This work was supported by NIH&#39;s OD grant R24OD031956. We thank Samantha Jalbert, Jill Ralston, and Cora Anderson for their assistance and support as well as our editor and anonymous peer reviewers for their helpful feedback

All experiments were performed in accordance with the rules and regulations of the Harvard Medical School IACUC (Institutional Animal Care and Use Committee) (IS 00001365_3). The representative results are shown for both a perfused and unperfused mature albino male Xenopus laevis.
1. Experimental preparation 
NOTE: If perfusion protocol8 is being followed before sampling, skip to step 2.2.
<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). The volume must be greater than the volume required to cover the animals being euthanized completely. Check the pH to ensure that it is &#8805;7.</li>
	<li>Perform primary euthanasia by placing the Xenopus in the euthanasia solution; the animal will remain submerged for a total of 1 h.</li>
	<li>Set up the dissection station so that immediately after sampling, all tissues can be rinsed in chilled PBS or 0.7x PBS9 (depending on experimental needs), checked, and trimmed under a 5x (or greater) magnification light. This station must also enable the user to either replace all forceps and scissors or wipe them clean between uses.</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 appropriate details for the animal, such as species, strain, sex, age, and health status as well as if it was perfused. Weigh the Xenopus and take any additional measurements, such as snout-vent length.</li>
	<li>Place the frog on its back and pin down the limbs proximal to the body (Figure 1).</li>
	<li>Using dissection scissors, cut through the skin, up the midline, and then laterally, making two flaps.</li>
	<li>Referring to Figure 2, identify the linea alba and use forceps to grasp it and pull it away from the coelomic cavity. Cut up through the musculature carefully using scissors. Make two flaps out of the cavity wall. Cut or pin all flaps out of the way.</li>
	<li>Identify the heart which will still be beating. Use dissecting scissors to reduce the coracoid bones (Figure 2) to gain better access to the heart. 
	NOTE: If the heart has stopped beating before sampling, it should be noted that the sample freshness has been compromised.</li>
</ol>
2. Sampling
NOTE: If the animal has been perfused, skip to step 2.2.
<ol>
	<li>Identify the thin pericardium and pull it taut with tissue forceps (Figure 3).</li>
	<li>Using the tip of the iridectomy scissors, gently perforate the pericardium, being careful not to cut the underlying tissues. Peel the pericardium up away from the 3 chambers of the heart.</li>
	<li>Use forceps to grasp the ventricle by the apex, identify where it attaches to the auricles and arterial trunk (Figure 4), and cut it below these attachments (Figure 5). If required, trim the ventricle so that no tissues from the auricles or arterial trunk are visible, and light-colored valve tissue will still be visible inside the ventricle. 
	NOTE: In unperfused animals, the removal of the ventricle may qualify as secondary euthanasia.</li>
	<li>The 3 lobes of the liver will be visible (Figure 6 and Figure 7). 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 (Figure 8). Sample the bottom 1/3 of the lobe below these attachments (Figure 9).</li>
	<li>To gain better access to the tissues of a female frog, it is helpful to remove the ovary. 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 10). These attachments are directly ventral to the paired kidney.</li>
	<li>Using scissors, remove the ovaries as close to the kidneys as possible without damaging them (Figure 11).</li>
	<li>Inspect the medial lobe (also called the anterior lobe) of the liver and note how it connects to the stomach and duodenum through the mesentery and hepatopancreatic duct (also called the common bile duct) (Figure 6, Figure 7, and Figure 8).&#160;</li>
	<li>Sever the mesentery, hepatoduodenal ligament using iridectomy scissors as well as the hepatopancreatic duct where it meets the duodenum. Sever the connection of the pancreas and hepatopancreatic duct to the medial lobe of the liver so that no dark liver tissue is attached (Figure 12).</li>
	<li>Grasp the stomach with toothed forceps and the superior end of the pancreas with tissue forceps. Under 5x magnification, gently tease the pancreas off of the stomach (Figure 13). 
	NOTE: If it does not come away cleanly, the remaining pancreatic tissue will be visible and can be picked off in fragments. Alternatively, the pancreas can be methodically detached using iridectomy scissors and tissue forceps.</li>
	<li>Referring to Figure 14A, identify the urinary bladder and remove it, cutting as close to the cloaca as possible. Discard this tissue.</li>
	<li>Referring to Figure 14B, identify the large intestine and pull it taut to sever the large intestine as close to the cloaca as possible. Remove and discard the entire alimentary canal, severing the peritoneum where it attaches to the spleen. The fat bodies will now be fully accessible.</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 the left fat body (on the viewer&#39;s right) and use scissors to cut it away from the peritoneum, leaving a small margin so that the kidney is not damaged (Figure 15).</li>
	<li>Remove and discard the remaining fat body. The paired kidneys will now be fully visible.</li>
	<li>In female frogs or males with distinct vestigial oviducts, grasp an oviduct and pull it away from the kidney and cloaca (Figure 16). Cut the oviduct where it meets the cloaca and continue to pull it away from the kidney, cutting any clear peritoneal attachments as they become apparent. Discard this tissue.</li>
	<li>Repeat this process with the remaining oviduct.</li>
	<li>The kidneys are still covered with clear peritoneum (retroperitoneal)10. Use forceps to grasp the kidneys and sever the peritoneum at their inferior end.</li>
	<li>Lift the kidneys out of the coelomic cavity, using scissors to sever the peritoneum as close to the kidneys as possible without damaging them (Figure 17).</li>
	<li>Under 5x magnification, cut away excess peritoneum and any other remaining tissue (fat bodies, spleen). If the frog is female, ensure any remaining ovary tissue is removed (Figure 18). If the frog is male, carefully remove the testis and check for a vestigial oviduct, which may not be visible without magnification (Figure 19).</li>
	<li>Remove the pins from the animal, flip it onto its ventrum, and re-pin the animal&#39;s limbs.</li>
	<li>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 20).</li>
</ol>

Standardized Dissection and Sampling Protocol for Adult Xenopus Organs

<ol>
	<li>Porro, L. B., Richards, C. T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Digital+dissection+of+the+model+organism+Xenopus+laevis+using+contrast-enhanced+computed+tomography.">Digital dissection of the model organism Xenopus laevis using contrast-enhanced computed tomography.</a> J Anat. 231 (2), 169-191 (2017).</li><li>Ruehl-Fehlert, C., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Revised+guides+for+organ+sampling+and+trimming+in+rats+and+mice--part+1.">Revised guides for organ sampling and trimming in rats and mice--part 1.</a> Exp Toxicol Pathol. 55 (23), 91-106 (2003).</li><li>Kittel, B., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Revised+guides+for+organ+sampling+and+trimming+in+rats+and+mice--Part+2.+A+joint+publication+of+the+RITA+and+NACAD+groups.">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, 413-431 (2004).</li><li>Morawietz, G., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Revised+guides+for+organ+sampling+and+trimming+in+rats+and+mice+-+Part+3+-+A+joint+publication+of+the+RITA+and+NACAD+groups.">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, 433-449 (2004).</li><li>Patmann, M. D., Shewade, L. H., Schneider, K. A., Buchholz, D. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Xenopus+tadpole+tissue+harvest.">Xenopus tadpole tissue harvest.</a> Cold Spring Harb Protoc. 2017 (11), 097675 (2017).</li><li>L&#337;w, P., Moln&#225;r, K., Kriska, G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Dissection+of+a+Frog+(Rana+sp.).">Dissection of a Frog (Rana sp.).</a> Atlas of Animal Anatomy and Histology. , 213-263 (2016).</li><li>O'Rourke, D. P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Amphibians+used+in+research+and+teaching.">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/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Effective+rapid+blood+perfusion+in+Xenopus.">Effective rapid blood perfusion in Xenopus.</a> JoVE. (issue), e65287 (2023).</li><li>Balls, M., Worley, R. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Amphibian+cells+in+vitro.+II.+Effects+of+variations+in+medium+osmolarity+on+a+permanent+cells+line+isolated+from+Xenopus.">Amphibian cells in vitro. II. Effects of variations in medium osmolarity on a permanent cells line isolated from Xenopus.</a> Exp Cell Res. 76 (2), 333-336 (1973).</li><li>Holz, P. H., Raidal, S. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Comparative+renal+anatomy+of+exotic+species.">Comparative renal anatomy of exotic species.</a> Vet North Am Exot Anim Pract. 9 (1), 1-11 (2006).</li><li>Trott, K. A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Characterization+of+a+Mycobacterium+ulcerans-like+infection+in+a+colony+of+African+tropical+clawed+frogs+(Xenopus+tropicalis).">Characterization of a Mycobacterium ulcerans-like infection in a colony of African tropical clawed frogs (Xenopus tropicalis).</a> Comp Med. 54 (3), 309-317 (2004).</li><li>Fremont-Rahl, J. J., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Mycobacterium+liflandii+outbreak+in+a+research+colony+of+Xenopus+(Silurana)+tropicalis+frogs.">Mycobacterium liflandii outbreak in a research colony of Xenopus (Silurana) tropicalis frogs.</a> Vet Pathol. 48 (4), 856-867 (2011).</li><li>Green, S. L., Parker, J., Davis, C., Bouley, D. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Ovarian+hyperstimulation+syndrome+in+gonadotropin-treated+laboratory+South+African+clawed+frogs+(Xenopus+laevis).">Ovarian hyperstimulation syndrome in gonadotropin-treated laboratory South African clawed frogs (Xenopus laevis).</a> J Am Assoc Lab Anim Sci. 46 (3), 64-67 (2007).</li><li>Vitt, L. J., Caldwell, J. P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Anatomy+of+amphibians+and+reptiles.+Herpetol.">Anatomy of amphibians and reptiles. Herpetol.</a> Herpetol. , 35-81 (2009).</li></ol>

The authors declare no competing interests.

As this protocol aims to maximize freshness, some samples may include undesired tissues. For example, the hepatopancreatic duct and some mesentery are sampled with the pancreas, and some peritoneal tissue, adrenal glands, and ureters will always be sampled with the paired kidneys.&#160; If freshness is not a concern, then more precise sampling can be achieved using modified techniques.
The appearance and location of organs are comparable between sexes and species of Xenopus. However, the color of tissues varies significantly based on whether or not the animals have been perfused. It is for this reason that images of both perfused and unperfused animals are included.
A constraint of this protocol is that speed and reproducibility are prioritized over collecting samples that best represent the entirety of the desired tissue. For example, the section of the left lobe of the liver sampled here cannot adequately represent all three lobes of liver tissue. If there are errors in sampling, options for troubleshooting are affected by the potential for variation between different sections of tissue. For example, it is not known if the right lobe of the liver, the right fat body, or a different portion of skin would be functional alternatives to the desired tissues. In these cases, discretion should be used, based on the needs of the research, before substituting sections of tissue.
Another limitation of this protocol is that if the animals being sampled have drastic anatomical defects or clinically significant health issues, the organs in the coelomic cavity may not appear as described here. Granulomas have been found in the tissues of frogs infected with Mycobacterium spp.11,12, and previous cases of ovarian hyperstimulation syndrome appear to lead to an abnormal presentation of organs13.
Though this method has been developed for laboratory Xenopus, there are significant similarities in the appearance of these organs within many non-caecilian amphibians and limbed reptiles14. The sampling portion of this protocol could easily be modified for other models, such as axolotls or the green anole.

Though the &#34;digital dissection&#34; of adult Xenopus&#8221; is available1, replicable organ and tissue sampling of adult Xenopus remains challenging without the detailed instruction available for other adult models (e.g. mice2,3,4). This article aims to provide clear guidance for accurate and replicable organ sampling of adult Xenopus similar to what is currently available for their larvae5. An emphasis is placed on ease of completion to maintain maximum freshness and make the protocol accessible to all users.
Though there is a thorough dissection guide for Rana sp.6, as well as numerous classroom dissection guides for other anurans7, no Xenopus dissection and sampling guide is currently available. For those not familiar with sampling practices or amphibian anatomy, the small differences between Xenopus and other anurans render these resources suboptimal for replicable tissue sampling.
Many valuable tissues are not included and are even discarded in the present guide; this is to ensure tissue freshness. Six samples are limited enough to ensure that these tissues can be collected in under an hour after the heart starts beating, regardless of the experience or skill level of the user. More advanced and detailed guides for collecting many other tissues are under preparation as separate companion papers.
For less experienced users, it is always recommended that this protocol be first attempted on animals being euthanized for reasons other than experimentation before sampling any animals that are challenging to replace (i.e., transgenics, animals of advanced age, etc). Ideally, all animals sampled will be healthy and, if female, will not have been ovulated in the past two weeks.

<table><tbody><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 Scinetific</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 Scinetific</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>PBS 1x</td><td>Corning</td><td>21-040-CV</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>

techniques for rapidly sampling six crucial organs in adult xenopus

Xenopus has been a powerful model organism for understanding vertebrate development and disease for over a hundred years. While experimental analysis and dissection techniques of the embryo have been well documented, descriptions of adult Xenopus structures and organs, together with techniques for working with adults, have not been updated to take into consideration the requirements of such modern approaches as quantitative proteomics and single-cell transcriptomics. The cell-type and gene-centric perspectives require contrasting observations in embryonic stages to those in adult tissues. The organs of the larva undergo significant changes in their overall structure, morphology, and anatomical location all along the larval to adult transition, most notably during massive metamorphosis remodeling. Establishing robust standards for organ identification and dissection is crucial to ensure datasets resulting from studies performed at different laboratories can be consistent. The present protocol identifies six of the organs in the adult Xenopus, demonstrating methods for dissection and sampling of the heart ventricle, liver, fat body, pancreas, paired kidney, and skin of the adult Xenopus. Depending on the preservation methods, the dissected organs can be used for quantitative proteomics, single cell/nuclei transcriptomics, in situ hybridization, immunohistochemistry, histology, etc. This protocol aims to standardize tissue sampling and facilitate multi-lab investigations of the adult organ systems.

By utilizing Figure 1 to Figure 20 and following all steps of this protocol, the heart ventricle, the left lobe of the liver, the pancreas, the left fat bodies, paired kidneys, and a flap of skin were cleanly excised within an hour of euthanasia. Within this time, the samples are rinsed and trimmed so that they will appear, as shown in Figure 21.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/66489/66489fig1.JPG" /> 
Figure 1: Pinned Xenopus. A mature female X. tropicalis pinned through each limb. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/66489/66489fig1large.JPG" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/66489/66489fig2.jpg" /> 
Figure 2: Abdominal wall. The ventral skin of an X. tropicalis female is cut into flaps, making the linea alba and coracoid bones visible. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/66489/66489fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/66489/66489fig3.JPG" /> 
Figure 3: Pericardium enclosed heart. The apex of the heart ventricle is grasped through the pericardium. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/66489/66489fig3large.JPG" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/66489/66489fig4.JPG" /> 
Figure 4: Heart ventricle and arterial trunk. The ventricle of a perfused X laevis, being grasped, showing its attachment to the arterial trunk. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/66489/66489fig4large.JPG" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/66489/66489fig5.JPG" /> 
Figure 5: Heart diagram. A diagram of the relevant structures of the heart with a dashed line indicating where to sample the ventricle. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/66489/66489fig5large.JPG" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 6" class="xfigimg" src="/files/ftp_upload/66489/66489fig6.JPG" /> 
Figure 6: Hepatopancreatic diagram. A diagram of the 3 lobes of the liver, pancreas, and associated organs. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/66489/66489fig6large.JPG" target="_blank">Please click here to view a larger version of this figure.</a>

<img alt="Figure 7" class="xfigimg" src="/files/ftp_upload/66489/66489fig7.JPG" /> 
Figure 7: Hepatopancreatic organs. A perfused, albino X. laevis male with 3 lobes of liver, pancreas, and associated organs labeled. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/66489/66489fig7large.JPG" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 8" class="xfigimg" src="/files/ftp_upload/66489/66489fig8.JPG" /> 
Figure 8: Cystic and hepatic ducts. The left lobe of the liver is being lifted to show the cystic and hepatic ducts in perfused X. laevis. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/66489/66489fig8large.JPG" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 9" class="xfigimg" src="/files/ftp_upload/66489/66489fig9.JPG" /> 
Figure 9: Liver sampling. The left liver lobe of an unperfused X. tropicalis is severed under the attachments of the hepatic ducts. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/66489/66489fig9large.JPG" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 10" class="xfigimg" src="/files/ftp_upload/66489/66489fig10.JPG" /> 
Figure 10: Ovary attachment. With the ovary lobes on their respective sides, the continuity of the germinal epithelium to the peritoneal wall (over the kidneys) is visible. Two white dashed lines indicate where to sever these attachments.&#160; <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/66489/66489fig10large.JPG" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 11" class="xfigimg" src="/files/ftp_upload/66489/66489fig11.JPG" /> 
Figure 11: Ovary removal. The ovary of an unperfused X. laevis, is pulled away from the paired kidneys. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/66489/66489fig11large.JPG" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 12" class="xfigimg" src="/files/ftp_upload/66489/66489fig12.JPG" /> 
Figure 12: Mesentery incisions. The coelomic cavity of an unperfused X. laevis, following the sampling of the heart ventricle and left lobe of the liver as well as the removal of the ovary. A white dashed line indicates where to sever the hepatopancreatic ligament and duct, while a green dashed line indicates where to sever the pancreas from the medial lobe of the liver. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/66489/66489fig12large.JPG" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 13" class="xfigimg" src="/files/ftp_upload/66489/66489fig13.jpg" /> 
Figure 13: Pancreas sampling. The pancreas of an unperfused X. laevis is being teased off of the stomach. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/66489/66489fig13large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 14" class="xfigimg" src="/files/ftp_upload/66489/66489fig14.JPG" /> 
Figure 14: Organ removal. (A) The urinary bladder of an unperfused X. laevis is pulled away from the cloaca with a dashed line indicating where to cut it. (B) The large intestine of an unperfused X. laevis, is being pulled away from the cloaca with a dashed line indicating where to sever it. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/66489/66489fig14large.JPG" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 15" class="xfigimg" src="/files/ftp_upload/66489/66489fig15.JPG" /> 
Figure 15: Fat body sampling. The fat bodies, attached to the peritoneum at the superior end of the paired kidneys, are pulled out of the coelomic cavity with a dashed line showing where to cut them. Note that adjacent to this attachment, this male X. tropicalis has 1 testis as well as a pair of distinct vestigial oviducts. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/66489/66489fig15large.JPG" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 16" class="xfigimg" src="/files/ftp_upload/66489/66489fig16.JPG" /> 
Figure 16: Oviduct removal. The oviduct of a perfused X. laevis is tugged away from the paired kidney, making the clear peritoneum visible. A dashed line indicates where to incise the peritoneum. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/66489/66489fig16large.JPG" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 17" class="xfigimg" src="/files/ftp_upload/66489/66489fig17.JPG" /> 
Figure 17: Kidney sampling. The paired kidneys of an unperfused X. laevis are being lifted out of the coelomic cavity. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/66489/66489fig17large.JPG" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 18" class="xfigimg" src="/files/ftp_upload/66489/66489fig18.jpg" /> 
Figure 18: Kidney trimming. (A) A ventral view of an unperfused female X. laevis&#8217;s paired kidney with associated peritoneal organs attached. (B) The same kidney with associated organs removed but with some peritoneal tissue remaining. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/66489/66489fig18large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 19" class="xfigimg" src="/files/ftp_upload/66489/66489fig19.JPG" /> 
Figure 19: Testis removal. The paired kidneys of unperfused X. tropicalis with one testis removed. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/66489/66489fig19large.JPG" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 20" class="xfigimg" src="/files/ftp_upload/66489/66489fig20.jpg" /> 
Figure 20: Skin sampling. (A) The right leg of an X. tropicalis with a dashed line indicating the area of skin to be sampled. (B) The right leg of an X. tropicalis with a skin sample removed over the tibiofibula. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/66489/66489fig20large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 21" class="xfigimg" src="/files/ftp_upload/66489/66489fig21.jpg" /> 
Figure 21: Representative results of organ sampling. Samples of heart ventricle, liver, pancreas, fat body, paired kidney, and skin taken from a perfused and unperfused albino X. laevis. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/66489/66489fig21large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Author Spotlight: Standardizing Tissue Sampling in Proteomics and Immunochemistry Research at JoVE.com

Author Spotlight: Standardizing Tissue Sampling in Proteomics and Immunochemistry Research

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

Xenopus has been a powerful model organism for understanding vertebrate development and disease for over a hundred years. While experimental analysis and ...

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Nanyang Technological University

Research

JoVE Journal

Biology

827 Views.  Harvard Medical School. 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.

Video: Author Spotlight: Standardizing Tissue Sampling in Proteomics and Immunochemistry Research

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

Understanding the genetic programs underlying neural development is an important goal of developmental and stem cell biology. In the amphibian blastula, cells from the roof of the blastocoel are pluripotent. These cells can be isolated, and programmed to generate various tissues through manipulation of genes expression or induction by morphogens. In this manuscript protocols are described for the use of Xenopus laevis blastocoel roof explants as an assay system to investigate key in vivo and in vitro features of early neural development. These protocols allow the investigation of fate acquisition, cell migration behaviors, and cell autonomous and non-autonomous properties. The blastocoel roof explants can be cultured in a serum-free defined medium and grafted into host embryos. This transplantation into an embryo allows the investigation of the long-term lineage commitment, the inductive properties, and the behavior of transplanted cells in vivo. These assays can be exploited to investigate molecular mechanisms, cellular processes and gene regulatory networks underlying neural development. In the context of regenerative medicine, these assays provide a means to generate neural-derived cell types in vitro that could be used in drug screening.

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

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.

Understanding the genetic programs underlying neural development is an important goal of developmental and stem cell biology. In the amphibian blastula, ...

Developmental Biology

Technique to Target Microinjection to the Developing Xenopus Kidney

The embryonic kidney of Xenopus&#160;laevis (frog), the pronephros, consists of a single nephron, and can be used as a model for kidney disease. Xenopus embryos are large, develop externally, and can be easily manipulated by microinjection or surgical procedures. In addition, fate maps have been established for early Xenopus embryos. Targeted microinjection into the individual blastomere that will eventually give rise to an organ or tissue of interest can be used to selectively overexpress or knock down gene expression within this restricted region, decreasing secondary effects in the rest of the developing embryo. In this protocol, we describe how to utilize established Xenopus fate maps to target the developing Xenopus kidney (the pronephros), through microinjection into specific blastomere of 4- and 8-cell embryos. Injection of lineage tracers allows verification of the specific targeting of the injection. After embryos have developed to stage 38 - 40, whole-mount immunostaining is used to visualize pronephric development, and the contribution by targeted cells to the pronephros can be assessed. The same technique can be adapted to target other tissue types in addition to the pronephros.

Technique to Target Microinjection to the Developing Xenopus Kidney

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.

The embryonic kidney of Xenopus&#160;laevis (frog), the pronephros, consists of a single nephron, and can be used as a model for kidney disease. Xenopus ...

Visualization and Quantitative Analysis of Embryonic Angiogenesis in Xenopus tropicalis

Blood vessels supply oxygen and nutrients throughout the body, and the formation of the vascular network is under tight developmental control. The efficient in vivo visualization of blood vessels and the reliable quantification of their complexity are key to understanding the biology and disease of the vascular network. Here, we provide a detailed method to visualize blood vessels with a commercially available fluorescent dye, human plasma acetylated low density lipoprotein DiI complex (DiI-AcLDL), and to quantify their complexity in Xenopus tropicalis. Blood vessels can be labeled by a simple injection of DiI-AcLDL into the beating heart of an embryo, and blood vessels in the entire embryo can be imaged in live or fixed embryos. Combined with gene perturbation by the targeted microinjection of nucleic acids and/or the bath application of pharmacological reagents, the roles of a gene or of a signaling pathway on vascular development can be investigated within one week without resorting to sophisticated genetically engineered animals. Because of the well-defined venous system of Xenopus and its stereotypic angiogenesis, the sprouting of pre-existing vessels, vessel complexity can be quantified efficiently after perturbation experiments. This relatively simple protocol should serve as an easily accessible tool in diverse fields of cardiovascular research.

Visualization and Quantitative Analysis of Embryonic Angiogenesis in Xenopus tropicalis

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.

Blood vessels supply oxygen and nutrients throughout the body, and the formation of the vascular network is under tight developmental control. The ...

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

The quantification of small molecules in single cells raises new potentials for better understanding the basic processes that underlie embryonic development. To enable single-cell investigations directly in live embryos, new analytical approaches are needed, particularly those that are sensitive, selective, quantitative, robust, and scalable to different cell sizes. Here, we present a protocol that enables the in situ analysis of metabolism in single cells in freely developing embryos of the South African clawed frog (Xenopus laevis), a powerful model in cell and developmental biology. This approach uses a capillary microprobe to aspirate a defined portion from single identified cells in the embryo, leaving neighboring cells intact for subsequent analysis. The collected cell content is analyzed by a microscale capillary electrophoresis electrospray ionization (CE-ESI) interface coupled to a high-resolution tandem mass spectrometer. This approach is scalable to various cell sizes and compatible with the complex three-dimensional structure of the developing embryo. As an example, we demonstrate that microprobe single-cell CE-ESI-MS enables the elucidation of metabolic cell heterogeneity that unfolds as a progenitor cell gives rise to descendants during development of the embryo. Besides cell and developmental biology, the single-cell analysis protocols described here are amenable to other cell sizes, cell types, or animal models.

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

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.

The quantification of small molecules in single cells raises new potentials for better understanding the basic processes that underlie embryonic ...

Chemistry

Simultaneous Pre- and Post-synaptic Electrophysiological Recording from Xenopus Nerve-muscle Co-cultures

Much information about the coupling of presynaptic ionic currents with the release of neurotransmitter has been obtained from invertebrate preparations, most notably the squid giant synapse1. However, except for the preparation described here, few vertebrate preparations exist in which it is possible to make simultaneous measurements of neurotransmitter release and presynaptic ionic currents. Embryonic Xenopus motoneurons and muscle cells can be grown together in simple culture medium at room temperature; they will form functional synapses within twelve to twenty-four hours, and can be used to study nerve and muscle cell development and synaptic interactions for several days (until overgrowth occurs). Some advantages of these co-cultures over other vertebrate preparations include the simplicity of preparation, the ability to maintain the cultures and work at room temperature, and the ready accessibility of the synapses formed2-4. The preparation has been used widely to study the biophysical properties of presynaptic ion channels and the regulation of transmitter release5-8. In addition, the preparation has lent itself to other uses including the study of neurite outgrowth and synaptogenesis9-12, molecular mechanisms of neurotransmitter release13-15, the role of diffusible messengers in neuromodulation16,17, and in vitro synaptic plasticity18-19.

Simultaneous Pre- and Post-synaptic Electrophysiological Recording from Xenopus Nerve-muscle Co-cultures

This video demonstrates the procedures used to grow primary cultures of embryonic Xenopus nerve and muscle cells and the usefulness of this preparation for making simultaneous pre- and post-synaptic patch clamp recordings.

Much information about the coupling of  presynaptic ionic currents with the release of neurotransmitter has been obtained  from invertebrate preparations, ...

Neuroscience

Assessing Primary Neurogenesis in Xenopus Embryos Using Immunostaining

Primary neurogenesis is a dynamic and complex process during embryonic development that sets up the initial layout of the central nervous system. During this process, a portion of neural stem cells undergo differentiation and give rise to the first populations of differentiated primary neurons within the nascent central nervous system. Several vertebrate model organisms have been used to explore the mechanisms of neural cell fate specification, patterning, and differentiation. Among these is the African clawed frog, Xenopus, which provides a powerful system for investigating the molecular and cellular mechanisms responsible for primary neurogenesis due to its rapid and accessible development and ease of embryological and molecular manipulations. Here, we present a convenient and rapid method to observe the different populations of neuronal cells within Xenopus central nervous system. Using antibody staining and immunofluorescence on sections of Xenopus embryos, we are able to observe the locations of neural stem cells and differentiated primary neurons during primary neurogenesis.

Assessing Primary Neurogenesis in Xenopus Embryos Using Immunostaining

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.

Primary neurogenesis is a dynamic and complex process during embryonic development that sets up the initial layout of the central nervous system. During ...

Functional Evaluation of Olfactory Pathways in Living Xenopus Tadpoles

Xenopus tadpoles offer a unique platform to investigate the function of the nervous system. They provide multiple experimental advantages, such as accessibility to numerous imaging approaches, electrophysiological techniques and behavioral assays. The Xenopus tadpole olfactory system is particularly well suited to investigate the function of synapses established during normal development or reformed after injury. Here, we describe methodologies to evaluate the processing of olfactory information in living Xenopus larvae. We outline a combination of in vivo measurements of presynaptic calcium responses in glomeruli of the olfactory bulb with olfactory-guided behavior assays. Methods can be combined with the transection of olfactory nerves to study the rewiring of synaptic connectivity. Experiments are presented using both wild-type and genetically modified animals expressing GFP reporters in central nervous system cells. Application of the approaches described to genetically modified tadpoles can be useful for unraveling the molecular bases that define vertebrate behavior.

Functional Evaluation of Olfactory Pathways in Living Xenopus Tadpoles

Xenopus tadpoles offer a unique platform to investigate the function of the nervous system in vivo. We describe methodologies to evaluate the processing of olfactory information in living Xenopus larvae in normal rearing conditions or after injury.

Xenopus tadpoles offer a unique platform to investigate the function of the nervous system. They provide multiple experimental advantages, such as ...

The Use of Ex Vivo Whole-organ Imaging and Quantitative Tissue Histology to Determine the Bio-distribution of Fluorescently Labeled Molecules

Fluorescent labeling is a well-established process for examining the fate of labeled molecules under a variety of experimental conditions both in vitro and in vivo. Fluorescent probes are particularly useful in determining the bio-distribution of administered large molecules, where the addition of a small-molecule fluorescent label is unlikely to affect the kinetics or bio-distribution of the compound. A variety of methods exist to examine bio-distribution that vary significantly in the amount of effort required and whether the resulting measurements are fully quantitative, but using multiple methods in conjunction can provide a rapid and effective system for analyzing bio-distributions.
Ex vivo whole-organ imaging is a method that can be used to quickly compare the relative concentrations of fluorescent molecules within tissues and between multiple types of tissues or treatment groups. Using an imaging platform designed for live-animal or whole-organ imaging, fluorescence within intact tissues can be determined without further processing, saving time and labor while providing an accurate picture of the overall bio-distribution. This process is ideal in experiments attempting to determine the tissue specificity of a compound or for the comparison of multiple different compounds. Quantitative tissue histology on the other hand requires extensive further processing of tissues in order to create a quantitative measure of the labeled compounds. To accurately assess bio-distribution, all tissues of interest must be sliced, scanned, and analyzed relative to standard curves in order to make comparisons between tissues or groups. Quantitative tissue histology is the gold standard for determining absolute compound concentrations within tissues.
Here, we describe how both methods can be used together effectively to assess the ability of different administration methods and compound modifications to target and deliver fluorescently labeled molecules to the central nervous system1.

The Use of Ex Vivo Whole-organ Imaging and Quantitative Tissue Histology to Determine the Bio-distribution of Fluorescently Labeled Molecules

Ex vivo whole-organ imaging is a rapid method for determining the relative concentrations of fluorescently labeled compounds within and between tissues or treatment groups. Conversely, quantitative fluorescence histology, while more labor intensive, allows for the quantification of the absolute tissue levels of labeled molecules.

Fluorescent labeling is a well-established process for examining the fate of labeled molecules under a variety of experimental conditions both in vitro ...

An Apical Resection Model in the Adult Xenopus tropicalis Heart

It is known that in adult mammals, the heart has lost its regenerative capacity, making heart failure one of the leading causes of death worldwide. Previous research has demonstrated the regenerative ability of the heart of the adult Xenopus tropicalis, an anuran amphibian with a diploid genome and a close evolutionary relationship with mammals. Additionally, studies have shown that following ventricular apex resection, the heart can regenerate without scarring in X. tropicalis. Consequently, these previous results suggest that X. tropicalis is an appropriate alternative vertebrate model for the study of adult heart regeneration. A surgical model of cardiac regeneration in the adult X. tropicalis is presented herein. Briefly, the frogs were anesthetized and fixed; then, a small incision was made with iridectomy scissors, penetrating the skin and pericardium. Gentle pressure was applied to the ventricle, and the apex of the ventricle was then cut out with scissors. Cardiac injury and regeneration were confirmed by histology at 7-30 days post resection (dpr). This protocol established an apical resection model in adult X. tropicalis, which&#160;can be employed to elucidate the mechanisms of adult heart regeneration.

An Apical Resection Model in the Adult Xenopus tropicalis Heart

Xenopus tropicalis is an ideal model for regenerative research as many of its organs possess a remarkable regenerative capacity. Here, we present a method for constructing a heart injury model in X. tropicalis via apex resection.

It is known that in adult mammals, the heart has lost its regenerative capacity, making heart failure one of the leading causes of death worldwide. ...

Medicine

Blood Perfusion in African Clawed Frogs for Transcriptomics and Proteomics

Effective Rapid Blood Perfusion in Xenopus

Xenopus have been powerful model organisms for understanding vertebrate development and disease for over 100 years. Here, a rapid blood perfusion protocol in Xenopus, aimed at a consistent and drastic reduction of blood within all tissues, is defined. Perfusion is carried out by inserting a needle directly into the ventricle of the heart and pumping heparinized phosphate-buffered saline (PBS) through the vascular system. The procedure can be completed in approximately 10 min per animal. The blood is dominated by a few highly abundant proteins and cell types, creating numerous issues as these proteins mask most other molecules and cell types of interest. The reproducible characterization of adult Xenopus tissues with quantitative proteomics and single-cell transcriptomics will benefit from applying this protocol prior to organ sampling. The protocols for tissue sampling are defined in companion papers. These procedures are aimed at the standardization of practices across Xenopus of different sex, age, and health status, specifically X. laevis and X. tropicalis.

Author Spotlight: Effective Rapid Blood Perfusion in Xenopus  

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

Xenopus have been powerful model organisms for understanding vertebrate development and disease for over 100 years. Here, a rapid blood perfusion protocol ...

Techniques for Rapidly Sampling Six Crucial Organs in Adult Xenopus

Summary

Explore More Videos

Chapters in this video

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

Technique to Target Microinjection to the Developing Xenopus Kidney

Visualization and Quantitative Analysis of Embryonic Angiogenesis in Xenopus tropicalis

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

Simultaneous Pre- and Post-synaptic Electrophysiological Recording from Xenopus Nerve-muscle Co-cultures

Assessing Primary Neurogenesis in Xenopus Embryos Using Immunostaining

Functional Evaluation of Olfactory Pathways in Living Xenopus Tadpoles

The Use of Ex Vivo Whole-organ Imaging and Quantitative Tissue Histology to Determine the Bio-distribution of Fluorescently Labeled Molecules

An Apical Resection Model in the Adult Xenopus tropicalis Heart

Effective Rapid Blood Perfusion in Xenopus