Abstract

Forming a small group of mainly marine meiofaunal slugs, the Acochlidia have recently been separated from the traditional opisthobranch gastropods and placed within a mixed clade of pulmonates, Sacoglossa and Pyramidelloidea on the basis of molecular data. In the light of this new phylogenetic framework, we examined several populations of a comparatively giant Strubellia (Acochlidiidae s. l.) found in rivers of the Solomon Islands and Vanuatu, combining microanatomical and molecular methods (interactive three-dimensional models are given in the online version). Novel features include an extended set of nerves, a ‘cephalic gland’ of unknown function and an osphradium, all detected here for the first time in Acochlidia. The protandric genital system is characterized by three receptacles in the male phase, a possibly secondary open seminal groove and a complete reduction of the elaborate cephalic copulatory apparatus during ontogeny. Combined evidence from copulatory features and DNA sequences indicate a specific separation between the type species S. paradoxa (Strubell, 1892) from Ambon and the eastern Melanesian Strubellia wawrai n. sp. Live observations show the species to feed on the highly mineralized egg capsules of limnic Neritidae using a special piercing radula. Limnic Pacific acochlidians are suggested to be amphidromic, as are their prey organisms. A unique type of adhesive larva, observed in an Acochlidium species, indicates a possible dispersive stage in Acochlidiidae. Molecular phylogeny confirms the morphology-based placement of Strubellia as sister taxon to other Acochlidiidae.

INTRODUCTION

The Acochlidia consist of about 30 described species of heterobranch slugs that are characterized by a rather uniform external morphology, showing a freely projecting and uncurled visceral sac (giving the order its name) and one or two pairs of head appendages. For long time considered as one of the classic orders of the ‘Opisthobranchia’, morphological studies have repeatedly failed to place the taxon conclusively (e.g. Dayrat & Tillier, 2002; Wägele & Klussmann-Kolb, 2005) and molecular studies of Heterobranchia have cast further doubt on this classification (Klussmann-Kolb et al., 2008). The most recent molecular studies with a direct focus on the group have consistently retrieved Acochlidia in a new monophylum comprising the Sacoglossa, Pyramidelloidea and the ‘pulmonate’ groups (all together called Panpulmonata), with acochlidians (including the recently described Aitengidae; Swennen & Buatip, 2009; Neusser et al., 2011a) as sister group to Eupulmonata (Jörger et al., 2010a). However, morphological synapomorphies of the panpulmonate group have not yet been identified.

Most acochlidian species are tiny inhabitants of worldwide marine interstitial sand habitats (Arnaud, Poizat & Salvini-Plawen, 1986). Internal phylogenetic relationships derived from morphology indicate a basal split into the completely meiofaunal Microhedylacea and partially meiofaunal Hedylopsacea, a relationship that has been confirmed by recent molecular approaches (Wawra, 1987; Jörger et al., 2010a; Schrödl & Neusser, 2010). The hedylopsaceans also contain—uniquely among shell-less Gastropoda—two independent lineages that have colonized freshwater streams of tropical volcanic islands: the minute Caribbean Tantulum elegansRankin, 1979 (from St Vincent; see Neusser & Schrödl, 2007) and the radiation of comparatively giant Indo-Pacific Acochlidiidae (sensuArnaud et al., 1986). The latter family comprises the genera Acochlidium and Strubellia, the first acochlidians discovered by the Austrian naturalist A. Strubell (1892); the type species for both genera were described from a stream on the island of Ambon (Amboina) in the Molucca archipelago of Indonesia (Bücking, 1933; Küthe, 1935). Together with the enigmatic PalliohedyleRankin, 1979, several acochlidiid species have been described from island streams of Indonesia, Palau, the Solomon Islands and Fiji (Bergh, 1895; Bayer & Fehlmann, 1960; Wawra, 1979, 1980; Haynes & Kenchington, 1991; own unpublished data).

Since the discovery of Strubellia paradoxa (Strubell, 1892) on Ambon (Küthe, 1935; original material redescribed by Brenzinger et al., 2011), populations of Strubellia have been discovered some 3,500 km away on Guadalcanal, Solomon Islands (Starmühlner, 1976). This geographically separate population was described as the “rediscovery of Strubellia paradoxa” by Wawra (1974, 1988). Further examinations of island stream malacofauna showed the genus to occur even further south on Efate and Espiritu Santo Islands, both Vanuatu (Haynes, 2000; present study). In all locations, Strubellia is known to share its habitat with numerous limnic Neritidae and can be found hiding under calcareous rocks in brackish water from close to the river's mouth to as far as 5 km upstream. A fifth population is presently known only from a single juvenile collected on Sulawesi, Indonesia (present study).

The Indo-Pacific limnic species are generally large-bodied (crawling individuals are up to at least 4 cm long, compared to the millimetre-scale marine mesopsammic acochlidians); they should thus be ideal candidates in the search for shared morphological characters uniting Acochlidia and their panpulmonate relatives. They are also relatively easy to keep in an aquarium; observations on their biology are nevertheless scarce and mostly limited to descriptions of habitat. Life history is unknown except for the observation that Acochlidium veligers do not survive in fresh water (Haynes & Kenchington, 1991; own observations). Assuming an amphidromous lifestyle as in many other invertebrates found in similar habitats (see McDowall, 2007; Kano, 2009), the questions how metamorphosed individuals manage to return and maintain reproductive populations, or how they have colonized widely separated islands, remain unanswered.

We observed and examined numerous specimens from Guadalcanal and Vanuatu, using three-dimensional (3D) microanatomical reconstruction from serial semithin sections and scanning electron microscopy (SEM). Molecular data from Strubellia specimens from all five known localities and from closely related hedylopsacean taxa were compared in order to reveal their origin and relationships. Based on morphological and molecular evidence, the eastern Melanesian Strubellia is described as a new species and the evolution of the genus is discussed in the light of these new data.

MATERIAL AND METHODS

Collection and cultivation

About 90 specimens of Strubellia wawrai n. sp. were collected on northwestern Guadalcanal, Solomon Islands, in October 2007; further specimens from Espiritu Santo Island, Vanuatu, were collected during the Santo Expedition in September 2006 (see Table 1 for collection localities). All specimens were collected by hand in shallow water of freshwater streams flowing into the sea. The slugs were most commonly found aggregating in small groups on the underside of loose limestone rocks at the river's edge, up to 5 km upstream. In most places the rocks showed covering of algae; freshwater neritids were abundant in most places.

Table 1.

Collection localities of Strubellia wawrai n. sp. on Guadalcanal, Solomon Islands (1–4) and Espiritu Santo, Vanuatu (5–8).

Number Locality Coordinates 
Mataniko River, near Tavaruhu (3 km upstream) S 9°27.377′, E 159°57.447′ 
Mataniko River, near Tavaruhu (3.5 km upstream) S 9°27.517′, E 159°57.490′ 
Kohove River, Tanasawa bridge (at sea level) S 9°25.333′, E 159°54.164′ 
Lungga River, near Mbetikama (6 km upstream) S 9°26.916′, E 160°02.448′ 
Wounaouss River, Tapuntari Cascades (800 m upstream) S 15°34.320′, E 167°00.159′ 
Puelapa River (Rowa River, 200 m upstream) S 15°34.664′, E 167°01.902′ 
Wenoui River (350 m upstream) S 15°34.826′, E 167°02.879′ 
Adson River (5 km upstream) S 15°33.397′, E 166°58.112′ 
Number Locality Coordinates 
Mataniko River, near Tavaruhu (3 km upstream) S 9°27.377′, E 159°57.447′ 
Mataniko River, near Tavaruhu (3.5 km upstream) S 9°27.517′, E 159°57.490′ 
Kohove River, Tanasawa bridge (at sea level) S 9°25.333′, E 159°54.164′ 
Lungga River, near Mbetikama (6 km upstream) S 9°26.916′, E 160°02.448′ 
Wounaouss River, Tapuntari Cascades (800 m upstream) S 15°34.320′, E 167°00.159′ 
Puelapa River (Rowa River, 200 m upstream) S 15°34.664′, E 167°01.902′ 
Wenoui River (350 m upstream) S 15°34.826′, E 167°02.879′ 
Adson River (5 km upstream) S 15°33.397′, E 166°58.112′ 

Living specimens were observed in petri dishes. Four specimens from Kohove River, Guadalcanal, were kept alive for several months in a small and shallow glass aquarium with a few flat rocks. Water was regularly replenished with tap water that had been allowed to stand for several days beforehand; the aquarium was ventilated by an aerating pump. Specimens were fed different types of fish feed, egg masses of Physa snails and egg capsules of freshwater neritids (Neritina cf. natalensis). The neritids were acquired from a zoo store and kept in a separate aquarium with added pieces of wood; chips of wood with freshly laid egg capsules were placed with the Strubellia specimens. Photographs of feeding specimens were made through a stereo microscope using a handheld digital camera.

For further studies, specimens were anaesthetized using menthol crystals sprinkled onto the water surface, fixed in 1.5% glutardialdehyde buffered with 0.2 M sodium cacodylate (pH 7.2) and stored in 75% ethanol for histological study or 96% ethanol for molecular analysis.

Serial sectioning and 3D reconstruction

Glutardialdehyde-fixed specimens were postfixed in 0.01 M cacodylate buffer/0.35 M sucrose (pH 7.2) and 1% osmium tetroxide. After decalcifying in 1% ascorbic acid, specimens were dehydrated in a graded acetone series and infiltrated overnight with Spurr's low-viscosity epoxy resin (Spurr, 1969) diluted with one part 100% acetone. Infiltrated specimens were placed on embedding grids, covered with pure epoxy resin and left to polymerize for 24 h at 60°C.

Serial sections of 1.5 µm were cut with Ralph glass knives (first half of series ZSM Mol-20071895) or a Histo Jumbo diamond knife (Diatome, Biel, Switzerland—all other series) with a Microm HM 360-rotation microtome (Zeiss, Germany) (Table 2). Serial sections were collected on cleaned microscopy slides, stained with methylene blue/azure II (Richardson, Jarett & Finke, 1960) and sealed with araldite. Slides were then mapped from 600-dpi greyscale scans; single sections were photographed through a Leica DMB-RBE microscope (Leica Microsystems, Wetzlar, Germany) with mounted Spot CCD camera (Spot Insight, Diagnostic Instruments, Sterling Heights, MI, USA). Series of photographs were downsized to c. 400 megabytes by conversion to 8-bit greyscale and a resolution of 800 × 600 pixels and then imported to amira 4.1 software (TGS Europe, Mercury Computer Systems, Mérignac, France) for 3D reconstruction. Labeling of organ systems was done manually, with interpolation and surface-smoothing features applied to create 3D surfaces, in general following the method described by Ruthensteiner (2008). Reconstructions of four specimens are used herein: one ‘male’ from Vanuatu (every eighth section was photographed for the model, resulting in a virtual stack of 871 photos; Figs 4A; 9C–E), one ‘female’ from the Solomon Islands (693 photos, every 4th; Figs 4E; 9A, B, F) and two further specimens for the CNS (Solomon Islands: 439 photos, every section photographed, Fig. 4B, D, F; Vanuatu: 479 photos, every 2nd; Fig. 4C). All sections are deposited in the Mollusca Department, Bavarian State Collection of Zoology, Munich, Germany (see Table 2 for museum numbers).

Table 2.

Material used for morphological and phylogenetic analyses.

Species Locality Museum number of voucher and use of specimens
 
Strubellia wawrai n. sp. Solomons, loc. 1 ZSM Mol-20071895 (used for 3D); 20071881, 20071883, 20071886, 20071887, 20071890 (further serial sections) 
 Solomons, loc. 2 ZSM Mol-20071796 (entire lot used for SEM) 
 Solomons, loc. 3 ZSM Mol-20071894 (used for 3D); 20071877, 20071880, 20071892 (further serial sections) 
 Vanuatu, loc. 5 ZSM Mol-20071105 (used for 3D) 
 Vanuatu, loc. 6 ZSM Mol-20071106 (used for 3D) 
 
  Museum number of voucher
 
DNA voucher DNA Bank
 
GenBank accession number
 
    16S rRNA COI 
 Solomons, loc. 3 ZSM Mol-20080014 AB34404271 JF819728* JF819756* 
 Solomons, loc. 3 ZSM Mol-20080015 AB34404208 JF819729* JF819757* 
 Solomons, loc. 3 ZSM Mol-20080016 AB34404250 JF819730* JF819758* 
 Solomons, loc. 1 ZSM Mol-20080017 AB34404264 JF819731* JF819759* 
 Solomons, loc. 1 ZSM Mol-20080018 AB34404255 JF819732* JF819760* 
 Solomons, loc. 1 ZSM Mol-20080019 AB34404256 JF819733* JF819761* 
 Solomons, loc. 4 ZSM Mol-20071810 AB34404212 JF819734* JF819762* 
 Vanuatu, loc. 7 ZSM Mol-20071117 AB34404234 JF819735* JF819763* 
 Vanuatu, loc. 7 ZSM Mol-20080150 AB34404205 JF819736* JF819764* 
 Vanuatu, loc. 5 ZSM Mol-20080072 AB34404207 JF819737* — 
 Vanuatu, loc. 5 ZSM Mol-20080148 AB34404251 JF819738* — 
Strubellia paradoxa Kemeri, Ambon, Indonesia Berlin Moll 193943 AB35081823 JF819739* — 
Watatiri, Ambon, Indonesia Berlin Moll 193944 AB34858174 HQ168419 HQ168457 
Strubellia sp. Tambala River, Manado, Sulawesi, Indonesia ZSM-Mol 20100339 AB35081762 JF819740* JF819765* 
Palliohedyle sp. Tambala River, Manado, Sulawesi, Indonesia ZSM-Mol 20100356 AB35081794 JF828040 JF828032 
Acochlidium fijiense Lami River, Viti Levu, Fiji ZSM-Mol 20080063 AB34404244 HQ168420 HQ168458 
Pseudunela espiritusanta SE Espiritu Santo, Vanuatu ZSM-Mol 20080117 AB34404289 JF819750 JF819775 
Pseudunela marteli Neusser Oyster Island, Vanuatu ZSM-Mol 20080393 AB35081809 HQ168418 HQ168456 
Hedylopsis ballantinei ‘INMO’ Reef, Dahab, Egypt, Red Sea ZSM-Mol 20090244 AB34858170 HQ168416 HQ168454 
Species Locality Museum number of voucher and use of specimens
 
Strubellia wawrai n. sp. Solomons, loc. 1 ZSM Mol-20071895 (used for 3D); 20071881, 20071883, 20071886, 20071887, 20071890 (further serial sections) 
 Solomons, loc. 2 ZSM Mol-20071796 (entire lot used for SEM) 
 Solomons, loc. 3 ZSM Mol-20071894 (used for 3D); 20071877, 20071880, 20071892 (further serial sections) 
 Vanuatu, loc. 5 ZSM Mol-20071105 (used for 3D) 
 Vanuatu, loc. 6 ZSM Mol-20071106 (used for 3D) 
 
  Museum number of voucher
 
DNA voucher DNA Bank
 
GenBank accession number
 
    16S rRNA COI 
 Solomons, loc. 3 ZSM Mol-20080014 AB34404271 JF819728* JF819756* 
 Solomons, loc. 3 ZSM Mol-20080015 AB34404208 JF819729* JF819757* 
 Solomons, loc. 3 ZSM Mol-20080016 AB34404250 JF819730* JF819758* 
 Solomons, loc. 1 ZSM Mol-20080017 AB34404264 JF819731* JF819759* 
 Solomons, loc. 1 ZSM Mol-20080018 AB34404255 JF819732* JF819760* 
 Solomons, loc. 1 ZSM Mol-20080019 AB34404256 JF819733* JF819761* 
 Solomons, loc. 4 ZSM Mol-20071810 AB34404212 JF819734* JF819762* 
 Vanuatu, loc. 7 ZSM Mol-20071117 AB34404234 JF819735* JF819763* 
 Vanuatu, loc. 7 ZSM Mol-20080150 AB34404205 JF819736* JF819764* 
 Vanuatu, loc. 5 ZSM Mol-20080072 AB34404207 JF819737* — 
 Vanuatu, loc. 5 ZSM Mol-20080148 AB34404251 JF819738* — 
Strubellia paradoxa Kemeri, Ambon, Indonesia Berlin Moll 193943 AB35081823 JF819739* — 
Watatiri, Ambon, Indonesia Berlin Moll 193944 AB34858174 HQ168419 HQ168457 
Strubellia sp. Tambala River, Manado, Sulawesi, Indonesia ZSM-Mol 20100339 AB35081762 JF819740* JF819765* 
Palliohedyle sp. Tambala River, Manado, Sulawesi, Indonesia ZSM-Mol 20100356 AB35081794 JF828040 JF828032 
Acochlidium fijiense Lami River, Viti Levu, Fiji ZSM-Mol 20080063 AB34404244 HQ168420 HQ168458 
Pseudunela espiritusanta SE Espiritu Santo, Vanuatu ZSM-Mol 20080117 AB34404289 JF819750 JF819775 
Pseudunela marteli Neusser Oyster Island, Vanuatu ZSM-Mol 20080393 AB35081809 HQ168418 HQ168456 
Hedylopsis ballantinei ‘INMO’ Reef, Dahab, Egypt, Red Sea ZSM-Mol 20090244 AB34858170 HQ168416 HQ168454 

The table lists the species names, collecting localities (number refers to Table 1), reference numbers of museum vouchers (ZSM, Bavarian State Collection of Zoology; Berlin, Museum of Natural History, Berlin), DNA vouchers deposited in the DNA Bank of the ZSM and GenBank accession numbers. Numbers in italics indicate designated paratypes; asterisks mark the sequences generated for the present study.

Interactive 3D model

The interactive 3D models in the online PDF version were prepared according to Ruthensteiner & Heß (2008), although using the 3D tools of Deep Exploration v. 5.5 (Right Hemisphere EMEA, Germany) and Adobe Acrobat v. 9.0 Professional Extended (Adobe Systems GmbH, Germany) to create interactive models of the original Amira surface files. Separate surface files of each organ were exported into the former program, then grouped into a complex model and rendered. An interactive figure was then created by importing these rendered models as backdrops of Figure 4; different views of the organ systems were prefabricated to allow the reader rapidly to get a general idea of the models. Click on the interactive Figure 4A–D for models of the general anatomy and on Figure 4E, F for a more detailed model of the CNS.

Scanning electron microscopy

Several specimens were dissected and spicules, radulae and copulatory stylets were removed and cleared from tissue in diluted KOH or Proteinase K (20 µl in 180 µl ATL Tissue lysis buffer; Qiagen, Hilden, Germany; after Holznagel, 1998). The undissolved sheath of radulae was removed using tungsten minutien needles before flattening the radula. After rinsing with distilled water, samples were mounted on aluminum stubs with sticky carbon tabs, sputter coated with gold (120 s at 2.4 kV) and examined in a LEO 1430 VP scanning electron microscope (15 kV; 2 × 10–5 mbar).

Molecular analysis

Genomic DNA was extracted from tissue samples of the foot or entire specimens using the DNeasy Blood and Tissue Kit (Qiagen), according to the manufacturer's instructions. Two mitochondrial markers, partial 16S rRNA (400 bp) and cytochrome c oxidase subunit I (COI; 650 bp), respectively, were amplified using PCR (for PCR protocols and primers, see Table 3). PCR products were purified using ExoSapIT (USB, Affymetrix, Inc.); cycle sequencing and the sequencing reaction were performed by the sequencing service of the Department of Biology Genomic Service Unit (GSU) of the Ludwig-Maximilians-University Munich, using Big Dye 3.1 kit and an ABI 3730 capillary sequencer. All fragments were sequenced on forward and reverse strand. DNA vouchers are stored at the DNAbank of the Bavarian State Collection of Zoology; sequences are deposited at GenBank (see Table 2 for accession numbers). Sequences were edited using Sequencer (Gene Codes Corporation). We applied a Blast search (Altschul et al., 1990) on each sequence to check for potential contamination (http://blast.ncbi.nlm.nih.gov/Blast.cgi). MUSCLE v. 3.8.31 (Edgar, 2004) was used to create the alignments of each marker, subsequently the COI alignment was checked manually according to the translation into amino acids. Maximum-likelihood analyses of the concatenated dataset (in two partitions) were performed using RAxML v. 7.0.3 (Stamatakis, 2006) under the GTR + G model (selected for the concatenated dataset under the Akaike information criterion with jModeltest; Posada, 2008) and 1,000 bootstrap replicates were generated. Outgroups were chosen according to previous morphological and molecular hypotheses on acochlidian phylogeny (Jörger et al., 2010a; Schrödl & Neusser, 2010) and retrieved from GenBank (Table 2). Hedylopsis ballantineiSommerfeldt & Schrödl, 2005 was defined as outgroup.

Table 3.

PCR protocols and primers used for the sequences generated within this study.

Gene Primer Sequence 5′–3′ Reference PCR program 
16S 16S-H CGC CTG TTT ATC AAA AAC AT Simon et al. (1994) 98°C 30 s (98°C 5 s, 48–55°C 5 s, 72°C 25 s) × 35–40, 72°C 60 s (Phire polymerase, New England Biolabs) 
16S-R CCG GTC TGA ACT CAG ATC ACG T Simon et al. (1994) 
COI LCO1490 GGT CAA CAA ATC ATA AAG ATA TTG G Folmer et al. (1994) 94°C 3 min (94°C 60 s, 45–48°C 60 s, 72°C 90 s) × 35–40, 72°C 3 min (Taq polymerase, Sigma) 
HCO2198 TAA ACT TCA GGG TGA CCA AAA AAT CA Folmer et al. (1994) 
COI long r TAA AGA AAG AAC ATA ATG AAA ATG Stothard & Rollinson (1997) 
Gene Primer Sequence 5′–3′ Reference PCR program 
16S 16S-H CGC CTG TTT ATC AAA AAC AT Simon et al. (1994) 98°C 30 s (98°C 5 s, 48–55°C 5 s, 72°C 25 s) × 35–40, 72°C 60 s (Phire polymerase, New England Biolabs) 
16S-R CCG GTC TGA ACT CAG ATC ACG T Simon et al. (1994) 
COI LCO1490 GGT CAA CAA ATC ATA AAG ATA TTG G Folmer et al. (1994) 94°C 3 min (94°C 60 s, 45–48°C 60 s, 72°C 90 s) × 35–40, 72°C 3 min (Taq polymerase, Sigma) 
HCO2198 TAA ACT TCA GGG TGA CCA AAA AAT CA Folmer et al. (1994) 
COI long r TAA AGA AAG AAC ATA ATG AAA ATG Stothard & Rollinson (1997) 

For both markers, intra- and inter-specific variation was evaluated using Species Identifier, available from TaxonDNA (http://taxondna.sourceforge.net; Meier et al., 2006) and used to cluster sequences based on pairwise distances (testing thresholds from 1 to 10%). Additionally, we calculated haplotype networks for both markers using TCS 1.21 (Clement, Posada & Crandall, 2000); the COI alignment was shortened, until all sequences had the same length; default settings (95% probability of parsimony) were used.

SYSTEMATIC DESCRIPTION

Heterobranchia sensuHaszprunar, 1985a

Panpulmonata Jörger et al., 2010a

Acochlidia sensu Wawra, 1987

Hedylopsacea sensuWawra, 1987

ACOCHLIDIIDAE sensuArnaud et al., 1986

StrubelliaOdhner, 1937

Strubellia wawrai n. sp.

Type material: Holotype: ZSM Mol-20100718; complete specimen stored in 75% ethanol; 7 mm preserved body length; collected in Mataniko River, Guadalcanal, Solomon Islands (locality 1, Table 1), 8/9 October 2007 by K. Jörger & Y. Kano. Paratypes: nine complete specimens stored in 75% ethanol (lot: ZSM Mol-20071797), same lot as the holotype; six serially sectioned specimens mounted on microscope slides [Mataniko River ZSM Mol-20071881, 20071883 (partial series), 20071886, 20071895; Kohove River: 20071892 (partial series), 20071894]; all paratypes collected 8/9 October 2007, together with holotype (Table 2).

Etymology: Named in honour of Erhard Wawra (1945–1994) for his pioneering work on the biology and systematics of Acochlidia and particularly the Strubellia of the Solomon Islands.

Interactive model: In addition to the 3D images (Figs 4, 9), see also the interactive 3D models of Strubellia wawrai n. sp. that can be accessed by clicking onto Figure 4A–D (general anatomy) and E, F (CNS) in the online PDF version of this article.

External morphology: External appearance is of a typical hedylopsacean acochlidian: elongate head–foot complex with two pairs of pointed head appendages; foot separated from body by longitudinal cephalopedal groove; uncoiled, shell-less visceral sac projecting freely behind foot, especially in fully grown specimens (Fig. 1). Epidermis appearing velvety smooth under stereo microscope; visceral sac slightly grainier. Body coloration orange to rusty brown in living specimens; foot, head appendages and translucent patch above the eye (Fig. 1A, E: ew) brighter, pale yellow; large specimens appear darker. Eyes visible externally as black dots, digestive gland as orange tube. Spicules in foot and head appendages visible as refracting bodies. Osphradium a keyhole-shaped brighter spot on right side of head–foot (Fig. 1A). Alcohol-fixed material light yellow-brown.

Figure 1.

Live specimens and general schematic overview of the anatomy of Strubellia wawrai n. sp. A–D. External morphology of living specimens from Kohove River, Guadalcanal, Solomon Islands (A–C) and Tapuntari Cascades, Wounaouss River, Espiritu Santo, Vanuatu (D). A. Young specimen, c. 8 mm, right view. B. 20 mm specimen, dorsal view. C. Juvenile feeding on egg capsule of Neritina cf. natalensis attached to wood (experimental setting). D. Adult, at least 30 mm, dorsal view. E. Overview of external morphology, based on young specimen A, right view. F. Composite of internal anatomy, female phase. Abbreviations: an, anus; ao, aorta; au, auricle; bg, buccal ganglion; bm, buccal mass; cg, cerebral ganglion; cgl, “cephalic gland”; cpg, cephalopedal groove; dg, digestive gland; dp, diaphragm separating body cavities of head–foot complex and visceral sac; es, esophagus; ey, eye; ew, translucent patch over eye (‘eye-window’); fgl, female gland mass; ft, foot; go, gonad; gp, genital pore; hf, head–foot complex; kd, kidney; lt, labial tentacle; mb, anterior border of mantle; mh, mantle ‘hood’; mo, mouth opening; nd, nephroduct; np, nephropore; ogl, oral glands; osp, osphradium; ot, oral tube; pc, pericardium; pg, pedal ganglion; r, radula; rh, rhinophore; rpd, renopericardioduct funnel; sgl, salivary glands; sp, salivary pump; spc, spicule; vs, visceral sac; vt, ventricle. Arrowheads: position of nephropore/anus (left) and genital pore (right).

Figure 1.

Live specimens and general schematic overview of the anatomy of Strubellia wawrai n. sp. A–D. External morphology of living specimens from Kohove River, Guadalcanal, Solomon Islands (A–C) and Tapuntari Cascades, Wounaouss River, Espiritu Santo, Vanuatu (D). A. Young specimen, c. 8 mm, right view. B. 20 mm specimen, dorsal view. C. Juvenile feeding on egg capsule of Neritina cf. natalensis attached to wood (experimental setting). D. Adult, at least 30 mm, dorsal view. E. Overview of external morphology, based on young specimen A, right view. F. Composite of internal anatomy, female phase. Abbreviations: an, anus; ao, aorta; au, auricle; bg, buccal ganglion; bm, buccal mass; cg, cerebral ganglion; cgl, “cephalic gland”; cpg, cephalopedal groove; dg, digestive gland; dp, diaphragm separating body cavities of head–foot complex and visceral sac; es, esophagus; ey, eye; ew, translucent patch over eye (‘eye-window’); fgl, female gland mass; ft, foot; go, gonad; gp, genital pore; hf, head–foot complex; kd, kidney; lt, labial tentacle; mb, anterior border of mantle; mh, mantle ‘hood’; mo, mouth opening; nd, nephroduct; np, nephropore; ogl, oral glands; osp, osphradium; ot, oral tube; pc, pericardium; pg, pedal ganglion; r, radula; rh, rhinophore; rpd, renopericardioduct funnel; sgl, salivary glands; sp, salivary pump; spc, spicule; vs, visceral sac; vt, ventricle. Arrowheads: position of nephropore/anus (left) and genital pore (right).

Crawling specimens usually between 6 and 12 mm, up to 20 mm (Solomon Islands specimens; Fig. 1B) or 35 mm long (Vanuatu; Fig. 1D). In younger specimens, visceral sac straight and slightly shorter than foot with foot tip visible in dorsal view; larger specimens with visceral sac longer and appearing somewhat ragged and bent, with tip often pointing to right side. Pericardial space and beating of heart sometimes visible (‘heart-bulb’) at anterior right of visceral sac. Spacious haemocoel cavity into which head–foot can be retracted located between ‘heart-bulb’ and anterior mantle border (mantle ‘hood’ just anterior to position of diaphragm separating head–foot from visceral sac; Fig. 1). When disturbed, animals retract head–foot into this cavity and contract, visceral sac then curved, foot folded and tucked into concave side of visceral sac, head appendages project partially from underneath mantle ‘hood’.

Front end of foot semicircular, edges slightly flaring; posterior end with pointed tip; foot sole wider than dorsal head–foot. Head appendages of about equal length; each appendage showing rod-like spicules sorted longitudinally. Labial tentacles slightly flattened in cross-section, held parallel to ground in crawling specimens, medially forming upper lip. Rhinophores round in cross-section, held erect.

General histology: Musculature consisting of blue staining fibres either spanning body cavity independently, or associated closely with organs. Body wall musculature a mesh of outer circular and inner longitudinal fibres. All parts of digestive system surrounded by longitudinal muscle fibres; circular fibres apparent only around salivary ducts. Transversal muscular diaphragm (Fig. 1F: dp) is punctured by aorta, oesophagus and visceral nerve, and is located at base of visceral sac, separating body cavities of head–foot and visceral sac (see mantle ‘hood’ above).

Connective tissue fills most spaces in foot (dense aggregates of cells), and flanks of head–foot and anterior visceral sac (less dense aggregates). Aggregates separated from central body cavity by thin longitudinal sheath of connective tissue; aggregates consisting of rather large, irregularly shaped cells staining homogeneously light blue, filled with darker grains and few yellow-stained vesicles.

Calcareous spicules embedded in most of connective tissue. In serial sections of decalcified animals only spicule cavity remaining, apparently enclosing spicule in living animals; chamber usually containing remnants of dissolved spicules visible as smaller, translucent body consisting of concentric layers of undissolved matter. Spicules themselves cylindrical, straight or slightly bent with slightly thickened, rounded tips, giving a dumb-bell-like shape. Spicules glassy transparent but strongly refracting (Fig. 2H) under light microscope. Spicule surface smooth (Fig. 2J), interior slightly yellowish to brown in phase-contrast due to organic material (Fig. 2G). Concentric lamination evident in broken spicules viewed with SEM. Spicules size differing greatly: very small and short spicules around oral opening and oesophagus; long and thin ones arranged longitudinally inside cephalic appendages, forming continuous row from labial tentacles into upper lip. Highest number of spicules (80–120 µm long) embedded in dense connective tissue of foot. Largest spicules (up to 300 μm) sorted in at least two parallel strips dorsolaterally of central nervous system (CNS) and buccal mass, forming a grid of interdigitating pieces (‘cephalic spicule grid’; Fig. 4E).

Figure 2.

Microscopic views of radula (SEM), stylet of basal finger (SEM) and spicules surrounding the buccal mass (SEM, light microscopy) of Strubellia wawrai n. sp. A, F, F′. Vanuatu specimen; others: Solomon Islands. A. Functional part of radula. B. Complete hook-shaped radula. C. Right lateral teeth. D. Left lateral teeth. E. Rhachidian teeth, left view. F. Stylet of basal finger. F′. Detail of stylet tip. G. Spicule, phase contrast. H. Spicule, lateral illumination. J. Spicule, SEM. Abbreviations: dt, denticle; gr, groove; llp, left lateral plate; nt, notch; rlp 1 and 2, first and second right lateral plates; rt, rhachidian tooth; *, opening of hollow stylet. Scale bars: A, C–E = 20 µm; B = 100 µm; F = 150 µm; F′ = 3 µm; G, H, J = 50 µm. This figure appears in colour in the online version of Journal of Molluscan Studies.

Figure 2.

Microscopic views of radula (SEM), stylet of basal finger (SEM) and spicules surrounding the buccal mass (SEM, light microscopy) of Strubellia wawrai n. sp. A, F, F′. Vanuatu specimen; others: Solomon Islands. A. Functional part of radula. B. Complete hook-shaped radula. C. Right lateral teeth. D. Left lateral teeth. E. Rhachidian teeth, left view. F. Stylet of basal finger. F′. Detail of stylet tip. G. Spicule, phase contrast. H. Spicule, lateral illumination. J. Spicule, SEM. Abbreviations: dt, denticle; gr, groove; llp, left lateral plate; nt, notch; rlp 1 and 2, first and second right lateral plates; rt, rhachidian tooth; *, opening of hollow stylet. Scale bars: A, C–E = 20 µm; B = 100 µm; F = 150 µm; F′ = 3 µm; G, H, J = 50 µm. This figure appears in colour in the online version of Journal of Molluscan Studies.

Large anterior pedal gland located in anterior body cavity, ventrally to pharynx and CNS; distal part consists of paired lobes of thick glandular epithelium surrounding central lumen; cells filled with very small granules staining dark or light blue. Lobes of this gland merge anteriorly, connecting to short and wide epidermal duct leading into strongly ciliated, V-shaped longitudinal groove on dorsal side of anterior foot margin, ventrally to mouth opening. Further clusters of round foot glands located in entire foot ventrally to connective tissue, between dorsoventral muscle fibres; glands most numerous in anterior foot. Glandular cells containing many small dark blue grains, some yellow vesicles; cells open onto foot sole through very thin ducts.

Digestive system: Digestive system closely resembling that of other acochlidians: oral tube elongate, followed by bulbous pharynx containing hook-shaped radula, followed by paired salivary glands and oesophagus; direct connection into large digestive gland filling large part of visceral sac; intestine short with anal opening on right anterior side of visceral sac (Fig. 1E). No histologically detectable differentiated stomach. Ciliation of digestive tract detectable only in two places: at short strip in proximal part of oesophagus (where it projects from pharynx) and inside intestine.

Mouth opening a vertical slit located underneath upper lip; the following rather long oral tube surrounded by lateral clusters of oral glands opening into oral tube through thin ducts; oral gland cells staining dark blue (peripheral) or pale pink (closer to oral tube). Strong pair of pharynx protractors running from posterior end of oral tube to rhinophores; another pair running posteroventrally. Posterior end of oral tube is lined with thin cuticle. Pharynx egg-shaped, complex mass of muscle surrounding pharyngeal cavity; muscle surrounds posterior tip of radula (Fig. 4E, F). Pharynx protrusible anteriorly in slightly sucker-like fashion, surrounded by circular margin of epidermal tissue. Haemocoel lacunae present within pharynx, between fibres of pharyngeal muscles, supporting radula laterally and ventrally. Pharyngeal cavity lined with thin epithelium covered by equally thick, clear blue-staining cuticle (up to 15 µm thick); cavity with three longitudinal furrows, appearing as three-pointed star in cross-section (vertical furrow extending dorsally of radula). Radula originates in posterior tip of pharynx; ribbon originally still folded, embedded between large cells. Folded, upper branch runs anteriorly, emerging into pharyngeal cavity and spreading open. Radula then curves down, open part with old and worn teeth leading posteriorly again for about half length of upper branch (Fig. 2B). Radula asymmetric: single left lateral plate, prominent rhachidian tooth, two right lateral plates per row. Radular formula 40–60 × 1.1.2 (number of tooth rows in small Solomon Islands to large Vanuatu specimens, respectively). Rhachidian teeth with rectangular base and very slender, blade-like and pointed median cusp, its margins serrated (c. 30 or more small denticles per side) (Fig. 2A, E). Under light microscope, youngest rhachidian teeth appearing more translucent and with slimmer base than following teeth; median cusps of oldest rhachidian teeth generally worn down to stumps. First lateral plates of both sides flat and rectangular; each plate equipped with strong denticle on border to next younger plate, this border with notch into which denticle of other plate fits (Fig. 2A). Small and diamond-shaped second lateral plate on right side of radula; inner border straight, right first lateral plate appearing equally cut-off (Fig. 2A, C). Left lateral plates slightly wider than right ones (65 vs 50 µm in same row), outer border more rounded (Fig. 2A, D).

Salivary glands paired, connecting to posterior end of pharynx via thin salivary ducts. Each gland with two longitudinal lobes (resembling figure-of-eight in cross-section) formed by columnar cells densely filled with dark blue-stained granules. Central collecting duct strongly ciliated, showing bulbous salivary pump distal to glandular tissue (Fig. 1F: sp); spindle-shaped pumps and following salivary ducts surrounded by circular muscle fibres (contrasting with all other muscular linings of digestive system); salivary ducts opening anteriorly into lateral folds of pharyngeal cavity.

Oesophagus a simple tube projecting from posterodorsal side of pharynx; distal oesophagus widens gradually before connecting to lumen of digestive gland. Digestive gland a long sac usually filling most of visceral sac (in mature specimens gonad more voluminous). Outer surface of gland with irregular transverse folds; inner surface highly enlarged by glandular epithelium with high columnar cells forming bundles projecting into lumen. Epithelial cells filled with numerous small blue-stained vesicles; large, spherical, yellow-stained vacuoles in an apical position make up large part of glandular mass (Fig. 7A, C). Intestine rather short and thick, emerging from digestive gland dextrolaterally to distal oesophagus. Inner surface of intestine folded longitudinally, strongly ciliated. Intestine gradually thinning towards anal opening; opening hard to detect in most specimens but very close to renal pore, both openings sometimes forming an invaginated and ciliated common cavity (possibly an artifact due to fixation).

Central nervous system—cerebral nerve ring: CNS euthyneurous, slightly epiathroid (i.e. pleural ganglia closer to cerebral than to pedal ganglia), following general acochlidian bauplan (Fig. 3). Prepharyngeal nerve ring consisting of paired cerebral, pedal and pleural ganglia; three ganglia on visceral nerve cord plus osphradial ganglion; paired buccal ganglia posterior to pharynx. Further elements: paired optic and rhinophoral ganglia (on anteroventral sides of cerebral ganglia), paired gastro-oesophageal ganglia dorsally on each buccal ganglion. Serial sections reveal numerous nerves (Figs 3, 4).

Figure 3.

Schematic overview of the CNS (pedal nerves omitted for clarity) of Strubellia wawrai n. sp., dorsal view. Abbreviations: acn, anterior cerebral nerve; aon, aortic nerve; bg, buccal ganglion; cg, cerebral ganglion; esn, esophageal nerves; ey, eye; gog, gastroesophageal ganglion; hnc, Hancock's organ; hnn, Hancock's organ nerve; ln, labial tentacle nerve; mpn, median pedal nerve; on, optic nerve; opt, optical ganglion; orn, oral nerve; osg, osphradial ganglion; osn, osphradial nerve; osp, osphradium; pag, parietal ganglion; pg, pedal ganglion; plg, pleural ganglion; psn, penial sheath nerve; rhg, rhinophoral ganglion; rhl, rhinoporal looping nerve; rhn, rhinophoral nerve; rn, radular nerve; sc, statocyst; sdn, salivary duct nerve; subg, subintestinal ganglion; supg, supraintestinal ganglion; vcn, ventral cerebral nerve; vg, visceral ganglion; vn, visceral nerve. Not to scale.

Figure 3.

Schematic overview of the CNS (pedal nerves omitted for clarity) of Strubellia wawrai n. sp., dorsal view. Abbreviations: acn, anterior cerebral nerve; aon, aortic nerve; bg, buccal ganglion; cg, cerebral ganglion; esn, esophageal nerves; ey, eye; gog, gastroesophageal ganglion; hnc, Hancock's organ; hnn, Hancock's organ nerve; ln, labial tentacle nerve; mpn, median pedal nerve; on, optic nerve; opt, optical ganglion; orn, oral nerve; osg, osphradial ganglion; osn, osphradial nerve; osp, osphradium; pag, parietal ganglion; pg, pedal ganglion; plg, pleural ganglion; psn, penial sheath nerve; rhg, rhinophoral ganglion; rhl, rhinoporal looping nerve; rhn, rhinophoral nerve; rn, radular nerve; sc, statocyst; sdn, salivary duct nerve; subg, subintestinal ganglion; supg, supraintestinal ganglion; vcn, ventral cerebral nerve; vg, visceral ganglion; vn, visceral nerve. Not to scale.

Figure 4.

Three-dimensional reconstruction of general anatomy and CNS of Strubellia wawrai n. sp. from Vanuatu (A, C) and Solomon Islands (B, D–F). A. General anatomy, right view. B. Main ganglia, left view. C. CNS, anterior right view. D. Main ganglia, posterodorsal view. E. CNS with spicule grid and rudimentary penial sheath, dorsal view. F. CNS and buccal mass, anterior right view. Abbreviations: an, anus; bg, buccal ganglion; bm, buccal mass; bvd, posterior-leading vas deferens; cbc, cerebrobuccal connective; ccm, cerebral commissure; cg, cerebral ganglion; cgl, ‘cephalic gland’; cns, central nervous system; cop, copulatory apparatus; dg, digestive gland; es, esophagus; ey, eye; fgl, nidamental glands; ft, foot; go, gonad; gog, gastroesophageal ganglion; gp, genital pore; hnc, Hancock's organ; hnn, Hancock's nerve; ht, heart; kd, kidney; ln, labial tentacle nerve; mo, mouth opening; nd, nephroduct; on, optic nerve; opt, optical ganglion; orn, oral nerve; osg, osphradial ganglion; osn, osphradial nerve; osp, osphradium; ot, oral tube; pag, parietal ganglion; pc, pericardium; pcm, pedal commissure; pg, pedal ganglion; plg, pleural ganglion; pn, pedal nerve; ppc, parapedal commissure; pr, prostate; ps, penial sheath; psn, penial sheath nerve; r, radula; rhg, rhinophoral ganglion; rhn, rhinophoral nerve; rm, retractor muscle of penial sheath; rn, radular nerve; sc, statocyst; sg, sperm groove; spc, spicules; subg, subintestinal ganglion; supg, supraintestinal ganglion; vg, visceral ganglion; vn, visceral nerve; asterisks: branching points of nerves. Scale bars: A = 2 mm; B, D = 100 µm; C, F = 200 µm; E = 500 µm. The interactive 3D models of S. wawrai n. sp. can be accessed by clicking onto AD (general anatomy) and E, F (CNS) in the online PDF version of this article. Rotate model by dragging with left mouse button pressed; shift model: same action + ctrl (or dragging with left and right mouse buttons pressed); zoom: use mouse wheel. Select or deselect (or change transparency of) components in the model tree, switch between prefab views or change surface visualization (e.g. lighting, render mode, crop etc.). Interactive manipulation requires Adobe Reader 7 or higher.

Figure 4.

Three-dimensional reconstruction of general anatomy and CNS of Strubellia wawrai n. sp. from Vanuatu (A, C) and Solomon Islands (B, D–F). A. General anatomy, right view. B. Main ganglia, left view. C. CNS, anterior right view. D. Main ganglia, posterodorsal view. E. CNS with spicule grid and rudimentary penial sheath, dorsal view. F. CNS and buccal mass, anterior right view. Abbreviations: an, anus; bg, buccal ganglion; bm, buccal mass; bvd, posterior-leading vas deferens; cbc, cerebrobuccal connective; ccm, cerebral commissure; cg, cerebral ganglion; cgl, ‘cephalic gland’; cns, central nervous system; cop, copulatory apparatus; dg, digestive gland; es, esophagus; ey, eye; fgl, nidamental glands; ft, foot; go, gonad; gog, gastroesophageal ganglion; gp, genital pore; hnc, Hancock's organ; hnn, Hancock's nerve; ht, heart; kd, kidney; ln, labial tentacle nerve; mo, mouth opening; nd, nephroduct; on, optic nerve; opt, optical ganglion; orn, oral nerve; osg, osphradial ganglion; osn, osphradial nerve; osp, osphradium; ot, oral tube; pag, parietal ganglion; pc, pericardium; pcm, pedal commissure; pg, pedal ganglion; plg, pleural ganglion; pn, pedal nerve; ppc, parapedal commissure; pr, prostate; ps, penial sheath; psn, penial sheath nerve; r, radula; rhg, rhinophoral ganglion; rhn, rhinophoral nerve; rm, retractor muscle of penial sheath; rn, radular nerve; sc, statocyst; sg, sperm groove; spc, spicules; subg, subintestinal ganglion; supg, supraintestinal ganglion; vg, visceral ganglion; vn, visceral nerve; asterisks: branching points of nerves. Scale bars: A = 2 mm; B, D = 100 µm; C, F = 200 µm; E = 500 µm. The interactive 3D models of S. wawrai n. sp. can be accessed by clicking onto AD (general anatomy) and E, F (CNS) in the online PDF version of this article. Rotate model by dragging with left mouse button pressed; shift model: same action + ctrl (or dragging with left and right mouse buttons pressed); zoom: use mouse wheel. Select or deselect (or change transparency of) components in the model tree, switch between prefab views or change surface visualization (e.g. lighting, render mode, crop etc.). Interactive manipulation requires Adobe Reader 7 or higher.

Cerebral ganglia largest ganglia, largely spherical; cerebral commissure strong (Figs 4D, 5G). Cerebropleural connective slightly shorter than cerebropedal one; static nerve very thin, emerging close to base of cerebropleural connective and running parallel to it to paired statocysts. Statocysts embedded in top of each pedal ganglion. Cerebrobuccal connectives thin, very long, running posteriorly within pharyngeal musculature laterally to dorsal branch of radula (Fig. 4F).

Figure 5.

Semithin sections of the CNS and sensory organs (Solomon Islands specimens) of Strubellia wawrai n. sp. A–C. Cerebral ganglion and double cerebro-optic connectives. D. Hancock's organ. E. Osphradium. F. Pedal ganglion and statocyst. G. Cephalic gland dorsally to cerebral ganglia. Abbreviations: acn, anterior cerebral nerve; ao, aorta; ccm, cerebral commissure; cg, cerebral ganglion; cgl, cephalic gland; ep, epidermis; ey, eye; ln, labial tentacle nerve; on, optic nerve; opt, optic ganglion; osn, osphradial nerve; ot, otal tube; pcm, pedal commissure; pg, pedal ganglion; pnd, dorsal pedal nerve; pnl, lateral pedal nerve; rhl, rhinophoral looping nerve; sc, statocyst; spc, spicules; 1, multiciliated cells; 2, microvillous border; 3, vacuolate cells. All scale bars = 50 µm. This figure appears in colour in the online version of Journal of Molluscan Studies.

Figure 5.

Semithin sections of the CNS and sensory organs (Solomon Islands specimens) of Strubellia wawrai n. sp. A–C. Cerebral ganglion and double cerebro-optic connectives. D. Hancock's organ. E. Osphradium. F. Pedal ganglion and statocyst. G. Cephalic gland dorsally to cerebral ganglia. Abbreviations: acn, anterior cerebral nerve; ao, aorta; ccm, cerebral commissure; cg, cerebral ganglion; cgl, cephalic gland; ep, epidermis; ey, eye; ln, labial tentacle nerve; on, optic nerve; opt, optic ganglion; osn, osphradial nerve; ot, otal tube; pcm, pedal commissure; pg, pedal ganglion; pnd, dorsal pedal nerve; pnl, lateral pedal nerve; rhl, rhinophoral looping nerve; sc, statocyst; spc, spicules; 1, multiciliated cells; 2, microvillous border; 3, vacuolate cells. All scale bars = 50 µm. This figure appears in colour in the online version of Journal of Molluscan Studies.

Labiotentacular nerves very thick (diameter c. 50 µm), emerging medioventrally from each cerebral ganglion; nerve splits early into thinner oral branch (running to upper lip) and thick part (to tip of labial tentacles, with thinner branches repeatedly running to anterior side of tentacles; Fig. 4C, F). Right labial nerve of some specimens with further branch extending posterodorsally, innervating penial sheath (Fig. 4C: psn).

Rhinophoral ganglion located at anteroventral part of cerebral ganglion between labiotentacular nerve and optic ganglion (Fig. 4B). Rhinophoral ganglion elongate and pear-shaped; thicker portion containing few peripheral cell bodies and connecting to cerebral ganglion by short connective, thinner part running smoothly into rhinophoral nerve. Rhinophoral nerve splitting into three branches close to its origin: thickest part continues into rhinophores (without much further branching); second, thinner part innervates Hancock's organs posterior to rhinophoral bases; third (thinnest) branch looping backwards and apparently connecting to anteroventral side of optic ganglion (Fig. 3: rhl).

Optic ganglion hemispherical, attached to cerebral ganglion laterally but separated by independent layer of connective tissue (Fig. 5B). Double, very short cerebro-optic connectives, posterior one stronger (Fig. 5A, C); third connective detected in single specimen. Optic nerve thin, rather long, joining to posteroventral portion of eye; thin and looping second nerve connecting to Hancock's organ's branch of rhinophoral nerve (see above).

Two further cerebral nerves detectable: (1) thin nerve leaving cerebral ganglion medially (Figs 3, 5A: acn), running anteroventrally along paired cephalic blood vessels before splitting into branches running towards rhinophores and to the mouth opening; (2) thin nerve emerging from posteroventral side of cerebral ganglion (Fig. 3: vcn), running into muscular lining of cephalic blood vessels.

Mass of loosely aggregated and apparently glandular cells in body cavity above cerebral ganglia and cerebral commissure (‘cephalic gland’); containing numerous vacuoles staining light yellow. Gland mass without detectable connection to ganglia except for some thin fibers (connective tissue?); symmetric lobes extending slightly down sides of cerebral ganglia (Figs 4F, 5G).

Pedal ganglia spherical, only slightly smaller than cerebral ganglia; joined by thick pedal commissure (Fig. 5F) and thinner, longer parapedal commissure; very thin nerve splitting off parapedal commissure just left of midline (Fig. 4D), running to median part of foot sole and anterior pedal gland.

Six further pairs of pedal nerves detected, all running to body flanks: anteroventral, ventrolateral, posteroventral and posterodorsal nerves rather thick and running along body sides in posterior direction (except for first one); additional thin antero- and posterodorsal nerves running to sides, the former one apparently joining to anteroventral pedal nerve close to eye.

Central nervous system—visceral loop and buccal ganglia: Visceral cord with three medium-sized to large ganglia, connecting beneath anterior part of pharynx (Fig. 4B, D; nomenclature after Haszprunar, 1985a; Sommerfeldt & Schrödl, 2005): (1) left parietal ganglion (small, thin nerve running to left body side); (2) fused subintestinal/visceral ganglion (large, left of midline; giant nerve cells and very thick visceral nerve running posteriorly); (3) fused supraintestinal/right parietal ganglion (medium sized, thin nerve running to right body side). Latter ganglion with osphradial ganglion (small, cap-shaped) on posterodorsal side (Fig. 5D), both ganglia enclosed by common sheath of connective tissue. Osphradial ganglion with two nerves, one looping upwards first before running posteriorly; second: osphradial nerve innervating osphradium on anterior right body side (Fig. 4F). Ganglia on visceral nerve cord joined by short to very short connectives, only ganglia (2) and (3) with long connective passing obliquely between pharynx and aorta; thin nerve emerging from left third of long connective running downward into musculature of aorta (Fig. 4D: asterisk).

Visceral nerve strongest nerve posterior to CNS (diameter c. 25 µm) and running posteriorly into visceral sac, slightly left of midline (Fig. 4C: vn); nerve identifiable by surrounding longitudinal muscle fibres throughout entire length; nerve passes through diaphragm close to aorta and oesophagus.

Buccal ganglia paired, medium-sized, situated on posterodorsal side of pharynx at emerging point of oesophagus. Buccal commissure short, running ventrally to oesophagus; thin, apparent radular nerve emerging from middle of commissure, leading forward into muscular mass of pharynx (Fig. 4C, F).

Gastro-oesophageal ganglia (small, bean-shaped) on top of each buccal ganglion, connected by short vertical connective; thin oesophageal nerve from upper part of connective leading medially into muscular sheath of oesophagus; another thin nerve running from base of each gastro-oesophageal ganglion into sheath surrounding salivary ducts (Fig. 3: esn, sdn).

Sensory organs: Eyes located dorsolaterally to slightly anteriorly to cerebral ganglia, underneath translucent patch of epidermis visible in living animals (Fig. 1A, B); eyes bean-shaped, c. 130 µm long, facing anterolaterally (Fig. 4C, F), surrounded by thin layer of connective tissue; innervation by thin optic nerves. Prismatic (sensory?) cells with distinct nuclei form cup-shaped outer layer of eye, followed by layer of grainy black pigment; grey-blue staining irregular band (possibly sensory microvilli) between pigment layer and otherwise acellular and light blue-staining lens (Fig. 5A). Lens covered distally by cornea consisting of single layer of flat cells.

Statocysts paired, hollow spheres (diameter 25 µm) with flat, slightly ciliated cells forming outer wall (Fig. 5F); remnants of layered single statolith inside fluid-filled cavity visible in some sections. Statocysts embedded in dorsal part of each pedal ganglion (Fig. 4B); static nerve originating in cerebral ganglia.

Hancock's organs posterior to base of each rhinophore, located inside zone of brighter epidermis over eyes; exact dimensions of organs detectable only in serial sections, there appearing as shallow patches of thin epidermis, resembling osphradium in histology (dense microvillous border, several multiciliated cells), differing in presence of rounded, apparently glandular cells with clear lumen (Fig. 5D); innervation by lateral branches of rhinophoral nerves.

Osphradium a small pit on right body side, visible in living animals as keyhole-shaped spot paler than surrounding epidermis (Fig. 1A); in serial sections a pit about 40 µm deep and 60 µm long, lined with very thin epidermis showing strong microvillous border (Figs 4F, 5E); several cells with bundles of cilia c. 25 µm long found inside pit but mainly close to rim; osphradial nerve emerging from osphradial ganglion, splitting up distally.

Multiciliated cells similar to putative sensory cells in Hancock's organs and osphradium found interspersed within normal epidermal cells on labial tentacles and rhinophores.

Circulatory and excretory systems: Pericardial complex located in anterior right of visceral sac, with externally visible ‘heart bulb’ indicated by beating of heart in living animals (Fig. 1B). Pericardial complex formed by spacious pericardium enveloping two-chambered heart; elongate kidney and looping nephroduct extending posteriorly along right side of visceral sac (Figs 4A, 6). Renal pore situated on anteroventral right, close to anal opening. Aorta extending into head–foot, passing between pharynx and pedal commissure, distally dividing into paired vessels (Figs 5F, 7); vessels terminating laterally of oral tube. In large Vanuatu specimens, second branch of aorta detectable, running posteriorly into visceral sac.

Figure 6.

Schematic overview of the circulatory and excretory systems of Strubellia wawrai n. sp., right view. Abbreviations: ao, aorta; au, auricle; cv, paired cephalic vessels; dkd, distal kidney lumen; nd, nephroduct; ndl, nephroduct loop; np, nephropore; pc, pericardium; pkd, proximal kidney lumen; rpd, renopericardioduct; vac, vacuolated epicardium on ventricular wall; ve, venous opening; vt, ventricle; *, ciliated intersection between kidney and nephroduct. Not to scale.

Figure 6.

Schematic overview of the circulatory and excretory systems of Strubellia wawrai n. sp., right view. Abbreviations: ao, aorta; au, auricle; cv, paired cephalic vessels; dkd, distal kidney lumen; nd, nephroduct; ndl, nephroduct loop; np, nephropore; pc, pericardium; pkd, proximal kidney lumen; rpd, renopericardioduct; vac, vacuolated epicardium on ventricular wall; ve, venous opening; vt, ventricle; *, ciliated intersection between kidney and nephroduct. Not to scale.

Figure 7.

Semithin sections of the circulatory and excretory systems of Strubellia wawrai n. sp. (Solomon Islands specimens). A. Heart, longitudinal section. B. Pericardium and heart, cross-section. C. Anterior portion of excretory system, longitudinal section. D. Anterior portion of excretory system, cross-section. E. Wall of ventricle, cross-section. F. Proximal and distal kidney lumina, cross-section. G. Nephroduct, suspended by muscle fiber, cross-section. Abbreviations: ao, aorta; dg, digestive gland; dkd, distal kidney lumen; ht, heart; it, intestine; lc, hemocoel lacunae dorsally to pericardium; nd, nephroduct; pc, lumen of pericardium; pkd, proximal kidney lumen; rpd, renopericardioduct; black arrowheads: wall of pericardium; white arrowhead: peritoneal membrane; *, venous opening of heart to hemocoel lacunae; **, loose cells inside heart; ***, ciliated intersection between kidney and nephroduct; 1, vacuolate epicardium; 2, muscular wall of ventricle; 3, cells containing spicule-like body; 4, muscle fibers spanning ventricle. Scale bars: A–C = 100 µm; D, E = 50 µm; F, G = 25 µm. This figure appears in colour in the online version of Journal of Molluscan Studies.

Figure 7.

Semithin sections of the circulatory and excretory systems of Strubellia wawrai n. sp. (Solomon Islands specimens). A. Heart, longitudinal section. B. Pericardium and heart, cross-section. C. Anterior portion of excretory system, longitudinal section. D. Anterior portion of excretory system, cross-section. E. Wall of ventricle, cross-section. F. Proximal and distal kidney lumina, cross-section. G. Nephroduct, suspended by muscle fiber, cross-section. Abbreviations: ao, aorta; dg, digestive gland; dkd, distal kidney lumen; ht, heart; it, intestine; lc, hemocoel lacunae dorsally to pericardium; nd, nephroduct; pc, lumen of pericardium; pkd, proximal kidney lumen; rpd, renopericardioduct; black arrowheads: wall of pericardium; white arrowhead: peritoneal membrane; *, venous opening of heart to hemocoel lacunae; **, loose cells inside heart; ***, ciliated intersection between kidney and nephroduct; 1, vacuolate epicardium; 2, muscular wall of ventricle; 3, cells containing spicule-like body; 4, muscle fibers spanning ventricle. Scale bars: A–C = 100 µm; D, E = 50 µm; F, G = 25 µm. This figure appears in colour in the online version of Journal of Molluscan Studies.

Pericardium formed by very thin wall breached in three places: (1) dorsally at venous connection between haemocoel and atrial lumen; (2) anteroventrally, where aorta extends from ventricle into body; (3) posterolaterally to heart where ciliated renopericardioduct drains off into kidney (Fig. 6). Pericardial lumen free of cells, except for few vacuolated cells at anteroventral wall which appears to wrap around distal part of nephroduct.

Heart consisting of thin-walled auricle and muscular, ovoid ventricle. Haemocoel on right side of visceral sac connected to auricle by small hole (diameter 10 µm); opening visible only in single series where auricle clearly distinguishable from ventricle (Fig. 7A); auricle collapsed in most other cases. Ventricle continuous with auricle in its wall, ovoid form appearing more constant; ventricular wall much thicker, formed by mesh of striated muscle fibres staining blue-grey, some fibres appearing to cross ventricular lumen, forming muscular bridges (Fig. 7B).

Inside of ventricular wall covered with irregular cells, some staining darker blue or with yellow-stained vacuole; conspicuous large cells embedded in former layer and interspersed freely in the ventricular lumen: cells elongate and ovoid, showing a central body stained light grey, with concentric layers somewhat resembling a spicule.

Outer wall of ventricle covered with irregularly bordered, conspicuous lining at least as thick as muscular layer of wall. Epicardial lining consisting of vacuolate cells staining light blue to grey, with flat nuclei sorted apically staining slightly darker (Fig. 7E).

Tip of ventricle continuing into thick aorta, wall consisting of longitudinal muscle fibres, internal surface smooth. Aorta leaving pericardium on medioventral side, running anteriorly and passing through diaphragm close to oesophagus and visceral nerve, splitting into paired vessels formed only by strips of muscle fibres and membranous wall ventrally to cerebral nerve ring; cephalic vessels spacious, running parallel to oral tube (Figs 5F, 6), terminating close to mouth.

Excretory system consisting of short but well-developed renopericardioduct, elongate kidney and long and looping nephroduct. Renopericardioduct longitudinally folded, connecting to pericardium via funnel-shaped opening containing conspicuous ciliary flame; cuboidal lining with bundles of strong cilia projecting into pericardium and renopericardial duct (Fig. 7C, E), leading into kidney.

Kidney elongate, extending along two thirds of visceral sac; longitudinal interior wall separating lumen into hairpin-like loop connected only at kidney's posterior end (Figs 6, 9A, B): proximal part of lumen (running front to back) lying more ventrally, lined with regular epithelium with dense microvillous border (Fig. 7C, F); distal part of kidney lumen (running back to front) lying dorsally, more voluminous and lined with epithelium with shorter microvillous border, conspicuous unstained vacuoles giving wall spongy appearance (Fig. 7D) and accounting for much of kidney's volume. Connection to nephroduct through constriction of only about 3 µm diameter (in direct proximity to the renopericardioduct funnel), followed by short patch of dense ciliation (Fig. 7C: triple asterisk). Undulating nephroduct running posterior to tip of kidney and looping forward again; nephroduct interconnected by single muscle fibres in at least one place; lined with smooth epithelium staining light blue, with interspersed yellow-stained vesicles and a slight microvillous border (Fig. 7G). Distal loop of nephroduct differing slightly in histology (epithelium staining darker, showing fewer yellow vesicles but possibly colorless, irregular vacuoles), arching upward before running downward again towards nephropore (Fig. 9A, B); appearing to be closely associated with fold of pericardium.

Nephropore formed by ciliated and invaginated part of epidermis, situated next to anal opening or inside invaginated cloaca (artifact?), on dextroventral anterior visceral sac.

Genital system: Presence of allosperm receptacles in males, and females with rudimentary ‘male’ features indicate protandric hermaphroditism (as in S. paradoxa from Ambon). Examined specimens from Solomon Islands only juveniles and two functional ‘females’ (one with vestigial bursa copulatrix and penial sheath; Figs 4E, 9F); Vanuatu specimens containing one juvenile and one female (gonad filled with oocytes, nidamental glands developed) but with apparently functional cephalic copulatory apparatus and two allosperm receptacles (Figs 4A, 9C, D, E).

Posterior genital system consisting of acinar gonad, proximal receptaculum seminis filled with sorted spermatozoa and glandular gonoduct leading to genital opening on anterior right of visceral sac. Ampulla thin-walled, wide; detected only in single specimen. Gonad consisting of numerous almost spherical acini, filling much of visceral sac in functionally female specimens. Each acinus formed by thin epithelial wall, filled with large spherical oocytes containing high numbers of vesicles staining brilliantly blue, with colorless vesicles filling gaps in between; acini connected to gonoduct by thin ducts (Fig. 8A). Collecting gonoduct surrounded by muscle fibres but collapsed in both specimens; strong ciliation apparent; following last acinus a very short piece of gonoduct from which receptaculum seminis (thick-walled and blind-ending sac) emerges laterally. Receptacle lined with simple blue-staining epithelium forming an undulated inner wall; numerous spermatozoa are embedded with their heads into wall. Heads of spermatozoa visible only at high magnifications as stronger refracting bodies; head short, not screw-shaped, diameter about 1 µm; flagella forming pink-stained, dense and streaked mass inside receptacle (Fig. 10B: arrowheads and asterisk).

Figure 8.

Schematic overview of the genital system and copulatory apparatus of Strubellia wawrai n. sp. A. Genital system, dark grey areas indicate organs that become reduced in the female phase. B. Copulatory apparatus. Abbreviations: alg, albumen gland; am, ampulla; bc, bursa copulatrix; bf, basal finger; bvd, posterior-leading vas deferens; dv, diverticle; ed, ejaculatory duct; go, gonad; gp, genital pore; meg, membrane gland; mug, mucus gland; p, penis; ppd, paraprostatic duct; ppr, paraprostate; pr, prostate; ps, penial sheath; rm, retractor muscle; rs, receptaculum seminis; sg, sperm groove; st, stylet of basal finger; th, spine. Not to scale.

Figure 8.

Schematic overview of the genital system and copulatory apparatus of Strubellia wawrai n. sp. A. Genital system, dark grey areas indicate organs that become reduced in the female phase. B. Copulatory apparatus. Abbreviations: alg, albumen gland; am, ampulla; bc, bursa copulatrix; bf, basal finger; bvd, posterior-leading vas deferens; dv, diverticle; ed, ejaculatory duct; go, gonad; gp, genital pore; meg, membrane gland; mug, mucus gland; p, penis; ppd, paraprostatic duct; ppr, paraprostate; pr, prostate; ps, penial sheath; rm, retractor muscle; rs, receptaculum seminis; sg, sperm groove; st, stylet of basal finger; th, spine. Not to scale.

Following receptaculum seminis another short piece of gonoduct, leading into female gland mass. Glandular mass tubular throughout, forming several stout loops in anterior visceral sac; strongly stained, columnar glandular cells surround lumen only from one side (Fig. 10A); lumen a longitudinal fold projecting in between glandular cells. Glandular cells up to almost 100 µm high, filled with granular secretions. Three differently staining zones along glandular gonoduct: (1) proximal part staining dark blue; (2) distal part blue with strong pinkish tone; (3) part in between appearing blue with slightly greenish hue (Fig. 9D, F). Distal part of glandular epithelium becomes thinner with diameter of strongly ciliated gonoduct lumen appearing to increase before opening to outside through genital pore.

Figure 9.

Three-dimensional reconstructions of the excretory, circulatory and reproductive systems of Strubellia wawrai n. sp. from Solomon Islands (A, B, F) and Vanuatu (C–E). A. Excretory system, left view. B. Excretory and circulatory systems, right view. C. Anterior male copulatory organs and (para-)prostatic glandular systems, left view. D. Nidamental glands and bursa copulatrix, right view. E. Penis and basal finger, left view. F. Nidamental glands and rudimental bursa copulatrix, ventral view. Abbreviations: am, ampulla; alg, albumen gland; ao, aorta; bc, bursa copulatrix; bf, basal finger; bvd, posterior-leading vas deferens; dkd, distal kidney lumen; dv, diverticle; ed, ejaculatory duct; go, gonad; gp, genital pore; ht, heart; meg, membrane gland; mug, mucus gland; nd, nephroduct, ndl, nephroduct loop; np, nephropore; p, penis; pc, pericardium; pkd, proximal kidney lumen; pr, prostate; ppd, paraprostatic duct; ppr, paraprostate; rm, retractor muscle; rpd, renopericardioduct; rs, receptaculum seminis; sg, sperm groove; st, stylet; th, spine; *, connection between proximal and distal kidney lumina; **, connection between distal kidney lumen and nephroduct; ***, position of ejaculatory duct opening. Scale bars: A, B = 500 µm; C = 600 µm; D, F = 400 µm; E = 200 µm.

Figure 9.

Three-dimensional reconstructions of the excretory, circulatory and reproductive systems of Strubellia wawrai n. sp. from Solomon Islands (A, B, F) and Vanuatu (C–E). A. Excretory system, left view. B. Excretory and circulatory systems, right view. C. Anterior male copulatory organs and (para-)prostatic glandular systems, left view. D. Nidamental glands and bursa copulatrix, right view. E. Penis and basal finger, left view. F. Nidamental glands and rudimental bursa copulatrix, ventral view. Abbreviations: am, ampulla; alg, albumen gland; ao, aorta; bc, bursa copulatrix; bf, basal finger; bvd, posterior-leading vas deferens; dkd, distal kidney lumen; dv, diverticle; ed, ejaculatory duct; go, gonad; gp, genital pore; ht, heart; meg, membrane gland; mug, mucus gland; nd, nephroduct, ndl, nephroduct loop; np, nephropore; p, penis; pc, pericardium; pkd, proximal kidney lumen; pr, prostate; ppd, paraprostatic duct; ppr, paraprostate; rm, retractor muscle; rpd, renopericardioduct; rs, receptaculum seminis; sg, sperm groove; st, stylet; th, spine; *, connection between proximal and distal kidney lumina; **, connection between distal kidney lumen and nephroduct; ***, position of ejaculatory duct opening. Scale bars: A, B = 500 µm; C = 600 µm; D, F = 400 µm; E = 200 µm.

Single female Solomon Island specimen with vestigial bursa copulatrix consisting of very thin duct (10 µm diameter; emerging from gonoduct close to genital opening) and almost spherical terminal bulb close to upper intestine (Fig. 9F); bulb stained very dark blue inside. Same individual with distal gonoduct containing several oval bodies with pink-stained and grainy vesicle and fully developed ciliated sperm groove running from genital opening to base of right rhinophore. Thin tube entering body and running posteriorly from anterior end of sperm groove: posterior-leading vas deferens passing cerebral commissure dorsally and terminating in elongate blind sac (an empty and reduced penial sheath); reduced, thread-like penial retractor muscle extending posteriorly from sac, ending freely in body cavity (Fig. 4E).

Cephalic male copulatory organs: One Vanuatu specimen with elaborate male and female features: external sperm groove between female genital opening and base of right rhinophore, connecting to fully developed male copulatory organs surrounded by penial sheath at left of pharynx. Copulatory organs consisting of muscular basal finger, considerably smaller penis and their associated paraprostatic and prostatic glandular systems, respectively (Figs 4A, 8B).

Posterior-leading vas deferens connected to voluminous, tubular prostate gland; prostate continuing into long and curled ejaculatory duct, entering muscular penis at its base; ejaculatory duct opening to exterior through penial papilla at tip of penis. Solid spine of c. 150 µm width situated next to penial papilla (Fig. 9E). Blind ending glandular paraprostate a longer and thinner tube than prostate, strongly coiled (Fig. 9C: ppr). Paraprostatic duct emerging from paraprostate and connecting to muscular basal finger, entering basal finger approximately in middle of curved muscle; duct opening apically via curved hollow stylet of about 750 µm length. Stylet with cuticular groove running along its side (Figs 2F, 10D–H). Penis and basal finger muscles interconnected at their base; both structures surrounded by thin-walled penial sheath meeting posterior-leading vas deferens before opening to exterior at base of right rhinophore.

Figure 10.

Semithin sections of the genital system of Strubellia wawrai n. sp. from Solomon Islands (A, B; posterior genital system in female phase) and Vanuatu (C–H; parts of copulatory apparatus). A. Membrane gland showing acentral lumen. B. Receptaculum seminis filled with spermatozoa, heads along the wall. C. (Para-)prostatic glandular system. D. Hollow stylet of basal finger (tip on the left, close to the base on the right). E. Basal finger at base of stylet. F, G. Penis with ejaculatory duct and thorn embedded in epithelium. H. Trumpet-shaped penial papilla and tip of thorn. Abbreviations: bf, basal finger; ed, ejaculatory duct; es, esophagus; gr, groove of basal finger stylet; lu, lumen of basal finger stylet; p, penis; ppd, paraprostatic duct; ppr, paraprostate; pr, prostate; ps, lumen of penial sheath; sgl, salivary gland; st, stylet of basal finger; th, spine of penis; arrowheads: sperm heads; asterisk: mass of flagella. Scale bars: A, B = 50 µm; C–H = 100 µm. This figure appears in colour in the online version of Journal of Molluscan Studies.

Figure 10.

Semithin sections of the genital system of Strubellia wawrai n. sp. from Solomon Islands (A, B; posterior genital system in female phase) and Vanuatu (C–H; parts of copulatory apparatus). A. Membrane gland showing acentral lumen. B. Receptaculum seminis filled with spermatozoa, heads along the wall. C. (Para-)prostatic glandular system. D. Hollow stylet of basal finger (tip on the left, close to the base on the right). E. Basal finger at base of stylet. F, G. Penis with ejaculatory duct and thorn embedded in epithelium. H. Trumpet-shaped penial papilla and tip of thorn. Abbreviations: bf, basal finger; ed, ejaculatory duct; es, esophagus; gr, groove of basal finger stylet; lu, lumen of basal finger stylet; p, penis; ppd, paraprostatic duct; ppr, paraprostate; pr, prostate; ps, lumen of penial sheath; sgl, salivary gland; st, stylet of basal finger; th, spine of penis; arrowheads: sperm heads; asterisk: mass of flagella. Scale bars: A, B = 50 µm; C–H = 100 µm. This figure appears in colour in the online version of Journal of Molluscan Studies.

Behaviour and feeding: Living specimens collected by hand under rocks in shallow water at sides of streams. Aggregations of up to 25 individuals found under single calcareous rocks, hidden in grooves and pits of undersurfaces. Exposure to light causes animals to move; specimens kept in a Petri dish moved around without pause until hiding place was presented. On smooth surfaces, movement was fast, about 7 mm/s, with head moving from left to right, labial tentacles held parallel to ground. Movement appeared to be caused by ciliary motion (visible in animals crawling upside down at water surface: fine particles on water surface were quickly drawn away from front margin of foot) and supported by clear mucus as observable in specimens suspended by thread of mucus from water surface.

Three small individuals (probably juveniles) were cultivated in a small aquarium for about 5 months. When supplied with calcareous egg capsules of freshwater neritids Strubellia individuals were observed to aggregate on the egg capsules after a few minutes. Other types of food (fish feed, algae tabs, gelatinous egg masses of Physa sp.) did not lead to any reaction. Individuals remained on egg capsule with anterior border of foot and mouth pressed onto capsule's surface, head appearing slightly contracted (head appendages bent backwards, eyes not visible; Fig. 1C). Slow peristaltic dilatations of entire visceral sac observed during this apparent feeding posture, accompanied by slow but strong pumping motions of heart. Each feeding period up to 15 min; between two and three egg capsules fed on per individual. Some egg capsules fed on by more than one individual, others were ignored. Continuous supply of neritid eggs over longer period of time proved difficult; specimens shrank during time in aquarium.

Molecular phylogeny: The RAxMC-tree based on 16S rRNA and COI genes recovers the monophyletic genus Strubellia (bootstrap support, BS = 100) as sister taxon to the genera Acochlidium and Palliohedyle (Fig. 11), all three genera forming the large-bodied and limnic family Acochlidiidae (sensuArnaud et al., 1986). Sampling of 13 Strubellia individuals reveals three clades: a basal and yet undescribed branch from Sulawesi (known only from single individual) as sister taxon to a clade formed of S. paradoxa from Ambon (BS = 100) and a clade consisting of all sampled individuals from Solomon Islands and Vanuatu (BS = 96). Specimens from Vanuatu are nested within populations from the Solomons.

Figure 11.

RAxML tree of the genus Strubellia, based on a concatenated dataset of mitochondrial COI and 16S rRNA (1113 bp) and colour coded distribution map of the different Strubellia species. Bootstrap values given above nodes.

Figure 11.

RAxML tree of the genus Strubellia, based on a concatenated dataset of mitochondrial COI and 16S rRNA (1113 bp) and colour coded distribution map of the different Strubellia species. Bootstrap values given above nodes.

Statistical parsimony analyses generate two independent haplotype networks (not shown) for partial 16S rRNA: S. paradoxa and a network uniting S. wawrai n. sp. populations from Solomons and Vanuatu (no 16S rRNA sequence was available for Strubellia from Sulawesi). Four independent networks were generated based on partial COI (reduced to 571 bp, to analyse sequences of same length): S. paradoxa, Strubellia sp. (Sulawesi), S. wawrai n. sp. from Solomons, and from Vanuatu.

Intraspecific variation is generally very low: in 16S rRNA (438 bp) 0.0% in S. paradoxa (n = 2), 0.68–0.91% in S. wawrai n. sp. from Solomons (n = 7) and 0.45–0.68% in S. wawrai n. sp. from Vanuatu (n = 4). Uniting both populations of S. wawrai n. sp. (n = 11), intraspecific variation ranges from 0.45 to 1.14% in 16S rRNA. Lowest interspecific variation in 16S rRNA between S. paradoxa and S. wawrai n. sp. is 4.1%; a higher selected threshold clusters both species together. In COI (571 bp) intraspecific variation ranges between 1.57 and 1.92% for S. wawrai n. sp. from the Solomons (n = 7) and 0.7% for S. wawrai n. sp. from Vanuatu (n = 2); uniting both populations (n = 9) the variation ranges between 2.1 and 2.8%. Interspecific variation is comparably high ranging between 12.25 and 13.31% in S. paradoxa and S. wawrai n. sp. and between 14.36 and 15.06% in Strubellia sp. from Sulawesi and S. wawrai n. sp.

DISCUSSION

Comparative morphology of the cerebral nerve ring

The CNS of Strubellia wawrai n. sp. has been described briefly from dissected material by Wawra (1974, as S. paradoxa). The general organization of ganglia resembles that of S. paradoxa and other hedylopsacean acochlidian species, e.g. Pseudunela espiritusanta (Neusser & Schrödl, 2009; Brenzinger et al., 2011). Examination of serially sectioned specimens revealed several additional features, such as the previously undetected parapedal commissure and several thin cerebral nerves that complement the set of nerves beyond what is generally detectable in small mesopsammic acochlidians. Among the usually present nerves are three comparatively large anterior cerebral nerves also shown in Wawra's drawing (1974: fig. 7); we regard the nerves numbered 1.1–1.3 therein to be the labial tentacle nerve, the Hancock's and the rhinophoral nerve, respectively. Strubellia shows two small ganglia attached to the cerebral ganglia, as do all other hedylopsaceans: The “procerebral lobe” described by Wawra (but not depicted) can be suspected at the base of the rhinophoral and Hancock's nerve and likely refers to the rhinophoral ganglion herein. The optic ganglion appears to have been overlooked by Wawra; his “optic” nerve is shown to arise directly from the cerebral ganglion and thus might alternatively be the oral nerve which extends close to the labial tentacle nerve.

The homology of opisthobranch rhinophoral or optic ganglia and the pulmonate procerebrum (with double connectives to the cerebral ganglion) has been suggested previously (e.g. Haszprunar, 1988; Haszprunar & Huber, 1990; Huber, 1993) and a general homology of the sensory innervation among Euthyneura appears more and more likely (Jörger et al., 2010a, b). Comparison of these ganglia among Acochlidia might however hint at a common anlage of both the optic and rhinophoral ganglion: the presence of a looping nerve interconnecting both (present in S. wawrai n. sp. and Tantulum elegans; Neusser & Schrödl, 2007), the variable origin of the optic nerve (usually from the optic ganglion, in P. cornuta it splits off from the rhinophoral nerve; Neusser, Heß & Schrödl, 2009a) and finally the presence of double connectives in one ganglion or the other. A double cerebro-rhinophoral connective is present in Tantulum, the microhedylacean Pontohedyle milaschewitchii (Kowalevsky, 1901) and Microhedyle glandulifera (Kowalevsky, 1901) (Jörger et al., 2008; Neusser & Schrödl, 2009; own unpublished data); S. wawrai n. sp. is the only known species with a double cerebro-optic connective.

Differences from the CNS of S. paradoxa involve the apparent lack of the small cerebral nerves, the Hancock's nerve and Hancock's organs, but are likely to be artefacts (see Brenzinger et al., 2011). The only evident difference between the CNS of S. wawrai n. sp. from the Solomon Islands and from Vanuatu is the ‘penial’ nerve in the examined specimen from Vanuatu, which might be present only in mature male specimens and could therefore not be detected in the female specimens from the Solomon Islands.

Visceral loop, osphradial ganglion and arrangement of buccal ganglia

Wawra (1974) described the typical acochlidian visceral nerve cord with three separate ganglia; we identify the ganglia herein as a left parietal, a fused subintestinal/visceral and a fused right parietal/supraintestinal ganglion, respectively. Nerves splitting from the connective joining the latter two ganglia and from the parietal ganglia have not been reported for any other acochlidian so far.

The additional ganglion attached to the fused right parietal/supraintestinal ganglion was mentioned by Wawra (1974); it is known for all hedylopsacean species examined in detail and also for the microhedylacean Parhedyle cryptophthalma (Westheide & Wawra, 1974; Jörger et al., 2010b; Schrödl & Neusser, 2010). Judging from its position on the right side of the visceral loop, the ganglion was hypothesized to be homologous with the osphradial ganglion of other euthyneurans (Wawra, 1989; Huber, 1993; Sommerfeldt & Schrödl, 2005); this interpretation can be confirmed with the detection of an osphradium in S. wawrai n. sp. The presence of two nerves in S. wawrai n. sp. and a bifurcating nerve in Pseudunela espiritusanta suggests more than one function of the osphradial ganglion. In Tantulum elegans, the single nerve leaving the osphradial ganglion was mentioned to terminate close to the copulatory apparatus and hence assumed to be a “genital” or “penial” nerve (Neusser & Schrödl, 2007); innervation of the copulatory apparatus in S. wawrai n. sp., however, appears to be mainly by the nerve of cerebral origin mentioned above.

Buccal ganglia posterior to the pharynx are present in all Acochlidia, and associated gastro-oesophageal ganglia are known from several hedylopsacean species but not (yet) Hedylopsis ballantinei (Sommerfeldt & Schrödl, 2005; Wawra, 1988, 1989; Schrödl & Neusser, 2010) and also the microhedylaceans Asperspina murmanica (Kudinskaya & Minichev, 1978) and Microhedyle glandulifera (Neusser, Martynov & Schrödl, 2009b; Eder, Schrödl & Jörger, 2011). In S. wawrai n. sp., this arrangement of ganglia innervates the salivary ducts, musculature of the oesophagus and the radula as can be shown from three pairs of nerves plus the unpaired radular nerve, again most of which have not been detected in other acochlidians.

The detection of a high number of previously unknown cerebral features, all possibly bearing useful phylogenetic information, again highlights the usefulness of serial sectioning and accompanying 3D reconstruction.

Osphradium

The observation of a pit-shaped osphradium is the first mention of this sensory organ in Acochlidia. In living S. wawrai n. sp. from Guadalcanal, the osphradium is clearly visible as a paler spot on the right body side. A similar spot is also visible in living Acochlidium sp. from the same locality, in this case rather inconspicuously on the anterior border of an otherwise darkly pigmented bar (own unpublished data). Interestingly, two previous accounts on the aforementioned genera mention body openings in the position of the osphradium: S. paradoxa was erroneously displayed to have the anus in the position of the osphradium (Rankin, 1979: 72) and the original account of A. amboinenseStrubell, 1892 described the copulatory apparatus to open in this place (Bücking, 1933: fig. 2), contradicting observations from other sources or species (e.g. Küthe, 1935; Haase & Wawra, 1996; Brenzinger et al., 2011).

The position of the osphradium—far anterior to what can be considered the mantle border (see Fig. 1A)—appears strange, since the chemosensory organ is usually part of the mantle cavity organs including the gill, anus, genital opening and nephropore (e.g. Thompson, 1976). Apparently the osphradium has moved to a more anterior position after the loss of the mantle cavity in acochlidians. While it appears possible that the osphradium as a discrete organ is expressed only in the large-bodied species, it is also likely to have simply been overlooked so far in the minute interstitial species. These taxa should be critically (re-)investigated regarding the presence of a possible osphradium by searching for the osphradial nerve and its targeted epithelium as part of the epidermis.

Judging from light-microscopical observations, the osphradium of S. wawrai n. sp. resembles the organ of the cephalaspidean Philine (a pit-like structure with a flat bottom; Edlinger, 1980) and can accordingly be divided into two zones: a microvillous inner zone and a ciliated border forming the rim (Fig. 5E), similar to the condition described for the cephalaspidean Scaphander lignarius (Linnaeus, 1758) by Haszprunar (1985b). Since ultrastructural research on the osphradial sensory epithelia has been used to test phylogenetic hypotheses, examination of the organ in Strubellia might reveal features shared with other Panpulmonata that have retained the osphradium.

Hancock's organs

Hancock's organs are cerebrally innervated chemosensory organs situated on the sides of the head; they are present in most shelled opisthobranch gastropods (see Göbbeler & Klussmann-Kolb, 2007). Previously assumed to be missing in Acochlidia (see e.g. Wawra, 1987; Sommerfeldt & Schrödl, 2005), the organs were detected in four mesopsammic species including one Pseudunela species (Edlinger, 1980; see Neusser, Jörger & Schrödl, 2007; Neusser et al., 2011b; own unpublished data). As in the latter species, the Hancock's organs of S. wawrai n. sp. are ciliated epidermal depressions located posterior to the labial tentacles; each organ is innervated by a lateral branch of the rhinophoral nerve. The organs can only be reliably detected in specimens where the head is not strongly retracted into the visceral sac and are thus likely to be overlooked, as was probably the case in S. paradoxa.

Oophagy and radular characters

An asymmetric radula with a formula of n × 1.1.2 is present in most hedylopsaceans and has been regarded as a possible synapomorphy of all Acochlidia (Schrödl & Neusser, 2010). Wawra (1974) described the radula of Solomon Island S. wawrai n. sp. (as S. paradoxa) with a formula of n × 2.1.2, but later corrected this to n × 1.1.2 (Wawra, 1979); the latter can be confirmed by our study. Strubellia paradoxa was also originally described with a formula of n × 2.1.2 (Küthe, 1935). Reexamination of S. paradoxa showed that on the left side there is just a single tooth (Brenzinger et al., 2011). The genus shares with Acochlidium (and Aiteng aterSwennen & Buatip, 2009) the finely serrated rhachidian teeth (e.g. Haynes & Kenchington, 1991; Swennen & Buatip, 2009; Neusser et al., 2011a), however the very elongate rhachidians appear to be a synapomorphy for Strubellia. There are no clear differences in tooth morphology separating S. paradoxa and the Solomon Islands/Vanuatu populations. Counts of radular rows do not show consistent differences among populations and the only connection appears to be with size or ontogenetic stage: very large individuals of S. wawrai n. sp. from Vanuatu had c. 55–60 rows of teeth, medium-sized specimens from the Solomon Islands showed between 48–51 rows (Wawra, 1974, 1979) and 40–46 rows (this study).

The observation of cultured S. wawrai n. sp. feeding on egg capsules of Neritina cf. natalensis is the first direct observation of feeding in Acochlidia. Only Acochlidium amboinense has been mentioned to have “animal remains in the stomach” (Bergh, 1895), while the meiofaunal Pontohedyle milaschewitchii was suggested to be an unspecialized detritus grazer due to its preference of substrates with microbial mats (Hadl et al., 1969; Edlinger, 1980; see Schrödl & Neusser, 2010).

Clusters of neritid egg capsules were seen on rocks at most sampling localities in the Solomon Islands and are an energy-rich potential food source. However, these capsules are strongly reinforced by conchiolin and diatoms or sand grains derived from the food (Andrews, 1935), a fact that appears to deter predation effectively. Only recently have other neritids been shown to feed facultatively on egg capules of other species (Kano & Fukumori, 2010). Strubellia wawrai n. sp. appears to be equipped with a radula specialized for piercing the hard-shelled capsules: the rhachidian teeth are more elongate than in any other acochlidian genus and show considerable wear in the older part of all examined radulae. The finely serrated rhachidians are most likely used to create a slit in the egg capsules through which the contents of the capsules are sucked out, as is suggested by the peristaltic movement of the visceral sac during feeding and the duration of each feeding interval. The sucker-like aspect of the lips surrounding the protruding pharynx is probably related to this mode of feeding. An oophagous habit can also be assumed for S. paradoxa, which shows no major differences in microhabitat or radular morphology (Brenzinger et al., 2011). The closely related Acochlidium species all share the same habitat (as far as can be deduced from the literature) and exhibit highly similar radular morphology (the rhachidian teeth are wider and less dagger-shaped). One might suggest a similar mode of feeding in this genus, perhaps involving niche differentiation with regard to the durability of egg capsules that are fed on; not all egg capsules are equally reinforced and most harden further after their deposition on the rock (Kano & Fukumori, 2010). During the feeding experiment, a specimen of Acochlidium from Guadalcanal was attracted to the presented egg capsules but did not feed (own observations).

Spicules

Subepidermal spicules are found in a number of shell-less heterobranchs and are there considered to be an adaptation to life in the marine interstitial environment (see Rieger & Sterrer, 1975 for a review), functioning as either protective or stabilizing skeletal elements. In some doridoidean nudibranchs, defensive calcareous spicules have also been suggested to be calcium reservoirs (Cattaneo-Vietti et al., 1995). Spicules are present in most acochlidians (Jörger et al., 2008; Schrödl & Neusser, 2010); members of the mesopsammic Asperspina and Hedylopsis have evolved a secondary spicule ‘shell’ that surrounds the visceral sac (e.g. Swedmark, 1968; Schrödl & Neusser, 2010). Wherever present, acochlidian spicules are calcareous, more or less elongate or forming concrements of irregularly formed grainy material.

In form, relative size and distribution, Strubellia spicules resemble those of Pseudunela or Acochlidium (see Bayer & Fehlmann, 1960; Neusser & Schrödl, 2009) but, judging from their location within the body, they do not function as protective elements (the lowest density of spicules is found in the dorsal surface of the visceral sac, the part of the body which remains most prominent in contracted animals). Rod-shaped spicules with blunt ends are found most numerously in the foot, in the head appendages, at the base of the visceral sac and in parallel strips dorsolaterally of the pharynx (‘cephalic spicule grid’). A skeletal function appears likely for the former three examples, in a position where the spicules might well function, in bulk, as stabilizing agents. A protective function (for the CNS) seems reasonable only for the cephalic spicules, as has already been suggested for S. paradoxa by Küthe (1935). We hypothesize an additional function of this spicular arrangement, namely acting as a supporting structure during feeding: the spicules might interlock and thus stabilize the pharyngeal region, while the head is pressed hard onto the neritid egg capsules in order to puncture their walls with the radula. Similar aggregations of spicules close to the pharynx have been reported in other acochlidian genera: in the microhedylacean Asperspina and Pontohedyle (Jörger et al., 2008) and as a “postpharyngeal spicule collar” in the hedylopsacean Tantulum elegans (Neusser & Schrödl, 2007; Schrödl & Neusser, 2010); in Acochlidium bayerfehlmanni Wawra, 1980 (Bayer & Fehlmann, 1960; as A. amboinense) spicules are stated to form “a ring around the esophagus” similar to the situation found in Strubellia.

Cephalic gland

The loose aggregation of cells covering the cerebral ganglia was present in all individuals examined in this study, but has not been reported for any acochlidian species, including S. paradoxa. Neusser et al. (2007, 2009b) mention both “cells above the cerebral commissure” and “lateral bodies” attached to the cerebral ganglia in the interstitial acochlidians Asperspina murmanica and Hedylopsis ballantinei; these cells were, however, embedded within the connective sheath of the cerebral commissure. Supposedly endocrine “dorsal bodies”—surrounded by a connective sheath and associated with the cerebral ganglia—are common among pulmonates, where there is considerable diversity regarding structure and innervation (e.g. Boer, Slot & van Andel, 1968); they have been shown to be more active during female reproduction (Saleuddin, Ashton & Khan, 1997). In S. wawrai n. sp. there appears to be no connective sheath and there are no histologically detectable differences between juveniles and mature specimens.

In histology (loose tissue with yellow-stained vesicles visible in serial sections) and position the structure also resembles the ‘blood’ gland found in some anthobranch nudibranchs, e.g. the doridoidean Corambe lucea Marcus, 1959 (Schrödl & Wägele, 2001) and the dexiarchian Doridoxa (Schrödl, Wägele & Willan, 2001). However, the presence of apparently osmiophilic, yellow-staining vesicles indicates fatty substances, as are present in the digestive gland, possibly implying a function as an additional fat-storing structure. Ultrastructural research on cell anatomy and affiliation to the CNS is needed for conclusive identification of this organ, which might represent an apomorphy for either Strubellia or Acochlidiidae.

Heart and kidney

Only few shell-less heterobranchs venture into habitats that are regularly influenced by freshwater, e.g. some nudibranchs and sacoglossans (Barnes, 1994). The excretory system of the sacoglossan Alderia modesta (Lovén, 1844), found on partially brackish intertidal mudflats (Evans, 1951), has been examined in detail but lacks a heart and shows no apparent elaboration of its sac-like kidney (Fahrner & Haszprunar, 2001). Members of the recently described Aitengidae (also Acochlidia) live amphibiously among mangroves or coastal rocks, and show an elaborate system of branched dorsal vessels (resembling the condition found in many plakobranchioid sacoglossans) which might originate from a histologically similar and sac-like kidney (Swennen & Buatip, 2009; Neusser et al., 2011a). Neither condition appears very similar to that found in Strubellia.

The circulatory and excretory systems of S. wawrai n. sp. show several apparent morphological adaptations to permanent life in fresh water, namely specialized cell types in the heart, elongated lumina of the kidney and nephroduct, and possibly the loop in the distal nephroduct. A strongly vacuolated epicardium and discrete thick-walled cells inside the lumen of the heart have been described only from S. paradoxa and the brackish-water Pseudunela espiritusanta (Neusser & Schrödl, 2009; Brenzinger et al., 2011). These cells possibly involve a novel site of ultrafiltration (on the ventricle) and aggregations of rhogocytes, however in both cases ultrastructural investigation is needed to identify those cell types. Muscular bridges spanning the lumen of the heart, presumably an aspect of an enhanced circulation, have been mentioned for Acochlidium amboinense (Bücking, 1933) and S. paradoxa.

In Strubellia there appears to be a functional division of otherwise elongated excretory lumina, judging from the separation of at least three histologically different zones (proximal and distal kidney lumina and nephroduct). The presence of the conspicuous upward loop of the distal nephroduct, which is closely associated with the pericardial wall, hints at the presence of a fourth zone involved in the modification of the primary urine. Again, ultrastructural studies are needed to test these observations derived from light microscopy.

Elongation of excretory lumina has been shown to be a feature of hedylopsaceans and is conspicuously present in the coastal mesopsammic Pseudunela cornuta (Challis, 1970) and P. espiritusanta (Neusser & Schrödl, 2009; Neusser et al., 2009a, Neusser, Jörger & Schrödl, 2011b) and the more basal but limnic Tantulum elegans (Neusser & Schrödl, 2007). All of these species display an elongate kidney with divided lumina and U-shaped nephroduct with distal loop. Members of the marine mesopsammic genus Hedylopsis also show the elongate, complex kidney, but have a short nephroduct (Fahrner & Haszprunar, 2002; Sommerfeldt & Schrödl, 2005). This means that the elaborate excretory system found in Strubellia is already more or less present in marine or brackish-water Pseudunela species (Neusser et al., 2011b) and thus further adaptations to life in freshwater are likely to have happened on an ultrastructural level.

There is only scarce information on the circulatory and excretory systems of Acochlidium species, although it appears to be more sophisticated. Bücking (1933) mentioned a branching vessel on the dorsal side of the visceral sac (superficially similar to that found in sacoglossans or Aitengidae) and the presence of multiple renopericardial funnels. It should be critically compared with the condition found in Strubellia to trace the evolution of characters in these organ systems that are crucially important in the colonization of limnic habitats.

Genital ontogeny

As was confirmed for S. paradoxa by Brenzinger et al. (2011), S. wawrai n. sp. appears to be a sequential, protandric hermaphrodite, as is otherwise known only for Tantulum elegans and Hedylopsis species among Acochlidia (Wawra, 1989; Neusser & Schrödl, 2007; Kohnert et al., 2011). The change of sex during ontogeny can be deduced (1) from the presence of two allosperm receptacles in otherwise male specimens and (2) the presence of intermediate stages (females with bursa copulatrix, seminal groove and copulatory apparatus still present but in various stages of reduction) (Wawra, 1988; present study). Sperm transfer appears to be via copulation and mainly in the male phase, after which the sex changes to a female state (gonad producing oocytes; female gland mass developed) while the strictly male genital features become reduced. This change is likely to be rapid since intermediate stages have rarely been found in previous studies of Strubellia species (Küthe, 1935; Wawra, 1988).

Genital system (posterior part)

The genital system of S. wawrai n. sp. was largely described by Wawra (1974, 1988; as S. paradoxa), assuming first gonochorism and then sequential hermaphroditism. We can confirm the description of the posterior genital system with a full set of sperm storing organs, i.e. the ampulla for autosperm and two allosperm receptacles (receptaculum seminis, bursa copulatrix), which is a condition known from the marine Pseudunela cornuta and the brackish-water P. espiritusanta, among Acochlidia. However, in both the latter species the receptaculum seminis is situated more proximally to the gonad than the sac-like ampulla (Neusser & Schrödl, 2009; Neusser et al., 2009a); this is in contrast to S. paradoxa and S. wawrai n. sp. where the receptaculum seminis is distal to the tubular ampulla. Except for its functional change during ontogeny, the gonad of Strubellia varies from the aforementioned genus by the separation into distinct follicles and the high number of eggs, both features shared with Acochlidium fijiense (Haynes & Kenchington, 1991; Haase & Wawra, 1996), probably reflecting a higher reproductive potential per individual. The female gland mass, developed from the very long gonoduct in ‘males’, is tubular all along and shows three histologically separable parts. This organ system is highly variable among Pseudunela and other acochlidians (but see Neusser et al., 2011b), where usually at least some of the glands are sac-like extensions and sometimes there appear to be only two different glands; the situation in Acochlidium species is unclear (see Schrödl & Neusser, 2010; Brenzinger et al., 2011).

The bursa copulatrix, reduced in the female phase, is similar to that of the marine hedylopsaceans in its morphology (bulbous, with thinner stalk) and its location next to the genital opening. Acochlidium on the other hand has been described to lack any allosperm receptacles due to its supposedly hypodermal mode of insemination (Haase & Wawra, 1996). The genital diverticulum next to the genital opening is a feature known also from S. paradoxa (Brenzinger et al., 2011); its variability in size (largest in one specimen from Vanuatu) and reduction in females hint at a function in copulation.

Strubellia shares the supposedly ‘primitive’ open seminal groove connecting to the genital opening distal to the bursa with Hedylopsis spiculifera Kowalevsky, 1901 (see Wawra, 1989). Other hedylopsaceans have been described to have a closed vas deferens that splits off the distal gonoduct proximal to the bursa and runs below the epidermis of the right body side (e.g. Neusser et al., 2009a; see Schrödl & Neusser, 2010). We suggest that the open seminal groove is not a plesiomorphic character per se, but is likely connected with ontogenetic sex change; as a transient feature, the duct remains only as a groove and is not sunk below the epidermis.

Cephalic copulatory apparatus

We disagree with Wawra's (1974) description of the cephalic copulatory apparatus which was based on dissected material missing the penis and associated glands; as in the description of S. paradoxa by Küthe (1935), the basal finger was erroneously interpreted as the penis. The copulatory organs of S. wawrai n. sp. consist of two distinct muscles with connected (para-)prostatic glandular systems as in S. paradoxa, resembling the Pseudunela species known in detail (Neusser & Schrödl 2009; Neusser et al., 2009a, 2011b). Strubellia, however, lacks the hollow penial stylet and instead features a solid spine near the penial opening, precluding sperm transfer by hypodermal injection which is believed to occur in Pseudunela, Acochlidium and a number of heterobranchs that possess one or several hollow penial stylets as an extension of the distalmost vas deferens (see Gascoigne, 1974; Haase & Wawra, 1996; Neusser et al., 2009a).

The long and hollow stylet of the basal finger, however, appears to be used for (hypodermal) injection of the paraprostatic secretion; only in Strubellia does the stylet have the longitudinal groove. Both muscle and chitinous elements are more pronounced in Strubellia than in other genera, which imply a relatively higher importance of the paraprostatic system in this genus. Stylet morphology (and perhaps that of the penial spine) may also present a possibility to distinguish at least male specimens from the two Strubellia species by SEM: the basal finger stylet of S. wawrai n. sp. appears to be more elongate than that of S. paradoxa and shows a bent or slightly hooked tip (Table 4). This distinction is however only based on few male specimens and disregards the possibility of the stylet being flexible, as is mentioned for the chitinous penial stylets of some sacoglossan species (Gascoigne, 1974).

Table 4.

Comparison of morphological data of Strubellia wawrai n. sp. and S. paradoxa.

 S. wawrai n. sp.
 
S. paradoxa (Strubell, 1892)
 
Reference Wawra (1974, 1979, 1988Present study Present study Küthe (1935) Brenzinger et al. (2011) 
Collecting site Guadalcanal, Solomon Is Guadalcanal, Solomon Is Espiritu Santo, Vanuatu Ambon, Indonesia Ambon, Indonesia 
Max. recorded body size ∼2.5 cm ∼2.0 cm ∼3.5 cm ∼2 cm ∼1 cm 
Radula formula 48–51 × 1.1.2 43–46 × 1.1.2 59 × 1.1.2 48–56 × 2.1.2 38 × 1.1.2 
1st lateral tooth denticle Present Present Present Absent Present 
Length of basal finger stylet 1 mm 0.75–1 mm 0.5 mm 0.6 mm 
Stylet form Elongate, tip hooked Elongate, tip bent Rather stout Rather stout 
Distal paraprostatic duct Divided (Wawra, 1974: fig. 4) Undivided Divided Divided 
Genital diverticle Small Large Small 
Penial thorn ?, curved Concave, curved Flat (?), curved Flat, curved 
 S. wawrai n. sp.
 
S. paradoxa (Strubell, 1892)
 
Reference Wawra (1974, 1979, 1988Present study Present study Küthe (1935) Brenzinger et al. (2011) 
Collecting site Guadalcanal, Solomon Is Guadalcanal, Solomon Is Espiritu Santo, Vanuatu Ambon, Indonesia Ambon, Indonesia 
Max. recorded body size ∼2.5 cm ∼2.0 cm ∼3.5 cm ∼2 cm ∼1 cm 
Radula formula 48–51 × 1.1.2 43–46 × 1.1.2 59 × 1.1.2 48–56 × 2.1.2 38 × 1.1.2 
1st lateral tooth denticle Present Present Present Absent Present 
Length of basal finger stylet 1 mm 0.75–1 mm 0.5 mm 0.6 mm 
Stylet form Elongate, tip hooked Elongate, tip bent Rather stout Rather stout 
Distal paraprostatic duct Divided (Wawra, 1974: fig. 4) Undivided Divided Divided 
Genital diverticle Small Large Small 
Penial thorn ?, curved Concave, curved Flat (?), curved Flat, curved 

The paraprostatic duct has been mentioned to split at the base of the stylet in S. paradoxa and S. wawrai n. sp. from Guadalcanal (Küthe, 1935; Wawra, 1974; Brenzinger et al., 2011), whereas it is undivided in the specimen from Vanuatu. This feature is of unclear function and may again be related to the individual stage of ontogeny, but is hard to detect and deserves reexamination.

Species-level relationships

Molecular data indicate that there are three separate lineages in the genus Strubellia, the first offshoot known only from the single juvenile specimen from Sulawesi examined herein. More material is needed to establish this population as a new species.

The eastern Melanesian specimens of S. wawrai n. sp. form a clade that is sister group to S. paradoxa from Ambon, Indonesia. Both clades receive strong bootstrap support and sequence divergence in COI (c. 12–13%) is relatively high. Both Species Identifier and parsimony network analyses indicate specific differences between S. paradoxa and S. wawrai n. sp. Given the 3,500-km distance between Ambon and the Solomon islands, this divergence is not surprising. Separation of S. wawrai n. sp. by only morphological means is not straightforward, since most organ systems previously used to separate acochlidian species are highly similar. However, there are some differences in parts of the copulatory apparatus, including length and curvature of the basal finger stylet (elongate and apically curved vs. rather stout and short in S. paradoxa; Brenzinger et al., 2011) and form of the penial spine that might be useful features discernible by SEM. In both cases these differences refer to few mature individuals only, so ranges of intraspecific or ontogenetic variations remain poorly known. Variations in radular row counts, as already mentioned, are likely to be attributable to the size of the individuals examined. The presence of a second lateral plate in S. paradoxa has to be formally confirmed (Brenzinger et al., 2011).

Summing up, potential differences in relevant parts of the copulatory organs, together with genetic evidence, leave little doubt that the populations from Ambon and Melanesia represent distinct species.

On a population level, the observed size disparity between mature specimens of S. wawrai n. sp. from the Solomon Island and Vanuatu is an obvious morphological difference, especially since female individuals from Vanuatu with remaining male genitalia were larger than already fully female specimens from Guadalcanal (Table 4). This observed delay in ontogeny is hard to explain given knowledge of the genetic similarity between the populations, but is perhaps attributable to ecological factors. Observed differences in the size of the genital diverticulum and the distal division of the paraprostatic duct (present/absent) are also likely to be variable during ontogeny. Analysis of molecular divergence shows that the Guadalcanal and Espiritu Santo populations of S. wawrai n. sp. are very similar, with the clade comprising the latter population nested inside the former, indicating that the split is too recent to be obvious from COI divergence. We therefore regard the two populations as a single species that might be close to separating into two species, with geographic separation preventing regular gene flow.

Habitats and dispersal

The localities discovered in this study fit well with the described habitat regarding physical and chemical properties, i.e. limestone slabs at the edge of shallow streams carrying mineral-rich and slightly alkaline water. Strubellia species co-occur with neritid gastropods (Starmühlner, 1976; Haynes, 2000). This is significant, since we observed S. wawrai n. sp. feeding on neritid eggs, resolving a longstanding mystery. In addition we know that different species and populations occur in limnic systems of more or less distant islands and archipelagos surrounded by sea.

So, how do limnic slugs, generally hiding away under rocks during the day, disperse to or maintain gene flow between different localities, as is implied by the molecular analysis? Other stream gastropods with similar lifestyles, such as the numerous neritid species occurring in the rivers of Indo-West Pacific islands, reach distant islands by means of planktonic larvae (Haynes, 1988; Myers, Meyer & Resh, 2000) and regularly recolonize them; juveniles of at least one species even migrate by sometimes ‘hitchhiking’ upstream on the shell of larger individuals (Kano, 2009). Assuming a similarly amphidromic life with larvae hatching in freshwater and returning to it after a period of time and metamorphosis in the sea (see McDowall, 2007) would explain the observed distribution in Strubellia—but there are yet no observations of eggs or larvae of Strubellia. However, Acochlidium fijiense is known to produce gelatinous egg masses from which veligers hatch that are apparently not able to survive in fresh water (Haynes & Kenchington, 1991). In seawater, these veligers quickly metamorphosize into ‘adhesive’-type larvae which remain alive for at least 2 months and glue themselves e.g. to the wall of the Petri dish they are kept in (own observations on Acochlidium sp.). This shows that limnic Acochlidium, and possibly already the common ancestor with Strubellia, have evolved a specialized larval type that might be able to disperse between islands of archipelagos leading to the colonization of rivers, involving a neritid-like amphidromic lifestyle. On one hand, these adhesive larvae, if quickly glued to a substratum outside the river, could avoid being drifted away too far into the ocean. Following juvenile neritids on their necessary movement upstream (possibly while glued to a shell during metamorphosis) and then feeding on their eggs would be a novel and efficient kind of symbiosis. On the other hand, it seems possible that this type of larva is able to use more mobile and far-ranging organisms as vectors between islands (planktonic organisms, fish, birds, boats). While acochlidiid larvae can survive in the laboratory for months without any movement or food uptake, metamorphosized juveniles would have to feed. Such juveniles would still be in the size range of most marine acochlidians (1–2 mm) and are not likely to prey on adult food, i.e. strongly mineralized neritid egg capsules. A juvenile stage feeding on microbial mats, mucus, algae or detritus is thus hypothesized. Field observations and laboratory experiments are needed to confirm the hypothesized life-history traits of Strubellia.

Larvae sticking to floating or swimming objects may therefore be the ‘missing’ dispersive stages explaining interisland dispersal, such as from the Solomon Islands to Vanuatu in the case of S. wawrai n. sp., or the colonization of Palau or Fiji in the case of Acochlidium bayerfehlmanni and A. fijiense (Bayer & Fehlmann, 1960; Haynes & Kenchington, 1991). Since limnic Acochlidiidae are estimated to have originated in the Palaeogene (Jörger et al., 2010a), this long period would present a timeframe to have enabled dispersal via island-hopping, facilitated by lower sea levels and shorter distances between islands in Indonesia during much of the period. Dispersal to the west might have been limited by deeper-water currents being deflected at the border of the Southeast Asian continental shelf, as is indicated by Wallace's-line distributional patterns of marine organisms with pelagic larval stages (Barber et al., 2000). The lack of records of acochlidiids west of the Wallace line hints at a similar limitation. On the other hand, it appears likely that numerous populations of acochlidiids are yet to be discovered and also that many have become extinct.

Phylogeny of Strubellia and evolution of characters

The molecular phylogeny of the acochlidiids shows Strubellia to have originated in Indonesia. The genus is sister group to the morphologically more derived Acochlidium and Palliohedyle, these in turn being sister group to the marine interstitial Pseudunelidae. This configuration is congruent with the previously proposed phylogenies of Acochlidia, based on morphology (Schrödl & Neusser, 2010) or molecular markers (Jörger et al., 2010a).

According to the new results, the apomorphies for Acochlidiidae are the limnic habitat, benthic and probably amphidromic lifestyle, accompanied by large body size and distinct epidermal pigmentation, and the finely serrated rhachidian teeth. The visible distinction of the anterior mantle border and heart ‘bulb’, complex kidneys and the bipartite copulatory organs with spines and associated glands are already present in the mesopsammic Pseudunela species (Neusser & Schrödl, 2009; Neusser et al., 2009a, 2011b).

Presence of an osphradium and oophagy might represent further apomorphies; however, we suggest that the presence of epidermal sensory cells is likely at least in the hedylopsacean species with an osphradial ganglion. Furthermore, we suggest that a piercing-and-sucking mode of feeding is typical for Acochlidia, since all share the muscular pharynx, a slender radula that appears ill-equipped for grazing, and sometimes arrays of spicules surrounding the pharynx. For the meiofaunal species, instead of grazing, sucking liquid contents from soft, encapsulated food such as large-bodied protists or eggs of sand-dwelling organisms might explain the coloration of some species' digestive glands (e.g. brown or green in Pontohedyle milaschewitchii; Jörger et al., 2008), the lack of both abraded teeth and mineral residues in the digestive system. The sacoglossan-like monostich radula of the microhedylacean Ganitidae (Challis, 1968) would thus be specialized for a specific type of food, but not a unique mode of feeding within the group. Given the similarity of the pharynx and radula (slender ribbon, triangular median tooth with serrated margins, flat or reduced laterals) in Sacoglossa (especially the basal Cylindrobulla; Mikkelsen, 1998), Aitengidae (Swennen & Buatip, 2009; Neusser et al., 2011a), Amphibolidae (Golding, Ponder & Byrne, 2007) and Glacidorbidae (Ponder, 1986; Ponder & Avern, 2000), the suggested mode of feeding by piercing and sucking might represent a basal panpulmonate feature. Somewhat similar to Strubellia, both Sacoglossa and Aiteng ater are known to feed by puncturing internally soft food (siphonal algae and insect pupae, respectively) and sucking out the contents (Jensen, 1980, 1981; Swennen & Buatip, 2009); some Sacoglossa are also known to feed on the more or less gelatinous egg masses of opisthobranch gastropods (see Jensen, 1980; Coelho, Malaquias & Calado, 2006). However, some Euopisthobranchia sensuJörger et al. (2010a) show similar, narrow radulae with serrated rhachidian and flat lateral teeth, e.g. species of the cephalaspidean Toledonia (Marcus, 1976; Golding, 2010), and also several nudibranchs (such as the oophagous aeolidioidean Favorinus; Schmekel & Portmann, 1982), making it difficult to detect phylogenetic patterns. An example is the proposed relationship of Toledonia and Acochlidia on the basis of radular morphology (Gosliner, 1994), which according to more recent hypotheses clearly represents a case of convergent evolution (Jensen, 1996; Sommerfeldt & Schrödl, 2005; Jörger et al., 2010a; Schrödl et al., 2011). Furthermore, a slender piercing radula is also present in Omalogyra atomus (Philippi, 1841) (‘lower Heterobranchia’; Bäumler et al., 2008).

Synapomorphies of Strubellia appear to be the reddish-brown pigmentation, very slender rhachidian teeth, three receptacles in the male phase, the genital diverticulum, the enhancement of the basal finger with the stylet having a lateral groove, and the possession of a single flat hook on the penis instead of a hollow penial stylet.

The organization of the posterior genital system of Strubellia essentially conforms to the ‘classic’ idea of plesiomorphic monauly that was suggested to be the condition found in the hermaphroditic “opisthobranch common ancestor” (Ghiselin, 1966; Gosliner & Ghiselin, 1984), however the condition of Strubellia is fundamentally different. All hedylopsaceans are (special) androdiaulic hermaphrodites (Schrödl & Neusser, 2010; Schrödl et al., 2011) except for Strubellia (and Hedylopsis species; see Wawra, 1989; Sommerfeldt & Schrödl, 2005; Kohnert et al., 2011). The derived phylogenetic position of Strubellia (Jörger et al., 2010a; Schrödl & Neusser, 2010) suggests either a reversal to a monaulic system (with sperm and oocyte pathways not separated anatomically but in time, with a secondarily open seminal groove) or multiple developments of diauly among Acochlidia. The presence of allosperm receptacles already in the male phase might have led to the evolution of defined breeding seasons in Strubellia, hinted at by the strong skew among sexes revealed from sampling in all known localities: specimens were either predominantly juvenile, or only either male or female (Küthe, 1935; Wawra, 1988; present study). This might also be related to the observation that Strubellia generally aggregates in groups: If Strubellia has defined breeding seasons (possibly the rainy seasons accompanied by changes in riverine water levels) then aggregations of numerous specimens might mate after which the specimens change sex synchronously, spawn and then either die or fully reduce their genital organs, as was suggested for A. fijiense (Haynes & Kenchington, 1991). This appears at least possible, since complete reduction of the large copulatory apparatus during ontogeny can be deduced from the observations presented here, and a strong reduction of body size likely connected with a reduction of organs has been observed after periods of starvation in the specimens maintained in aquaria for this study.

Strubellia differs externally from Acochlidium and Palliohedyle by its more slender body, elongate visceral sac (versus leaf-shaped and flattened) and uniform reddish coloration (vs greenish-brown and black pigmentation), making it externally more similar to the aforementioned Pseudunela species (e.g. Haynes & Kenchington, 1991; own observations). According to the literature, internal differences from the better-known Acochlidium species include shape of the rhachidian teeth (very elongate in Strubellia vs triangular), morphology of the penis (relatively small with single apical thorn in Strubellia vs large and multi-spined; e.g. Wawra, 1979, 1980; Haase & Wawra, 1996) and basal finger (larger than the penis and with long stylet in Strubellia), the mode of genital ontogeny (protandric hermaphroditism in Strubellia vs hermaphroditism; Haynes & Kenchington, 1991) and the elaboration of visceral organs (multiple renopericardial funnels, digestive gland lobes, praeampullary gonoducts and branched, dorsally situated vessels connected to the heart in Acochlidium; Bücking, 1933; Haase & Wawra, 1996). Since the only comprehensive anatomical description of an Acochlidium species is very old (Bücking, 1933) and the only detailed study of the genital system includes characters that are still unclear (e.g. a connection between the ampulla and the digestive gland; Haase & Wawra, 1996), revision of the aforementioned anatomical features is urgently needed to trace the evolution of these unique limnic slugs.

ACKNOWLEDGEMENTS

Many thanks to Alison Haynes (Suva) for sharing specimens from Efate and Matthias Glaubrecht (Berlin) for sharing material collected on Ambon. Yasunori Kano (Tokyo) is thanked for his help during the field trips to Espiritu Santo and Guadalcanal. We would like to acknowledge Eva Lodde for her help with the histological methods and Roland Melzer, Enrico Schwabe and Jens Bohn for their help with the SEM (all ZSM). Many thanks go to Martin Heß (LMU Munich) for his help in creating the interactive figures. This study was financed by a grant of the German Research Foundation (DFG SCHR 667/4-3 to M.S.) and a PhD scholarship from the VW Foundation to K.M.J. Three-dimensional reconstruction was financed by the GeoBioCenter/LMU München. T.P.N. is grateful to Philippe Bouchet (Paris) for the opportunity to join the Mission MNHN/PNI/IRD Santo 2006 to Vanuatu. The SANTO 2006 Expedition was organized by Museum National d'Histoire Naturelle, Paris, Pro Natura International (PNI) and Institut de Recherche pour le Développement (IRD). It operated under a permit granted to Philippe Bouchet (MNHN) by the Environment Unit of the Government of Vanuatu. The Marine Biodiversity part of the expedition, a part of Census of Marine Life's CReefs programme, was specifically funded by grants from the Total Foundation and the Sloan Foundation. Finally, we would like to thank two anonymous referees for their helpful comments on the manuscript.

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