

  • Smaller inbred J-strain females require less hormone and can be primed with 30 U of PMSG while larger wild type females can be given 50 U of PMSG. hCG can also be used to prime at 30 U for a smaller J-strain female and 50 U for a larger wild type female. Although not necessary, males are also sometimes primed to presumably promote sperm maturation. If priming a male, use half the amount of the hormone as used for the female.
• 17.
  Pick up and restrain the frog with one hand in such a way that the dorsal surface of the frog is rested against palm of the hand with the head of the frog pointed towards the wrist (Figure 2A, B). Place the thumb and the middle finger along the posterior sides of the animal abdomen and use the index finger to abduct one of the frog’s hindlimbs (Figure 2C).

  • It is typically easier to handle the animals with one’s dominant hand. If possible, the palm of the hand should cover the frog’s eyes which will help keep it calm. This hold makes the dorsal lymph sac pocket easily accessible while restraining the animal.
• 18.
  With the other hand, insert the needle subcutaneously through the dorsal surface into the dorsal lymph sac in the posterior medial region of the animal proximal to the lateral line "stich marks” and slowly inject the hormone (Figure 2C-E).

  • Insert the needle at a shallow angle so the penetration is superficial to prevent injection into the muscle. Intramuscular injection should have no detrimental effect on animal health but may reduce the desired effect of the hormone.
• 19.
  Return the frog to its permanent tank and do not feed it prior to the boosting injection.

  • Adult frogs can regurgitate food following hormone injection and the presence of solid waste can reduce oocyte durability.

Figure 2. Restraint of an X. laevis female during gonadotropin injection into the dorsal lymph sac.

(A) A dorsal view of a restrained female. (B) A view of the ventral side of a restrained female giving a better picture of the optimal finger position. (C) A female with one of its legs braced against the body resulting in better access for injection into the dorsal lymph sac. (D) A dorsal view of a sexually mature X. laevis female with the general location for hormone injection outlined in red. (E) A female in a more relaxed hold with a needle positioned for injection.

Inducing ovulation and fertilization for natural mating

• 20.
  Perform this the evening before you would like to collect eggs (see note 24 for timing details)

  • The boosting injection promotes the final steps of oocyte maturation and is necessary for induction of ovulation. Females given a boosting injection are likely to spawn, even if they’ve not been given a prior priming injection. The priming injection serves to ensure consistent egg laying response to the boosting injection. Furthermore, in males, the boosting injection seems to encourage the amplexus and the mating behavior, necessary for proper juxtaposition of the male and female cloacas during mating and therefore for rapid and efficient fertilization of eggs as they are being laid by the female.
• 21.Remove oLH or hCG stocks from −20°C freezer and allow to thaw at room temperature or 37°C.
• 22.
  Fill a 1ml syringe with 2 μg of oLH per 1 g of body mass (typically ~200 μg of 0.4 mg/mL oLH) or with 500 U of 1000 U/mL hCG.

  • Smaller inbred J-strain females require less hormone and can be boosted with 120 μg of oLH or 300 U of hCG while larger wild type females will require 200 μg of oLH or 500 U of hCG. Males should be given half the amount of hormone as the females.
• 23.Inject both the male and female (See steps 16-18).
• 24.
  Place both animals in a tank filled with system water and put in an 18°C incubator. Ovulation will begin approximately 8 to 12 hours post injection.

  • When checking on a natural mating in progress, if the male is in amplexus, avoid unnecessarily moving or disturbing the pair as this may cause the male to release the female thus aborting the mating.
• 25.Collect embryos throughout the day and place them in 0.1x MMR with 10 μg/mL of gentamicin in sterile petri dishes.
• 26.
  At the end of the day, transfer the frogs to a clean tank overnight and allow them 24 hours to recover prior to returning them back to the system.

  • Males and females should be isolated from each other to prevent prolonged amplexus which may sometimes cause sores or stress to the female.

Discussion

Healthy and mature X. laevis females can produce in excess of 1500 eggs per spawning that can be easily fertilized via the described methods to generate embryos for use in experiments (Wlizla et al. 2017). The main advantage of using IVF is that it results in synchronized development of the embryos within the batch, which greatly simplifies certain aspects of experimental design and staging as the embryos continue to develop. When using IVF, only a small piece of the testis is required; the rest can be stored in 1x MMR or 1x MBS at 4°C and will maintain viability for at least a week. This permits the same male to be used for several experiments through the week, even after its been euthanized. Nevertheless, use of natural mating may be necessary if the parental male being used is rare or precious. In such a case, it is useful for two investigators to work as a pair if precisely staged embryos during early development are required for the experiment, such as for microinjection at one or two cell stage. One investigator can collect the embryos from the mating tank at regular intervals, select those at the appropriate stage, and perform any additional necessary treatments such as removal of the jelly coat, while the other investigator performs the experimental manipulations such as microinjection. This permits for the maximum use of the embryos generated through natural mating as it may not be trivial for a single person to collect, stage, prepare and perform experiments on embryos as they are being continuously generated. Alternatively, when using precious males, users should also consider the extensive transgenic and mutant stocks available at the National Xenopus Resource (NXR) and the European Xenopus Resource Centre (EXRC) (Horb et al. 2019). Both resource centers can rapidly fulfill requests and ship adult frogs, embryos, isolated testes, and cryopreserved sperm. Finally, regarding potentially limited resources, it is worth pointing out that several different mammalian gonadotropins can be used for the priming and boosting injections. Historically, hCG has been used for both injections, and may be the more humane option when considering effects on animal welfare associated with extraction and purification of gonadotropins from animal sources. However, during a recent commercial shortage of hCG we have demonstrated that use of PMSG for priming and oLH for boosting is just as effective in inducing ovulation (Wlizla et al. 2017). Awareness of such alternatives is necessary for preventing interruptions in research resulting from supply disruptions.