Surgical Removal of a Complex Sensory Organ in Highly Regenerative Ctenophores

Summary

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

Abstract

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.

Introduction

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.

Protocol

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.

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 clos...

Discussion

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 pipet...

Materials

List of materials used in this article

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