The authors would like to thank Michelle Deochand for perfecting this eye ablation technique, Taylor Birkholz for assistance with the functional assay, Michael Levin for the anti-arrestin antibody, and Junji Morokuma for information on the Peltier plates. This work was supported by an SFSA grant from Western Michigan University to WSB.

1. Animal Culture and Handling
NOTE: This protocol uses Schmidtea mediterranea, a diploid planarian species with a sequenced genome16,17 that is commonly used for regeneration research. However, the assay is equally successful with other species, such as Girardia tigrina and Girardia dorotocephala (which are commercially available).
<ol>
	<li>Maintain worms in &#34;worm water&#34; made from 0.5 g/L sea salts in ultrapure (or filtered deionized) water. Use sterile polypropylene or glass containers to hold the worm water. See <a href="https://cloudfront-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/55594/Supplementary_File_1.docx" target="_blank">Supplementary File 1</a> for details on worm water preparation. 
	NOTE: Never use soap to clean containers (or other supplies) as soap is toxic to worms; instead wipe with 70% ethanol and allow to air dry.
	<ol>
		<li>Wear gloves while working with planaria to protect them from contamination. 
		NOTE: Worms are very sensitive to environmental toxins and chemicals, including soap, bleach, shampoo, conditioner, and hand lotion.</li>
	</ol>
	</li>
	<li>House colonies in polypropylene food containers, with the lid left partially open for air exchange at ~20 &#176;C (room temperature). Store experimental worms in either petri dishes (20 worms per 100 mm dish) or non-treated tissue-culture plates (1 worm per well, for 12-well plates). To reduce stress, keep both colonies and experiments in the dark. 
	NOTE: Colonies should have no more than 500-1,000 worms per liter of worm water. For experiments, leave lids fully in place, since airflow is designed into these dishes.</li>
	<li>Feed non-experimental worms once a week with pureed organic beef or chicken liver by dropping puree from a transfer pipette, placing drops of food throughout the container. Allow worms 2 h to consume food in the dark before cleaning out container (see step 1.4). Prior to use, store puree at -20 &#176;C for short term (weeks) or -80 &#176;C for long term (months); do not refreeze puree for re-use (as bacteria may bloom and harm the worms). 
	NOTE: Liver should contain no hormones or antibiotics and preferably not be previously frozen. Centrifuge puree prior to freezing (or prior to feeding) to remove air and prevent food from floating. Food should sink to the bottom of the container. 1 mL of puree is sufficient for 500-1,000 worms.</li>
	<li>Exchange water once per week with fresh worm water for both colonies and experiments. Colony containers should also be wiped down with an unbleached paper towel (to remove mucus residues that trap wastes) once a week for starving worms, or twice a week for feeding worms (2 h after feeding and again 2 days later). In order to retain the biofilm, do not wipe more than 80% of the container.</li>
	<li>Move worms between containers (or surgery setup) using a transfer pipette. To free worms adhered to container surfaces, squirt water over/behind the worm to release it from the surface. Then suck up the worm into the pipette, while trying to keep the worm within the bottom inch (thinner) portion of the pipette.
	<ol>
		<li>Backload pipettes with a small amount of worm water before sucking them up.</li>
		<li>Try to move the water around the worm, rather than the worm itself, to help prevent tearing soft-bodied worms by touching them with the pipette.</li>
		<li>Keep experimental dishes covered unless transferring worms or replacing water.</li>
	</ol>
	</li>
</ol>
2. Preparation
<ol>
	<li>Stop feeding worms at least one week prior to experiments.
	<ol>
		<li>Select worms that have been not been fed for &#8805;1 week and are at least 5-7 mm in length. Transfer the worms to a Petri dish 2/3 full of worm water, and confirm worm length by sliding a ruler underneath the dish. Measure worms while fully extended and moving. 
		NOTE: While smaller (5-7 mm) worms work better when performing subsequent immunofluorescence and in situ hybridization analyses, larger worms (8-10 mm) are easier to work with &#8211; especially when learning to ablate.</li>
		<li>Ensure that worms are whole, undamaged, and not recently regenerating. 
		NOTE: Regenerating worms will have much lighter (or no) pigment in the head and/or tail region as compared to normal worms.</li>
		<li>If performing RNAi or pharmacological treatments on worms, do so at this point. If worms were fed as part of the treatment (as for RNAi), wait at least 7 days after the last feeding before continuing to step 2.2.</li>
	</ol>
	</li>
	<li>Clean the work space and dissecting microscope base with 70% ethanol and allow it to dry completely. Place the dish of selected worms to the left of the scope. To the right of the scope, place a pair of #5 forceps, a 31 G 5/16-inch insulin needle with a 1 mL syringe, and a clean transfer pipette.
	<ol>
		<li>Ensure that the instruments are kept from touching the bench (which could introduce contaminants). Use a rigid item (such as a chopstick rest) to keep the forceps, needle, and pipette clean. Alternatively, place instruments on top of a fresh brown (unbleached) paper towel. 
		NOTE: These and subsequent instructions are written as they pertain to right-handed individuals. Left-handed individuals should reverse the right/left directions.</li>
	</ol>
	</li>
	<li>Next to the work space, place a box of lint-free tissue wipes, a source of fresh worm water, and some extra transfer pipettes (protected from the bench top). Place worm water in a plastic wash bottle for ease of dispensing. Also place a labeled 12-well plate (or 100 mm Petri dish) to the right of the scope for collecting ablated animals.</li>
	<li>If using a custom-made Peltier plate to immobilize worms, position the Peltier plate into the depression in the base of the dissecting microscope and adjust the output on the DC power source until the working surface of the Peltier plate is sufficiently cool (typically ~5 V).Alternatively, make a cold plate by filling a 100 mm Petri dish &#189; full with water and freeze for at least 24 h. Discard the lid and place the bottom of the 100 mm dish on the base of the dissecting scope, then place the lid of a 60 mm Petri dish upside down directly on the surface of the ice (Figure 1A).
	<ol>
		<li>After the water has frozen in the 100 mm dish, a &#34;volcano&#34; of ice can appear in the center of the plate. If this happens, use a razor blade to scrape the ice flat, to remove any irregularities from the surface. 
		NOTE: During surgeries the ice will melt, making ablations much more difficult. Prepare several ice dishes in advance, and replace frozen ones for melting ones during the assay as soon as the lid of the 60 mm dish starts floating.</li>
	</ol>
	</li>
	<li>Prepare the surgery surface. Cut a 5 cm x 10 cm piece of plastic paraffin film (4 cm2 if using the petri dish cold plate) and place it on the center of the immobilization device. Fold a lint-free tissue wipe into a square roughly 2 cm2 and place it on top of the film. Cut a piece of white filter paper 1.5-2 cm2 and place it on top of the wipe. See Figure 1B-1E.
	<ol>
		<li>Hold the folded wipe in place with a gloved finger, and use worm water to lightly dampen the wipe to ensure that it remains folded. Use the side of a transfer pipette to roll the wipe flat. 
		NOTE: Cold temperature helps to keep worms from moving. Working &#34;dry&#34; also helps in immobilization. The tissue wipe acts to wick water through the filter paper, keeping the worm moist during surgery without allowing worms to dry out completely (which is lethal).</li>
	</ol>
	</li>
</ol>
3. Surgical Ablation
<ol>
	<li>Use a transfer pipette to place one worm dorsal side up onto the filter paper. Turn the light source on and direct the goosenecks so that the light is focused on the worm. Adjust the dissecting scope focus and magnification so that the eyes are clearly visible (the entire worm does not need to be in view); 5X magnification is a good starting point. Orient the worm by rotating the paraffin film so that the head is pointed towards the researcher (towards the front of the microscope), angled 30-40 degrees to the right.
	<ol>
		<li>Avoid use of bright light to prevent excess worm movement. Planarians display negative phototaxis and will attempt to evade bright light.</li>
		<li>If the worm is positioned ventral side facing up (with pharyngeal opening visible), use the dull side of the forceps to gently reposition the worm dorsal side up (with eyes visible). Avoid approaching the animal with the sharp tip of the forceps to minimize the chance of tearing through soft tissues when repositioning the animal.</li>
		<li>Adjust the microscope focus so that the eyes are clearly in focus. Also ensure that both the eyes and surrounding head tissues are in view.</li>
	</ol>
	</li>
	<li>Hold a fresh needle/syringe in the right hand as though it were a pen (between thumb and index finger). Brace the left thumb against the Peltier plate (or Petri dish cold plate), and use the left thumb as a fulcrum on which the bottom portion of the syringe will rest (Figure 2A). Look through the microscope, and make sure the bevel of the needle is visible. 
	NOTE: Needles are very sharp. Be cautious when handling them. Wearing latex or nitrile gloves can offer some protection.
	<ol>
		<li>Use the left thumb to help hold the needle/syringe steady throughout the procedure and avoid shaking hands.</li>
		<li>Make an angle of about 40&#176; with the left thumb and left index finger (with the index finger on the surgery surface) to help hold the surgery surface stabile.</li>
	</ol>
	</li>
	<li>With the bevel of the needle facing upward (like a spoon), position the needle at right angles to the eye (Figure 2B). Use the tip of the needle to gently penetrate the thin layer of tissue overlying the optic cup (white, unpigmented region) of the eye, scooping from right to left. Very gently remove any pigmented tissue located within the optic cup, taking care not to puncture the bottom or destroy the borders of the optic cup.
	<ol>
		<li>If the worm has been on the filter paper for &#62;1-2 min at any point during the procedure, add one drop of worm water to rehydrate the worm. 
		NOTE: Dehydration will increase the worm&#39;s susceptibility to ripping injuries.</li>
	</ol>
	</li>
	<li>Repeat step 3.3 until all of the black pigmented tissues (pigment cells), as well as all of the white tissues of the optic cup (dendrites of the photoreceptor cells), are removed. Remove excised tissues stuck to the end of the needle by carefully wiping with a tissue wipe. If double eye ablation is desired, ablate the second eye at this point. 
	NOTE: To avoid injury, needles can be cleaned by grasping the needle with a clean tissue wipe held with the thumb and forefinger, then using the other hand to pull the needle through the wipe away from the thumb and forefinger. Alternatively, the needle can be wiped onto the surface of a wet tissue wipe and examined for cleanliness.</li>
	<li>When finished, use a transfer pipette to move the worm to a labeled dish containing fresh worm water. If analyzing group data, place up to 20 worms in a single 100 mm Petri dish. If tracking regeneration in the same individuals over time, place 1 worm into each well of a 12-well plate. Once all worms are transferred, rinse worms by removing all worm water from the dishes/wells and replacing with more fresh worm water.
	<ol>
		<li>To remove worms from the filter, backload the pipette with a small amount of worm water and release the water onto the worm. This should lift the worm off the filter paper, to be immediately sucked into the pipette for transfer.</li>
	</ol>
	</li>
	<li>Incubate experiments at ~20 &#176;C (room temperature) protected from light and follow the regenerative process. 
	NOTE: Worms can be stored in a temperature-controlled incubator to maintain a constant temperature, but this is not strictly necessary. Most room temperatures vary only by +5 &#176;C, which will not affect the worm&#39;s ability to regenerate.</li>
	<li>If desired, fix worms for immunohistochemistry (see Figures 3-4) and/or in situ hybridization analyses of gene expression. Different fixation methods exist for planarian immunohistochemistry18,19,20 and in situ hybridization21,22,23 protocols.</li>
	<li>When experiments are concluded, sacrifice live worms by removing worm water from the dish and replacing with 70% ethanol. Incubate worms for 3-5 min, checking for worms to lyse and turn grayish.</li>
	<li>Place non-treated worms in the normal waste stream once sacrificed. Avoid introducing live worms into the environment, particularly non-native species.</li>
</ol>

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A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Chapter+3.">Chapter 3.</a> Animal Models in Eye Research. , 15-26 (2008).</li><li>Lapan, S. W., Reddien, P. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=dlx+and+sp6-9+Control+optic+cup+regeneration+in+a+prototypic+eye.">dlx and sp6-9 Control optic cup regeneration in a prototypic eye.</a> PLoS Genet. 7 (8), e1002226 (2011).</li><li>Lapan, S. W., Reddien, P. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Transcriptome+analysis+of+the+planarian+eye+identifies+ovo+as+a+specific+regulator+of+eye+regeneration.">Transcriptome analysis of the planarian eye identifies ovo as a specific regulator of eye regeneration.</a> Cell Rep. 2 (2), 294-307 (2012).</li><li>Inoue, T., 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=Morphological+and+functional+recovery+of+the+planarian+photosensing+system+during+head+regeneration.">Morphological and functional recovery of the planarian photosensing system during head regeneration.</a> Zoolog Sci. 21 (3), 275-283 (2004).</li><li>Pineda, D., 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+genetic+network+of+prototypic+planarian+eye+regeneration+is+Pax6+independent.">The genetic network of prototypic planarian eye regeneration is Pax6 independent.</a> Development. 129 (6), 1423-1434 (2002).</li><li>Paskin, T. R., Jellies, J., Bacher, J., Beane, W. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Planarian+Phototactic+Assay+Reveals+Differential+Behavioral+Responses+Based+on+Wavelength.">Planarian Phototactic Assay Reveals Differential Behavioral Responses Based on Wavelength.</a> PLoS One. 9 (12), e114708 (2014).</li><li>Raffa, R. B., Martley, A. F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Amphetamine-induced+increase+in+planarian+locomotor+activity+and+block+by+UV+light.">Amphetamine-induced increase in planarian locomotor activity and block by UV light.</a> Brain Res. 1031 (1), 138-140 (2005).</li><li>Sandmann, T., Vogg, M. C., Owlarn, S., Boutros, M., Bartscherer, K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+head-regeneration+transcriptome+of+the+planarian+Schmidtea+mediterranea.">The head-regeneration transcriptome of the planarian Schmidtea mediterranea.</a> Genome Biol. 12 (8), R76 (2011).</li><li>Vasquez-Doorman, C., Petersen, C. P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+NuRD+complex+component+p66+suppresses+photoreceptor+neuron+regeneration+in+planarians.">The NuRD complex component p66 suppresses photoreceptor neuron regeneration in planarians.</a> Regeneration (Oxf). 3 (3), 168-178 (2016).</li><li>Deochand, M. E., Birkholz, T. R., Beane, W. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Temporal+regulation+of+planarian+eye+regeneration.">Temporal regulation of planarian eye regeneration.</a> Regeneration. 3 (4), 209-221 (2016).</li><li>Sakai, F., Agata, K., Orii, H., Watanabe, K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Organization+and+regeneration+ability+of+spontaneous+supernumerary+eyes+in+planarians+-eye+regeneration+field+and+pathway+selection+by+optic+nerves.">Organization and regeneration ability of spontaneous supernumerary eyes in planarians -eye regeneration field and pathway selection by optic nerves.</a> Zoolog Sci. 17 (3), 375-381 (2000).</li><li>Asano, Y., Nakamura, S., Ishida, S., Azuma, K., Shinozawa, T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Rhodopsin-like+proteins+in+planarian+eye+and+auricle:+detection+and+functional+analysis.">Rhodopsin-like proteins in planarian eye and auricle: detection and functional analysis.</a> J Exp Biol. 201 (Pt 9), 1263-1271 (1998).</li><li>Cross, S. 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A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Generation+of+cell+type-specific+monoclonal+antibodies+for+the+planarian+and+optimization+of+sample+processing+for+immunolabeling.">Generation of cell type-specific monoclonal antibodies for the planarian and optimization of sample processing for immunolabeling.</a> BMC Dev Biol. 14, 45 (2014).</li><li>Ross, K. G., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Novel+monoclonal+antibodies+to+study+tissue+regeneration+in+planarians.">Novel monoclonal antibodies to study tissue regeneration in planarians.</a> BMC Dev Biol. 15, 2 (2015).</li><li>Cardona, A., Fernandez, J., Solana, J., Romero, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=An+in+situ+hybridization+protocol+for+planarian+embryos:+monitoring+myosin+heavy+chain+gene+expression.">An in situ hybridization protocol for planarian embryos: monitoring myosin heavy chain gene expression.</a> Dev Genes Evol. 215 (9), 482-488 (2005).</li><li>King, R. S., Newmark, P. 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The authors have nothing to disclose.

This eye ablation technique improves on the current methods (such as hole punch) by excluding brain tissues and restricting excision mainly to the eye tissues. With practice, this technique can be performed by most individuals, from technicians experienced in microsurgeries to inexperienced but conscientious undergraduate students. It is recommended that this technique be practiced many times prior to using ablations in experiments, including (when possible) confirmation of complete removal of all eye tissues by immunohistochemistry or in situ hybridization for eye marker(s)4,5,13. The most critical step in this protocol is the scooping out of tissues. It is important to not remove either too much or too little tissue. This can be avoided by not scooping too deeply with the needle, to avoid penetrating through to the ventral side of the worm. The tip of the needle should only be inserted about 0.4 mm deep, and the beveled part of the needle should never go all the way through the eye. Special care should be taken to not damage the fragile tissue between the two eyes (the medial edge of the black pigmented region of each eye represents the borders of each eye). Damage between the eye and the distal margin of the worm should also be avoided. Smaller worms are more likely to rip or be unintentionally injured, so the use of larger worms (particularly for practice) is recommended. The main limitations of this technique are that it is relatively time intensive (and not appropriate for high-throughput screens), and it requires an initial investment in practicing to ensure the accuracy of the results. However, with practice 10 eyes can be ablated in ~15 min.
A major area for troubleshooting is in keeping planarians still during the procedure. The two main reasons worms are not immobilized sufficiently to perform ablations are (a) too much water and (b) loss of cold temperature. (A) Excess planarian water that pools on the surgery surface while transferring worms will allow worms to move and complicate the surgery. The surgery surface (lint-free tissue wipe and filter paper) should remain moist only. If at any point during the procedure the wipe becomes saturated with worm water, it should either be replaced or a transfer pipette can be used to gently remove excess fluid from the tissue wipe (always draw from the wipe and not the filter paper). Tissue wipes can also be used to remove excess water pooling on the surgery surface. This process may need to be repeated many times, depending on the number of worms being ablated. (B) If using the petri dish cold plate set up, remember to replace the cold plate as soon as the ice starts melting and the 60 mm Petri dish lid begins to float. This will ensure both a stable surgery surface and an appropriately cold surface. Alternatively, if using the Peltier plate set up, be aware that after 45 min to 1 h of use, the heat in the outer rings will overcome the cold plate in the center and all cooling will be lost. If this happens, turn off the Peltier plate for 5-10 min to allow it to come to room temperature before resuming ablations.
Another area of troubleshooting is during worm transfer between the dish and the surgery surface and back again. If ablated worms are hard to remove from the filter paper, place a drop of worm water on the worm until the liquid pools on top of the filter paper. Worms can then be sucked into the transfer pipette as usual. If problems persist, try flipping the worm onto its ventral side first (using the dull side of the forceps). If a worm gets stuck inside the transfer pipette, this is usually a sign that too much water has been taken up (worms should be drawn into the transfer pipette no farther than about an inch). To remove a worm stuck in a transfer pipette, draw up 1-2 mL of worm water into the pipette, so that the worm is covered with water. Firmly flick the side of the pipette until the worm detaches from the wall of the pipette and is floating.
If the needle becomes dull, replace with a new needle/syringe. Consider replacing the needle after using it to ablate &#8805;30 eyes. Excessive removal of resected tissues by wiping with a tissue wipe can also dull the needle, making ablations more difficult. It is okay if resected tissues stay within the beveled part of the needle throughout the procedure (even across multiple animals). Always use a fresh needle/syringe when changing between worms of differing conditions (e.g. wildtype vs. RNAi worms), as well as when beginning new ablation sessions (i.e. on different days).
This surgical eye ablation technique is a powerful means to investigate light-induced behaviors (and their genetic and neurological basis) in planarians, particularly when combined with pharmacological treatments or RNA interference. Eye ablation is also a great means by which to elucidate the mechanisms that regulate endogenous planarian eye regeneration (and eye stem cell maintenance and differentiation) in vivo. As most vertebrates have very limited abilities to regenerate eye tissues, understanding how planarians are able to regenerate their eyes will be important in identifying possible mechanisms for translation into future therapies.

Planarians are a powerful model organism for studying adult stem cell-mediated regeneration. These non-parasitic freshwater flatworms possess the ability to regenerate any and all missing tissues, including their central nervous system and brain1. Studied as far back as the 1700s2, technological advances in the planarian field during the past 10-15 years (such as a sequenced genome, in situ hybridization, immunohistochemistry, RNA interference (RNAi), and transcriptomics) have updated this historical model organism. Specifically, planarians have recently gained popularity as an emerging model for eye research3.
Planarians have prototypic eyes with only two tissue types, the photoreceptor neurons and pigment cells; this has enabled the characterization of its eye stem-cell population and demonstrated that many of the same genes regulating vertebrate eye development are conserved in planarians4,5. The optic cups are located dorsally and comprised of the white, unpigmented dendrites of the photoreceptor neurons and the semi-lunar black pigment cells, and the eyes innervate the brain via an optic chiasm. In addition to being a model for elucidating regenerative processes6, the planarian eye is well suited to studying the evolution of visual mechanisms7, behavioral responses to light (planarians display negative phototaxis)8, and the neurological basis of behavior9.
Eye regeneration in planarians has been largely studied in two main contexts: as part of head regeneration following decapitation4,10, and following excision of just the eye tissues11,12. Most planarian studies on eye regeneration have used the decapitation method, as it is simple and straightforward. The most common planarian eye excision method to date has been via hole punch with a fine glass capillary tube13,14, although some studies have also performed amputations just behind the eyes (partial decapitation)15. However, all of these methods involve the loss of many tissues other than just the eye (such as brain, intestines, and nephridia), potentially complicating interpretation of results. The eye ablation protocol presented here restricts excision to the eye tissues (specifically excluding brain), resulting in data that are more specific to the eye. Additionally, unlike decapitated worms that take 7-14 days to begin feeding, eye ablated worms will feed within 24 h of ablation12, allowing RNAi experiments (where RNAi is delivered via food) to be performed concurrently.
Although eye ablation is technically harder to successfully perform than decapitation, current studies involving eye excision have not included detailed instructions on their procedures. The goal of this video article is to enable researchers to consistently remove the planarian optic cup without disturbing the underlying brain tissues and removing as few other tissues as possible. This method can be used for both single and double eye ablation and is applicable to a wide range of investigations. Like most regeneration assays, eye ablation is well suited for combination with both pharmacological and genetic (RNAi) screens, as well as behavioral studies. Here we describe methods for the handling of worms, maintaining a planarian colony, and the eye ablation technique itself.

<table><tbody><tr><td>Instant Ocean sea salts</td><td>Spectrum Brands&amp;nbsp;</td><td>SS15-10</td><td>&quot;10 Gallon&quot; box&amp;nbsp; (net weight 3 lbs)</td></tr><tr><td>Kimwipes EX-L lint-free tissue wipe</td><td>Kimberly-Clark</td><td>34155</td><td>4.5 x 8.5 in&amp;nbsp;</td></tr><tr><td>Whatman #2 filter paper</td><td>Sigma&amp;nbsp;</td><td>WHA1002125</td><td>Circles, 125 mm diameter, white</td></tr><tr><td>Easy Touch Insulin syringe (with needle)</td><td>Pet Health Market</td><td>17175-04</td><td>&amp;nbsp;U-100 1 cc syringe, 31 G 5/16 in needle</td></tr><tr><td>100 mm Petri dish</td><td>VWR</td><td>25384-342</td><td>100 mm x 15 mm</td></tr><tr><td>60 mm Petri dish</td><td>VWR</td><td>25384-092</td><td>60 mm x 15 mm</td></tr><tr><td>Dumont #5&amp;nbsp; forceps&amp;nbsp;</td><td>Fine Science Tools</td><td>11254-20</td><td>Inox, straight tip , 11 cm</td></tr><tr><td>Transfer pipettes&amp;nbsp;</td><td>Samco Scientific&amp;nbsp;</td><td>225</td><td>Graduated, large bulb, 7.5 mL, non sterile</td></tr><tr><td>Parafilm M paraffin film</td><td>Brand&amp;nbsp;</td><td>701606</td><td>&amp;nbsp;4 in x 125 ft roll</td></tr><tr><td>12-well untreated tissue culture plate</td><td>VWR</td><td>15705-059</td><td>Untreated, flat bottom, sterile, Falcon brand</td></tr><tr><td>Plastic food containers (for colony)&amp;nbsp;</td><td>Ziploc</td><td>Large rectangle</td><td>2.25 qt (2.12 L), 10&quot; x 6 -3/4 &quot; x 3 -3/16&quot;&amp;nbsp;</td></tr><tr><td>Planaria (&lt;em&gt;Girardia tigrina&lt;/em&gt;)</td><td>Carolina Biological</td><td>132954</td><td>Sold as &quot;Brown&quot; Planaria; most often they are &lt;em&gt;G. tigrina&lt;/em&gt; (aka &lt;em&gt;Dugesia tigrina&lt;/em&gt;), but sometimes are &lt;em&gt;G. dorotocephala&lt;/em&gt; (aka &lt;em&gt;Dugesia dorotocephala&lt;/em&gt;); either will work.</td></tr><tr><td>Planaria (&lt;em&gt;Schmidtea mediterranea&lt;/em&gt;)</td><td>n/a</td><td>n/a</td><td>&lt;em&gt;S. mediterranea&lt;/em&gt; are not commercially available. At this time animals are only obtainable from laboratories that use them and have extra animals.</td></tr><tr><td>Brown paper towels&amp;nbsp;</td><td>Grainger</td><td>2U229</td><td>9-3/16 x 9-3/8&quot; 1-Ply Multifold Paper Towel, UNBLEACHED</td></tr><tr><td>Wash bottle (for worm water), optional</td><td>VWR</td><td>16650-275</td><td>Wash Bottles, Low-Density Polyethylene, Wide Mouth, 500 mL</td></tr><tr><td>Anti-synapsin antibody, optional</td><td>Developmental Studies Hybridoma Bank</td><td>3C11</td><td>Supernatant</td></tr><tr><td>Anti-arrestin antibody, optional</td><td>n/a</td><td>n/a</td><td>Not commercially available. Kind gift from Michael Levin, Tufts University</td></tr><tr><td>Nalgene Lowboy carboy with spigot (for storing worm water), optional</td><td>Nalge Nunc International Corporation</td><td>2324-0015</td><td>15 L,&amp;nbsp; polypropylene, low profile makes it easier to fill plastic colony containers&amp;nbsp;</td></tr><tr><td>Custom Peltier plate, optional</td><td>Williams Machine, Foxboro, MA&amp;nbsp;</td><td>n/a</td><td>Design specifics courtesy of Junji Morokuma, Tufts University:&amp;nbsp; Peltier plate is constructed of a standard thermoelectric heat pump (for example, All Electronics Corp Catalog # PJT-1, 30 mm&lt;sup&gt;2&lt;/sup&gt;).&amp;nbsp; The square heat pump is covered with a thin mirrored surface, then placed inside a 30 mm&lt;sup&gt;2&lt;/sup&gt; square hole in a circular plexiglass form (~50 mm in diameter). This form is of similar thickness to the heat pump, and fits flush into a well tooled in the center of a round heat sink (~115 mm in diameter). The form/heat pump is&amp;nbsp; &quot;anchored&quot; to the sink with silicone base heat sink compound. The leads are threaded through holes drilled through both the form and the the heat sink. The bottom half of the heat sink is tooled into a &quot;foot&quot; that fits into the opening of your microscope&#039;s base plate.&amp;nbsp;</td></tr><tr><td>DC power source (for Peltier plate), optional</td><td>B &amp;amp; K Precision</td><td>1665</td><td>Regulated Low Voltage DC Power Supply, 1-18 V (DC), 1-10 amps.</td></tr><tr><td>Other common supplies</td><td></td><td></td><td></td></tr><tr><td>Gloves</td><td></td><td></td><td></td></tr><tr><td>Razor blade&amp;nbsp;</td><td></td><td></td><td></td></tr><tr><td>Scissors</td><td></td><td></td><td></td></tr><tr><td>Dissecting scope with gooseneck lighting</td><td></td><td></td><td></td></tr><tr><td>Chopstick rests, optional</td><td></td><td></td><td></td></tr></tbody></table>

surgical ablation assay for studying eye regeneration in planarians

In the study of adult stem cells and regenerative mechanisms, planarian flatworms are a staple in vivo model system. This is due in large part to their abundant pluripotent stem cell population and ability to regenerate all cell and tissue types after injuries that would be catastrophic for most animals. Recently, planarians have gained popularity as a model for eye regeneration. Their ability to regenerate the entire eye (comprised of two tissue types: pigment cells and photoreceptors) allows for the dissection of the mechanisms regulating visual system regeneration. Eye ablation has several advantages over other techniques (such as decapitation or hole punch) for examining eye-specific pathways and mechanisms, the most important of which is that regeneration is largely restricted to eye tissues alone. The purpose of this video article is to demonstrate how to reliably remove the planarian optic cup without disturbing the brain or surrounding tissues. The handling of worms and maintenance of an established colony is also described. This technique uses a 31 G, 5/16-inch insulin needle to surgically scoop out the optic cup of planarians immobilized on a cold plate. This method encompasses both single and double eye ablation, with eyes regenerating within 1-2 weeks, allowing for a wide range of applications. In particular, this ablation technique can be easily combined with pharmacological and genetic (RNA interference) screens for a better understanding of regenerative mechanisms and their evolution, eye stem cells and their maintenance, and phototaxic behavioral responses and their neurological basis.

For the first 1-2 h post surgery, animals may exhibit decreased movement compared to intact worms (however they will still move). If desired, worms will eat within 24 h of surgery (for instance, for RNAi feeding). When following eye regeneration in the same individuals over time, make sure to take a photograph of each worm both prior to surgery (intact) and at 1 h post ablation (hpa). By 4 days post ablation (dpa) regenerating pigment cells should be visible, and by 14 dpa the entire eye will have fully regenerated (Figure 3A-B). This includes the photoreceptor neurons and their innervation to the brain (Figure 3C), as well as functional recovery of the visual system (Figure 4A-C). Successful ablations will not disturb the underlying tissues of the brain (Figure 4D-E), nor introduce other injuries to the animal (Figure 5A). Unsuccessful ablations include animals with: an overly large excision that connects the two eyes (Figure 5B), tears to the lateral margin of the animal (Figure 5C), and/or wound sites that poke through the ventral epidermis (Figure 5D). Furthermore, unsuccessful ablations include incomplete removal of the entire optic cup, comprised of both the white unpigmented tissues and the black pigment cells (Figure 5E-F).
<img alt="Figure 1" src="/files/ftp_upload/55594/55594fig1.jpg" /> 
Figure 1: Surgery preparation. (A) Diagram of cold plate construction, using the bottom of a 100 mm Petri dish (filled with ice) and the lid of a 60 mm Petri dish (placed upside down on the ice surface). (B) Diagram of the surgery surface, made from a stack of (top to bottom) white filter paper, a damp folded tissue wipe, and a piece of paraffin film. (C) The surgery set up (for the right-handed). A: dissecting scope, B: gooseneck lighting, C: surgery surface, D: Peltier plate, E: dish of 5-7 mm worms ready for ablation, F: wash bottle of worm water, G: chopstick rest holding clean transfer pipette, H: chopstick rest holding needle/syringe, I: chopstick rest holding forceps, J: container of extra transfer pipettes. (D) Custom Peltier plate configuration. (E) Petri dish cold plate configuration. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/55594/55594fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" src="/files/ftp_upload/55594/55594fig2.jpg" /> 
Figure 2: Position of hands and needle/syringe during ablation. (A) Placement of hands (for the right-handed). Note that the left thumb is used to support the syringe (held in the right hand) to minimize movement. (B) Placement of the needle in relation to the worm&#39;s eye. Note that the beveled surface of the needle faces upwards. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/55594/55594fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" src="/files/ftp_upload/55594/55594fig3.jpg" /> 
Figure 3: Ablated planarian eyes regenerate. Morphology of Schmidtea mediterranea planarians regrowing eyes following (A) double eye ablation and (B) single eye ablation. (C) Immunohistochemistry showing regeneration of the photoreceptor neurons (anti-arrestin) following single eye ablation. Intact worms are shown before surgery, and regenerates are shown at days 4 and 14 after ablation. For single eye ablations, the left eye was ablated and the right eye serves as an internal (uninjured) control. Red arrowheads: ablated eyes. Scale bars = 100 &#181;m. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/55594/55594fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" src="/files/ftp_upload/55594/55594fig4.jpg" /> 
Figure 4: Functional recovery of the visual system after ablation. (A-C) Functional assay to test planarian photophobia. (A) Intact worms avoid traveling through areas of light, such as a spot from a green laser pointer. (B) At 24 h post ablation, &#34;blind&#34; double eye ablated worms travel through light spots. (C) At 7 days post ablation, double eye ablated worms have regenerated their visual system sufficiently to recover light avoidance. Yellow arrowhead: aberrant behavioral response. (D-E) Eye ablation is restricted to the tissues of the optic cup, and does not disturb the underlying brain tissues. (D) Immunohistochemistry showing brain architecture (anti-synapsin) is unchanged from before ablation to 1 h post ablation. (E) Hematoxylin and eosin (H&#38;E) staining at 1 h post ablation in a transverse section showing damage is largely restricted to the site of the ablated optic cup. Right eye (with black pigment cells) serves as internal control. Red arrowheads: ablated eyes. Scale bars = 1 mm in (A-C) 1 mm, 100 &#181;m in (D-E). <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/55594/55594fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" src="/files/ftp_upload/55594/55594fig5.jpg" /> 
Figure 5: Hallmarks of successful and unsuccessful ablations. (A) Successful single and double eye ablations (at 5 min post ablation) have wounds that are roughly similar in size to the original optic cup. (B-E) Unsuccessful ablations: (B) too much tissue is removed or the wound connects the two eyes, (C) the wound site tears and extends to the margin, (D) the wound is too deep and pokes through from the dorsal (D) to the ventral (D&#39;) side, and (E-F) not all of the optic cup tissues are removed, visualized both morphologically (E) and by anti-arrestin staining of the photoreceptor neurons (F). Yellow arrowheads: aberrant ablations. Yellow circles: ablation site. Scale bars = 100 &#181;m. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/55594/55594fig5large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Surgical Ablation Assay for Studying Eye Regeneration in Planarians at JoVE.com

Surgical Ablation Assay for Studying Eye Regeneration in Planarians

This protocol shows how to consistently excise planarian eyes (optic cups) without disturbing surrounding tissues. Using an insulin needle and syringe, either one or both eyes can be ablated to facilitate investigations into the mechanisms regulating eye regeneration, the evolution of visual regeneration, and the neural basis of light-induced behavior.

In the study of adult stem cells and regenerative mechanisms, planarian flatworms are a staple in vivo model system. This is due in large part to their ...

surgical-ablation-assay-for-studying-eye-regeneration-in-planarians

Nanyang Technological University

Research

JoVE Journal

Developmental Biology

10.9K Views.  Western Michigan University. This protocol shows how to consistently excise planarian eyes (optic cups) without disturbing surrounding tissues. Using an insulin needle and syringe, either one or both eyes can be ablated to facilitate investigations into the mechanisms regulating eye regeneration, the evolution of visual regeneration, and the neural basis of light-induced behavior.

Video: Surgical Ablation Assay for Studying Eye Regeneration in Planarians

Chemical Amputation and Regeneration of the Pharynx in the Planarian Schmidtea mediterranea

Planarians are flatworms that are extremely efficient at regeneration. They owe this ability to a large number of stem cells that can rapidly respond to any type of injury. Common injury models in these animals remove large amounts of tissue, which damages multiple organs. To overcome this broad tissue damage, we describe here a method to selectively remove a single organ, the pharynx, in the planarian Schmidtea mediterranea. We achieve this by soaking animals in a solution containing the cytochrome oxidase inhibitor sodium azide. Brief exposure to sodium azide causes extrusion of the pharynx from the animal, which we call &#34;chemical amputation.&#34;&#160;Chemical amputation removes the entire pharynx, and generates a small wound where the pharynx attaches to the intestine. After extensive rinsing, all amputated animals regenerate a fully functional pharynx in approximately one week. Stem cells in the rest of the body drive regeneration of the new pharynx. Here, we provide a detailed protocol for chemical amputation, and describe both histological and behavioral methods to assess successful amputation and regeneration.

Chemical Amputation and Regeneration of the Pharynx in the Planarian Schmidtea mediterranea

The planarian Schmidtea mediterranea is an excellent model for studying stem cells and tissue regeneration. This publication describes a method to selectively remove one organ, the pharynx, by exposing animals to the chemical sodium azide. This protocol also outlines methods for monitoring pharynx regeneration.

Planarians are flatworms that are extremely efficient at regeneration. They owe this ability to a large number of stem cells that can rapidly respond to ...

A Planarian Motility Assay to Gauge the Biomodulating Properties of Natural Products

A straightforward, controllable means of using the non-parasitic planarian, Dugesia tigrina, a free-living aquatic flatworm, to study the stimulant and withdrawal properties of natural products is described. Experimental assays benefitting from unique aspects of planarian physiology have been applied to studies on wound healing, regeneration, and tumorigenesis. In addition, because planarians exhibit sensitivity to a variety of environmental stimuli and are capable of learning and developing conditioned responses, they can be used in behavioral studies examining learning and memory. Planarians possess a basic bilateral symmetry and a central nervous system that uses neurotransmitter systems amenable to studies examining the effects of neuromuscular biomodulators. Consequently, experimental systems monitoring planarian movement and motility have been developed to examine substance addiction and withdrawal. Because planarian motility offers the potential for a sensitive, easily standardized motility assay system to monitor the effect of stimuli, the planarian locomotor velocity (pLmV) test was adapted to monitor both stimulation and withdrawal behaviors by planarians through the determination of the number of grid lines crossed by the animals with time. Here, the technique and its application are demonstrated and explained.

Planarian motility is used to gauge the stimulant and withdrawal properties of natural products when compared to the movement of the animals in spring water alone.

A straightforward, controllable means of using the non-parasitic planarian, Dugesia tigrina, a free-living aquatic flatworm, to study the stimulant and ...

Biochemistry

Confocal Microscope-Based Laser Ablation and Regeneration Assay in Zebrafish Interneuromast Cells

Hair cells are mechanosensory cells that mediate the sense of hearing. These cells do not regenerate after damage in humans, but they are naturally replenished in non-mammalian vertebrates such as zebrafish. The zebrafish lateral line system is a useful model for characterizing sensory hair cell regeneration. The lateral line is comprised of hair cell-containing organs called neuromasts, which are linked together by a string of interneuromast cells (INMCs). INMCs act as progenitor cells that give rise to new neuromasts during development. INMCs can repair gaps in the lateral line system created by&#160;cell death. A method is described here for selective INMC ablation using a conventional laser-scanning confocal microscope and transgenic fish that express green fluorescent protein in INMCs. Time-lapse microscopy is then used to monitor INMC regeneration and determine the rate of gap closure. This represents an accessible protocol for cell ablation that does not require specialized equipment, such as a high-powered pulsed ultraviolet laser. The ablation protocol may serve broader interests, as it could be useful for the ablation of additional cell types, employing a tool set that is already available to many users. This technique will further enable the characterization of INMC regeneration under different conditions and from different genetic backgrounds, which will advance the understanding of sensory progenitor cell regeneration.

Laser ablation is a widely applicable technology for studying regeneration in biological systems. The presented protocol describes use of a standard laser-scanning confocal microscope for laser ablation and subsequent time-lapse imaging of regenerating interneuromast cells in the zebrafish lateral line.

Hair cells are mechanosensory cells that mediate the sense of hearing. These cells do not regenerate after damage in humans, but they are naturally ...

Eye Removal in Living Zebrafish Larvae to Examine Innervation-dependent Growth and Development of the Visual System

Zebrafish exhibit remarkable life-long growth and regenerative abilities. For example, specialized stem cell niches established during embryogenesis support continuous growth of the entire visual system, both in the eye and the brain. Coordinated growth between the retinae and the optic tectum ensures accurate retinotopic mapping as new neurons are added in the eyes and brain. To address whether retinal axons provide crucial information for regulating tectal stem and progenitor cell behaviors such as survival, proliferation, and/or differentiation, it is necessary to be able to compare innervated and denervated tectal lobes within the same animal and across animals. 
Surgical removal of one eye from living larval zebrafish followed by observation of the optic tectum achieves this goal. The accompanying video demonstrates how to anesthetize larvae, electrolytically sharpen tungsten needles, and use them to remove one eye. It next shows how to dissect brains from fixed zebrafish larvae. Finally, the video provides an overview of the protocol for immunohistochemistry and a demonstration of how to mount stained embryos in low-melting-point agarose for microscopy.

The article explains how to surgically remove eyes from living zebrafish larvae as the first step toward investigating how retinal input influences optic tectum growth and development. In addition, the article provides information about larval anesthetization, fixation, and brain dissection, followed by immunohistochemistry and confocal imaging.

Zebrafish exhibit remarkable life-long growth and regenerative abilities. For example, specialized stem cell niches established during embryogenesis ...

Laser Ablation of the Zebrafish Pronephros to Study Renal Epithelial Regeneration

Acute kidney injury (AKI) is characterized by high mortality rates from deterioration of renal function over a period of hours or days that culminates in renal failure1. AKI can be caused by a number of factors including ischemia, drug-based toxicity, or obstructive injury1. This results in an inability to maintain fluid and electrolyte homeostasis. While AKI has been observed for decades, effective clinical therapies have yet to be developed. Intriguingly, some patients with AKI recover renal functions over time, a mysterious phenomenon that has been only rudimentally characterized1,2. Research using mammalian models of AKI has shown that ischemic or nephrotoxin-injured kidneys experience epithelial cell death in nephron tubules1,2, the functional units of the kidney that are made up of a series of specialized regions (segments) of epithelial cell types3. Within nephrons, epithelial cell death is highest in proximal tubule cells. There is evidence that suggests cell destruction is followed by dedifferentiation, proliferation, and migration of surrounding epithelial cells, which can regenerate the nephron entirely1,2. However, there are many unanswered questions about the mechanisms of renal epithelial regeneration, ranging from the signals that modulate these events to reasons for the wide variation of abilities among humans to regenerate injured kidneys.
The larval zebrafish provides an excellent model to study kidney epithelial regeneration as its pronephric kidney is comprised of nephrons that are conserved with higher vertebrates including mammals4,5. The nephrons of zebrafish larvae can be visualized with fluorescence techniques because of the relative transparency of the young zebrafish6. This provides a unique opportunity to image cell and molecular changes in real-time, in contrast to mammalian models where nephrons are inaccessible because the kidneys are structurally complex systems internalized within the animal. Recent studies have employed the aminoglycoside gentamicin as a toxic causative agent for study of AKI and subsequent renal failure: gentamicin and other antibiotics have been shown to cause AKI in humans, and researchers have formulated methods to use this agent to trigger kidney damage in zebrafish7,8. However, the effects of aminoglycoside toxicity in zebrafish larvae are catastrophic and lethal, which presents a difficulty when studying epithelial regeneration and function over time. Our method presents the use of targeted cell ablation as a novel tool for the study of epithelial injury in zebrafish. Laser ablation gives researchers the ability to induce cell death in a limited population of cells. Varying areas of cells can be targeted based on morphological location, function, or even expression of a particular cellular phenotype. Thus, laser ablation will increase the specificity of what researchers can study, and can be a powerful new approach to shed light on the mechanisms of renal epithelial regeneration. This protocol can be broadly applied to target cell populations in other organs in the zebrafish embryo to study injury and regeneration in any number of contexts of interest.

Acute kidney injury (AKI) in humans is a common clinical problem caused by damage to the epithelial cells that comprise kidney nephrons, and AKI is associated with high mortality rates of 50-70%1. Following epithelial cell destruction, nephrons have a limited ability to regenerate, though the mechanisms and limitations that guide this phenomenon remain poorly understood. In this video article, we describe our technique for targeted laser ablation of kidney nephron cells in the zebrafish embryo kidney, or pronephros. Our new method can be used to complement nephrotoxicity-induced models of AKI and gain a high-resolution understanding of the cell and molecular alterations that are associated with epithelial regeneration in the kidney nephron.

Acute kidney injury (AKI) is characterized by high mortality rates from deterioration of renal function over a period of hours or days that culminates in ...

Biology

Pharmacological and Functional Genetic Assays to Manipulate Regeneration of the Planarian Dugesia japonica

Free-living planarian flatworms have a long history of experimental usage owing to their remarkable regenerative abilities1. Small fragments excised from these animals reform the original body plan following regeneration of missing body structures. For example if a 'trunk' fragment is cut from an intact worm, a new 'head' will regenerate anteriorly and a 'tail' will regenerate posteriorly restoring the original 'head-to-tail' polarity of body structures prior to amputation (Figure 1A).
Regeneration is driven by planarian stem cells, known as 'neoblasts' which differentiate into ~30 different cell types during normal body homeostasis and enforced tissue regeneration. This regenerative process is robust and easy to demonstrate. Owing to the dedication of several pioneering labs, many tools and functional genetic methods have now been optimized for this model system. Consequently, considerable recent progress has been made in understanding and manipulating the molecular events underpinning planarian developmental plasticity2-9.
The planarian model system will be of interest to a broad range of scientists. For neuroscientists, the model affords the opportunity to study the regeneration of an entire nervous system, rather than simply the regrowth/repair of single nerve cell process that typically are the focus of study in many established models. Planarians express a plethora of neurotransmitters10, represent an important system for studying evolution of the central nervous system11, 12 and have behavioral screening potential13, 14. 
Regenerative outcomes are amenable to manipulation by pharmacological and genetic apparoaches. For example, drugs can be screened for effects on regeneration simply by placing body fragments in drug-containing solutions at different time points after amputation. The role of individual genes can be studied using knockdown methods (in vivo RNAi), which can be achieved either through cycles of microinjection or by feeding bacterially-expressed dsRNA constructs8, 9, 15. Both approaches can produce visually striking phenotypes at high penetrance- for example, regeneration of bipolar animals16-21. To facilitate adoption of this model and implementation of such methods, we showcase in this video article protocols for pharmacological and genetic assays (in vivo RNAi by feeding) using the planarian Dugesia japonica.

Pharmacological and Functional Genetic Assays to Manipulate Regeneration of the Planarian Dugesia japonica

An attractive model for studying stem cell differentiation within a live animal is the planarian flatworm. Regeneration is studied by simple amputation experiments that are easily performed in a basic laboratory and are amenable to pharmacological and genetic (in vivo RNAi) manipulation as detailed by protocols in this article.

Free-living planarian flatworms have a long history of experimental usage owing to their remarkable regenerative abilities1. Small fragments excised from ...

M&#252;ller Glia Cell Activation in a Laser-induced Retinal Degeneration and Regeneration Model in Zebrafish

A fascinating difference between teleost and mammals is the lifelong potential of the teleost retina for retinal neurogenesis and regeneration after severe damage. Investigating the regeneration pathways in zebrafish might bring new insights to develop innovative strategies for the treatment of retinal degenerative diseases in mammals. Herein, we focused on the induction of a focal lesion to the outer retina in adult zebrafish by means of a 532 nm diode laser. A localized injury allows investigating biological processes that take place during retinal degeneration and regeneration directly at the area of damage. Using non-invasive optical coherence tomography (OCT), we were able to define the location of the damaged area and monitor subsequent regeneration in vivo. Indeed, OCT imaging produces high-resolution, cross-sectional images of the zebrafish retina, providing information which was previously only available with histological analyses. In order to confirm the data from real-time OCT, histological sections were performed and regenerative response after the induction of the retinal injury was investigated by immunohistochemistry.

The zebrafish is a popular animal model to study mechanisms of retinal degeneration/regeneration in vertebrates. This protocol describes a method to induce localized injury disrupting the outer retina with minimal damage to the inner retina. Subsequently, we monitor in vivo the retinal morphology and the M&#252;ller glia response throughout retinal regeneration.

A fascinating difference between teleost and mammals is the lifelong potential of the teleost retina for retinal neurogenesis and regeneration after ...

Neuroscience

Planarian Immobilization, Partial Irradiation, and Tissue Transplantation

The planarian, a freshwater flatworm, has proven to be a powerful system for dissecting metazoan regeneration and stem cell biology1,2. Planarian regeneration of any missing or damaged tissues is made possible by adult stem cells termed neoblasts3. Although these stem cells have been definitively shown to be pluripotent and singularly capable of reconstituting an entire animal4, the heterogeneity within the stem cell population and the dynamics of their cellular behaviors remain largely unresolved. Due to the large number and wide distribution of stem cells throughout the planarian body plan, advanced methods for manipulating subpopulations of stem cells for molecular and functional study in vivo are needed.
Tissue transplantation and partial irradiation are two methods by which a subpopulation of planarian stem cells can be isolated for further study. Each technique has distinct advantages. Tissue transplantation allows for the introduction of stem cells, into a na&iuml;ve host, that are either inherently genetically distinct or have been previously treated pharmacologically. Alternatively, partial irradiation allows for the isolation of stem cells within a host, juxtaposed to tissue devoid of stem cells, without the introduction of a wound or any breech in tissue integrity. Using these two methods, one can investigate the cell autonomous and non-autonomous factors that control stem cell functions, such as proliferation, differentiation, and migration.
Both tissue transplantation5,6 and partial irradiation7 have been used historically in defining many of the questions about planarian regeneration that remain under study today. However, these techniques have remained underused due to the laborious and inconsistent nature of previous methods. The protocols presented here represent a large step forward in decreasing the time and effort necessary to reproducibly generate large numbers of grafted or partially irradiated animals with efficacies approaching 100 percent. We cover the culture of large animals, immobilization, preparation for partial irradiation, tissue transplantation, and the optimization of animal recovery. Furthermore, the work described here demonstrates the first application of the partial irradiation method for use with the most widely studied planarian, Schmidtea mediterranea. Additionally, efficient tissue grafting in planaria opens the door for the functional testing of subpopulations of na&iuml;ve or treated stem cells in repopulation assays, which has long been the gold-standard method of assaying adult stem cell potential in mammals8. Broad adoption of these techniques will no doubt lead to a better understanding of the cellular behaviors of adult stem cells during tissue homeostasis and regeneration.

An effective method for grafting tissue of defined and consistent size between planaria is described. Also included is a description of how the immobilization technique used for transplantation can be adapted, in conjunction with lead shields, for the partial irradiation of live animals.

The planarian, a freshwater flatworm, has proven to be a powerful system for dissecting metazoan regeneration and stem cell biology1,2. Planarian ...

An Assay for Lateral Line Regeneration in Adult Zebrafish

Due to the clinical importance of hearing and balance disorders in man, model organisms such as the zebrafish have been used to study lateral line development and regeneration. The zebrafish is particularly attractive for such studies because of its rapid development time and its high regenerative capacity. To date, zebrafish studies of lateral line regeneration have mainly utilized fish of the embryonic and larval stages because of the lower number of neuromasts at these stages. This has made quantitative analysis of lateral line regeneration/and or development easier in the earlier developmental stages. Because many zebrafish models of neurological and non-neurological diseases are studied in the adult fish and not in the embryo/larvae, we focused on developing a quantitative lateral line regenerative assay in adult zebrafish so that an assay was available that could be applied to current adult zebrafish disease models. Building on previous studies by Van Trump et al.17&#160;that described procedures for ablation of hair cells in adult Mexican blind cave fish and zebrafish (Danio rerio), our assay was designed to allow quantitative comparison between control and experimental groups. This was accomplished by developing a regenerative neuromast standard curve based on the percent of neuromast reappearance over a 24 hr time period following gentamicin-induced necrosis of hair cells in a defined region of the lateral line. The assay was also designed to allow extension of the analysis to the individual hair cell level when a higher level of resolution is required.

Because many zebrafish models of neurological and non-neurological diseases are studied in the adult fish rather than the embryo/larvae, we developed a quantitative lateral line regenerative assay that can be applied to adult zebrafish disease models. The assay involved resolution at the 1) neuromast and 2) individual hair cell levels.

Due to the clinical importance of hearing and balance disorders in man, model organisms such as the zebrafish have been used to study lateral line ...

Nitroreductase/Metronidazole-Mediated Ablation and a MATLAB Platform (RpEGEN) for Studying Regeneration of the Zebrafish Retinal Pigment Epithelium

The retinal pigment epithelium (RPE) resides at the back of the eye and performs functions essential for maintaining the health and integrity of adjacent retinal and vascular tissues. At present, the limited reparative capacity of mammalian RPE, which is restricted to small injuries, has hindered progress to understanding in vivo RPE regenerative processes. Here, a detailed methodology is provided to facilitate the study of in vivo RPE repair utilizing the zebrafish, a&#160;vertebrate model capable of robust tissue regeneration. This protocol describes a transgenic nitroreductase/metronidazole (NTR/MTZ)-mediated injury paradigm (rpe65a:nfsB-eGFP), which results in ablation of the central two-thirds of the RPE after 24 h treatment with MTZ, with subsequent tissue recovery. Focus is placed on RPE ablations in larval zebrafish and methods for testing the effects of pharmacological compounds on RPE regeneration are also outlined. Generation and validation of RpEGEN, a MATLAB script created to automate quantification of RPE regeneration based on pigmentation, is also discussed. Beyond active RPE repair mechanisms, this protocol can be expanded to studies of RPE degeneration and injury responses as well as the effects of RPE damage on adjacent retinal and vascular tissues, among other cellular and molecular processes. This zebrafish system holds significant promise in identifying genes, networks, and processes that drive RPE regeneration and RPE disease-related mechanisms, with the long-term goal of applying this knowledge to mammalian systems and, ultimately, toward therapeutic development.

Nitroreductase/Metronidazole-Mediated Ablation and a MATLAB Platform RpEGEN for Studying Regeneration of the Zebrafish Retinal Pigment Epithelium

This protocol describes the methodology to genetically ablate the retinal pigment epithelium (RPE) using a transgenic zebrafish model. Adapting the protocol to incorporate signaling pathway modulation using pharmacological compounds is extensively detailed. A MATLAB platform for quantifying RPE regeneration based on pigmentation was developed and is presented and discussed.

The retinal pigment epithelium (RPE) resides at the back of the eye and performs functions essential for maintaining the health and integrity of adjacent ...

Surgical Ablation Assay for Studying Eye Regeneration in Planarians

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An Assay for Lateral Line Regeneration in Adult Zebrafish

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