Surgical Removal of a Complex Sensory Organ in Highly Regenerative Ctenophores

Ctenophores, also known as comb jellies, are cosmopolitan to the Earth'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 regeneration^1,2. Their ability to regenerate any missing organ or cell type arises towards the end of embryogenesis and persists throughout their postembryonic lifetime^3,4 (~0.1 to >1,000 mm in body size). They are also transparent throughout their life and are amenable to diverse live and fixed imaging techniques. The most recent common ancestor 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' placement as the likely sister group to all other living animals^5,6,7,8. 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 uncertain^9,10,11,12.

The aboral sensory organ complex includes the statocyst, which includes many cell types, including multiple types of putative neurosensory cells^{13,14,15,16,17,18}, 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 "polar fields," which contain putative sensory structures of unknown function¹⁴ and probable photosensory cells¹⁶, are adjacent to the statocyst.

Experimental investigation of M. leidyi's regenerative capacity has spanned the smallest animals at the embryo-to-hatchling transition (~100 µm) to the free-living macroscopic adults (>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.

1. Prepare tools and supplies

1. 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 μl size works well across the focal range of 0.5-3 mm.
   1. Establish a sharp blue cone of flame in the Bunsen burner.
   2. Hold the center point of the glass pipette in the flame until it is softened enough to move under gentle pressure.
   3. 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.
      ​ALTERNATIVE: 26 G ½ inch needles also work well, especially at the larger end of the focal size range.
2. Prepare sterile seawater by passing natural or artificial seawater through a 0.22 µm filter.
3. Create measuring pipettes.
   1. 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.

2. Prepare animals for the experiment

1. 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.
2. 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.

3. Perform cuts

1. 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.
2. 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.
3. Gently hold the animal in place with one glass needle using the non-dominant hand.
4. 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.
5. 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.

4. Check that the cuts are correct

NOTE: Even carefully checking under the dissecting microscope, it is occasionally possible to miss surgical errors.

1. 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.

5. Live imaging

1. Mounting
   1. Treat a standard glass microscope slide with a silanizing agent, such as a water repellant for glass treatment, and allow to dry completely.
   2. Transfer one postsurgical animal to a prepared slide with a transfer pipette. Confirm that the water beads up with the animal inside.
   3. Place a small amount of modeling clay (~1.5 mg) on each corner of a coverglass to form "feet" that will serve as a spacer to lift the coverglass away from the animal.
   4. 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.
   5. 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.

6. Culturing

1. Cydippid culture conditions
   1. Culture surgically manipulated animals and controls in filtered natural or artificial seawater near room temperature. Use an incubator set to 20-22 °C to ensure consistent results.

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 closure of the wound results in an overall smaller body diameter, as shown by before- and after-surgery images (Figure 3).

Using cydippid stage M. leidyi in the size range of 0.5-3 mm, undergraduate students in our lab have been able to produce regeneration assays following this protocol, with 53% of animals fully regenerating their aboral organ by 48 h post removal and 100% 72 h post removal (N = 30).

Following surgery, follow-up assays tailored to the user's hypothesis might include time-lapse imaging, fixation, and labeling (such as RNA in situ hybridization or immunohistochemistry), or extraction of biomolecules for further characterization using published methods.

Figure 1: Mnemiopsis leidyi cydippid with key body axes and organs identified. Top panel: Adesophageal view, aboral-oral axis oriented vertically with oral side down. Bottom panel: Aboral view, adesophageal axis oriented vertically. Scale bar = 0.1 mm. Please click here to view a larger version of this figure.

Figure 2: Hand-pulled needles. (A) Examples of good hand-pulled needles. (B) (a) A well-pulled needle alongside various mistakes that can occur when hand-pulling, including (b) a pipette that did not separate due to slow pulling and/or low temperature, (c) a needle with too much extra material caused by lingering in the flame or multiple attempts using the same glass pipette without removing excess, (d) a needle that is not sharp enough, (e) a needle with a short handle resulting from using the same pipette to make multiple needles, (f) a needle with a hooked end due to pulling the pipette apart at an angle, (g) needle with excess material in the body caused by repeated attempts, (h) a needle with a very short and blunt tip, and (i) a needle with a very long and thin tip. Please click here to view a larger version of this figure.

Figure 3: Measuring pipettes. (A) One uncut disposable pipette and two disposable pipettes with tips cut at different points up the pipette to create larger openings, with the cut-off pieces on the side showing the different internal diameters. (B) Disposable pipettes with different internal diameters, some cut and some not, to create a range of size options. (C) Three different measuring pipettes made from the same size disposable pipette shown at the bottom. Please click here to view a larger version of this figure.

Figure 4: Aboral organ removal and subsequent wound closure in M. leidyi cydippid, lateral view. Top panel: Surgical approach to aboral organ removal. Dashed red line shows the area to remove from a diagrammatic representation of an intact cydippid. Bottom panels: Time series at 10, 30, and 60 min following surgery during wound closure. The wound is closed within minutes of injury. Scale bar = 0.1 mm. Please click here to view a larger version of this figure.

Figure 5: Aboral view of successful aboral organ removal and subsequent wound closure in cydippid stage M.leidyi. Top panel shows an uncut animal; dashed red circle indicates the area to be removed. Middle panel: The same animal immediately after surgery, dashed red circle indicates the approximate wound margins. Exposed mesoglea in the open wound appears as a slightly out-of-focus area in the middle, in which the animal's gut can be seen in section. Bottom panel: The same animal 20 min after surgery. The wound margins have shrunk significantly, and tissue can be seen in the plane of focus across the site above the animal's gut, which was exposed at 0 min. Please click here to view a larger version of this figure.

Figure 6. Lateral view of successful aboral organ removal and subsequent wound closure in cydippid stage M.leidyi. Top left panel: Aboral organ of a cydippid prior to its surgical removal. All other panels: The same cydippid immediately after the removal of the aboral organ (0 min) and the time series of the wound closure (shown clearly at 60 min) and full regeneration of the aboral organ (shown at 72 h). Scale bar = 10 µm. Please click here to view a larger version of this figure.

Figure 7: Lateral view of successful aboral organ removal and subsequent wound closure in lobate stage M. leidyi. Top left panel: Aboral organ of a lobate prior to its surgical removal (full body image inset). All other panels: The same lobate immediately after the removal of the aboral organ (0 min) and the time series of the wound closure (shown at 240 min) and full regeneration of the aboral organ (shown at 120 h). Insets in the time series panels show the aboral view of the wound closing. Scale bar = 1 mm for all panels. Please click here to view a larger version of this figure.

Similar experiments could be performed on any size of M. leidyi desired; the organ's morphogenesis is completed concurrently with the onset of regenerative ability⁴. Animals may be collected from the wild¹⁹ or reared in the lab^20,21. 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 pipettes, made in protocol step 1.3, cut to the desired internal diameter (Figure 3). Using one pipette that is smaller and one that is larger allows for a pool of animals in a size range to be gathered. Handling becomes easier with bigger animals, but immobilization and imaging become more difficult, and animals over ~5 mm in body size may require extensive modifications to this profile. We find animals ~1 week old (or 0.5-3 mm body diameter, as shown in the above protocol) to be in a convenient size range. However, with minimal adjustments to the protocol, any body size is feasible. For example, the AO may be removed from large lobates with standard dissection tools or hypodermic needles (which were used to generate the sample shown in Figure 7). Mounting such samples for imaging could present a greater challenge than performing the surgery, depending on the temporal and spatial resolution desired, and these modifications would go beyond the scope of this protocol; however, very large cydippids and small lobates can be mounted on depression slides.

The handmade dissection needles used in this protocol balance strength and flexibility for animals ≲5 mm, do not require the addition of a handle, and are easy to make, reshape, and replace. However, depending on personal preference and animal size, a wide range of sharp dissecting tools will work, including beveled hypodermic needles, tungsten needles, ophthalmological or other small scalpels. We have tried this protocol on another ctenophore species, Beroe ovata (unpublished), and found that their denser tissue benefits from metallic instruments such as hypodermic needles and microscalpels rather than glass needles.

Clean removals of the aboral organ have a very high rate of regenerative success (virtually 100% in our lab), as do operations that leave that region intact while removing any other structures^4,22,23. However, animals with a fragmented AO show much lower rates of regeneration. If only the AO is removed, there typically is not enough remaining tissue to allow regeneration of that fragment. While most individuals regenerate successfully, nutritional status²⁴, and possibly other environmental factors, influence whether an individual regenerates missing body parts or merely heals the wound. If regeneration fails for many of the individuals in an experiment, the experimental animals' diet or other aspects of culture may be the cause.

Extensions of this protocol and other similar surgeries may be combined with other assays to examine cell cycle dynamics²⁵ or gene expression²⁶ during regeneration or behavioral changes arising from ablation or supernumerary grafts^23,27. The extremely high success rate in the controls provides an excellent launching point for further investigation.