We thank Dartmouth College&#39;s Center for Comparative Medicine and Research for providing daily husbandry for the animals used in this protocol. We also thank Leah Jacob and Adwaita Bose for their help in testing the protocol and photographing animals. Lastly, we thank Ann Miller&#39;s lab for training in the practice of tattooing. This work was supported by NIH grant R00 GM147826 to J.L.

All animal procedures described were approved by Dartmouth College&#39;s Institutional Animal Care and Use Committee.
1. Equipment setup
NOTE: A workflow for the procedure and an example bench setup are included (Figure 1).
<ol>
	<li>Connect the tattoo gun and foot pedal to the power supply. Position the foot pedal underneath the working surface.</li>
	<li>Assembling the tattoo gun
	<ol>
		<li>Using an Allen wrench or hex screwdriver, loosen the screws in the grip (Figure 2A, encircled in red).</li>
		<li>Insert the plastic tip all the way into the grip. Tighten the screw to hold it in place.</li>
		<li>Insert the metal tube in the back of the grip (Figure 2A). This will be adjusted, so insert it to half its length. Tighten the screw to hold it into place.</li>
		<li>Insert a tattoo needle through the back of the grip so that it sits comfortably in the plastic tip at the front of the grip (Figure 2B).</li>
		<li>Slide the back of the needle and metal tube through the tube clamp. Adjust the metal tube so that the needle fits comfortably in the plastic tip (Figure 2B). Hand-tighten the tube clamp until the tube is unable to move (Figure 2C).</li>
		<li>Remove the black O-ring and hook the needle onto the rotor arm; replace the O-ring to secure the needle into place (Figure 2C).</li>
	</ol>
	</li>
	<li>Set the voltage on the power supply so that there is enough power to pierce the skin but not so powerful that it is difficult to control. Once the power supply is plugged in and connected, press the foot pedal to make sure the machine is working. Take care not to point the needle at yourself or other personnel. 
	NOTE: The tattoo machine used in this protocol works best between 6.0 and 9.0 V, but this may vary between tattoo machines and voltage supplies and should be determined empirically.</li>
	<li>Fill the cap of a 1.5 mL microcentrifuge tube or a provided plastic inkpot with black tattoo ink to about three-quarters full.</li>
</ol>
2. Anesthesia
<ol>
	<li>Prepare anesthesia in a tank large enough to submerge one adult female X. laevis frog.
	<ol>
		<li>Using frog-safe (chlorine-free) water, add tricaine to the final concentration of 1.5 g/L and sodium bicarbonate to the final concentration of 3.5 g/L. Mix to dissolve. The pH of this solution is 7.15. 
		CAUTION: Tricaine is an irritant.</li>
	</ol>
	</li>
	<li>Submerge one frog in the anesthesia tank. Ensure that the frog stays submerged in the tricaine solution until it is anesthetized (7-8 min).
	<ol>
		<li>To check if the frog is fully anesthetized, pick it up, hold it upside down, and give it a firm squeeze on one foot. If the frog does not flinch or respond, it can be tattooed.</li>
		<li>If frogs consistently take more than 10 min to be anesthetized, prepare fresh tricaine solution buffered with sodium bicarbonate. 
		NOTE: Frogs should not spend more than 30 min in tricaine solution15.</li>
	</ol>
	</li>
</ol>
3. Tattooing
NOTE: Before tattooing a live animal, it can be useful to practice on a piece of fruit with a firm peel (such as a lemon or banana).
<ol>
	<li>Prepare a recovery tank for the frogs before tattooing. Fill a tank with 10-15 L of fresh frog-safe water and add a Styrofoam island. This will provide a surface for the frog to wake up without drowning. 
	NOTE: A Styrofoam island can be made using the lid of a Styrofoam cold shipping container placed inside a zip-top bag.</li>
	<li>Place the anesthetized frog onto its back on a dry paper towel or bench paper (Figure 3A). Use unbleached (brown) paper towels for all work involving frogs.</li>
	<li>Using a dry lint-free wipe, wipe away the water and mucus from the frog&#39;s chest.</li>
	<li>Find the sternum in the center of the frog&#39;s chest with fingers and, using the non-dominant hand, hold the skin taut.</li>
	<li>Marking the frog
	<ol>
		<li>Hold the assembled tattoo gun vertically relative to the working surface and dip the needle into the ink (Figure 3B).</li>
		<li>Keeping the assembled tattoo gun vertical, press the tip of the needle onto the frog&#39;s skin before pressing the foot pedal. Draw lines onto the frog&#39;s skin while applying even pressure. 
		NOTE: Some minor bleeding or redness is normal, as is minor skin sloughing.</li>
		<li>If the needle is getting caught on the frog&#39;s skin, clear excess ink or skin from the needle with a wipe. Excess ink is normal during tattooing.</li>
		<li>If the frog&#39;s chest is covered in too much ink to clearly see the area being marked (Figure 3C), clear excess ink with frog-safe water and a wipe (Figure 3C&#39;). Then, wick away moisture with a dry wipe and continue inking. 
		NOTE: A squeeze bottle of frog-safe water can be prepared before tattooing.</li>
		<li>Continue to ink the same area until dark, legible numbers remain after wiping away excess ink (Figure 3D). Instead of drawing the entire marking or number in one stroke, repeatedly make smaller strokes, especially for curves.</li>
	</ol>
	</li>
</ol>
4. Recovery
<ol>
	<li>Wet a paper towel with frog-safe water and place it flat on top of the Styrofoam island (Figure 4A).</li>
	<li>Returning frogs to standing tanks
	<ol>
		<li>Lay the frog belly-down on the towel facing the water (Figure 4B). Fold half of the paper towel over the back half of the frog and, using cupped hands, wet the top of the frog. Once a frog emerges from anesthesia (about 1 h after tattooing), they will enter water independently and swim normally.</li>
		<li>Return frogs to long-term housing after 24 h, once the tattoo has fully healed.</li>
	</ol>
	</li>
</ol>
5. Cleaning and equipment maintenance
NOTE: Although needles can be reused for multiple frogs in the same sitting, best practice is to replace or sterilize needles between tattooing frogs.
<ol>
	<li>Reusing and autoclaving tattooing needles
	<ol>
		<li>Rinse needles in 100% isopropanol and deionized water to loosen the ink and scrub off the remaining ink that has not dissolved.</li>
		<li>After scrubbing, place the needles into a metal autoclaving case or wrap them with aluminum foil and autoclave for 30 min on a dry cycle.</li>
	</ol>
	</li>
	<li>If there is ink on pieces of the tattoo machine or other equipment, wipe it with 100% ethanol and a paper towel.</li>
	<li>The anesthetic solution has a final tricaine concentration of approximately 0.5%. Dispose of this according to institutional hazardous waste management guidelines.</li>
	<li>Equipment storage and maintenance
	<ol>
		<li>If possible, store the tattoo machine in a dry room. Excess moisture will damage the electric components and reduce the longevity of the machine.</li>
		<li>To keep the tattoo machine in good working order, check the bearing on the rotor (Figure 2C) to make sure it has not come loose from vibrations22. This can be tightened with an Allen wrench or hex screwdriver.</li>
		<li>About once a month, lubricate the bearing on the rotor arm using a thick lubricating grease22.</li>
		<li>On some tattoo machines, the rotor arm can get stuck and will not vibrate. To correct this, twist the rotor arm or compress the spring so that the needle is in the lowered position (Figure 2C).</li>
	</ol>
	</li>
</ol>

Optimizing Amphibian Marking: A Guide to Tattooing Xenopus

<ol>
	<li>Hubrecht, R. C., Carter, E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+3Rs+and+humane+experimental+technique:+Implementing+change.">The 3Rs and humane experimental technique: Implementing change.</a> Animals (Basel). 9 (10), 754 (2019).</li><li>Elkan, E. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+Xenopus+pregnancy+test.">The Xenopus pregnancy test.</a> British Med J. 2, 1253-1256 (1938).</li><li>Vize, P. D., Zorn, A. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Xenopus+genomic+data+and+browser+resources.">Xenopus genomic data and browser resources.</a> Dev Biol. 426 (2), 194-199 (2017).</li><li>Sive, H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Xenopus: A Laboratory Manual. , (2023).</li><li>Borodinsky, L. N. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Xenopus+laevis+as+a+model+organism+for+the+study+of+spinal+cord+formation,+development,+function+and+regeneration.">Xenopus laevis as a model organism for the study of spinal cord formation, development, function and regeneration.</a> Front Neural Circuits. 11, 90 (2017).</li><li>Hoppler, S., Conlon, F. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Xenopus:+Experimental+access+to+cardiovascular+development,+regeneration+discovery,+and+cardiovascular+heart-defect+modeling.">Xenopus: Experimental access to cardiovascular development, regeneration discovery, and cardiovascular heart-defect modeling.</a> Cold Spring Harb Perspect Biol. 12 (6), a037200 (2020).</li><li>Fisher, M., 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:+key+features+and+resources+of+the+Xenopus+model+organism+knowledgebase.">Xenbase: key features and resources of the Xenopus model organism knowledgebase.</a> Genetics. 224 (1), 018 (2023).</li><li>Slater, P. G., Hayrapetian, L., Lowery, L. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Xenopus+laevis+as+a+model+system+to+study+cytoskeletal+dynamics+during+axon+pathfinding.">Xenopus laevis as a model system to study cytoskeletal dynamics during axon pathfinding.</a> Genesis. 55 (1-2), 22994 (2017).</li><li>Bermudez, J. G., Chen, H., Einstein, L. C., Good, M. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Probing+the+biology+of+cell+boundary+conditions+through+confinement+of+Xenopus+cell-free+cytoplasmic+extracts.">Probing the biology of cell boundary conditions through confinement of Xenopus cell-free cytoplasmic extracts.</a> Genesis. 55 (1-2), 23013 (2017).</li><li>Stooke-Vaughan, G. A., Davidson, L. A., Woolner, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Xenopus+as+a+model+for+studies+in+mechanical+stress+and+cell+division.">Xenopus as a model for studies in mechanical stress and cell division.</a> Genesis. 55 (1-2), 23004 (2017).</li><li>Blackburn, A. T. M., Miller, R. K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Modeling+congenital+kidney+diseases+in+Xenopus+laevis.">Modeling congenital kidney diseases in Xenopus laevis.</a> Dis Model Mech. 12 (4), 038604 (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> Dev Biol. 426 (2), 325-335 (2017).</li><li>Wheeler, G. N., Br&#228;ndli, A. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Simple+vertebrate+models+for+chemical+genetics+and+drug+discovery+screens:+Lessons+from+zebrafish+and+Xenopus.">Simple vertebrate models for chemical genetics and drug discovery screens: Lessons from zebrafish and Xenopus.</a> Dev Dyn. 236 (6), 1287-1308 (2009).</li><li>Carron, M., 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=Evolutionary+origin+of+Hox13-dependent+skin+appendages+in+amphibians.">Evolutionary origin of Hox13-dependent skin appendages in amphibians.</a> Nat Commun. 15 (1), 2328 (2024).</li><li>Schultz, T. W., Dawson, D. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Housing+and+husbandry+of+Xenopus+for+oocyte+production.">Housing and husbandry of Xenopus for oocyte production.</a> Lab Animal. 32 (2), 34-39 (2003).</li><li>Cadillac, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Animal identification systems used for mice. , (2006).</li><li>Donnelly, M. A., Guyer, C., Juterbock, J. E., Alford, R. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Techniques+for+marking+amphibians.">Techniques for marking amphibians.</a> Meas Monitor Biol Div: Std Meth Amphibians. , 277-284 (1994).</li><li>Hutchens, S. J., Deperno, C. S., Matthews, C. E., Pollock, K. H., Woodward, D. K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Visible+implant+fluorescent+marker:+A+reliable+marking+alternative+for+snakes.">Visible implant fluorescent marker: A reliable marking alternative for snakes.</a> Herpetol Rev. 39 (3), 301-303 (2008).</li><li>Hoogstraten-Miller, S., Dunham, D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Practical+identification+methods+for+African+clawed+frogs+(Xenopus+laevis).">Practical identification methods for African clawed frogs (Xenopus laevis).</a> Lab Animal. 26 (7), 36-38 (1997).</li><li>Loopstra, J. A., Zwart, P., Verhoeff-de Fremery, R., Vervoordeldonk, F. J. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Marking+of+African+clawed+toads+(Xenopus+laevis).+Improvement+of+a+skin+autograft+technique.">Marking of African clawed toads (Xenopus laevis). Improvement of a skin autograft technique.</a> New Dev Biosci: Their Implicat Lab Animal Sci. , 311-316 (1988).</li><li>Rodel, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Xenopus+tropicalis.">Xenopus tropicalis.</a> AmphibiaWeb. , (2001).</li><li>Worldwide Tattoo Supply. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Stealth+2+Rotary+Tattoo+Machine+Intro+&amp;+Maintenance.">Stealth 2 Rotary Tattoo Machine Intro &amp; Maintenance.</a> Worldwide Tattoo Supply. , (2015).</li><li>Khan, S. U., Mufti, O. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+hot+history+and+cold+future+of+brands.">The hot history and cold future of brands.</a> J Manager Sci. 1 (1), 75-84 (2007).</li><li>Catlett, T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=All+About+Lip+Tattoo:+what+to+Expect,+Cost,+Risks,+Touch-ups+&amp;+More.">All About Lip Tattoo: what to Expect, Cost, Risks, Touch-ups &amp; More.</a> Healthline. , (2019).</li></ol>

The authors declare no competing interests.

Tattooing humans is an art form dating back thousands of years, and for as long as humans have tattooed themselves, they have also tattooed or branded animals23. The equipment and techniques for marking animals, particularly mammals, are well-established, well-documented, and widely accessible. While marking animals was originally for distinguishing livestock and theft deterrents23, its role in biomedical research has become just as important.
A ...

Animal models are useful tools for investigating questions pertaining to human health. In practice, biomedical research using animal models requires careful organization and maintenance of a healthy animal colony. Best practices for ethical animal handling and husbandry aim to reduce the number of animals needed for experimentation and refine practices to ensure animal welfare1. The clawed frog genus, including Xenopus laevis (X. laevis; African clawed frog) and Xenopus tropicalis (X. tropicalis; Western clawed frog), have been used in biomedical research since the 1930s when X. laevis were used by South African doctors to conduct the first pregnancy tests2. While modern pregnancy tests no longer require frogs, the role of Xenopus research persists. Advantages of using Xenopus for biomedical research include their well-annotated genomes3, year-round inducible ovulation of large clutches of eggs4, and externally laid eggs amenable to in vitro fertilization. These features make them a useful asset for vertebrate embryology and development5,6,7, basic molecular and cellular biology7,8,9,10, and for modeling human disease7,11,12,13.
Reliable methods for tracking individual Xenopus animals are essential for recording biological replicates and improving rigor and reproducibility in research. As Xenopus are frequently housed in groups, animal marking allows researchers to easily track individual animals4. Maintaining an accurate record of animals can save time and resources and improve the ability to track animals&#39; health. For example, individual identification of animals can improve organization workflows for generating transgenic Xenopus lines, as this requires multiple generations of frogs with specific genotypes verified by sequencing14, which requires organization and individual identification of animals. This is particularly true when these mutations lack easily discernible adult phenotypes. Similarly, the use of Xenopus oocytes and embryos to study basic cellular and developmental biology benefits from tracking individual animals. After inducing ovulation, animals need to rest for a minimum of 3 months to prevent health complications such as hyper-ovulation syndrome15. Individual identification methods ensure that animals are not induced to ovulate too frequently.
Marking and tracking animals also enables lab personnel to track animals&#39; health concerns. Animals of the same genotype falling ill can indicate excessive inbreeding or unanticipated health concerns associated with the transgene. Similarly, animals falling ill after recent ovulation can indicate issues with reagents, materials, or techniques. Tracking animals and their health enables lab personnel and veterinarians to follow up when concerns resurface and take preventative measures to prevent future illness. In mammals, there are numerous identification methods. Permanent methods for mice include ear punching, ear tags, tattooing, and subcutaneous microchips16. These can clearly and reliably differentiate animals within a colony or cage and can be easily administered by laboratory personnel. Methods such as ear punching are minimally invasive, require only one piece of specialized equipment, and work for animals of most ages. While these systems are straightforward and useful for mice, their use in frogs presents a unique set of challenges. Frogs and other amphibians lack a pinna (external ear structure). Some researchers have attached tags to the animal&#39;s jaw, toe, or hind limb17,18. This approach resulted in various problems: jaw tags caused irritation, and agitated frogs attempted to pull off tags with their forelimbs17. Toe tags pierced the webbing between toes, impairing movement and carrying the risk of becoming lost. As such, amphibians require their own methods for identification. Historically, toe-clipping has also been used to mark amphibians17,19. The toe is clipped with a sharp pair of scissors, and the animal can be identified by the length of toes on the forefeet and hind feet or by the angle at which the toe has grown back (in salamanders). However, this method poses the ethical concern that toe clipping may impair the animal&#39;s movement17. In addition, this can cause bleeding and introduce a risk of infection. Another established marking system is skin autografts, in which skin is taken from one part of the frog and surgically attached to another part. For example, a method is described for marking a frog&#39;s back or shoulder using a light-colored skin graft from its chest20. Skin grafts also come with limitations and risks: the procedure is invasive and introduces the risk of Aeromonas hydrophila infection, or red leg, a potentially fatal affliction; complete healing of the autograft takes up to 6 weeks; and with the methods described, only 6 frogs can be housed together because of limited places to put an autograft20.
Less invasive marking approaches include glass beads and transponder chips17,19. In the glass bead method, glass beads are threaded onto a small suture and sewn into the frog&#39;s skin. This provides greater variability than skin autografts, with at least 60 distinctive color combinations. There is, however, a risk that the suture can come out and result in the beads being lost. Alternatively, a microchip transponder can be implanted under the skin in the frog&#39;s dorsal lymph sac. This is considered the most permanent marking method and enables a potentially infinite number of animals to be individually identified and cataloged. However, this is also the most expensive method, as individual microchips are expensive, and a large colony would be costly to mark. Microchips also require a special scanner to read19. One common approach for Xenopus identification is referring to animals&#39; natural coloring and patterning. This works especially well for frogs such as X. laevis, which have distinct patterns that remain throughout adulthood. However, these patterns can change over time with stress, and coloring can appear different when frogs are moved between transparent and tinted containers15. Additionally, this identification method is less useful for X. tropicalis, which has less distinct marking patterns compared to X. laevis, or for albino animals, which have no color markings21. Even for species with distinct markings, lab personnel can interpret the placement and size of markings differently, which can cause errors in identification. Because of this, photographing animals is most reliable in conjunction with an additional identification method. Therefore, we seek to mark and identify Xenopus animals using a technique that is easily discernable, permanent, and minimally invasive.
There are limited published resources describing methods for tattooing amphibians. Tattooing has been described alongside other branding techniques, including heat brands, silver nitrate brands, and freeze brands17. In the same resource, tattooing was done by drawing numerals with a 27G hypodermic needle, and the process was noted not to cause infection, in contrast to the other branding techniques, which used a wire shaped into a numeral or other mark. In another source, an electric tattooing machine (described as a vibrating needle) was used to mark frogs, but little detail was provided on the technique17,19. The authors warn that by disturbing the frog&#39;s protective slime layer, this procedure increases the risk of red leg. While there is no marking or identification method that is both completely noninvasive (such as photographs) and permanent (such as microchips), tattooing provides an effective compromise. Tattooing is relatively straightforward compared to other techniques, such as skin autografts. Additional benefits include a smaller learning curve and relatively inexpensive equipment. Tattooing aquatic amphibians comes with certain challenges, which can intimidate researchers and impair successful animal marking. This paper aims to provide researchers with well-documented methods for tattooing adult Xenopus with a rotary tattoo machine.

<table><tbody><tr><td>3 needle round liners</td><td>Worldwide Tattoo Supply</td><td>1203RLB</td><td>Packaged sterile</td></tr><tr><td>5 Needle Round Disposable ULTRA</td><td>Worldwide Tattoo Supply</td><td>HTIPRS5-U</td><td>Packaged sterile</td></tr><tr><td>5 needle round liners</td><td>Worldwide Tattoo Supply</td><td>1205RLB</td><td>Packaged sterile</td></tr><tr><td>7 needle round liners</td><td>Worldwide Tattoo Supply</td><td>1207RLB</td><td>Packaged sterile</td></tr><tr><td>Clip Cord</td><td>Worldwide Tattoo Supply</td><td>N/A</td><td></td></tr><tr><td>Foot pedal</td><td>Worldwide Tattoo Supply</td><td>N/A</td><td></td></tr><tr><td>Inkpots</td><td>Worldwide Tattoo Supply</td><td>N/A</td><td></td></tr><tr><td>Kimwipes, delicate task wipes</td><td>Fisher Scientific</td><td>06-666A</td><td></td></tr><tr><td>RCA Connection</td><td>Worldwide Tattoo Supply</td><td>N/A</td><td></td></tr><tr><td>Scream Ink Pitch Black, 1oz</td><td>Worldwide Tattoo Supply</td><td>SI101</td><td></td></tr><tr><td>Sodium bicarbonate (NaHCO3)</td><td>Sigma-Aldrich</td><td>S5761</td><td></td></tr><tr><td>Stainless steel grips</td><td>Worldwide Tattoo Supply</td><td>N/A</td><td></td></tr><tr><td>Stealth 2.0 Rotary Tattoo Machine</td><td>Worldwide Tattoo Supply</td><td>N/A</td><td></td></tr><tr><td>Stealth 2.0 Rotary Tattoo Machine Box Set</td><td>Worldwide Tattoo Supply</td><td>STEALTH2-SET</td><td></td></tr><tr><td>Styrofoam island</td><td>N/A</td><td>N/A</td><td>This is the lid of a styrofoam cold shipping container</td></tr><tr><td>Tricaine (ethyl 3-aminobenzonate methanesulfate)</td><td>Sigma-Aldrich</td><td>E10521</td><td>CAUTION: IRRITANT</td></tr><tr><td>Unbleached paper towels</td><td>Grainger</td><td>2U229</td><td>Paper towels MUST be unbleached, bleach is toxic to amphibians</td></tr><tr><td>Voltage Supply</td><td>Worldwide Tattoo Supply</td><td>N/A</td><td></td></tr><tr><td>Wash bottle (with frog-safe water)</td><td>Fisher Scientific</td><td>FB0340923T</td><td>Frog safe water is dechlorinated, pH 7.0-8.5, conductivity 1200-1800 uS</td></tr><tr><td>&lt;em&gt;X. laevis&lt;/em&gt; adult female</td><td>Xenopus1</td><td>N/A</td><td></td></tr><tr><td>Zip-top plastic bag</td><td>N/A</td><td>N/A</td><td>This bag should be large enough to hold the styrofoam island</td></tr></tbody></table>

methods for tattooing xenopus laevis with a rotary tattoo machine

Animal models expand the scope of biomedical research, furthering our understanding of developmental, molecular, and cellular biology and enabling researchers to model human disease. Recording and tracking individual animals allows researchers to reduce the number of animals required for study and refine practices to improve animal wellbeing. Several well-documented methods exist for marking and tracking mammals, including ear punching and ear tags. However, methods for marking aquatic amphibian species are limited, with the existing resources being outdated, ineffective, or prohibitively costly. In this manuscript, we outline methods and best practices for marking Xenopus laevis with a rotary tattoo machine. Proper tattooing results in high-quality tattoos, making individuals easily distinguishable for researchers and posing minimal risk to animals' health. We also highlight the causes of poor-quality tattoos, which can result in tattoos that fade quickly and cause unnecessary harm to animals. This approach allows researchers and veterinarians to mark amphibians, enabling them to track biological replicates and transgenic lines and to keep accurate records of animal health.

High-quality tattoos will have dark, legible strokes on the frog&#39;s chest and can be clearly differentiated from several feet away (Figure 5A). In general, larger numbers and markings are better for readability, but longer names and numbers can be made smaller to fit comfortably on the frog&#39;s chest. Tattoo longevity is more difficult to judge, but high-quality frog tattoos should remain dark and legible for at least 3-6 months (Figure 5D-D&#39;</s...

Author Spotlight: Refining Xenopus laevis Marking Techniques for Biomedical Studies

Here, we describe methods for tattooing adult Xenopus laevis (African clawed frog) with a rotary tattoo machine. Proper tattooing results in dark, easily legible numerals that last for several months and make animals easily distinguishable for research and recordkeeping purposes.

Methods for Tattooing Xenopus laevis with a Rotary Tattoo Machine

Animal models expand the scope of biomedical research, furthering our understanding of developmental, molecular, and cellular biology and enabling ...

methods-for-tattooing-xenopus-laevis-with-a-rotary-tattoo-machine

Research

JoVE Journal

Biology

567 Views.  Dartmouth College. Here, we describe methods for tattooing adult Xenopus laevis (African clawed frog) with a rotary tattoo machine. Proper tattooing results in dark, easily legible numerals that last for several months and make animals easily distinguishable for research and recordkeeping purposes.

Video: Author Spotlight: Refining Xenopus laevis Marking Techniques for Biomedical Studies

Using Confocal Analysis of Xenopus laevis to Investigate Modulators of Wnt and Shh Morphogen Gradients

This protocol describes a method to visualise ligands distributed across a field of cells. The ease of expressing exogenous proteins, together with the large size of their cells in early embryos, make Xenopus laevis a useful model for visualising GFP-tagged ligands. Synthetic mRNAs are efficiently translated after injection into early stage Xenopus embryos, and injections can be targeted to a single cell. When combined with a lineage tracer such as membrane tethered RFP, the injected cell (and its descendants) that are producing the overexpressed protein can easily be followed. This protocol describes a method for the production of fluorescently tagged Wnt and Shh ligands from injected mRNA. The methods involve the micro dissection of ectodermal explants (animal caps) and the analysis of ligand diffusion in multiple samples. By using confocal imaging, information about ligand secretion and diffusion over a field of cells can be obtained. Statistical analyses of confocal images provide quantitative data on the shape of ligand gradients. These methods may be useful to researchers who want to test the effects of factors that may regulate the shape of morphogen gradients.

Using Confocal Analysis of Xenopus laevis to Investigate Modulators of Wnt and Shh Morphogen Gradients

The manuscript here provides a simple set of methods for analysing the secretion and diffusion of fluorescently tagged ligands in Xenopus. This provides a context for testing the ability of other proteins to modify ligand distribution and allowing experiments that may give insight into mechanisms regulating morphogen gradients.

This protocol describes a method to visualise ligands distributed across a field of cells. The ease of expressing exogenous proteins, together with the ...

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 ...

Skin Tattooing As A Novel Approach For DNA Vaccine Delivery

Nucleic acid-based vaccination is a topic of growing interest, especially plasmid DNA (pDNA) encoding immunologically important antigens. After the engineered pDNA is administered to the vaccines, it is transcribed and translated into immunogen proteins that can elicit responses from the immune system. Many ways of delivering DNA vaccines have been investigated; however each delivery route has its own advantages and pitfalls. Skin tattooing is a novel technique that is safe, cost-effective, and convenient. In addition, the punctures inflicted by the needle could also serve as a potent adjuvant. Here, we a) demonstrate the intradermal delivery of plasmid DNA encoding enhanced green fluorescent protein (pCX-EGFP) in a mouse model using a tattooing device and b) confirm the effective expression of EGFP in the skin cells using confocal microscopy.

Skin tattooing is a potent and safe way to delivery DNA vaccine intradermally. Here, a DNA plasmid encoding EGFP is delivered by tattooing to the skin of a laboratory mouse, and the expression of EGFP in the skin cells is then inspected by confocal microscopy.

Nucleic acid-based vaccination is a topic of growing interest, especially plasmid DNA (pDNA) encoding immunologically important antigens. After the ...

Bioengineering

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 ...

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 ...

Neural Explant Cultures from Xenopus laevis

The complex process of axon guidance is largely driven by the growth cone, which is the dynamic motile structure at the tip of the growing axon. During axon outgrowth, the growth cone must integrate multiple sources of guidance cue information to modulate its cytoskeleton in order to propel the growth cone forward and accurately navigate to find its specific targets1. How this integration occurs at the cytoskeletal level is still emerging, and examination of cytoskeletal protein and effector dynamics within the growth cone can allow the elucidation of these mechanisms. Xenopus laevis growth cones are large enough (10-30 microns in diameter) to perform high-resolution live imaging of cytoskeletal dynamics (e.g.2-4 ) and are easy to isolate and manipulate in a lab setting compared to other vertebrates. The frog is a classic model system for developmental neurobiology studies, and important early insights into growth cone microtubule dynamics were initially found using this system5-7 . In this method8, eggs are collected and fertilized in vitro, injected with RNA encoding fluorescently tagged cytoskeletal fusion proteins or other constructs to manipulate gene expression, and then allowed to develop to the neural tube stage. Neural tubes are isolated by dissection and then are cultured, and growth cones on outgrowing neurites are imaged. In this article, we describe how to perform this method, the goal of which is to culture Xenopus laevis growth cones for subsequent high-resolution image analysis. While we provide the example of +TIP fusion protein EB1-GFP, this method can be applied to any number of proteins to elucidate their behaviors within the growth cone.

Neural Explant Cultures from Xenopus laevis

Culturing neural explants from dissected Xenopus laevis embryos that express fluorescent fusion proteins allows for imaging of growth cone cytoskeletal dynamics.

The complex process of axon guidance is largely driven by the growth cone, which is the dynamic motile structure at the tip of the growing axon. During ...

Neuroscience

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

The primary visual projection of tadpoles of the aquatic frog Xenopus laevis serves as an excellent model system for studying mechanisms that regulate the development of neuronal connectivity. During establishment of the retino-tectal projection, optic axons extend from the eye and navigate through distinct regions of the brain to reach their target tissue, the optic tectum. Once optic axons enter the tectum, they elaborate terminal arbors that function to increase the number of synaptic connections they can make with target interneurons in the tectum. Here, we describe a method to express DNA encoding green fluorescent protein (GFP), and gain- and loss-of-function gene constructs, in optic neurons (retinal ganglion cells) in Xenopus embryos. We explain how to microinject a combined DNA/lipofection reagent into eyebuds of one day old embryos such that exogenous genes are expressed in single or small numbers of optic neurons. By tagging genes with GFP or co-injecting with a GFP plasmid, terminal axonal arbors of individual optic neurons with altered molecular signaling can be imaged directly in brains of intact, living Xenopus tadpoles several days later, and their morphology can be quantified. This protocol allows for determination of cell-autonomous molecular mechanisms that underlie the development of optic axon arborization in vivo.

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

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

The primary visual projection of tadpoles of the aquatic frog Xenopus laevis serves as an excellent model system for studying mechanisms that regulate the ...

Production of Transgenic Xenopus laevis by Restriction Enzyme Mediated Integration and Nuclear Transplantation

Stable integration of cloned gene products into the Xenopus genome is necessary to control the time and place of expression, to express genes at later stages of embryonic development, and to define how enhancers and promoters regulate gene expression within the embryo. The protocol demonstrated here can be used to efficiently produce transgenic Xenopus laevis embryos. This transgenesis approach involves three parts: 1. Sperm nuclei are isolated from adult X. laevis testis by treatment with lysolecithin, which permeabilizes the sperm plasma membrane. 2. Egg extract is prepared by low speed centrifugation, addition of calcium to cause the extract to progress to interphase of the cell cycle, and a high-speed centrifugation to isolate interphase cytosol. 3. Nuclear transplantation: the nuclei and extract are combined with the linearized plasmid DNA to be introduced as the transgene and a small amount of restriction enzyme. During a short reaction, egg extract partially decondenses the sperm chromatin and the restriction enzyme generates chromosomal breaks that promote recombination of the transgene into the genome. The treated sperm nuclei are then transplanted into unfertilized eggs. Integration of the transgene usually occurs prior to the first embryonic cleavage such that the resulting embryos are not chimeric. These embryos can be analyzed without any need to breed to the next generation, allowing for efficient and rapid generation of transgenic embryos for analyses of promoter and gene function. Adult X. laevis resulting from this procedure also propagate the transgene through the germline and can be used to generate lines of transgenic animals for multiple purposes.

Production of Transgenic Xenopus laevis by Restriction Enzyme Mediated Integration and Nuclear Transplantation

This video protocol demonstrates a method for generating transgenic Xenopus laevis by introduction of transgenes into sperm nuclei followed by nuclear transplantation into unfertilized eggs.

Stable integration of cloned gene products into the Xenopus genome is necessary to control the time and place of expression, to express genes at later ...

Polarized Translocation of Fluorescent Proteins in Xenopus Ectoderm in Response to Wnt Signaling

Cell polarity is a fundamental property of eukaryotic cells that is dynamically regulated by both intrinsic and extrinsic factors during embryonic development 1, 2. One of the signaling pathways involved in this regulation is the Wnt pathway, which is used many times during embryogenesis and critical for human disease3, 4, 5. Multiple molecular components of this pathway coordinately regulate signaling in a spatially-restricted manner, but the underlying mechanisms are not fully understood. Xenopus embryonic epithelial cells is an excellent system to study subcellular localization of various signaling proteins. Fluorescent fusion proteins are expressed in Xenopus embryos by RNA microinjection, ectodermal explants are prepared and protein localization is evaluated by epifluorescence. In this experimental protocol we describe how subcellular localization of Diversin, a cytoplasmic protein that has been implicated in signaling and cell polarity determination6, 7 is visualized in Xenopus ectodermal cells to study Wnt signal transduction8. Coexpression of a Wnt ligand or a Frizzled receptor alters the distribution of Diversin fused with red fluorescent protein, RFP, and recruits it to the cell membrane in a polarized fashion 8, 9. This ex vivo protocol should be a useful addition to in vitro studies of cultured mammalian cells, in which spatial control of signaling differs from that of the intact tissue and is much more difficult to analyze.

Polarized Translocation of Fluorescent Proteins in Xenopus Ectoderm in Response to Wnt Signaling

Xenopus embryonic ectoderm has become an attractive model for studies of cell polarity. An assay is described, in which subcellular distribution of fluorescent proteins is assessed in ectoderm cells. This protocol will help address questions related to spatial control of signaling.

Cell polarity is a fundamental property of eukaryotic cells that is dynamically regulated by both intrinsic and extrinsic factors during embryonic ...

Xenopus Oocytes: Optimized Methods for Microinjection, Removal of Follicular Cell Layers, and Fast Solution Changes in Electrophysiological Experiments

The Xenopus oocyte as a heterologous expression system for proteins, was first described by Gurdon et al.1 and has been widely used since its discovery (References 2 - 3, and references therein). A characteristic that makes the oocyte attractive for foreign channel expression is the poor abundance of endogenous ion channels4. This expression system has proven useful for the characterization of many proteins, among them ligand-gated ion channels.
The expression of GABAA receptors in Xenopus oocytes and their functional characterization is described here, including the isolation of oocytes, microinjections with cRNA, the removal of follicular cell layers, and fast solution changes in electrophysiological experiments. The procedures were optimized in this laboratory5,6 and deviate from the ones routinely used7-9. Traditionally, denuded oocytes are prepared with a prolonged collagenase treatment of ovary lobes at RT, and these denuded oocytes are microinjected with mRNA. Using the optimized methods, diverse membrane proteins have been expressed and studied with this system, such as recombinant GABAA receptors10-12, human recombinant chloride channels13, Trypanosome potassium channels14, and a myo-inositol transporter15, 16.
The methods detailed here may be applied to the expression of any protein of choice in Xenopus oocytes, and the rapid solution change can be used to study other ligand-gated ion channels.

Xenopus Oocytes: Optimized Methods for Microinjection, Removal of Follicular Cell Layers, and Fast Solution Changes in Electrophysiological Experiments

Optimized procedures for the isolation of single follicles, cytoplasmic RNA microinjections, the removal of surrounding cell layers, and protein expression in Xenopus oocytes are described. In addition, a simple method for fast solution changes in electrophysiological experiments with ligand-gated ion channels is presented.

The Xenopus oocyte as a heterologous expression system for proteins, was first described by Gurdon et al.1 and has been widely used since its discovery ...

Methods for Tattooing Xenopus laevis with a Rotary Tattoo Machine

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Chapters in this video

Using Confocal Analysis of Xenopus laevis to Investigate Modulators of Wnt and Shh Morphogen Gradients

Technique to Target Microinjection to the Developing Xenopus Kidney

Skin Tattooing As A Novel Approach For DNA Vaccine Delivery

Visualization and Quantitative Analysis of Embryonic Angiogenesis in Xenopus tropicalis

Microinjection of Xenopus Laevis Oocytes

Neural Explant Cultures from Xenopus laevis

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

Production of Transgenic Xenopus laevis by Restriction Enzyme Mediated Integration and Nuclear Transplantation

Polarized Translocation of Fluorescent Proteins in Xenopus Ectoderm in Response to Wnt Signaling

Xenopus Oocytes: Optimized Methods for Microinjection, Removal of Follicular Cell Layers, and Fast Solution Changes in Electrophysiological Experiments