Jun 16, 2026

Producing Xenopus eggs and embryos

DOI
https://dx.doi.org/10.17504/protocols.io.81wgboqjnlpk/v1
  • Alice Blythe1,2,
  • Matt Guille1,2,
  • Esther Pearl3,
  • Billie Dolphin1,2,
  • Gretel Nicholson1,2,
  • Anna Noble1,2
  • 1University of Portsmouth;
  • 2EXRC;
  • 3The National Centre for the Replacement, Refinement and Reduction of Animals in Research
  • Alice Blythe: https://orcid.org/0009-0004-2629-2475;
  • Esther Pearl: https://orcid.org/0000-0001-6510-0959;
matthew.guille
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DOI: https://dx.doi.org/10.17504/protocols.io.81wgboqjnlpk/v1
Protocol Citation: Alice Blythe, Matt Guille, Esther Pearl, Billie Dolphin, Gretel Nicholson, Anna Noble 2026. Producing Xenopus eggs and embryos. protocols.io https://dx.doi.org/10.17504/protocols.io.81wgboqjnlpk/v1
License: This is an open access  protocol  distributed under the terms of the  Creative Commons Attribution License,  which permits unrestricted use, distribution, and reproduction in any medium, provided the original author and source are credited
Protocol status: Working
This protocol is currently in use at the European Xenopus Resource Centre (EXRC) as of 2026.
Created: February 11, 2026
Last Modified: June 16, 2026
Protocol  Integer ID: 243033
Keywords: Xenopus, Ovulation, Post-ovulation monitoring, IVF, Natural mating, collecting xenopus embryo, embryos for both xenopus laevi, xenopus embryo, xenopus tropicalis embryo, embryos xenopus laevi, producing xenopus egg, xenopus egg, selection of healthy xenopus female, healthy xenopus female, embryos for the experimental use, tropicalis embryo, xenopus tropicali, consistent embryo production, ethical use of xenopus, embryo, egg collection via ivf, xenopus laevi, decades by the european xenopus resource centre, xenopus, european xenopus resource centre, collection of fresh sperm, hormonal induction for ovulation, fresh sperm, positive welfare environment for the frog, natural mating, fertilisation, frozen sperm, ovulation, frog, egg collection, hormonal induction, ivf, developmental biology
Disclaimer
This is a preprint of the following chapter
Authors: Alice Blythe, Anna Noble, Gretel Nicholson, Billie Dolphin, Esther J. Pearl and Matthew Guille
Chapter: Chapter 2 - Producing Xenopus eggs and embryos
Book title: Xenopus protocols (part of the Methods in Molecular Biology series)
Edited by Caroline W. Beck
Publisher: Springer Nature.
Expected publication in 2026. Once the book is published a link to the final version will be added to a version of this protocol.
This is the version of the author's manuscript prior to acceptance for publication and has not undergone editorial and/ or peer review on behalf of the publisher.
Abstract
Xenopus laevis and Xenopus tropicalis embryos are widely used in developmental biology, genetics and disease modelling, and can be generated through in vitro fertilisation (IVF), fertilising with either fresh or cryogenically frozen sperm, or by natural matings. This chapter describes protocols developed and used successfully over two decades by the European Xenopus Resource Centre (EXRC) in Portsmouth, United Kingdom to consistently and reliably induce ovulation and generate embryos for both Xenopus laevis and Xenopus tropicalis.
The following protocols describe the complete workflow for collecting Xenopus embryos, beginning from the selection of healthy Xenopus females, through to the collection of fresh sperm, hormonal induction for ovulation, egg collection via IVF and natural matings, and preparation and selection of embryos for the experimental uses described in this volume. The chapter also contains post-ovulation monitoring for both species of Xenopus females to promote a positive welfare environment for the frogs, as well as good welfare standards for the community.
The following methods support consistent embryo production across a range of experimental applications, including those listed in this volume, whilst promoting best practice in husbandry and ethical use of Xenopus.
Guidelines
These are the current protocols used by the European Xenopus Resource Centre (EXRC) based in Portsmouth, UK. The EXRC protocols are periodically reviewed as best practices evolve, this chapter represents the protocols in use in 2025.
Materials
Selecting and Ovulating Xenopus
  1. Nitrile Gl1. Nitrile gloves (for handling frogs)
  2. hCG (human chorionic gonadotrophin) freeze-dried powder, stock prepared at 1500 IU/mL (we use Chorulon from Intervet). Store this at 4oC for up to 48 hours and warmed to ambient before use.
  3. 1 mL syringes
  4. 27-gauge syringe needle (X. laevis). (The EXRC are currently transitioning to use 27-guage needles from 23G).
  5. 30-gauge syringe needle (X. tropicalis)
  6. Modified Marc’s Modified Ringer’s (1x MMR): 0.1 M NaCl, 2 mM KCl, 1 mM MgSO4, 2 mM CaCl2, 5 mM HEPES (pH 7.8), adjust to pH 7.4 with NaOH, OR
  7. Modified Barth’s Saline (1x MBS): 88 mM NaCl, 1 mM KCl, 2.4 mM NaHCO3, 0.82 mM MgSO4 2H2O, 0.33 mM Ca(NO3)2.2H2O, 0.41 mM CaCl2.6H2O, 10 mM HEPES, adjust pH to 7.5 with NaOH
  8. Opaque 16 L bucket and lid (with holes in the lid for air)
  9. 10 cm petri dishes
  10. 50 mL centrifuge (Falcon) tubes

Fertilising and preparing Xenopus eggs and embryos for experiments (IVF and Natural Matings)
  1. Sperm/testes (IVF only) see Chapter 8 to prepare cryogenically frozen sperm.
  2. Surgical scissors (if isolating testes for fresh sperm)
  3. Forceps (if isolating testes for fresh sperm)
  4. 1x MBS (if using fresh sperm only)
  5. 2% (w/v) Tricaine/MS222 (killing Xenopus males for fresh sperm)
  6. Wide-end 200 µL pipette tips
  7. 0.1x MMR/MBS (X. laevis) or 0.05x MMR (X. tropicalis). See above
  8. 2% (w/v) L-cysteine, pH to 8 with NaOH
  9. Incubator at 18°C (X. laevis) or 24°C (X. tropicalis)
  10. 3 mL disposable plastic transfer pipettes (sterile) OR 3 mL reusable glass pasture pipettes (sterile)
  11. Petri dishes lined with 1% agarose in 0.05x MMR (X. tropicalis only)
  12. Stereo dissecting microscope, with a cold light source.
Before start
Please ensure you have reviewed this protocol thoroughly and wear appropriate PPE.

This protocol contains references to other Chapters due to be published in Xenopus protocols (part of the Methods in Molecular Biology series, Springer Nature) in 2026. In the final, edited publication, there will also be reference to figures that show appropriate frog handling/squeezing techniques, location of testes and how to determine egg quality. Please contact the corresponding author if you wish for more details and access to these figures.
Introduction
X. laevis and X. tropicalis have been used as research animals for decades. One of the benefits of the model system is the ability to produce many embryos per clutch that can be grown in petri dishes. Ex utero development of embryos enables research on different developmental stages without having to kill multiple females at different points in pregnancy, reducing the number of adult animals required for each experiment. The large number of embryos produced per female also reduces the number of females needed to generate sufficient embryos for large experiments. Embryos can be generated via natural matings, where male and female frogs are induced to mate and embryos are collected afterwards, or via in vitro fertilisation, where a male frog is killed and his sperm used to fertilise eggs collected from a female frog. For all procedures, the animal welfare standards agreed by the appropriate ethical review body must be applied. The embryos generated are robust and can then be manipulated during early development for experiments described throughout this volume. This includes, but is not limited to, microinjection (for gain and loss of function experiments), tissue explants, microsurgery, and disease modelling.

In vitro fertilisation (IVF) has long been considered the gold standard for obtaining synchronously dividing embryos, suitable for microinjection or micromanipulation. Female Xenopus frogs can be induced to lay eggs by hormone injection; human chorionic gonadotrophin (hCG) is most common (1,2) but luteinising hormone can also be used (3). Post-injection the females naturally begin to ovulate, and the eggs can be collected on demand by gently massaging the sides of the body (colloquially referred to as “squeezing”) (4). Alternatively, for X. laevis, female frogs can be placed in a salt solution where they will spontaneously lay eggs. The eggs are easily collected in petri dishes and can be fertilised using either freshly harvested or cryopreserved sperm (5–8) from a humanely killed male.

Natural matings are another method to obtain Xenopus embryos. Using this method, both a male and a female Xenopus are hormonally primed and placed together. The male will clasp on to the female for the duration of a day and will fertilise the eggs as she lays them throughout the day (amplexus). This results in clutches of embryos at various developmental stages.

Natural matings are a gentler method than IVF, since no abdominal massage is involved and male frogs are not killed. It is often used for generating wild-type stock or F1 animals from genetically altered lines. This method is recommended when there are a limited number of males or if synchronous embryos at early developmental stages are not required, it is appropriate for experiments like late-stage whole-mount in situ.

This chapter describes the current and complete ovulation and mating protocols used by the European Xenopus Resource Centre (EXRC) based in Portsmouth, UK. It outlines the selection of healthy Xenopus females for ovulation and provides a clear comparison of injection protocols for X. laevis and X. tropicalis, supported by comparative tables. Methods for isolating and using fresh sperm for IVF to produce embryos are described, alongside the less invasive method of natural matings. In support of downstream experiments presented in this volume, this chapter describes the preparation and identification of fertilised eggs and early-stage embryos. Finally, the EXRC have provided details of post-ovulation monitoring in both Xenopus species, with the aim of promoting high welfare standards within the Xenopus community. The EXRC periodically reviews protocols as best practices evolve; this chapter represents the protocols in use in 2025.
Methods
Selecting female X. laevis or X. tropicalis to ovulate
Ensure that female Xenopus are ready to be ovulated. All frogs should be identifiable for health records (see MiMB Chapter 1 sections 3.5.2 and 3.5.3). X. laevis can be easily identified by their dorsal skin patterns. X. tropicalis, however, look extremely similar to one another. Instead, they can have microchips implanted to distinguish individuals (see MiMB Chapter 1 section 3.5.3). Individual animals can be tracked in a database or logbook with the timing of past ovulations, weight, overall health, as well as the quantity and quality of eggs recorded.

  1. Ensure all frogs are individually identifiable (see MiMB Chapter 1 sections 3.2.5 and 3.5.2) and tracked in a database or logbook with health status, the timing of past ovulations as well as the quantity and quality of eggs recorded.
  2. Use the database or logbook to make sure that the chosen frogs have been rested for >90 days since their last ovulation and have been ovulated 16 or fewer times (or equivalent limits from local regulations/ethical review bodies).
  3. Check the weight of the frog against its history (frogs can be weighed in a suitable bucket or beaker of water that is slightly smaller than their outstretched legs, they quickly stay still). If a X. laevis has lost more than 20% of its body weight, they should not be ovulated. If a X. tropicalis has lost more than 10% of its body weight, they should not be ovulated.
  4. Compare the frog’s body shape with its history, a healthy frog should appear rounded, almost plump. Females should appear pear shaped.
  5. Check the frog’s skin texture. It should be smooth and free from abrasions, sores and redness.
  6. If you are in any doubt about the health status of the frog, contact an experienced user, the facility manager or a veterinary surgeon.
  7. All frog use should be recorded in the facility database or logbook.
Isolating Xenopus testes for fresh sperm
If you are using fresh sperm, prepare it in advance. X. laevis testis can be prepared up to two weeks in advance. X. tropicalis testis should be prepared as close to egg collection as possible, ideally no more than 1 hour. Once the male Xenopus has been humanely killed, the dissected testes are put into 1X MBS (see Note 1). More details on dissecting males for their testis can be found in MiMB Chapter 8.

  1. Always wear gloves when handling the frogs.
  2. Humanely kill a male Xenopus with darkened nuptial pads (indicating sexual maturity) using Tricaine/MS222 overdose, followed by an appropriate secondary method of killing (see MiMB Chapter 1 section 3.2) before sperm collection.
  3. Using a pair of fine scissors, cut through both layers of skin to expose the abdominal cavity. Pull the fat bodies to either side of the cavity to expose the testes. See Figure 1.
  4. Testes are usually oval and a pale yellow/off-white located near the kidneys. Remove the testes by dissection, roll them gently on a fresh paper towel to remove lipid and blood vessels and remnants of other non-testicular tissues.
  5. X. laevis testes can be stored at 4°C in 1x MBS until needed and can be kept up to 2 weeks with no recorded loss of fertility. X. tropicalis testes should be stored at 14°C and used as soon as possible (ideally immediately) since they lose ability to fertilise eggs quickly, an effect that is exacerbated at lower temperatures.

Figure 1: Dissection of male Xenopus testes for isolation of fresh sperm. A dead Xenopus male is rested in the supine position, ready for dissection (A). Once the body cavity has been opened (B) and the fat bodies (labelled ‘F’) pulled to the side, the testes (labelled ‘T’) are visible, located just above the kidney. Once dissected out, each testis is cleaned to remove remnants of non-testicular tissue and blood (C).

Ovulating Xenopus females for IVF
To induce ovulation in Xenopus females, hCG hormone is injected into the dorsal lymph sac. The person performing the injection must be trained and signed off as competent to comply with local regulations. It is important not to reuse needles as each use blunts the needle; therefore, reusing a needle to inject a frog can cause more damage and pain than a new needle would. Piercing the vial of Chorulon (hCG) counts as one use.

Table 1: Hormone doses, needle guages and sperm requirements for IVF in X. laevis and X. tropicalis
ABCDE
X. laevis (18.5 °C) X. tropicalis (25.5 °C)
Concentration of hormone Time/Day Concentration of hormone Time/day
Priming dose N/A N/A 10 IUThe evening before egg collection (4pm)
Boosting dose 600 IUThe evening before egg collection (4pm) 100 IUThe morning of egg collection (8am)
Laying begins 7:30am 12:30pm
Needle size 27-gauge Drawing up hCG stock/solvent AND All frog injections 27-guage To draw up hCG solution/solvent only
30-guageAll frog injections
Testes collection (for fresh sperm) 1/5 to 1/3 of a testis in 1X MBS Can be collected up to 2 weeks in advance. 1 whole testis in 1X MBS Should be collected as close to egg collection as possible.
Table 1: A table to describe and compare the injection concentration of hCG, timings, needle sizes and fresh sperm collection between Xla and Xtr for IVF. When following the protocols listed in 3.3. Ovulating Xenopus females, please refer to this table to administer the correct dose, at the correct time, with the correct needles. This table can be used to adapt the timings of injections if you need to make a more suitable time, i.e. egg collection for Xtr to be 2 hours later, females can be e administered the boost at 10:30 am.


Inducing ovulation in Xenopus females

  1. Always wear gloves when handling the frogs.
  2. The evening before egg collection, prepare and administer one full dose (see Note 2) of hCG for X. laevis OR the priming dose for X. tropicalis. Add 1 mL of solvent to 1 vial of freeze dried hCG (1500 IU/mL stock solution) using a 27-gauge needle attached to a 1 mL syringe and mix. Let the solution warm to room temperature.
  3. Dilute the stock to the correct dose concentration. See Table 2.1 and Table 2.3 for guidance.
  4. Remove the syringe (see Note 3) and swap the needle (piercing the vial blunts the needle and counts as one use) for a suitable needle for injection. See Table 2.1.
  5. Remove any air bubbles from the syringe. See Note 4.
  6. Gently remove the frog from the tank, holding it firmly. The dorsal side of the frog should rest against the palm, with its face towards the wrist. Restrain the frog with the thumb and middle finger wrapped around the posterior sides of the abdomen. Put the index finger between the legs. The little finger can be gently placed ventrally at the throat to further restrain the female. See Figure 2.

Figure 2: An appropriate handling technique of Xla (A&B) and Xtr (C&D) from dorsal (A&C) and ventral (B&D) view. The frog is being held firmly, but is relaxed in the hand and not struggling. The thumb and the middle finger are wrapped around the abdomen, and the index finger is placed between the legs. For Xla, little finger is placed on the throat to further restrain the frog.

7. Lie the frog on a damp paper towel and fold up the sides to cover the head and the legs. Always wrap the head first, to relax the frog. Tear a hole in the paper towel to expose the site of injection.
8. Insert the needle at a shallow angle, subcutaneously through the dorsal surface, proximal to the lateral lines. See Figure 3. Inject the hCG solution slowly into the dorsal lymph sac. See Table 2.1.
9. Ensure a new needle is used for every injection.

Figure 3: The location for injection of hCG into the dorsal lymph sac of a Xenopus is easily found on the dorsal side (A). Just above the junction of the back leg (labelled 'L') and the body is a semicircle of slits (B) called lateral lines (labelled 'LL'), or ‘stitching’. Below this, a membrane layer joins the outer skin to the body wall, when the frog’s skin is gently pinched for injection (C), it is easy to differentiate the two layers. The needle is inserted at a shallow angle, almost parallel to the frog proximal to the lateral line. This ensures the needle stays between the two layers. The figure demonstrates the rotation of the frog (craniocaudal and dorsoventral rotation).

10. If the injection site bleeds after injection, hold a piece of paper towel over the area and apply gentle pressure for 30 seconds until the bleeding has stopped.
11. Place the frogs into an allocated, clearly marked tank, in which they can be identified, ready for egg collection. (See Chapter 1, husbandry labels).
12. Freeze any leftover hCG stock. See Note 5. For X. laevis go to Section 2.4, for X. tropicalis, continue to Step 13.
13. The boosting dose can be prepared to the correct dilution the day before or the same day it is administered. See Table 2.1. and Table 2.3. for guidance. If pre-prepared, store in the fridge at 4°C.
14. Warm the hCG to room temperature in the morning, before injecting into the frog. See Note 6.
15. Remove the syringe (see Note 3) and swap needle to a 30-gauge needle. Remove any air bubbles in the syringe.
16. Using the handling and injection techniques described in Steps 6-10 slowly administer the hCG.
17. Place the frog into a singly housed temporary holding tank and wait for her to start laying and an indication that her eggs are ready to be collected. See Note 7.
Egg Collection

When the frogs are ready to lay, their cloaca will appear swollen and red. Eggs are collected hourly by gently massaging the abdomen of the female frog to induce her to release her eggs, known as a ‘squeeze’. X. laevis females can be squeezed up to 6 times. Each X. tropicalis female should be squeezed no more than three times. Allow 45 minutes between squeezes for X. laevis, and at least an hour between squeezes for X. tropicalis. Alternatively, for X. laevis, place each female into a 1.5 L bath of 1x MMR (or 100 mM NaCl) to allow natural laying, and collect her eggs hourly with a pipette. Either way, collect eggs into a Petri dish and ensure most of the liquid is removed before fertilising. Ensure fresh testes or frozen sperm has been acquired in advance and is ready ahead of egg collection. See 5.2 Isolating Xenopus testes for fresh sperm or MiMB Chapter 8 Cryopreservation of Xenopus sperm.

  1. Label clean Petri dishes with the Frog ID/ strain number for identification.
  2. Always wear gloves when handling the frogs.
  3. Remove the injected frogs (using the handling instructions from 5.2. Inducing ovulation, Step 6) from their tank and put them into short-term housing. For X. laevis we recommend putting them into 16 L buckets, filled ¾ with system water (no more than 2 large females per bucket). For X. tropicalis, we recommend individually housing in small 3 L tanks. Ensure you can identify the frogs to check each animal and record their health and productivity in the colony database or logbook.
  4. When ready to collect eggs, gently but firmly pick up the female from the dorsal side, covering its face with the palm. Use the thumb and middle finger to hold the posterior sides of the abdomen. Use both index fingers (dominant and non-dominant hands) to gently push out and restrain the legs. Use the thumbs to apply gentle pressure to the abdomen.
  5. Massage each frog for no more than 30 seconds, or until the flow of eggs slows down (see Note 8). Allow the eggs to drop from a shallow height into the labelled petri dishes, making small mounds of eggs. See Note 9.
  6. Return the frog to her temporary housing to rest before subsequent ‘squeezes’. Once the frog has been ‘squeezed’ the maximum number of times (6 for X. laevis, 3 for X. troplicalis), put her in a recovery tank to be monitored (see section 3.8. Post-ovulation monitoring).
  7. Use a 3.5 mL plastic transfer pipette to remove any system water that may have dropped into the Petri dish during collection.


Figure 4: Collecting eggs via ‘squeezes’. The frog is held using a similar handling technique (Figure 2), but the index fingers on both hands are used to restrain the legs whilst the abdomen is massaged (A&B), releasing the Xenopus’ eggs into a clean Petri dish with the liquid removed (C). Frogs can be left to lay their eggs naturally in their tank (D), and the eggs can be collected with a Pasteur or transfer pipette.

Recognising healthy eggs is similar between X. laevis and X. tropicalis. Eggs that are laid in stringy clumps, or milky in colour are likely to be poor quality. Eggs should not have a ‘cracked top’ or large patches of white pigment. Healthy eggs have a clearly defined dark animal pole (top) and an off-white vegetal pole (bottom). Any eggs that are completely white are dead and not usable.
In vitro fertilisation of Xenopus eggs

Eggs are fertilised with either fresh (Step 1) or cryogenically frozen (Step 2) sperm (see Note 10), which is dispersed equally over the eggs using a Pasteur or widened “yellow” pipette tip (see Note 11). See MiMB Chapter 8 for protocols to preparing cryogenically frozen sperm. IVF is similar between both species, though it is important to use the correct concentration of MMR, 0.1x for X. laevis and 0.05x for X. tropicalis. For X. laevis, at any point, 0.1x MMR can be substituted for 0.1x MBS.

  1. Dissociate the testis in 0.5 to 1 mL of 1x MBS using two pairs of broad forceps to tear the tissue in an angled Petri dish OR macerate 1 testis with a pellet pestle in a microcentrifuge tube containing 0.5 mL 1x MBS (see Note 12). If you have more than one Petri dish of eggs, the sperm can be diluted (up to 2 mL) with 1x MBS, to ensure there is enough volume. Go to Step 3.
  2. OR if using frozen sperm, follow the Steps listed in MiMB Chapter 8. Then go to Step 3.
  3. Add the sperm suspension to the eggs collected in a Petri dish using a pipette. If you are using a micropipette with a yellow tip, cut off the end to reduce damage to the sperm.
  4. For X. laevis, gently shake the petri dish to spread the eggs or for X. tropicalis gently rearrange the eggs with the ends of two 200 µL pipette tips, to spread the eggs out until they are a monolayer to enable the sperm to access all eggs.
  5. Place into an incubator set to the appropriate temperature for the species. For X. laevis this should be set at 18.5°C, for X. tropicalis this should be set at 25°C.
6. After 5-10 minutes, flood eggs with room temperature 0.1x MMR (X. laevis) or 0.05x MMR (X. tropicalis). Return to the incubator for a further 20 minutes.
De-jellying fertilised Xenopus eggs for experiments that require early developmental stages

Fertilised X. laevis and X. tropicalis eggs that will be used in experiments requiring early developmental stages (e.g. NF1-8) must have the jelly coat surrounding the embryos removed. The method is similar between both species, though it is important to use the correct concentration of MMR, 0.1x for X. laevis and 0.05x for X. tropicalis. For X. laevis, at any point, 0.1x MMR can be substituted for 0.1x MBS.

  1. After 20 minutes in the incubator, the animal pole (brown/pigmented side) will have turned to face up. This is most consistent for X. laevis. For X. tropicalis, it is more likely to see the animal pole contract under a light microscope.
  2. Pour off the MMR and pour on 2% cysteine pH 7.8-8.
  3. Gently swirl the dish until the eggs unstick from the base of the petri dish.
  4. Using a 3 mL plastic transfer pipette, carefully transfer the eggs into a 50 mL conical tube.
  5. Top up the cysteine to 30 mL and gently invert the tube for 5 minutes.
  6. Stop inverting and allow the eggs to settle regularly. Once the gaps between the eggs caused by the jelly coat have disappeared, pour off the cysteine. See Note 13.
  7. To wash the eggs, top up the conical tube to 40 mL with MMR, once the eggs have settled, pour it away.
  8. Repeat Step 7 four more times to remove the cysteine.
  9. Top up the conical tube to 25 mL with MMR and carefully pour the fertilised, de-jellied eggs into a clean petri dish. De-jellied X. tropicalis eggs are very sticky and should be poured into an agarose-lined petri dish.

Successfully fertilised eggs have a consistent pigment colour (this can range from light to dark brown) and are “fuller”, more rounded and firmer than unfertilised eggs. X. laevis eggs will rotate so that the animal pole is facing up (4) and the animal pole of X. tropicalis eggs will contract. The sperm entry point may be visible, a small dark spot on the animal pole, encased by a dark ring (9), but this is also seen upon prick activation when something sharp has nicked the egg eliciting a similar reaction. Both prick activation and sperm entry induce an influx of calcium and harden the outside of the egg (10–14). If an egg has been prick activated without sperm entry, it is no longer able to be fertilised and will not divide (15). You may, however, see a dark line that looks like the egg is about to cleave, but it will progress no further than this. Unfertilised eggs that have not undergone prick activation lack firmness, are misshapen and may not roll so the animal pole faces up.

Figure 5: Differentiating fertilised, one-cell eggs from unfertilised eggs. After the initial fertilisation (A), healthy eggs have a distinct dark ‘speckled’ animal pole (top), an off-white vegetal pole (bottom) and a band separating the animal and vegetal pole. Dying eggs are consistently white and labelled ‘D’. Dark, pigmented eggs (B) visibly show the maturation spot (labelled ‘M’), a white spot that indicates that the egg has successfully undergone meiosis and is viable for fertilisation, and have clearly distinct animal and vegetal poles. The sperm entry point (labelled ‘SE’) is more visible on pale Xenopus eggs (C), characterised as a small dark spot within the area of the animal pole. Those marked ‘U’ are unfertilised and would be unsuitable for micro-injection of mRNA/CRISPR, for example.

After either IVF or natural mating, poorly developing or dead eggs should be removed since they interfere with the development of healthy embryos (16). Development of embryos can be monitored under a light microscope and the developmental stages determined using the Nieuwkoop and Faber Table of Normal Xenopus Development (17), or the updated Zahn images (18) This will also be helpful to recognise fertilised eggs that are cleaving abnormally or progressing through development poorly. For normal development, fertilised eggs will undergo several rapid, synchronous cleavages, progressing from one-cell to two-cell to four-cell, and so on. Cleavages appear first as a dark line, before fully bisecting the embryos. The first cleavage establishes the dorsal-ventral axis during early blastula by establishing the mid-sagittal plane. For some embryos, the first cleavage furrow may look incomplete or off-centre. These embryos will progress through development poorly and, as the Xenopus fate map is well characterised, off-centre will negatively affect downstream experiments that rely on fate restriction. During blastula stages, some embryos may lose their full and spherical shape, becoming flat and disk-shaped. These embryos will usually continue to progress through development poorly.
Natural Matings of Xenopus

Natural matings enable embryos to be generated without killing a male frog or massaging a female. This is recommended when there are not many males of a specific genetically altered line or if embryos are not required at early developmental stages for experiments. Though uncommon, X. tropicalis females are more prone to squeeze-related-death, regardless of how diligently the protocol is followed, and tend to recover more consistently after natural matings.

To prepare Xenopus frogs for natural matings, a priming and boosting dose of hCG hormone is injected into the dorsal lymph sac of both males and females. The person performing the injection must be trained and signed off as competent to comply with local regulations. It is important not to reuse needles as each use blunts the needle. Reusing a needle to inject a frog will cause more damage and pain than using a new needle. Piercing the vial of hCG counts as one use.

The following protocol to prepare male and female Xenopus for natural matings is similar to the female Xenopus ovulation protocol, using the same handling and injection techniques. It is important to note the dose changes of hCG.

Table 2: Hormone doses, needle gauges and sperm requirements for natural matings in X. laevis and X. tropicalis
ABCDEF
X. laevis (18.5 °C) X. tropicalis (25.5 °C)
Concentration of hormone Timing Concentration of hormone Timing
Priming Female 100 IU 1-3 days before the natural mating 10 IU The evening before the natural mating (around 4:30pm)
Male 50 IU 10 IU
Boosting Female 200 IU The evening of the natural mating (around 4:30pm) 100 IUThe morning of the natural mating (around 8:30am)
Male 100 IU100 IU
Time of Natural mating Frogs are left to mate overnight (6pm-8am) Frogs are left to mate throughout the day (10am-5pm)
Needle size for injections 27-gauge needle Drawing up the stock/solvent AND All frog injections 27-gauge needle Drawing up the stock/solvent
30-gauge needle All frog injections
Table 2: This table describes and compares the injection concentration of hCG, timings, and needle sizes between Xla and Xtr for natural matings. When following the protocols listed in 3.5. Natural Matings, please refer to this table to administer the correct dose, at the correct time, with the correct needles. This table can be used to adapt the timings of injections if you need to make a more suitable time, i.e. egg collection for Xtr to be 2 hours later, males and females can be administered the boost at 10:30 am.

Priming
  1. Always wear gloves when handling the frogs.
  2. Prepare and administer the priming dose. Add 1 mL of solvent to 1 vial of freeze-dried hCG (1500 IU/mL of stock solution) using a 27-gauge needle attached to a 1 mL syringe. Let the solution warm to room temperature.
  3. Dilute the stock and draw up the volume to administer the correct dose. See Table 2.2. and Table 2.3. for guidance.
  4. Detach the syringe (see Note 3) and swap the needle (piercing the vial blunts the needle and counts as one use) for a suitable needle for injection. See Table 2.2. Remove any air bubbles. See Note 3.
  5. Gently remove the frog from its tank, holding it firmly. The dorsal side of the frog should rest against the palm with its face towards the wrist. Restrain the frog by wrapping the thumb and middle finger around the posterior sides of the abdomen. Put the index finger between the legs. The little finger can be placed ventrally under the throat to further restrain the frog.
  6. Lie the frog on a damp paper towel and fold up the sides to cover the head and the legs. Always wrap headfirst to relax the frog. Tear a holein the paper towel to expose the site of injection.
  7. Insert the needle at a shallow angle, subcutaneously through the dorsal surface, proximal to the lateral lines. Inject the hCG solution slowly into the dorsal lymph sac. See Table 2.2.
  8. Ensure a new needle is used for each injection.
  9. If the injection site bleeds after injection, hold a piece of paper towel over the area and apply gentle pressure for 30 seconds until the bleeding has stopped.
  10. The frogs can be returned to the system, but must be easily identifiable (see MiMB Chapter 1, husbandry labels). Males and females should be housed separately from each other, however we tend to inject animals in small groups so that they do not have to be kept singly. See Note 14.
  11. The hCG stock solution can be stored in the fridge overnight. See Note 5. Boosting
  12. Prepare and administer the boosting dose. Remove the hCG stock solution(1500 UI/mL) from the fridge and bring to room temperature.
  13. Dilute the stock and draw up the volume to administer the correct dose. See Table 2.2. and 2.3. for guidance.
  14. Remove the syringe (see Note 3) and swap the needle (piercing the vial blunts the needle and counts as one use) for a suitable needle for injection. See Table 2.2. Remove any air bubbles.
  15. Following the handling and injection techniques in Steps 5-9.
  16. Pair male and female frogs in a labelled 16 litre bucket housed in a temperature-controlled room holding at a suitable temperature. For X. laevis, 18-20°C. For X. tropicalis, 23-26°C.
  17. Leave frogs to mate. Amplexus (where the male latches on to the female around the dorsal region anterior of her back legs using his arms) can often occur for several hours.
  18. When amplexus has finished (see Table 2.2. for timings), remove the frogs from the bucket and place them in tanks for post-ovulation monitoring (see section 3.9. Post ovulation care and monitoring of Xenopus after both IVF and Natural Mating).
Collecting embryos from Natural Matings

If possible, wait until the frogs have finished mating and have been removed from the bucket before collecting the embryos to reduce stress to the animals. It is, however, possible to collect embryos generated from natural matings at intervals carefully without disturbing amplexus, if specific early developmental stages are needed. If many, synchronously developing embryos at early developmental stages are required then consider using IVF as an alternative.

The EXRC employs two methods to collect embryos from natural matings. Please follow either Step 1 OR Step 2.

  1. Remove 80-90% of the system water from the bucket. The embryos will remain in clumps and stick to the bottom of the bucket. Use a 3.5 mL plastic transfer pipette to remove the embryos from the bucket into clearly labelled 50 mL conical tubes. Continue to Step 3.
  2. OR pour off all the water from the bucket and swirl 2% cysteine pH7.8-8 around the bottom of the bucket until the eggs detach, collect them in a beaker, pour off excess cysteine then pour them into clearly labelled 50 mL conical tubes. Continue to Step 3.
  3. Fill each conical tube with no more than 5 mL of embryos.
  4. Follow Steps 5-9 from section 5.6. Preparing Xenopus eggs for experiments

Post-ovulating care and monitoring of Xenopus after both IVF and Natural Matings

Post ovulation care and monitoring differ between X. laevis and X. tropicalis, as there is a higher risk of mortality for X. tropicalis in the two weeks following ovulation due to becoming more frequently egg-bound.

After ovulation, X. laevis should be returned to their home tank. X. tropicalis, however, should be placed in a higher salt tank or tank system to help encourage release of any remaining loose eggs, and should not be fed during this time.

Once female X. tropicalis squeezes have finished, and are they are no longer being used to produce eggs, they should be housed in a high salt tank (0.3g/L of marine salt) for 48 hours. They do not (and should not) need to be singly housed if they are easily identifiable. The EXRC has a recirculating rack system with a high salt conductivity (1300 µS). It is possible to use individual static tanks, ensuring the water has the same conductivity, as an alternative. The water temperature in the tank should be lowered to 23-24°C (around room temperature), as the EXRC have found this to be more conducive to encouraging the females to lay their remaining eggs. If you are using individual static tanks, ensure the water temperature is maintained between 23 and 24°C. This can be achieved by placing tanks on a heat mat or putting them in a temperature-controlled room (24°C).

X. tropicalis post-ovulation salt treatment
  1. Put females into a clean salt tank with enrichment. If you are using a recirculating system, turn the water flow to this tank off. This allows the water temperature to drop to 24°C.
  2. Move the females to a new, clean salt tank every day for 48 hours as they will continue laying their eggs, making the water murky (see Note 15). If you are using a recirculating system, turn off the water flow. Ensure the water temperature is maintained between 23 and 24°C.
  3. When they have completed their 48 hours in higher salt, return them to their home tank.
  4. Once a week, donate a biofilter from a regular (not high salt) X. tropicalis system to the high salt system; this will maintain healthy bacterial levels in the dump.

After ovulation, Xenopus, regardless of species, should be monitored at least twice daily, ideally four times daily, for two weeks for signs of poor health. If possible, house the frogs in a location that is eye level and easy to see. Key signs of an egg-bound frog include bloating, lethargy, swimming or floating on one side, fluid accumulation under the skin, eggs stuck in their cloaca or being too buoyant and unable to swim to the bottom of the tank. These events are rare, but if any of these are observed, humanely kill the frog. Some frogs will show signs of sickness or bloating even if the protocol is followed diligently.
Notes
  1. There are references to storing X. laevis and X. tropicalis testes in 14oC 60% (v/v) Leibovitz’s 15 medium, 10% calf serum, for up to two weeks. This has not been found to be as consistent in maintaining the integrity of the sperm at the EXRC.
  2. The EXRC finds it is not necessary to give 2 (priming and boosting) injections. This reduces the number a of time a frog needs to be handled and injected. It is recommended to follow this protocol to promote good ethical standards
  3. If you are experienced with injections and handling needles, you can leave the needle in the vial for subsequent withdrawals of the stock. This minimises the number of needles you will have to use.
  4. The easiest way to remove bubbles is to draw up more solution than you will administer. For example, if you plan to administer 0.1 mL, draw up 0.2 mL. Once removed from the vial, hold the syringe with the needle facing upwards and pull the plunger back, creating a gap of air. Tap the sides of the syringe so the bubbles rise to the top. Gently press the plunger to expel the bubbles (and additional liquid) via the needle.
  5. The EXRC do not store hCG stock as liquid at 4°C for more than 24 hours, or at -20°C for more than 4 weeks. However, other labs have found that hCG suspended in the supplied buffer is viable up to 1 month at 4 °C.
  6. Frogs should never be injected with a cold solution for priming and/or boosting. It is imperative that you administer the injection solution at room temperature. You can use your hands to warm the vial.
  7. Frogs should start naturally producing eggs 3-4 hours after the boosting dose. Their cloaca will be red and more prominent than normal. If not, this is a good indication that the frog will not lay any eggs.
  8. Some females may not lay, do not over squeeze them, as it could be the result of jelly clogging the oviduct or cloaca or that she is not ready to release her eggs. Gently massage the abdomen to encourage the release of the jelly plug. If the jelly plug does not pass, return the frog to her tank to allow it to pass naturally, monitoring continuously. It is important to monitor the health of every frog post-ovulation. Any mature eggs left inside the body can die, resulting in frog death in a process related to septic shock. For X. tropicalis, housing the female in a high salt concentration tank will encourage the passing of eggs.
  9. Try not to pile all the eggs in one part of the petri dish. Smaller clumps of eggs are easier to spread out and allow a more even dispersion of sperm. During IVF, spread the eggs into a monolayer after the sperm suspension has been applied.
  10. EXRC staff have found that fresh X. tropicalis sperm does not fertilise well if not used immediately. For this reason, the EXRC frequently use frozen sperm for X. tropicalis fertilisations (see Chapter 8 for sperm cryopreservation protocols). This helps reduce the number male X. tropicalis killed for in vitro fertilisation.
  11. Use wide-end micropipette tips to avoid damaging the sperm. These can be bought pre-cut or simply cut 1.5-2 cm off the end of a standard 200 µL micropipette tip.
  12. Check that your pellet pestle fits to the bottom of the microcentrifuge tube to ensure thorough maceration of the testis.
  13. Males and females can be housed with other frogs of the same sex who have not been primed. It is not recommended to singly house frogs unnecessarily.
  14. There should be no visible gaps between the embryos, they should fall quickly to the bottom and not visibly stick together.
  15. You may have to move the females more than once per day, they will lay a significant number of eggs, which makes the water murky.

Table 3 Dilutions of hCG for injection concentrations.
ABCDEF
Stock solution (1500 IU/mL) Species and type of mating In Vitro Fertilisation (IVF) or Natural Mating (NM) Desired amount of hCG to inject Stock:Solvent Final volume and conc/mL Volume to inject per frog
Xla female full ovulation dose (IVF) 600 IU No dilution needed 0.4 mL 1500 IU/mL 0.4 mL
Xla female boosting dose (NM) 200 IU 0.4 mL stock 0.2 mL solvent 2:1 0.6 mL 1000 IU/mL 0.2 mL
Xla female priming dose (NM) 100 IU 0.2 mL stock 0.4 mL solvent 1:2 0.6 mL 500 IU/mL 0.2 mL
Xtr female boosting dose (IVF and NM) Xtr male boosting dose (NM) 0.2 mL stock 0.1 mL solvent 2:1 0.3 mL 1000 IU/mL 0.1 mL
Xla male priming dose (NM) 50 IU 0.2 mL stock 0.4 mL solvent 1:2 0.6 mL 500 IU/mL 0.1 mL
Xtr female priming dose (IVF and NM) Xtr male priming dose (NM) 10 IU 0.02 mL stock 0.28 mL solvent 1:14 0.3 mL 100 IU/mL 0.1 mL
Table 3: This table describes how to make the correct dilutions from a hCG stock concentration of 1500 IU/mL. The recommended volumes are used by the EXRC currently, but these may be altered using the given ratios for ease or number of frogs injected, depending on experience.

Figure Legends
Figure 1: Dissection of male Xenopus testes for isolation of fresh sperm. A dead Xenopus male is rested in the supine position, ready for dissection (A). Once the body cavity has been opened (B) and the fat bodies (labelled ‘F’) pulled to the side, the testes (labelled ‘T’) are visible, located just above the kidney. Once dissected out, each testis is cleaned to remove remnants of non-testicular tissue and blood (C).
Figure 2: An appropriate handling technique of Xla (A&B) and Xtr (C&D) from dorsal (A&C) and ventral (B&D) view. The frog is being held firmly, but is relaxed in the hand and not struggling. The thumb and the middle finger are wrapped around the abdomen, and the index finger is placed between the legs. For Xla, little finger is placed on the throat to further restrain the frog.
Figure 3: The location for injection of hCG into the dorsal lymph sac of a Xenopus is easily found on the dorsal side (A). Just above the junction of the back leg (labelled 'L') and the body is a semicircle of slits (B) called lateral lines (labelled 'LL'), or ‘stitching’. Below this, a membrane layer joins the outer skin to the body wall, when the frog’s skin is gently pinched for injection (C), it is easy to differentiate the two layers. The needle is inserted at a shallow angle, almost parallel to the frog proximal to the lateral line. This ensures the needle stays between the two layers. The figure demonstrates the rotation of the frog (craniocaudal and dorsoventral rotation).
Figure 4: Collecting eggs via ‘squeezes’. The frog is held using a similar handling technique (Figure 2), but the index fingers on both hands are used to restrain the legs whilst the abdomen is massaged (A&B), releasing the Xenopus’ eggs into a clean Petri dish with the liquid removed (C). Frogs can be left to lay their eggs naturally in their tank (D), and the eggs can be collected with a Pasteur or transfer pipette.
Figure 5: Differentiating fertilised, one-cell eggs from unfertilised eggs. After the initial fertilisation (A), healthy eggs have a distinct dark ‘speckled’ animal pole (top), an off-white vegetal pole (bottom) and a band separating the animal and vegetal pole. Dying eggs are consistently white and labelled ‘D’. Dark, pigmented eggs (B) visibly show the maturation spot (labelled ‘M’), a white spot that indicates that the egg has successfully undergone meiosis and is viable for fertilisation, and have clearly distinct animal and vegetal poles. The sperm entry point (labelled ‘SE’) is more visible on pale Xenopus eggs (C), characterised as a small dark spot within the area of the animal pole. Those marked ‘U’ are unfertilised and would be unsuitable for micro-injection of mRNA/CRISPR, for example.
Table Legends
Table 2.1: A table to describe and compare the injection concentration of hCG, timings, needle sizes and fresh sperm collection between Xla and Xtr for IVF. When following the protocols listed in 3.3. Ovulating Xenopus females, please refer to this table to administer the correct dose, at the correct time, with the correct needles. This table can be used to adapt the timings of injections if you need to make a more suitable time, i.e. egg collection for Xtr to be 2 hours later, females can be e administered the boost at 10:30 am.

Table 2.2: This table describes and compares the injection concentration of hCG, timings, and needle sizes between Xla and Xtr for natural matings. When following the protocols listed in 3.5. Natural Matings, please refer to this table to administer the correct dose, at the correct time, with the correct needles. This table can be used to adapt the timings of injections if you need to make a more suitable time, i.e. egg collection for Xtr to be 2 hours later, males and females can be administered the boost at 10:30 am.

Table 2.3: This table describes how to make the correct dilutions from a hCG stock concentration of 1500 IU/mL. The recommended volumes are used by the EXRC currently, but these may be altered using the given ratios for ease or number of frogs injected, depending on experience.
Protocol references
  1. Shapiro HA, Zwarenstein H. A Rapid Test for Pregnancy on Xenopus lævis. Nature. 1934 May 19;133(3368):762–762.
  2. Bellerby CW. A Rapid Test for the Diagnosis of Pregnancy. Nature. 1934 Mar;133(3361):494–5.
  3. Wlizla M, Falco R, Peshkin L, Parlow AF, Horb ME. Luteinizing Hormone is an effective replacement for hCG to induce ovulation in Xenopus. Dev Biol. 2017 Jun;426(2):442–8.
  4. Guille M. Molecular Methods in Developmental Biology. Vol. 127. New Jersey: Humana Press; 1999.
  5. Sargent MG, Mohun TJ. Cryopreservation of sperm of Xenopus laevis and Xenopus tropicalis. genesis. 2005 Jan;41(1):41–6.
  6. Mansour N, Lahnsteiner F, Patzner RA. Optimization of the cryopreservation of African clawed frog (Xenopus laevis) sperm. Theriogenology. 2009 Dec;72(9):1221–8.
  7. Pearl E, Morrow S, Noble A, Lerebours A, Horb M, Guille M. An optimized method for cryogenic storage of Xenopus sperm to maximise the effectiveness of research using genetically altered frogs. Theriogenology. 2017 Apr;92:149–55.
  8. Noble A, Abu-Daya A, Guille M. Cryopreservation of Xenopus Sperm and In Vitro Fertilization Using Frozen Sperm Samples. Cold Spring Harb Protoc. 2022 Feb;2022(2):pdb.prot107029.
  9. Zahn N, James-Zorn C, Ponferrada VG, Adams DS, Grzymkowski J, Buchholz DR, et al. Normal Table of Xenopus development: a new graphical resource. Development. 2022 Jul 15;149(14).
  10. Wolf DP. The cortical response in Xenopus laevis ova. Dev Biol. 1974 Sep;40(1):102–15.
  11. Stewart-Savage J, Grey RD. Fertilization of investment-free Xenopus eggs. Exp Cell Res. 1984 Oct;154(2):639–42.
  12. Takeichi T, Kubota HY. Structural basis of the activation wave in the egg of Xenopus laevis. Development. 1984 Jun 1;81(1):1–16.
  13. Cheer A, Vincent JP, Nuccitelli R, Oster G. Cortical activity in vertebrate eggs I: The activation waves. J Theor Biol. 1987 Feb;124(4):377–404.
  14. Kline D, Nuccitelli R. The wave of activation current in the Xenopus egg. Dev Biol. 1985 Oct;111(2):471–87.
  15. Grey RD, Working PK, Hedrick JL. Evidence that the fertilization envelope blocks sperm entry in eggs of Xenopus laevis: Interaction of sperm with isolated envelopes. Dev Biol. 1976 Nov;54(1):52–60.
  16. Coleman A. Transcription and Translation, a Practical Approach. Hames BD, Higgins SJ, editors. Oxford: IRL Press; 1984.
  17. Nieuwkoop PD, Faber J. Normal Table of Xenopus laevis (Daudin); a Systematical and Chronological Survey of the Development from the Fertilized Egg Till the End of Metamorphosis. 1956;
  18. Zahn N, Levin M, Adams DS. The Zahn drawings: new illustrations of Xenopus embryo and tadpole stages for studies of craniofacial development. Development. 2017 Aug 1;144(15):2708–13.
Acknowledgements
The authors would like to thank Caroline W Beck for her comments on the protocols.