Opposed bands of cilia in the nonfeeding larvae of the serpulid tribranchiata

Bruno Pernet

Department of Biological Sciences, California State University, Long Beach, 1250 Bellflower Blvd, Long Beach, CA 90840, USA

E-mail: [email protected] Tel: 562-985-5378

Additional key words: metatroch; ; Salmacina;

2

Abstract. The nonfeeding planktonic larvae of marine invertebrates typically lack larval feeding structures. One puzzling exception to this generalization is the annelid clade , in which nonfeeding larvae possess ciliary bands (specifically, food groove and metatroch) that, to the best of our knowledge, have no function other than in feeding. Nishi and Yamasu (1992b) published a scanning electron micrograph showing that nonfeeding larvae of the serpulid annelid

Salmacina dysteri also possess food groove and metatrochal cilia. Here I demonstrate that nonfeeding larvae of S. tribranchiata also bear ciliary bands identifiable as food groove and metatroch by position. High-speed video of ciliary beat patterns shows that, together with the prototrochal cilia, these bands function in an opposed band system. The presence of feeding structures in nonfeeding annelid larvae is thus more widely distributed than previously recognized. The presence of feeding structures may make evolutionary transitions to planktotrophy more likely, and may underlie an inferred origin of larval feeding in the common ancestor of one of the two major clades of serpulid , Serpulinae.

1. INTRODUCTION

Planktotrophic (feeding) larvae of marine invertebrates have morphological features used solely to capture food particles, but lecithotrophic (nonfeeding) larvae usually lack such structures (Emlet 1991; Strathmann 1993; Collin and Moran 2018). In many cases, lecithotrophy appears to be the derived condition (Hart 2000; Collin and Moran 2018). In these cases, the loss of feeding structures in nonfeeding larvae is generally interpreted as a consequence of relaxed selection on feeding performance. Feeding structures may be lost because remaining sources of selection – on developmental rate, energetic efficiency, or other aspects of larval performance – act to remove them, or simply as a result of the accumulation of neutral mutations that affect 3 their development (Wray 1996; Lahti et al. 2009). Though we have little quantitative information on the rate of loss of feeding structures following the evolution of lecithotrophy (but see Wray

1992; Emlet 1995), the rarity of nonfeeding larvae that bear particle capture structures suggests that this transition is relatively rapid (Wray and Raff 1991; Hart 1996; Wray 1996). This logical framework appears general, applying at the least to the two major clades best known in terms of larval biology, the echinoderms and molluscs (e.g., Hofstee and Pernet 2011; Li and O’Foighil

2016; Wray 1996).

However, the larvae of one major clade of annelids, Sabellidae, fit into this framework only uncomfortably. Sabellidae (a term used here in the narrow sense of Kupriyanova and Rouse

2008) is a group of 250-400 of mostly large-bodied “featherduster worms” (Rouse and

Pleijel 2001). Most of these species have planktonic larvae, but no sabellid is known to have feeding larvae (Rouse and Fitzhugh 1994). Kupriyanova and Rouse (2008) suggest that

Sabellidae is sister to a clade comprised of Fabriciidae (a group of ~75 species of small-bodied featherduster worms, all with intratubular brooding, nonfeeding development) and Serpulidae

(the “fanworms”, a group of ~500 species, some of which have feeding larvae, and others of which have nonfeeding larvae) (Fig. 1). This clade is known as . Sister to Sabellida is

Spionida, a clade composed of Sabellariidae and Spionidae (Struck et al. 2015; Weigert and

Bleidorn 2016; Struck 2019). All sabellariids whose development has been described have feeding larvae; spionids include some species with feeding larvae, and some with nonfeeding larvae (Rouse and Pleijel 2001).

The feeding larvae of sabellariids, at least one spionid, and serpulids capture food particles between “opposed bands” of cilia (Strathmann et al. 1972; Pernet and McArthur 2006; Pernet and Strathmann 2011). Such opposed band feeding systems involve three equatorial bands of 4

cilia – a pre-oral prototroch, a post-oral metatroch, and an intermediate (peri-oral) food groove.

Prototrochal cilia beat from anterior to posterior, generating a swimming current as well as

capturing food particles; metatrochal cilia beat from posterior to anterior, serving uncertain

functions in trapping particles. Particles trapped between these two “opposed” bands are carried

to the mouth by the cilia of the food groove. Metatrochal and food groove cilia are not known to

have functions other than in food capture and transport.

To my knowledge, no formal comparative analysis of the evolution of larval ciliation or

larval nutritional mode in sabellidans and spionidans has been carried out in the context of our

current understanding of their relationships (Fig. 1; for earlier analyses, see Rouse 2000;

Kupriyanova 2003). However, the distribution of metatrochal and food groove cilia suggests that

these ciliary bands were present in the common ancestor of the clade (Sabellida + Spionida).

Since these ciliary bands are only known to function in larval feeding, it is possible that this (or

an earlier) ancestor had feeding larvae. After the evolutionary loss of larval feeding, one might

expect the relatively rapid loss of metatroch and food groove. However, Pernet (2003) showed

that nonfeeding larvae of diverse sabellids – members of at least eight genera, distributed across

the three major clades of sabellids recovered by Kupriyanova and Rouse (2008) – possess these

ciliary bands. In all four sabellid species directly observed by Pernet (2003), metatrochal and

food groove cilia beat in the expected directions, and in the species observed in the most detail

(Schizobranchia insignis BUSH 1905), the opposed band system captured suspended particles and

transported them to the mouth (where they were rejected, as the larval mouth of S. insignis does

not connect to the rest of the digestive system).

It is unclear why the nonfeeding larvae of sabellids possess apparently functional opposed

band systems. The beat of metatrochal cilia may negatively affect larval swimming performance 5

(Strathmann and Grunbaum 2006), so maintaining this ciliary band may be energetically costly

and its presence in nonfeeding larvae may be selectively unfavorable. Sabellids may have had

lecithotrophic development for several hundred million years ago, so there has likely been

substantial time for the accumulation of neutral mutations that might affect the development of

metatroch and food groove (Pernet 2003). The presence of structures known only to function in

larval feeding in an old, diverse clade of species with nonfeeding larvae is thus unexpected and

puzzling.

As noted by Pernet (2003), this phenomenon might not be limited only to Sabellidae;

nonfeeding larvae of some serpulids may also possess metatrochal and food groove cilia. Hints

of this can be found in two papers. In the first, Nishi and Yamasu (1992a) provide a drawing

(their Figure 5C) of the larva of Rhodopsis pusilla BUSH 1905; the drawing shows a partial equatorial ciliary band in what is likely a post-oral position, suggesting it is a metatroch. No food groove cilia are shown in the drawing. Scanning electron micrographs in that paper are of poor quality and do not show identifiable metatrochal or food groove cilia, and neither ciliary band is mentioned in the text of the paper. Much stronger evidence is provided in a second paper, on the development of (HUXLEY 1855) (Nishi and Yamasu 1992b). In that paper the authors provide a scanning electron micrograph of a larva (their Figure 3C) that unambiguously shows peri- and post-oral ciliary bands (i.e., food groove and metatroch). The authors do not mention these ciliary bands in the text of the paper.

Here I describe the nonfeeding larvae of Salmacina tribranchiata (MOORE 1923). Like the

larvae of S. dysteri figured by Nishi and Yamasu (1992b), larvae of this species bear bands of

cilia identifiable as metatroch and food groove by position. I show that metatrochal cilia beat

from posterior to anterior, and food groove cilia beat equatorially towards the mouth, as expected 6 in an opposed band system. I also use observations of larval anatomy and metamorphic competence to support the assertion that these larvae are indeed lecithotrophic. The puzzling presence of feeding structures in nonfeeding annelid larvae is thus more widely distributed than previously recognized.

2. METHODS

2.1 Collection and maintenance

Aggregations of Salmacina tribranchiata were collected by divers from ~10 m depth near

White Point, Los Angeles County, California, on 1 Nov 2019. Aggregations were placed in an insulated cooler and transported to the CSU Long Beach Marine Laboratory. There they were transferred to an aquarium holding recirculating seawater at 16°C, where they were held for the next few weeks without supplementary food. When needed, larvae were obtained by breaking off a small (~2-3 cm diameter) piece of an aggregation, placing it into a small dish of seawater, and crushing most of the adult tubes, freeing any brooded embryos or larvae. Embryos and early larvae could be picked by pipet from the bottom of the dish; late larvae were active swimmers and strongly phototactic, so could be easily gathered by pipet from the side of the dish nearest a source of illumination.

2.2 Microscopy

Brightfield microscopy. Living larvae were mounted on slides and prevented from swimming by gently compressing them under a coverslip supported by Plasticine modelling material at its corners. They were observed and photographed with a Nikon E600 compound microscope equipped with a QImaging MicroPublisher 6 camera. For high-speed video 7

microscopy, larvae were prepared in the same way but observed on an Olympus BX51

compound microscope onto which an Edgertronic SC1 color high-speed video camera had been

mounted. High-speed sequences were captured at 300 frames per second, with a shutter speed of

1/500 second.

Details of gut structure were observed in semi-thin sections. Late larvae (those with three

chaetigerous segments) were briefly anaesthetized in a 1:1 solution of 7.5% MgCl2 and filtered seawater (FSW), then fixed for 2 hr in 2.5% paraformaldehyde in FSW. After rinsing in FSW, they were post-fixed in 1.5% osmium tetroxide in FSW. They were again rinsed in FSW, dehydrated in an ascending series of ethanol concentrations, transferred through three changes of propylene oxide, then infiltrated with and embedded in Araldite. Semi-thin sections (~200 nm) were cut using a diamond knife, mounted on glass slides, and heat-stained with toluidine blue.

Confocal microscopy. Additional information on gut structure was obtained by confocal microscopy. Late larvae were anaesthetized as above, then fixed for 1 hr in 2.5% paraformaldehyde in FSW. They were rinsed several times in phosphate buffered saline (PBS), then incubated in PBS with 0.1% Triton-X (PBT) for 1 hr. Filamentous actin was labelled for 2 hr using BODIPY-FL phallacidin (Molecular Probes B-607; 6 U/ml in PBT). Labelled larvae were rinsed three times in PBS, then mounted on slides coated with poly-L-lysine. Slides were dehydrated in an ascending concentration series of 2-propanol (to 100%), then cleared in three changes of BABB (one part benzyl alcohol and two parts benzyl benzoate). Larvae were mounted in BABB, with coverslips supported by tape and sealed using five-minute epoxy.

Larvae were Z-sectioned with 2-µm steps using an Olympus Fluoview 1000 confocal microscope with a 20X, 0.75 NA objective. Stacks of images were viewed, and maximum intensity Z-projections produced, in ImageJ 2.0.0. 8

Scanning electron microscopy. Larvae of a variety of stages were anaesthetized as above,

then fixed for 2 hr in 1.5% osmium tetroxide in seawater. Fixed larvae were rinsed in filtered

seawater, dehydrated in an ascending series of ethanol concentrations, and critical point dried

using CO2 as a transitional fluid. They were mounted on copper conductive tape with conductive adhesive on both sides, sputter-coated with gold-palladium, and imaged using an FEI Quanta

200.

2.3 Larval feeding

To determine if late larvae could ingest particles, several hundred late larvae were incubated in a beaker in the dark with high (but unmeasured) concentrations of cells of both Isochrysis galbana and Rhodomonas lens. Cells of both of these species are captured and ingested by the feeding larvae of sabellariids and serpulids (Pernet and Strathmann 2011; Pernet, pers. obs.). The beaker was stirred using a paddle system (M. Strathmann 1987). After two hours, the larvae were concentrated on a Nitex mesh. Twenty haphazardly selected living larvae were immediately compressed under a coverslip on a glass slide and observed using epifluorescence microscopy on an Olympus BX51 compound microscope. The remaining larvae were anaesthetized as above, then killed by addition of paraformaldehyde to a final concentration of 4%. Later, an additional

40 haphazardly selected fixed larvae were observed using epifluorescence microscopy.

2.4 Metamorphic competence

To determine whether or not larvae of S. tribranchiata require particulate food to develop to metamorphic competence, I collected 40 late larvae from parental tubes (where they have no access to particulate food) and immediately placed them in 5 ml of FSW in a small bowl. I added 9 several small fragments of clean (free of juveniles) adult tube material to the bowl and incubated it in the dark. No suspended food particles were available for these larvae. I checked for juveniles in the bowl after 3 and 22 hr.

3. RESULTS

Tubes of adult Salmacina tribranchiata contained embryos and larvae of a diversity of stages, from zygotes to three chaetiger larvae. Early larvae were roughly pear-shaped and had no chaetae. At the widest part of the body, they bore a band of prototrochal cilia that was clearly visible with light microscopy (Fig. 2A). This band was used for swimming, which early larvae did only weakly. Anterior to the prototroch, larvae bore two dark red eyespots. The majority of the body of early larvae was filled with a large mass of orange spheres, each ~5-15 µm in diameter. These were presumably lipid droplets. Late larvae were slightly elongated relative to early larvae (Fig. 2B). Posterior to the prototroch, these larvae had annulations that suggested segmentation, and bore three pairs of chaetae. The center of the body remained filled with a mass of orange spheres, as in early larvae. Late larvae swam vigorously using the prototroch and were strongly phototactic.

Scanning electron microscopy showed that both early and late larvae bore not only a prototroch, but also peri- and post-oral bands of cilia (i.e., food groove and metatroch: Fig. 2C-

F). The positions of these bands of cilia relative to the mouth could be identified because the mouth was visible as a small mid-ventral depression. At both stages, the prototroch, which completely encircled the body, was composed of three tiers of cilia – an anterior tier of short cilia, a central tier of long (~35-50 µm) compound cilia, and a posterior tier of short cilia. In early larvae, the food groove and metatrochal cilia were sparse (Fig. 2E). In fixed specimens, 10 food groove cilia typically rested against the body, with their tips pointing dorsally, away from the mouth. The width of the food groove was ~4 µm on the sides of the body. Metatrochal cilia also rested against the body, with their tips pointed posteriorly; these cilia were about 15 µm in length. Early larvae also bore a large tuft of cilia midventrally near the posterior end of the body but did not have a neurotroch.

In late larvae, food groove and metatrochal cilia were much denser, but otherwise similar in position and orientation to those in early larvae (Fig. 2F). At this stage, metatrochal cilia were

15-25 µm in length; the food groove was still about 4 µm wide on the sides of the body. Late larvae had a neurotroch running from the mouth to the midventral posterior tuft of cilia.

High speed video showed that the active strokes of prototrochal cilia were from anterior to posterior, and that those of metatrochal cilia were from posterior to anterior (Video S1). It was more difficult to identify the active strokes of food groove cilia, but they appeared to be equatorial, towards the mouth (Video S1, starting at ~22 sec). Though the prototrochal cilia beat continuously in the specimens I observed, both food groove and metatrochal cilia were often arrested (e.g., Video S1, 5-15 sec).

Swimming larvae of S. tribranchiata did not ingest potential food particles. After two hours of incubation with cells of Isochrysis galbana and Rhodomonas lens, none of the 20 larvae observed alive or the 40 larvae observed after fixation contained algal cells in their bodies, as determined by the absence of chlorophyll autofluorescence in the larvae. In both living and fixed samples, algal cells floating freely in suspension were strongly autofluorescent and easily visible.

Confocal microscopy indicated that the digestive systems of late larvae were usually not complete. In the ten larvae imaged, the mouth appeared to connect to the midgut, but in nine of the ten larvae the midgut clearly did not connect to the hindgut (Fig. 3A,B) (in the 10th larva, it 11

was unclear if those two digestive compartments were connected). In specimens prepared for

confocal microscopy, the orange spheres clearly visible in living larvae (Fig. 2A,B) were not

visible, so their position relative to the digestive system could not be determined. I suspect that

their contents were extracted by the dehydration and clearing steps during preparation for

confocal microscopy. These spheres were visible in semi-thin sections, however, where their

contents were presumably stabilized by secondary fixation with osmium tetroxide. In serial

sections of three larvae (one frontal, two sagittal), the mouth or its connection to the midgut were

not visible. The midgut epithelium was ~20-25 µm thick, and its cells filled with large spherical

vacuoles (Fig. 3C). The midget lumen was ciliated. The midgut ended blindly at the posterior

end. The hindgut and anus were not seen in serial sections.

At least some larvae were competent to metamorphose soon after release from the parental

tube, in the absence of suspended particulate food. Three hours after larvae were freed from adult

tubes and placed in a small bowl, seven of 40 had settled on the small fragments of adult tube

provided. These new juveniles had formed mucus tubes. Nineteen hours later, a total of 12 larvae

had settled on the fragments of adult tube; none had settled elsewhere in the dish. These

juveniles had already formed calcium carbonate tubes and had developed the buds of adult

.

4. DISCUSSION

The larvae of Salmacina tribranchiata clearly bear the three ciliary bands – prototroch, food groove, and metatroch – that make up an opposed band system. These ciliary bands are identifiable by position (Fig. 2), as well as by behavior (Video S1). These larvae have long been considered as nonfeeding (e.g., Kupriyanova 2001), and results of my observations of gut 12 morphology, feeding experiments, and settlement experiments are consistent with that interpretation. The observations reported here extend those of Nishi and Yamasu (1992b), whose

Figure 3C clearly show the presence of all three ciliary bands in a larva of S. dysteri, but who did not mention these cilia or their beat patterns in their paper. Together, these observations demonstrate that the presence of opposed bands of cilia in nonfeeding larvae is not restricted only to sabellids, but also to at least some serpulid species.

It is unclear how broadly distributed opposed bands are in the nonfeeding larvae of serpulids.

It is possible that they are widespread, as suggested by Kupriyanova (2001, p. 71), who noted that “in serpulids, there is no significant difference in morphology of feeding and non-feeding larvae: both the metatroch and prototroch are present in lecithotrophic larvae which have guts filled with yolk.” However, evidence supporting this assertion is sparse. For example, the only published description of the development of the larvae of Protula spp. is that of Tampi (1960), who does not describe metatroch or food groove in the text of the paper and does not show either ciliary band in the figures (but see below). No metatroch or food groove are obvious in scanning electron micrographs of the brooded, nonfeeding larvae of Paraprotis dendrova UCHIDA 1978

(Nishi and Yamasu 1992c). Larvae of spirorbid serpulids, all of which are nonfeeding, do not have obvious metatroch or food groove cilia (though at least some species have a short transverse post-oral band of cilia which might be considered a metatroch: Okuda 1946).

Despite the rarity of evidence of opposed bands in nonfeeding serpulid larvae, such larvae merit closer attention to determine if metatroch and food groove cilia are present but have been overlooked. Relatively little work has been done on the morphology of nonfeeding larvae of serpulids. Further, metatroch and food groove cilia were often overlooked by earlier workers, even where present in feeding annelid larvae (Pernet and Strathmann 2011), perhaps in part 13

because they are quite inconspicuous relative to the large, active prototroch, and perhaps in part

because their functional significance was not understood until fairly recently (Strathmann et al.

1972). Thus Kupriyanova (2001) may be correct in stating that nonfeeding larvae of serpulids

generally bear a metatroch (and perhaps also a food groove), but additional work is needed to

verify this assertion.

A specific example concerns the serpulid Protula. As noted above, Tampi (1960) did

not mention or illustrate metatroch or food groove in his brief description of the development of

P. tubularia (MONTAGU 1803). However, I have reared early embryos and larvae of Protula spp. from adults collected on the east coast of Florida and the Caribbean coast of Panama. In neither case could I identify adults to species; they may represent two different species, or they may be conspecific. In both cases, starting about two days after fertilization, larvae appeared to bear a post-oral ciliary band corresponding in position to a metatroch (Fig. 4). Unfortunately, in neither case did I have access to scanning electron or high-speed video microscopy, so could not examine these ciliary bands or their behavior in more detail. These observations do, however, suggest that metatroch and food groove might be present in more than just one genus of serpulids with nonfeeding larvae.

Whether or not the nonfeeding larvae of additional serpulid species bear opposed bands of cilia, the observations reported here and by Nishi and Yamasu (1992b) show that the presence of feeding structures in nonfeeding annelid larvae is more widely distributed than previously recognized. Metatroch and food groove cilia might be present in nonfeeding sabellid and serpulid larvae if an evolutionary loss of larval feeding happened recently; if maintenance of these structures incurs no or little cost to the larvae; if these structures have functions other than in 14 feeding; or if their presence is a pleiotropic effect of another, functionally important gene network.

None of these conditions is obviously true, however. Serpulids and sabellids are preserved in the fossil record as adult tubes, but we know of no way to infer larval nutritional mode from adult structures in these annelids (unlike in some echinoderms and mollusks: Jablonski & Lutz

1983; Emlet 1989). Thus it is difficult to make strong inferences on the timing of loss of larval feeding in particular clades. However, the timing of origin of these clades suggests the possibility of an ancient origin of nonfeeding larval development in serpulids and sabellids. The fossil record of serpulids stretches back 244 Ma to the early Mesozoic, and some of the earliest definitive serpulid fossils are classified as members of the genus Filograna (which is perhaps synonymous with Salmacina: Zibrowius 1968). All extant species of Filograna/Salmacina whose development has been described have nonfeeding larval development (Kupiyanova 2001).

The oldest sabellid fossils known are those of calcifying sabellids, present in the late Paleozoic

(~300 Ma). All known extant sabellids whose development has been described have nonfeeding larval development. These data are consistent with an ancient origin of nonfeeding larval development in these groups, though other interpretations (e.g., multiple recent parallel origins of lecithotrophy) are certainly possible.

The energetic cost of the development, maintenance, and operation of metatroch and food groove cilia is not known, but seems unlikely to be high. Operation of metatrochal cilia may incur other costs, however. Strathmann and Grunbaum (2006) suggest that in larvae with opposed bands, beating metatrochal cilia might steepen shear gradients around prototrochal cilia

(which generate a swimming current), thus reducing larval swimming speed. In feeding larvae, this might represent a trade-off between feeding and swimming performance. However, in 15 nonfeeding larvae, the beating of metatrochal cilia would hamper swimming performance without any known benefit. Nonfeeding larvae of serpulids (and sabellids) might reduce this cost by arresting metatrochal beat (Strathmann and Grunbaum 2006), which larvae of sabellids and serpulids are certainly capable of doing (Strathmann et al. 2019). It is also possible that reducing the width of the food groove reduces this cost; in nonfeeding serpulid and sabellid larvae with opposed bands, the food groove is narrower than it typically is in feeding larvae with opposed bands (Pernet 2003; this paper).

In annelids with opposed bands of cilia, there is no evidence that food groove and metatrochal cilia have any function other than in feeding. In an attempt to understand their presence in nonfeeding sabellid larvae, Pernet (2003) speculated about other possible functions of these ciliary bands, including serving as the site of uptake of dissolved organic material; playing a role in assessing potential settlement sites; being involved in forming the juvenile tube; or being used in feeding by recently settled juveniles while the definitive feeding structures develop. An additional hypothesis was suggested by R. Strathmann (pers. comm.): nonfeeding larvae might use an opposed band system to capture microbial cells that will later serve as symbionts in juveniles and adults. I know of no evidence suggesting that any of these hypotheses for alternative functions of food groove and metatrochal cilia are correct.

Larvae of Salmacina spp. and sabellids are not the only nonplanktotrophic larvae that bear functional opposed band systems. In gastropods, there are several examples of clades in which some species develop to the larval stage in benthic capsules then hatch out as feeding, planktonic larvae, and other species pass their entire larval developmental period in the capsule, hatching out as juveniles. Character reconstruction analyses usually suggest that the former strategy is ancestral, and that fully encapsulated, nonplanktonic development has been derived numerous 16 times (e.g., Collin 2004). In some of these derived species with nonplanktonic larval development, larvae retain functional opposed bands of cilia (e.g., Collin et al. 2007; Hookham and Page 2016). In at least some of these cases, however, opposed bands of cilia do seem to have an “alternative” function – instead of being used to capture planktonic food, they are used to capture maternally provided food particles in the capsule (e.g., nurse eggs: Collin et al. 2007).

This seems likely to explain the retention of food groove and metatrochal cilia in these nonplanktonic larvae.

Several workers have noted that the presence of functional particle capture structures in nonfeeding (or at least nonplanktonic) larvae might make evolutionary transitions to planktotrophy more likely. Indeed, possible examples of such transitions have been identified in calyptraeid and muricid gastropods (Collin et al. 2007; Pappalardo et al. 2014). The presence of functional opposed band systems in nonfeeding larvae might increase the likelihood of transitions to planktotrophy in sabellidan annelids, as well. An inferred origin of larval feeding in the common ancestor of one clade of serpulid annelids – the Serpulinae – may be an example of just such a transition (Kupriyanova 2003; Kupriyanova et al. 2006). Additional efforts to reconstruct sabellidan relationships (especially those within Serpulidae) and to describe larval form and function are needed to further elucidate patterns in the evolution of larval nutritional mode and larval form in the clade.

Acknowledgments. I thank Yvette Ralph and Dr. Darren Johnson for collecting adults of

Salmacina tribranchiata; Dr. Elizabeth Eldon for allowing me to use her compound microscope for imaging living larvae; Dr. Misty Paig-Tran for generously loaning me her high-speed video camera; Michael Sullivan for assistance in editing video footage; and Dr. Thomas Douglass for 17 advice and assistance in embedding and sectioning larvae. I also thank two anonymous reviewers and Dr. Greg Rouse for comments that greatly improved the manuscript. This work was supported by National Science Foundation grant OCE-1756531.

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Figure legends

Fig. 1. The relationships of Serpulidae to some other sabellidan and spionidan families, as described by Kupriyanova and Rouse (2008), Struck et al. (2015), Weigert & Bleidorn (2016), and Struck (2019). “F” and “NF” refer to the presence of feeding and nonfeeding (respectively) larvae in the clade. Note that prior analyses typically viewed the “Sabellidae” and “Fabriciidae” described here as subfamilies (Sabellinae and Fabriciinae) of the family Sabellidae sensu latu). 22

Sabellidae s.l. was generally considered sister to Serpulidae. See Kupriyanova and Rouse (2008) for a full discussion of the phylogeny of these taxa.

Fig. 2. Larvae of Salmacina tribranchiata. A. Light micrograph showing a right lateral view of an early larva. B. Light micrograph showing a dorsal view of a late (three-chaetiger) larva. C.

Scanning electron micrograph showing a right lateral view of an early larva. D. Scanning electron micrograph showing a ventral view of a late larva. E. Detail of the opposed bands from the specimen shown in C. F. Detail of the opposed bands from the specimen shown in D. fg, food groove; mo, mouth; mt, metatroch; ne, neurotroch; pt, prototroch.

Fig. 3. The digestive system of a late larva of Salmacina tribranchiata. A. A Z-projection of 11 confocal images showing a ventral view of a late larva labelled for filamentous actin. B. The same image as in A, except that the outlines of the digestive system are highlighted in red. C. A semi-thin sagittal section of a late larva. The midgut lumen is surrounded by a thick epithelium containing many spherical vacuoles. The midgut is not connected to either the foregut or hindgut. lu, midgut lumen.

Fig. 4. Larvae of Protula spp. (Serpulidae). A. A larva of an unidentified Protula sp. from

Florida. Adults were collected by dredge off of Fort Pierce, Florida, in 2002, and eggs were fertilized and larvae reared in the laboratory. The larva is approximately 2 d old. B. A 43 hr old larva of an unidentified Protula sp. from Bocas del Toro, Panama, in 2003. Adults were collected by snorkel, and eggs were fertilized and larvae reared in the lab. Exact scalebars are not 23 available for either image, but each larva is approximately 125 µm in length. mt?, possible metatroch.

Supplementary files

Video S1. High-speed video clip of prototrochal, food groove, and metatrochal cilia in a late larva of Salmacina tribranchiata. Prototrochal cilia beat continuously throughout the clip, but metatrochal and food groove cilia are sometimes beating, and sometimes arrested. The width of the food groove is ~4 µm.