This work was supported by NIH&#39;s OD R24 grant OD031956 and NICHD R01 grant HD073104. We thank Darcy Kelly for helpful discussions and initial input on this protocol. We would also like to thank Samantha Jalbert, Jill Ralston, and Wil Ratzan for their assistance and support as well as our three anonymous peer reviewers for their 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).
NOTE: Though the primary method of euthanasia described is deemed an acceptable technique for euthanasia by the American Veterinary Medical Association12, it has not been found to lead to the cessation of a heartbeat13. Even the frequently used secondary method of double pithing does not prevent this, nor does removing the heart from the animal. Exsanguination of anesthetized animals is considered a humane and effective method for successful euthanasia12. As maintaining fresh tissues through euthanasia is the goal of this protocol, it is beneficial that the heart continues to beat through primary euthanasia with MS-222, and that perfusion is itself a secondary euthanasia method through exsanguination.
1. Preparation
<ol>
	<li>Ensure that the research institution has approved the euthanasia and perfusion technique described in this protocol.</li>
	<li>Prepare a solution of 5 g/L MS-222 (tricaine methanesulfonate) and 5 g/L sodium bicarbonate. The volume should be greater than the volume required to completely cover the animals being euthanized. Check the pH to ensure that it is &#8805;7.</li>
	<li>Prepare 500 &#181;L of 180 U/mL heparin in phosphate-buffered saline (PBS) per X. laevis, or 200 &#181;L per X. tropicalis.</li>
	<li>Perform primary euthanasia by placing the Xenopus in this solution (from step 1.2); the animal will remain submerged for a total of 1 h.</li>
	<li>Confirm that the Xenopus has lost its pain response by pinching the foot 15 min into euthanasia. If the animal is reactive, return it to the euthanasia solution until this response is lost.</li>
	<li>Weigh the Xenopus and take any additional measurements required before sampling.</li>
	<li>Using a 31 G needle, inject X. laevis with 250 &#181;L and X. tropicalis with 100 &#181;L of 180 U/mL heparin in PBS (from step 1.3) into the musculature of each forelimb.</li>
	<li>Prepare a solution of 1 mL/g of the animal weight of 54 U/mL heparin in PBS perfusate. Individuals more experienced with this protocol may find that less media is required to complete perfusion.</li>
	<li>Use a 22 G hypodermic needle to perfuse X. laevis, and a 25 G hypodermic needle for X. tropicalis. Blunt the perfusion needle by trimming off the tip with wire cutters (Figure 1)14. 
	NOTE: This reduces the likelihood of the needle perforating through the ventricle if shifted. In addition to blunting, the needle may be ground down slightly with a sharpening stone or file, but still remaining sharp enough to pierce the ventricle.</li>
	<li>Prepare the pump by attaching the trimmed needle and circulating 54 U/mL heparinized PBS perfusate (from step 1.8). Ensure to purge all air bubbles from the tubing to eliminate the possibility of air embolism, which leads to decreased perfusion efficiency or failure (see Table 1). Keep the perfusion media on ice for the duration of the procedure.</li>
	<li>If the pump is not programmable, with the needle in place, measure the pumped volume of media under the different settings to determine which settings are closest to 5 mL/min and 10 mL/min. These flow rates will be used regardless of the species. If the perfusion pump is programmable, calibrate it with the needle in place, following the manufacturer&#39;s instructions.</li>
	<li>Place the dissection surface (tray or foam sheet) at an incline within a secondary container, or 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 recheck the loss of pain response by performing a foot pinch.</li>
	<li>Place the frog on its back and pin down each limb (Figure 2). If preserving the tissue of the limbs is required, thin pins may be placed through the digits or U-shaped staples around the limbs.</li>
	<li>Using dissection scissors, cut through the skin, up the midline, and then laterally, making two flaps. (Figure 2)</li>
	<li>Use forceps to grasp the linea alba and pull it away from the coelomic cavity (Figure 3). Carefully use scissors to cut up through the musculature. Make two flaps out of the cavity wall and cut or pin all the flaps out of the way.</li>
	<li>Use dissection scissors to cut up through the coracoid bones and cut away excess tissue to gain better access to the heart (Figure 3).</li>
	<li>The heart should still be beating. If the heart has stopped beating prior to perfusion, note that the sample freshness has been compromised.</li>
</ol>
2. Perfusion
<ol>
	<li>Identify the stomach and gently shift it so that it is on top of the left lobe of the liver (on the viewer&#39;s right), with its vasculature visible for the duration of the procedure. Identify a lung and grasp it by its tip using tissue forceps. Pull the lung outside of the coelomic cavity and pin it through the tip (Figure 4). Do this gently, as broken blood vessels do not perfuse well. Note if blood is visible within the lobe, as this will affect the ability to determine the completion of the procedure.</li>
	<li>Take an image of the coelomic cavity to better assess perfusion efficiency and potentially identify abnormal tissues at a later date.</li>
	<li>Identify the thin pericardium and pull it taught with tissue forceps (Figure 5). Gently perforate the pericardium using the tip of the iridectomy scissors, being careful not to cut the underlying tissues. Peel the pericardium up away from the three chambers of the heart.</li>
	<li>Use forceps to gently grasp the ventricle by its apex. Apply limited pressure so that there is enough room between the traction surfaces of the forceps for the perfusion needle to pass (Figure 6).</li>
	<li>Insert the needle through the closure of the forceps into the chamber of the ventricle, being careful not to perforate through the ventricle (Figure 7). Clamp the tissue forceps in place using a needle holder by using a hemostat. 
	NOTE: This technique stabilizes the position of the needle, which is still sharp. Clamping the needle directly to the ventricle will also cause unnecessary damage, making reclamping more difficult should it be required (see Table 1).</li>
	<li>Start the flow of the pump at approximately 5 mL/min. The three chambers of the heart and arterial trunk will engorge (Figure 8; see Table 1).</li>
	<li>With scissors, carefully lance the right auricle (on the viewer&#39;s left); blood will pour out. Set the flow rate at 5 mL/min or increase it to 10 mL/min.</li>
	<li>Continue until the vasculature of the stomach blanches (see Table 1), then lance the left auricle of the heart (on the viewer&#39;s right). If the flow rate is still 5 mL/min, increase it to approximately 10 mL/min.</li>
	<li>Use a transfer pipette to rinse the coelomic cavity in perfusion media, to help maintain visibility and to better assess the color of the perfusate flowing from the auricles.</li>
	<li>Keep the needle in place until the perfusate flowing from the auricles is clear (see Table 1) and the lung has lost its red hue (see Table 1; Figure 9).</li>
</ol>

Blood Perfusion in African Clawed Frogs for Transcriptomics and Proteomics

<ol>
	<li>Saltman, A. J., Barakat, M., Bryant, D. M., Brodovskaya, A., Whited, J. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=DiI+perfusion+as+a+method+for+vascular+visualization+in+Ambystoma+mexicanum.">DiI perfusion as a method for vascular visualization in Ambystoma mexicanum.</a> Journal of Visualized Experiments. (124), e55740 (2017).</li><li>Lametschwandtner, A., Minnich, B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Microvascular+anatomy+of+the+brain+of+the+adult+pipid+frog,+Xenopus+laevis+(Daudin):+A+scanning+electron+microscopic+study+of+vascular+corrosion+casts.">Microvascular anatomy of the brain of the adult pipid frog, Xenopus laevis (Daudin): A scanning electron microscopic study of vascular corrosion casts.</a> Journal of Morphology. 279 (7), 950-969 (2018).</li><li>Lametschwandtner, A., Minnich, B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Microvascular+anatomy+of+the+urinary+bladder+in+the+adult+African+clawed+toad,+Xenopus+laevis:+A+scanning+electron+microscope+study+of+vascular+casts.">Microvascular anatomy of the urinary bladder in the adult African clawed toad, Xenopus laevis: A scanning electron microscope study of vascular casts.</a> Journal of Morphology. 282 (3), 368-377 (2021).</li><li>Lametschwandtner, 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=Microvascular+anatomy+of+the+gallbladder+of+the+adult+South+African+clawed+toad,+Xenopus+laevis+Daudin:+A+scanning+electron+microscope+study+of+vascular+corrosion+casts.">Microvascular anatomy of the gallbladder of the adult South African clawed toad, Xenopus laevis Daudin: A scanning electron microscope study of vascular corrosion casts.</a> Microscopy and Microanalysis. 13, 492-493 (2007).</li><li>Lametschwandtner, A., Spornitz, U., Minnich, B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Microvascular+anatomy+of+the+non-lobulated+liver+of+adult+Xenopus+laevis:+A+scanning+electron+microscopic+study+of+vascular+casts.">Microvascular anatomy of the non-lobulated liver of adult Xenopus laevis: A scanning electron microscopic study of vascular casts.</a> Anatomical Record. 305 (2), 243-253 (2022).</li><li>Miodo&#324;ski, A. J., B&#228;r, T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Arterial+supply+of+the+choriocapillaris+of+anuran+amphibians+(Rana+temporaria,+Rana+esculenta).+Scanning+electron-microscopic+(SEM)+study+of+microcorrosion+casts.">Arterial supply of the choriocapillaris of anuran amphibians (Rana temporaria, Rana esculenta). Scanning electron-microscopic (SEM) study of microcorrosion casts.</a> Cell and Tissue Research. 249 (1), 101-109 (1987).</li><li>Nenni, M. 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=Xenbase:+Facilitating+the+use+of+Xenopus+to+model+human+disease.">Xenbase: Facilitating the use of Xenopus to model human disease.</a> Frontiers in Physiology. 10, 154 (2019).</li><li>Tandon, P., Conlon, F., Furlow, J. D., Horb, M. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Expanding+the+genetic+toolkit+in+Xenopus:+Approaches+and+opportunities+for+human+disease+modeling.">Expanding the genetic toolkit in Xenopus: Approaches and opportunities for human disease modeling.</a> Developmental Biology. 426 (2), 325-335 (2017).</li><li>Peshkin, L., 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=The+protein+repertoire+in+early+vertebrate+embryogenesis.">The protein repertoire in early vertebrate embryogenesis.</a> bioRxiv. , (2019).</li><li>Briggs, J. 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=The+dynamics+of+gene+expression+in+vertebrate+embryogenesis+at+single-cell+resolution.">The dynamics of gene expression in vertebrate embryogenesis at single-cell resolution.</a> Science. 360 (6392), (2018).</li><li>Liao, Y., 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=Cell+landscape+of+larval+and+adult+Xenopus+laevis+at+single-cell+resolution.">Cell landscape of larval and adult Xenopus laevis at single-cell resolution.</a> Nature Communications. 13 (1), 4306 (2022).</li><li>AVMA (American Veterinary Medical Association). <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=AVMA+guidelines+for+the+euthanasia+of+animals,+2020+edition.">AVMA guidelines for the euthanasia of animals, 2020 edition.</a> AVMA. , 37 (2020).</li><li>Navarro, K., Jampachaisri, K., Chu, D., Pacharinsak, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Bupivacaine+as+a+euthanasia+agent+for+African+Clawed+Frogs+(Xenopus+laevis).">Bupivacaine as a euthanasia agent for African Clawed Frogs (Xenopus laevis).</a> PLoS One. 17 (12), e0279331 (2022).</li><li>Wu, 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=Transcardiac+perfusion+of+the+mouse+for+brain+tissue+dissection+and+fixation.">Transcardiac perfusion of the mouse for brain tissue dissection and fixation.</a> Bio-Protocol. 11 (5), e3988 (2021).</li><li>Heinz-Taheny, K. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cardiovascular+physiology+and+diseases+of+amphibians.+Veterinary+clinics+of+North+America.">Cardiovascular physiology and diseases of amphibians. Veterinary clinics of North America.</a> The Veterinary Clinics of North America. Exotic Animal Practice. 12 (1), 39-50 (2009).</li><li>Stephenson, A., Adams, J. W., Vaccarezza, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+vertebrate+heart:+an+evolutionary+perspective.">The vertebrate heart: an evolutionary perspective.</a> Journal of Anatomy. 231 (6), 787-797 (2017).</li><li>Hoops, D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+perfusion+protocol+for+lizards,+including+a+method+for+brain+removal.">A perfusion protocol for lizards, including a method for brain removal.</a> MethodsX. 2, 165-173 (2015).</li></ol>

The authors declare no competing interests.

This protocol describes traditional dissection techniques for accessing the coelomic cavity. Other techniques are also acceptable, provided they cause minimal damage to the tissues, the heart is accessible, and the lung and stomach are visible. Similarly, most dissection tools listed can be easily substituted with comparable items.
While attempts have been made to optimize the efficacy of this procedure, results may vary depending on one's experience and variability between individual frogs. One interesting aspect of blood perfusion that remained outside of the scope of this paper is how this procedure compares to alternative ways of perfusion for animals that undergo surgery. Another unexplored variable is how blood perfusion would work in very young animals or animals of advanced age where vasculature might be excessively fragile. Additional remarks are provided to facilitate the application of this protocol. Several typical problems, their possible causes, and suggested remedial actions are provided in Table 1.
A limitation of this procedure is that the perfusion efficiency can be negatively impacted by its speed. If perfusion efficiency takes precedence over rapid perfusion, adapting an axolotl technique is recommended1 (Saltman et al. use the term aorta to refer to the arterial trunk). 
The duration of the procedure and volume of media used are dependent on a number of variables. Generally, X. tropicalis males take between 2-3 min to successfully perfuse with 15-25 mL of media, while X. tropicalis females take between 3-4 min with 25-40 mL of media. Significantly more animal-to-animal variation was found when perfusing X. laevis. Though a higher flow rate would lessen the length of time required to perfuse larger animals, the increased line pressure can easily lead to the tube fittings becoming dislodged and pump failure.
Naturally, it is much easier to assess perfusion efficiency in albino animals. The difference is especially apparent in the lung and liver tissues (Figure 12). Thus, the use of albinos is recommended, especially when first attempting perfusion or undergoing training. 
By adjusting the flow rate and needle size, the protocol is adaptable for all species of Xenopus. Owing to the homology in heart anatomy and blood circulation between Xenopus and most other amphibians15, as well as non-crocodilian reptiles, this technique may be modified for full-body rapid perfusion of other models with three-chambered hearts16. If a non-crocodilian reptile model is used that exclusively requires the perfusion of one of the aortic arches, other protocols are recommended17.

The whole body perfusion of amphibians is routinely completed for the purposes of preservation and fixation1,2,3,4,5,6. However, these procedures occur at a rate that limits the number of fresh samples that can be taken per animal. The goal of this work is to develop an effective blood perfusion protocol in Xenopus, prioritizing the speed of the technique. The protocol takes less than 10 min per animal for X. tropicalis and less than 15 min per X. laevis animal. The secondary priorities are the ease of replication and the use of easily acquired equipment so that high-quality samples can be shared widely between Xenopus labs.
Xenopus frogs are used widely in biomedical research to study fundamental biological and pathological processes conserved across species. This tetrapod has a closer evolutionary relationship with mammals than other aquatic models, having lungs, a three-chambered heart, and limbs with digits. The international community effectively uses Xenopus to gain a deeper understanding of human disease through in-depth disease modeling and molecular analysis of disease-related gene function. The numerous advantages of Xenopus as an animal model make them invaluable tools to study the molecular basis of human development and disease; these advantages include: large oocyte and embryo size, high fecundity, ease of housing, rapid external development, and ease of genomic manipulation. Xenopus have been estimated to share ~80% of the identified human disease genes7.
Compared to popular mammalian models, Xenopus is a rapid, cost-effective model, with the ease of morpholino knock-down and availability of efficient transgenics and targeted gene mutations using CRISPR8. Quantitative mass spectrometry and single-cell transcriptomics have been successfully applied to Xenopus embryos9,10,&#160;but a recent cell atlas from Xenopus laevis shows that the composition of most tissues is dominated by blood cell types11. By developing a technique that exsanguinates tissue at a rapid rate and using chilled media, the sample freshness is minimally affected by perfusion. This is particularly important for applications where the goal is to profile physiologically unperturbed mRNA or protein expression.

<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>&lt;em&gt;414004-036&lt;/em&gt;</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</td><td>Item&amp;nbsp;2860</td><td>alternative opaque containers acceptable</td></tr><tr><td>Euthanasia container lid</td><td>US Plastic</td><td>Item&amp;nbsp;3047</td><td></td></tr><tr><td>Fine dissection pins</td><td>Living Systems Instrumentation</td><td>PIN-#3</td><td></td></tr><tr><td>General use hypodermic needles, 22 G</td><td>Fisher Scientific</td><td>14-826-5A</td><td>for &lt;em&gt;X. laevis&lt;/em&gt;</td></tr><tr><td>General use hypodermic needles, 25 G</td><td>Fisher Scientific</td><td>14-826AA</td><td>for &lt;em&gt;X. tropicalis&lt;/em&gt;</td></tr><tr><td>Heparin, porcine intestinal mucosa</td><td>MilliporeSigma</td><td>37-505-410MG</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>Luer-to-barb adapter male Luer with lock ring</td><td>amazon.com</td><td>B09PTX6M2Z</td><td>size will be dependant on the hosing of the pump used</td></tr><tr><td>Mayo-Hegar needle holder</td><td>Fisher Scinetific</td><td>08-966</td><td>mosquito forceps 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>Peristaltic liquid pump dosing pump 5&amp;ndash;100 mL/min</td><td>amazon.com</td><td>B07PWY4SM6</td><td>any peristaltic pump capable of pumping 5-10mL/min is acceptable</td></tr><tr><td>Sharpening stone</td><td>VWR</td><td>470150-112</td><td>optional; for dulling needles</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><tr><td>Ultra-fine short insulin syringes, 31 G</td><td>VWR</td><td>BD328438</td><td></td></tr><tr><td>Wire flush cutters, 6-inch ultra sharp &amp;amp; powerful side cutter clippers</td><td>amazon.com</td><td>B087P191LP</td><td></td></tr></tbody></table>

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.

Following successful perfusion, all tissues (excluding the liver in pigmented Xenopus) will be distinctly lighter and less saturated with blood. Major blood vessels will become less noticeable (Figure 10), and tissues (excluding the liver) will rinse cleanly in the buffer after being sampled. While the successful execution of the protocol can ultimately only be confirmed by the quality of the data from exsanguinated tissue samples, several typical problems, their possible causes, and suggested remedial actions are provided in the troubleshooting table (Table 1 and Figure 11).
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/65287/65287fig01.jpg" /> 
Figure 1: Untrimmed and trimmed needles14. Using wire clippers, blunt the needle by cutting off its tip. It will be sharp enough to pierce the heart, yet perforating the ventricle will be less likely in the event of human error. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/65287/65287fig01large.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/65287/65287fig02.jpg" /> 
Figure 2: Mature female X. tropicalis pinned through each limb. Use toothed dissecting forceps to pull the skin near the cloaca taught to perforate it with dissection scissors and create two flaps. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/65287/65287fig02large.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/65287/65287fig03.jpg" /> 
Figure 3: Muscular wall. With the ventral skin open but the muscular wall intact, the linea alba is visible. To reduce the likelihood of damaging the underlying tissues, grasp the linea alba and pull it taught prior to cutting. The coracoid bones are visible through the peritoneum. Once the coelomic cavity has been opened, these bones should be reduced to give better access to the heart. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/65287/65287fig03large.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/65287/65287fig04.jpg" /> 
Figure 4: The coelomic cavity of a mature X. tropicalis male. The coracoid bones have been reduced, providing access to the pericardium-enclosed heart. The stomach has been shifted in front of the left lobe of the liver, and its vasculature is clearly visible. The left lung has been pulled out of the coelomic cavity by its tip and pinned to ensure that it does not retract during the rinsing process. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/65287/65287fig04large.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/65287/65287fig05.jpg" /> 
Figure 5: Pericardium. The pericardium is a thin, tough membrane enclosing the heart. Using tissue forceps, gently grasp the pericardium and then use the tip of iridectomy scissors to perforate it. Once perforated, peel it up, away from the heart. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/65287/65287fig05large.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/65287/65287fig06.jpg" /> 
Figure 6: Heart anatomy needle placement diagrams. (A) Ventral diagram of an X. laevis heart. (B) Heart diagram, with the pericardium removed, showing the correct needle and clamp placement. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/65287/65287fig06large.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/65287/65287fig07.jpg" /> 
Figure 7: Heart anatomy and needle placement photograph. With the pericardium removed, the three chambers of the heart and arterial trunk are easily visible. Use forceps to gently grasp the ventricle by its apex and then insert the needle through the forceps. Be careful not to cause unnecessary damage to the ventricle or other chambers as this will compromise the perfusion efficiency. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/65287/65287fig07large.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/65287/65287fig08.jpg" /> 
Figure 8: Perfusion is underway. The right auricle has been lanced, and the ventricle, arterial trunk, and left auricle are visibly engorged. The stomach is blanching and both the media running from the animal and the lung tissue are heavily saturated with blood. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/65287/65287fig08large.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/65287/65287fig09.jpg" /> 
Figure 9: The coelomic cavity following successful rapid perfusion and rinsing. The vasculature of the stomach and other organs is no longer easily visible. Unless the Xenopus is albino, the liver will remain heavily pigmented. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/65287/65287fig09large.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/65287/65287fig10.jpg" /> 
Figure 10: Tissues samples from an unperfused and perfused albino male X. laevis. The differences in pigmentation and visibility of the vasculature are pronounced. All samples are within 3.5 cm diameter wells. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/65287/65287fig10large.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/65287/65287fig11.jpg" /> 
Figure 11: Troubleshooting diagrams of a Xenopus heart. (A) The ventricle has perforation (in red); this perforation is isolated by the forceps and will not affect the perfusion efficiency. (B) A heart with a severely damaged ventricle. The needle can be guided into the arterial trunk and clamped in place. It is especially important to ensure the needle is well-blunted when using this technique14. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/65287/65287fig11large.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/65287/65287fig12.jpg" /> 
Figure 12: Assessing perfusion efficiency in albinos. An albino X. laevis both before (A) and after (B) rapid perfusion. The albinism makes it easier to determine the proficiency of the perfusion than it would be in a pigmented animal. This is especially apparent in the lung and liver tissues. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/65287/65287fig12large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Table 1: Troubleshooting table. Several typical problems, their possible causes, and suggested remedial actions are provided. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/65287/Table 1.xlsx" target="_blank">Please click here to download this Table.</a>

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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.

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 ...

effective-rapid-blood-perfusion-in-xenopus

Nanyang Technological University

Research

JoVE Journal

Biology

4.5K Views.  Harvard Medical School. Deze procedure draait om het humane gebruik van amfibieënmodellen op een manier die de efficiëntie maximaliseert. Standaardisatiepraktijken kunnen worden toegepast op monsters die zijn genomen en bewaard in de kolonie van oorsprong, waardoor bioveiligheidsrisico's worden beperkt en stress bij het transport van dieren wordt geëlimineerd. Profusie kan snel worden voltooid, waardoor de beschikbare tijd om weefsels te bemonsteren terwijl ze nog vers zijn, wordt gemaximaliseerd.Deze fris...

Video: Auteur in de schijnwerpers: effectieve snelle bloedperfusie bij Xenopus  

Comparative in vivo Study of gp96 Adjuvanticity in the Frog Xenopus laevis

We have developed in the amphibian Xenopus laevis a unique non-mammalian model to study the ability of certain heat shock proteins (hsps) such as gp96 to facilitate cross-presentation of chaperoned antigens and elicit innate and adaptive T cell responses. Xenopus skin graft rejection provides an excellent platform to study the ability of gp96 to elicit classical MHC class Ia (class Ia) restricted T cell responses. Additionally, the Xenopus model system also provides an attractive alternative to mice for exploring the ability of gp96 to generate responses against tumors that have down-regulated their class Ia molecules thereby escaping immune surveillance. Recently, we have developed an adoptive cell transfer assay in Xenopus clones using peritoneal leukocytes as antigen presenting cells (APCs), and shown that gp96 can prime CD8 T cell responses in vivo against minor histocompatibility skin antigens as well as against the Xenopus thymic tumor 15/0 that does not express class Ia molecules. We describe here the methodology involved to perform these assays including the elicitation, pulsing and adoptive transfer of peritoneal leukocytes, as well as the skin graft and tumor transplantation assays. Additionally we are also describing the harvesting and separation of peripheral blood leukocytes used for flow cytometry and proliferation assays which allow for further characterization of the effector populations involved in skin rejection and anti-tumor responses.

Comparative in vivo Study of gp96 Adjuvanticity in the Frog Xenopus laevis

The frog Xenopus laevis provides an attractive alternative non-mammalian model for exploring the ability of heat shock protein such as gp96 to promote antigen-specific CD8 T cell responses. We present methods to study in vivo facilitation of cross-presentation of skin and tumor antigens by gp96.

We have developed in the amphibian Xenopus laevis a unique non-mammalian model to study the ability of certain heat shock proteins (hsps) such as gp96 to ...

Immunology and Infection

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 ...

Developmental Biology

Ex Vivo Perfusion of the Rodent Placenta

The placenta is a key organ during pregnancy that serves as a barrier to fetal xenobiotic exposure and mediates the exchange of nutrients for waste. An assay is described here to perfuse an isolated rat placenta and evaluate the maternal-to-fetal translocation of xenobiotics ex vivo. In addition, the evaluation of physiological processes such as fluid flow to the fetus and placental metabolism may be conducted with this methodology. This technique is suitable for evaluating maternal-to-fetal kinetics of pharmaceutical candidates or environmental contaminants. In contrast to current alternative approaches, this methodology allows the evaluation of the isolated maternal-fetal vasculature, with the systemic neural or immune involvement removed, allowing any observed changes in physiological function to be attributable to local factors within the isolated tissue.

Here is presented a protocol of ex vivo maternal-fetal vascular perfusion to enable the administration of a test article into maternal vasculature and to evaluate placental transfer of xenobiotic particles or pharmacological agents in addition to alterations in placental physiology.

The placenta is a key organ during pregnancy that serves as a barrier to fetal xenobiotic exposure and mediates the exchange of nutrients for waste. An ...

Patch Clamp and Perfusion Techniques for Studying Ion Channels Expressed in Xenopus oocytes

The protocol presented here is designed to study the activation of the large conductance, voltage- and Ca2+-activated K+ (BK) channels. The protocol may also be used to study the structure-function relationship for other ion channels and neurotransmitter receptors1. BK channels are widely expressed in different tissues and have been implicated in many physiological functions, including regulation of smooth muscle contraction, frequency tuning of inner hair cells and regulation of neurotransmitter release2-6. BK channels are activated by membrane depolarization and by intracellular Ca2+ and Mg2+ 6-9. Therefore, the protocol is designed to control both the membrane voltage and the intracellular solution. In this protocol, messenger RNA of BK channels is injected into Xenopus laevis oocytes (stage V-VI) followed by 2-5 days of incubation at 18&deg;C10-13. Membrane patches that contain single or multiple BK channels are excised with the inside-out configuration using patch clamp techniques10-13. The intracellular side of the patch is perfused with desired solutions during recording so that the channel activation under different conditions can be examined. To summarize, the mRNA of BK channels is injected into Xenopus laevis oocytes to express channel proteins on the oocyte membrane; patch clamp techniques are used to record currents flowing through the channels under controlled voltage and intracellular solutions.

Patch Clamp and Perfusion Techniques for Studying Ion Channels Expressed in Xenopus oocytes

Ionic current of BK channels is recorded using patch clamp techniques. BK channels are expressed in Xenopus oocytes by injecting messenger RNA. The intracellular solution during patch clamp recordings is controlled by a perfusion system.

The protocol presented here is designed to study the activation of the large conductance, voltage- and Ca2+-activated K+ (BK) channels. The protocol may ...

An In Vivo Blood-brain Barrier Permeability Assay in Mice Using Fluorescently Labeled Tracers

Blood-brain barrier (BBB) is a specialized barrier that protects the brain microenvironment from toxins and pathogens in the circulation and maintains brain homeostasis. The principal sites of the barrier are endothelial cells of the brain capillaries whose barrier function results from tight intercellular junctions and efflux transporters expressed on the plasma membrane. This function is regulated by pericytes and astrocytes that together form the neurovascular unit (NVU). Several neurological diseases such as stroke, Alzheimer&#39;s disease (AD), brain tumors are associated with an impaired BBB function. Assessment of the BBB permeability is therefore crucial in evaluating the severity of the neurological disease and the success of the treatment strategies employed.
We present here a simple yet robust permeability assay that have been successfully applied to several mouse models both, genetic and experimental. The method is highly quantitative and objective in comparison to the tracer fluorescence analysis by microscopy that is commonly applied. In this method, mice are injected intraperitoneally with a mix of aqueous inert fluorescent tracers followed by anesthetizing the mice. Cardiac perfusion of the animals is performed prior to harvesting brain, kidneys or other organs. Organs are homogenized and centrifuged followed by fluorescence measurement from the supernatant. Blood drawn from the cardiac puncture just before perfusion serves for normalization purpose to the vascular compartment. The tissue fluorescence is normalized to the wet weight and serum fluorescence to obtain a quantitative tracer permeability index. For additional confirmation, the contralateral hemi-brain preserved for immunohistochemistry can be utilized for tracer fluorescence visualization purposes.

An In Vivo Blood-brain Barrier Permeability Assay in Mice Using Fluorescently Labeled Tracers

Here we present a mouse brain vascular permeability assay using intraperitoneal injection of fluorescent tracers followed by perfusion that is applicable to animal models of blood-brain barrier dysfunction. One hemi-brain is used for assessing permeability quantitatively and the other for tracer visualization/immunostaining. The procedure takes 5 - 6 h for 10 mice.

Blood-brain barrier (BBB) is a specialized barrier that protects the brain microenvironment from toxins and pathogens in the circulation and maintains ...

Neuroscience

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 ...

Improvement of a Closed Chest Porcine Myocardial Infarction Model by Standardization of Tissue and Blood Sampling Procedures

Myocardial ischemia reperfusion (I/R) injury contributes to almost half of the necrotic area after myocardial infarction. To date there is no approved drug to prevent or reduce myocardial I/R injury. The study and understanding of the pathophysiological mechanisms of myocardial I/R injury is essential to develop successful treatments. Large animal experiments are an important step in translational methods. The porcine model of acute myocardial infarction has been established and described by ourselves and others. We aimed to further improve the value of the model by focusing in detail on the sampling techniques for use in future experiments. Furthermore, we emphasize small but important steps that can affect the quality of the final results. To mimic the clinical situation of myocardial I/R injury, a percutaneous coronary intervention (PCI) catheter was inserted into the left anterior descending coronary artery (LAD) of an anesthetized pig. &#176;&#176;&#176;This model mimics acute myocardial infarction and PCI treatment in humans with the possibility of accurately determining the area at risk as well as the necrotic- and viable ischemic tissue. Here the model was used to investigate the effect of a bicyclic peptide inhibitor of FXIIa. The model can also be modified to allow longer reperfusion times to study later effects of myocardial infarction.

Here we demonstrate a protocol to standardize sampling procedures of an established porcine model of acute myocardial infarction in order to increase its translational value in the understanding of the pathophysiology of myocardial ischemia/reperfusion injury and to test novel drug candidates.

Myocardial ischemia reperfusion (I/R) injury contributes to almost half of the necrotic area after myocardial infarction. To date there is no approved ...

Medicine

Establishing In Situ Closed Circuit Perfusion of Lower Abdominal Organs and Hind Limbs in Mice

Ex vivo perfusion is an important physiological tool to study the function of isolated organs (e.g. liver, kidneys). At the same time, due to the small size of mouse organs, ex vivo perfusion of bone, bladder, skin, prostate, and reproductive organs is challenging or not feasible. Here, we report for the first time an in situ lower body perfusion circuit in mice that includes the above tissues, but bypasses the main clearance organs (kidney, liver, and spleen). The circuit is established by cannulating the abdominal aorta and inferior vena cava above the iliac artery and vein and cauterizing peripheral blood vessels. Perfusion is performed via a peristaltic pump with perfusate flow maintained for up to 2 h. In situ staining with fluorescent lectin and Hoechst solution confirmed that the microvasculature was successfully perfused. This mouse model can be a very useful tool for studying pathological processes as well as mechanisms of drug delivery, migration/metastasis of circulating tumor cells into/from the tumor, and interactions of immune system with perfused organs and tissues.

A protocol is described for in situ perfusion of the mouse lower body, including the bladder, the prostate, sex organs, bone, muscle and foot skin.

Ex vivo perfusion is an important physiological tool to study the function of isolated organs (e.g. liver, kidneys). At the same time, due to the small ...

Development of a Selective Aortic Arch Perfusion System in a Porcine Model of Exsanguination Cardiac Arrest

Hemorrhage constitutes the majority of potentially preventable deaths from trauma. There is growing interest in endovascular resuscitation techniques such as selective aortic arch perfusion (SAAP) for patients in cardiac arrest. This involves active perfusion of the coronary circulation via a thoracic aortic balloon catheter and is approaching clinical application. However, the technique is complex and requires refinement in animal models before human use can be considered. This paper describes a large animal model of exsanguination cardiac arrest treated with a bespoke SAAP system.
Swine were anesthetized, instrumented and a splenectomy was performed before a controlled, logarithmic exsanguination was initiated. Animals were heparinized and the shed blood collected in a reservoir. Once cardiac arrest was observed, the blood was pumped through an extra-corporeal circuit into an oxygenator and then delivered through a 10 Fr balloon catheter placed in the thoracic aorta.
This resulted in the return of a spontaneous circulation (ROSC) as demonstrated by ECG and aortic root pressure waveform. This model and accompanying SAAP system allow for standardized and reproducible recovery from exsanguination cardiac arrest.

The goal of this protocol is to demonstrate a porcine exsanguination cardiac arrest model and a specifically built selective aortic arch perfusion circuit for translational research.

Hemorrhage constitutes the majority of potentially preventable deaths from trauma. There is growing interest in endovascular resuscitation techniques such ...

DiI Perfusion as a Method for Vascular Visualization in Ambystoma mexicanum

Perfusion techniques have been used for centuries to visualize the circulation of tissues. Axolotl (Ambystoma mexicanum) is a species of salamander that has emerged as an essential model for regeneration studies. Little is known about how revascularization occurs in the context of regeneration in these animals. Here we report a simple method for visualization of the vasculature in axolotl via perfusion of 1,1&#8217;-Dioctadecy-3,3,3&#8217;,3&#8217;-tetramethylindocarbocyanine perchlorate (DiI). DiI is a lipophilic carbocyanine dye that inserts into the plasma membrane of endothelial cells instantaneously. Perfusion is done using a peristaltic pump such that DiI enters the circulation through the aorta. During perfusion, dye flows through the axolotl&#8217;s blood vessels and incorporates into the lipid bilayer of vascular endothelial cells upon contact. The perfusion procedure takes approximately one hour for an eight-inch axolotl. Immediately after perfusion with DiI, the axolotl can be visualized with a confocal fluorescent microscope. The DiI emits light in the red-orange range when excited with a green fluorescent filter. This DiI perfusion procedure can be used to visualize the vascular structure of axolotls or to demonstrate patterns of revascularization in regenerating tissues.

DiI Perfusion as a Method for Vascular Visualization in Ambystoma mexicanum

Using a lipophilic 1,1&#39;-Dioctadecy-3,3,3&#39;,3&#39;-tetramethylindocarbocyanine perchlorate (DiI) staining technique, Ambystoma mexicanum can undergo vascular perfusion to allow for easy visualization of the vasculature.

Perfusion techniques have been used for centuries to visualize the circulation of tissues. Axolotl (Ambystoma mexicanum) is a species of salamander that ...

Effective Rapid Blood Perfusion in Xenopus

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