Efficient Techniques for Comprehensive Sampling of Accessible Tissues in Adult Xenopus

Summary

This is part one of a comprehensive Xenopus sampling protocol. The tissues sampled are the heart ventricle, arterial trunk, left liver lobe, gallbladder, lung, pancreas, spleen, larynx, esophagus, stomach, intestines, testes, fat bodies, oviduct, paired kidneys, sciatic plexus, skin, thymus, and whole eye.

Abstract

Xenopus has long been a pivotal model organism for investigating vertebrate development and disease, offering deep insights into cellular processes and gene function. Despite the wealth of information on embryonic Xenopus, there remains a significant gap in standardized methods for adult tissue sampling, especially for modern approaches like quantitative proteomics. This study introduces a comprehensive protocol for rapid, precise, and efficient sampling of multiple tissues in adult Xenopus. The protocol addresses challenges associated with the subtle anatomical differences compared to other anurans, ensuring reproducibility even for those with limited experience in frog dissection. This protocol is optimized for high-quality biochemical analyses by prioritizing sample freshness. We are facilitating the rapid collection of up to 18 tissues within an hour. Additionally, the methods apply to perfused and unperfused conditions, providing flexibility for a range of experimental needs. This work not only fills a critical methodological gap for Xenopus laevis and tropicalis but also serves as a valuable resource for researchers adapting techniques to similar amphibian models, thereby enhancing the scope and reliability of comparative biological and evolutionary studies.

Introduction

Xenopus offers unique advantages over mice, zebrafish, and other model organisms for studying human disease, particularly in developmental and evolutionary contexts. Its external embryonic development and large, easily manipulable embryos enable real-time observation and experimental interventions (e.g., tissue grafting, gene editing) that are challenging in mice, which develop internally and require invasive procedures. Additionally, Xenopus’ three-chambered heart and tetrapod limbs provide a more direct evolutionary bridge for studying congenital defects¹, limb regeneration², and cardiac development³. Unlike mice, Xenopus tadpoles can regenerate limbs and organs, modeling regenerative processes absent in mammals, while their aquatic life stage allows non-invasive study of environmental or toxin impacts. Combined with faster breeding, lower costs, and fewer ethical constraints, Xenopus is superior for high-throughput, mechanistic studies of organogenesis and stress responses that are difficult to replicate in mammalian systems.

This paper aims to offer clear guidance for precise and reproducible organ sampling of adult Xenopus tissues, as is available for Xenopus tadpoles⁴. While a "digital dissection"⁵ and “micro-CT imaging”⁶ of adult Xenopus is available, achieving consistent organ and tissue sampling in adult Xenopus remains difficult without the detailed instructions provided for other adult models, such as mice^7,8,9.

Although a sample guide for six crucial tissues is available¹⁰, no specific and extensive dissection and sampling guide for Xenopus is currently available. A comprehensive dissection guide exists for Rana sp.¹¹ and various classroom dissection manuals are available for other anurans¹², However, for individuals who are inexperienced in sampling techniques or amphibian anatomy, the subtle differences between Xenopus and other anurans make existing resources inadequate for consistent and replicable tissue sampling.

This guide was developed to prioritize tissue freshness for proteomics and immunochemistry. To an experienced user, all tissues can be collected from one individual in under an hour. It is recommended that if a user lacks experience, they attempt this protocol on specimens that were euthanized for other reasons before sacrificing any animal that is more challenging to replace.

Protocol

All experiments were approved by and performed in accordance with the rules and regulations of the Harvard Medical School IACUC (Institutional Animal Care and Use Committee IS 00001365_3). If perfusion protocol¹³ is followed prior to sampling, skip to step 2.3.

1. Preparation (without perfusion)

1. Ensure that the research institution has approved the euthanasia technique described in this protocol.
2. Prepare a solution of 5 g/L MS-222 (tricaine methanesulfonate) and 5 g/L sodium bicarbonate (see Table of Materials). Check the pH to ensure that it is ≥7.
   NOTE: The volume must be greater than the volume required to completely cover the animals being euthanized.
3. Perform primary euthanasia by placing the Xenopus in the MS-222 solution (from step 1.2), keeping the animal submerged for a total of 1 h¹⁴.
4. Place the dissection surface (tray or foam sheet) at an incline within a secondary container or otherwise arrange it to facilitate blood drainage.
5. Once the frog has been in the solution for 1 h, primary euthanasia has been completed. Remove the frog and check the loss of pain response by performing a foot pinch.
6. Record the age (if known), sex, and health status of the animal being sampled, as well as if it has been perfused. Weigh the Xenopus and take any additional measurements required before sampling.
7. Place the frog on its back and pin down the forelimbs proximal to the body (Figure 1A).
8. Using dissection scissors cut through the skin, up the midline, and then laterally, making two flaps.
9. Identify the linea alba (Figure 1B). Use forceps to grasp the linea alba and pull it away from the coelomic cavity. Carefully use scissors to cut towards the head, through the musculature. Make two flaps out of the cavity wall and cut or pin all flaps out of the way.
10. Identify the heart, which should still be beating. If the heart is not easily accessible, use dissecting scissors to reduce the coracoid bones (Figure 1B).
    NOTE: If the heart has stopped beating prior to sampling, the sample freshness has been compromised.
11. If desired, complete the rapid perfusion protocol⁹.

2. Sampling

NOTE: If the animal has been perfused, skip to step 2.3. All margins should be approximately 25 mm. Forceps and scissors should be wiped clean or replaced between samples. All tissues should be sampled within 1 h of euthanasia, for maximum freshness.

1. Using a transfer pipette, rinse the coelomic cavity with chilled PBS or 0.7x PBS¹⁵ to clear blood and maintain visibility.
2. Identify the thin pericardium (Figure 2A) and pull it taut with tissue forceps. Using the tip of the iridectomy scissors, gently perforate it, being careful not to cut the underlying tissues. Peel the pericardium up away from the three chambers of the heart (Figure 2B).
3. Trim away the auricles by cutting them where they meet the ventricle and arterial trunk.
4. Use forceps to grasp the ventricle by the apex and cut it below where it meets the arterial trunk, leaving a margin (Figure 2C).
5. Visually inspect the ventricle under magnification to ensure no unwanted tissues are attached.
   NOTE: The center of the ventricle may include light-colored spiral valve tissue; this may be kept. In unperfused animals, the removal of the ventricle may qualify as secondary euthanasia (also called terminal anesthesia).
6. Immediately after sampling, rinse the tissue samples in chilled PBS or 0.7x PBS, depending on experimental needs¹⁵.
   NOTE: In perfused animals, it is expected that all tissues excluding the liver will rinse cleanly. If this is not the case, then a note should be made. In unperfused tissues, all rinsing media will be saturated with blood. Steps 2.5–2.6 should be followed after each tissue is sampled.
7. Observe that the arterial trunk bifurcates into two aortic trunks (also called lateral aortae) and that each splits into three arterial arches (Figure 2D). Pull the arterial trunk taut and cut the aortic trunks immediately after where these arches split.
8. Observe the three lobes of the liver (Figure 3A). Grasp the lip of the left lobe (on the viewer's right) and gently lift it so that the hepatic and cystic ducts are visible. Sample the bottom 2/3 of the lobe, below these attachments (Figure 3B).
9. Identify the gallbladder, which may vary drastically in color. Grasp the gallbladder, pull it away from the right lobe of the liver, and sever its attachment (Figure 3C). If it does not burst, then rupture it with iridectomy scissors.
10. Grasp the apex of the lung and pull it taut. Note the texture of the alveoli, and determine where that texture ends and the bronchus begins. Cut at this margin (Figure 3D). If the lung is fused to the peritoneum, carefully detach it.
11. Identify the ovary, which is enveloped in a layer of visceral peritoneum called the germinal epithelium. Gently shift the lobes until they are on their respective sides to make the area of attachment visible (Figure 4A); these attachments are directly ventral to the paired kidney. Using scissors, remove the ovaries as close to the kidneys as possible, without damaging them (Figure 4B). During this process, if oocytes leak out, sample them and then rinse them off.
    NOTE: Removing the ovary is helpful to gain better access to a female frog's tissues.
12. Inspect the anterior lobe (also called the median lobe) of the liver and note how it connects to the stomach and duodenum through the mesentery (Figure 3A and Figure 4C). Sever the mesentery and hepatoduodenal ligament using iridectomy scissors. The remaining fleshy mass is the pancreas and common bile duct.
13. Sever the connection of the pancreas and common bile duct to the anterior lobe of the liver, leaving a margin. Grasp the stomach with toothed forceps and one end of the pancreas with tissue forceps. Under magnification, gently tease the pancreas off of the stomach (Figure 4D).
    1. Should the pancreas not come away cleanly, pick off the remaining pancreatic tissue. Alternatively, detach the pancreas methodically using iridectomy scissors and tissue forceps.
       NOTE: In addition to the common bile duct, it is expected that some mesentery and the pancreatic duct will be included in this sample.
14. Identify the spleen and sample it by cutting its attachment to the peritoneum (Figure 4E).
15. Remove and dispose of the remaining liver tissues.
16. Remove the other lung and the remaining bronchus where they attach to the larynx. Grasping the larynx by its inferior end, begin to lift it out of the cavity, severing clear attachments as they become apparent.
17. As the larynx is lifted, its attachment to the esophagus will become noticeable. Carefully peel and cut the esophageal tissue away from the larynx using curved iridectomy scissors (Figure 5A). Then, pull the larynx down and trim it away from the mouth cavity. Cut any clear attachments that become apparent.
18. Notice the curvature of the stomach, which differentiates it from the esophagus, as well as the pyloric restriction that differentiates the stomach from the intestines (Figure 5B). Cut the alimentary canal immediately anterior to the stomach's curvature, separating it from the esophagus.
19. Open the mouth and identify the shared opening of the eustachian tubes on its roof¹⁶. With the mouth open, tug on the esophagus to note its attachment (Figure 5C). Lift the esophagus out of the cavity, severing clear attachments along the way. Sever the esophagus from the buccal cavity immediately below the eustachian opening.
20. Grasp the stomach and cut it where it meets the duodenum, at the pyloric restriction (Figure 5A,B).
21. Pull the colon taut and cut it as close to the cloaca as possible (Figure 5D). Pull the intestines out, severing any clear peritoneal attachments.
22. If the frog is male, identify the testes and remove one using iridectomy scissors, being careful not to damage the underlying kidney (Figure 6A).
23. Tease apart the fat bodies so that they are on their respective sides; the area over the kidney, where the fat body connects to the peritoneum, will be visible. Grasp the base of either fat body and use scissors to cut it away from the peritoneum, leaving a margin (Figure 6B).
24. Identify and remove the urinary bladder, cutting as close to the cloaca as possible (Figure 6C).
25. If the frog is female or has a distinct vestigial oviduct, grasp the inferior end of the oviduct, pull it away from the cloaca, and cut it as close to the cloaca as possible (Figure 6D). Pull the oviduct out of the coelomic cavity, severing any peritoneal attachments as they become apparent (Figure 6E).
26. Remove and dispose of the remaining fat bodies and testes or oviduct.
27. Use forceps to grasp the kidneys at their inferior base and note that they are covered by the clear peritoneum (retroperitoneal)¹⁷. Sever the peritoneum at the base of the kidney (Figure 7A) and then lift the kidneys out of the coelomic cavity (Figure 7B), using scissors to sever the peritoneum as needed.
28. Under magnification, cut away excess peritoneum and any remaining fat body, ovary, or testes tissue.
29. Observe the bundle of peripheral nerves attaching the terminus of the spinal column to each leg; this is the sciatic plexus. Grasp the bundle of nerves with forceps and sever them where they exit the coelomic cavity (Figure 7C). Lift the bundle out of the cavity and cut it where the nerves leave the vertebral column (Figure 7D). 
30. Remove the pins from the animal, flip it onto its ventrum, and re-pin the animal's limbs. Select either hindlimb to sample from and pin the foot of that limb.
31. Remove an almond-shaped flap of skin from over the gastrocnemius/tibiofibula (Figure 8A,B).
32. Grasp the skin of the dorsum and cut laterally so there is a continuous cut around the midsection of the animal (Figure 8C).
33. Remove the pins, secure the legs in one hand, and grasp the anterior dorsal skin with toothed forceps. Skin the animal by pulling the dorsal skin up over the head and arms, using force (Figure 8D).
34. Identify the fatty mass covering the jaw joint, sever the posterior margin of the mass from where it is fused to pink muscle tissues, and then use iridectomy scissors to cut it away from the underlying bone (Figure 9A). Depending on the animal's age and maturity, a cluster of melanophores will become apparent (Figure 9B). Trim the mass down so that no muscular attachments or cartilage are attached.
35. Repin the animal and select an eye. Insert curved iridectomy scissors into the orbit around the curvature of the eye (Figure 9C). Gently cut around the eye, severing muscular attachments.
36. Slide the scissors deeper into the orbit and detach the optic nerve, behind the eye. Use these scissors to dislodge the eye out of the orbit and then further reduce any muscular attachments.

Results

By following all steps of this protocol, the heart ventricle, arterial trunk, left liver lobe, gallbladder, lung, pancreas, spleen, larynx, esophagus, stomach, intestines, testes, fat bodies, oviduct, paired kidneys, sciatic plexus, skin, thymus, and whole eye were cleanly excised within an hour of euthanasia (see Figure 1, Figure 2, Figure 3, Figure 4,

Discussion

The protocol presented here is designed to provide flexibility for users at various experience levels while ensuring the collection of essential tissues in a timely and efficient manner. New users are advised to follow each step in the protocol as outlined to ensure the systematic collection of tissues. More experienced users, however, may opt to skip certain steps and sample only the tissues they require. For instance, accessing the spleen, which is located behind the mesentery, can be achieved without sampling the panc...

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
1x PBS	Corning	21-040-CV
5x Magnifying Glass with LED Light and Stand	amazon.com	B08QJ6J8P1	light must not produce heat
Disposable Transfer Pipets	VWR	414004-036
Dissecting Fine-Pointed Forceps	Fisher Scientific	08-875
Dissecting scissors sharp piont, straight 6.5"	VWR	76457-374
Dissection Tray	Fisher Scientific	14-370-284	styrofoam sheets are an acceptable alternative
Euthanasia container	US Plastic	Item 2860	alternative opaque containers acceptable
Euthanasia container lid	US Plastic	Item 3047
Iridectomy Scissors 6"	VWR	470018-938	iris scissors are an acceptable alternative
MS-222: Syncaine (formerly tricaine)	Pentair AES	TRS1
Rat Tooth Tissue Forceps 5.5 in., Stainless Steel	Fisher Scientific	S08100
Sodium Bicarbonate, Powder, USP	Fisher Scientific	18-606-333
Specimen Forceps, Serrated	VWR	82027-442
T-Pins for Dissecting	Fisher Scinetific	S99385