Zoomorphology (2009) 128:327338DOI 10.1007/s00435-009-0093-9ORIGINAL PAPER

Larval apical sensory organ in a neritimorph gastropod, an ancient gastropod lineage with feeding larvae

Louise R. Page S. C. Kempf

Received: 5 September 2008 / Revised: 25 January 2009 / Accepted: 27 February 2009 / Published online: 27 March 2009 Springer-Verlag 2009

Abstract The Neritimorpha is an ancient clade of gastro-pods that may have acquired larval planktotrophy indepen-dently of the evolution of this developmental mode in othergastropods (caenogastropods and heterobranchs). Neri-timorphs are therefore centrally important to questionsabout larval evolution within the Gastropoda, but there isvery little information about developmental morphologythrough metamorphosis for this group. We used immunola-beling (antibodies binding to acetylated -tubulin and sero-tonin) and serial ultrathin sections for transmission electronmicroscopy to characterize the apical sensory organ inplanktotrophic larvae of a marine neritimorph. The apicalsensory organ of gastropod larvae is a highly conservedmulticellular sensory structure that includes an apical gan-glion and often an associated ciliary structure. Surprisingly,the apical ganglion of Nerita melanotragus (Smith, 1884)does not have typical ampullary neurons, a type of sensoryneuron consisting of a cilia Wlled inpocketing that has beendescribed in all other major gastropod groups. N. melanotr-agus has cilia-Wlled pockets embedded within the apicalganglion, but these so-called sensory cups are cassettes ofmultiple cells: one supporting cell and up to three multicili-ated sensory cells. We suggest that an internalized pocketthat is Wlled with cilia and open to the exterior via a narrowpore may be essential architectural features for whateversensory cues are detected by ampullary neurons and

sensory cups; however, morphogenesis of these features atthe cellular level has undergone evolutionary change. Wealso note a correlation between the number of sensory ele-ments consisting of cilia-Wlled pockets within the larvalapical sensory organ of gastropods and morphological com-plexity of the velum or length of the trochal ciliary bands.

Keywords Apical ganglion Neurodevelopment Immunolabeling Planktotrophy Electron microscopy

Introduction

Multicellular sensory complexes within major taxa of ani-mals typically carry signatures of both homology and adap-tive function. As a result, the sensory organs of marineinvertebrate larvae have attracted widespread researchinterest because their morphology can be a source of phylo-genetically relevant characters and a challenge for func-tional interpretations (Lacalli 1994; Hay Schmidt 2000;Byrne et al. 2007). Gastropod veliger larvae are well provi-sioned with multicellular sensory structures, but only thestatocysts and apical sensory organ (ASO) are typicallypresent throughout the planktonic stage of both feedingveligers (Chia and Koss 1981, 1984; Ulthe 1995; Lin andLeise 1996; Kempf et al. 1997; Marois and Carew 1997a;Dickinson et al. 1999; Page and Parries 2000; Ruthensteinerand Schaefer 2002; Dickinson and Croll 2003; LaForge andPage 2007) and non-feeding veligers (Barlow and Truman1992; Schaefer and Ruthensteiner 2001) (see Ruthensteinerand Schaefer 2002 for the only reported exception). Thissuggests that gravity perception (Chia and Koss 1981) andwhatever other sensory information may be detected bythe ASO (Page 2002b) are functionally important duringthe entire larval stage of most gastropods, regardless of the

L. R. Page (&)Department of Biology, University of Victoria, P.O. Box 3020, STN CSC, Victoria, BC V8W 3N5, Canadae-mail: lpage@uvic.ca

S. C. KempfDepartment of Biological Sciences, 331 Funchess Hall, University of Auburn, Auburn, AL 36849-5407, USA123

328 Zoomorphology (2009) 128:327338length of larval life or capacity for feeding. Furthermore,although the statocysts of gastropod larvae are carried intothe benthic juvenile and adult stages, the ASO is destroyedat metamorphosis in species where this has been studied(Lin and Leise 1996; Marois and Carew 1997b; Gifondorwaand Leise 2006) and therefore has functions unique tolarvae.

Previous studies have revealed both similarities anddiVerences for the larval ASO among members of variousgastropod clades (LaForge and Page 2007; others reviewedby Page 2002b; Ruthensteiner and Schaefer 2002), but it isoften unclear if the character states are best explained byancestry or as adaptations associated with derived larvallifestyles. This situation is exacerbated by the completelack of information about a possible ASO in larvae of theNeritimorpha (Neritopsina). The Neritimorpha is a criti-cally important group for the topic of gastropod larval evo-lution because it features two attributes that do not co-occurwithin any other gastropod clade: a relatively basalalthough unresolved position within the gastropod phyloge-netic tree (Fig. 1) (Haszprunar 1988; Ponder and Lindberg1997; Winnepenninckx et al. 1998; McArthur and Koop1999; Colgan et al. 2007; Yoon and Kim 2007) togetherwith a feeding larval stage in the life history of many mem-bers (Scheltema 1971; Bandel 1982; Kano 2006).

A number of phylogenetic reconstructions (Haszprunar1988; Ponder and Lindberg 1997) and interpretations offossil data (Ntzel et al. 2006) have suggested that feedinggastropod larvae evolved from a pelagic, but non-feeding,larval form (see also Haszprunar et al. 1995). Certainly, themajority of species within the two most derived gastropodclades, the Caenogastropoda and its sister group the Hetero-branchia (collectively known as the Apogastropoda) havefeeding larvae (Fig. 1), and larval ASOs of species in these

two groups show many similarities (reviewed by Page2002b; Ruthensteiner and Schaefer 2002). By contrast,members of more basal clades, the Patellogastropoda andVetigastropoda, have non-feeding larvae (HadWeld et al.1997), and the ASO of the patellogastopod Tectura scutum(Rathke, 1833) shows substantial diVerences from those offeeding gastropod larvae (Page 2002a). The phylogeneticposition of neritimorphs has long been problematic, butrecent information has resurrected Bournes (1908) thelong ago opinion that neritimorphs are an ancient gastropodgroup that diverged independently of other basal clades(Ntzel et al. 2007 and references cited within). If ongoingeVorts to resolve the phylogenetic tree topology for gastro-pods reinforce the claim that larval planktotrophy evolvedindependently in neritimorphs and apogastropods (i.e., neri-timorphs arose from nodes a or b in Fig. 1), then larvalmorphology should be mined for independent corrobora-tion of that hypothesis (Ponder and Lindberg 1997). Fur-thermore, if larval planktotrophy evolved at least two timesindependently from larval lecithotrophy among basal gas-tropods, then elaborations of ASO structure that coevolvedwith larval planktotrophy might provide clues about func-tional roles for this enigmatic larval structure.

We adopt Conklins (1897) term of apical sensoryorgan for the structural complex that includes both an api-cal ganglion and one or more distinctive ciliary structuresarising from epidermal cells overlying or adjacent to theganglion (Page 2002b). The ganglion includes: (1) sensoryneurons that each have a dendrite extending to the exposedepidermal surface at the anterior end of the larva, (2) addi-tional neurons that may be interneurons or motorneurons,and (3) a neuropil of interwoven neurites having large, ves-icle-Wlled varicosities. In those species previously exam-ined, the most distinctive type of sensory neuron, termedampullary receptor cell (Chia and Koss 1984), has a deepinvagination of the apical membrane to form a spaciouspocket Wlled with cilia that opens to the exterior in at leastsome species (Kempf and Page 2005). A subset of neuronalsomata and neurites within the neuropil express serotonin-like immunoreactivity (reviewed by Page 2002b) and someof these neurites extend into the ciliated velar lobes (Kempfet al. 1997; Marois and Carew 1997c; Dickinson et al.1999; Page and Parries 2000; Dickinson and Croll 2003),structures that allow swimming and feeding by veligerlarvae.

There has been much speculation about the function ofthe ASO in larval gastropods. Experimental evidence indi-cates a role in reception or processing of sensory informa-tion triggering larval metamorphosis (Lin and Leise 1996;HadWeld et al. 2000; Leise et al. 2001; Thavaradhara andLeise 2001;). Nevertheless, neurons within the apical gan-glion are among the Wrst cells to express a neuronal pheno-type during embryogenesis (Kempf et al. 1997; Marois

Fig. 1 Hypotheses on phylogenetic relationships among major gastro-pod groups showing three proposed nodes (ac) for the origin of theNeritimorpha; asterisks indicate groups with feeding larvae. a Molec-ular phylogeny of Yoon and Kim (2007). b Morphological phylogeniesof Haszprunar (1988) and Ponder and Lindberg (1997). c Molecularphylogeny of Winnepenninckx et al. (1998) and phylogeny of Lind-berg and Guralnick (2003) based on embryological cleavage patterns123

Zoomorphology (2009) 128:327338 329et al. 1997b; Dickinson et al. 1999; Dickinson and Croll2003), which suggests that the ganglion has additionalfunctions prior to metamorphosis.

We provide a Wrst ever characterization of the ASO infeeding larvae of a neritimorph gastropod and compare it tothe ASO of other gastropods for insights into its evolutionand function. We studied Nerita melanotragus (Smith,1884) collected from Sydney Harbour, Australia. In a pre-vious study on larvae obtained from this population ofneritids (Lesoway and Page 2008), the species was calledNerita atramentosa (Reeve, 1855), the conventional namefor many previous years. However, a recent taxonomicreassessment (Spencer et al. (2007) revised the speciesname to N. melanotragus.

Materials and methods

Source of animals

Small pebbles with attached egg capsules of Nerita mela-notragus (Smith, 1884) (Neritimorpha, Gastropoda) werecollected from high intertidal rock pools bordering thenorth side of Edwards Bay, Sydney Harbour, Australia.These were maintained in the laboratory within a 20 laquarium of Weld-collected seawater provided with aerationand an activated charcoal Wlter. Larvae did not hatch fromundisturbed egg capsules. Instead, capsules were mechani-cally opened and if released larvae showed active swim-ming behavior when they were placed into culture.Throughout this paper, we use the term hatching stage lar-vae for larvae that were capable of feeding when artiW-cially released from the egg capsule.

Larvae were maintained at an initial density of 0.3 larvae/ml in glass beakers containing 1000 ml coarse-Wltered sea-water at 2023C. Larval cultures were provided with a mixof Pavlova lutheri and Isochrysis sp. (TIso) at 4 104 cells/ml. Inocula of these microalgae were obtained from CSIROMicroalgae Supply Service (Tasmania, Australia) and theywere grown in Guillards f/2 enrichment medium withoutsilicates (Guillard 1975) under constant illumination. Larvaewere transferred to fresh culture medium every other dayusing a combination of sieving and hand pipetting.

Transmission electron microscopy

Prior to chemical Wxation, larvae were anesthetized usingthe method previously described by Page (2002a). Theywere then Wxed in 2.5% glutaraldehyde in 0.2 M Millonigsphosphate buVer (pH 7.6) with 0.14 M sodium chloride toadjust osmolality. Specimens were stored in this Wxative forup to 2 weeks at 68C prior to further processing. AfterdecalciWcation in a 1:1 mix of the glutaraldehyde Wxative

and 10% ethylenediaminotetraacetic acid (disodium salt)and several rinses in 2.5% sodium bicarbonate (pH 7.2), thespecimens were post-Wxed in 2% osmium tetroxide inbicarbonate buVer for 1 h at room temperature. Specimenswere then dehydrated in a graded acetone dilution seriesand embedded in Procure 812 (ProSciTech), a substitute forEpon 812.

Thick sections of 0.75 m thickness were cut with a Dia-tome histoknife and stained with a mix of methylene blue,azure II and sodium borate (Richardson et al. 1960). Imagesof sections were recorded with a digital camera mounted ona Zeiss Axioskop microscope and operated by NorthernEclipse software (version 5, Empix Imaging Inc.).

Ultrathin, non-serial sections and ribbons of serial ultra-thin sections were cut using a Diatome Ultra diamond knifeand picked-up on Formvar-coated slot grids. These werestained in ethanolic uranyl acetate followed by lead citrate.Images of sections were acquired with a Hitachi 7000 trans-mission electron microscope Wtted with an Optronics digitalcamera. Contrast, brightness and sharpness of images wereadjusted using Adobe Photoshop software (version 7). Intotal, we examined ultrathin sections through the ASO oftwo larvae at hatching stage, two at 18 dph, two at 26 dph,and one at 61 dph. Serial sections through Wve entireampullary cup neurons were imaged: three from a hatchingstage larva and two from a larva at 18 dph.

Immunolabeling and Xuorescence microscopy

Anesthetized specimens (hatching stage larvae and larvaeat 61 days post-hatching) were Wxed for 512 h at 68C infreshly prepared 4% paraformaldehyde in 0.1 M Millonigsphosphate buVer (pH 7.6) and 0.1 M sodium chloride. Theywere rinsed several times in phosphate buVered saline(PBS) + 0.1% sodium azide and stored in this solution forup to a month before being immunolabeled.

All antibodies were diluted in 20 mM phosphatebuVered saline with 5% heat inactivated goat serum, 0.1%sodium azide, and 0.1% Triton X-100. Antibody incuba-tions and rinses were done at 5C on an orbital shaker.Specimens were initially incubated for 48 h in a mixture oftwo primary antibodies: a mouse monoclonal raised againstacetylated -tubulin (Sigma, T-6793) at 1:500 dilution anda rabbit polyclonal raised against serotonin (Immunostar,20080), at 1:1,000. After rinsing, specimens were incubatedin a mixture of secondary antibodies consisting of Alexa-Xuor 488 conjugated, goat anti-mouse IgG at 1:300, andAlexaXuor 594 conjugated, goat anti-rabbit IgG at 1:500(Molecular Probes/Invitrogen). Labeled larvae were rinsed,dehydrated through an ethanol series to absolute ethanol,transferred to toluene and inWltrated with DPX mountant(Electron Microscopy Sciences). Whole, inWltrated larvaewere mounted in DPX under coverslips supported by small123

330 Zoomorphology (2009) 128:327338shards of broken coverslips and examined with a Bio-RadMRC 1024, confocal laser scanning system mounted on aZeiss Axioskop microscope. Maximum intensity projec-tions at z-intervals of 0.5 or 1.0 m were collected usingBio-Rad Lasersharp software.

A negative control for the immunolabeling procedureconsisted of specimens that were processed and visualizedin an identical manner, but without initial incubation in pri-mary antibodies. A positive control consisted of incubatingspecimens in a mouse monoclonal raised against small car-dioactive peptide (SCPB) at 1:20 dilution for the primaryantibody (Masinovsky et al. 1988), followed by incubationin the AlexaXuor 488 conjugated, goat anti-mouse IgG at1:300 dilution.

Results

Overview of larval development

During 56 weeks of larval culture, the biomineralizedshell of N. melanotragus larvae grew from 186 m(6.3 m, N = 5) to 556 m (12.1 m; N = 5) in length(maximum lateral diameter) (Lesoway and Page 2008) andwas accompanied by growth of the soft tissues. One of themost dramatic events of larval development began at2 weeks after the hatching stage, when each of the initiallysemi-circular velar lobes (Fig. 2a) began subdividing intowhat eventually became a long, dorsally projecting lobeand a shorter, ventrally projecting lobe on each side of thehead (Fig. 2b). As in all feeding larvae of gastropods, aband of long prototrochal (pre-oral) cilia and a parallelband of shorter metatrochal (post-oral) cilia ran around theperiphery of each velar lobe. These ciliary bands allow thelarvae to swim and capture microalgal food. The fastestgrowing larvae in our cultures became capable of intermit-tent crawling at 36 days after the hatching stage, a behaviormade possible by gradual enlargement and elaboration ofthe foot during larval development. Larvae allowed tocrawl on organically Wlmed pebbles collected from theadult habitat underwent metamorphosis beginning at42 days post-hatching.

Larvae of N. melanotragus had a number of multicellu-lar sensory structures. Deeply pigmented eyes, one oneither side of the head, were present at the hatching stage(Fig. 3a), but cephalic tentacles were not visible until4 weeks later. Hatching larvae also had a pair of statocystswithin the base of the foot and a small ciliated osphradiumembedded in epithelium along the left side of the roof of themantle cavity. The statocysts and osphradium were obviousin sections prepared for light microscopy, but were diYcultto resolve in live larvae. Similarly, the apical sensory organ(ASO) could not be discerned in whole mounts of live

larvae, but sectioned and immunolabeled specimensrevealed the presence of this sensory structure in all larvalstages between hatching stage and metamorphosis. Asdescribed below, the ASO consisted of a ganglion and aciliary structure.

Apical ganglion

The apical ganglion was essentially intraepithelial. Gangli-onic neurons existed as a multilayered thickening of apicalepithelium at the center of the velar Weld, immediatelybetween the two eyes (Fig. 3a). The basal surface of theganglion sat upon the cerebral commissure that connectedthe two cerebral ganglia of the developing central nervoussystem (Fig. 3a). Many cells of this intraepithelial ganglionextended narrow cytoplasmic processes, presumably den-drites, to the exposed surface of the epithelium (Fig. 3b).Each dendrite gave rise to one or two cilia. Fortuitous sec-tions revealed proWles of neurites arising from neuronalsomata of the apical ganglion (Fig. 3c). In addition, spa-cious pockets Wlled with cilia were embedded within theapical ganglion of hatching larvae (Fig. 3b). These were

Fig. 2 Apical views of live larvae of Nerita melanotragus. a Hatchingstage larva with two semi-circular velar lobes (arrowheads). b Larvaat 5 days after onset of crawling ability (43 days after hatching stage)showing elaborate velum with four elongate lobes (arrowheads) andleft cephalic tentacle (t). Both images are shown at the same scale123

Zoomorphology (2009) 128:327338 331Fig. 3 General features of the apical ganglion of Nerita melanotragusas revealed by TEM (ad) and light microscopical sections (f). a Fron-tal section through a hatching stage larvae showing the intraepithelialapical ganglion (ag) located between the eyes (e) and above the cere-bral commissure (cc). b Apical, exposed surface of the apical ganglionshowing terminals of sensory dendrites (arrowheads) and part of a cil-ia-Wlled sensory cup (sc). c Neuronal soma (asterisk) of an apical gan-glion neuron extending a neurite (arrowheads) into the apical neuropil

(anp). d Frontal section of a hatching stage larva showing neurites ofthe apical neuropil (anp) lying between the cerebral commissure (cc)and a neuronal soma of the apical ganglion (ag); a portion of the fore-gut wall (fg) is also visible. e Dense cored vesicles within a neuriticswelling of the apical neuropil. f Frontal histological section througha larva at 26 days after hatching showing the bilobed apical ganglion(ag) overlying the cerebral commissure (cc) and cerebral ganglia (cg);also note the foregut (fg)123

332 Zoomorphology (2009) 128:327338superWcially similar to ampullary neurons, a distinctiveneuronal type identiWed within the larval apical ganglion ofother gastropods (Bonar 1978; Chia and Koss 1984; Kempfand Page 2005). However, as described in greater detailbelow, these pockets had a diVerent cytoarchitecture thanthat of conventional ampullary neurons. We have thereforeelected to call them sensory cups.

A zone of distinctive neurites was evident between cellbodies of the ganglion and neurites of the cerebral commis-sure. The distinctive neurites had large swellings Wlled withdense concentrations of vesicles (Fig. 3d). The ultrastruc-tural appearance of these vesicles was variable, but oftentheir content was moderately to strongly electron dense(Fig. 3e). By contrast, the neuritic proWles of the cerebralcommissure showed fewer enlarged areas, with vesiclesand their trajectories less convoluted (Fig. 3d). We identi-Wed the zone of convoluted neurites with vesicle-Wlledswellings as the neuropil of the apical ganglion. Neuritesarising from neuronal somata of the apical ganglion enteredthis neuropil (Fig. 3c).

As larval development progressed, the number of neu-rons within the apical ganglion of N. melanotragusincreased greatly. By 26 days after hatching, the enlargingganglion had acquired the form of a bilobed cap overlyingthe cerebral ganglia and commissure (Fig. 3f). However,the ganglion remained broadly connected to the exposedsurface of the epithelium by the apical dendrites arisingfrom neuronal cell bodies.

Ciliary structure

The ciliary structure of the ASO in the hatching stage lar-vae of N. melanotragus was a discrete tuft of long, denselypacked cilia arising from the apical epithelium at the dorsalside of the apical ganglion. Cilia of this tuft were labeled byantibodies against acetylated -tubulin, as were the motilecilia of the velar lobes (Fig. 4a). The cilia had a typical9 + 2 axoneme with dynein arms on the doublet microtu-bules and they were anchored by very long, striated rootlets(Fig. 5ac). As larval growth and development proceeded,the cells expressing the apical ciliary tuft extended laterallyso as to form a long strip of cilia bordering the entire dorsalside of the bilobed apical ganglion (Fig. 4b). We failed toWnd evidence of neurites arising from the multiciliated cellsthat gave rise to the ciliary tuft and later the ciliary strip.These large ciliated cells were separated from the apicalganglion proper by a bundle of muscle Wbers extendingbetween the right and left sides of the velum (Fig. 5a, b).

Serial ultrathin sections clearly showed that the ciliarytuft of young larvae and the ciliary strip of older larvaewere not part of the prototrochal ciliary band that ran alongthe periphery of each velar lobe.

Fig. 4 Neurons within the larval apical ganglion of Nerita melanotra-gus as labeled with antibodies against acetylated -tubulin (green) andserotonin (red). a Hatching stage larva; immunolabeling identiWes thecilia-Wlled pockets of six sensory cups (asterisks) and serotonin-likeantigenicity in neurites and somata of two non-sensory neurons(arrowheads). Also note the ciliary tuft (ct); other labeled cilia are partof the prototrochal ciliary band (pt). b 61-day-old larva; many sensorycups are labeled with antibodies against -tubulin and additional non-sensory neurons expressing serotonin-like antigenicity (arrowheads)have appeared; also note the strip of dense cilia (cst) along the dorsalmargin of the ASO. c Details from (b) showing sensory cups in a61-day-old larva. Cilia can be seen within the cups123

Zoomorphology (2009) 128:327338 333Sensory cups

Cilia within the sensory cups of the larval apical ganglionof N. melanotragus were also labeled with antibodiesagainst acetylated -tubulin (Fig. 4ac). This facilitated thecounts of their number during the course of larval develop-ment. Hatching stage larvae had only six sensory cups, withthe cilia-Wlled pockets having a tubular shape (Fig. 4a). At61 days post-hatching, a linear array of over 30 sensorycups was present within the apical ganglion (Fig. 4b). Afew of the cups had a tubular shape, but most had anexpanded, globose shape (Fig. 4c). Although the cups oflater stage larvae were distributed along the full extent ofthe apical ganglion, they tended to be concentrated withintwo lateral clusters. Increase in number of sensory cupsduring the larval phase was accompanied by a marked

increase in the size of the neuropil of the apical ganglion(Fig. 5a).

Ultrastructural analysis revealed that, unlike the ampul-lary neurons within the apical ganglion of other larval gas-tropods, the cilia-Wlled pockets (sensory cups) embeddedwithin the apical ganglion of N. melanotragus larvae weremulticellular epithelial invaginations, rather than invagin-ations within single cells. As shown in the sketches com-paring a sensory cup with a typical ampullary neuron(Fig. 6a. b), the internalized pocket of each sensory cup waslined by the apices of multiple cells: a single supporting celland from one to three multiciliated cells. Each cilia-Wlledcup narrowed toward the epithelial surface and opened tothe exterior via a small pore (Fig. 5d).

Figure 7a shows a section passing through three adjacentmulti-ciliated cells lining the lumen of a sensory cup. These

Fig. 5 TEMs of a longitudinally sectioned larva of Nerita melanotra-gus at 18 days after hatching stage showing the ciliary structure andsensory cups. a Apical ganglion and adjacent multiciliated tuft cell(ctc) and ciliary tuft (ct); also note the proWles of two sensory cups (sc)within the apical ganglion overlying the apical neuropil (anp) and cere-

bral commissure (cc). a anterior, p posterior, d dorsal, v ventral. b De-tails of long ciliary rootlets (arrowheads) within a ciliary tuft cell (ctc);also note the underlying muscle bundle (m). c Details of ciliary rootletsshowing striations. d Apex of a sensory cup (sc) showing the narrowpore (arrowhead) communicating to the exterior123

334 Zoomorphology (2009) 128:327338multiciliated cells appeared to be neurons, because theygave rise to a basal neurite that entered the neuropil of theapical ganglion (Fig. 7b). Cilia arising from the multicili-ated cells of the sensory cups had an axoneme consisting ofnine outer doublet microtubules with dynein arms and twocentral singlet microtubules (non-modiWed axoneme;Fig. 7c).

Our observations on sensory cups within the apical gan-glion of N. melanotragus were mostly based on images ofserial, ultrathin sections through Wve cups: three from ahatching stage larva and two from a larva at 18 days post-hatching. The three cups from the hatching stage larvaincluded one or two multiciliated cells, whereas the twocups from the larva at 18 days post-hatching both had threemulticiliated cells.

In addition to multiciliated cells, the complex of cellsforming each sensory cup included a single supporting cell.The apical membrane of this cell formed most of the wall ofthe cup. In hatching stage larvae, the nucleus of the sup-porting cell was positioned close to the base of the cilia-Wlled cup. However, as illustrated in Fig. 6a, in larvae aged18 days post-hatching and older, the cytoplasm forming thecup was spatially more distant from the perinuclear cyto-plasm and these two regions were connected by only a nar-row cytoplasmic stalk. Electron-dense microWbers ranthrough the cytoplasm of the supporting cell (Fig. 8ac),

forming a particularly large bundle within the narrow cyto-plasmic stalk. At the apical end of the stalk, the Wber bundlebroke up into Wnger-like extensions that embraced the sen-sory cup (Fig. 8a, b). At the basal end of the stalk, the Wberbundle split into individual tracts that ran around thenucleus before anchoring on to the basal membrane of thesupport cell at the periphery of the apical ganglion(Fig. 8c).

Neurons with serotonin-like immunoreactivity

Surprisingly, only two neurons within the apical ganglionof hatching larvae of N. melanotragus were labeled withantibodies against serotonin (Fig. 4a). Both these neuronslacked a dendrite extending to the epithelial surface, sug-gesting that they were not sensory neurons. Antibodies

Fig 6 Sketches comparing the cellular architecture of sensory cupsand ampullary neurons within the apical ganglia of gastropod larvae.a Sensory cups in larvae of Nerita melanotragus; the cilia-Wlled pockethas a pore (p) to the exterior and is formed by a supporting cell (spc)containing Wber bundles (arrowheads) and the multiciliated apices ofone to three presumed sensory neurons (sn). b Ampullary neurons pre-viously described within the larval apical ganglion of caenogastropods,opisthobranchs and a patellogastropod; the pocket is a deep invagina-tion of a single sensory neuron (sn)

Fig. 7 TEMs of sensory cup cells within the apical ganglion of Neritamelanotragus at 18 days after hatching. a Three adjacent multiciliatedcells (asterisks) lining the lumen of a sensory cup. b Putative neurite(arrowheads) arising from a multiciliated cell (asterisk) of a sensorycup. c Details of cilia within a sensory cup showing unmodiWed 9 + 2axonemes123

Zoomorphology (2009) 128:327338 335against serotonin also labeled a tract of multiple neuritesthat extended between these two cell bodies and the tractscontinued laterally into the velar lobes on each side. By

61 days post-hatching, one to two additional non-sensoryneurons with serotonin-like immunoreactivity were addedto both the left and right sides and these contributed neu-rites to the tract between the two cells (Fig. 4b).

Specimens processed without primary antibodies (nega-tive control) showed no Xuorescent labeling of neurons,whereas specimens processed with primary antibodiesagainst SCPB (positive control) revealed labeled neuritesand neuronal somata associated with the pleurovisceralconnectives, as well as in neurites within the neuropils andcommissure of the cerebral ganglia (not shown).

Discussion

Comparative context for the apical sensory organ of Nerita melanotragus

Our study provides the Wrst description of the ASO in thelarval stage of a neritimorph gastropod, thereby extendingthe taxonomic occurrence of the larval ASO to all majorextant lineages of gastropods, a class of Molluscs that aroseduring the early Paleozoic (Wagner 2002), and all lineageswith planktotrophic larvae. The hypothesis of shared ances-try for this larval sensory structure is supported by: (1) sim-ilar location at the anterior pole of the larva, (2) similarjuxtaposition and connections with other structures such asthe cerebral commissure and velar lobes, (3) similar com-position of cellular phenotypes, including sensory andnon-sensory neurons contributing neurites to an associatedneuropil, (4) expression of serotonin-like antigenicity insome of the neurons and in neurites extending from theneuropil into the velum, and (5) internalized, multiciliatedstructures in which ciliated neurons extend neurites into anapical neuropil. In addition, there is frequently a close asso-ciation between the ganglion and ciliary tuft(s) arisingfrom adjacent epidermal cells (see reviews by Page 2002b;Ruthensteiner and Schaefer 2002). Opinions vary concern-ing the neuronal nature of the cells giving rise to the ciliarytufts, but to date there have been no published imagesshowing a neurite arising from the ciliary tuft cells or a syn-apse onto these cells in any gastropod larva. The ASO inlarvae of N. melanotragus exhibits all six of the foregoingcharacteristics of gastropod ASOs, yet several of its fea-tures are highly unusual.

The most extraordinary feature of the ASO of N. mela-notragus is what appears, at Wrst glance, to be a variantform of the ampullary neuron, a distinctive type of sensoryneuron within the apical ganglion of caenogastropods andheterobranchs (reviewed by Kempf and Page 2005) and atleast one species of patellogastropod (Page 2002a). Thedendritic membrane of ampullary sensory neurons is deeplyinvaginated to form a large, cilia-Wlled pocket that retains a

Fig. 8 a Base of a sensory cup (sc) in an 18-day larva of Nerita mela-notragus showing cytoplasmic Wbrous rods (arrowheads) within thesupporting cell. b Fibrous rods (arrowheads) bundling together as asingle stalk. c Fibrous rods (arrowheads) anchoring on the neurolami-na at the dorsal side of the apical ganglion123

336 Zoomorphology (2009) 128:327338narrow opening to the external environment. However, inN. melanotragus, the cilia-Wlled pockets embedded withinthe apical ganglion of the ASO are delineated by multiplecells (Fig. 6a).

At the hatching stage, larvae of N. melanotragus possesssix tubular-shaped sensory cups consisting of one support-ing cell and one to two ciliated cells (see below). As the lar-vae grow toward larval competence, additional sensorycups are added. Most of these cups have a globose mor-phology characterized by a larger number of ciliated cells,while a few cups remain tubular shaped. We suspect thatthe globose shape of most sensory cups in more mature lar-vae is a reXection of the cup morphogenesis. It is likely thatnewly formed cups consist of two to three cells and have atubular shape, whereas the globose morphology thatappears in older larvae is the result of recruitment of addi-tional ciliated sensory cells that increase the number of ciliawithin each cup as they mature.

Two other features of the ASO of N. melanotragus lackprecedent among the ASOs of other pelagic gastropod lar-vae. These are: (1) the complete absence of sensory neuronswith an apical dendritic process that express serotonin-likeimmunoreactivity within the apical ganglion, and (2) theelaboration of the ciliary tuft of hatching stage larvae into anelongate strip of cilia extending along the entire dorsal mar-gin of the apical ganglion. The only previous report of theabsence of ASO sensory neurons expressing serotonin-like immunoreactivity has been for the direct developingembryo of the trochacean vetigastropod, Lirularia succincta(Carpenter, 1864), which hatches as a crawl-away juvenile(Page 2002b). The functional signiWcance of these unique fea-tures of the ASO of N. melanotragus is currently unknown.

Evolutionary derivation and phylogenetic signiWcance of sensory cups

Each sensory cup is a cassette of several multi-ciliated sen-sory cells and a supporting cell that collectively delineate acilia-Wlled pocket that has a pore to the external environ-ment. A spacious pocket that is open to the environmentand Wlled with presumed sensory cilia is also characteristicof ampullary neurons, but the pocket of ampullary neuronsis formed within a single cell. A previous study on thepatellogastropod Tectura scutum identiWed a single, unicel-lular ampullary neuron within the larval apical ganglion(Page 2002a), although no ampullary neurons were foundin cited unpublished observations on the patellogastropodPatella caerulea Linnaeus, 1857; Haszprunar et al. (2002).It is possible that the sensory cups of N. melanotragus andthe ampullary neurons of other gastropod larvae are derivedfrom a common ancestral sensory cell, with evolutionarydivergence resulting in variation in the construction of thecilia-Wlled pocket. Under this scenario, an internalized

pocket lined by sensory cilia and having an external pore iscritical to function, but how cellular components aredeployed to form this cytoarchitectural feature is not criti-cal and has undergone evolutionary divergence.

Alternatively, the sensory cups of N. melanotragus maybe iterated homologs of the so-called apical pit, as previ-ously identiWed within the apical ganglion of the patello-gastropod, Tectura scutum (Page 2002a). Although anapical pit has also been described for older larvae of twocaenogastropods (Page and Parries 2000), these latter sortsof pits are formed when the entire apical epidermis pene-trated by sensory dendrites of the apical ganglion descendsinto a pocket. This is qualitatively diVerent from the apicalpits of the patellogastropod T. scutum and the sensory cupsof N. melanotragus. Instead, the apical pit of the patellogas-tropod T. scutum and the sensory cups of N. melanotragusare inpocketings of discrete cassettes of cells; they do notinvolve the entire apical epithelium.

If planktotrophic gastropod larvae evolved from a non-feeding larval form that was similar to that of extant patel-logastropods, as represented by T. scutum, then the transitionmay have been accompanied by great proliferation of apicalganglion sensory neurons having a cilia-Wlled pocketdesign. However, the signiWcant structural diVerencesbetween ampullary neurons of apogastropods and the sen-sory cups of N. melanotragus should be recognized as amajor diVerence between planktotrophic larvae of apogas-tropods and neritimorphs and may be a signature ofindependent origin of larval planktotrophy in these two gas-tropod clades. Nevertheless, interspeciWc diVerences indetails of ASO structure are certainly not suYciently pro-found to suggest that the whole ASO evolved indepen-dently within individual clades of gastropods.

The functional signiWcance of an increase in sensory ele-ments having a cilia-Wlled pocket design in conjunctionwith the transition to planktotrophy among gastropod lar-vae is not clear. However, the extent of the increase appearsto be correlated with the size of the velum and length of theprototrochal ciliary band running along the periphery of thevelar lobes. Patellogastropod larvae have the smallestvelum among pelagic gastropod larvae and their apical gan-glion includes only one ampullary neuron and one apicalpit. Planktotrophic opisthobranch and pulmonate larvaehave larger velar lobes, but their larval shell sizes betweenhatching and metamorphic competence bracket those ofpatellogastropods (HadWeld and Miller 1987). This grouphas four to six ampullary neurons within the larval apicalganglion (Chia and Koss 1984; Marois and Carew 1997a;Schaefer and Ruthensteiner 2001;Ruthensteiner and Schaefer2002; Kempf and Page 2005; LaForge and Page 2007).Caenogastropods may hatch at small size, but they growmuch larger and the prototrochal ciliary band displays allo-metric increase in length (Lesoway and Page 2008).123

Zoomorphology (2009) 128:327338 337Ampullary neurons in at least one species of caenogastro-pod increase in number from 8 to 10 at hatching to 20 atonset of metamorphic competence (Dickinson and Croll2003). In parallel with the caenogastropod situation, theprototrochal ciliary band of N. melanotragus (formerlycalled N. atramentosa) also shows allometric growth(Lesoway and Page 2008; Fig. 2a, b) and the sensory cupsincrease from 6 to 30 during the course of the larval phase.The correlation between prototroch length or complexity ofvelar shape (i.e., more than two lobes) and number of cilia-Wlled pockets within the ampullary ganglion may havefunctional signiWcance, particularly if further studies con-Wrm an independent origin and thus convergence for thesecorrelated traits in gastropod larvae. A parsimonioushypothesis is that ampullary neurons and sensory cups areinvolved in the reception of sensory stimuli that are criticalfor the proper modulation of the activities of velar cilia and/or musculature. Larger and more complex velar lobesrequire greater modulatory innervation and, thus, a largernumber of sensory receptors in the form of ampullary neu-rons or sensory cups.

Acknowledgments We thank Dr. Maria Byrne, University of Syd-ney, Australia for generously allowing use of her lab facilities withinthe Discipline of Anatomy and Histology, where larval neritimorphswere cultured. Mr. Brent Gowen did an expert job of cutting serialultrathin sections used in this study. Funding for this research was pro-vided by a Discovery Grant to LRP from NSERC of Canada.

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Larval apical sensory organ in a neritimorph gastropod, an ancient gastropod lineage with feeding larvaeAbstractIntroductionMaterials and methodsSource of animalsTransmission electron microscopyImmunolabeling and Xuorescence microscopy

ResultsOverview of larval developmentApical ganglionCiliary structureSensory cupsNeurons with serotonin-like immunoreactivity

DiscussionComparative context for the apical sensory organ of Nerita melanotragusEvolutionary derivation and phylogenetic signiWcance of sensory cups

References

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