Name: Tecniche efficienti per il campionamento completo di tessuti accessibili in Xenopus adulto (Video) | JoVE Journal
Uploaded: 2025-06-10T14:30:00.000+00:00
Duration: 8 min 36 s
Description: Impara tecniche efficienti di campionamento tessutale in Xenopus adulto per analisi biochimiche. Questo metodo ottimizza la freschezza dei tessuti e riduce l'uso di animali in meno di un'ora.

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This work was supported by NIH&#8217;s OD R24 grant ODO31956. We would also like to thank Samantha Jalbert and Jill Ralston for their assistance and support.

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)
<ol>
	<li>Ensure that the research institution has approved the euthanasia technique described in this protocol.</li>
	<li>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 &#8805;7. 
	NOTE: The volume must be greater than the volume required to completely cover the animals being euthanized.</li>
	<li>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.</li>
	<li>Place the dissection surface (tray or foam sheet) at an incline within a secondary container or otherwise arrange it to facilitate blood drainage.</li>
	<li>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.</li>
	<li>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.</li>
	<li>Place the frog on its back and pin down the forelimbs proximal to the body (Figure 1A).</li>
	<li>Using dissection scissors cut through the skin, up the midline, and then laterally, making two flaps.</li>
	<li>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.</li>
	<li>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.</li>
	<li>If desired, complete the rapid perfusion protocol9.</li>
</ol>
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.
<ol>
	<li>Using a transfer pipette, rinse the coelomic cavity with chilled PBS or 0.7x PBS15 to clear blood and maintain visibility.</li>
	<li>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).</li>
	<li>Trim away the auricles by cutting them where they meet the ventricle and arterial trunk.</li>
	<li>Use forceps to grasp the ventricle by the apex and cut it below where it meets the arterial trunk, leaving a margin (Figure 2C).</li>
	<li>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).</li>
	<li>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&#8211;2.6 should be followed after each tissue is sampled.</li>
	<li>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.</li>
	<li>Observe the three lobes of the liver (Figure 3A). Grasp the lip of the left lobe (on the viewer&#39;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).</li>
	<li>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.</li>
	<li>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.</li>
	<li>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&#39;s tissues.</li>
	<li>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.</li>
	<li>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).
	<ol>
		<li>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.</li>
	</ol>
	</li>
	<li>Identify the spleen and sample it by cutting its attachment to the peritoneum (Figure 4E).</li>
	<li>Remove and dispose of the remaining liver tissues.</li>
	<li>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.</li>
	<li>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.</li>
	<li>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&#39;s curvature, separating it from the esophagus.</li>
	<li>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.</li>
	<li>Grasp the stomach and cut it where it meets the duodenum, at the pyloric restriction (Figure 5A,B).</li>
	<li>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.</li>
	<li>If the frog is male, identify the testes and remove one using iridectomy scissors, being careful not to damage the underlying kidney (Figure 6A).</li>
	<li>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).</li>
	<li>Identify and remove the urinary bladder, cutting as close to the cloaca as possible (Figure 6C).</li>
	<li>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).</li>
	<li>Remove and dispose of the remaining fat bodies and testes or oviduct.</li>
	<li>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.</li>
	<li>Under magnification, cut away excess peritoneum and any remaining fat body, ovary, or testes tissue.</li>
	<li>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).&#160;</li>
	<li>Remove the pins from the animal, flip it onto its ventrum, and re-pin the animal&#39;s limbs. Select either hindlimb to sample from and pin the foot of that limb.</li>
	<li>Remove an almond-shaped flap of skin from over the gastrocnemius/tibiofibula (Figure 8A,B).</li>
	<li>Grasp the skin of the dorsum and cut laterally so there is a continuous cut around the midsection of the animal (Figure 8C).</li>
	<li>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).</li>
	<li>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&#39;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.</li>
	<li>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.</li>
	<li>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.</li>
</ol>

<ol><li>Duncan, A. R., Khokha, M. K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Xenopus+as+a+model+organism+for+birth+defects%E2%80%94Congenital+heart+disease+and+heterotaxy%5BTitle%5D)+AND+%22Semin+Cell+Biol%22%5BJournal%5D)">Xenopus as a model organism for birth defects—Congenital heart disease and heterotaxy.</a> Semin Cell Biol. 51, 73(2016).</li>
<li>Kawasumi-Kita, A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((hoxc12%2Fc13+as+key+regulators+for+rebooting+the+developmental+program+in+Xenopus+limb+regeneration%5BTitle%5D)+AND+%22Nat+Commun%22%5BJournal%5D)">hoxc12/c13 as key regulators for rebooting the developmental program in Xenopus limb regeneration.</a> Nat Commun. 15, 3340(2024).</li>
<li>Hoppler, S., Conlon, F. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Xenopus%3A+experimental+access+to+cardiovascular+development%2C+regeneration+discovery%2C+and+cardiovascular+heart-defect+modeling%5BTitle%5D)+AND+%22Cold+Spring+Harb+Perspect+Biol%22%5BJournal%5D)">Xenopus: experimental access to cardiovascular development, regeneration discovery, and cardiovascular heart-defect modeling.</a> Cold Spring Harb Perspect Biol. 12 (6), a037200(2020).</li>
<li>Patmann, M. D., Shewade, L. H., Schneider, K. A., Buchholz, D. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Xenopus+tadpole+tissue+harvest%5BTitle%5D)+AND+%22Cold+Spring+Harb+Protoc%22%5BJournal%5D)">Xenopus tadpole tissue harvest.</a> Cold Spring Harb Protoc. 2017 (11), (2017).</li>
<li>Porro, L. B., Richards, C. T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Digital+dissection+of+the+model+organism+Xenopus+laevis+using+contrast-enhanced+computed+tomography%5BTitle%5D)+AND+%22J+Anat%22%5BJournal%5D)">Digital dissection of the model organism Xenopus laevis using contrast-enhanced computed tomography.</a> J Anat. 231 (2), 169-191 (2017).</li>
<li>Laznovsky, J., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Unveiling+vertebrate+development+dynamics+in+frog+Xenopus+laevis+using+micro-CT+imaging%5BTitle%5D)+AND+%22Gigascience%22%5BJournal%5D)">Unveiling vertebrate development dynamics in frog Xenopus laevis using micro-CT imaging.</a> Gigascience. 13, giae037(2024).</li>
<li>Ruehl-Fehlert, C., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Revised+guides+for+organ+sampling+and+trimming+in+rats+and+mice%E2%80%94Part+1%5BTitle%5D)+AND+%22Exp+Toxicol+Pathol%22%5BJournal%5D)">Revised guides for organ sampling and trimming in rats and mice—Part 1.</a> Exp Toxicol Pathol. 55 (2-3), 91-106 (2003).</li>
<li>Kittel, B., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Revised+guides+for+organ+sampling+and+trimming+in+rats+and+mice%E2%80%94Part+2.+A+joint+publication+of+the+RITA+and+NACAD+groups%5BTitle%5D)+AND+%22Exp+Toxicol+Pathol%22%5BJournal%5D)">Revised guides for organ sampling and trimming in rats and mice—Part 2. A joint publication of the RITA and NACAD groups.</a> Exp Toxicol Pathol. 55 (6), 413-431 (2004).</li>
<li>Morawietz, G., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Revised+guides+for+organ+sampling+and+trimming+in+rats+and+mice%E2%80%94Part+3.+A+joint+publication+of+the+RITA+and+NACAD+groups%5BTitle%5D)+AND+%22Exp+Toxicol+Pathol%22%5BJournal%5D)">Revised guides for organ sampling and trimming in rats and mice—Part 3. A joint publication of the RITA and NACAD groups.</a> Exp Toxicol Pathol. 55 (6), 433-449 (2004).</li>
<li>Jonas-Closs, R. A., Peshkin, L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Techniques+for+rapidly+sampling+six+crucial+organs+in+adult+Xenopus%5BTitle%5D)+AND+%22J+Vis+Exp%22%5BJournal%5D)">Techniques for rapidly sampling six crucial organs in adult Xenopus.</a> J Vis Exp. (204), (2024).</li>
<li>Lőw, P., Molnár, K., Kriska, G. Dissection of a frog (Rana sp.). <a target="_blank" href="http://scholar.google.com/scholar?hl=en&safe=off&q=author%3aL%C5%91w%2C+P+%22Atlas+of+animal+anatomy+and+histology%22">Atlas of animal anatomy and histology</a>. , Springer. Cham. 213-263 (2016).</li>
<li>O'Rourke, D. P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Amphibians+used+in+research+and+teaching%5BTitle%5D)+AND+%22ILAR+J%22%5BJournal%5D)">Amphibians used in research and teaching.</a> ILAR J. 48 (3), 183-187 (2007).</li>
<li>Jonas-Closs, R. A., Peshkin, L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Effective+rapid+blood+perfusion+in+Xenopus%5BTitle%5D)+AND+%22J+Vis+Exp%22%5BJournal%5D)">Effective rapid blood perfusion in Xenopus.</a> J Vis Exp. (195), e65287(2023).</li>
<li>AVMA guidelines for the euthanasia of animals. 37, 2020 edition, AVMA Veterinary Medical Association. 
 <a target="_blank" href="https://www.avma.org/resources-tools/avma-policies/avma-guidelines-euthanasia-animals">https://www.avma.org/resources-tools/avma-policies/avma-guidelines-euthanasia-animals</a> (2020).</li>
<li>Balls, M., Worley, R. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Amphibian+cells+in+vitro.+II.+Effects+of+variations+in+medium+osmolarity+on+a+permanent+cell+line+isolated+from+Xenopus%5BTitle%5D)+AND+%22Exp+Cell+Res%22%5BJournal%5D)">Amphibian cells in vitro. II. Effects of variations in medium osmolarity on a permanent cell line isolated from Xenopus.</a> Exp Cell Res. 76 (2), 333-336 (1973).</li>
<li>Mason, M., Wang, M., Narins, P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Structure+and+function+of+the+middle+ear+apparatus+of+the+aquatic+frog%2C+Xenopus+laevis%5BTitle%5D)+AND+%22Proc+Inst+Acoust%22%5BJournal%5D)">Structure and function of the middle ear apparatus of the aquatic frog, Xenopus laevis.</a> Proc Inst Acoust. 31, 13-21 (2009).</li>
<li>Holz, P. H., Raidal, S. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Comparative+renal+anatomy+of+exotic+species%5BTitle%5D)+AND+%22Vet+Clin+North+Am+Exot+Anim+Pract%22%5BJournal%5D)">Comparative renal anatomy of exotic species.</a> Vet Clin North Am Exot Anim Pract. 9 (1), 1-11 (2006).</li>
<li>Adamson, R. H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Permeability+of+frog+mesenteric+capillaries+after+partial+pronase+digestion+of+the+endothelial+glycocalyx%5BTitle%5D)+AND+%22J+Physiol%22%5BJournal%5D)">Permeability of frog mesenteric capillaries after partial pronase digestion of the endothelial glycocalyx.</a> J Physiol. 428, 1-13 (1990).</li>
<li>O'Rourke, K. P., Ackerman, S., Dow, L. E., Lowe, S. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Isolation%2C+culture%2C+and+maintenance+of+mouse+intestinal+stem+cells%5BTitle%5D)+AND+%22Bio+Protoc%22%5BJournal%5D)">Isolation, culture, and maintenance of mouse intestinal stem cells.</a> Bio Protoc. 6 (4), e1733(2016).</li>
<li>Vitt, L. J., Caldwell, J. P. Anatomy of amphibians and reptiles. <a target="_blank" href="http://scholar.google.com/scholar?hl=en&safe=off&q=author%3aVitt%2C+LJ+%22Herpetology%3A+An+Introductory+Biology+of+Amphibians+and+Reptiles%22">Herpetology: An Introductory Biology of Amphibians and Reptiles</a>. , Academic Press. 35-81 (2009).</li>
</ol>

The authors have no competing interests to declare.

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...

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&#8217; 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 &#34;digital dissection&#34;5 and &#8220;micro-CT imaging&#8221;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 mice7,8,9.
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.

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

efficient techniques for comprehensive sampling of accessible tissues in adult xenopus

Efficient Techniques for Comprehensive Sampling of Accessible Tissues in Adult Xenopus

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.

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, <strong ...

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.

Xenopus has long been a pivotal model organism for investigating vertebrate development and disease, offering deep insights into cellular processes and ...

This work aims to create a replicable method for rapidly collecting the Xenopus heart ventricle, arterial trunk, left liver lobe, gallbladder, lung, pancreas, spleen, larynx, esophagus, stomach, intestines, testes, fat body, oviduct, paired kidneys, sciatic plexus, skin, thymus, and the whole eye in under one hour. An experimental challenge is optimizing tissue freshness. For this reason, the protocol has been designed to be completed in under one hour. By prioritizing tissue freshness, these tissues can be used to complete high quality biochemical analyses such as proteomics, while minimizing the number of animals used. In the future, we'll write a protocol on tissues that are more challenging to access, such as the brain, muscle, and bone.[Instructor] To begin, position the euthanized frog for the dissection procedure and pin down the limbs. Using dissection scissors, cut through the skin along the midline, and then laterally to create two skin flaps. Using forceps, grasp the linea alba and pull it away from the coelomic cavity. Carefully cut through the musculature towards the head and create two flaps of the cavity wall. Reduce the coracoid bones to gain better access to the he

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Animal caps overexpressing gene product(s) are transplanted to the flank of developing Xenopus laevis embryos in order to establish whether tissue is determined.

Tissue Determination Using the Animal Cap Transplant (ACT) Assay in Xenopus laevis

Xenopus embryonic epithelia are an ideal model system to study cell behaviors such as polarity development and shape change during epithelial morphogenesis. Traditional histology of fixed samples is increasingly being complemented by live-cell confocal imaging. Here we demonstrate methods to isolate frog tissues and visualize live epithelial cells and their cytoskeleton using live-cell confocal microscopy.

Live-cell Imaging and Quantitative Analysis of Embryonic Epithelial Cells in Xenopus laevis

In Xenopus embryos, cells from the roof of the blastocoel are pluripotent and can be programmed to generate various tissues. Here, we describe protocols to use amphibian blastocoel roof explants as an assay system to investigate key in vivo and in vitro features of early neural development.

Stem cell-like Xenopus Embryonic Explants to Study Early Neural Developmental Features In Vitro and In Vivo

Here, we present a protocol to use fate maps and lineage tracers to target injections into individual blastomeres that give rise to the kidney of Xenopus laevis embryos.

Technique to Target Microinjection to the Developing Xenopus Kidney

This article presents a convenient and rapid method for visualizing different neuronal cell populations in the central nervous system of Xenopus embryos using immunofluorescent staining on sections.

Assessing Primary Neurogenesis in Xenopus Embryos Using Immunostaining

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

We describe steps that enable fast in situ sampling of a small portion of an individual cell with high precision and minimal invasion using capillary-based micro-sampling, to facilitate chemical characterization of a snapshot of metabolic activity in live embryos using a custom-built single cell capillary electrophoresis and mass spectrometry platform.

Microprobe Capillary Electrophoresis Mass Spectrometry for Single-cell Metabolomics in Live Frog (Xenopus laevis) Embryos

This protocol aims to demonstrate how to microinject a DNA/DOTAP mixture into eyebuds of one day old Xenopus laevis embryos, and how to image and reconstruct individual green fluorescent protein (GFP) expressing optic axonal arbors in tectal midbrains of intact, living Xenopus tadpoles.

Microinjection of DNA into Eyebuds in Xenopus laevis Embryos and Imaging of GFP Expressing Optic Axonal Arbors in Intact, Living Xenopus Tadpoles

We present a two-part protocol that combines fluorescent calcium imaging with in situ hybridization, allowing the experimenter to correlate patterns of calcium activity with gene expression profiles on a single-cell level.

Fluorescent Calcium Imaging and Subsequent In Situ Hybridization for Neuronal Precursor Characterization in Xenopus laevis

Presented here is an effective rapid blood perfusion protocol to prepare tissue samples from African clawed frogs for transcriptomics and proteomics studies.

Effective Rapid Blood Perfusion in Xenopus

This article presents a guide for sampling six significant and diverse organs in adult Xenopus that can be rapidly and easily accessed: the heart ventricle, liver lobe, pancreas, fat bodies, paired kidneys, and skin.

Techniques for Rapidly Sampling Six Crucial Organs in Adult Xenopus

Impara tecniche efficienti di campionamento tessutale in Xenopus adulto per analisi biochimiche. Questo metodo ottimizza la freschezza dei tessuti e riduce l'uso di animali in meno di un'ora.

Efficient Techniques for Comprehensive Sampling of Accessible Tissues in Adult Xenopus

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