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5 minutes

We thank Susan Carroll, Eva Flores, and the Animal Resources Center staff at Montana State University for expert frog care. This work used the Animal Resources Center (RRID:SCR_026351), which is supported by the Office of Research and Economic Development at Montana State University. This work was funded by NSF grant HRD-2054276.

All methods were approved and performed in accordance with the rules and regulations of the Montana State University IACUC (Institutional Animal Care and Use Committee).
The following steps detail the necessary preparation, surgical procedure, and post-operative care for performing unilateral gonadectomy in adult male X. laevis.
1. Instrument and surgical site asepsis
<ol>
	<li>Use aseptic techniques for all procedures.</li>
	<li>Sterilize instruments by autoclaving or using a glass-bead sterilizer. Ensure instrument tips are completely cooled before contact with tissues if using a glass-bead sterilizer.</li>
	<li>Wear sterile, powderless gloves, and change gloves prior to making the surgical incision.</li>
</ol>
2. Presurgical preparation and anesthesia
<ol>
	<li>Prepare 200-250 mL of a 0.15% (1.5 g/L) solution of Tricaine methane sulfonate (MS-222), buffered to pH 7.0 with Sodium Bicarbonate. 
	CAUTION: MS-222 is toxic; handle with appropriate Personal Protective Equipment and avoid skin contact or inhalation.</li>
	<li>Partially submerge the frog's body in the buffered MS-222 solution, ensuring its head remains above water (Figure 2A) and keep the frog moist. Observe the frog continuously during induction of anesthesia to prevent hypoxemia and oversedation. 
	NOTE: We found that a simple 24 fl oz “deep dish” storage container works well (Figure 2A).</li>
	<li>Confirm a surgical level of anesthesia by gently pinching the fleshy part of both rear feet with forceps or a hemostat to ensure non-responsiveness.</li>
	<li>Place the frog on its back on sterile paper towels wetted with the MS-222 solution, on a large piece of sterile aluminum foil. Apply strips of paper towel moistened in MS-222 solution to the frog’s legs and torso to prevent the skin from drying and to maintain an adequate anesthetic plane throughout the procedure.</li>
</ol>
3. Administration of analgesia
<ol>
	<li>Inject Meloxicam at a dose of 0.1 mg/kg intramuscularly into a calf muscle (gastrocnemius). Proceed right away. 
	NOTE: See Kawano et al.9 for the stability of diluted meloxicam. 
	CAUTION: Meloxicam is toxic; handle with appropriate Personal Protective Equipment and avoid skin and eye contact.</li>
	<li>Apply strips of sterile paper towel wetted in the MS222 solution to the periphery of the surgical site to keep the frog moist and maintain anesthesia.</li>
	<li>If desired, place a surgical drape over the frog.</li>
	<li>Rinse the incision site with 0.9% sterile saline for at least 5 s using a sterile bulb pipette.</li>
	<li>Using a sterile cotton swab, dab Bupivacaine diluted to 0.25% with sterile water onto the skin in the incision area and allow 5 min to absorb prior to incision. 
	NOTE: Do not exceed 5 mg/kg Bupivacaine. 
	CAUTION: Bupivacaine is toxic; handle with appropriate Personal Protective Equipment and avoid skin contact.</li>
</ol>
4. Surgical procedure
<ol>
	<li>Use a number 15 scalpel and sharp scissors to cut through the skin and muscle layers respectively. Make a 1 cm incision (Figure 2B).</li>
	<li>Use the spay hook (Figure 3) to gently pull out the yellow/orange abdominal fat body along with the small intestine. Follow the abdominal fat body medially to locate and remove the testis (Figure 2C). 
	NOTE: Do not cut across the large abdominal blood vessels when cutting through the muscle layer. Similarly, avoid damaging any blood vessels when pulling out the abdominal fat body and intestines, and when removing the testis, as this poses a significant risk of hemorrhage.</li>
	<li>Store testis in 1x Modified Barth’s Saline (MBS or similar) at 4°C.</li>
	<li>Guide the intestines and the abdominal fat body back into the body cavity using a blunt instrument (spay hook).</li>
	<li>Close the muscle layer and skin layer separately using an absorbable Poliglecaprone 25 monofilament suture in two or three simple interrupted sutures (Figure 2D,E). 
	NOTE: Tying square knots is recommended as described in Schneider et al.10.</li>
</ol>
5. Postsurgical monitoring and housing
<ol>
	<li>Rinse the frog in clean, chlorine-free water to remove residual MS-222 (e.g., frog system water).</li>
	<li>Place the frog into a container with fresh chlorine-free water. Support the head above water and ensure skin remains moist until the frog is moving normally (similar to Figure 2A). Observe the frog at all times during recovery, which may take 20–30 min.</li>
	<li>Once the frog is moving normally, transfer the frog into a stand-alone recovery tank with enough water to just fully submerge the frog for 24 h (Figure 2F).</li>
	<li>After 24 hours, return the frog to a standard housing tank (Figure 2G), separate from other frogs, and monitor the frog daily for normal eating and swimming, signs of skin lesions or loss of skin, swollen or red legs, and incisional healing. After surgery, frogs return to a normal feeding schedule right away. 
	NOTE: The frog is fed 3 days after surgery and after that 2x per week. The standard housing tank is part of a recirculating system (e.g., a Techniplast Xenoplus system, Figure 2G). </li>
</ol>
<img alt="Frog dissection: surgical procedure, testis identification, recovery tank, laboratory setup." class="xfigimg" src="/files/ftp_upload/71258/71258fig2.jpg" data-altupdatedat="1790049645000" data-alt="Figure 2"> 
Figure 2: Male X. laevis survival surgery for single testis collection. (A) Frog body partially submerged in an anesthetic at the beginning or in water at the end of the procedure, ensuring that the head remains above the water level. (B) Small incision, approximately 1 cm in length. (C) Intestines and abdominal fat body are gently pulled out until the testis can be seen. The kidney-shaped testis is dissected out. (D) For suturing, square knots are recommended. Muscle and skin layers are sutured separately. (E) Three sutures are appropriate for the incision length. (F) Frog is returned into a separate housing tank with low water for 24 h to enable easy breathing. (G) The frog is returned to a standard housing tank on the recirculating system. <a href="https://www.jove.com/files/ftp_upload/71258/71258fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Surgical instruments setup; includes forceps, scalpel, surgical scissors; medical tools overview." class="xfigimg" src="/files/ftp_upload/71258/71258fig3.jpg" data-altupdatedat="1790049645000" data-alt="Figure 3"> 
Figure 3: Surgical tools. 1- Adson dressing forceps, 2- Rat tooth forceps, 3- Spay hook, 4- Scissors, 5- Surgical needle driver. <a href="https://www.jove.com/files/ftp_upload/71258/71258fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

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M., et al. <a target="_blank" href="https://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+Xenopus+ORFeome:+a+resource+that+enables+functional+genomics.">The Xenopus ORFeome: a resource that enables functional genomics.</a> Dev Biol. 408 (2), 345-357 (2015).</li><li>Harland, R. M., Grainger, R. M. <a target="_blank" href="https://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Xenopus+research:+metamorphosed+by+genetics+and+genomics.">Xenopus research: metamorphosed by genetics and genomics.</a> Trends Genet. 27 (12), 507-515 (2011).</li><li>Blum, M., Ott, T. <a target="_blank" href="https://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Xenopus:+an+undervalued+model+organism+to+study+and+model+human+genetic+disease.">Xenopus: an undervalued model organism to study and model human genetic disease.</a> Cells Tissues Organs. 205 (5-6), 303-313 (2018).</li><li>Wessely, O., Kim, J. I., Geissert, D., Tran, U., De Robertis, E. M. <a target="_blank" href="https://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Analysis+of+Spemann+organizer+formation+in+Xenopus+embryos+by+cDNA+macroarrays.">Analysis of Spemann organizer formation in Xenopus embryos by cDNA macroarrays.</a> Dev Biol. 269 (2), 552-566 (2004).</li><li>Anderson, C., Stern, C. D. <a target="_blank" href="https://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Organizers+in+development.">Organizers in development.</a> Current topics in developmental biology. , 435-454 (2016).</li><li>Wlizla, M., McNamara, S., Horb, M. E. <a target="_blank" href="https://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Generation+and+care+of+Xenopus+laevis+and+Xenopus+tropicalis+embryos.">Generation and care of Xenopus laevis and Xenopus tropicalis embryos.</a> Methods Mol Biol. 1865, 19-32 (2018).</li><li>Kawano, H. K., Simonek, G. D., Moffitt, A. D., Tahara, J. M., Brignolo, A. L. <a target="_blank" href="https://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Sterility+and+stability+of+diluted+meloxicam+in+compounded+multi-dose+vial+after+365+days.">Sterility and stability of diluted meloxicam in compounded multi-dose vial after 365 days.</a> J Am Assoc Lab Anim Sci. 58 (5), 594-596 (2019).</li><li>Schneider, P. N., Hulstrand, A. M., Houston, D. W. <a target="_blank" href="https://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Fertilization+of+Xenopus+oocytes+using+the+host+transfer+method.">Fertilization of Xenopus oocytes using the host transfer method.</a> J Vis Exp. (45), e1864 (2010).</li><li>Balls, M. <a target="_blank" href="https://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+principles+of+humane+experimental+technique:+timeless+insights+and+unheeded+warnings.">The principles of humane experimental technique: timeless insights and unheeded warnings.</a> ALTEX. 27, 144-148 (2010).</li><li>Philips, B. H., et al. <a target="_blank" href="https://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Evaluation+of+presurgical+skin+preparation+agents+in+African+clawed+frogs+(Xenopus+laevis).">Evaluation of presurgical skin preparation agents in African clawed frogs (Xenopus laevis).</a> J Am Assoc Lab Anim Sci. 54 (6), 788-798 (2015).</li><li>Ramlochansingh, C., et al. <a target="_blank" href="https://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Efficacy+of+tricaine+methanesulfonate+(MS-222)+as+an+anesthetic+agent+for+blocking+sensory-motor+responses+in+Xenopus+laevis+tadpoles.">Efficacy of tricaine methanesulfonate (MS-222) as an anesthetic agent for blocking sensory-motor responses in Xenopus laevis tadpoles.</a> PLoS One. 9 (7), e101606 (2014).</li><li>Baumgartner, C., et al. <a target="_blank" href="https://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+comparison+of+the+analgesic+effects+of+fentanyl+and+butorphanol+in+African+clawed+frogs+(Xenopus+laevis)+under+tricaine+methanesulfonate+anesthesia.">A comparison of the analgesic effects of fentanyl and butorphanol in African clawed frogs (Xenopus laevis) under tricaine methanesulfonate anesthesia.</a> SOJ Anesthesiol Pain Manag. 5 (2), 1-10 (2018).</li><li>. Surgical oocyte collection from Xenopus laevis Available from: <a target="_blank" href="https://animalcare.ubc.ca/sites/default/files/2025-12/acc_sop_tech24_surgical_oocyte_collection_from_xenopus_laevis_20251117.pdf">https://animalcare.ubc.ca/sites/default/files/2025-12/acc_sop_tech24_surgical_oocyte_collection_from_xenopus_laevis_20251117.pdf</a> (2013)</li><li>. Policy on major invasive surgery (oocyte harvest) in frogs Available from: <a target="_blank" href="https://www.urmc.rochester.edu/medialibraries/urmcmedia/ucar/policies/documents/frogpolicy-majorinvasivesurger.pdf">https://www.urmc.rochester.edu/medialibraries/urmcmedia/ucar/policies/documents/frogpolicy-majorinvasivesurger.pdf</a> (2017)</li><li>. Guidelines for egg and oocyte harvesting in Xenopus laevis Available from: <a target="_blank" href="https://oacu.oir.nih.gov/system/files/media/file/2025-04/b11_xenopus_oocyte_collection.pdf">https://oacu.oir.nih.gov/system/files/media/file/2025-04/b11_xenopus_oocyte_collection.pdf</a> (2025)</li><li>Leung, G., Diaz, L. L., Monette, S., Jiang, C. S., Tolwani, R., et al. <a target="_blank" href="https://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Effectiveness,+safety,+and+pharmacokinetics+of+meloxicam+formulations+in+African-clawed+frogs,+Xenopus+laevis.">Effectiveness, safety, and pharmacokinetics of meloxicam formulations in African-clawed frogs, Xenopus laevis.</a> Comp Med. 74 (5), 336-343 (2024).</li><li>Warwick, C., Bates, G., Arena, P. C., Steedman, C. <a target="_blank" href="https://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Reevaluating+the+use+of+hypothermia+for+anesthetizing+and+euthanizing+amphibians+and+reptiles.">Reevaluating the use of hypothermia for anesthetizing and euthanizing amphibians and reptiles.</a> J Am Vet Med Assoc. 253 (12), 1536-1539 (2018).</li><li>Lalonde-Robert, V., Beaudry, F., Vachon, P. <a target="_blank" href="https://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Pharmacologic+parameters+of+MS222+and+physiologic+changes+in+frogs+(Xenopus+laevis)+after+immersion+at+anesthetic+doses.">Pharmacologic parameters of MS222 and physiologic changes in frogs (Xenopus laevis) after immersion at anesthetic doses.</a> J Am Assoc Lab Anim Sci. 51 (4), 464-468 (2012).</li><li>. Guidelines on amphibian anesthesia analgesia and surgery Available from: <a target="_blank" href="https://az.research.umich.edu/animalcare/guidelines/guidelines-amphibian-anesthesia-analgesia-and-surgery/">https://az.research.umich.edu/animalcare/guidelines/guidelines-amphibian-anesthesia-analgesia-and-surgery/</a> (2023)</li><li>Reed, B. T. <a target="_blank" href="https://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Guidance on the housing and care of the African clawed frog Xenopus laevis. , (2005).</li><li>Mansour, N., Lahnsteiner, F., Patzner, R. A. <a target="_blank" href="https://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Optimization+of+the+cryopreservation+of+African+clawed+frog+(Xenopus+laevis)+sperm.">Optimization of the cryopreservation of African clawed frog (Xenopus laevis) sperm.</a> Theriogenology. 72 (9), 1221-1228 (2009).</li><li>Sargent, M. G., Mohun, T. J. <a target="_blank" href="https://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cryopreservation+of+sperm+of+Xenopus+laevis+and+Xenopus+tropicalis.">Cryopreservation of sperm of Xenopus laevis and Xenopus tropicalis.</a> Genesis. 41 (1), 41-46 (2005).</li><li>Pearl, E., Morrow, S., Noble, A., Lerebours, A., Horb, M., et al. <a target="_blank" href="https://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=An+optimized+method+for+cryogenic+storage+of+Xenopus+sperm+to+maximise+the+effectiveness+of+research+using+genetically+altered+frogs.">An optimized method for cryogenic storage of Xenopus sperm to maximise the effectiveness of research using genetically altered frogs.</a> Theriogenology. 92, 149-155 (2017).</li></ol>

The authors have no conflicts of interest to declare.

The unilateral gonadectomy technique addresses a need in the management of laboratory X. laevis by maximizing the utility of male animals while adhering to high welfare standards. The core benefit of unilateral gonadectomy is survival of the male, thus preserving the remaining gonad for subsequent fertilization needs, while fulfilling the &#8220;3Rs&#8221; (replacement, reduction, refinement) mission for animal research11.
An important factor in successful frog surgery is the avoidance of certain antiseptic agents. Frogs&#39; skin has a protective mucus layer containing antimicrobial agents. A study evaluating presurgical skin preparation found that 0.75% chlorhexidine had superior antimicrobial activity but was associated with significant drawbacks, including a higher incidence of illness, epidermal degeneration, and necrosis. In contrast, simple rinses with 0.9% sterile saline or with 0.5% povidone&#8211;iodine were well tolerated12. Therefore, this protocol uses a saline rinse to protect the integrity of the frogs&#8217; highly sensitive skin.
MS-222 has been shown to be a safe and effective anesthetic that blocks the activity of both sensory and motor nerves in Xenopus13. However, MS-222 alone was shown insufficient for analgesia14. Given that amphibians perceive pain, it is assumed that X. laevis benefits from analgesics. Thus, the combined use of anesthetics and analgesics during surgical procedures is recommended14. The analgesic agents Meloxicam and Bupivacaine have been standard in X. laevis survival surgery to obtain oocytes15,16,17. Meloxicam is classified as a nonsteroidal anti-inflammatory drug, which was reported to be ineffective in providing analgesia in frogs18. However, the NIH guidelines for survival surgery include the use of meloxicam as an analgesic17. Hypothermia has been shown to mimic analgesia by reducing nociceptor firing rates in amphibians, with concerns that it results in discomfort, shivering, and prolonged recovery rather than reliable pain control19. Cooling or hypothermia is therefore not recommended for anesthesia or analgesia17,19.
Despite the advantages of the presented method, it requires familiarity with X. laevis anatomy and appropriate training and veterinary oversight. A concentration of 1&#8211;2 g/L MS-222 typically results in an anesthesia duration of 30&#8211;60 min20. Should the surgical frog begin to recover prior to completion of the procedure, either place the frog back into the MS-222 solution if there is no open incision, or reapply strips of paper towel moistened in MS-222 directly to the skin, taking care to avoid the incision site21. Cutting across the large abdominal blood vessels within the muscles poses a significant risk due to their central role in systemic circulation10. This can cause hemorrhage or vascular fragility. These vessels are visible and can be avoided. Should the vessel be severed, the frog needs to be euthanized. Further, great care must be taken to ensure the intestines are not damaged as they are pulled out and guided back into the body cavity. If the intestines are accidentally severed, the frog needs to be euthanized. As the frog recovers from anesthesia, it needs to be continuously monitored to prevent drowning and keep its head moist. Finally, the frog needs to be observed for post-surgical complications, including infection, suture tearing, skin sloughing, or redness. These adverse effects can be treated by placing the frog in a salt bath for up to 3&#8211;5 days22. If the condition worsens, euthanasia may need to be performed.
Cryopreservation methods exist for archiving sperm from X. tropicalis, for example, to preserve genetically altered lines, and these protocols have been adapted to X. laevis23,24. While cryopreservation of X. laevis sperm was successful, fertilization success after thawing was significantly lower than for X. tropicalis and was a fraction of that with fresh sperm. Fertilization success was also quite inconsistent, attributed to individual differences among frogs25.
The protocol described here provides an additional method for reducing the number of male X. laevis used for research. Standard protocols typically employ in vitro fertilization using macerated testis to generate synchronized embryos for studies of morphogenesis, early patterning, and gene function. The ability to collect sperm twice from the same male saves animals and reduces cost. It is expected that this procedure can easily be adapted to X. tropicalis. For laboratories working with scarce lines, such as transgenic strains, obtaining sperm from a single male more than once extends the productive lifespan of each frog and conserves resources without resorting exclusively to cryogenic preservation. In summary, this protocol contributes to more sustainable colony management.

Xenopus laevis, the African clawed frog, is an excellent model organism for studying embryonic development. Its usefulness stems from the accessibility of its stages, ranging from zygotic to adult1, allowing easy, reproducible manipulations such as microinjections and dissections. The conservation of genes and of cellular and molecular mechanisms between Xenopus and humans is remarkably high, with approximately 80% of Xenopus genes possessing a corresponding ortholog in humans2,3. As a result, fundamental concepts in cell and developmental biology were derived from X. laevis embryos, including the identification of different vertebrate developmental genes and core signaling pathways, such as those in the primary organizer4,5,6,7. Due to its many advantages as a model organism, X. laevis is widely employed in laboratories worldwide. Thus, improving methods for its use as a laboratory model is desirable.
Current standard fertilization protocols are performed either through in vitro fertilization, which yields a large number of synchronously developing embryos, or through natural mating, which offers the advantage of preserving the male for repeated use, but the resulting embryos are fertilized asynchronously. Thus, in vitro fertilization remains the most widely used approach for X. laevis. In vitro fertilization traditionally requires the euthanasia of the male frog to isolate its testes. Sperm collection from these testes yields high fertilization rates, but viability declines sharply within a few days8.
To reduce the number of required males, this protocol presents a specialized surgical approach: unilateral gonadectomy in X. laevis adult males (Figure 1). This procedure involves the removal of a single testis, followed by suturing the incision to allow the male to fully recover and survive. This method allows for the immediate use of viable sperm from the removed testis, while preserving the remaining gonad for subsequent fertilization needs.
<img alt="Frog gonadectomy diagram; pre and post-surgery comparison with surgical scissors for research." class="xfigimg" src="/files/ftp_upload/71258/71258fig1.jpg" data-altupdatedat="1790049645000" data-alt="Figure 1"> 
Figure 1: Schematic diagram for surgical removal of one testis. <a href="https://www.jove.com/files/ftp_upload/71258/71258fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

<table><tbody><tr><td>Adson Tissue Forceps, Delicate, 4.75&quot;</td><td>Vetone</td><td>615008</td><td></td></tr><tr><td>Bupivacaine Hydrochloride Injection 2.5 mg/mL, 50 mL</td><td>Meitheal Pharmaceuticals</td><td>121952</td><td></td></tr><tr><td>On-Cryl Poliglecaprone (4-0) SH 30 in</td><td>Vetone</td><td>603108</td><td>Pharmaceutical grade</td></tr><tr><td>Operating Scissors, Curved, Sharp-Sharp, 4.5&quot;</td><td>Securos Surgical</td><td>45217</td><td></td></tr><tr><td>Operating Scissors, Straight, Sharp-Sharp, 4.5&quot;</td><td>Securos Surgical</td><td>48468</td><td></td></tr><tr><td>OstiLox (Meloxicam Injection) 5 mg/mL</td><td>Vetone</td><td>501080</td><td></td></tr><tr><td>Spay hook</td><td>Vetone</td><td>39016</td><td></td></tr><tr><td>Surgical Drape</td><td>MWI Animal Health</td><td>600302</td><td></td></tr><tr><td>Surgical needle driver</td><td>Cynatech via Amazon</td><td>CNHB-STC</td><td></td></tr><tr><td>Techniplast Xenoplus system</td><td></td><td></td><td>commercial Xenopus housing/recirculating water system</td></tr><tr><td>Tricaine methane sulfonate&amp;nbsp;</td><td>Sigma</td><td>A5040</td><td></td></tr><tr><td>Vantage Brown-Adson Tissue Forceps, 7 x 7 Teeth, 4.75&quot;</td><td>Integra Miltex</td><td>22240</td><td></td></tr></tbody></table>

unilateral gonadectomy in adult male xenopus laevis

Unilateral Gonadectomy in Adult Male Xenopus laevis

In vitro fertilization is a reliable laboratory technique for generating large numbers of synchronously developing embryos. This synchronicity enhances Xenopus laevis as a powerful model organism for studying vertebrate embryonic development, molecular, cellular, and disease mechanisms. In conventional in vitro fertilization protocols for X. laevis, male frogs are euthanized, and both testes are harvested. In order to reduce the number of required males, a procedure was developed for unilateral gonadectomy in male X. laevis, which allows sequential use of male frogs&#8217; gonads for in vitro fertilizations. This approach has been successfully used to generate embryos, yielding equally good fertilization outcomes with sperm from either the surgically removed or the contralateral testis. The surgical process involves the administration of MS-222 anesthesia, use of Meloxicam and Bupivacaine analgesia, and post-operative care and monitoring, thus allowing reduction in animal use and supporting best practices in animal care across multiple experimental conditions and laboratory setting.

To date, this procedure has been performed on 25 male frogs. One of the earliest frogs died as a result of cutting a major abdominal blood vessel during surgery. The other 24 frogs had successful surgeries. None of these frogs showed any sign of infection or other distress post surgery.
Surgically removed and contralateral testes yielded equally successful in vitro fertilizations, which were consistently observed with all 24 males so far employed. Embryos derived from both testes developed equivalently through cleavage, gastrulation, neurulation, and tailbud stages. Two representative examples for embryos resulting from fertilization with each (surgically removed or contralateral) testis are shown in Figure 4A,B. All embryos exhibited normal morphology with no differences in developmental progression or viability (Figure 4A,B). In all 24 males, the remaining contralateral testis showed no morphological changes or hypertrophy.
Unilateral gonadectomies significantly reduced male frog usage. Figure 4C illustrates frog usage in two-month intervals from January 2023 until April 2026. Frog usage was an average of 3.2 frogs per month from January 2023 to October 2024. Unilateral gonadectomies were begun in November 2024. Since then, an average of 1.7 frogs were used per month. This shows a clear reduction in animal use.
<img alt="Frog embryo development stages; diagram; gonadectomy impact on frog usage over time; data chart." class="xfigimg" src="/files/ftp_upload/71258/71258fig4.jpg" data-altupdatedat="1790049645000" data-alt="Figure 4"> 
Figure 4: Equally effective development resulting from use of surgically removed and contralateral gonads and reduction in frog usage. (A) Independent clutches of embryos derived from surgically removed gonads. (a,b) Fifteen representative 4-cell stage embryos from two clutches. (a’,b’) The same fifteen embryos at tailbud stage. (B) Embryos derived from using contralateral gonads. (c,d) Fifteen representative four-cell embryos. (c’,d’) The same fifteen embryos at tailbud stage. (C) Total number of male frogs used in 2 month periods from January 2023 to April 2026. Gonadectomies were begun in November 2024 and resulted in a significant reduction in male frog usage. <a href="https://www.jove.com/files/ftp_upload/71258/71258fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

We present a protocol for the surgical removal of one testis from adult male X. laevis, which allows the use of the second testis at a later time. This protocol aims to reduce the number of male frogs required for in vitro fertilization.

In vitro fertilization is a reliable laboratory technique for generating large numbers of synchronously developing embryos. This synchronicity enhances ...

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Research

JoVE Journal

Developmental Biology

The eggs of Xenopus laevis intact, lysed, and/or fractionated are useful for a wide variety of experiments. This protocol shows how to induce egg laying, collect and dejelly the eggs, and sort the eggs to remove any damaged eggs.

Obtaining Eggs from Xenopus laevis Females

We describe a method to quantify the activity of K+-countertransporting P-type ATPases by heterologous expression of the enzymes in Xenopus oocytes and measuring Rb+ or Li+ uptake into individual cells by atomic absorption spectrophotometry. The method is a sensitive and safe alternative to radioisotope flux experiments facilitating complex kinetic studies.

Measuring Cation Transport by Na,K- and H,K-ATPase in Xenopus Oocytes by Atomic Absorption Spectrophotometry: An Alternative to Radioisotope Assays

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

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

Protein synthesis control occurs mainly at the translation initiation step, deficiencies in which are linked to diverse disorders. To better understand their etiology, we described here a protocol using Xenopus laevis oocytes assessing the translation of mos transcript in the presence of a mutant of translation initiation factor eIF4G1.

Xenopus laevis as a Model to Identify Translation Impairment

Here, we present a protocol for isolating gonadal tissue of larval zebrafish, which will facilitate investigations of zebrafish sex differentiation and maintenance.

Dissection of Larval Zebrafish Gonadal Tissue

This protocol demonstrates a fluorescence-based method to visualize the vasculature and to quantify its complexity in Xenopus tropicalis. Blood vessels can be imaged minutes after the injection of a fluorescent dye into the beating heart of an embryo after genetic and/or pharmacological manipulations to study cardiovascular development in vivo.

Visualization and Quantitative Analysis of Embryonic Angiogenesis in Xenopus tropicalis

The study of amphibian biology provides valuable information on the reproductive, physiological, embryological and developmental processes that drive organisms of many taxonomic groups. Here, we present a comprehensive guide on different methodologies that can be used to study ovarian control and monitoring in amphibians.

Reproductive Techniques for Ovarian Monitoring and Control in Amphibians

Here, we provide a dissection protocol required to live-image the late embryonic Drosophila male gonad. This protocol will permit observation of dynamic cellular processes under normal conditions or after transgenic or pharmacological manipulation.

Dissection and Live-Imaging of the Late Embryonic Drosophila Gonad

The article describes a quick protocol to gonadectomize and sample blood from the small teleost fish, using Japanese medaka (Oryzias latipes) as a model, to investigate the role of sex steroids in animal physiology.

Gonadectomy and Blood Sampling Procedures in the Small Size Teleost Model Japanese Medaka (Oryzias latipes)

When Transforming Growth Factor ß family precursor proteins are ectopically expressed in Xenopus laevis embryos, they dimerize, get cleaved and are secreted into the blastocoele, which begins at the late blastula to early gastrula stage. We describe a method for aspirating cleavage products from the blastocoele cavity for immunoblot analysis.

Analysis of Transforming Growth Factor &#223; Family Cleavage Products Secreted Into the Blastocoele of Xenopus laevis Embryos

This manuscript describes a consistent way to quickly perform survival rodent orchiectomies and ovariectomies.

Murine Orchiectomy and Ovariectomy to Reduce Sex Hormone Production

Montana State University.  We present a protocol for the surgical removal of one testis from adult male X. laevis, which allows the use of the second testis at a later time. This protocol aims to reduce the number of male frogs required for in vitro fertilization.

Unilateral Gonadectomy in Adult Male Xenopus laevis

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