Within-host speciation events in yoyo clams, obligate commensals with mantis shrimps, including one that involves a change in microhabitat and a loss of specialized traits

Within-host speciation events in yoyo clams, obligate commensals with mantis shrimps, including... Abstract Compared to host shifts, the importance of within-host cladogenesis in the diversification of symbionts remains less well understood in marine systems. Yoyo clams (Galeommatidae: Vasconiellinae) are a clade of marine bivalves that live commensally with burrowing mantis shrimp. Almost all yoyo clams byssally-attach to the host burrow wall via a specialized hanging foot structure bearing a thread-like posterior extension. In contrast, Parabornia squillina (Vasconiellinae) byssally-attaches directly to the host shrimp and lacks a hanging foot structure. In this study, we examine phylogenetic relationships among vasconiellines by performing molecular analyses based on five genes (28S and 16S rRNA, H3, COI and ANT). We found evidence for two within-host speciation events among Floridian vasconiellines commensal with the same mantis shrimp host, Lysiosquilla scabricauda. One involved a cryptic sister species pair of burrow-wall commensals. The other involved the ectocommensal P. squillina and its somewhat unexpected sister taxon, the burrow-wall commensal Divariscintilla octotentaculata. This latter result suggests that a habitat shift from host burrow wall to host body surface occurred while retaining the same host species and led to the loss of the specialized hanging foot structure. Our findings suggest that ostensibly modest within-host ecological shifts can lead to major morphological changes in these clams. adaptation, burrow, commensalism, Galeommatoidea, habitat shift, host shift, specialization, Stomatopoda, symbiosis INTRODUCTION Symbiotic and parasitic organisms represent a large fraction of the Earth’s biodiversity (Windsor, 1998; Poulin & Morand, 2004; Moran, 2006). Host shifting, an evolutionary change in host species, has been recognized as the major driver of speciation in these organisms both in terrestrial (Coyne & Orr, 2004; Matsubayashi, Ohshima & Nosil, 2010) and in marine realms (Duffy, 1996; Munday, van Herwerden & Dudgeon, 2004; Faucci, Toonen & Hadfield, 2007; Tsang et al., 2009; Goto et al., 2012; Hurt et al., 2013). However, the role of host shifts in diversification may be less important than previously thought (Winkler & Mitter, 2008; Imada, Kawakita & Kato, 2011; Nakadai & Kawakita, 2016), and alternative speciation processes, such as allopatric speciation and within-host speciation, can also play an important role in the diversification of parasites and symbionts (Imada et al., 2011; Nakadai & Kawakita, 2016; Jahner et al., 2017). Within-host speciation driven by ecological shifts has been studied mainly in phytophagus insects (e.g. gall-inducing insects) (Cook et al., 2002; Joy & Crespi, 2007). In these systems, speciation is probably initiated by adaptations to different host tissues (Cook et al., 2002; Joy & Crespi, 2007; Althoff, 2014) or different life history stages (Zhang et al., 2015). Similar speciation patterns have also been reported in other systems, such as avian malaria (Pérez-Tris et al., 2007) and freshwater fish parasites (Vanhove et al., 2016). However, evidence for within-host speciation driven by ecological shifts remains limited in marine systems. The superfamily Galeommatoidea is a group of small-bodied bivalves that exhibit high species diversity in shallow-water environments (Bouchet et al., 2002; Paulay, 2003). Many galeommatoidean species are commensal with benthic invertebrates in soft sediments (Boss, 1965a; Morton & Scott, 1989; Goto et al., 2012; Li, Ó Foighil & Middelfart, 2012). Galeommatoideans associate with various animal phyla (e.g. Arthropoda, Echinodermata and Annelida) (Boss, 1965a; Morton & Scott, 1989), with many species demonstrating high fidelity to a particular host species or genus (Sato et al., 2011), with some exceptions (Li & Ó Foighil, 2012). Although recent molecular phylogenies suggest that host shifts between distantly related taxa occurred frequently in some clades of Galeommatoidea (Goto et al., 2012; Li, Ó Foighil & Strong, 2016), some congeneric galeommatoidean species share the same host (Mikkelsen & Bieler, 1989, 1992; Goto & Kato, 2012; Goto et al., 2014; Goto, Ishikawa & Hamamura, 2016), suggesting the possibility of within-host speciation in these lineages. The subfamily Vasconiellinae is a group of galeommatoideans that includes seven genera (Huber, 2015). Most species have reduced shells covered by hypertrophied mantles with highly developed sensory tentacles (Popham, 1939; Mikkelsen & Bieler, 1989, 1992; Fig. 1). Among them, four genera (Divariscintilla, Phlyctaenachlamys, Parabornia and Ephippodontomorpha) are known as symbionts of burrowing mantis shrimp (Stomatopoda: Lysiosquillidae) (Boss, 1965b; Judd, 1971; Mikkelsen & Bieler, 1989, 1992; Simone, 2001; Middelfart, 2005; Yamashita, Haga & Lützen, 2011). Except for Parabornia species, these vasconiellines live suspended from the burrow walls of their stomatopod hosts by means of a specialized hanging foot structure having a thread-like posterior extension (Judd, 1971; Mikkelsen & Bieler, 1989, 1992; Middelfart, 2005; Yamashita et al., 2011; Fig. 1). In association with this posture, these clams engage in a characteristic ‘yo-yo’ up and down motion by contracting and relaxing the posterior foot (Mikkelsen & Bieler, 1989, 1992; Fig. 1B; see Supporting Information, Movie S1), hence the informal ‘yoyo clam’ name (Mikkelsen & Bieler, 1989). Figure 1. View largeDownload slide Diversity of Floridian yoyo clams (Galeommatoidea: Galeommatidae: Vasconiellinae: Divariscintilla) collected from the burrow of Lysiosquilla scabricauda. A. Divariscintilla yoyo. B. Hanging behaviour of D. yoyo. C. D. aff. yoyo. D. D. troglodytes. E. D. octotentaculata. F. D. luteocrinita. Arrows indicate posterior foot extension (hanging foot structure). Figure 1. View largeDownload slide Diversity of Floridian yoyo clams (Galeommatoidea: Galeommatidae: Vasconiellinae: Divariscintilla) collected from the burrow of Lysiosquilla scabricauda. A. Divariscintilla yoyo. B. Hanging behaviour of D. yoyo. C. D. aff. yoyo. D. D. troglodytes. E. D. octotentaculata. F. D. luteocrinita. Arrows indicate posterior foot extension (hanging foot structure). Divariscintilla includes seven described species that have been recorded from Florida, New Zealand and Japan (Judd, 1971; Mikkelsen & Bieler, 1989, 1992; Yamashita et al., 2011). Interestingly, five species (D. yoyo Mikkelsen & Bieler, 1989, D. troglodytes Mikkelsen & Bieler, 1989, D. octotentaculata Mikkelsen & Bieler, 1992, D. luteocrinita Mikkelsen & Bieler, 1992 and D. cordiformis Mikkelsen & Bieler, 1992) are at present known only from south-eastern Florida (Mikkelsen & Bieler, 1989, 1992), living exclusively in the burrows of a single stomatopod host, Lysiosquilla scabricauda (Lamarck, 1818) (Mikkelsen & Bieler, 1989, 1992). In addition to Divariscintilla, the ectocommensal vasconielline Parabornia squillina Boss, 1965 also utilizes L. scabricauda as a host in Florida (Boss, 1965b; Mikkelsen & Bieler, 1992), although this species has also been recorded from Mississippi (Boss, 1965b) and Panama (Moore & Boss, 1966). Unlike Divariscintilla spp., Parabornia spp. live attached to the host body surface (Boss, 1965b). Taken together, L. scabricauda hosts six vasconielline species in eastern Florida, thereby providing an invaluable opportunity to investigate the possibility of within-host speciation. Burrow-wall-commensal Divariscintilla spp. and ectocommensal Parabornia spp. are thought to be closely related because many possess flower-like organs near the base of the foot in addition to morphological similarity in the posterior foot structure (Bieler & Mikkelsen, 1992; Mikkelsen & Bieler, 1992), although the posterior foot extension of the genus Parabornia is much shorter than that of the genus Divariscintilla (Mikkelsen & Bieler, 1992). Unlike the genus Parabornia, the shells of Divariscintilla spp. are partially to fully covered by mantle tissue that bears highly developed sensory tentacles (Boss, 1965b; Mikkelsen & Bieler, 1989, 1992; Simone, 2001; Fig. 1). These differences in mantle coverage and foot structure are thought to reflect the differences in host utilization between the two genera (burrow-wall-commensals vs. ectocommensals). However, the phylogenetic relationship of these genera remains unexamined. In this study, we addressed the following questions: (1) are the six vasconiellines associated with L. scabricauda in Florida monophyletic, and if so, (2) how have evolutionary transitions between burrow-wall-commensal and ectocommensal lifestyles occurred in this bivalve clade? We performed molecular analyses of Vasconiellinae based on two nuclear genes (28S rRNA and histone H3) and three mitochondrial genes [cytochrome c oxidase subunit I (COI), 16S rRNA and adenine nucleotide translocator (ANT)]. Because the ectocommensal lifestyle of Parabornia was only briefly mentioned in previous studies (Boss, 1965b; Mikkelsen & Bieler, 1992; Simone, 2001), we observed living P. squillina to further understand its ecological adaptations to an ectocommensal lifestyle. Lastly, morphological characteristics of Divariscintilla and Parabornia were compared to reveal if morphological differences between genera are associated with ecological shifts. MATERIAL AND METHODS Sample collection and observations Sampling was performed in intertidal sand flats in the Indian River lagoon (Fort Pierce, FL, USA), the type locality of the five Divariscintilla species (D. yoyo, D. troglodytes, D. octotentaculata, D. luteocrinita and D. cordiformis) (Mikkelsen & Bieler, 1989, 1992) during 30 May–4 June 2016 and 26–31 January 2017. We collected Divariscintilla species from L. scabricauda burrows using stainless steel bait pumps (‘yabby pumps’) and 1–2-mm mesh sieves. With the exception of D. cordiformis, a very rare species at this site (Mikkelsen & Bieler, 1992), all known Floridian Divariscintilla species were collected. We also collected P. squillina from the ventral body surface of it host L. scabricauda, which were captured manually using fish bait. The bivalves were kept for several days in aquaria for observations and then preserved in 100% ethanol for DNA analyses. Additionally, alcohol-fixed museum specimens of Divariscintilla spp. and close relatives were loaned from the Muséum National d’Histoire Naturelle, Paris, Field Museum, Florida Museum of Natural History and Museum of New Zealand, Te Papa Togarewa for DNA analyses (Table 1). The DNA sequences of Divariscintilla and closely related species used in previous phylogenetic studies were obtained from GenBank (Table 1). For outgroups, we used several galeommatoideans that were identified to be closely related to Vasconiellinae by Li et al. (2016). Table 1. Species used for molecular phylogenetic analyses with museum catalogue number or private specimen ID, sampling localities and GenBank accession numbers Species  Specimen ID  Sampling locality  28S rRNA  16S rRNA  COI  H3  ANT  Divariscintilla luteocrinita Mikkelsen & Bieler, 1992  FMNH F318896  Fort Pierce, FL, USA  LC375966  KX376063  LC375982  KX375835  KX361301  Divariscintilla octotentaculata Mikkelsen & Bieler, 1992  SMBL Mol2001  Fort Pierce, FL, USA  LC375967  LC375976  LC375983  LC375991  LC375999  Divariscintilla toyohiwakensis Yamashita, Haga & Lützen, 2011  SMBL Mol2002  Nakatsu, Oita, Japan  AB714788  LC375977  AB714869  AB714831  –  Divariscintilla troglodytes Mikkelsen & Bieler, 1989  SMBL Mol2003  Fort Pierce, FL, USA  LC375968  LC375978  LC375984  LC375992  LC376000  Divariscintilla yoyo Mikkelsen & Bieler, 1989  SMBL Mol2004  Fort Pierce, FL, USA  LC375969  LC375979  LC375985  LC375993  LC376001  Divariscintilla aff. yoyo  SMBL Mol2005  Fort Pierce, FL, USA  LC375970  LC375980  LC375986  LC375994  LC376002  Divariscintilla aff. maoria Powell, 1992  NMNZ M301615  Off Otago Peninsula, South Island, New Zealand  LC375971  KX376064  LC375987  LC375995  –  Ephippodontomorpha hirsuta Middelfart, 2005  AM C452337  Magnetic Island, Queensland, Australia  LC375972  KX376066  LC375988  KX375935  LC376003  Parabornia squillina Boss, 1965  FLMNH 446286  Rattle Snake Island, FL, USA  LC375973  LC375981  LC375989  LC375996  –  Phlyctaenachlamys lysiosquillina Popham, 1939  FLMNH 436851  Moorea Island, French Polynesia  LC375974  KX367605  LC375990  LC375997  KX361304  Phlyctaenachlamys sp.  FLMNH 436804  Moorea Island, French Polynesia  LC375975  KX376062  –  LC375998  KX361303  Outgroup                Lasaea adansoni (Gmelin, 1791)  GenBank  –  KC429472  KC429282  KC429124  KC429203  –  Galeommatoidea sp. 1  MNHN 16650  Off Aurora, Philippines  KX376127  KX376027  –  –  KX361300  Galeommatoidea sp. 2  MNHN 7676  Off Vella Lavella Island, Solomon Islands  KX376191  KX376057  –  –  –  Species  Specimen ID  Sampling locality  28S rRNA  16S rRNA  COI  H3  ANT  Divariscintilla luteocrinita Mikkelsen & Bieler, 1992  FMNH F318896  Fort Pierce, FL, USA  LC375966  KX376063  LC375982  KX375835  KX361301  Divariscintilla octotentaculata Mikkelsen & Bieler, 1992  SMBL Mol2001  Fort Pierce, FL, USA  LC375967  LC375976  LC375983  LC375991  LC375999  Divariscintilla toyohiwakensis Yamashita, Haga & Lützen, 2011  SMBL Mol2002  Nakatsu, Oita, Japan  AB714788  LC375977  AB714869  AB714831  –  Divariscintilla troglodytes Mikkelsen & Bieler, 1989  SMBL Mol2003  Fort Pierce, FL, USA  LC375968  LC375978  LC375984  LC375992  LC376000  Divariscintilla yoyo Mikkelsen & Bieler, 1989  SMBL Mol2004  Fort Pierce, FL, USA  LC375969  LC375979  LC375985  LC375993  LC376001  Divariscintilla aff. yoyo  SMBL Mol2005  Fort Pierce, FL, USA  LC375970  LC375980  LC375986  LC375994  LC376002  Divariscintilla aff. maoria Powell, 1992  NMNZ M301615  Off Otago Peninsula, South Island, New Zealand  LC375971  KX376064  LC375987  LC375995  –  Ephippodontomorpha hirsuta Middelfart, 2005  AM C452337  Magnetic Island, Queensland, Australia  LC375972  KX376066  LC375988  KX375935  LC376003  Parabornia squillina Boss, 1965  FLMNH 446286  Rattle Snake Island, FL, USA  LC375973  LC375981  LC375989  LC375996  –  Phlyctaenachlamys lysiosquillina Popham, 1939  FLMNH 436851  Moorea Island, French Polynesia  LC375974  KX367605  LC375990  LC375997  KX361304  Phlyctaenachlamys sp.  FLMNH 436804  Moorea Island, French Polynesia  LC375975  KX376062  –  LC375998  KX361303  Outgroup                Lasaea adansoni (Gmelin, 1791)  GenBank  –  KC429472  KC429282  KC429124  KC429203  –  Galeommatoidea sp. 1  MNHN 16650  Off Aurora, Philippines  KX376127  KX376027  –  –  KX361300  Galeommatoidea sp. 2  MNHN 7676  Off Vella Lavella Island, Solomon Islands  KX376191  KX376057  –  –  –  Abbreviations: AM, Australian Museum; FMNH, Field Museum of Natural History; NMNZ, Museum of New Zealand, Te Papa Tongarewa; MNHN, Muséum National d’Histoire Naturelle; SMBL, Seto Marine Laboratory; and FLMNH, Florida Museum of Natural History. View Large Table 1. Species used for molecular phylogenetic analyses with museum catalogue number or private specimen ID, sampling localities and GenBank accession numbers Species  Specimen ID  Sampling locality  28S rRNA  16S rRNA  COI  H3  ANT  Divariscintilla luteocrinita Mikkelsen & Bieler, 1992  FMNH F318896  Fort Pierce, FL, USA  LC375966  KX376063  LC375982  KX375835  KX361301  Divariscintilla octotentaculata Mikkelsen & Bieler, 1992  SMBL Mol2001  Fort Pierce, FL, USA  LC375967  LC375976  LC375983  LC375991  LC375999  Divariscintilla toyohiwakensis Yamashita, Haga & Lützen, 2011  SMBL Mol2002  Nakatsu, Oita, Japan  AB714788  LC375977  AB714869  AB714831  –  Divariscintilla troglodytes Mikkelsen & Bieler, 1989  SMBL Mol2003  Fort Pierce, FL, USA  LC375968  LC375978  LC375984  LC375992  LC376000  Divariscintilla yoyo Mikkelsen & Bieler, 1989  SMBL Mol2004  Fort Pierce, FL, USA  LC375969  LC375979  LC375985  LC375993  LC376001  Divariscintilla aff. yoyo  SMBL Mol2005  Fort Pierce, FL, USA  LC375970  LC375980  LC375986  LC375994  LC376002  Divariscintilla aff. maoria Powell, 1992  NMNZ M301615  Off Otago Peninsula, South Island, New Zealand  LC375971  KX376064  LC375987  LC375995  –  Ephippodontomorpha hirsuta Middelfart, 2005  AM C452337  Magnetic Island, Queensland, Australia  LC375972  KX376066  LC375988  KX375935  LC376003  Parabornia squillina Boss, 1965  FLMNH 446286  Rattle Snake Island, FL, USA  LC375973  LC375981  LC375989  LC375996  –  Phlyctaenachlamys lysiosquillina Popham, 1939  FLMNH 436851  Moorea Island, French Polynesia  LC375974  KX367605  LC375990  LC375997  KX361304  Phlyctaenachlamys sp.  FLMNH 436804  Moorea Island, French Polynesia  LC375975  KX376062  –  LC375998  KX361303  Outgroup                Lasaea adansoni (Gmelin, 1791)  GenBank  –  KC429472  KC429282  KC429124  KC429203  –  Galeommatoidea sp. 1  MNHN 16650  Off Aurora, Philippines  KX376127  KX376027  –  –  KX361300  Galeommatoidea sp. 2  MNHN 7676  Off Vella Lavella Island, Solomon Islands  KX376191  KX376057  –  –  –  Species  Specimen ID  Sampling locality  28S rRNA  16S rRNA  COI  H3  ANT  Divariscintilla luteocrinita Mikkelsen & Bieler, 1992  FMNH F318896  Fort Pierce, FL, USA  LC375966  KX376063  LC375982  KX375835  KX361301  Divariscintilla octotentaculata Mikkelsen & Bieler, 1992  SMBL Mol2001  Fort Pierce, FL, USA  LC375967  LC375976  LC375983  LC375991  LC375999  Divariscintilla toyohiwakensis Yamashita, Haga & Lützen, 2011  SMBL Mol2002  Nakatsu, Oita, Japan  AB714788  LC375977  AB714869  AB714831  –  Divariscintilla troglodytes Mikkelsen & Bieler, 1989  SMBL Mol2003  Fort Pierce, FL, USA  LC375968  LC375978  LC375984  LC375992  LC376000  Divariscintilla yoyo Mikkelsen & Bieler, 1989  SMBL Mol2004  Fort Pierce, FL, USA  LC375969  LC375979  LC375985  LC375993  LC376001  Divariscintilla aff. yoyo  SMBL Mol2005  Fort Pierce, FL, USA  LC375970  LC375980  LC375986  LC375994  LC376002  Divariscintilla aff. maoria Powell, 1992  NMNZ M301615  Off Otago Peninsula, South Island, New Zealand  LC375971  KX376064  LC375987  LC375995  –  Ephippodontomorpha hirsuta Middelfart, 2005  AM C452337  Magnetic Island, Queensland, Australia  LC375972  KX376066  LC375988  KX375935  LC376003  Parabornia squillina Boss, 1965  FLMNH 446286  Rattle Snake Island, FL, USA  LC375973  LC375981  LC375989  LC375996  –  Phlyctaenachlamys lysiosquillina Popham, 1939  FLMNH 436851  Moorea Island, French Polynesia  LC375974  KX367605  LC375990  LC375997  KX361304  Phlyctaenachlamys sp.  FLMNH 436804  Moorea Island, French Polynesia  LC375975  KX376062  –  LC375998  KX361303  Outgroup                Lasaea adansoni (Gmelin, 1791)  GenBank  –  KC429472  KC429282  KC429124  KC429203  –  Galeommatoidea sp. 1  MNHN 16650  Off Aurora, Philippines  KX376127  KX376027  –  –  KX361300  Galeommatoidea sp. 2  MNHN 7676  Off Vella Lavella Island, Solomon Islands  KX376191  KX376057  –  –  –  Abbreviations: AM, Australian Museum; FMNH, Field Museum of Natural History; NMNZ, Museum of New Zealand, Te Papa Tongarewa; MNHN, Muséum National d’Histoire Naturelle; SMBL, Seto Marine Laboratory; and FLMNH, Florida Museum of Natural History. View Large DNA extraction, PCR and sequencing Total genomic DNA was isolated from the mantle or foot tissue of each bivalve specimen, including museum specimens, with the Omega Bio-Tek E.Z.N.A. Mollusc DNA Kit (Omega Bio-Tek, Norcross, GA, USA). We sequenced fragments of 28S, 16S, COI and ANT genes. Polymerase chain reactions (PCRs) were used to amplify ~1030 bp of 28S, ~480 bp of 16S, ~690 bp of COI, ~330 bp of H3 and ~580 bp of ANT. Amplifications were performed in 12.5-μL mixtures consisting of 1.0 μL of forward and reverse primers (10 μM each; Table 2), 0.5 μL of template DNA, 6.25 μL of GoTaq Green master mix (Promega, Madison, WI, USA) and 3.75 μL of distilled water. Thermal cycling was performed with an initial denaturation of 3 min at 94 °C, followed by 30 cycles of 30 s at 94 °C, 30 s at a gene-specific annealing temperature (50–55 °C) and 2 min at 72 °C, with a final 3 min extension at 72 °C. All PCR products were directly sequenced at the University of Michigan Sequencing Core using PCR primers and internal primers (Table 2). The obtained sequences were deposited in the DDBJ/EMBL/GenBank databases with accession numbers LC375966–LC376003 (Table 1). Table 2. Information on primers used in this study Primer  Direction  Sequence 5′–3′  References  28S rRNA  PCR amplification and sequencing  D1  Forward  ACCCSCTGAAYTTAAGCAT  Colgan et al. (2003)  D3  Reverse  GACGATCGATTTGCACGTCA  Vonnemann et al. (2005)  Sequencing  D2F  Forward  CCCGTCTTGAAACACGGACCAAGG  Vonnemann et al. (2005)  C2R  Reverse  ACTCTCTCTTCAAAGTTCTTTTC  Dayrat et al. (2001)  16S rRNA  PCR amplification and sequencing  16SarL  Forward  CGCCTGTTTATCAAAAACAT  Palumbi et al. (1991)  16SbrH  Reverse  CCGGTCTGAACTCAGATCACGT  Palumbi et al. (1991)  H3  PCR amplification and sequencing  H3F  Forward  ATGGCTCGTACCAAGCAGACVGC  Colgan et al. (1998)  H3R  Reverse  ATATCCTTRGGCATRATRGTGAC  Colgan et al. (1998)  COI  PCR amplification and sequencing  LCO1490  Forward  GGTCAACAATCATAAAGATATTGG  Folmer et al. (1994)  HCO2198  Reverse  TAAACTTCAGGGTGACCAAAAAATC  Folmer et al. (1994)  ANT  PCR amplification and sequencing  ANTGF1  Forward  GCCAACTGCATTCGGTATTTCCC  Audzijonyte & Vrijenhoek (2010)  ANTR1  Reverse  TTCATCAAMGACATRAAMCCYTC  Audzijonyte & Vrijenhoek (2010)  Primer  Direction  Sequence 5′–3′  References  28S rRNA  PCR amplification and sequencing  D1  Forward  ACCCSCTGAAYTTAAGCAT  Colgan et al. (2003)  D3  Reverse  GACGATCGATTTGCACGTCA  Vonnemann et al. (2005)  Sequencing  D2F  Forward  CCCGTCTTGAAACACGGACCAAGG  Vonnemann et al. (2005)  C2R  Reverse  ACTCTCTCTTCAAAGTTCTTTTC  Dayrat et al. (2001)  16S rRNA  PCR amplification and sequencing  16SarL  Forward  CGCCTGTTTATCAAAAACAT  Palumbi et al. (1991)  16SbrH  Reverse  CCGGTCTGAACTCAGATCACGT  Palumbi et al. (1991)  H3  PCR amplification and sequencing  H3F  Forward  ATGGCTCGTACCAAGCAGACVGC  Colgan et al. (1998)  H3R  Reverse  ATATCCTTRGGCATRATRGTGAC  Colgan et al. (1998)  COI  PCR amplification and sequencing  LCO1490  Forward  GGTCAACAATCATAAAGATATTGG  Folmer et al. (1994)  HCO2198  Reverse  TAAACTTCAGGGTGACCAAAAAATC  Folmer et al. (1994)  ANT  PCR amplification and sequencing  ANTGF1  Forward  GCCAACTGCATTCGGTATTTCCC  Audzijonyte & Vrijenhoek (2010)  ANTR1  Reverse  TTCATCAAMGACATRAAMCCYTC  Audzijonyte & Vrijenhoek (2010)  View Large Table 2. Information on primers used in this study Primer  Direction  Sequence 5′–3′  References  28S rRNA  PCR amplification and sequencing  D1  Forward  ACCCSCTGAAYTTAAGCAT  Colgan et al. (2003)  D3  Reverse  GACGATCGATTTGCACGTCA  Vonnemann et al. (2005)  Sequencing  D2F  Forward  CCCGTCTTGAAACACGGACCAAGG  Vonnemann et al. (2005)  C2R  Reverse  ACTCTCTCTTCAAAGTTCTTTTC  Dayrat et al. (2001)  16S rRNA  PCR amplification and sequencing  16SarL  Forward  CGCCTGTTTATCAAAAACAT  Palumbi et al. (1991)  16SbrH  Reverse  CCGGTCTGAACTCAGATCACGT  Palumbi et al. (1991)  H3  PCR amplification and sequencing  H3F  Forward  ATGGCTCGTACCAAGCAGACVGC  Colgan et al. (1998)  H3R  Reverse  ATATCCTTRGGCATRATRGTGAC  Colgan et al. (1998)  COI  PCR amplification and sequencing  LCO1490  Forward  GGTCAACAATCATAAAGATATTGG  Folmer et al. (1994)  HCO2198  Reverse  TAAACTTCAGGGTGACCAAAAAATC  Folmer et al. (1994)  ANT  PCR amplification and sequencing  ANTGF1  Forward  GCCAACTGCATTCGGTATTTCCC  Audzijonyte & Vrijenhoek (2010)  ANTR1  Reverse  TTCATCAAMGACATRAAMCCYTC  Audzijonyte & Vrijenhoek (2010)  Primer  Direction  Sequence 5′–3′  References  28S rRNA  PCR amplification and sequencing  D1  Forward  ACCCSCTGAAYTTAAGCAT  Colgan et al. (2003)  D3  Reverse  GACGATCGATTTGCACGTCA  Vonnemann et al. (2005)  Sequencing  D2F  Forward  CCCGTCTTGAAACACGGACCAAGG  Vonnemann et al. (2005)  C2R  Reverse  ACTCTCTCTTCAAAGTTCTTTTC  Dayrat et al. (2001)  16S rRNA  PCR amplification and sequencing  16SarL  Forward  CGCCTGTTTATCAAAAACAT  Palumbi et al. (1991)  16SbrH  Reverse  CCGGTCTGAACTCAGATCACGT  Palumbi et al. (1991)  H3  PCR amplification and sequencing  H3F  Forward  ATGGCTCGTACCAAGCAGACVGC  Colgan et al. (1998)  H3R  Reverse  ATATCCTTRGGCATRATRGTGAC  Colgan et al. (1998)  COI  PCR amplification and sequencing  LCO1490  Forward  GGTCAACAATCATAAAGATATTGG  Folmer et al. (1994)  HCO2198  Reverse  TAAACTTCAGGGTGACCAAAAAATC  Folmer et al. (1994)  ANT  PCR amplification and sequencing  ANTGF1  Forward  GCCAACTGCATTCGGTATTTCCC  Audzijonyte & Vrijenhoek (2010)  ANTR1  Reverse  TTCATCAAMGACATRAAMCCYTC  Audzijonyte & Vrijenhoek (2010)  View Large Phylogenetic analyses In addition to the sequences obtained in this study, we also accessed sequence data of other galeommatoideans and outgroups from GenBank (Table 1). Sequences of the 28S and 16S genes were aligned using the Muscle program (Edgar, 2004) with default settings in the software Seaview (Galtier, Gouy & Gautier, 1996; Gouy, Guindon & Gascuel, 2010). We employed Gblocks v0.91b (Castresana, 2000; Talavera & Castresana, 2007) to eliminate the ambiguously aligned regions in the 28S and 16S genes. The sizes of 28S and 16S sequences prior to treatment with Gblocks were 1042 and 481 bp, respectively, whereas those after Gblocks treatment were 1032 and 343 bp, respectively. Phylogenetic trees were constructed using Bayesian and maximum likelihood (ML) methods. Bayesian analyses were performed using MrBayes 3.1.2 (Ronquist & Huelsenbeck, 2003) with substitution models chosen by Kakusan 4 (Tanabe, 2011). In the combined data set, substitution parameters were estimated separately for each gene partition [28S: GTR + Gamma, 16S: HKY85 + Gamma, COI: HKY85 + Gamma, GTR + Gamma, and F81 + Homogeneous (for each codon partition), H3: GTR + Gamma, K80 + Homogeneous, and JC69 + Homogeneous (for each codon partition), ANT: HKY85 + Gamma, F81 + Gamma, and JC69 + Homogeneous (for each codon partition)]. Two independent Metropolis-coupled Markov chain Monte Carlo runs were carried out simultaneously, sampling trees every 100 generations and calculating the average standard deviation of split frequencies (ASDSFs) every 1000 generations. Analyses were continued until ASDSF dropped below 0.01, at which point the two chains were considered to have achieved convergence. Because ASDSF was calculated based on the last 75% of the samples, we discarded the initial 25% of the sampled trees as burn-in. We confirmed that analyses reached stationarity well before the burn-in period by plotting the ln-likelihood of the sampled trees against generation time. ML analyses were performed using RAxML (Stamatakis, 2006) as implemented in raxmlGUI 1.31 (Silvestro & Michalak, 2012). The robustness of the ML tree was evaluated based on 1000 bootstrap replications. Datasets were partitioned by gene and the GTR + GAMMA model was implemented. RESULTS Observation of living Parabornia squillina Three individuals of P. squillina were collected from one male individual of L. scabricauda (Fig. 2). Each individual was attached by byssal threads to the host abdomen, specifically the lateral portion of the pleonal sternite (Fig. 2E, F). Two individuals were found between the 1st and 2nd pleopods, and one was found between 2nd and 3rd pleopods. We detached the bivalves from the host to observe the extension of the foot and mantle in the living state. The bivalves have numerous short papillae extended along the ventral and posterior–dorsal margins (Fig. 2A–C). One pair of longer papillae was observed anterodorsally (Fig. 2A). The clams were placed with their host in an aquarium to test if they would reattach after removal (Movie S2). They directly approached the host by crawling, and once below the host pleopods, each clam waved its foot upward towards the host (Fig. 2G). Once their foot touched the host pleon, the bivalves attached using newly secreted byssal threads. The bivalves then crawled across the host until they reached their original position on the lateral portion of the pleonal sternite. Figure 2. View largeDownload slide Parabornia squillina and its host Lysiosquilla scabricauda. A–C. A crawling individual of P. squillina (A, lateral side; B, dorsal view; C, ventral view). D. L. scabricauda. E and F. P. squillina attached to the lateral portion of the pleonal sternite. G. P. squillina extending its foot to attach to the host pleon. Arrows indicate the heel of P. squillina without posterior extension or hanging foot structure (A) and P. squillina (E–G). Figure 2. View largeDownload slide Parabornia squillina and its host Lysiosquilla scabricauda. A–C. A crawling individual of P. squillina (A, lateral side; B, dorsal view; C, ventral view). D. L. scabricauda. E and F. P. squillina attached to the lateral portion of the pleonal sternite. G. P. squillina extending its foot to attach to the host pleon. Arrows indicate the heel of P. squillina without posterior extension or hanging foot structure (A) and P. squillina (E–G). Molecular phylogenetic analyses Our results suggest that Vasconiellinae is monophyletic [Bayesian posterior probability (PP) = 1.00, bootstrap percentage (BS) = 92] (Fig. 3). Divariscintilla aff. maoria Powell, 1932 was sister to all of the remaining vasconiellines (PP = 1.00, BS = 80), including the other species of Divariscintilla, Phlyctaenachlamys, Ephippodontomorpha and Parabornia. The ectocommensal P. squillina was nested within the burrow-wall-commensal vasconiellines and was sister to D. octotentaculata (Fig. 3). Divariscintilla yoyo included one cryptic sister species (D. aff. yoyo) (Fig. 3). Floridian vasconiellines were not monophyletic; D. troglodytes was sister to a clade of Pacific and Floridian species, whereas all of the other Floridian taxa formed a crown clade that was well supported in Bayesian (PP = 0.99) but not in ML (BS = 28) phylogenetic analyses. Figure 3. View largeDownload slide Bayesian phylogenetic tree of yoyo clams (Galeommatoidea: Galeommatidae: Vasconiellinae) based on the combined data set of 28S, 16S, H3, COI and ANT genes. Numbers above branches indicate Bayesian posterior probabilities followed by maximum likelihood bootstrap support values. Six species collected from Florida are associated with Lysiosquilla scabricauda and Divariscintilla toyohiwakensis in Japan is associated with Bigelowina phalangium, whereas the other species were collected from mantis-shrimp burrows but the host species were not identified. Abbreviation: MNHN, Muséum National d’Histoire Naturelle. Figure 3. View largeDownload slide Bayesian phylogenetic tree of yoyo clams (Galeommatoidea: Galeommatidae: Vasconiellinae) based on the combined data set of 28S, 16S, H3, COI and ANT genes. Numbers above branches indicate Bayesian posterior probabilities followed by maximum likelihood bootstrap support values. Six species collected from Florida are associated with Lysiosquilla scabricauda and Divariscintilla toyohiwakensis in Japan is associated with Bigelowina phalangium, whereas the other species were collected from mantis-shrimp burrows but the host species were not identified. Abbreviation: MNHN, Muséum National d’Histoire Naturelle. DISCUSSION Within-host speciation in Floridian yoyo clams Our analysis discovered one previously unknown cryptic species (D. aff. yoyo) that is sister to D. yoyo (Fig. 3). They differed by 14.8% in their mitochondrial COI gene sequences, which is much higher than intraspecific variation levels reported for galeommatoideans [e.g. ~2% in Sato et al. (2011); ~5% in Li & Ó Foighil (2012)], or in our preliminary results for these two taxa [1.5% in D. yoyo (N = 2) and 0–0.2% in D. aff. yoyo (N = 3)] (unpublished data). They are superficially identical in external appearance but can be morphologically distinguished by their shell outlines: an angulate anterior shell margin is present in D. yoyo but not in D. aff. yoyo (our unpublished data). This means that in Florida, L. scabricauda hosts no fewer than seven vasconielline species including six burrow-wall-commensal species (Divariscintilla spp.) and one ectocommensal species (P. squillina). Our phylogenetic analyses included six Floridian vasconiellines except for D. cordiformis. Bayesian analyses suggested that Floridian vasconiellines are not monophyletic but are divided into two groups: D. troglodytes and the remaining five species (Fig. 3). The monophyly of five Floridian vasconielline species, except for D. troglodytes (Fig. 3), was supported by Bayesian posterior probabilities, suggesting that the diversity of Floridian vasconiellines is caused both by secondary contact of a distantly related linage (D. troglodytes) and by local diversification. However, bootstrap values supporting this topology are low (Fig. 3). Thus, a molecular analysis with more genetic data should be conducted in the future. Our phylogenetic analyses identified two sister-group pairs among Floridian yoyo clams: (1) D. octotentaculata and P. squillina, and (2) D. yoyo and D. aff. yoyo. In Florida, all of these species use a single host, L. scabricauda (Mikkelsen & Bieler, 1989, 1992; this study), suggesting that within-host speciation may have occurred in these two cases. Interestingly, these sister-group pairs have contrasting characteristics. Divariscintilla octotentaculata and P. squillina are ecologically and morphologically quite distinct. The former lives on host burrow walls, whereas the latter lives on the host body surface. There are differences in morphological characteristics between these two species as well, possibly corresponding to differences in host use patterns (see details below). Ecological shifts associated with host use mode probably played a key role in speciation events and led to dramatic morphological change. Divariscintilla yoyo and D. aff. yoyo, by contrast, are ecologically and morphologically very similar: both live on the host’s burrow walls and have two elongated anterior tentacle pairs (Fig. 1A, C). Lastly, an ecological shift is not apparent in this speciation event. Sympatrically distributed sister species are common among marine benthic invertebrates (Knowlton, 1993) and stem from either sympatric ecological speciation or allopatric speciation with subsequent range expansion and secondary contact (Bowen et al., 2013). These two mechanisms can be difficult to distinguish based upon existing patterns, and it is unclear whether sympatric speciation occurred in our two Floridian sister-group pairs (Fig. 3). If D. yoyo and D. aff. yoyo are not ecologically differentiated, it may be more likely that they speciated in allopatry prior to secondary contact. While Floridian Divariscintilla species have been recorded only from the Indian River Lagoon and areas nearby (Mikkelsen & Bieler, 1992; Mikkelsen, Mikkelsen & Karlen, 1995), this may be due to insufficient sampling (Mikkelsen & Bieler, 1992). Considering that L. scabricauda is distributed broadly from the Atlantic coast of the United States to Brazil (Reaka et al., 2009), allopatric speciation of yoyo clams within the host distribution range is plausible. To explore this question, further investigation of the distribution of each vasconielline species is necessary. Divariscintilla species are simultaneous hermaphrodites that brood their young to a straight-hinge ‘D’ veliger stage in the suprabranchial chamber as well as in the outer demibranch, and then release them to the water column through their exhalant siphon (Judd, 1971; Mikkelsen & Bieler, 1989, 1992; Yamashita et al., 2011). It remains unknown how long the planktotrophic larval stage endures in these species prior to metamorphosis and settlement in the host burrows. The shorter the duration of the planktonic stage for sedentary or sessile marine invertebrates, the lower the rate of gene flow among discontinuously distributed populations, and the greater the probability of allopatric speciation (but see Weersing & Toonen, 2009). Reproductive isolation mechanisms have been often considered to be necessary for the maintenance of coexistence of closely related species, because otherwise, hybridization may lead to the breakdown of species boundaries (Shine et al., 2002; Muthiga, 2003). Thus, how Floridian yoyo clams achieve reproductive isolation among co-occurring species is an intriguing question. Mikkelsen & Bieler (1992) observed an interesting copulatory-like behaviour in D. yoyo and D. octotentaculata. If this behaviour is actually copulatory in function and common in Divariscintilla species, it may allow them to engage in species-specific selective mating that can prevent interspecific hybridization. Ectocommensal lifestyle of Parabornia The genus Parabornia comprises two species, P. squillina and P. palliopapillata Simone, 2001 (Boss, 1965b; Simone, 2001). They are very similar in morphology (Simone, 2001) and both are ectocommensal on the same host, L. scabricauda (Boss, 1965b; Mikkelsen & Bieler, 1992; Simone, 2001). The former is distributed from Florida to Panama (Boss, 1965b; Moore & Boss, 1966; Mikkelsen & Bieler, 1992), whereas the latter is known only from Brazilian coasts (Simone, 2001). Previous studies briefly describe P. squillina as attached to the inner surface of the abdominal sclera of L. scabricauda (Mikkelsen & Bieler, 1992), whereas Simone (2001) mentioned that the other species, P. palliopapillata, lives attached to the pleopod base of the host. In this study, we found P. squillina attached to the lateral portion of the pleonal sternite of the host (Fig. 2). This is consistent with previous descriptions of P. squillina and P. palliopapillana, and suggests that these two species use the host in the same way. Simone (2001) mentioned that young individuals of P. palliopapillata occur on maxilipedal bases and under the carapace. This may be characteristic of P. squillina as well, although it was not confirmed in this study. During behavioural trials, P. squillina actively moved back to the lateral portion of the pleonal sternite after being detached from the host (Movie S2), suggesting that this species has a strong habitat preference for a specific part of the host abdomen. Habitat preference for a specific part of the host abdomen is common among galeommatoideans that are ectocommensal with mantis shrimp and mud shrimp (Morton, 1972; Ó Foighil, 1985; Kato & Itani, 1995), but these shared preferences in microhabitat are the result of convergent evolution (Goto et al., 2012). Gage (1968) suggested that some galeommatoideans detect hosts by using chemicals emitted from the host. We found that P. squillina can home back to a specific part of host body when it is detached from the host. Thus, it is probable that P. squillina recognizes L. scabricauda based on chemotaxis to host-emitted chemicals. Most members of Vasconiellinae, including Parabornia, have a flower-like organ near the foot, which is suggested to be a receptor of host chemicals (Mikkelsen & Bieler, 1989, 1992; Middelfart, 2005). Additionally, we found that this bivalve has numerous short papillae that occur densely along the ventral and dorsal edges of the mantle, and the former directly touch the host abdominal body surface (Fig. 2A–C). Such papillae are not known in other burrow-wall-commensal yoyo clams (Fig. 1). It is probable that these papillae have a sensory function, and that P. squillina uses them to locate its preferred position on the host body. Habitat shift from host burrow wall to host body surface Ectosymbionts that live on the body surface of burrowing invertebrates have evolved in various marine invertebrate lineages (Funch & Christensen, 1995; Kobayashi & Kato, 2003). However, the evolutionary processes that produce an ectosymbiotic lifestyle are not well understood. In this study, we show that ectocommensal Parabornia evolved from burrow-wall-commensal ancestors (Fig. 3), indicating that the burrow-wall-commensal lifestyle was an evolutionary stepping stone for an ectocommensal lifestyle in this case. Other than the genus Parabornia, ectocommensalism has evolved multiple times in Galeommatoidea (Goto et al., 2012). Evolutionary transitions from free-living to commensal lifestyles have occurred multiple times in Galeommatoidea, most of which are transitions from a free-living to burrow-wall-commensal lifestyle (Goto et al., 2012; Li et al., 2016). However, transitions from a free-living to an ectocommensal lifestyle have not previously been reported (Goto et al., 2012; Li et al., 2016), indicating that a burrow-wall-commensal lifestyle may be a prerequisite to attaining an ectocommensal lifestyle in these clams. Future in-depth phylogenetic studies of commensal galeommatoideans are likely to uncover additional cases of such evolutionary transitions. Competition for limited resources is frequently recognized as a selective pressure that promotes habitat shifts (Schluter, 2000; Munday et al., 2004; Losos, 2011; Hurt et al., 2013). However, whether resource competition has influenced the habitat shift from host burrow wall to host body surface in P. squillina is unclear, and based on the evidence to date, P. squillina and burrow-wall-commensal species do not co-occur (Mikkelsen & Bieler, 1992). While more evidence is needed, evolving an ectocommensal lifestyle may benefit P. squillina in several ways. For instance, in the case of burrow abandonment of one or both host shrimps, an ectocommensal can move to a new burrow with its host, although lysiosquillids may stay in the same burrow in monogamous pairs for up to 15 years (R. L. Caldwell, pers. comm.), making it unclear how large a factor this is in the ecology of P. squillina. Attachment to the host body may also add another level of protection. Parabornia squillina are hidden within the host’s pleopods (Fig. 2), whereas burrow-wall-commensals may be exposed to small predators accessing the burrow. Additionally, attachment to the host may provide additional food resources and a more consistent flow of oxygenated water due to the constant movement of the pleopods (Movie S2), as known in the galeommatoidean Borniopsis subsinuata ectocommensal with mantis shrimps (Morton, 1981). This positioning could be particularly useful when the burrow opening is capped during moulting or during low tides. However, there are potential disadvantages to the ectocommensal lifestyle including predation on the host, especially when outside of its burrow, being lethal to the P. squillina and the requirement for P. squillina to successfully reattach to the host after moulting events, as known in the galeommatoidean Peregrinamor ectocommensal with mud shrimps (Itani, Kato & Shirayama, 2002). Further research is required to evaluate the relative importance of these factors. Morphological changes associated with ecological shift Our results show that ectocommensal Parabornia evolved from burrow-wall-commensal ancestors (Fig. 3). The morphologies of P. squillina and Divariscintilla species have been well described in previous studies (Boss, 1965b; Mikkelsen & Bieler, 1989, 1992; Simone, 2001). By comparing their morphological characters, we detected four major morphological changes associated with the ecological shift in Parabornia: (1) loss of the thread-like posterior foot extension and associated hanging behaviour, (2) loss of covering of the shell by mantle tissue (i.e. shell externalization), (3) loss of developed sensory tentacles, and (4) acquisition of dense papillae along the mantle margin. We discuss these in detail below. Many members of Vasconiellinae, including Divariscintilla, Phlyctaenachlamys and Ephippodontomorpha, have a specialized foot with a thread-like posterior extension (Judd, 1971; Mikkelsen & Bieler, 1989, 1992; Middelfart, 2005; Yamashita et al., 2011; Fig. 1). This specialized foot, associated with the yoyo motion, is only known in Vasconiellinae and is suggested to be an adaptation to life on mantis shrimp burrow walls (Popham, 1939; Mikkelsen & Bieler, 1989). On the other hand, P. squillina has a more typical galeommatoidean foot lacking a thread-like posterior extension (Mikkelsen & Bieler, 1992; this study; Fig. 2A). Additionally, Parabornia has never been observed hanging from vertical wall surfaces or the host body in aquaria nor engaging in yoyo saltatory behaviour typical of Divariscintilla (R. Goto & T. A. Harrison, personal observations). While extreme morphological specialization of the foot has been reported in some bivalves (e.g. Dufour & Felbeck, 2003), significant morphological change in bivalve foot structure associated with a microhabitat shift is documented here for the first time. Burrow-wall-commensal yoyo clams have reduced shells fully or partially internalized by mantle tissue bearing highly developed tentacles (Popham, 1939; Judd, 1971; Mikkelsen & Bieler, 1989, 1992; Middelfart, 2005) (Fig. 1). Parabornia squillina has an externalized shell as is typical in most bivalve species (Boss, 1965b; Simone, 2001; Fig. 2A). Unlike typical bivalves, burrow-wall-commensal yoyo clams actively crawl on the burrow wall surfaces using their foot. The soft mantle and tentacles are probably useful in perceiving the surrounding environment during crawling behaviour as well as in perceiving and reacting to host movement within the burrow (Judd, 1971). Unlike burrow-wall-commensal yoyo clams, Parabornia spp. are basically sessile on the host body surface (Fig. 2E, F), probably reducing the necessity for sensory and defensive structures and hence the loss of a mantle shell covering, although P. squillina does have numerous short mantle papillae (Fig. 2), which touch the host body surface. Our results suggest that evolution from (semi)internalized to externalized shells occurred as a consequence of a change in microhabitat. Evolution of shell internalization is known in several molluscan lineages (e.g. Oposthobranchia and cephalopods) (Wägele & Klussmann-Kolb, 2005; Tanner et al., 2017). However, as far as we know, the evolution of shell externalization in molluscs has not been previously reported. The internalized shells, developed tentacles and specialized foot of burrow-wall-commensal yoyo clams are considered adaptations to the unique habitat of living on walls of mantis shrimp burrows (Popham, 1939; Mikkelsen & Bieler, 1989, 1992; this study). Our study suggests that these specialized morphological traits are lost as a consequence of colonization of the host body surface. Host shifts and subsequent specialization to different host taxa have been considered drivers of morphological evolution in Galeommatoidea (Goto et al., 2012; Li et al., 2016). Our study suggests that a microhabitat shift within a single host can also lead to significant morphological change. Goto et al. (2014) found that sister galeommatoidean species ectocommensal on Lingula brachiopods have significantly different shell shapes (elongated triangular shape vs. ovate shape) and suggested that this difference is due to adaptations for different posture on the host body. Goto et al. (2014) and the present study suggest that not only a host shift but also a ecological shift in association with the same host can play an important role in morphological evolution. However, burrow-wall-commensal yoyo clams show great morphological diversity, especially in their number of tentacles (Mikkelsen & Bieler, 1989, 1992; Fig. 1). The degree and functional significance of this tentacle diversity remains unknown. To answer this question, further examination of ecological differences among these species (e.g. niche partitioning within the host burrow) and of tentacle function is required. Taxonomic implications and remaining issues in Vasconiellinae Li et al. (2016) found that the genera Divariscintilla, Ephippodontomorpha and Phlyctaenachlamys formed a clade although their inter-relationships were not fully resolved. Our molecular analysis based on five genes showed that Ephippodontmorpha, Phlyctaenachlamys and Parabornia are nested within Divariscintilla (Fig. 3). According to Huber (2015), these genera are assigned to the same subfamily Vasconiellinae. The other four genera within this subfamily (i.e. Vasconiella, Bellascintilla, Ceratobornia and Aclistothyra) were not included in the present analysis. Ceratobornia also has hanging foot structure, but lives attached to the burrow walls of ghost shrimp on western Atlantic coasts (Dall, 1899; Narchi, 1966). Morphologically similar species with different hosts imply that host shifts have occurred in Vasconiellinae, although the hanging foot structure may have evolved multiple times in this group. Lastly, our results show that D. aff. maoria is sister to the remaining vasconiellines. Divariscintilla aff. maoria is distinguished from the other species used in this study in having a notch in the ventral side of shells. A ventral shell notch is also known in some other vasconiellines (i.e. Vasconiella, Bellascintilla and D. cordiformis) (Mikkelsen & Bieler, 1992; Huber, 2015). It is possible that Vasconiellinae may prove to be separable into two major groups discernible by the presence or absence of a notch in the ventral side of shells. To resolve these remaining issues, a further molecular analysis based on more taxon sampling is required. SUPPORTING INFORMATION Additional Supporting Information may be found in the online version of this article at the publisher’s web-site. Movie S1. Hanging behaviour of Divariscintilla yoyo. Movie S2. Parabornia squillina moving back to the abdomen of the mantis shrimp Lysiosquilla scabricauda. ACKNOWLEDGMENTS We thank Sherry Reed, Michael J. Boyle, William (Woody) Lee M.B.S. and David R. Branson (Smithsonian Marine Station at Fort Pierce) for helping to collect the specimens, Taehwan Lee (University of Michigan) for organizing the specimens used for this study, Jingchun Li (University of Colorado Boulder) for providing information on Galeommatoidea, Philippe Bouchet (Muséum National d’Histoire Naturelle, Paris), Gustav Paulay (Museum of Natural History, University of Florida) and Bruce Marshall (Museum of New Zealand, Te Papa Togarewa) for allowing us to use their museum specimens in this study, Arthur Anker (Universidade Federal de Goiás), Roy L. Caldwell (University of California, Berkeley), Maya S. deVries (University of California, San Diego) and Gyo Itani (Kochi University) for advice, and Paula M. Mikkelsen (Cornell University) and two anonymous referees for comments that improved the manuscript. This study was partially supported by an Overseas Research Fellowship grant (27-186) and a KAKENHI grant (17H06795) to R.G. from the Japan Society for the Promotion of Science and a Smithsonian Minority Awards Program grant to T.H. REFERENCES Althoff DM. 2014. Shift in egg-laying strategy to avoid plant defense leads to reproductive isolation in mutualistic and cheating yucca moths. Evolution  68: 301– 307. Google Scholar CrossRef Search ADS   Audzijonyte A, Vrijenhoek RC. 2010. Three nuclear genes for phylogenetic, SNP and population genetic studies of molluscs and other invertebrates. Molecular Ecology Resources  10: 200– 204. Google Scholar CrossRef Search ADS   Bieler R, Mikkelsen PM. 1992. Preliminary phylogenetic analysis of the bivalve family Galeommartidae. American Malacological Bulletin  9: 157– 164. Boss KJ. 1965a. Symbiotic erycinacean bivalves. Malacologia  3: 183– 195. Boss KJ. 1965b. A new mollusk (Bivalvia, Erycinidae) commensal on the stomatopod crustacean Lysiosquilla. American Museum Novitates  2215: 1– 11. Bouchet P, Lozouet P, Maestrati P, Heros V. 2002. Assessing the magnitude of species richness in tropical marine environments: exceptionally high numbers of molluscs at a New Caledonia site. Biological Journal of the Linnean Society  75: 421– 436. Google Scholar CrossRef Search ADS   Bowen BW, Rocha LA, Toonen RJ, Karl SA ; ToBo Laboratory. 2013. The origins of tropical marine biodiversity. Trends in Ecology & Evolution  28: 359– 366. Google Scholar CrossRef Search ADS   Castresana J. 2000. Selection of conserved blocks from multiple alignments for their use in phylogenetic analysis. Molecular Biology and Evolution  17: 540– 552. Google Scholar CrossRef Search ADS   Colgan DJ, McLauchlan A, Wilson GDF, Livingston S, Macaranas J, Edgecombe GD, Cassis G, Gray MR. 1998. Molecular phylogenetics of the Arthropoda: relationships based on histone H3 and U2 snRNA DNA sequences. Australian Journal of Zoology  46: 419– 437. Google Scholar CrossRef Search ADS   Colgan DJ, Ponder WF, Beacham E, Macaranas JM. 2003. Gastropod phylogeny based on six segments from four genes representing coding or non-coding and mitochondrial or nuclear DNA. Molluscan Research  23: 123– 148. Google Scholar CrossRef Search ADS   Cook JM, Rokas A, Pagel M, Stone GN. 2002. Evolutionary shifts between host oak sections and host-plant organs in Andricus gallwasps. Evolution  56: 1821– 1830. Google Scholar CrossRef Search ADS   Coyne JA, Orr HA. 2004. Speciation . Sunderland: Sinauer Associates. Dall WH. 1899. Synopsis of the recent and Tertiary Leptonacea of North America and the West Indies. Proceedings of the United States National Museum  21: 873– 897. Google Scholar CrossRef Search ADS   Dayrat B, Tillier A, Lecointre G, Tillier S. 2001. New clades of euthyneuran gastropods (Mollusca) from 28S rRNA sequences. Molecular Phylogenetics and Evolution  19: 225– 235. Google Scholar CrossRef Search ADS   Duffy JE. 1996. Resource-associated population subdivision in a symbiotic coral-reef shrimp. Evolution  50: 360– 373. Google Scholar CrossRef Search ADS   Dufour SC, Felbeck H. 2003. Sulphide mining by the superextensile foot of symbiotic thyasirid bivalves. Nature  426: 65– 67. Google Scholar CrossRef Search ADS   Edgar RC. 2004. MUSCLE: multiple sequence alignment with high accuracy and high throughput. Nucleic Acids Research  32: 1792– 1797. Google Scholar CrossRef Search ADS   Faucci A, Toonen RJ, Hadfield MG. 2007. Host shift and speciation in a coral-feeding nudibranch. Proceedings of the Royal Society B: Biological Sciences  274: 111– 119. Google Scholar CrossRef Search ADS   Folmer O, Black M, Hoeh W, Lutz R, Vrijenhoek R. 1994. DNA primers for amplification of mitochondrial cytochrome c oxidase subunit I from diverse metazoan invertebrates. Molecular Marine Biology and Biotechnology  3: 294– 299. Funch P, Christensen RM. 1995. Cycliophora is a new phylum with affinities to Entoprocta and Ectoprocta. Nature  378: 711– 714. Google Scholar CrossRef Search ADS   Gage J. 1968. The mode of life of Montacuta elevata, a bivalve ‘commensal’ with Clymenella torquata (Polychaeta). Canadian Journal of Zoology  46: 877– 892. Google Scholar CrossRef Search ADS   Galtier N, Gouy M, Gautier C. 1996. SEAVIEW and PHYLO_WIN: two graphic tools for sequence alignment and molecular phylogeny. Computer Applications in the Biosciences  12: 543– 548. Goto R, Kato M. 2012. Geographic mosaic of mutually exclusive dominance of obligate commensals in symbiotic communities associated with a burrowing echiuran worm. Marine Biology  159: 319– 330. Google Scholar CrossRef Search ADS   Goto R, Kawakita A, Ishikawa H, Hamamura Y, Kato M. 2012. Molecular phylogeny of the bivalve superfamily Galeommatoidea (Heterodonta, Veneroida) reveals dynamic evolution of symbiotic lifestyle and interphylum host switching. BMC Evolutionary Biology  12: 172. Google Scholar CrossRef Search ADS   Goto R, Ishikawa H, Hamamura Y. 2016. The enigmatic bivalve genus Paramya (Myoidea: Myidae): symbiotic association of an East Asian species with spoon worms (Echiura) and its transfer to the family Basterotiidae (Galeommatoidea). Journal of the Marine Biological Association of the United Kingdom  97: 1447– 1454. Google Scholar CrossRef Search ADS   Goto R, Ishikawa H, Hamamura Y, Sato S, Kato M. 2014. Evolution of symbiosis with Lingula (Brachiopoda) in the bivalve superfamily Galeommatoidea (Heterodonta), with description of a new species of Koreamya. Journal of Molluscan Studies  80: 148– 160 Google Scholar CrossRef Search ADS   Gouy M, Guindon S, Gascuel O. 2010. SeaView version 4: a multiplatform graphical user interface for sequence alignment and phylogenetic tree building. Molecular Biology and Evolution  27: 221– 224. Google Scholar CrossRef Search ADS   Huber M. 2015. Compendium of Bivalves 2. A full-color guide to the remaining seven families. A systematic listing of 8500 bivalve species and 10500 synonyms . Harxheim: ConchBooks. Hurt C, Silliman K, Anker A, Knowlton N. 2013. Ecological speciation in anemone-associated snapping shrimps (Alpheus armatus species complex). Molecular Ecology  22: 4532– 4548. Google Scholar CrossRef Search ADS   Imada Y, Kawakita A, Kato M. 2011. Allopatric distribution and diversification without niche shift in a bryophyte-feeding basal moth lineage (Lepidoptera: Micropterigidae). Proceedings of the Royal Society B: Biological Sciences  278: 3026– 3033. Google Scholar CrossRef Search ADS   Itani G, Kato M, Shirayama Y. 2002. Behaviour of the shrimp ectosymbionts, Peregrinamor ohshimai (Mollusca: Bivalvia) and Phyllodurus sp. (Crustacea: Isopoda) through ecdyses. Journal of Marine Biological Association of the United Kingdom  82: 69– 78. Jahner JP, Forister ML, Parchman TL, Smilanich AM, Miller JS, Wilson JS, Walla TR, Tepe EJ, Richards LA, Quijano-Abril MA, Glassmire AE, Dyer LA. 2017. Host conservatism, geography, and elevation in the evolution of a Neotropical moth radiation. Evolution  71: 2885– 2900. Google Scholar CrossRef Search ADS   Joy JB, Crespi BJ. 2007. Adaptive radiation of gall-inducing insects within a single host-plant species. Evolution  61: 784– 795. Google Scholar CrossRef Search ADS   Judd W. 1971. The structure and habits of Divariscintilla maoria Powell (Bivalvia: Galeommatidae). Proceedings of the Malacological Society of London  39: 343– 354. Kato M, Itani G. 1995. Commensalism of a bivalve, Peregrinamor ohshimai, with a thalasseinidean burrowing shrimp, Upogebia major. Journal of Marine Biological Association of the United Kingdom  75: 941– 947. Google Scholar CrossRef Search ADS   Knowlton N. 1993. Sibling species in the sea. Annual Reviews of Ecology and Systematics  24: 189– 216. Google Scholar CrossRef Search ADS   Kobayashi C, Kato M. 2003. Sex-biased ectosymbiosis of a unique cirripede, Octolamis unguisiformis sp. nov., that resembles the chelipeds of its host crab, Macrophthalmus milloti. Journal of Marine Biological Association of the United Kingdom  83: 925– 930. Google Scholar CrossRef Search ADS   Li J, Ó Foighil D. 2012. Host-specific morphologies but no host races in the commensal bivalve Nearomya rugifera. Invertebrate Biology  3: 197– 203. Google Scholar CrossRef Search ADS   Li J, Ó Foighil D, Middelfart P. 2012. The evolutionary ecology of biotic association in a megadiverse bivalve superfamily: sponsorship required for permanent residency in sediment. PLoS One  7: e42121. Google Scholar CrossRef Search ADS   Li J, Ó Foighil D, Strong E. 2016. Commensal associations and benthic habitats shape macroevolution of the bivalve clade Galeommatoidea. Proceedings of the Royal Society B: Biological Sciences  283: 20161006. Google Scholar CrossRef Search ADS   Losos JB. 2011. Lizards in an evolutionary tree: ecology and adaptive radiation of anoles . Berkeley: University of California Press. Matsubayashi KW, Ohshima I, Nosil P. 2010. Ecological speciation in phytophagous insects. Entomologia Experimentalis et Applicata  134: 1– 27. Google Scholar CrossRef Search ADS   Middelfart P. 2005. Review of Ephippodonta sensu lato (Galeommatidae: Bivalvia), with descriptions of new related genera and species from Australia. Molluscan Research  25: 129– 144. Mikkelsen PM, Bieler R. 1989. Biology and comparative anatomy of Divariscintilla yoyo and D. troglodytes, two new species of Galeommatidae (Bivalvia) from stomatopod burrows in eastern Florida. Malacologia  31: 175– 195. Mikkelsen PM, Bieler R. 1992. Biology and comparative anatomy of three new species of commensal Galeommatidae, with a possible case of mating behavior in bivalves. Malacologia  34: 1– 24. Mikkelsen PM, Mikkelsen PS, Karlen DJ. 1995. Molluscan biodiversity in the Indian River Lagoon, Florida. Bulletin of Marine Science  57: 94– 127. Moore DR, Boss KJ. 1966. Records for Parabornia squillina. The Nautilus  80: 34– 35. Moran NA. 2006. Symbiosis. Current Biology  16: R866– R871. Google Scholar CrossRef Search ADS   Morton B. 1972. Some aspects of the functional morphology and biology of Pseudopythina subsinuata (Bivalvia; Leptonacea) commensal on stomatopod crustaceans. Journal of Zoology, London  166: 79– 96. Google Scholar CrossRef Search ADS   Morton B. 1981. The biology and functional morphology of Chlamydoconcha orcutii with a discussion on the taxonomic status of the Chlamydoconchacea (Mollusca: Bivalvia). Journal of Zoology, London  195: 81– 121. Google Scholar CrossRef Search ADS   Morton B, Scott PH. 1989. Hong Kong Galeommatacea (Mollusca: Bivalvia) and their hosts, with descriptions of new species. Asian Marine Biology  6: 129– 160. Muthiga NA. 2003. Coexistence and reproductive isolation of the sympatric echinoids Diadema savignyi Michelin and Diadema setosum (Leske) on Kenyan coral reefs. Marine Biology  143: 669– 677. Google Scholar CrossRef Search ADS   Munday PL, van Herwerden L, Dudgeon CL. 2004. Evidence for sympatric speciation by host shift in the sea. Current Biology  14: 1498– 1504. Google Scholar CrossRef Search ADS   Nakadai R, Kawakita A. 2016. Phylogenetic test of speciation by host shift in leaf cone moths (Caloptilia) feeding on maples (Acer). Ecology and Evolution  6: 4958– 4970. Google Scholar CrossRef Search ADS   Narchi W. 1966. The functional morphology of Ceratobornia cema, new species of the Erycinacea (Mollusca, Eulamellibranchiata). Amaos da Academia Brasileira de Ciências  38: 513– 524. Ó Foighil D. 1985. Form, function, and origin of temporary dwarf males in Pseudopythina rugifera (Carpenter, 1864) (Bivalvia: Galeommatacea). The Veliger  27: 245– 252. Palumbi S, Martin A, Romano S, McMillan WO, Stice L, Grabowski G. 1991. The simple fool’s guide to PCR, Version 2.0 . Honolulu: Department of Zoology and Kewalo Marine Laboratory, University of Hawaii. Paulay G. 2003. Marine Bivalvia (Mollusca) of Guam. Micronesica  35–36: 218– 243. Pérez-Tris J, Hellgren O, Krizanauskiene A, Waldenström J, Secondi J, Bonneaud C, Fjeldså J, Hasselquist D, Bensch S. 2007. Within-host speciation of malaria parasites. PLoS One  2: e235. Google Scholar CrossRef Search ADS   Popham ML. 1939. On Phlyctaenachlamys lysiosquillina gen. and sp. nov., a lamellibranch commensal in the burrows of Lysiosquilla maculata. Great Barrier Reef Expedition 1928–1929. Scientific Reports  6: 61– 84. Poulin R, Morand S. 2004. Parasite biodiversity . Washington: Smithsonian Institution Press. Reaka ML, Camp DK, Álvarez F, Gracia AG, Ortiz M, Vázquez-Bader AR. 2009. Stomatopoda (Crustacea) of the Gulf of Mexico. In: Felder DL, Camp DK, eds. Gulf of Mexico–origins, waters, and biota: biodiversity . College Station: Texas A&M University Press, 901– 921. Ronquist F, Huelsenbeck JP. 2003. MrBayes 3: Bayesian phylogenetic inference under mixed models. Bioinformatics  19: 1572– 1574. Google Scholar CrossRef Search ADS   Sato S, Owada M, Haga T, Hong J-S, Lützen J, Yamashita H. 2011. Genus-specific commensalism of the galeommatoid bivalve Koreamya arcuata (A. Adams, 1856) associated with lingulid brachiopods. Molluscan Research  31: 95– 105. Schluter D. 2000. The ecology of adaptive radiation . Oxford: Oxford University Press. Shine R, Reed RN, Shetty S, Lemaster M, Mason RT. 2002. Reproductive isolating mechanisms between two sympatric sibling species of sea snakes. Evolution  56: 1655– 1662. Google Scholar CrossRef Search ADS   Simone LRL. 2001. Revision of the genus Parabornia (Bivalvia: Galeommatoidea: Galeommatidae) from the western Atlantic, with description of a new species from Brazil. Journal of Conchology  37: 159– 169. Silvestro D, Michalak I. 2012. RaxmlGUI: a graphical front-end for RAxML. Organism Diversity and Evolution  12: 335– 337. Google Scholar CrossRef Search ADS   Stamatakis A. 2006. RAxML-VI-HPC: maximum likelihood-based phylogenetic analyses with thousands of taxa and mixed models. Bioinformatics  22: 2688– 2690. Google Scholar CrossRef Search ADS   Talavera G, Castresana J. 2007. Improvement of phylogenies after removing divergent and ambiguously aligned blocks from protein sequence alignments. Systematic Biology  56: 564– 577. Google Scholar CrossRef Search ADS   Tanabe AS. 2011. Kakusan4 and Aminosan: two programs for comparing nonpartitioned, proportional and separate models for combined molecular phylogenetic analyses of multilocus sequence data. Molecular Ecology Resources  11: 914– 921. Google Scholar CrossRef Search ADS   Tanner AR, Fuchs D, Winkelmann IE, Gilbert MTP, Pankey MS, Ribeiro ÂM, Kocot KM, Halanych KM, Oakley TH, da Fonseca RR, Pisani D, Vinther J. 2017. Molecular clocks indicate turnover and diversification of modern coleoid cephalopods during the Mesozoic Marine Revolution. Proceedings of the Royal Society B: Biological Sciences  284: 20162818. Google Scholar CrossRef Search ADS   Tsang LM, Chan BKK, Shin F-L, Chu KH, Chen CA. 2009. Host-associated speciation in the coral barnacle Wanella milleporae (Cirripedia: Pyrgomatidae) inhabiting the Millepora coral. Molecular Ecology  18: 1463– 1475. Google Scholar CrossRef Search ADS   Vanhove MPM, Hablützel PI, Pariselle A, Šimková A, Huyse T, Raeymaekers JAM. 2016. Cichlids: a host of opportunities for evolutionary parasitology. Trends in Parasitology  32: 821– 832. Google Scholar CrossRef Search ADS   Vonnemann V, Schrödl M, Klussmann-Kolb A, Wägele H. 2005. Reconstruction of the phylogeny of the Opisthobranchia (Mollusca: Gastropoda) by means of 18S and 28S rRNA gene sequences. Journal of Molluscan Studies  71: 113– 125. Google Scholar CrossRef Search ADS   Wägele H, Klussmann-Kolb A. 2005. Opisthobranchia (Mollusca, Gastropoda) – more than just slimy slugs. Shell reduction and its implications on defence and foraging. Frontiers in Zoology  2: 3. Google Scholar CrossRef Search ADS   Weersing K, Toonen RJ. 2009. Population genetics, larval dispersal, and connectivity in marine systems. Marine Ecology Progress Series  393: 1– 12. Google Scholar CrossRef Search ADS   Windsor DA. 1998. Most of the species on Earth are parasites. International Journal for Parasitology  28: 1939– 1941. Google Scholar CrossRef Search ADS   Winkler IS, Mitter C. 2008. The phylogenetic dimension of insect–plant interactions: a review of recent evidence. In: Tilmon KJ, ed. Specialization, speciation, and radiation: the evolutionary biology of herbivorous insects . Berkeley: University of California Press, 203– 215. Google Scholar CrossRef Search ADS   Yamashita H, Haga T, Lützen J. 2011. The bivalve Divariscintilla toyohiwakensis n. sp. (Heterodonta: Galeommatidae) from Japan, a commensal with a mantis shrimp. Venus  69: 123– 133. Zhang B, Segraves KA, Xue H-J, Nie R-E, Li W-Z, Yang X-K. 2015. Adaptation to different host plant ages facilitates insect divergence without a host shift. Proceedings of the Royal Society B: Biological Sciences  282: 20151649. Google Scholar CrossRef Search ADS   © 2018 The Linnean Society of London, Biological Journal of the Linnean Society This article is published and distributed under the terms of the Oxford University Press, Standard Journals Publication Model (https://academic.oup.com/journals/pages/about_us/legal/notices) http://www.deepdyve.com/assets/images/DeepDyve-Logo-lg.png Biological Journal of the Linnean Society Oxford University Press

Within-host speciation events in yoyo clams, obligate commensals with mantis shrimps, including one that involves a change in microhabitat and a loss of specialized traits

Loading next page...
 
/lp/ou_press/within-host-speciation-events-in-yoyo-clams-obligate-commensals-with-cjAakhQD3l
Publisher
The Linnean Society of London
Copyright
© 2018 The Linnean Society of London, Biological Journal of the Linnean Society
ISSN
0024-4066
eISSN
1095-8312
D.O.I.
10.1093/biolinnean/bly044
Publisher site
See Article on Publisher Site

Abstract

Abstract Compared to host shifts, the importance of within-host cladogenesis in the diversification of symbionts remains less well understood in marine systems. Yoyo clams (Galeommatidae: Vasconiellinae) are a clade of marine bivalves that live commensally with burrowing mantis shrimp. Almost all yoyo clams byssally-attach to the host burrow wall via a specialized hanging foot structure bearing a thread-like posterior extension. In contrast, Parabornia squillina (Vasconiellinae) byssally-attaches directly to the host shrimp and lacks a hanging foot structure. In this study, we examine phylogenetic relationships among vasconiellines by performing molecular analyses based on five genes (28S and 16S rRNA, H3, COI and ANT). We found evidence for two within-host speciation events among Floridian vasconiellines commensal with the same mantis shrimp host, Lysiosquilla scabricauda. One involved a cryptic sister species pair of burrow-wall commensals. The other involved the ectocommensal P. squillina and its somewhat unexpected sister taxon, the burrow-wall commensal Divariscintilla octotentaculata. This latter result suggests that a habitat shift from host burrow wall to host body surface occurred while retaining the same host species and led to the loss of the specialized hanging foot structure. Our findings suggest that ostensibly modest within-host ecological shifts can lead to major morphological changes in these clams. adaptation, burrow, commensalism, Galeommatoidea, habitat shift, host shift, specialization, Stomatopoda, symbiosis INTRODUCTION Symbiotic and parasitic organisms represent a large fraction of the Earth’s biodiversity (Windsor, 1998; Poulin & Morand, 2004; Moran, 2006). Host shifting, an evolutionary change in host species, has been recognized as the major driver of speciation in these organisms both in terrestrial (Coyne & Orr, 2004; Matsubayashi, Ohshima & Nosil, 2010) and in marine realms (Duffy, 1996; Munday, van Herwerden & Dudgeon, 2004; Faucci, Toonen & Hadfield, 2007; Tsang et al., 2009; Goto et al., 2012; Hurt et al., 2013). However, the role of host shifts in diversification may be less important than previously thought (Winkler & Mitter, 2008; Imada, Kawakita & Kato, 2011; Nakadai & Kawakita, 2016), and alternative speciation processes, such as allopatric speciation and within-host speciation, can also play an important role in the diversification of parasites and symbionts (Imada et al., 2011; Nakadai & Kawakita, 2016; Jahner et al., 2017). Within-host speciation driven by ecological shifts has been studied mainly in phytophagus insects (e.g. gall-inducing insects) (Cook et al., 2002; Joy & Crespi, 2007). In these systems, speciation is probably initiated by adaptations to different host tissues (Cook et al., 2002; Joy & Crespi, 2007; Althoff, 2014) or different life history stages (Zhang et al., 2015). Similar speciation patterns have also been reported in other systems, such as avian malaria (Pérez-Tris et al., 2007) and freshwater fish parasites (Vanhove et al., 2016). However, evidence for within-host speciation driven by ecological shifts remains limited in marine systems. The superfamily Galeommatoidea is a group of small-bodied bivalves that exhibit high species diversity in shallow-water environments (Bouchet et al., 2002; Paulay, 2003). Many galeommatoidean species are commensal with benthic invertebrates in soft sediments (Boss, 1965a; Morton & Scott, 1989; Goto et al., 2012; Li, Ó Foighil & Middelfart, 2012). Galeommatoideans associate with various animal phyla (e.g. Arthropoda, Echinodermata and Annelida) (Boss, 1965a; Morton & Scott, 1989), with many species demonstrating high fidelity to a particular host species or genus (Sato et al., 2011), with some exceptions (Li & Ó Foighil, 2012). Although recent molecular phylogenies suggest that host shifts between distantly related taxa occurred frequently in some clades of Galeommatoidea (Goto et al., 2012; Li, Ó Foighil & Strong, 2016), some congeneric galeommatoidean species share the same host (Mikkelsen & Bieler, 1989, 1992; Goto & Kato, 2012; Goto et al., 2014; Goto, Ishikawa & Hamamura, 2016), suggesting the possibility of within-host speciation in these lineages. The subfamily Vasconiellinae is a group of galeommatoideans that includes seven genera (Huber, 2015). Most species have reduced shells covered by hypertrophied mantles with highly developed sensory tentacles (Popham, 1939; Mikkelsen & Bieler, 1989, 1992; Fig. 1). Among them, four genera (Divariscintilla, Phlyctaenachlamys, Parabornia and Ephippodontomorpha) are known as symbionts of burrowing mantis shrimp (Stomatopoda: Lysiosquillidae) (Boss, 1965b; Judd, 1971; Mikkelsen & Bieler, 1989, 1992; Simone, 2001; Middelfart, 2005; Yamashita, Haga & Lützen, 2011). Except for Parabornia species, these vasconiellines live suspended from the burrow walls of their stomatopod hosts by means of a specialized hanging foot structure having a thread-like posterior extension (Judd, 1971; Mikkelsen & Bieler, 1989, 1992; Middelfart, 2005; Yamashita et al., 2011; Fig. 1). In association with this posture, these clams engage in a characteristic ‘yo-yo’ up and down motion by contracting and relaxing the posterior foot (Mikkelsen & Bieler, 1989, 1992; Fig. 1B; see Supporting Information, Movie S1), hence the informal ‘yoyo clam’ name (Mikkelsen & Bieler, 1989). Figure 1. View largeDownload slide Diversity of Floridian yoyo clams (Galeommatoidea: Galeommatidae: Vasconiellinae: Divariscintilla) collected from the burrow of Lysiosquilla scabricauda. A. Divariscintilla yoyo. B. Hanging behaviour of D. yoyo. C. D. aff. yoyo. D. D. troglodytes. E. D. octotentaculata. F. D. luteocrinita. Arrows indicate posterior foot extension (hanging foot structure). Figure 1. View largeDownload slide Diversity of Floridian yoyo clams (Galeommatoidea: Galeommatidae: Vasconiellinae: Divariscintilla) collected from the burrow of Lysiosquilla scabricauda. A. Divariscintilla yoyo. B. Hanging behaviour of D. yoyo. C. D. aff. yoyo. D. D. troglodytes. E. D. octotentaculata. F. D. luteocrinita. Arrows indicate posterior foot extension (hanging foot structure). Divariscintilla includes seven described species that have been recorded from Florida, New Zealand and Japan (Judd, 1971; Mikkelsen & Bieler, 1989, 1992; Yamashita et al., 2011). Interestingly, five species (D. yoyo Mikkelsen & Bieler, 1989, D. troglodytes Mikkelsen & Bieler, 1989, D. octotentaculata Mikkelsen & Bieler, 1992, D. luteocrinita Mikkelsen & Bieler, 1992 and D. cordiformis Mikkelsen & Bieler, 1992) are at present known only from south-eastern Florida (Mikkelsen & Bieler, 1989, 1992), living exclusively in the burrows of a single stomatopod host, Lysiosquilla scabricauda (Lamarck, 1818) (Mikkelsen & Bieler, 1989, 1992). In addition to Divariscintilla, the ectocommensal vasconielline Parabornia squillina Boss, 1965 also utilizes L. scabricauda as a host in Florida (Boss, 1965b; Mikkelsen & Bieler, 1992), although this species has also been recorded from Mississippi (Boss, 1965b) and Panama (Moore & Boss, 1966). Unlike Divariscintilla spp., Parabornia spp. live attached to the host body surface (Boss, 1965b). Taken together, L. scabricauda hosts six vasconielline species in eastern Florida, thereby providing an invaluable opportunity to investigate the possibility of within-host speciation. Burrow-wall-commensal Divariscintilla spp. and ectocommensal Parabornia spp. are thought to be closely related because many possess flower-like organs near the base of the foot in addition to morphological similarity in the posterior foot structure (Bieler & Mikkelsen, 1992; Mikkelsen & Bieler, 1992), although the posterior foot extension of the genus Parabornia is much shorter than that of the genus Divariscintilla (Mikkelsen & Bieler, 1992). Unlike the genus Parabornia, the shells of Divariscintilla spp. are partially to fully covered by mantle tissue that bears highly developed sensory tentacles (Boss, 1965b; Mikkelsen & Bieler, 1989, 1992; Simone, 2001; Fig. 1). These differences in mantle coverage and foot structure are thought to reflect the differences in host utilization between the two genera (burrow-wall-commensals vs. ectocommensals). However, the phylogenetic relationship of these genera remains unexamined. In this study, we addressed the following questions: (1) are the six vasconiellines associated with L. scabricauda in Florida monophyletic, and if so, (2) how have evolutionary transitions between burrow-wall-commensal and ectocommensal lifestyles occurred in this bivalve clade? We performed molecular analyses of Vasconiellinae based on two nuclear genes (28S rRNA and histone H3) and three mitochondrial genes [cytochrome c oxidase subunit I (COI), 16S rRNA and adenine nucleotide translocator (ANT)]. Because the ectocommensal lifestyle of Parabornia was only briefly mentioned in previous studies (Boss, 1965b; Mikkelsen & Bieler, 1992; Simone, 2001), we observed living P. squillina to further understand its ecological adaptations to an ectocommensal lifestyle. Lastly, morphological characteristics of Divariscintilla and Parabornia were compared to reveal if morphological differences between genera are associated with ecological shifts. MATERIAL AND METHODS Sample collection and observations Sampling was performed in intertidal sand flats in the Indian River lagoon (Fort Pierce, FL, USA), the type locality of the five Divariscintilla species (D. yoyo, D. troglodytes, D. octotentaculata, D. luteocrinita and D. cordiformis) (Mikkelsen & Bieler, 1989, 1992) during 30 May–4 June 2016 and 26–31 January 2017. We collected Divariscintilla species from L. scabricauda burrows using stainless steel bait pumps (‘yabby pumps’) and 1–2-mm mesh sieves. With the exception of D. cordiformis, a very rare species at this site (Mikkelsen & Bieler, 1992), all known Floridian Divariscintilla species were collected. We also collected P. squillina from the ventral body surface of it host L. scabricauda, which were captured manually using fish bait. The bivalves were kept for several days in aquaria for observations and then preserved in 100% ethanol for DNA analyses. Additionally, alcohol-fixed museum specimens of Divariscintilla spp. and close relatives were loaned from the Muséum National d’Histoire Naturelle, Paris, Field Museum, Florida Museum of Natural History and Museum of New Zealand, Te Papa Togarewa for DNA analyses (Table 1). The DNA sequences of Divariscintilla and closely related species used in previous phylogenetic studies were obtained from GenBank (Table 1). For outgroups, we used several galeommatoideans that were identified to be closely related to Vasconiellinae by Li et al. (2016). Table 1. Species used for molecular phylogenetic analyses with museum catalogue number or private specimen ID, sampling localities and GenBank accession numbers Species  Specimen ID  Sampling locality  28S rRNA  16S rRNA  COI  H3  ANT  Divariscintilla luteocrinita Mikkelsen & Bieler, 1992  FMNH F318896  Fort Pierce, FL, USA  LC375966  KX376063  LC375982  KX375835  KX361301  Divariscintilla octotentaculata Mikkelsen & Bieler, 1992  SMBL Mol2001  Fort Pierce, FL, USA  LC375967  LC375976  LC375983  LC375991  LC375999  Divariscintilla toyohiwakensis Yamashita, Haga & Lützen, 2011  SMBL Mol2002  Nakatsu, Oita, Japan  AB714788  LC375977  AB714869  AB714831  –  Divariscintilla troglodytes Mikkelsen & Bieler, 1989  SMBL Mol2003  Fort Pierce, FL, USA  LC375968  LC375978  LC375984  LC375992  LC376000  Divariscintilla yoyo Mikkelsen & Bieler, 1989  SMBL Mol2004  Fort Pierce, FL, USA  LC375969  LC375979  LC375985  LC375993  LC376001  Divariscintilla aff. yoyo  SMBL Mol2005  Fort Pierce, FL, USA  LC375970  LC375980  LC375986  LC375994  LC376002  Divariscintilla aff. maoria Powell, 1992  NMNZ M301615  Off Otago Peninsula, South Island, New Zealand  LC375971  KX376064  LC375987  LC375995  –  Ephippodontomorpha hirsuta Middelfart, 2005  AM C452337  Magnetic Island, Queensland, Australia  LC375972  KX376066  LC375988  KX375935  LC376003  Parabornia squillina Boss, 1965  FLMNH 446286  Rattle Snake Island, FL, USA  LC375973  LC375981  LC375989  LC375996  –  Phlyctaenachlamys lysiosquillina Popham, 1939  FLMNH 436851  Moorea Island, French Polynesia  LC375974  KX367605  LC375990  LC375997  KX361304  Phlyctaenachlamys sp.  FLMNH 436804  Moorea Island, French Polynesia  LC375975  KX376062  –  LC375998  KX361303  Outgroup                Lasaea adansoni (Gmelin, 1791)  GenBank  –  KC429472  KC429282  KC429124  KC429203  –  Galeommatoidea sp. 1  MNHN 16650  Off Aurora, Philippines  KX376127  KX376027  –  –  KX361300  Galeommatoidea sp. 2  MNHN 7676  Off Vella Lavella Island, Solomon Islands  KX376191  KX376057  –  –  –  Species  Specimen ID  Sampling locality  28S rRNA  16S rRNA  COI  H3  ANT  Divariscintilla luteocrinita Mikkelsen & Bieler, 1992  FMNH F318896  Fort Pierce, FL, USA  LC375966  KX376063  LC375982  KX375835  KX361301  Divariscintilla octotentaculata Mikkelsen & Bieler, 1992  SMBL Mol2001  Fort Pierce, FL, USA  LC375967  LC375976  LC375983  LC375991  LC375999  Divariscintilla toyohiwakensis Yamashita, Haga & Lützen, 2011  SMBL Mol2002  Nakatsu, Oita, Japan  AB714788  LC375977  AB714869  AB714831  –  Divariscintilla troglodytes Mikkelsen & Bieler, 1989  SMBL Mol2003  Fort Pierce, FL, USA  LC375968  LC375978  LC375984  LC375992  LC376000  Divariscintilla yoyo Mikkelsen & Bieler, 1989  SMBL Mol2004  Fort Pierce, FL, USA  LC375969  LC375979  LC375985  LC375993  LC376001  Divariscintilla aff. yoyo  SMBL Mol2005  Fort Pierce, FL, USA  LC375970  LC375980  LC375986  LC375994  LC376002  Divariscintilla aff. maoria Powell, 1992  NMNZ M301615  Off Otago Peninsula, South Island, New Zealand  LC375971  KX376064  LC375987  LC375995  –  Ephippodontomorpha hirsuta Middelfart, 2005  AM C452337  Magnetic Island, Queensland, Australia  LC375972  KX376066  LC375988  KX375935  LC376003  Parabornia squillina Boss, 1965  FLMNH 446286  Rattle Snake Island, FL, USA  LC375973  LC375981  LC375989  LC375996  –  Phlyctaenachlamys lysiosquillina Popham, 1939  FLMNH 436851  Moorea Island, French Polynesia  LC375974  KX367605  LC375990  LC375997  KX361304  Phlyctaenachlamys sp.  FLMNH 436804  Moorea Island, French Polynesia  LC375975  KX376062  –  LC375998  KX361303  Outgroup                Lasaea adansoni (Gmelin, 1791)  GenBank  –  KC429472  KC429282  KC429124  KC429203  –  Galeommatoidea sp. 1  MNHN 16650  Off Aurora, Philippines  KX376127  KX376027  –  –  KX361300  Galeommatoidea sp. 2  MNHN 7676  Off Vella Lavella Island, Solomon Islands  KX376191  KX376057  –  –  –  Abbreviations: AM, Australian Museum; FMNH, Field Museum of Natural History; NMNZ, Museum of New Zealand, Te Papa Tongarewa; MNHN, Muséum National d’Histoire Naturelle; SMBL, Seto Marine Laboratory; and FLMNH, Florida Museum of Natural History. View Large Table 1. Species used for molecular phylogenetic analyses with museum catalogue number or private specimen ID, sampling localities and GenBank accession numbers Species  Specimen ID  Sampling locality  28S rRNA  16S rRNA  COI  H3  ANT  Divariscintilla luteocrinita Mikkelsen & Bieler, 1992  FMNH F318896  Fort Pierce, FL, USA  LC375966  KX376063  LC375982  KX375835  KX361301  Divariscintilla octotentaculata Mikkelsen & Bieler, 1992  SMBL Mol2001  Fort Pierce, FL, USA  LC375967  LC375976  LC375983  LC375991  LC375999  Divariscintilla toyohiwakensis Yamashita, Haga & Lützen, 2011  SMBL Mol2002  Nakatsu, Oita, Japan  AB714788  LC375977  AB714869  AB714831  –  Divariscintilla troglodytes Mikkelsen & Bieler, 1989  SMBL Mol2003  Fort Pierce, FL, USA  LC375968  LC375978  LC375984  LC375992  LC376000  Divariscintilla yoyo Mikkelsen & Bieler, 1989  SMBL Mol2004  Fort Pierce, FL, USA  LC375969  LC375979  LC375985  LC375993  LC376001  Divariscintilla aff. yoyo  SMBL Mol2005  Fort Pierce, FL, USA  LC375970  LC375980  LC375986  LC375994  LC376002  Divariscintilla aff. maoria Powell, 1992  NMNZ M301615  Off Otago Peninsula, South Island, New Zealand  LC375971  KX376064  LC375987  LC375995  –  Ephippodontomorpha hirsuta Middelfart, 2005  AM C452337  Magnetic Island, Queensland, Australia  LC375972  KX376066  LC375988  KX375935  LC376003  Parabornia squillina Boss, 1965  FLMNH 446286  Rattle Snake Island, FL, USA  LC375973  LC375981  LC375989  LC375996  –  Phlyctaenachlamys lysiosquillina Popham, 1939  FLMNH 436851  Moorea Island, French Polynesia  LC375974  KX367605  LC375990  LC375997  KX361304  Phlyctaenachlamys sp.  FLMNH 436804  Moorea Island, French Polynesia  LC375975  KX376062  –  LC375998  KX361303  Outgroup                Lasaea adansoni (Gmelin, 1791)  GenBank  –  KC429472  KC429282  KC429124  KC429203  –  Galeommatoidea sp. 1  MNHN 16650  Off Aurora, Philippines  KX376127  KX376027  –  –  KX361300  Galeommatoidea sp. 2  MNHN 7676  Off Vella Lavella Island, Solomon Islands  KX376191  KX376057  –  –  –  Species  Specimen ID  Sampling locality  28S rRNA  16S rRNA  COI  H3  ANT  Divariscintilla luteocrinita Mikkelsen & Bieler, 1992  FMNH F318896  Fort Pierce, FL, USA  LC375966  KX376063  LC375982  KX375835  KX361301  Divariscintilla octotentaculata Mikkelsen & Bieler, 1992  SMBL Mol2001  Fort Pierce, FL, USA  LC375967  LC375976  LC375983  LC375991  LC375999  Divariscintilla toyohiwakensis Yamashita, Haga & Lützen, 2011  SMBL Mol2002  Nakatsu, Oita, Japan  AB714788  LC375977  AB714869  AB714831  –  Divariscintilla troglodytes Mikkelsen & Bieler, 1989  SMBL Mol2003  Fort Pierce, FL, USA  LC375968  LC375978  LC375984  LC375992  LC376000  Divariscintilla yoyo Mikkelsen & Bieler, 1989  SMBL Mol2004  Fort Pierce, FL, USA  LC375969  LC375979  LC375985  LC375993  LC376001  Divariscintilla aff. yoyo  SMBL Mol2005  Fort Pierce, FL, USA  LC375970  LC375980  LC375986  LC375994  LC376002  Divariscintilla aff. maoria Powell, 1992  NMNZ M301615  Off Otago Peninsula, South Island, New Zealand  LC375971  KX376064  LC375987  LC375995  –  Ephippodontomorpha hirsuta Middelfart, 2005  AM C452337  Magnetic Island, Queensland, Australia  LC375972  KX376066  LC375988  KX375935  LC376003  Parabornia squillina Boss, 1965  FLMNH 446286  Rattle Snake Island, FL, USA  LC375973  LC375981  LC375989  LC375996  –  Phlyctaenachlamys lysiosquillina Popham, 1939  FLMNH 436851  Moorea Island, French Polynesia  LC375974  KX367605  LC375990  LC375997  KX361304  Phlyctaenachlamys sp.  FLMNH 436804  Moorea Island, French Polynesia  LC375975  KX376062  –  LC375998  KX361303  Outgroup                Lasaea adansoni (Gmelin, 1791)  GenBank  –  KC429472  KC429282  KC429124  KC429203  –  Galeommatoidea sp. 1  MNHN 16650  Off Aurora, Philippines  KX376127  KX376027  –  –  KX361300  Galeommatoidea sp. 2  MNHN 7676  Off Vella Lavella Island, Solomon Islands  KX376191  KX376057  –  –  –  Abbreviations: AM, Australian Museum; FMNH, Field Museum of Natural History; NMNZ, Museum of New Zealand, Te Papa Tongarewa; MNHN, Muséum National d’Histoire Naturelle; SMBL, Seto Marine Laboratory; and FLMNH, Florida Museum of Natural History. View Large DNA extraction, PCR and sequencing Total genomic DNA was isolated from the mantle or foot tissue of each bivalve specimen, including museum specimens, with the Omega Bio-Tek E.Z.N.A. Mollusc DNA Kit (Omega Bio-Tek, Norcross, GA, USA). We sequenced fragments of 28S, 16S, COI and ANT genes. Polymerase chain reactions (PCRs) were used to amplify ~1030 bp of 28S, ~480 bp of 16S, ~690 bp of COI, ~330 bp of H3 and ~580 bp of ANT. Amplifications were performed in 12.5-μL mixtures consisting of 1.0 μL of forward and reverse primers (10 μM each; Table 2), 0.5 μL of template DNA, 6.25 μL of GoTaq Green master mix (Promega, Madison, WI, USA) and 3.75 μL of distilled water. Thermal cycling was performed with an initial denaturation of 3 min at 94 °C, followed by 30 cycles of 30 s at 94 °C, 30 s at a gene-specific annealing temperature (50–55 °C) and 2 min at 72 °C, with a final 3 min extension at 72 °C. All PCR products were directly sequenced at the University of Michigan Sequencing Core using PCR primers and internal primers (Table 2). The obtained sequences were deposited in the DDBJ/EMBL/GenBank databases with accession numbers LC375966–LC376003 (Table 1). Table 2. Information on primers used in this study Primer  Direction  Sequence 5′–3′  References  28S rRNA  PCR amplification and sequencing  D1  Forward  ACCCSCTGAAYTTAAGCAT  Colgan et al. (2003)  D3  Reverse  GACGATCGATTTGCACGTCA  Vonnemann et al. (2005)  Sequencing  D2F  Forward  CCCGTCTTGAAACACGGACCAAGG  Vonnemann et al. (2005)  C2R  Reverse  ACTCTCTCTTCAAAGTTCTTTTC  Dayrat et al. (2001)  16S rRNA  PCR amplification and sequencing  16SarL  Forward  CGCCTGTTTATCAAAAACAT  Palumbi et al. (1991)  16SbrH  Reverse  CCGGTCTGAACTCAGATCACGT  Palumbi et al. (1991)  H3  PCR amplification and sequencing  H3F  Forward  ATGGCTCGTACCAAGCAGACVGC  Colgan et al. (1998)  H3R  Reverse  ATATCCTTRGGCATRATRGTGAC  Colgan et al. (1998)  COI  PCR amplification and sequencing  LCO1490  Forward  GGTCAACAATCATAAAGATATTGG  Folmer et al. (1994)  HCO2198  Reverse  TAAACTTCAGGGTGACCAAAAAATC  Folmer et al. (1994)  ANT  PCR amplification and sequencing  ANTGF1  Forward  GCCAACTGCATTCGGTATTTCCC  Audzijonyte & Vrijenhoek (2010)  ANTR1  Reverse  TTCATCAAMGACATRAAMCCYTC  Audzijonyte & Vrijenhoek (2010)  Primer  Direction  Sequence 5′–3′  References  28S rRNA  PCR amplification and sequencing  D1  Forward  ACCCSCTGAAYTTAAGCAT  Colgan et al. (2003)  D3  Reverse  GACGATCGATTTGCACGTCA  Vonnemann et al. (2005)  Sequencing  D2F  Forward  CCCGTCTTGAAACACGGACCAAGG  Vonnemann et al. (2005)  C2R  Reverse  ACTCTCTCTTCAAAGTTCTTTTC  Dayrat et al. (2001)  16S rRNA  PCR amplification and sequencing  16SarL  Forward  CGCCTGTTTATCAAAAACAT  Palumbi et al. (1991)  16SbrH  Reverse  CCGGTCTGAACTCAGATCACGT  Palumbi et al. (1991)  H3  PCR amplification and sequencing  H3F  Forward  ATGGCTCGTACCAAGCAGACVGC  Colgan et al. (1998)  H3R  Reverse  ATATCCTTRGGCATRATRGTGAC  Colgan et al. (1998)  COI  PCR amplification and sequencing  LCO1490  Forward  GGTCAACAATCATAAAGATATTGG  Folmer et al. (1994)  HCO2198  Reverse  TAAACTTCAGGGTGACCAAAAAATC  Folmer et al. (1994)  ANT  PCR amplification and sequencing  ANTGF1  Forward  GCCAACTGCATTCGGTATTTCCC  Audzijonyte & Vrijenhoek (2010)  ANTR1  Reverse  TTCATCAAMGACATRAAMCCYTC  Audzijonyte & Vrijenhoek (2010)  View Large Table 2. Information on primers used in this study Primer  Direction  Sequence 5′–3′  References  28S rRNA  PCR amplification and sequencing  D1  Forward  ACCCSCTGAAYTTAAGCAT  Colgan et al. (2003)  D3  Reverse  GACGATCGATTTGCACGTCA  Vonnemann et al. (2005)  Sequencing  D2F  Forward  CCCGTCTTGAAACACGGACCAAGG  Vonnemann et al. (2005)  C2R  Reverse  ACTCTCTCTTCAAAGTTCTTTTC  Dayrat et al. (2001)  16S rRNA  PCR amplification and sequencing  16SarL  Forward  CGCCTGTTTATCAAAAACAT  Palumbi et al. (1991)  16SbrH  Reverse  CCGGTCTGAACTCAGATCACGT  Palumbi et al. (1991)  H3  PCR amplification and sequencing  H3F  Forward  ATGGCTCGTACCAAGCAGACVGC  Colgan et al. (1998)  H3R  Reverse  ATATCCTTRGGCATRATRGTGAC  Colgan et al. (1998)  COI  PCR amplification and sequencing  LCO1490  Forward  GGTCAACAATCATAAAGATATTGG  Folmer et al. (1994)  HCO2198  Reverse  TAAACTTCAGGGTGACCAAAAAATC  Folmer et al. (1994)  ANT  PCR amplification and sequencing  ANTGF1  Forward  GCCAACTGCATTCGGTATTTCCC  Audzijonyte & Vrijenhoek (2010)  ANTR1  Reverse  TTCATCAAMGACATRAAMCCYTC  Audzijonyte & Vrijenhoek (2010)  Primer  Direction  Sequence 5′–3′  References  28S rRNA  PCR amplification and sequencing  D1  Forward  ACCCSCTGAAYTTAAGCAT  Colgan et al. (2003)  D3  Reverse  GACGATCGATTTGCACGTCA  Vonnemann et al. (2005)  Sequencing  D2F  Forward  CCCGTCTTGAAACACGGACCAAGG  Vonnemann et al. (2005)  C2R  Reverse  ACTCTCTCTTCAAAGTTCTTTTC  Dayrat et al. (2001)  16S rRNA  PCR amplification and sequencing  16SarL  Forward  CGCCTGTTTATCAAAAACAT  Palumbi et al. (1991)  16SbrH  Reverse  CCGGTCTGAACTCAGATCACGT  Palumbi et al. (1991)  H3  PCR amplification and sequencing  H3F  Forward  ATGGCTCGTACCAAGCAGACVGC  Colgan et al. (1998)  H3R  Reverse  ATATCCTTRGGCATRATRGTGAC  Colgan et al. (1998)  COI  PCR amplification and sequencing  LCO1490  Forward  GGTCAACAATCATAAAGATATTGG  Folmer et al. (1994)  HCO2198  Reverse  TAAACTTCAGGGTGACCAAAAAATC  Folmer et al. (1994)  ANT  PCR amplification and sequencing  ANTGF1  Forward  GCCAACTGCATTCGGTATTTCCC  Audzijonyte & Vrijenhoek (2010)  ANTR1  Reverse  TTCATCAAMGACATRAAMCCYTC  Audzijonyte & Vrijenhoek (2010)  View Large Phylogenetic analyses In addition to the sequences obtained in this study, we also accessed sequence data of other galeommatoideans and outgroups from GenBank (Table 1). Sequences of the 28S and 16S genes were aligned using the Muscle program (Edgar, 2004) with default settings in the software Seaview (Galtier, Gouy & Gautier, 1996; Gouy, Guindon & Gascuel, 2010). We employed Gblocks v0.91b (Castresana, 2000; Talavera & Castresana, 2007) to eliminate the ambiguously aligned regions in the 28S and 16S genes. The sizes of 28S and 16S sequences prior to treatment with Gblocks were 1042 and 481 bp, respectively, whereas those after Gblocks treatment were 1032 and 343 bp, respectively. Phylogenetic trees were constructed using Bayesian and maximum likelihood (ML) methods. Bayesian analyses were performed using MrBayes 3.1.2 (Ronquist & Huelsenbeck, 2003) with substitution models chosen by Kakusan 4 (Tanabe, 2011). In the combined data set, substitution parameters were estimated separately for each gene partition [28S: GTR + Gamma, 16S: HKY85 + Gamma, COI: HKY85 + Gamma, GTR + Gamma, and F81 + Homogeneous (for each codon partition), H3: GTR + Gamma, K80 + Homogeneous, and JC69 + Homogeneous (for each codon partition), ANT: HKY85 + Gamma, F81 + Gamma, and JC69 + Homogeneous (for each codon partition)]. Two independent Metropolis-coupled Markov chain Monte Carlo runs were carried out simultaneously, sampling trees every 100 generations and calculating the average standard deviation of split frequencies (ASDSFs) every 1000 generations. Analyses were continued until ASDSF dropped below 0.01, at which point the two chains were considered to have achieved convergence. Because ASDSF was calculated based on the last 75% of the samples, we discarded the initial 25% of the sampled trees as burn-in. We confirmed that analyses reached stationarity well before the burn-in period by plotting the ln-likelihood of the sampled trees against generation time. ML analyses were performed using RAxML (Stamatakis, 2006) as implemented in raxmlGUI 1.31 (Silvestro & Michalak, 2012). The robustness of the ML tree was evaluated based on 1000 bootstrap replications. Datasets were partitioned by gene and the GTR + GAMMA model was implemented. RESULTS Observation of living Parabornia squillina Three individuals of P. squillina were collected from one male individual of L. scabricauda (Fig. 2). Each individual was attached by byssal threads to the host abdomen, specifically the lateral portion of the pleonal sternite (Fig. 2E, F). Two individuals were found between the 1st and 2nd pleopods, and one was found between 2nd and 3rd pleopods. We detached the bivalves from the host to observe the extension of the foot and mantle in the living state. The bivalves have numerous short papillae extended along the ventral and posterior–dorsal margins (Fig. 2A–C). One pair of longer papillae was observed anterodorsally (Fig. 2A). The clams were placed with their host in an aquarium to test if they would reattach after removal (Movie S2). They directly approached the host by crawling, and once below the host pleopods, each clam waved its foot upward towards the host (Fig. 2G). Once their foot touched the host pleon, the bivalves attached using newly secreted byssal threads. The bivalves then crawled across the host until they reached their original position on the lateral portion of the pleonal sternite. Figure 2. View largeDownload slide Parabornia squillina and its host Lysiosquilla scabricauda. A–C. A crawling individual of P. squillina (A, lateral side; B, dorsal view; C, ventral view). D. L. scabricauda. E and F. P. squillina attached to the lateral portion of the pleonal sternite. G. P. squillina extending its foot to attach to the host pleon. Arrows indicate the heel of P. squillina without posterior extension or hanging foot structure (A) and P. squillina (E–G). Figure 2. View largeDownload slide Parabornia squillina and its host Lysiosquilla scabricauda. A–C. A crawling individual of P. squillina (A, lateral side; B, dorsal view; C, ventral view). D. L. scabricauda. E and F. P. squillina attached to the lateral portion of the pleonal sternite. G. P. squillina extending its foot to attach to the host pleon. Arrows indicate the heel of P. squillina without posterior extension or hanging foot structure (A) and P. squillina (E–G). Molecular phylogenetic analyses Our results suggest that Vasconiellinae is monophyletic [Bayesian posterior probability (PP) = 1.00, bootstrap percentage (BS) = 92] (Fig. 3). Divariscintilla aff. maoria Powell, 1932 was sister to all of the remaining vasconiellines (PP = 1.00, BS = 80), including the other species of Divariscintilla, Phlyctaenachlamys, Ephippodontomorpha and Parabornia. The ectocommensal P. squillina was nested within the burrow-wall-commensal vasconiellines and was sister to D. octotentaculata (Fig. 3). Divariscintilla yoyo included one cryptic sister species (D. aff. yoyo) (Fig. 3). Floridian vasconiellines were not monophyletic; D. troglodytes was sister to a clade of Pacific and Floridian species, whereas all of the other Floridian taxa formed a crown clade that was well supported in Bayesian (PP = 0.99) but not in ML (BS = 28) phylogenetic analyses. Figure 3. View largeDownload slide Bayesian phylogenetic tree of yoyo clams (Galeommatoidea: Galeommatidae: Vasconiellinae) based on the combined data set of 28S, 16S, H3, COI and ANT genes. Numbers above branches indicate Bayesian posterior probabilities followed by maximum likelihood bootstrap support values. Six species collected from Florida are associated with Lysiosquilla scabricauda and Divariscintilla toyohiwakensis in Japan is associated with Bigelowina phalangium, whereas the other species were collected from mantis-shrimp burrows but the host species were not identified. Abbreviation: MNHN, Muséum National d’Histoire Naturelle. Figure 3. View largeDownload slide Bayesian phylogenetic tree of yoyo clams (Galeommatoidea: Galeommatidae: Vasconiellinae) based on the combined data set of 28S, 16S, H3, COI and ANT genes. Numbers above branches indicate Bayesian posterior probabilities followed by maximum likelihood bootstrap support values. Six species collected from Florida are associated with Lysiosquilla scabricauda and Divariscintilla toyohiwakensis in Japan is associated with Bigelowina phalangium, whereas the other species were collected from mantis-shrimp burrows but the host species were not identified. Abbreviation: MNHN, Muséum National d’Histoire Naturelle. DISCUSSION Within-host speciation in Floridian yoyo clams Our analysis discovered one previously unknown cryptic species (D. aff. yoyo) that is sister to D. yoyo (Fig. 3). They differed by 14.8% in their mitochondrial COI gene sequences, which is much higher than intraspecific variation levels reported for galeommatoideans [e.g. ~2% in Sato et al. (2011); ~5% in Li & Ó Foighil (2012)], or in our preliminary results for these two taxa [1.5% in D. yoyo (N = 2) and 0–0.2% in D. aff. yoyo (N = 3)] (unpublished data). They are superficially identical in external appearance but can be morphologically distinguished by their shell outlines: an angulate anterior shell margin is present in D. yoyo but not in D. aff. yoyo (our unpublished data). This means that in Florida, L. scabricauda hosts no fewer than seven vasconielline species including six burrow-wall-commensal species (Divariscintilla spp.) and one ectocommensal species (P. squillina). Our phylogenetic analyses included six Floridian vasconiellines except for D. cordiformis. Bayesian analyses suggested that Floridian vasconiellines are not monophyletic but are divided into two groups: D. troglodytes and the remaining five species (Fig. 3). The monophyly of five Floridian vasconielline species, except for D. troglodytes (Fig. 3), was supported by Bayesian posterior probabilities, suggesting that the diversity of Floridian vasconiellines is caused both by secondary contact of a distantly related linage (D. troglodytes) and by local diversification. However, bootstrap values supporting this topology are low (Fig. 3). Thus, a molecular analysis with more genetic data should be conducted in the future. Our phylogenetic analyses identified two sister-group pairs among Floridian yoyo clams: (1) D. octotentaculata and P. squillina, and (2) D. yoyo and D. aff. yoyo. In Florida, all of these species use a single host, L. scabricauda (Mikkelsen & Bieler, 1989, 1992; this study), suggesting that within-host speciation may have occurred in these two cases. Interestingly, these sister-group pairs have contrasting characteristics. Divariscintilla octotentaculata and P. squillina are ecologically and morphologically quite distinct. The former lives on host burrow walls, whereas the latter lives on the host body surface. There are differences in morphological characteristics between these two species as well, possibly corresponding to differences in host use patterns (see details below). Ecological shifts associated with host use mode probably played a key role in speciation events and led to dramatic morphological change. Divariscintilla yoyo and D. aff. yoyo, by contrast, are ecologically and morphologically very similar: both live on the host’s burrow walls and have two elongated anterior tentacle pairs (Fig. 1A, C). Lastly, an ecological shift is not apparent in this speciation event. Sympatrically distributed sister species are common among marine benthic invertebrates (Knowlton, 1993) and stem from either sympatric ecological speciation or allopatric speciation with subsequent range expansion and secondary contact (Bowen et al., 2013). These two mechanisms can be difficult to distinguish based upon existing patterns, and it is unclear whether sympatric speciation occurred in our two Floridian sister-group pairs (Fig. 3). If D. yoyo and D. aff. yoyo are not ecologically differentiated, it may be more likely that they speciated in allopatry prior to secondary contact. While Floridian Divariscintilla species have been recorded only from the Indian River Lagoon and areas nearby (Mikkelsen & Bieler, 1992; Mikkelsen, Mikkelsen & Karlen, 1995), this may be due to insufficient sampling (Mikkelsen & Bieler, 1992). Considering that L. scabricauda is distributed broadly from the Atlantic coast of the United States to Brazil (Reaka et al., 2009), allopatric speciation of yoyo clams within the host distribution range is plausible. To explore this question, further investigation of the distribution of each vasconielline species is necessary. Divariscintilla species are simultaneous hermaphrodites that brood their young to a straight-hinge ‘D’ veliger stage in the suprabranchial chamber as well as in the outer demibranch, and then release them to the water column through their exhalant siphon (Judd, 1971; Mikkelsen & Bieler, 1989, 1992; Yamashita et al., 2011). It remains unknown how long the planktotrophic larval stage endures in these species prior to metamorphosis and settlement in the host burrows. The shorter the duration of the planktonic stage for sedentary or sessile marine invertebrates, the lower the rate of gene flow among discontinuously distributed populations, and the greater the probability of allopatric speciation (but see Weersing & Toonen, 2009). Reproductive isolation mechanisms have been often considered to be necessary for the maintenance of coexistence of closely related species, because otherwise, hybridization may lead to the breakdown of species boundaries (Shine et al., 2002; Muthiga, 2003). Thus, how Floridian yoyo clams achieve reproductive isolation among co-occurring species is an intriguing question. Mikkelsen & Bieler (1992) observed an interesting copulatory-like behaviour in D. yoyo and D. octotentaculata. If this behaviour is actually copulatory in function and common in Divariscintilla species, it may allow them to engage in species-specific selective mating that can prevent interspecific hybridization. Ectocommensal lifestyle of Parabornia The genus Parabornia comprises two species, P. squillina and P. palliopapillata Simone, 2001 (Boss, 1965b; Simone, 2001). They are very similar in morphology (Simone, 2001) and both are ectocommensal on the same host, L. scabricauda (Boss, 1965b; Mikkelsen & Bieler, 1992; Simone, 2001). The former is distributed from Florida to Panama (Boss, 1965b; Moore & Boss, 1966; Mikkelsen & Bieler, 1992), whereas the latter is known only from Brazilian coasts (Simone, 2001). Previous studies briefly describe P. squillina as attached to the inner surface of the abdominal sclera of L. scabricauda (Mikkelsen & Bieler, 1992), whereas Simone (2001) mentioned that the other species, P. palliopapillata, lives attached to the pleopod base of the host. In this study, we found P. squillina attached to the lateral portion of the pleonal sternite of the host (Fig. 2). This is consistent with previous descriptions of P. squillina and P. palliopapillana, and suggests that these two species use the host in the same way. Simone (2001) mentioned that young individuals of P. palliopapillata occur on maxilipedal bases and under the carapace. This may be characteristic of P. squillina as well, although it was not confirmed in this study. During behavioural trials, P. squillina actively moved back to the lateral portion of the pleonal sternite after being detached from the host (Movie S2), suggesting that this species has a strong habitat preference for a specific part of the host abdomen. Habitat preference for a specific part of the host abdomen is common among galeommatoideans that are ectocommensal with mantis shrimp and mud shrimp (Morton, 1972; Ó Foighil, 1985; Kato & Itani, 1995), but these shared preferences in microhabitat are the result of convergent evolution (Goto et al., 2012). Gage (1968) suggested that some galeommatoideans detect hosts by using chemicals emitted from the host. We found that P. squillina can home back to a specific part of host body when it is detached from the host. Thus, it is probable that P. squillina recognizes L. scabricauda based on chemotaxis to host-emitted chemicals. Most members of Vasconiellinae, including Parabornia, have a flower-like organ near the foot, which is suggested to be a receptor of host chemicals (Mikkelsen & Bieler, 1989, 1992; Middelfart, 2005). Additionally, we found that this bivalve has numerous short papillae that occur densely along the ventral and dorsal edges of the mantle, and the former directly touch the host abdominal body surface (Fig. 2A–C). Such papillae are not known in other burrow-wall-commensal yoyo clams (Fig. 1). It is probable that these papillae have a sensory function, and that P. squillina uses them to locate its preferred position on the host body. Habitat shift from host burrow wall to host body surface Ectosymbionts that live on the body surface of burrowing invertebrates have evolved in various marine invertebrate lineages (Funch & Christensen, 1995; Kobayashi & Kato, 2003). However, the evolutionary processes that produce an ectosymbiotic lifestyle are not well understood. In this study, we show that ectocommensal Parabornia evolved from burrow-wall-commensal ancestors (Fig. 3), indicating that the burrow-wall-commensal lifestyle was an evolutionary stepping stone for an ectocommensal lifestyle in this case. Other than the genus Parabornia, ectocommensalism has evolved multiple times in Galeommatoidea (Goto et al., 2012). Evolutionary transitions from free-living to commensal lifestyles have occurred multiple times in Galeommatoidea, most of which are transitions from a free-living to burrow-wall-commensal lifestyle (Goto et al., 2012; Li et al., 2016). However, transitions from a free-living to an ectocommensal lifestyle have not previously been reported (Goto et al., 2012; Li et al., 2016), indicating that a burrow-wall-commensal lifestyle may be a prerequisite to attaining an ectocommensal lifestyle in these clams. Future in-depth phylogenetic studies of commensal galeommatoideans are likely to uncover additional cases of such evolutionary transitions. Competition for limited resources is frequently recognized as a selective pressure that promotes habitat shifts (Schluter, 2000; Munday et al., 2004; Losos, 2011; Hurt et al., 2013). However, whether resource competition has influenced the habitat shift from host burrow wall to host body surface in P. squillina is unclear, and based on the evidence to date, P. squillina and burrow-wall-commensal species do not co-occur (Mikkelsen & Bieler, 1992). While more evidence is needed, evolving an ectocommensal lifestyle may benefit P. squillina in several ways. For instance, in the case of burrow abandonment of one or both host shrimps, an ectocommensal can move to a new burrow with its host, although lysiosquillids may stay in the same burrow in monogamous pairs for up to 15 years (R. L. Caldwell, pers. comm.), making it unclear how large a factor this is in the ecology of P. squillina. Attachment to the host body may also add another level of protection. Parabornia squillina are hidden within the host’s pleopods (Fig. 2), whereas burrow-wall-commensals may be exposed to small predators accessing the burrow. Additionally, attachment to the host may provide additional food resources and a more consistent flow of oxygenated water due to the constant movement of the pleopods (Movie S2), as known in the galeommatoidean Borniopsis subsinuata ectocommensal with mantis shrimps (Morton, 1981). This positioning could be particularly useful when the burrow opening is capped during moulting or during low tides. However, there are potential disadvantages to the ectocommensal lifestyle including predation on the host, especially when outside of its burrow, being lethal to the P. squillina and the requirement for P. squillina to successfully reattach to the host after moulting events, as known in the galeommatoidean Peregrinamor ectocommensal with mud shrimps (Itani, Kato & Shirayama, 2002). Further research is required to evaluate the relative importance of these factors. Morphological changes associated with ecological shift Our results show that ectocommensal Parabornia evolved from burrow-wall-commensal ancestors (Fig. 3). The morphologies of P. squillina and Divariscintilla species have been well described in previous studies (Boss, 1965b; Mikkelsen & Bieler, 1989, 1992; Simone, 2001). By comparing their morphological characters, we detected four major morphological changes associated with the ecological shift in Parabornia: (1) loss of the thread-like posterior foot extension and associated hanging behaviour, (2) loss of covering of the shell by mantle tissue (i.e. shell externalization), (3) loss of developed sensory tentacles, and (4) acquisition of dense papillae along the mantle margin. We discuss these in detail below. Many members of Vasconiellinae, including Divariscintilla, Phlyctaenachlamys and Ephippodontomorpha, have a specialized foot with a thread-like posterior extension (Judd, 1971; Mikkelsen & Bieler, 1989, 1992; Middelfart, 2005; Yamashita et al., 2011; Fig. 1). This specialized foot, associated with the yoyo motion, is only known in Vasconiellinae and is suggested to be an adaptation to life on mantis shrimp burrow walls (Popham, 1939; Mikkelsen & Bieler, 1989). On the other hand, P. squillina has a more typical galeommatoidean foot lacking a thread-like posterior extension (Mikkelsen & Bieler, 1992; this study; Fig. 2A). Additionally, Parabornia has never been observed hanging from vertical wall surfaces or the host body in aquaria nor engaging in yoyo saltatory behaviour typical of Divariscintilla (R. Goto & T. A. Harrison, personal observations). While extreme morphological specialization of the foot has been reported in some bivalves (e.g. Dufour & Felbeck, 2003), significant morphological change in bivalve foot structure associated with a microhabitat shift is documented here for the first time. Burrow-wall-commensal yoyo clams have reduced shells fully or partially internalized by mantle tissue bearing highly developed tentacles (Popham, 1939; Judd, 1971; Mikkelsen & Bieler, 1989, 1992; Middelfart, 2005) (Fig. 1). Parabornia squillina has an externalized shell as is typical in most bivalve species (Boss, 1965b; Simone, 2001; Fig. 2A). Unlike typical bivalves, burrow-wall-commensal yoyo clams actively crawl on the burrow wall surfaces using their foot. The soft mantle and tentacles are probably useful in perceiving the surrounding environment during crawling behaviour as well as in perceiving and reacting to host movement within the burrow (Judd, 1971). Unlike burrow-wall-commensal yoyo clams, Parabornia spp. are basically sessile on the host body surface (Fig. 2E, F), probably reducing the necessity for sensory and defensive structures and hence the loss of a mantle shell covering, although P. squillina does have numerous short mantle papillae (Fig. 2), which touch the host body surface. Our results suggest that evolution from (semi)internalized to externalized shells occurred as a consequence of a change in microhabitat. Evolution of shell internalization is known in several molluscan lineages (e.g. Oposthobranchia and cephalopods) (Wägele & Klussmann-Kolb, 2005; Tanner et al., 2017). However, as far as we know, the evolution of shell externalization in molluscs has not been previously reported. The internalized shells, developed tentacles and specialized foot of burrow-wall-commensal yoyo clams are considered adaptations to the unique habitat of living on walls of mantis shrimp burrows (Popham, 1939; Mikkelsen & Bieler, 1989, 1992; this study). Our study suggests that these specialized morphological traits are lost as a consequence of colonization of the host body surface. Host shifts and subsequent specialization to different host taxa have been considered drivers of morphological evolution in Galeommatoidea (Goto et al., 2012; Li et al., 2016). Our study suggests that a microhabitat shift within a single host can also lead to significant morphological change. Goto et al. (2014) found that sister galeommatoidean species ectocommensal on Lingula brachiopods have significantly different shell shapes (elongated triangular shape vs. ovate shape) and suggested that this difference is due to adaptations for different posture on the host body. Goto et al. (2014) and the present study suggest that not only a host shift but also a ecological shift in association with the same host can play an important role in morphological evolution. However, burrow-wall-commensal yoyo clams show great morphological diversity, especially in their number of tentacles (Mikkelsen & Bieler, 1989, 1992; Fig. 1). The degree and functional significance of this tentacle diversity remains unknown. To answer this question, further examination of ecological differences among these species (e.g. niche partitioning within the host burrow) and of tentacle function is required. Taxonomic implications and remaining issues in Vasconiellinae Li et al. (2016) found that the genera Divariscintilla, Ephippodontomorpha and Phlyctaenachlamys formed a clade although their inter-relationships were not fully resolved. Our molecular analysis based on five genes showed that Ephippodontmorpha, Phlyctaenachlamys and Parabornia are nested within Divariscintilla (Fig. 3). According to Huber (2015), these genera are assigned to the same subfamily Vasconiellinae. The other four genera within this subfamily (i.e. Vasconiella, Bellascintilla, Ceratobornia and Aclistothyra) were not included in the present analysis. Ceratobornia also has hanging foot structure, but lives attached to the burrow walls of ghost shrimp on western Atlantic coasts (Dall, 1899; Narchi, 1966). Morphologically similar species with different hosts imply that host shifts have occurred in Vasconiellinae, although the hanging foot structure may have evolved multiple times in this group. Lastly, our results show that D. aff. maoria is sister to the remaining vasconiellines. Divariscintilla aff. maoria is distinguished from the other species used in this study in having a notch in the ventral side of shells. A ventral shell notch is also known in some other vasconiellines (i.e. Vasconiella, Bellascintilla and D. cordiformis) (Mikkelsen & Bieler, 1992; Huber, 2015). It is possible that Vasconiellinae may prove to be separable into two major groups discernible by the presence or absence of a notch in the ventral side of shells. To resolve these remaining issues, a further molecular analysis based on more taxon sampling is required. SUPPORTING INFORMATION Additional Supporting Information may be found in the online version of this article at the publisher’s web-site. Movie S1. Hanging behaviour of Divariscintilla yoyo. Movie S2. Parabornia squillina moving back to the abdomen of the mantis shrimp Lysiosquilla scabricauda. ACKNOWLEDGMENTS We thank Sherry Reed, Michael J. Boyle, William (Woody) Lee M.B.S. and David R. Branson (Smithsonian Marine Station at Fort Pierce) for helping to collect the specimens, Taehwan Lee (University of Michigan) for organizing the specimens used for this study, Jingchun Li (University of Colorado Boulder) for providing information on Galeommatoidea, Philippe Bouchet (Muséum National d’Histoire Naturelle, Paris), Gustav Paulay (Museum of Natural History, University of Florida) and Bruce Marshall (Museum of New Zealand, Te Papa Togarewa) for allowing us to use their museum specimens in this study, Arthur Anker (Universidade Federal de Goiás), Roy L. Caldwell (University of California, Berkeley), Maya S. deVries (University of California, San Diego) and Gyo Itani (Kochi University) for advice, and Paula M. Mikkelsen (Cornell University) and two anonymous referees for comments that improved the manuscript. This study was partially supported by an Overseas Research Fellowship grant (27-186) and a KAKENHI grant (17H06795) to R.G. from the Japan Society for the Promotion of Science and a Smithsonian Minority Awards Program grant to T.H. REFERENCES Althoff DM. 2014. Shift in egg-laying strategy to avoid plant defense leads to reproductive isolation in mutualistic and cheating yucca moths. Evolution  68: 301– 307. Google Scholar CrossRef Search ADS   Audzijonyte A, Vrijenhoek RC. 2010. Three nuclear genes for phylogenetic, SNP and population genetic studies of molluscs and other invertebrates. Molecular Ecology Resources  10: 200– 204. Google Scholar CrossRef Search ADS   Bieler R, Mikkelsen PM. 1992. Preliminary phylogenetic analysis of the bivalve family Galeommartidae. American Malacological Bulletin  9: 157– 164. Boss KJ. 1965a. Symbiotic erycinacean bivalves. Malacologia  3: 183– 195. Boss KJ. 1965b. A new mollusk (Bivalvia, Erycinidae) commensal on the stomatopod crustacean Lysiosquilla. American Museum Novitates  2215: 1– 11. Bouchet P, Lozouet P, Maestrati P, Heros V. 2002. Assessing the magnitude of species richness in tropical marine environments: exceptionally high numbers of molluscs at a New Caledonia site. Biological Journal of the Linnean Society  75: 421– 436. Google Scholar CrossRef Search ADS   Bowen BW, Rocha LA, Toonen RJ, Karl SA ; ToBo Laboratory. 2013. The origins of tropical marine biodiversity. Trends in Ecology & Evolution  28: 359– 366. Google Scholar CrossRef Search ADS   Castresana J. 2000. Selection of conserved blocks from multiple alignments for their use in phylogenetic analysis. Molecular Biology and Evolution  17: 540– 552. Google Scholar CrossRef Search ADS   Colgan DJ, McLauchlan A, Wilson GDF, Livingston S, Macaranas J, Edgecombe GD, Cassis G, Gray MR. 1998. Molecular phylogenetics of the Arthropoda: relationships based on histone H3 and U2 snRNA DNA sequences. Australian Journal of Zoology  46: 419– 437. Google Scholar CrossRef Search ADS   Colgan DJ, Ponder WF, Beacham E, Macaranas JM. 2003. Gastropod phylogeny based on six segments from four genes representing coding or non-coding and mitochondrial or nuclear DNA. Molluscan Research  23: 123– 148. Google Scholar CrossRef Search ADS   Cook JM, Rokas A, Pagel M, Stone GN. 2002. Evolutionary shifts between host oak sections and host-plant organs in Andricus gallwasps. Evolution  56: 1821– 1830. Google Scholar CrossRef Search ADS   Coyne JA, Orr HA. 2004. Speciation . Sunderland: Sinauer Associates. Dall WH. 1899. Synopsis of the recent and Tertiary Leptonacea of North America and the West Indies. Proceedings of the United States National Museum  21: 873– 897. Google Scholar CrossRef Search ADS   Dayrat B, Tillier A, Lecointre G, Tillier S. 2001. New clades of euthyneuran gastropods (Mollusca) from 28S rRNA sequences. Molecular Phylogenetics and Evolution  19: 225– 235. Google Scholar CrossRef Search ADS   Duffy JE. 1996. Resource-associated population subdivision in a symbiotic coral-reef shrimp. Evolution  50: 360– 373. Google Scholar CrossRef Search ADS   Dufour SC, Felbeck H. 2003. Sulphide mining by the superextensile foot of symbiotic thyasirid bivalves. Nature  426: 65– 67. Google Scholar CrossRef Search ADS   Edgar RC. 2004. MUSCLE: multiple sequence alignment with high accuracy and high throughput. Nucleic Acids Research  32: 1792– 1797. Google Scholar CrossRef Search ADS   Faucci A, Toonen RJ, Hadfield MG. 2007. Host shift and speciation in a coral-feeding nudibranch. Proceedings of the Royal Society B: Biological Sciences  274: 111– 119. Google Scholar CrossRef Search ADS   Folmer O, Black M, Hoeh W, Lutz R, Vrijenhoek R. 1994. DNA primers for amplification of mitochondrial cytochrome c oxidase subunit I from diverse metazoan invertebrates. Molecular Marine Biology and Biotechnology  3: 294– 299. Funch P, Christensen RM. 1995. Cycliophora is a new phylum with affinities to Entoprocta and Ectoprocta. Nature  378: 711– 714. Google Scholar CrossRef Search ADS   Gage J. 1968. The mode of life of Montacuta elevata, a bivalve ‘commensal’ with Clymenella torquata (Polychaeta). Canadian Journal of Zoology  46: 877– 892. Google Scholar CrossRef Search ADS   Galtier N, Gouy M, Gautier C. 1996. SEAVIEW and PHYLO_WIN: two graphic tools for sequence alignment and molecular phylogeny. Computer Applications in the Biosciences  12: 543– 548. Goto R, Kato M. 2012. Geographic mosaic of mutually exclusive dominance of obligate commensals in symbiotic communities associated with a burrowing echiuran worm. Marine Biology  159: 319– 330. Google Scholar CrossRef Search ADS   Goto R, Kawakita A, Ishikawa H, Hamamura Y, Kato M. 2012. Molecular phylogeny of the bivalve superfamily Galeommatoidea (Heterodonta, Veneroida) reveals dynamic evolution of symbiotic lifestyle and interphylum host switching. BMC Evolutionary Biology  12: 172. Google Scholar CrossRef Search ADS   Goto R, Ishikawa H, Hamamura Y. 2016. The enigmatic bivalve genus Paramya (Myoidea: Myidae): symbiotic association of an East Asian species with spoon worms (Echiura) and its transfer to the family Basterotiidae (Galeommatoidea). Journal of the Marine Biological Association of the United Kingdom  97: 1447– 1454. Google Scholar CrossRef Search ADS   Goto R, Ishikawa H, Hamamura Y, Sato S, Kato M. 2014. Evolution of symbiosis with Lingula (Brachiopoda) in the bivalve superfamily Galeommatoidea (Heterodonta), with description of a new species of Koreamya. Journal of Molluscan Studies  80: 148– 160 Google Scholar CrossRef Search ADS   Gouy M, Guindon S, Gascuel O. 2010. SeaView version 4: a multiplatform graphical user interface for sequence alignment and phylogenetic tree building. Molecular Biology and Evolution  27: 221– 224. Google Scholar CrossRef Search ADS   Huber M. 2015. Compendium of Bivalves 2. A full-color guide to the remaining seven families. A systematic listing of 8500 bivalve species and 10500 synonyms . Harxheim: ConchBooks. Hurt C, Silliman K, Anker A, Knowlton N. 2013. Ecological speciation in anemone-associated snapping shrimps (Alpheus armatus species complex). Molecular Ecology  22: 4532– 4548. Google Scholar CrossRef Search ADS   Imada Y, Kawakita A, Kato M. 2011. Allopatric distribution and diversification without niche shift in a bryophyte-feeding basal moth lineage (Lepidoptera: Micropterigidae). Proceedings of the Royal Society B: Biological Sciences  278: 3026– 3033. Google Scholar CrossRef Search ADS   Itani G, Kato M, Shirayama Y. 2002. Behaviour of the shrimp ectosymbionts, Peregrinamor ohshimai (Mollusca: Bivalvia) and Phyllodurus sp. (Crustacea: Isopoda) through ecdyses. Journal of Marine Biological Association of the United Kingdom  82: 69– 78. Jahner JP, Forister ML, Parchman TL, Smilanich AM, Miller JS, Wilson JS, Walla TR, Tepe EJ, Richards LA, Quijano-Abril MA, Glassmire AE, Dyer LA. 2017. Host conservatism, geography, and elevation in the evolution of a Neotropical moth radiation. Evolution  71: 2885– 2900. Google Scholar CrossRef Search ADS   Joy JB, Crespi BJ. 2007. Adaptive radiation of gall-inducing insects within a single host-plant species. Evolution  61: 784– 795. Google Scholar CrossRef Search ADS   Judd W. 1971. The structure and habits of Divariscintilla maoria Powell (Bivalvia: Galeommatidae). Proceedings of the Malacological Society of London  39: 343– 354. Kato M, Itani G. 1995. Commensalism of a bivalve, Peregrinamor ohshimai, with a thalasseinidean burrowing shrimp, Upogebia major. Journal of Marine Biological Association of the United Kingdom  75: 941– 947. Google Scholar CrossRef Search ADS   Knowlton N. 1993. Sibling species in the sea. Annual Reviews of Ecology and Systematics  24: 189– 216. Google Scholar CrossRef Search ADS   Kobayashi C, Kato M. 2003. Sex-biased ectosymbiosis of a unique cirripede, Octolamis unguisiformis sp. nov., that resembles the chelipeds of its host crab, Macrophthalmus milloti. Journal of Marine Biological Association of the United Kingdom  83: 925– 930. Google Scholar CrossRef Search ADS   Li J, Ó Foighil D. 2012. Host-specific morphologies but no host races in the commensal bivalve Nearomya rugifera. Invertebrate Biology  3: 197– 203. Google Scholar CrossRef Search ADS   Li J, Ó Foighil D, Middelfart P. 2012. The evolutionary ecology of biotic association in a megadiverse bivalve superfamily: sponsorship required for permanent residency in sediment. PLoS One  7: e42121. Google Scholar CrossRef Search ADS   Li J, Ó Foighil D, Strong E. 2016. Commensal associations and benthic habitats shape macroevolution of the bivalve clade Galeommatoidea. Proceedings of the Royal Society B: Biological Sciences  283: 20161006. Google Scholar CrossRef Search ADS   Losos JB. 2011. Lizards in an evolutionary tree: ecology and adaptive radiation of anoles . Berkeley: University of California Press. Matsubayashi KW, Ohshima I, Nosil P. 2010. Ecological speciation in phytophagous insects. Entomologia Experimentalis et Applicata  134: 1– 27. Google Scholar CrossRef Search ADS   Middelfart P. 2005. Review of Ephippodonta sensu lato (Galeommatidae: Bivalvia), with descriptions of new related genera and species from Australia. Molluscan Research  25: 129– 144. Mikkelsen PM, Bieler R. 1989. Biology and comparative anatomy of Divariscintilla yoyo and D. troglodytes, two new species of Galeommatidae (Bivalvia) from stomatopod burrows in eastern Florida. Malacologia  31: 175– 195. Mikkelsen PM, Bieler R. 1992. Biology and comparative anatomy of three new species of commensal Galeommatidae, with a possible case of mating behavior in bivalves. Malacologia  34: 1– 24. Mikkelsen PM, Mikkelsen PS, Karlen DJ. 1995. Molluscan biodiversity in the Indian River Lagoon, Florida. Bulletin of Marine Science  57: 94– 127. Moore DR, Boss KJ. 1966. Records for Parabornia squillina. The Nautilus  80: 34– 35. Moran NA. 2006. Symbiosis. Current Biology  16: R866– R871. Google Scholar CrossRef Search ADS   Morton B. 1972. Some aspects of the functional morphology and biology of Pseudopythina subsinuata (Bivalvia; Leptonacea) commensal on stomatopod crustaceans. Journal of Zoology, London  166: 79– 96. Google Scholar CrossRef Search ADS   Morton B. 1981. The biology and functional morphology of Chlamydoconcha orcutii with a discussion on the taxonomic status of the Chlamydoconchacea (Mollusca: Bivalvia). Journal of Zoology, London  195: 81– 121. Google Scholar CrossRef Search ADS   Morton B, Scott PH. 1989. Hong Kong Galeommatacea (Mollusca: Bivalvia) and their hosts, with descriptions of new species. Asian Marine Biology  6: 129– 160. Muthiga NA. 2003. Coexistence and reproductive isolation of the sympatric echinoids Diadema savignyi Michelin and Diadema setosum (Leske) on Kenyan coral reefs. Marine Biology  143: 669– 677. Google Scholar CrossRef Search ADS   Munday PL, van Herwerden L, Dudgeon CL. 2004. Evidence for sympatric speciation by host shift in the sea. Current Biology  14: 1498– 1504. Google Scholar CrossRef Search ADS   Nakadai R, Kawakita A. 2016. Phylogenetic test of speciation by host shift in leaf cone moths (Caloptilia) feeding on maples (Acer). Ecology and Evolution  6: 4958– 4970. Google Scholar CrossRef Search ADS   Narchi W. 1966. The functional morphology of Ceratobornia cema, new species of the Erycinacea (Mollusca, Eulamellibranchiata). Amaos da Academia Brasileira de Ciências  38: 513– 524. Ó Foighil D. 1985. Form, function, and origin of temporary dwarf males in Pseudopythina rugifera (Carpenter, 1864) (Bivalvia: Galeommatacea). The Veliger  27: 245– 252. Palumbi S, Martin A, Romano S, McMillan WO, Stice L, Grabowski G. 1991. The simple fool’s guide to PCR, Version 2.0 . Honolulu: Department of Zoology and Kewalo Marine Laboratory, University of Hawaii. Paulay G. 2003. Marine Bivalvia (Mollusca) of Guam. Micronesica  35–36: 218– 243. Pérez-Tris J, Hellgren O, Krizanauskiene A, Waldenström J, Secondi J, Bonneaud C, Fjeldså J, Hasselquist D, Bensch S. 2007. Within-host speciation of malaria parasites. PLoS One  2: e235. Google Scholar CrossRef Search ADS   Popham ML. 1939. On Phlyctaenachlamys lysiosquillina gen. and sp. nov., a lamellibranch commensal in the burrows of Lysiosquilla maculata. Great Barrier Reef Expedition 1928–1929. Scientific Reports  6: 61– 84. Poulin R, Morand S. 2004. Parasite biodiversity . Washington: Smithsonian Institution Press. Reaka ML, Camp DK, Álvarez F, Gracia AG, Ortiz M, Vázquez-Bader AR. 2009. Stomatopoda (Crustacea) of the Gulf of Mexico. In: Felder DL, Camp DK, eds. Gulf of Mexico–origins, waters, and biota: biodiversity . College Station: Texas A&M University Press, 901– 921. Ronquist F, Huelsenbeck JP. 2003. MrBayes 3: Bayesian phylogenetic inference under mixed models. Bioinformatics  19: 1572– 1574. Google Scholar CrossRef Search ADS   Sato S, Owada M, Haga T, Hong J-S, Lützen J, Yamashita H. 2011. Genus-specific commensalism of the galeommatoid bivalve Koreamya arcuata (A. Adams, 1856) associated with lingulid brachiopods. Molluscan Research  31: 95– 105. Schluter D. 2000. The ecology of adaptive radiation . Oxford: Oxford University Press. Shine R, Reed RN, Shetty S, Lemaster M, Mason RT. 2002. Reproductive isolating mechanisms between two sympatric sibling species of sea snakes. Evolution  56: 1655– 1662. Google Scholar CrossRef Search ADS   Simone LRL. 2001. Revision of the genus Parabornia (Bivalvia: Galeommatoidea: Galeommatidae) from the western Atlantic, with description of a new species from Brazil. Journal of Conchology  37: 159– 169. Silvestro D, Michalak I. 2012. RaxmlGUI: a graphical front-end for RAxML. Organism Diversity and Evolution  12: 335– 337. Google Scholar CrossRef Search ADS   Stamatakis A. 2006. RAxML-VI-HPC: maximum likelihood-based phylogenetic analyses with thousands of taxa and mixed models. Bioinformatics  22: 2688– 2690. Google Scholar CrossRef Search ADS   Talavera G, Castresana J. 2007. Improvement of phylogenies after removing divergent and ambiguously aligned blocks from protein sequence alignments. Systematic Biology  56: 564– 577. Google Scholar CrossRef Search ADS   Tanabe AS. 2011. Kakusan4 and Aminosan: two programs for comparing nonpartitioned, proportional and separate models for combined molecular phylogenetic analyses of multilocus sequence data. Molecular Ecology Resources  11: 914– 921. Google Scholar CrossRef Search ADS   Tanner AR, Fuchs D, Winkelmann IE, Gilbert MTP, Pankey MS, Ribeiro ÂM, Kocot KM, Halanych KM, Oakley TH, da Fonseca RR, Pisani D, Vinther J. 2017. Molecular clocks indicate turnover and diversification of modern coleoid cephalopods during the Mesozoic Marine Revolution. Proceedings of the Royal Society B: Biological Sciences  284: 20162818. Google Scholar CrossRef Search ADS   Tsang LM, Chan BKK, Shin F-L, Chu KH, Chen CA. 2009. Host-associated speciation in the coral barnacle Wanella milleporae (Cirripedia: Pyrgomatidae) inhabiting the Millepora coral. Molecular Ecology  18: 1463– 1475. Google Scholar CrossRef Search ADS   Vanhove MPM, Hablützel PI, Pariselle A, Šimková A, Huyse T, Raeymaekers JAM. 2016. Cichlids: a host of opportunities for evolutionary parasitology. Trends in Parasitology  32: 821– 832. Google Scholar CrossRef Search ADS   Vonnemann V, Schrödl M, Klussmann-Kolb A, Wägele H. 2005. Reconstruction of the phylogeny of the Opisthobranchia (Mollusca: Gastropoda) by means of 18S and 28S rRNA gene sequences. Journal of Molluscan Studies  71: 113– 125. Google Scholar CrossRef Search ADS   Wägele H, Klussmann-Kolb A. 2005. Opisthobranchia (Mollusca, Gastropoda) – more than just slimy slugs. Shell reduction and its implications on defence and foraging. Frontiers in Zoology  2: 3. Google Scholar CrossRef Search ADS   Weersing K, Toonen RJ. 2009. Population genetics, larval dispersal, and connectivity in marine systems. Marine Ecology Progress Series  393: 1– 12. Google Scholar CrossRef Search ADS   Windsor DA. 1998. Most of the species on Earth are parasites. International Journal for Parasitology  28: 1939– 1941. Google Scholar CrossRef Search ADS   Winkler IS, Mitter C. 2008. The phylogenetic dimension of insect–plant interactions: a review of recent evidence. In: Tilmon KJ, ed. Specialization, speciation, and radiation: the evolutionary biology of herbivorous insects . Berkeley: University of California Press, 203– 215. Google Scholar CrossRef Search ADS   Yamashita H, Haga T, Lützen J. 2011. The bivalve Divariscintilla toyohiwakensis n. sp. (Heterodonta: Galeommatidae) from Japan, a commensal with a mantis shrimp. Venus  69: 123– 133. Zhang B, Segraves KA, Xue H-J, Nie R-E, Li W-Z, Yang X-K. 2015. Adaptation to different host plant ages facilitates insect divergence without a host shift. Proceedings of the Royal Society B: Biological Sciences  282: 20151649. Google Scholar CrossRef Search ADS   © 2018 The Linnean Society of London, Biological Journal of the Linnean Society This article is published and distributed under the terms of the Oxford University Press, Standard Journals Publication Model (https://academic.oup.com/journals/pages/about_us/legal/notices)

Journal

Biological Journal of the Linnean SocietyOxford University Press

Published: May 10, 2018

There are no references for this article.

You’re reading a free preview. Subscribe to read the entire article.


DeepDyve is your
personal research library

It’s your single place to instantly
discover and read the research
that matters to you.

Enjoy affordable access to
over 18 million articles from more than
15,000 peer-reviewed journals.

All for just $49/month

Explore the DeepDyve Library

Search

Query the DeepDyve database, plus search all of PubMed and Google Scholar seamlessly

Organize

Save any article or search result from DeepDyve, PubMed, and Google Scholar... all in one place.

Access

Get unlimited, online access to over 18 million full-text articles from more than 15,000 scientific journals.

Your journals are on DeepDyve

Read from thousands of the leading scholarly journals from SpringerNature, Elsevier, Wiley-Blackwell, Oxford University Press and more.

All the latest content is available, no embargo periods.

See the journals in your area

DeepDyve

Freelancer

DeepDyve

Pro

Price

FREE

$49/month
$360/year

Save searches from
Google Scholar,
PubMed

Create lists to
organize your research

Export lists, citations

Read DeepDyve articles

Abstract access only

Unlimited access to over
18 million full-text articles

Print

20 pages / month

PDF Discount

20% off