Name: Surgical Removal of a Complex Sensory Organ in Highly Regenerative Ctenophores (Video) | JoVE Journal
Uploaded: 2025-08-08T12:40:00.000+00:00
Duration: 5 min 4 s
Description: Learn surgical removal of the statocyst in Mnemiopsis leidyi ctenophores for live imaging studies. This protocol enables detailed observation of regeneration processes and life history strategies.

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5 minutes

The authors gratefully acknowledge Jovita Joseph.

1. Prepare tools and supplies
<ol>
	<li>Using a standard Bunsen burner, hand-pull glass dissection needles from glass capillary micropipettes. For animals over 1 mm, use calibrated soda lime glass micropipettes intended for liquid handling. 
	NOTE: The 100 &#956;l size works well across the focal range of 0.5-3 mm.
	<ol>
		<li>Establish a sharp blue cone of flame in the Bunsen burner.</li>
		<li>Hold the center point of the glass pipette in the flame until it is softened enough to move under gentle pressure.</li>
		<li>Remove the pipette from the flame and immediately pull the two sides straight apart to create a short, strong needle. 
		NOTE: The glass needlestock may be re-melted if a satisfactory result was not obtained in the first attempt, and broken needles may be saved and re-pulled. See Figure 2 for examples of usable and unusable needles pulled by this method. 
		&#8203;ALTERNATIVE: 26 G &#189; inch needles also work well, especially at the larger end of the focal size range.</li>
	</ol>
	</li>
	<li>Prepare sterile seawater by passing natural or artificial seawater through a 0.22 &#181;m filter.</li>
	<li>Create measuring pipettes.
	<ol>
		<li>Using sharp scissors or a razor blade, cut a plastic transfer pipette to produce an opening with the internal diameter of the preferred size. Keep ready at least one pipette for the top of the desired range and one for the bottom of the desired range. See Figure 3 for an example set of measuring pipettes. 
		NOTE: These pipettes work well up to ~8 mm. It is also possible to estimate size with a micrometer or measurement tools from photographs.</li>
	</ol>
	</li>
</ol>
2. Prepare animals for the experiment
<ol>
	<li>Select morphologically normal individuals of the desired size and stage. 
	NOTE: We find animals of ~0.5-3 mm in diameter to be optimal for mounting on slides. Check that animals do not fit in the bottom-of-the-range sizing pipette but do fit comfortably in the top-of-the-range sizing pipette.</li>
	<li>Starve experimental animals for 8-16 h. 
	NOTE: While M. leidyi are optically transparent and not highly autofluorescent, food circulating through their digestive system is generally opaque and often autofluorescent.</li>
</ol>
3. Perform cuts
<ol>
	<li>Use a transfer pipet wider than the body size of the experimental animal to transfer it to a 35 mm polystyrene dish containing sterile seawater.</li>
	<li>Visualize the sample using the dissecting microscope. Roll one animal onto its adesophageal side (Figure 1). This view gives the easiest access to remove the aboral organ. Focus on the aboral organ. 
	NOTE: The most useful magnification may vary by animal size and user preference. Generally 12-20x magnification works well.</li>
	<li>Gently hold the animal in place with one glass needle using the non-dominant hand.</li>
	<li>Using the dominant hand, insert the tip of a needle slightly below the aboral organ. Enter obliquely to make one cut from one visible edge of the aboral organ to below its base. Withdraw the needle and make a second cut to remove a wedge-shaped piece of tissue containing all structures of the aboral organ (see Figure 4, top panel). Examine the animal and the excised tissue carefully to ensure the operation was successful.</li>
	<li>Use the same transfer pipet to move the freshly cut animal into a fresh dish containing sterile seawater. 
	NOTE: The wound will close quickly, so depending on experimental goals (such as for multiple synchronously regenerating samples or to image early stages of healing and regeneration), one must move quickly to generate several samples. Use a timer to limit surgical time to keep cohorts of regenerating animals closer together.</li>
</ol>
4. Check that the cuts are correct
NOTE: Even carefully checking under the dissecting microscope, it is occasionally possible to miss surgical errors.
<ol>
	<li>Follow the mounting steps below to temporarily mount each postsurgical individual on a glass slide to check under a compound microscope with DIC that all the structures intended were correctly removed. Perform this check even if not conducting further imaging at this time. 
	NOTE: The excellent healing abilities of ctenophores lend them well to grafting experiments; we always remove the excised aboral organs from the culture vessel to prevent the possibility that the cut sides of a removed AO and the rest of the animal it was removed from may stick together and fuse. 
	If not proceeding directly to long-term imaging, do not seal the slide. This will facilitate the quick recovery and return to the culture of this experimental animal. Checking surgical accuracy under high magnification is recommended. In particular, the statolith can be knocked out of the dome without completing the surgery.</li>
</ol>
5. Live imaging
<ol>
	<li>Mounting
	<ol>
		<li>Treat a standard glass microscope slide with a silanizing agent, such as a water repellant for glass treatment, and allow to dry completely.</li>
		<li>Transfer one postsurgical animal to a prepared slide with a transfer pipette. Confirm that the water beads up with the animal inside.</li>
		<li>Place a small amount of modeling clay (~1.5 mg) on each corner of a coverglass to form &#34;feet&#34; that will serve as a spacer to lift the coverglass away from the animal.</li>
		<li>Gently place the coverglass, clay-down, on top of the animal. Holding each corner of the coverglass, roll the animal into position for imaging. Press down to compress the animal slightly, holding it in place. Reposition the sample as much as needed to reach a satisfactory view.</li>
		<li>For imaging longer than ~30 min, apply a thin layer of petroleum jelly on each edge of the coverglass to prevent drying of the specimen. 
		NOTE: M. leidyi mounted in this way can be recovered easily for further culture or other assays. However, animals will generally free and reposition themselves within 1-2 h, so a continuous time-lapse capture longer than 2 h would require a different imaging approach.</li>
	</ol>
	</li>
</ol>
6. Culturing
<ol>
	<li>Cydippid culture conditions
	<ol>
		<li>Culture surgically manipulated animals and controls in filtered natural or artificial seawater near room temperature. Use an incubator set to 20-22 &#176;C to ensure consistent results.</li>
	</ol>
	</li>
</ol>

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<li>Whelan, N. V., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Ctenophore+relationships+and+their+placement+as+the+sister+group+to+all+other+animals%5BTitle%5D)+AND+%22Nat+Ecol+Evol%22%5BJournal%5D)">Ctenophore relationships and their placement as the sister group to all other animals.</a> Nat Ecol Evol. 1 (11), 1737-1746 (2017).</li>
<li>Schultz, D. T., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Ancient+gene+linkages+support+ctenophores+as+sister+to+other+animals%5BTitle%5D)+AND+%22Nature%22%5BJournal%5D)">Ancient gene linkages support ctenophores as sister to other animals.</a> Nature. 618, 110-117 (2023).</li>
<li>Burkhardt, P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Ctenophores+and+the+evolutionary+origin%28s%29+of+neurons%5BTitle%5D)+AND+%22Trends+Neurosci%22%5BJournal%5D)">Ctenophores and the evolutionary origin(s) of neurons.</a> Trends Neurosci. 45 (12), 878-880 (2022).</li>
<li>Sachkova, M. Y., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Neuropeptide+repertoire+and+3D+anatomy+of+the+ctenophore+nervous+system%5BTitle%5D)+AND+%22Curr+Biol%22%5BJournal%5D)">Neuropeptide repertoire and 3D anatomy of the ctenophore nervous system.</a> Curr Biol. 31 (23), 5274-5285.e6 (2021).</li>
<li>Moroz, L. L., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((The+ctenophore+genome+and+the+evolutionary+origins+of+neural+systems%5BTitle%5D)+AND+%22Nature%22%5BJournal%5D)">The ctenophore genome and the evolutionary origins of neural systems.</a> Nature. 510 (7503), 109-114 (2014).</li>
<li>Burkhardt, P., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Syncytial+nerve+net+in+a+ctenophore+adds+insights+on+the+evolution+of+nervous+systems%5BTitle%5D)+AND+%22Science%22%5BJournal%5D)">Syncytial nerve net in a ctenophore adds insights on the evolution of nervous systems.</a> Science. 380 (6642), 293-297 (2023).</li>
<li>Tamm, S. L., Tamm, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Novel+bridge+of+axon-like+processes+of+epithelial+cells+in+the+aboral+sense+organ+of+ctenophores%5BTitle%5D)+AND+%22J+Morphol%22%5BJournal%5D)">Novel bridge of axon-like processes of epithelial cells in the aboral sense organ of ctenophores.</a> J Morphol. 254 (2), 99-120 (2002).</li>
<li>Jager, M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((New+insights+on+ctenophore+neural+anatomy%3A+Immunofluorescence+study+in+Pleurobrachia+pileus+%28M%C3%BCller%2C+1776%29%5BTitle%5D)+AND+%22J+Exp+Zool+B+Mol+Dev+Evol%22%5BJournal%5D)">New insights on ctenophore neural anatomy: Immunofluorescence study in Pleurobrachia pileus (Müller, 1776).</a> J Exp Zool B Mol Dev Evol. 316B (3), 171-187 (2011).</li>
<li>Hernandez-Nicaise, M. -L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((The+nervous+system+of+ctenophores+III.+Ultrastructure+of+synapses%5BTitle%5D)+AND+%22J+Neurocytol%22%5BJournal%5D)">The nervous system of ctenophores III. Ultrastructure of synapses.</a> J Neurocytol. 2 (3), 249-263 (1973).</li>
<li>Schnitzler, C. E., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Genomic+organization%2C+evolution%2C+and+expression+of+photoprotein+and+opsin+genes+in+Mnemiopsis+leidyi%3A+a+new+view+of+ctenophore+photocytes%5BTitle%5D)+AND+%22BMC+Biol%22%5BJournal%5D)">Genomic organization, evolution, and expression of photoprotein and opsin genes in Mnemiopsis leidyi: a new view of ctenophore photocytes.</a> BMC Biol. 10 (1), 107(2012).</li>
<li>Harrison, F. W., Westfall, J. A. Ctenophora. <a target="_blank" href="http://scholar.google.com/scholar?hl=en&safe=off&q=author%3aHernandez-Nicaise%2C+ML+%22Microscopic+Anatomy+of+Invertebrates.+2%3A+Placozoa%2C+Porifera%2C+Cnidaria%2C+and+Ctenophora%22">Microscopic Anatomy of Invertebrates. 2: Placozoa, Porifera, Cnidaria, and Ctenophora</a>. , Wiley. New York. 359-418 (1991).</li>
<li>Horridge, G. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Presumed+photoreceptive+cilia+in+a+ctenophore%5BTitle%5D)+AND+%22J+Cell+Sci%22%5BJournal%5D)">Presumed photoreceptive cilia in a ctenophore.</a> J Cell Sci. S3-105 (71), 311-317 (1964).</li>
<li>Ramon-Mateu, J., Edgar, A., Mitchell, D., Martindale, M. Q. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Experimental+methods+to+study+ctenophore+whole-body+regeneration%5BTitle%5D)+AND+%22Methods+Mol+Biol%22%5BJournal%5D)">Experimental methods to study ctenophore whole-body regeneration.</a> Methods Mol Biol. 2450, 95-119 (2022).</li>
<li>Soto-Àngel, J. J., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Stable+laboratory+culture+system+for+the+ctenophore+Mnemiopsis+leidyi%5BTitle%5D)+AND+%22Methods+Mol+Biol%22%5BJournal%5D)">Stable laboratory culture system for the ctenophore Mnemiopsis leidyi.</a> Methods Mol Biol. 2757, 123-145 (2022).</li>
<li>Presnell, J. S., Bubel, M., Knowles, T., Patry, W., Browne, W. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Multigenerational+laboratory+culture+of+pelagic+ctenophores+and+CRISPR-Cas9+genome+editing+in+the+lobate+Mnemiopsis+leidyi%5BTitle%5D)+AND+%22Nat+Protoc%22%5BJournal%5D)">Multigenerational laboratory culture of pelagic ctenophores and CRISPR-Cas9 genome editing in the lobate Mnemiopsis leidyi.</a> Nat Protoc. 17 (8), 1868-1900 (2022).</li>
<li>Coonfield, B. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Apical+dominance+and+polarity+in+Mnemiopsis+leidyi%2C+Agassiz%5BTitle%5D)+AND+%22Biol+Bull%22%5BJournal%5D)">Apical dominance and polarity in Mnemiopsis leidyi, Agassiz.</a> Biol Bull. 70 (3), 460-471 (1936).</li>
<li>Freeman, G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Studies+on+regeneration+in+the+creeping+ctenophore%2C+Vallicula+multiformis%5BTitle%5D)+AND+%22J+Morphol%22%5BJournal%5D)">Studies on regeneration in the creeping ctenophore, Vallicula multiformis.</a> J Morphol. 123, 71-84 (1967).</li>
<li>Bading, K. T., Kaehlert, S., Chi, X., Jaspers, C., Martindale, M. Q., Javidpour, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Food+availability+drives+plastic+self-repair+response+in+a+basal+metazoan-case+study+on+the+ctenophore+Mnemiopsis+leidyi+A.+Agassiz%5BTitle%5D)+AND+%22Sci+Rep%22%5BJournal%5D)">Food availability drives plastic self-repair response in a basal metazoan-case study on the ctenophore Mnemiopsis leidyi A. Agassiz.</a> Sci Rep. 7 (1), 16419(2017).</li>
<li>Mateu, J. R., Ellison, S. T., Angelini, T. E., Martindale, M. Q. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Regeneration+in+the+ctenophore+Mnemiopsis+leidyi+occurs+in+the+absence+of+a+blastema%2C+requires+cell+division%2C+and+is+temporally+separable+from+wound+healing%5BTitle%5D)+AND+%22BMC+Biol%22%5BJournal%5D)">Regeneration in the ctenophore Mnemiopsis leidyi occurs in the absence of a blastema, requires cell division, and is temporally separable from wound healing.</a> BMC Biol. 17 (1), 80(2019).</li>
<li>Mitchell, D. G., Edgar, A., Mateu, J. R., Ryan, J. F., Martindale, M. Q. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((The+ctenophore+Mnemiopsis+leidyi+deploys+a+rapid+injury+response+dating+back+to+the+last+common+animal+ancestor%5BTitle%5D)+AND+%22Commun+Biol%22%5BJournal%5D)">The ctenophore Mnemiopsis leidyi deploys a rapid injury response dating back to the last common animal ancestor.</a> Commun Biol. 7 (1), 203(2024).</li>
<li>Jokura, K., Anttonen, T., Rodriguez-Santiago, M., Arenas, O. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Rapid+physiological+integration+of+fused+ctenophores%5BTitle%5D)+AND+%22Curr+Biol%22%5BJournal%5D)">Rapid physiological integration of fused ctenophores.</a> Curr Biol. 34 (19), R889-R890 (2024).</li>
</ol>

The authors have no conflicts of interest to disclose.

Similar experiments could be performed on any size of M. leidyi desired; the organ&#39;s morphogenesis is completed concurrently with the onset of regenerative ability4. Animals may be collected from the wild19 or reared in the lab20,21. If a particular body size is desired, animals can be allowed to spawn, and the offspring raised to the desired size. Body size can be approximated using the measuring pipet...

Ctenophores, also known as comb jellies, are cosmopolitan to the Earth&#39;s marine environments and can be major players in ecosystems, particularly as part of the food web. The ctenophore Mnemiopsis leidyi is a historied and increasingly popular animal model of whole-body regeneration1,2. Their ability to regenerate any missing organ or cell type arises towards the end of embryogenesis and persists throughout their postembryonic lifetime3,4 (~0.1 to &#62;1,000 mm in body size). They are also transparent throughout their life and are amenable to diverse live and fixed imaging techniques. The&#160;most recent common ancestor&#160;of all ctenophores had many cell types and complex organs; most extant ctenophores possess most of these during at least one life stage: an aboral, gravity-sensing organ (historically called the apical organ and here called the aboral organ, AO), specialized locomotory structures composed of highly organized macrocilia that beat in a coordinated manner (comb rows), retractable feeding tentacles, and photocytes that bioluminesce in response to noxious stimuli (Figure 1).
Despite their numerous cell types and complex, context-dependent behaviors, a robust body of genomic data supports ctenophores&#39; placement as the likely sister group to all other living animals5,6,7,8. Major differences in the cellular and functional basis of the ctenophore nervous system from other animals possessing a nervous system (bilaterians and cnidarians) have suggested the hypothesis that the ctenophore nervous system represents an independent origin; however, this remains uncertain9,10,11,12.
The aboral sensory organ complex includes the statocyst, which includes many cell types, including multiple types of putative neurosensory cells13,14,15,16,17,18, and is under constant maintenance in unmanipulated ctenophores. Mechanosensory cells bear large ciliary bundles, which are deflected by a statolith made up of biomineralized cells (called lithocytes) to detect gravity. Two &#34;polar fields,&#34; which contain putative sensory structures of unknown function14 and probable photosensory cells16, are adjacent to the statocyst.
Experimental investigation of M. leidyi&#39;s regenerative capacity has spanned the smallest animals at the embryo-to-hatchling transition (~100 &#181;m) to the free-living macroscopic adults (&#62;2 cm). M. leidyi of ~1 mm in body diameter are large enough to permit manual surgical manipulation without special instruments such as micromanipulators but are small enough to be mounted on a standard microscope slide for imaging with high-powered objectives. This protocol demonstrates surgical removal of the aboral sensory organ and associated structures, along with culturing and imaging methods appropriate to document the processes of wound healing and regeneration that take place in the following hours to days.

<table><tbody><tr><td>0.2 &amp;mu;m SFCA syringe filter</td><td>Thermo/Nalgene</td><td>723-2520</td><td>create sterile seawater</td></tr><tr><td>35 mm polystyrene petri dish</td><td>Falcon/Corning</td><td>351008</td><td>perform cuts; culture small animals</td></tr><tr><td>60 mL disposable luer-lock syringe</td><td>BH Supplies</td><td>BH60LL</td><td>create sterile seawater</td></tr><tr><td>Calibrated glass pipets, 100 &amp;mu;L</td><td>Drummond</td><td>2-000-100</td><td>perform cuts</td></tr><tr><td>cover glasses, #1.5, 18 mm square</td><td>VWR</td><td>16004-326</td><td>mount for imaging</td></tr><tr><td>ctenophores</td><td>Gulf Specimens</td><td></td><td>live animals</td></tr><tr><td>Double cavity glass depression slides, 1.3 mm</td><td>VWR</td><td>470200-930</td><td>mount for imaging</td></tr><tr><td>glass microscope slides</td><td>VWR</td><td>16004-398</td><td>mount for imaging</td></tr><tr><td>Instant Ocean Sea Salt</td><td>Instant Ocean</td><td></td><td>animal culture medium</td></tr><tr><td>mini ruler</td><td>Ted Pella</td><td>13623</td><td>measure plastic pipettes</td></tr><tr><td>modeling clay (plasticine)</td><td>Pepy Plastilina</td><td></td><td>mount for imaging</td></tr><tr><td>petroelum jelly</td><td>Vaseline</td><td></td><td>mount for imaging</td></tr><tr><td>plastic transfer pipettes (assorted sizes)</td><td>Cole-Parmer</td><td>06226-13, 06226-12, 06226-01</td><td>move small animals</td></tr><tr><td>Rain-X glass treatment</td><td>Rain-X</td><td></td><td>silanize microscope slides</td></tr><tr><td>ruler</td><td>Westcott</td><td>10564</td><td>measure plastic pipettes</td></tr><tr><td>scissors</td><td>OXO</td><td></td><td>cut plastic pipettes</td></tr></tbody></table>

surgical removal of a complex sensory organ in highly regenerative ctenophores

Surgical Removal of a Complex Sensory Organ in Highly Regenerative Ctenophores

The ctenophore Mnemiopsis leidyi is a classic animal model of whole-body regeneration. Increased tractability for many laboratory techniques and their phylogenetic placement as the likely sister group to the remaining Metazoa has led to a recent renewal of scientific interest in working with ctenophores. They can regenerate any missing organ or cell type, including complete whole-body regeneration from a fragment as small as ~15% of the body, over the course of a few days. Like most ctenophores, M. leidyi have an aborally located, gravity-sensing organ that links sensory input to motor output to control their body position and orientation. This protocol demonstrates surgical removal of the aboral organ (AO) complex and associated structures in M. leidyi along with culturing, handling, and mounting methods appropriate to image the processes of wound healing and regeneration that take place in the following hours to days. These straightforward techniques are broadly adaptable to different experimental paradigms and laboratory contexts.

Cydippid-stage M. leidyi typically complete wound closure within ~20 min after injury (Figure 4 and Figure 5), while complete regeneration of the organ can take 1-3 days. This timing has some biological variability but is almost invariably completed by 72 h after surgery (Figure 6).
The surgical removal of cells and mesoglea from the aboral organ complex and nearby tissues, followed by rapid clos...

We describe a simple protocol for removing the statocyst and associated tissues in the ctenophore Mnemiopsis leidyi, which is amenable to live imaging.

The ctenophore Mnemiopsis leidyi is a classic animal model of whole-body regeneration. Increased tractability for many laboratory techniques and their ...

surgical-removal-complex-sensory-organ-highly-regenerative

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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.

Surgical Ablation Assay for Studying Eye Regeneration in Planarians

Here we present a protocol for minimally invasive surgical lesioning of muscles intrinsic to the feeding apparatus of the marine mollusk Aplysia californica to understand the roles of these muscles during feeding behavior.

A Minimally Invasive Lesion Technique for Muscles Intrinsic to the Odontophore of Aplysia californica

This protocol presents a method for imaging neuronal population activity with single-cell resolution in non-transgenic invertebrate species using absorbance voltage-sensitive dyes and a photodiode array. This approach enables a rapid workflow, wherein imaging and analysis can be pursued over the course of a single day.

Photodiode-Based Optical Imaging for Recording Network Dynamics with Single-Neuron Resolution in Non-Transgenic Invertebrates

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.

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

This is part one of a comprehensive Xenopus sampling protocol. The tissues sampled are the heart ventricle, arterial trunk, left liver lobe, gallbladder, lung, pancreas, spleen, larynx, esophagus, stomach, intestines, testes, fat bodies, oviduct, paired kidneys, sciatic plexus, skin, thymus, and whole eye.

Efficient Techniques for Comprehensive Sampling of Accessible Tissues in Adult Xenopus

Learn surgical removal of the statocyst in Mnemiopsis leidyi ctenophores for live imaging studies. This protocol enables detailed observation of regeneration processes and life history strategies.

Surgical Removal of a Complex Sensory Organ in Highly Regenerative Ctenophores

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