Development of a lecithotrophic pilidium larva illustrates convergent evolution of trochophore-like morphology
© The Author(s). 2017
Received: 25 August 2016
Accepted: 3 January 2017
Published: 8 February 2017
The pilidium larva is an idiosyncrasy defining one clade of marine invertebrates, the Pilidiophora (Nemertea, Spiralia). Uniquely, in pilidial development, the juvenile worm forms from a series of isolated rudiments called imaginal discs, then erupts through and devours the larval body during catastrophic metamorphosis. A typical pilidium is planktotrophic and looks like a hat with earflaps, but pilidial diversity is much broader and includes several types of non-feeding pilidia. One of the most intriguing recently discovered types is the lecithotrophic pilidium nielseni of an undescribed species, Micrura sp. “dark” (Lineidae, Heteronemertea, Pilidiophora). The egg-shaped pilidium nielseni bears two transverse circumferential ciliary bands evoking the prototroch and telotroch of the trochophore larva found in some other spiralian phyla (e.g. annelids), but undergoes catastrophic metamorphosis similar to that of other pilidia. While it is clear that the resemblance to the trochophore is convergent, it is not clear how pilidium nielseni acquired this striking morphological similarity.
Here, using light and confocal microscopy, we describe the development of pilidium nielseni from fertilization to metamorphosis, and demonstrate that fundamental aspects of pilidial development are conserved. The juvenile forms via three pairs of imaginal discs and two unpaired rudiments inside a distinct larval epidermis, which is devoured by the juvenile during rapid metamorphosis. Pilidium nielseni even develops transient, reduced lobes and lappets in early stages, re-creating the hat-like appearance of a typical pilidium. Notably, its two transverse ciliary bands can be ontogenetically linked to the primary ciliary band spanning the larval lobes and lappets of the typical planktotrophic pilidium.
Our data shows that the development of pilidium nielseni differs remarkably from that of the trochophore, even though their larval morphology is superficially similar. Pilidium nielseni’s morphological and developmental features are best explained by transition from planktotrophy to lecithotrophy in the context of pilidial development, rather than by retention of or reversal to what is often assumed to be the spiralian ancestral larval type — the trochophore. Development of pilidium nielseni is a compelling example of convergent evolution of a trochophore-like body plan within Spiralia.
Nemerteans (ribbon worms) are a phylum of ~ 1300 described species  of primarily marine spiralians (lophotrochozoans) characterized by an eversible proboscis within a rhynchocoel. Like most benthic marine invertebrates, nemerteans have a biphasic life history with benthic adults and planktonic larvae. Their larvae are usually classified as either planuliform larvae (“direct developers”) or pilidia (“indirect developers”), but these two categories encompass a diverse array of developmental modes.
Planuliform larvae are named for their superficial resemblance to cnidarian planulae (uniform ciliation, specifically), and are found in the Hoplonemertea and the Palaeonemertea (e.g. [2, 3]), two of the three major lineages. Their development is comparatively “direct,” with the larva gradually becoming more worm-like as it transitions into its adult form, although the two groups display significant differences in development, and certain characteristics of indirect development are found in hoplonemerteans [4–6]. The third major lineage, the Pilidiophora [7, 8], which comprises the sister taxa Heteronemertea and Hubrechtiidae, is named for its idiosyncratic pilidium larva, a long-lived planktotroph which typically resembles a deer-stalker cap with the earflaps pulled down (from Greek pilos (πῖλος), or pilidion (πιλίδιον) — a type of brimless conical cap). Pilidial development is “maximally-indirect” ; the juvenile is formed by a series of discrete paired invaginations of the larval epidermis, called imaginal discs, as well as unpaired juvenile rudiments possibly derived from the mesenchyme. A total of eight juvenile rudiments, including three pairs of imaginal discs and two unpaired rudiments, gradually fuse together around the larval gut to form the complete juvenile, which ultimately emerges from—while it simultaneously ingests—the larval body in a dramatic catastrophic metamorphosis [10 and references therein].
The basic elements of pilidial development are conserved in all pilidia; each one develops via a sequence of imaginal discs and rudiments and undergoes catastrophic metamorphosis. However, the shape of the pilidium, the orientation of the juvenile anteroposterior (AP) axis relative to the larval AP axis, and the reported number and sequence of rudiments vary [2, 5, 10–18]. That said, the reported variation in the number of juvenile rudiments formed during the development of different species may be partially attributed to ambiguity in terminology; “imaginal discs” and “juvenile rudiments” are often used interchangeably in the literature (e.g. [16, 19]) as, historically, it was believed that the juvenile developed via seven imaginal discs, all of which invaginated from the larval epidermis (e.g. [20, 21] and references therein). Note here, that we use “imaginal discs” only to describe the paired discs formed by epidermal invaginations, while “juvenile rudiments” will include imaginal discs, as well as the unpaired rudiments not formed by invaginations . These terms indicate tissue origin and formation, so it is important to distinguish between them.
Beyond alterations in morphology, it is also increasingly clear that pilidiophorans have transitioned from a planktotrophic pilidium to a lecithotrophic pilidium repeatedly [2, 15, 22]. Since 2005, the number of pilidiophoran species known (or suspected) to have a non-feeding larva has increased from three (i.e. Desor’s larva, Schmidt’s larva and Iwata’s larva) to twenty [2, 15, 22–24]. Some of these are uniformly ciliated, while others, in addition to a complete covering of short cilia, have one or two circumferential ciliary bands of longer cilia which superficially resemble the prototroch and telotroch of trochophore larvae of other spiralians, e.g. annelids and molluscs [2, 15, 23, 24].
The subject of this study, a trochophore-like pilidium with an anterior “prototroch” and posterior “telotroch,” was dubbed pilidium nielseni  in honor of Claus Nielsen, for his provocative theories on the evolution of marine larval forms, in which the trochophore is considered the ancestral larva of spiralians [25–30]. Pilidium nielseni, which resembles a trochophore, is a lecithotrophic larva of an undescribed lineiform species (Lineidae, Heteronemertea, Pilidiophora) provisionally referred to as Micrura sp. “dark” . Its mere existence prompts a central question in the trochophore debate — is the widespread occurrence of the trochophore morphology among spiralians due to the retention of an ancestral larval form, as Nielsen suggests, or did this larval body plan evolve multiple times independently [31–36]?
Developmental timeline of Micrura sp. “dark”
Earliest appearance (16 °C)
Earliest appearance (8 °C)
Embryo furrows at five to seven sites (Fig. 3c)
Embryo cleaves equally
Blastula is slightly flattened along the animal-vegetal axis
Gastrula is somewhat flattened along animal-vegetal axis, becomes ciliated, and develops an apical tuft and a vegetal invagination (blastopore). Gastrulae may swim freely in advanced stages (Fig. 3e)
Paired cephalic discs invaginate (Fig. 4)
Cephalic and trunk discs
Paired trunk discs invaginate (Fig. 4)
Larva develops transient lobes and lappets, the gut curves backward, the paired cerebral organ discs invaginate from the gut, and the proboscis and dorsal rudiment appear (Figs. 3f and 5). Ciliary bands appear as four segments which span each transient lobe and lappet (Fig. 6)
The head and trunk rudiments fuse around the base of the gut (Fig. 8). Ciliary band segments are re-arranged to form two complete transverse ciliary bands
Epidermis of trunk rudiment extends over the proboscis, but has not yet fused with the epidermis of the head rudiment, leaving a dorsal gap (Fig. 9)
The head and trunk rudiments fused to form a complete juvenile (Fig. 10). Juvenile erupts from and devours larval body in catastrophic metamorphosis
Larvae begin to exhibit a distinctive start-stop swimming behavior between the third and fourth day of development; pilidium nielseni spiral forward led by the apical tuft, then abruptly stop and flare out the cilia, halting ciliary motion for a brief moment (Fig. 3g) before continuing on. At about the same time, the larval ciliary cirrus and an amniotic “larval pore” become apparent below what used to be the posterior larval lobe (now located between the two transverse ciliary bands) (Figs. 5a and e2). The larval pore is located just vegetal to the larval cirrus, and opens through the larval epidermis to the outside (Figs. 6a and 7f).
In as few as eight days, the juvenile begins to move within the larval body, pushing against it, and retracting from it. At the earliest, metamorphosis occurred in only nine days, and most individuals metamorphosed in fewer than 20 days. During its catastrophic metamorphosis, the juvenile extends against the larval body, distorting it, as its tail jabs between the ciliary bands near the lateral cirrus, as described by Maslakova and von Dassow . Confocal imaging exposed a small larval pore open to the outside near the larval cirrus, which suggests that the juvenile may use this pore as an “escape hatch” during metamorphosis (Figures 6a2, 7f and 9a1, 9b.
Newly metamorphosed juveniles have a length of ~500-600 μm in gliding, including a distinct caudal cirrus of ~50 μm (Fig. 2i). The cirrus is sometimes used as a sticky anchor while the juvenile extends its anterior end and writhes in the water. Micrura sp. “dark” juveniles have a pair of longitudinal cephalic slits, as is characteristic of adults of this species (and the entire family Lineidae), and a slight constriction separates the head from the rest of the body, which can appear somewhat bulbous while the stomach is engorged with the larval body (Fig. 3i).
One of the most obvious differences between the typical pilidium and pilidium nielseni is lecithotrophy. In a typical hat-like planktotrophic pilidium, the ciliary bands generate currents while the lobes and lappets perform specialized movements to capture unicellular algae . Likely, other kinds of planktotrophic pilidia, such as the mitten-shaped pilidium auriculatum and sock-shaped pilidium recurvatum, have developed feeding mechanisms suited to their individual morphologies . The elaborate feeding structures and mechanisms required by planktotrophic pilidia are, of course, unnecessary for non-feeding pilidia, which begins to explain their simplified body plans.
All described free-swimming non-feeding pilidia are uniformly ciliated, or have one or two circumferential ciliary bands of long cilia in addition to short cilia covering the rest of the surface (e.g. [2, 12, 15, 22, 23]). They are also more streamlined, shaped like a prolate spheroid. Similar patterns of simplification and modification (with uniform ciliation or circumferential ciliary bands) are seen in the derived non-feeding larvae of some other taxa which ancestrally had a more complex planktotrophic larva, such as bryozoans (e.g. [46–49]), hemichordates (e.g. [50–52]), and echinoderms (e.g. [53–56]). It is thought that these patterns of ciliation and a streamlined body shape improve swimming ability, while complex larval feeding structures, such as ciliated bands extended on lobes and arms, increase hydrodynamic drag, thereby reducing swimming ability [53, 54, 57]. For non-feeding larvae, any pressure to feed efficiently is removed, and may be replaced by pressure to swim efficiently . Accordingly, pilidium nielseni, much like the derived non-feeding larvae in other marine invertebrates, has reduced its feeding structures and reorganized its ciliary bands, converging on a trochophore-like body plan, and likely lessening the energy spent swimming.
Still, the trochophore-like appearance of pilidium nielseni is provocative, and there may be an impulse to draw a direct connection to the hypothetical ancestral trochophore larva of Spiralia (Lophotrochozoa or Trochozoa, depending on the interpretation). However, the “prototroch” and “telotroch” of pilidium nielseni are both positioned anterior to the blastopore (i.e. vestigial mouth), which retains its posterior/vegetal position. The prototroch of a true trochophore would also be anterior to the mouth, but the telotroch, if present, would surround the anus at the posterior end. Furthermore, the “prototroch” and “telotroch” of pilidium nielseni can be ontogenetically linked with the primary ciliary band of a planktotrophic pilidium, as they initially form along the lobes and lappets before wrapping around the larva as two circumferential ciliary bands (Fig. 11a, c). This is analogous to the re-organization of the ciliary bands during the auricularia-to-doliolaria transition in development of holothuroids [58, 59]. The ciliary band of typical pilidia functions very differently from the prototroch-metatroch pair in the opposed-band feeding mechanism described for some trochophores [38, 60–63] and is not homologous to the prototroch as a differentially ciliated band, so we can infer that the circumferential ciliary bands of pilidium nielseni are not homologous to the prototroch and telotroch in a typical spiralian trochophore, either (see Fig. 1). Additional substantiation may be provided by a cell lineage study of pilidium nielseni, which would clarify the relationship between its ciliary bands and those of a typical pilidium and trochophore, and determine which cell lineages contribute to the formation of the pilidium nielseni “trochs.”
Another distinction between planktotrophic pilidia and pilidium nielseni is the size of the eggs from which they arise. The eggs produced by Micrura sp. “dark” are ~250 μm in diameter, much larger than the 75–160 μm eggs of planktotrophic nemertean species  (Fig. 2b). The larger egg size is likely due to the proportionate abundance of yolk, which is later doled out into lipid granules dotting the larval epidermis . The yolk provides enough nutrition for pilidium nielseni to develop a complete juvenile without ever needing to feed, which accounts for pilidium nielseni’s accelerated development to metamorphosis compared to that of a typical planktotrophic pilidium [10, 18, 45, 56, 64]. Relatively large eggs (150–350 μm) are also characteristic of other non-planktotrophic pilidia ( and references therein), and evolution of larger eggs is associated with lecithotrophy in other taxa as well . Though these yolk-rich, relatively short-lived larvae are energetically more expensive to produce, the benefits of a shortened planktonic stage must outweigh the costs in these cases .
Comparison of juvenile rudiment development in several lecithotrophic and one planktotrophic pilidium.
Rudiments reported as imaginal discs
Other reported juvenile rudiments
Rudiments reported with uncertain origin
Total # of juvenile rudiments
Paired cd and td. Unpaired dd
Paired cd, td and cod.
Iwata 1958 
Paired cd and td
Paired cod. Unpaired pb
Paired cd and td
Martîn-Durán et al. 2015 
Paired cd, td, and cod.
Unpaired dr and pb
Paired cd and td
Schmidt 1964 
Schwartz and Norenburg 2005 
Schwartz 2009 
Micrura sp. 803
Schwartz 2009 
Micrura sp. “dark”
Paired cd, td and cod
Unpaired pb and dr
Paired cd, td and cod
Maculaura alaskensis a
Paired cd, td and cod
Unpaired pb and dr
Paired cd, td and cod
Maslakova 2010 
Schmidt’s larva was originally described to form a juvenile via eight imaginal discs—paired cephalic, trunk and cerebral organ discs, and unpaired proboscis and dorsal rudiments—but Schmidt did not specify which invaginate from the epidermis . Recent analysis with confocal microscopy shows only two pairs of imaginal discs, the cephalic and trunk discs, and a separate proboscis rudiment . This technique also revealed a cluster of mesenchymal cells which, based on location, may contribute to the dorsal side of the juvenile, and similarly, another cluster of cells which appear to be associated with the formation of the cerebral organs . More recently discovered non-feeding free-swimming pilidia have been studied in less detail. Micrura rubramaculosa and M. verrilli are thought to develop via five and Micrura sp. 803 via six imaginal discs, but development was only observed through their yolky epidermis, the discs were not identified, and disc formation was not described [15, 23]. So, while published literature suggests there are many possible departures from typical pilidial development in non-feeding larvae, this may be an artifact of the methods employed (e.g. histology vs. confocal microscopy), the depth of study, and interpretation by the author (e.g. which rudiments are counted as imaginal discs and which are not), rather than a representation of true developmental variation.
In this study, the first to document in depth the development of a free-swimming non-feeding pilidium with modern microscopy methods, we have demonstrated that fundamental aspects of pilidial development (8 juvenile rudiments, catastrophic metamorphosis) are conserved in pilidium nielseni, the larva of a pilidiophoran species. Notably, its ciliary bands first form in segments along the transient lobes and lappets, resembling a planktotrophic pilidium, before connecting and encircling the larva as two transverse ciliary bands (Fig. 11) resembling a prototroch and telotroch of some spiralian trochophores. Patterns associated with the transition from planktotrophy to lecithotrophy predict its departures from typical pilidial development, including a larger egg size, an accelerated developmental timeline, a reduction in feeding structures (reduced lobes and lappets), and the rearrangement and repurposing of the ciliary bands (from feeding to locomotion). We suggest that transition from planktotrophy to lecithotrophy explains the trochophore-like morphology of pilidium nielseni, a compelling example of evolutionary convergence on a larval body plan often assumed to be widely homologous.
Collection of adults
We collected a total of 129 adults of Micrura sp. “dark” in rocky intertidal areas around Cape Arago in Charleston, Oregon (especially Middle Cove, 43.305 ̊N, 124.400 ̊ W) during or just prior to their fall/winter reproductive season. Of these, 33 were collected from October 2013 to March 2014, 41 were collected from July 2014 to February 2015, and 55 individuals were collected from July 2015 to March 2016. The increases in individuals collected from one spawning season to the next are likely due to our improved skill in locating them, rather than an increase in population. Fertile adults were observed from September through February. Some were fertile when collected, and others (particularly those collected in July and August) developed gametes in the laboratory following collection. Interestingly, despite being kept unfed in the laboratory for a year, a few males and one female developed gametes the next reproductive season. However, we were unable to start cultures with these males, and their sperm appeared somewhat lackadaisical. Micrura sp. “dark” were primarily found intertwined with the dense root masses of Phyllospadix spp. growing in shell hash, though several individuals were wedged between rocks, or in surf grass rooted in finer sand. Most individuals were collected from root masses of the most dominant surf grass, Phyllospadix serrulatus, but also from the root masses of P. torreyi, and possibly P. scouleri. Micrura sp. “dark” may be confused with two other common co-occurring and also undescribed lineiform nemertean species, which are similar in size (several centimeters long) and color (pinkish, reddish or brownish). Possible misidentifications include Lineus sp. “red,” which has considerably smaller oocytes (~100 μm) than Micrura sp. “dark” and develops via a planktotrophic pilidium , and Lineidae gen. sp. “large eggs,” a species with considerably larger oocytes (~600 μm) and encapsulated lecithotrophic development (, Maslakova, pers. obs.). Micrura sp. “dark” can be distinguished from these two species by the presence of a distinct caudal cirrus (a tail-like extension of the posterior end) (Fig. 2a), and by the nearly constant, pronounced peristaltic motion, which is especially apparent in the foregut region (Fig. 2b). One or two of these dramatic anterior to posterior peristaltic waves can be readily observed at nearly any given time, and their distinct margins conjure up images of a cartoon worm swallowing a series of doughnuts whole.
Initially, individuals were visually identified in the field prior to collection, and subsequently their identity was confirmed via DNA-barcoding (sequencing a 460–537 bp region of the 16S rDNA gene). Once we were confident and consistent in our identifications, confirming identification with DNA sequence data was no longer necessary. Adult individuals were photographed, and kept in 150 ml glass dishes in a flow-through sea table at ambient sea temperature, where their water was changed weekly.
Obtaining gametes and rearing larvae
Gametes were dissected from gravid male and female Micrura sp. “dark” individuals when reproductive pairs were available, and we established a total of 18 cultures over three reproductive seasons from 2013–2016. In three instances, sperm was dissected from a male to fertilize naturally spawned oocytes, and in two others, naturally spawned oocytes and sperm were used. The 13 other cultures resulted from dissected oocytes and sperm. Observations are based on eleven embryonic cultures maintained through metamorphosis, including two started with spawned oocytes and one started with both spawned oocytes and sperm, as well as seven other cultures maintained through early developmental stages (two to three days), including one started with spawned oocytes, and another started with spawned oocytes and sperm. Oocytes were fertilized by a dilute suspension of sperm in filtered sea water (FSW, 0.2 μm), and cultures were maintained in 150 ml glass dishes of FSW placed in flowing sea tables at ambient seawater temperature. The water in their dishes was changed every one to two days. Because the first few cultures suffered a high mortality rate due to bacterial infestation, subsequent cultures were established and maintained in FSW re-filtered through a bottle-top vacuum system (Corning), and an antibiotic solution (a mixture of penicillin and streptomycin at a concentration of 5–50 μg/ml each) was added to the cultures.
Adult specimens of Micrura sp. “dark” were examined and photographed live using a Leica DF400 digital camera mounted to a Leica MZ10F dissecting microscope. Gametes and larval specimens were photographed, trapped between a glass slide and a coverslip supported by clay feet, using a Leica DF400 digital camera mounted to an Olympus BX51 compound microscope equipped with DIC.
Fluorescent labeling and confocal microscopy
Larvae were relaxed in a 1:1 mixture of 0.34 M MgCl2 and FSW for 15 min, then in 100% 0.34 M MgCl2 for 15 min prior to fixation. They were fixed in 4% paraformaldehyde prepared from 16% or 20% ultrapure paraformaldehyde (Electron Microscopy Sciences) and filtered sea water. Fixed specimens were rinsed in three 10-min changes of phosphate buffered saline (PBS, pH 7.4, Fisher Scientific), then stored in PBS at 4 °C, or immediately permeabilized and stained. Larvae were permeabilized with three changes of PBS with 0.1% or 0.5% Triton X-100 (PBT) and rinsed in three 10-min changes of PBS. Specimens were stained with Bodipy FL phallacidin (Molecular Probes) at a concentration of 5 U/ml, propidium iodide (Sigma) at a 0.1% concentration, or a combination of both in 0.1% or 0.5% PBT. Stained specimens were rinsed in three 10-min changes of PBS, then stored in PBS at 4 °C, or immediately mounted. To view internal structures, specimens were mounted onto Poly-L-lysine (Sigma) coated coverslips, dehydrated through an isopropyl alcohol series (70%, 80%, 90%, 100% I, 100% II) for 40 s-1 min at each step, then cleared with three 10-min changes of Murray Clear (a 2:1 mixture of benzyl benzoate and benzyl alcohol). Slides were prepared with strips of foil tape to support the coverslip. After mounting, the coverslips were filled with Murray Clear, which has a refractive index close to that of the immersion oil (~1.5) used for imaging, then sealed with nail polish and imaged immediately, or stored at 4 °C. To view surface features, stained specimens were placed in a glass-bottom microwell dish filled with PBS, and covered with a coverslip.
Specimens were imaged with an Olympus Fluoview 1000 laser scanning confocal mounted on an Olympus IX81 inverted microscope. Specimens mounted in Murray Clear were imaged with a UPlanFLN 40× 1.3 NA oil lens. Uncleared specimens mounted in PBS were imaged using a UPlanFLN 40× 1.15 water lens. Stacks of 0.65 μm optical sections were imported into ImageJ 1.47v (Wayne Rasband, Nations Institute of Health, Bethesda, MD, USA) for further processing. Channels were false-colored and levels adjusted in Adobe Photoshop CS6. In figures, we refer to stacks of a subset of optical sections of a specimen (most often projections of three sections) as “slabs.”
This work was supported by the NSF grant IOS-1120537 to SAM.
Availability of data and materials
Data generated during this study and necessary to interpret and build upon the findings are included in this published article and its supplementary data files. Additional materials (confocal stacks, tissue samples, DNA sequence data) can be made available by the corresponding author on reasonable request.
MKH collected worms, maintained larval cultures, collected all images, prepared most figures, and drafted the manuscript. SAM conceived of the study, assisted in figure preparation, edited drafts and revised manuscript. All authors read and approved the final manuscript.
The authors declare that they have no competing interests.
Consent for publication
Open AccessThis article is distributed under the terms of the Creative Commons Attribution 4.0 International License (http://creativecommons.org/licenses/by/4.0/), which permits unrestricted use, distribution, and reproduction in any medium, provided you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons license, and indicate if changes were made. The Creative Commons Public Domain Dedication waiver (http://creativecommons.org/publicdomain/zero/1.0/) applies to the data made available in this article, unless otherwise stated.
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