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. O...

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 border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<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&#34;</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&#160;2860</td>
			<td>alternative opaque containers acceptable</td>
		</tr>
		<tr>
			<td>Euthanasia container lid</td>
			<td>US Plastic</td>
			<td>Item&#160;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 X. laevis</td>
		</tr>
		<tr>
			<td>General use hypodermic needles, 25 G</td>
			<td>Fisher Scientific</td>
			<td>14-826AA</td>
			<td>for X. tropicalis</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&#34;</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&#8211;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 &#38; 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, an...

Watch this Scientific Journal Video about Effective Rapid Blood Perfusion in Xenopus at JoVE.com

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 ...

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4.2K Views.  Harvard Medical School. Presented here is an effective rapid blood perfusion protocol to prepare tissue samples from African clawed frogs for transcriptomics and proteomics studies. 

Video: Author Spotlight: Effective Rapid Blood Perfusion in Xenopus  

Obtaining Eggs from Xenopus laevis Females

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

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

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

In Vitro Nuclear Assembly Using Fractionated Xenopus Egg Extracts

Nuclear membrane assembly is an essential step in the cell division cycle; this process can be replicated in the test tube by combining Xenopus sperm chromatin, cytosol, and light membrane fractions. Complete nuclei are formed, including nuclear membranes with pore complexes, and these reconstituted nuclei are capable of normal nuclear processes.

Nuclear membrane assembly is an essential step in the cell division cycle; this process can be replicated in the test tube by combining Xenopus sperm chromatin, cytosol, and light membrane fractions. Complete nuclei are formed, including nuclear membranes with pore complexes, and these reconstituted nuclei are capable of normal nuclear processes.

Nuclear membrane assembly is an essential step in the cell division cycle; this process can be replicated in the test tube by combining Xenopus sperm ...

Microinjection of Xenopus Laevis Oocytes

Microinjection of Xenopus laevis oocytes followed by thin-sectioning electron microscopy (EM) is an excellent system for studying nucleocytoplasmic transport. Because of its large nucleus and high density of nuclear pore complexes (NPCs), nuclear transport can be easily visualized in the Xenopus oocyte. Much insight into the mechanisms of nuclear import and export has been gained through use of this system (reviewed by Pant&eacute;, 2006). In addition, we have used microinjection of Xenopus oocytes to dissect the nuclear import pathways of several viruses that replicate in the host nucleus. 
Here we demonstrate the cytoplasmic microinjection of Xenopus oocytes with a nuclear import substrate. We also show preparation of the injected oocytes for visualization by thin-sectioning EM, including dissection, dehydration, and embedding of the oocytes into an epoxy embedding resin. Finally, we provide representative results for oocytes that have been microinjected with the capsid of the baculovirus Autographa californica nucleopolyhedrovirus (AcMNPV) or the parvovirus Minute Virus of Mice (MVM), and discuss potential applications of the technique.

Here we demonstrate cytoplasmic microinjection of Xenopus laevis oocytes with a nuclear import substrate, as well as preparation of the injected oocytes for visualization by thin-sectioning electron microscopy.

Microinjection of Xenopus laevis oocytes followed by thin-sectioning electron microscopy (EM) is an excellent system for studying nucleocytoplasmic ...

Visualizing RNA Localization in Xenopus Oocytes

RNA localization is a conserved mechanism of establishing cell polarity. Vg1 mRNA localizes to the vegetal pole of Xenopus laevis oocytes and acts to spatially restrict gene expression of Vg1 protein. Tight control of Vg1 distribution in this manner is required for proper germ layer specification in the developing embryo. RNA sequence elements in the 3' UTR of the mRNA, the Vg1 localization element (VLE) are required and sufficient to direct transport. To study the recognition and transport of Vg1 mRNA in vivo, we have developed an imaging technique that allows extensive analysis of trans-factor directed transport mechanisms via a simple visual readout. 
To visualize RNA localization, we synthesize fluorescently labeled VLE RNA and microinject this transcript into individual oocytes. After oocyte culture to allow transport of the injected RNA, oocytes are fixed and dehydrated prior to imaging by confocal microscopy. Visualization of mRNA localization patterns provides a readout for monitoring the complete pathway of RNA transport and for identifying roles in directing RNA transport for cis-acting elements within the transcript and trans-acting factors that bind to the VLE (Lewis et al., 2008, Messitt et al., 2008). We have extended this technique through co-localization with additional RNAs and proteins (Gagnon and Mowry, 2009, Messitt et al., 2008), and in combination with disruption of motor proteins and the cytoskeleton (Messitt et al., 2008) to probe mechanisms underlying mRNA localization.

Visualizing RNA Localization in Xenopus Oocytes

Visualization of in vivo RNA transport is accomplished by microinjection of fluorescently labeled RNA transcripts into Xenopus oocytes, followed by confocal microscopy.

RNA localization is a conserved mechanism of establishing cell polarity.  Vg1 mRNA localizes to the vegetal pole of Xenopus laevis oocytes and acts to ...

Rapid Homogeneous Detection of Biological Assays Using Magnetic Modulation Biosensing System

A magnetic modulation biosensing system (MMB) [1,2] rapidly and homogeneously detected biological targets at low concentrations without any washing or separation step. When the IL-8 target was present, a 'sandwich'-based assay attached magnetic beads with IL-8 capture antibody to streptavidin coupled fluorescent protein via the IL-8 target and a biotinylated IL-8 antibody. The magnetic beads are maneuvered into oscillatory motion by applying an alternating magnetic field gradient through two electromagnetic poles. The fluorescent proteins, which are attached to the magnetic beads are condensed into the detection area and their movement in and out of an orthogonal laser beam produces a periodic fluorescent signal that is demodulated using synchronous detection. The magnetic modulation biosensing system was previously used to detect the coding sequences of the non-structural Ibaraki virus protein 3 (NS3) complementary DNA (cDNA) [2]. The techniques that are demonstrated in this work for external manipulation and condensation of particles may be used for other applications, e.g. delivery of magnetically-coupled drugs in-vivo or enhancing the contrast for in-vivo imaging applications.

Magnetic modulation biosensing system is utilized to rapidly, sensitively and simply detect biological assays, such as DNA molecules and proteins.

A magnetic modulation biosensing system (MMB) [1,2] rapidly and homogeneously detected biological targets at low concentrations without any washing or ...

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 ...

Isolation and Culture of Pulmonary Endothelial Cells from Neonatal Mice

Endothelial cells provide a useful research model in many areas of vascular biology. Since its first isolation 1, human umbilical vein endothelial cells (HUVECs) have shown to be convenient, easy to obtain and culture, and thus are the most widely studied endothelial cells. However, for research focused on processes like angiogenesis, permeability or many others, microvascular endothelial cells (ECs) are a much more physiologically relevant model to study 2. Furthermore, ECs isolated from knockout mice provide a useful tool for analysis of protein function ex vivo. Several approaches to isolate and culture microvascular ECs of different origin have been reported to date 3-7, but consistent isolation and culture of pure ECs is still a major technical problem in many laboratories. Here, we provide a step-by-step protocol on a reliable and relatively simple method of isolating and culturing mouse lung endothelial cells (MLECs). In this approach, lung tissue obtained from 6- to 8-day old pups is first cut into pieces, digested with collagenase/dispase (C/D) solution and dispersed mechanically into single-cell suspension. MLECS are purified from cell suspension using positive selection with anti-PECAM-1 antibody conjugated to Dynabeads using a Magnetic Particle Concentrator (MPC). Such purified cells are cultured on gelatin-coated tissue culture (TC) dishes until they become confluent. At that point, cells are further purified using Dynabeads coupled to anti-ICAM-2 antibody. MLECs obtained with this protocol exhibit a cobblestone phenotype, as visualized by phase-contrast light microscopy, and their endothelial phenotype has been confirmed using FACS analysis with anti-VE-cadherin 8 and anti-VEGFR2 9 antibodies and immunofluorescent staining of VE-cadherin. In our hands, this two-step isolation procedure consistently and reliably yields a pure population of MLECs, which can be further cultured. This method will enable researchers to take advantage of the growing number of knockout and transgenic mice to directly correlate in vivo studies with results of in vitro experiments performed on isolated MLECs and thus help to reveal molecular mechanisms of vascular phenotypes observed in vivo.

Here, we describe a protocol for isolation and culture of murine pulmonary endothelial cells. This method comprises mechanic and enzymatic lung tissue dissociation as well as a 2-step purification process using anti-PECAM-1 and anti-ICAM-2 antibodies conjugated to magnetic beads, which produces a pure endothelial cell population of mostly microvascular origin.

Endothelial cells provide a useful research model in many areas of vascular biology. Since its first isolation 1, human umbilical vein endothelial cells ...

Rapid Genetic Analysis of Epithelial-Mesenchymal Signaling During Hair Regeneration

Hair follicle morphogenesis, a complex process requiring interaction between epithelia-derived keratinocytes and the underlying mesenchyme, is an attractive model system to study organ development and tissue-specific signaling. Although hair follicle development is genetically tractable, fast and reproducible analysis of factors essential for this process remains a challenge. Here we describe a procedure to generate targeted overexpression or shRNA-mediated knockdown of factors using lentivirus in a tissue-specific manner. Using a modified version of a hair regeneration model 5, 6, 11, we can achieve robust gain- or loss-of-function analysis in primary mouse keratinocytes or dermal cells to facilitate study of epithelial-mesenchymal signaling pathways that lead to hair follicle morphogenesis. We describe how to isolate fresh primary mouse keratinocytes and dermal cells, which contain dermal papilla cells and their precursors, deliver lentivirus containing either shRNA or cDNA to one of the cell populations, and combine the cells to generate fully formed hair follicles on the backs of nude mice. This approach allows analysis of tissue-specific factors required to generate hair follicles within three weeks and provides a fast and convenient companion to existing genetic models.

Tissue-specific analysis of a hair follicle regeneration model using lentivirus to mediate gain- or loss-of-function.

Hair follicle morphogenesis, a complex process requiring interaction between epithelia-derived keratinocytes and the underlying mesenchyme, is an ...

Facial Transplants in Xenopus laevis Embryos

Craniofacial birth defects occur in 1 out of every 700 live births, but etiology is rarely known due to limited understanding of craniofacial development. To identify where signaling pathways and tissues act during patterning of the developing face, a 'face transplant' technique has been developed in embryos of the frog Xenopus laevis. A region of presumptive facial tissue (the "Extreme Anterior Domain" (EAD)) is removed from a donor embryo at tailbud stage, and transplanted to a host embryo of the same stage, from which the equivalent region has been removed. This can be used to generate a chimeric face where the host or donor tissue has a loss or gain of function in a gene, and/or includes a lineage label. After healing, the outcome of development is monitored, and indicates roles of the signaling pathway within the donor or surrounding host tissues. Xenopus is a valuable model for face development, as the facial region is large and readily accessible for micromanipulation. Many embryos can be assayed, over a short time period since development occurs rapidly. Findings in the frog are relevant to human development, since craniofacial processes appear conserved between Xenopus and mammals.

Facial Transplants in Xenopus laevis Embryos

A technique for transplanting "Extreme Anterior Domain" facial tissue between Xenopus laevis embryos has been developed. Tissue can be moved from one gene expression background into another, allowing the study of local requirements for craniofacial development and for signaling interactions between facial regions.

Craniofacial birth defects occur in 1 out of every 700 live births, but etiology is rarely known due to limited understanding of craniofacial development. ...

The Xenopus Oocyte Cut-open Vaseline Gap Voltage-clamp Technique With Fluorometry

The cut-open oocyte Vaseline gap (COVG) voltage clamp technique allows for analysis of electrophysiological and kinetic properties of heterologous ion channels in oocytes. Recordings from the cut-open setup are particularly useful for resolving low magnitude gating currents, rapid ionic current activation, and deactivation. The main benefits over the two-electrode voltage clamp (TEVC) technique include increased clamp speed, improved signal-to-noise ratio, and the ability to modulate the intracellular and extracellular milieu.
Here, we employ the human cardiac sodium channel (hNaV1.5), expressed in Xenopus oocytes, to demonstrate the cut-open setup and protocol as well as modifications that are required to add voltage clamp fluorometry capability.
The properties of fast activating ion channels, such as hNaV1.5, cannot be fully resolved near room temperature using TEVC, in which the entirety of the oocyte membrane is clamped, making voltage control difficult. However, in the cut-open technique, isolation of only a small portion of the cell membrane allows for the rapid clamping required to accurately record fast kinetics while preventing channel run-down associated with patch clamp techniques.
In conjunction with the COVG technique, ion channel kinetics and electrophysiological properties can be further assayed by using voltage clamp fluorometry, where protein motion is tracked via cysteine conjugation of extracellularly applied fluorophores, insertion of genetically encoded fluorescent proteins, or the incorporation of unnatural amino acids into the region of interest1. This additional data yields kinetic information about voltage-dependent conformational rearrangements of the protein via changes in the microenvironment surrounding the fluorescent molecule.

The Xenopus Oocyte Cut-open Vaseline Gap Voltage-clamp Technique With Fluorometry

The cut-open Vaseline gap approach is used to obtain low noise recordings of ionic and gating currents from voltage-dependent ion channels expressed in Xenopus oocytes with high resolution of fast channel kinetics. With minor modification, voltage clamp fluorometry can be coupled to the cut-open oocyte protocol.

The cut-open oocyte Vaseline gap (COVG) voltage clamp technique allows for analysis of electrophysiological and kinetic properties of heterologous ion ...